Open access

Nitrogen Fixation Outside and Inside Plant Tissues

Written By

C.P. Chanway, R. Anand and H. Yang

Submitted: 16 October 2013 Published: 29 January 2014

DOI: 10.5772/57532

From the Edited Volume

Advances in Biology and Ecology of Nitrogen Fixation

Edited by Takuji Ohyama

Chapter metrics overview

4,934 Chapter Downloads

View Full Metrics

1. Introduction

Nitrogen is one of the most important elements in biological systems, comprising the main building blocks of nucleic acids, enzymes and proteins among its multiple functions. In nature, it exists primarily in the gaseous form and constitutes approximately 78% of the atmosphere. Despite its abundance, nitrogen (N) is one of the most growth-limiting nutrients in terrestrial and aquatic ecosystems (Dalton & Krammer, 2006) because its gaseous form is inert and unusable by most living organisms except for nitrogen fixing microorgansisms. For it to become biologically available, atmospheric nitrogen must be transformed or “fixed” from its inert gaseous form (N2) to ammonia (NH3), which can then be assimilated into a variety of important biochemicals. This transformation, which requires a large amount of energy to break apart the triple-bonded N atoms that comprise gaseous N2, is called ‘nitrogen fixation’ (NF). Nitrogen is fixed naturally through energy-releasing abiotic processes such as lightening, forest fires and volcanic activity. These processes produce oxides of N in the atmosphere that subsequently dissolve in rain and descend to the ground as NH3 molecules. Approximately 12% of annual global NF is fixed in this way (Bezdicek & Kennedy, 1998). Fertilizer production using high temperatures and pressures in the Haber-Bosch process occurs widely and accounts for approximately 20% of annual global NF (Bezdicek & Kennedy, 1998). However, the process is fossil-fuel intensive and consumes 3-5% of the world’s natural gas annually (Myrold & Bottomley, 2007). Alternatively, NF occurs through the normal metabolic activity of many prokaryotic microorganisms, known as diazotrophs, through a process commonly referred to as biological nitrogen fixation (BNF). This essentially “free” process is responsible for the addition of almost all biologically available N that enters terrestrial ecosystems, some 140 million metric tons per year (Bezdicek & Kennedy, 1998; Galloway et al., 2008). Biological nitrogen fixation is an ATP-demanding process that is catalyzed by the enzyme complex known as nitrogenase, which is found in many members of the Bacteria and Archaea (Galloway et al., 2008). Terrestrial BNF occurs primarily in the soil, by either free-living diazotrophs or those associated to varying degrees with plants (see below).


2. Free-living bacteria

Free-living diazotrophic bacteria are those that do not associate with plants (cf. rhizosphere bacteria below) and are found in soils that are free from the direct influence of plant roots. These microorganisms are ubiquitous in terrestrial and aquatic environments and are physiologically very diverse (Reed et al., 2011). Since most soils are C and N limited, the amount of N2 they fix in soil is restricted by access to energy sources, i.e., substrates to generate adenosine triphosphate (ATP) and micronutrients required for the synthesis and functioning of nitrogenase (Reed et al., 2011). BNF by free-living diazotrophs is also limited by the severe oxygen sensitivity of nitrogenase (Postgate, 1998), which is a problem that has been at least partially overcome in different ways by diazotrophs participating in nitrogen-fixing symbioses (see below). Antagonistic microbial interactions such as parasitism and competition for nutrients (Cacciari et al.,1986; Bashan & Holguin, 1997; Bashan et al., 2004) further reduce the amount of N2 they fix. While the general belief is that free-living diazotrophs do not contribute large quantities of fixed N to most terrestrial ecosystems, perhaps 3-5 kg/ha/yr (Postgate, 1998; Newton, 2007), their cumulative N contributions are thought to be important in some tropical and temperate forest ecosystems (Cleveland et al., 1999; Gehring et al., 2005; Reed et al., 2007; 2008).


3. Rhizosphere bacteria

The rhizosphere is defined as soil that surrounds plant roots and is under their direct metabolic influence (Curl & Truelove, 1987). Proximity to plant roots is important for soil organisms as actively growing plant roots deposit approximately 20% of annual photosynthate in the rhizosphere (Nguyen, 2003), but depending on the type of plant and its growth stage, more than 50% of newly-fixed carbon may be deposited in the rhizosphere at any given time. The soluble carbon compounds that plants deposit through their root systems are known as root exudates. These comprise a wide range of carbon compounds (amino acids, peptides, proteins, enzymes, “growth factors”, vitamins and phytohormones) (Grayston et al., 1997; Jones et al., 2004; Shi et al., 2012) and are released continuously during the growing season through a process known as root exudation (Jones et al., 2004). This process has been shown to significantly stimulate growth and population sizes of most soil microorganisms, but effects are particularly noticeable in soil bacteria and fungi. The degree of stimulation is significant: in comparison with bulk soil not under the influence of plant roots, the rhizosphere typically supports 5-100 x larger bacterial and fungal populations than non-rhizosphere or “bulk” soil (Warembourg, 1997; Dobelaere et al., 2003). Due to their ability to fix N2, diazotrophs can have a competitive advantage over non-N2 fixing bacteria in the rhizosphere and prevail in it particularly when soil N is limited (Döbereiner & Pedrosa, 1987). In addition to stimulating their own growth, rhizosphere diazotrophs representing several genera (e.g., Acetobacter, Azoarcus, Azospirillum, Azotobacter, Beijerinckia, Burkholderia, Enterobacter, Herbaspirillum, Klebsiella, Paenibacillus and Pseudomonas) have been shown to enhance the growth of the plants that generate a suitable rhizosphere. These include agriculturally-important species such as rice, wheat, barley, potato and several vegetable crops (Dobelaere et al., 2003). While it is logical to assume diazotrophic rhizosphere bacteria enhance plant growth through BNF, these microorganisms are often capable of producing plant growth-enhancing phytohormones and pathogen-suppressing antibiotics (Chanway, 2002). Many are also able to enhance the availability of N, P and S in the rhizosphere enzymatically, rendering it difficult to conclude with certainty the mechanism by which plant growth is stimulated.


4. Phyllosphere

The leaf surface, or phyllosphere is another microsite known to be colonized by a wide range of microorganisms, including diazotrophic bacteria (Lindow & Brandl, 2003). While comparatively little work has been done on phyllosphere-colonizing diazotrophs, it is likely that their contribution to plant N nutrition is modest, owing to the energy constraints and problems associated with oxygen toxicity of nitrogenase such N2 fixing microorganisms would experience. Nevertheless, phyllosphere-colonizing diazotrophs should be evaluated further for possible contributions to plant nutrition.


5. Cyanobacterial associations

Cyanobacteria are prokaryotes belonging to the domain Bacteria that are capable of fixing carbon, through oxygenic photosynthesis, as well as nitrogen through BNF. As a result, cyanobacteria are nutritionally independent to a large degree (Meeks, 1988) and occupy diverse habitats, ranging from freshwater and oceanic ecosystems, temperate soils to extreme environments such as hot springs and deserts (Herrero et al., 2001). To counter the adverse effects of oxygen evolution from photosynthesis on BNF, diazotrophic cyanobacteria have developed strategies such as temporal separation of oxygenic photosynthesis and micro-oxic NF (Cerveny et al., 2013) and formation of thick-walled, non-photosynthetic heterocysts with microaerobic interiors that are conducive for NF (Wolk et al., 1994).

Many diazotrophic cyanobacterial species enter into symbioses with eukaryotes including phytoplankton, fungi and terrestrial plants. Cyanobacterial symbionts (cyanobionts) in these associations may contribute a significant portion of N required for growth of both organisms through BNF in N-limited aquatic and terrestrial environments (Schell & Alexander,1973; Hobara et al., 2006). In the ocean, they are frequently found in association with diatoms (Ferrario et al., 1995) and brown algae (Carpenter, 1972), while in fresh water, the cyanobiont Anabaena azollae forms a symbiotic association with water ferns belonging to the genus Azolla (Talley et al., 1977). The Azolla-Anabaena symbiosis often occurs in rice culture, where it results in enhanced rice performance due to a N fertilizer effect from BNF (Yanni, 1992). Diazotrophic cyanobacteria have also been found in the coralloid roots of cycads (Gehringer et al., 2010), auricles of liverworts (Adams & Duggan, 2008), slime cavities of hornworts (Adams & Duggan, 2008), stem glands of Gunnera (Bergman et al., 1992), hyphae of lichens (Jayasinghearachchi & Seneviratne, 2004) and on the leaf surfaces of mosses (Solheim & Zielke, 2002), where they fix and transfer N to the non-N2 fixing partner. To facilitate locating a suitable symbiotic partner, some cyanobacteria belonging to the order Nostocales can differentiate into motile segments termed hormogonia that travel chemotactically toward a potential partner (Campbell & Meeks, 1989). If the partner is suitable, hormogonia differentiate back into vegetative cells and enter into an active diazotrophic symbiosis where they fix and provide N for both partners.


6. Legume x Rhizobium symbiosis

The legume x Rhizobium symbioses is a well-known mutualism involving plants from the angiosperm family Leguminosae (synonym Fabaceae) and bacteria belonging to the family Rhizobiaceae (Postgate 1998). In this symbiosis, diazotrophic soil bacteria belonging to the genus Rhizobium (or closely related genera) seek out and infect roots of suitable legume plant hosts using a complex chemical signaling system. Bacteria then colonize certain root cortex cells and initiate formation of a new plant organ, the root nodule. Bacteria proliferate within root nodule cells and then differentiate into a nitrogen fixing form called a bacteroid, to fix N2. The plant vascular system is continuous with that of the root nodule, which enables newly fixed N to be rapidly translocated to other parts of the plant most in need of N. Energy for BNF is provided to bacteroids from plant photosynthesis and the oxygen concentration in root nodule cells is tightly regulated by an iron containing protein very similar in composition to hemoglobin called leghemoglobin. Bacteroids in this coadapted symbiosis are capable of high rates of BNF, i.e., up to 600 kg/ha/yr, particularly when compared to BNF in the rhizosphere (15-25 kg/ha/yr) or by free-living diazotrophs (3-5 kg/ha/yr).

The legume x Rhizobium symbiosis has been studied widely from ecological, agronomic and molecular biological perspectives to not only enhance the nitrogen-fixing efficacy of existing symbioses but to determine if similar associations might also be developed with non-legume crops (Oldroyd et al., 2011, Udvardi & Poole, 2013). Notwithstanding impressive gains in our understanding of this symbiosis, its inherent complexity currently precludes our ability to extend effective nodulation or nitrogen fixing capacity to non-legume species (Beatty & Good, 2011). However, systemic infection of non-legumes with endophytic diazotrophic bacteria has been observed in several plant species and in some cases, appears to satisfy the majority of the “host” plant’s N requirement with amounts of BNF similar to that of legume root nodules (see Sugarcane and Other Crops and Nitrogen Fixation in Gymnosperms below).


7. Actinorhizal symbiosis

The actinorhizal symbiosis refers to a root nodule-forming, nitrogen-fixing symbiotic relationship that is functionally analogous to the legume x Rhizobium symbiosis but distinct in most of the details. It is restricted to members of a small group of woody, non-legume pioneer species known as Actinorhizal plants and diazotrophs belonging to a single genus, Frankia, in a phylum of mostly filamentous soil bacteria, the Actinobacteria. Actinorhizal plants comprise approximately 200 plant species in 24 genera belonging to 8 plant families: Casuarinaceae, Betulaceae, Myricaceae, Elaeagnaceae, Coriariaceae, Rhamnaceae, Datiscaceae and Rosaceae (Huss-Danell, 1997). All but some members of Datiscaceae are shrubs or trees, and all are relatively shade-intolerant pioneering species that are able to colonize N-poor sites due to their ability to enter into the actinorhizal symbiosis (Crocker & Major, 1955; Chapin et al., 1994). Frankia spp. are notoriously difficult to grow in culture hence our understanding of these microorganisms and the actinorhizal symbioses lags far behind that of the well-characterized legume x Rhizobium associations. Owing to their recalcitrance to growth in culture, taxonomy of the genus Frankia is poorly developed, with only one species designation since a member of this genus was first reproducibly isolated over 30 years ago (Callaham et al., 1978). Numerous strains of Frankia have since been isolated from inside surface-sterilized root nodules and are designated as Frankia spp. followed by a strain number or name.

In contrast to the easily cultured, gram negative rod shaped cells that typify Rhizobium (and relatives) species, Frankia spp. strains possess gram-variable cells of three distinct types: filamentous vegetative hyphae, reproductive spores and N2-fixing vesicles (Benson & Silvester, 1993). The infection of plant roots and subsequent formation of root nodules bear some similarity to these processes in Rhizobium-infected legumes, but the origin and composition of Frankia-induced root nodules differs significantly (Wall & Berry, 2008). Unlike the de novo structure of a legume root nodule, actinorhizal root nodules are comprised of numerous tightly or loosely packed lobes, each of which originates from the pericycle indicating that it is a modified lateral root. Unlike Rhizobium bacteroids, the filamentous Frankia hyphae differentiate into an active nitrogen fixing form by developing numerous vesicles inside the root nodule or they may differentiate into spores and contribute nothing to the N economy of the plant. The tendency to sporulate in the nodule appears to be a strain specific characteristic that is influenced by the identity of plant host species. The oxygen concentration in root nodules is regulated primarily by varying the number and thickness of nitrogen fixing vesicle wall layers, however, the elegant heme protein leghemoglobin, which facilitates oxygen diffusion and supply in legume root nodules, has also been detected in nodules of some actinorhizal plants. Physiological and structural differences between actinorhizal and legume root nodules suggest that actinorhizal symbioses are less evolved and less efficacious in NF compared to legume x Rhizobium symbioses, but an effective actinorhizal symbiosis can fix several hundred kilograms of N per hectare annually, an amount which is similar to an effective legume symbiosis.


8. Bacterial endophytes

The presence of microorganisms in plant tissues might reasonably be considered an indication that a disease state is imminent, however this is not necessarily the case. Several decades ago, Trevet and Hollis (1948) reported the occurrence of bacteria within tissues of healthy potato plants and several studies have since demonstrated that internal tissues of healthy plants are colonized by bacteria. The term ‘bacterial endophytes’ has been used to describe bacteria that reside within living plant tissues without causing disease (Wilson, 1995; van der Lelie et al., 2009), however it does not differentiate whether such bacteria are (i) truly harmless, (ii) latent pathogens (Sinclair & Cerkauskas, 1996) or (iii) able to elicit production of symbiotic structures such as root nodules on the host. We use the term ‘endophyte’ in this paper to describe bacteria that ‘can be detected at a particular moment within the tissue of apparently healthy plant hosts’ (Schultz & Boyle, 2005) without inducing disease or organogenesis (Iniguez et al., 2005). In contrast to free-living, rhizosphere or phyllosphere microorganisms, bacterial endophytes are better protected from abiotic stresses such as extreme variations in temperature, pH, nutrient and water availability as well as biotic stresses such as competition (Loper et al., 1985; Cocking, 2003; Rosenblueth & Martinez-Romero, 2006). In addition, bacterial endophytes colonize niches that are more conducive to forming mutualistic relationships with plants through NF, for example, as suggested in sugarcane and other crops (see below) (Richardson et al., 2009).

8.1. Bacterial endophytes of sugarcane and other agricultural crops

In the 1980’s, Brazilian researchers were perplexed by the consistently high yields of field-grown sugarcane, an N-demanding crop, without exogenous N fertilizer application and looked for a microbiological explanation for this apparently anomalous observation. After it was determined that rhizospheric NF did not occur at sufficient rates to facilitate high sugarcane yields, Cavalcante & Döbereiner (1988) looked for microorganisms within sugarcane tissues that might be involved and isolated a diazotrophic bacterium, Gluconoacetobacter diazotrophicus, previously known as Acetobacter diazotrophicus. Its ability to establish high endophytic populations and to fix N2 in high sucrose concentrations (Boddey et al., 1991) at low pH (Boddey et al., 1991; Stephan et al., 1991), conditions which typify sugarcane tissues, led to the suggestion that this diazotrophic bacterium could satisfy almost all of the sugarcane N requirements. Several other N-contributing diazotrophic endophytes were subsequently found to associate with sugarcane including two Herbaspirillum species (Cavalcante & Döbereiner, 1988; Baldani et al., 1992; 2002), Azoarcus spp. (Reinhold et al., 1993) and Azospirillum brasilense (de Bellone and Bellone, 2006). However, more recent studies indicate that the primary source of diazotrophy may involve a consortium of bacteria, possibly including uncultured strains, that live on or inside plant tissues (Burbano et al., 2011; Taulé et al., 2012). Since the discovery of diazotrophic endophytes in sugarcane (Saccharum officinarum L.) (Ruschel et al., 1975), several other agriculturally important crop species including rice (Oryza sativa) (Shrestha & Ladha, 1996), maize (Zea mays L.) (Montañez et al., 2009) and kallar grass (Leptochloa fusca L.) (Malik et al., 1997) have been postulated to receive significant amounts of fixed N2 in this way.

Despite sugarcane’s apparent potential to derive much of its N from BNF, it has not been proven that G. diazotrophicus or any endophytic diazotroph is the primary causal agent of N accumulation by this plant in the field. Non-culturable endophytes with plant growth-promoting potential have been detected in sugarcane tissues (Hallmann et al., 1997; Mendes et al., 2007). In addition, all known culturable bacterial endophytes including G. diazotrophicus colonize the rhizosphere as well as internal plant tissues, rendering it difficult if not impossible to distinguish endophytic from non-endophytic BNF. There is also no clear evidence that N is transferred directly from endophytic diazotrophs to the host plant, unlike other symbiotic N2 fixing systems where specific nitrogenous compounds such as ureides in soybean (Herridge, 1982) and citrulline in alder (Leaf et al., 1958) are synthesized to transport fixed N from the site of NF to other host tissues. It is generally accepted that the N transfer to the host from rhizospheric NF results from the release of mineralized N from dead bacterial cells (Dobbelaere et al., 2003; Momose et al., 2009; Mia & Shamsuddin, 2010). Whether or not a similar mechanism operates with endophytic diazotrophs remains unanswered.

If endophytic diazotrophs are ultimately proven to be the primary cause of BNF and growth promotion of their host plants, such a plant x microbe association would represent another type of mutualistic symbiosis where the plant provides photosynthate and a competition-free, microaerobic environment for effective N2 fixation (Hallman et al., 1997; Reinhold-Hurek & Hurek, 1998a,b; Santi et al., 2013) for microorganisms in return for plant growth-promoting amounts of N from BNF. In contrast to the legume x Rhizobium and actinorhizal symbioses, no symbiosis-specific structures (e.g., nodules or chambers housing bacteria) have been detected in infected host, though Anand & Chanway (2013 a)observed occasional plant cells filled with diazotrophic Paenibacillus polymyxa in inoculated lodgepole pine tissues (Fig. 1b).

8.2. Nitrogen fixation in gymnosperms: Effects on seedling growth and N content

Lodgepole pine (Pinus contorta var. latifolia (Dougl.) Engelm.), a commercially important gymnosperm species indigenous to western North America, is capable of growing in very rocky substrates and is notable for its ability to thrive on nutrient poor, N-limited soils (Weetman et al., 1988; Chapman & Paul, 2012). Based on earlier work with lodgepole pine suggesting that rhizospheric BNF contributed only small amounts of N to seedlings (Chanway & Holl, 1991) as well as reports that BNF in sugarcane was endophytic, we searched for endophytic diazotrophs in lodgepole pine as a possible explanation for the ability of this species to grow on N-deficient substrates. We successfully isolated several Paenibacillus strains that possessed significant acetylene reduction activity from extracts of surface-sterilized lodgepole pine seedling and tree tissues (Bal et al., 2012) and tested them for BNF with lodgepole pine and western red cedar seedlings in a 15N soil dilution assay (Bal & Chanway, 2012a,b). When pine was reintroduced to one of the strains, Paenibacillus polymyxa strain P2b-2R, and grown in a very N-limited soil, seedlings were found to derive more than half (66%) of their foliar N from BNF, but their growth was inhibited compared to non-inoculated controls 9 months after planting (Bal & Chanway, 2012a). Similar effects, i.e., BNF and seedling growth inhibition, were observed with western red cedar (Bal & Chanway, 2012b).

Based on these results, we hypothesized that soil N depletion would eventually restrict the growth rate of control seedlings to a point where they would be outperformed by N2-fixing seedlings, and set up longer term growth experiments to evaluate this possibility. After a 13- month growth period, pine seedlings treated with P2b-2R were observed to derive most of their foliar N (79%) from BNF (Anand et al., 2013), which was confirmation of the ability of lodgepole pine to fix N after colonization by P. polymyxa. When compared with previous results, we found that inoculated seedlings grown for 7, 9 and 13 months derived 30%, 66% (Bal & Chanway, 2012a) and 79% (Anand et al., 2013), respectively, of their foliar N from the atmosphere. The progressive increase in the proportion of N derived from BNF i.e., %Ndfa, with seedling age suggests that BNF is an important component of N nutrition of pine in N-limited soil. Urquiaga et al. (1992) also observed an increasing reliance on BNF with seedling age, and concomitant decreasing soil N, in sugarcane cv. Krakatau. The %Ndfa rose from 6% to 55% during the interval 100 - 250 days after emergence. In addition to high %Ndfa, 13-month old seedlings treated with P2b-2R also accumulated significantly more biomass (78%) than controls, and had overcome the growth inhibition observed in seedlings of younger ages. While P. polymyxa possesses several characteristics that can result in plant growth promotion (Chanway, 2002), the enhanced performance of 13-month old seedlings was likely caused by an increase in the amount of N derived from BNF because the proportion of foliar N from BNF (68%-79%) (Anand et al., 2013) in older seedlings was only marginally higher than that of nine-month old seedlings (64%-66%) (Bal & Chanway, 2012a), but the concentration of foliar N (2.35%) (Anand et al., 2013) was 5-fold greater than nine-month old seedlings. The difference in total foliar N content, i.e., foliar N concentration x foliar biomass, was even greater and represented an elevation of foliar N concentration from a level considered to be very severely N deficient in all nine-month old seedlings (Bal & Chanway, 2012a) and the 13-month old controls (Anand et al., 2013) to one that is adequate for healthy lodgepole pine (Ballard & Carter, 1986). The comparatively early onset of BNF and delayed seedling growth response suggest that development of a fully effective N2-fixing bacterial population, able to enhance foliar N concentration in pine, is not a rapid process. Establishment of fully effective BNF in pine does not appear to depend on the population size of endophytic diazotrophs because P2b-2R colonization of root, stem and needle tissues in older seedlings did not differ significantly from younger seedlings (Anand et al., 2013), though it is possible that external root colonization was quantitatively related to BNF. However, if BNF is endophytic and requires physiological modifications of bacteria, e.g., differentiation of Paenibacillus into a N2-fixing form similar to Rhizobium bacteroids (Postgate, 1998), or plants, e.g., the establishment of specialized sites of BNF within plant tissues or cells, to be fully effective, some time may be required for them to complete. Our observations of intracellular colonization by P2b-2Rgfp (Figure 1; Anand & Chanway, 2013a) are consistent with this idea but further research is required to evaluate the physiology of P. polymyxa in association with pine as well as the relationship of bacterial population size to BNF.

Figure 1.

Colonization of cortex cells from a pine stem by green fluorescent protein (GFP) labeled Paenibacillus polymyxa strain P2b-2Rgfp using confocal laser scanning microscropy. Stem sections were viewed using green channel light (Figure 1a & b, panel 1) to visualize bright green GFP-labeled bacteria or red channel light (Figure 1a & b, panel 2) to visualize chloroplasts within pine cells. Using green channel light, GFP-labeled bacteria appear as green dots in a single pine cell (Figure 1a & b, panel 1). Under red light, chloroplasts appear as larger red organelles in the same single pine cells (Figure 1a & b, panel 2). Cells viewed using green and red channel light simultaneously (Figure 1a & b, panel 3) show GFP-labeled bacteria and chloroplasts within a pine cell. The white circle in Figure 1a, panel 3 encloses (i) several GFP-labeled bacterial cells, (ii) a single chloroplast in Figure 1a, panel 2 and GFP-labeled bacteria associated very closely with the chloroplast in Figure 1a, panel 3. Complete methodology is described in Anand and Chanway (2013a).

8.3. Nitrogen fixation in gymnosperms: Colonization of seedlings

The ability of P. polymyxa to colonize gymnosperm seedlings internally and externally has been evaluated in several greenhouse experiments (Bent & Chanway, 2002; Bal & Chanway, 2012a,b; Anand et al., 2013), but the ability of a green fluorescent protein-tagged derivative of P2b-2R to colonize internal pine tissues (Anand & Chanway, 2013a) provided convincing evidence that P2b-2R is endophytic, notwithstanding possible differences in behavior of GFP-tagged and wild type cells (van der Lelie et al., 2009). The population densities of P2b-2R inside pine tissues were observed to be comparable to densities reported for endophytic diazotrophs of crop plants such as rice (Oryza sativa L.), 104 - 105 cfu/g tissue (Elbeltagy et al., 2001), surgarcane, 105 - 107 (Sevilla et al., 2001) and grape (Vitis vinifera L.), 105 -109 (Compant et al., 2005) as well as hybrid spruce (Picea glauca x P. engelmannii), 103 – 105 (Shishido et al., 1999) and other tree species, 101 – 107 (Izumi, 2011). Recovery of P2b-2R from stem and needle tissues suggests bacteria migrate from the roots and soil to aerial plant organs: the observation that root and stem populations decreased while needle populations increased (Anand et al., 2013) supports this idea. Compant et al. (2005) observed a similar trend in grape plantlets inoculated with Burkholderia sp. PsJN and suggested that the stem acts as a transportation corridor for bacteria to reach leaves, which they considered a sink for endophytes. However, it is also possible that primordial pine stem and needle tissues were colonized at germination by P2b-2R residing on the seed coat and in the spermosphere, resulting in growth of bacterial populations in situ after shoots expanded, but the mechanism and patterns by which P2b-2R colonizes aerial pine tissues cannot be affirmed without further study.

We also observed intact cells from different pine stem sections that were colonized internally by GFP-labeled bacteria (Anand & Chanway, 2013a). In some cases, GFP-labeled bacteria were observed in pine cells in close proximity to chloroplasts (Fig. 1a), which raises the possibility that bacteria colonized microsites near these energy-generating organelles. GFP labelling were also observed tightly packed within other pine cells (Fig. 1b). Whether bacteria in either of these pine cells (Fig. 1) were fixing N cannot be determined, but these unique endophytic colonization patterns warrant further study. Endophytic bacteria have also been observed inside cells of grape (Compant et al., 2005), grasses (Hurek et al., 1994), sugarcane (James & Olivares, 1998) and poplar (Populus deltoides x P. nigra) (van der Lelie et al., 2009). Intracellular colonization of the pine stem cortex was rare and we were unable to inspect needle tissues for P2b-2Rgfp so it is unclear if colonized pine cells are of biological importance. GFP-labeled bacteria may simply have detected micro-colonies of bacteria feeding on dead or dying plant cells in the stem cortex. Alternatively, it is tempting to hypothesize a possible role for intracellular bacteria in BNF considering the foliar 15N dilution observed in colonized seedlings (Bal and Chanway, 2012a,b; Anand et al., 2013; Anand & Chanway, 2013b). Could these cells be specialized seats of N2-fixation, perhaps similar to Rhizobium bacteroids inside cells of legume root nodules? You et al. (1991) reported root cortex cells of rice, a non-leguminous plant, contained diazotrophic bacteria and some species of algae have been found to harbor endophytic bacteria near their chloroplasts (Colombo, 1978; Preisig & Hibberd, 1984). While this is a very interesting possibility, it is premature to focus on intracellular BNF in pine without more convincing supporting data.

Results with lodgepole pine strongly suggest that seedlings can fix N2 when colonized by P. polymyxa and demonstrate that N2-fixing seedlings develop healthy foliar N levels and enhanced growth in very N-limited soil. Western red cedar may also benefit from treatment with this bacterium (Bal & Chanway, 2012b; Anand & Chanway, 2013b), but amount of BNF and plant growth promotion were reduced compared to pine. Because P. polymyxa P2b-2R originated from pine seedling tissue, it is tempting to speculate that some degree of plant x microbe specificity may exist in this type of association. Research with pine and cedar raises several other questions regarding the ecological importance of endophytic diazotrophs in gymnosperms. For example, is P. polymyxa unique in its ability to enter into this type of interaction with gymnosperms or is this a comparatively common trait in conifer forests? If P. polymyxa is unique, how widespread is its distribution? How long must pine and diazotroph coexist before effective N fixation occurs and is BNF restricted to the seedling stage? What are the effects of soil N on colonization and BNF by P. polymyxa? It is possible that endophytic diazotrophs play a key role in natural regeneration of gymnosperms at N-poor sites but whether this is true and their importance in forest ecosystems can only be ascertained through future studies.


9. Conclusions and future research

Biological nitrogen fixation is a free and environmentally benign process through which biologically useful N can be generated for plant growth. Because nitrogen fixation is most effective when bacteria associate with plants to some degree, future research should focus on the various known plant associative and symbiotic nitrogen fixing systems. However, owing to the complexity of root-nodule based symbioses, colonization of plants by endophytic diazotrophs seems to hold the greatest potential for expansion of BNF to plant species that do not normally fix N2. Initial steps might include mass screening of known endophytic diazotrophs with commercially important plant species while continuing studies of effective nitrogen fixing associations involving plants that are colonized by endophytic diazotrophs. Such studies will help to elucidate the molecular, physiological and ecological details of this potentially useful plant x microbe interaction.



Funding for this study was provided by an NSERC Discovery Grant to CPC.


  1. 1. Adams, D.G. & Duggan, P.S. (2008). Cyanobacteria-bryophyte symbioses. Journal of Experimental Botany, Vol.59, pp.1047-1058
  2. 2. Anand, R & Chanway, C.P. (2013a). Detection of GFP-labeled Paenibacillus polymyxa in auto-fluorescing pine seedling tissues. Biology & Fertility of Soils, Vol.49, pp.111-118 doi: 10.1007/s0037401207279
  3. 3. Anand, R. & Chanway, C.P. (2013b). N2-fixation and growth promotion in cedar colonized by an endophytic strain of Paenibacillus polymyxa. Biology & Fertility of Soils, Vol.49, pp.235-239 doi: 10.1007/s0037401207359
  4. 4. Anand, R.; Grayston, S. & Chanway C.P (2013). N2-fixation and seedling growth promotion of lodgepole pine by endophytic Paenibacillus polymyxa. Microbial Ecology, Vol.66, pp.369-374 doi: 10.1007/s0024801301961
  5. 5. Bal A.S.; Anand, R.; Berge, O. & Chanway, C.P. (2012). Isolation and identification of diazotrophic bacteria from internal tissues of Pinus contorta and Thuja plicata. Can J For Res., Vol.42, pp.807–813
  6. 6. Bal, A.S. & Chanway, C.P. (2012a). Evidence of nitrogen fixation in lodgepole pine inoculated with diazotrophic Paenibacillus polymyxa. Botany, Vol.90, pp.891–896
  7. 7. Bal, A. & Chanway, C.P. (2012b). 15N foliar dilution of western red cedar in response to seed inoculation with diazotrophic Paenibacillus polymyxa. Biology & Fertility of Soils, Vol.48, pp.967–971 doi: 10.1007/s0037401206999
  8. 8. Baldani, J.I.; Reis, V.M.; Baldani, V.L.D. & Döbereiner, J. (2002). A brief story of nitrogen fixation in sugarcane—reasons for success in Brazil. Func Plant Biol., Vol.29, pp.417–423
  9. 9. Baldani, V.L.D.; Baldani, J.L.; Olivares, F. & Dobereiner, J. (1992). Identification and ecology of Herbaspirillum seropedicae and the closely related Pseudomonas rubrisubalbicans. Symbiosis, Vol.13, pp.65–73
  10. 10. Ballard, T.M. & Carter, R.E. (1986). Evaluating forest stand nutrient status. B.C. Min. For., Victoria. Land Manage., Rep.29
  11. 11. Bashan, Y. & Holguin, G. (1997). Azospirillum plant relationships : environmental and physiological advances (1990-1996). Can. J. Microbiol., Vol.43, pp.103-121
  12. 12. Bashan, Y.; Holguin, G. & de-Bashan, L.E. (2004). Azospirillum-plant relationships : physiological, molecular, agricultural, and environmental advances (1997-2003). Can. J. Microbiol., Vol. 50, pp.521-577
  13. 13. Beatty, P.H. & Good A.G. (2011). Future prospects for cereals that fix nitrogen. Science, Vol.333, pp.416-417
  14. 14. Benson, D.R. & Silvester, W.B. (1993). Biology of Frankia strains, actinomycete symbionts of actinorhizal plants. Microbiol Rev., Vol.57, pp.293–319
  15. 15. Bent, E. & Chanway, C.P. (2002). Potential misidentification of a spore-forming Paenibacillus polymyxa isolate as an endophyte by using culture-based methods. Applied and Environmental Microbiology, Vol.68, pp.4650-4652
  16. 16. Bergman, B.; Johansson, C. & Söderback, E. (1992). The Nostoc-Gunnera symbiosis. New Phytol., Vol.122, pp.379–400
  17. 17. Bezdicek, D.F. & Kennedy A.C. (1998) In Lynch JM, Hobbie JE (eds), Microorganisms in Action. Blackwell, Oxford, UK
  18. 18. Boddey, R.M.; Urquiaga, S.; Reis, V. & Dobereiner, J. (1991). Biological nitrogen fixation associated with sugar cane. Plant and Soil, Vol.137, pp.111-117
  19. 19. Burbano, C.S.; Liu, Y.; Rösner, K.L.; Reis, V.M.; Caballero-Mellado, J.; Reinhold-Hurek B. & Hurek, T. (2011). Predominant nifH transcript phylotypes related to Rhizobium rosettiformans in field-grown sugarcane plants and in Norway spruce. Environ Microbiol Rpt., Vol.3, pp.383–389
  20. 20. Cacciari, I.; Del Gallo, M.; Ippoliti, S.; Lippi, D.; Pietrosanti, T. & Pietrosanti, W. (1986). Growth and survival of Azospirillum brasilense and Arthrobacter giacomelloi in binary continuous culture. Plant Soil., Vol. 90, pp. 107-116
  21. 21. Callaham, D.; Del Tredici, P. & Torrey, J.G. (1978). Isolation and cultivation in vitro of the actinomycete causing root nodulation in Comptonia. Science Vol.199, pp.899–902
  22. 22. Campbell, E.L. & Meeks, J.C. (1989). Characteristics of hormogonia formation by symbiotic Nostoc spp. in response to the presence of Anthoceros punctatus or its extracellular products. Applied and Environmental Microbiology, Vol.55, pp.125-131
  23. 23. Carpenter, E.J. (1972). Nitrogen fixation by a blue-green epiphyte on pelagic Sargassum. Science, Vol.178, pp.1207-1208
  24. 24. Cavalcante, V.A. & Döbereiner, J. (1988). A new acid-tolerant nitrogen-fixing bacterium associated with sugarcane. Plant Soil., Vol.108, pp.23-31
  25. 25. Cerveny, J.; Sinetova, M.A.; Valledor, L.; Sherman, L.A. & Nedbal, L. (2013). Ultradian metabolic rhythm in the diazotrophic cyanobacterium Cyanothece sp. ATCC 51142. Proc Natl Acad Sci U.S.A., Vol.110, pp.13210-13215
  26. 26. Chanway, C.P. & Holl, F.B. (1991). Biomass increase and associative nitrogen fixation of mycorrhizal Pinus contorta seedlings inoculated with a plant growth promoting Bacillus strain. Can J Bot., Vol.69, pp.507–511
  27. 27. Chanway, C.P. (2002). Plant growth promotion by Bacillus and relatives, pp. 219-235. In Berkeley R, Heyndrickx M, Logan N, De Vos, P (eds), Applications and systematics of Bacillus and relatives. Blackwell, Oxford, UK
  28. 28. Chapin, F.S.; Walker, L.R.; Fastie, C.L. & Sharman, L.C. (1994). Mechanisms of primary succession following deglaciation at glacier bay, Alaska. Ecological Monographs, Vol.64, pp.149-175
  29. 29. Chapman, W.K. & Paul, L. (2012). Evidence that northern pioneering pines with tuberculate mycorrhizae are unaffected by varying soil nitrogen levels. Microb Ecol., Vol.64, pp.964–972
  30. 30. Cleveland, C.; Townsend, A.; Schimel, D.; Fisher, H.; Howarth, R.; Hedin, L.; Perakis, S.; Latty, E.; Von Fischer, J.; Elseroad, A. & Wasson, M. (1999). Global patterns of terrestrial biological nitrogen (N2) fixation in natural ecosystems. Global Biogeochem. Cycles., Vol.13, pp.623-645
  31. 31. Cocking, E. (2003). Endophytic colonization of plant roots by nitrogen-fixing bacteria. Plant and Soil, Vol.252, pp.169-175
  32. 32. Colombo, P.M. (1978). Occurrence of endophytic bacteria in siphonous algae. Phycologia, Vol.17, pp.148–151. doi: 10.1093/biomet/78.3.691
  33. 33. Compant, S.; Reiter, B.; Sessitsch, A.; Nowak, J.; Clement, C. & Ait Barka, E. (2005). Endophytic colonization of Vitis vinifera L. by plant growth-promoting bacterium Burkholderia sp. strain PsJN. Appl. Environ. Microbiol., Vol.71, pp.1685-1693
  34. 34. Crocker, R.L. & Major, J. (1955). Soil development in relation to vegetation and surface age at glacier bay, Alaska. The Journal of Ecology, Vol.43, pp.427-448
  35. 35. Curl, E.A. & Truelove, B. (1986). The Rhizosphere. Springer, New York
  36. 36. Dalton, D.A. & Kramer, S. (2006). Nitrogen-fixing bacteria in non-legumes. In Springer Netherlands, Dordrecht pp. 105-130
  37. 37. de Bellone S.C. & Bellone, C.H. (2006). Presence of endophytic diazotrophs in sugarcane juice. World Journal of Microbiology and Biotechnology, Vol.22, pp.1065-1068
  38. 38. Dobbelaere, S.; Vanderleyden, J. & Okon, Y. (2003). Plant growth-promoting effects of diazotrophs in the rhizosphere. Critical Reviews in Plant Sciences, Vol.22, pp.107-149
  39. 39. Döbereiner, J. &Pedrosa, F.O. (1987). Nitrogen-fixing bacteria in non-leguminous crop plants. Science Tech, Inc., Madison, Wisconsin
  40. 40. Elbeltagy, A.; Nishioka, K.; Sato, T.; Suzuki, H.; Ye, B.; Hamada, T.; Isawa, T.; Mitsui, H. & Minamisawa, K. (2001). Endophytic colonization and in planta nitrogen fixation by a Herbaspirillum sp. isolated from wild rice species. Appl Environ Microbiol., Vol.67, pp.5285–5293
  41. 41. Galloway, J.N.; Townsend, A.R.; Erisman, J.W.; Bekunda, M.; Cai, Z.; Freney, J.R.; Martinelli, L.A.; Seitzinger, S.P. & Sutton, M.A. (2008). Transformation of the nitrogen cycle: recent trends, questions, and potential solutions. Science, Vol.320, pp.889-892
  42. 42. Gehring, C.; Vlek, P.L.G.; de Souza, L.A.G. & Denich, M. (2005). Biological nitrogen fixation in secondary regrowth and mature rainforest of central Amazonia. Agric Ecosyst Environ., 111: 237-252
  43. 43. Gehringer, M.M.; Pengelly, J.J.; Cuddy, W.S.; Fieker, C.; Forster, P.I. & Neilan, B.A. (2010). Host selection of symbiotic cyanobacteria in 31 species of the Australian cycad genus: Macrozamia (Zamiaceae). Mol Plant Microbe Interact., Vol.23, pp.811-22
  44. 44. Grayston, S.J.; Vaughan, D. & Jones, D. (1997). Rhizosphere carbon flow in trees, in comparison with annual plants: the importance of root exudation and its impact on microbial activity and nutrient availability. Applied Soil Ecology, Vol.5, pp.29–56
  45. 45. Hallmann, J.; Quadt-Hallmann, A.; Mahaffee, W.F. & Kloepper, J.W., 1997. Bacterial endophytes in agricultural crops. Can. J. Microbiol., 43: 895-914
  46. 46. Hallmann, J.; Quadt‐Hallmann, A.; Mahaffee, W.F. & Kloepper, J.W. (1997). Bacterial endophytes in agricultural crops. Canadian Journal of Microbiology, Vol.43, pp.895–914
  47. 47. Herrero, A.; Muro-Pastor, A. & Flores, E. (2001). Nitrogen control in cyanobacteria. J.Bacteriol., Vol.183, pp.411-425
  48. 48. Herridge, D.F. (1982). Relative abundance of ureides and nitrate in plant tissues of soybean as a quantitative assay of nitrogen fixation. Plant Physiol., Vol.70, pp.1–6
  49. 49. Hobara S.; McCalley C.; Koba K.; Giblin A.E.; Weiss, M.S.; Gettel, G.M. & Shaver, G.R. (2006). Nitrogen fixation in surface soils and vegetation in an Arctic tundra watershed: A key source of atmospheric nitrogen. Arct Antarct Alp Res., Vol.38, pp.363–372
  50. 50. Hurek, T.B.; Reinhold-Hurek, B.; Montagu, M.B. & Kellenberger, E. (1994). Root colonization and systematic spreading of Azoarcus sp strain BH72 in grasses. J Bacteriol., Vol.176, pp.1913-1923
  51. 51. Huss-Danell, K. (1997). Actinorhizal symbioses and their N2 fixation. New Phytologist, Vol.136, pp.375-405
  52. 52. Iniquez, A.L.; Dong, Y.; Carter, H.D.; Ahmer, B.M.; Stone, J.M. & Triplett, E.W. (2005). Regulation of enteric endophytic bacterial colonization by plant defenses. Mol Plant Microbe Interact., Vol.18, pp.169-178
  53. 53. Izumi, H. (2011). Diversity of endophytic bacteria in forest trees. In: Pirttilä AM, Frank AC (eds) Endophytes of forest trees. Biology and Applications Series: Forestry Sciences Vol. 80. Springer, Heidelberg, pp 95–105
  54. 54. James, E.K. & Olivares, F.L.(1998). Infection and colonization of sugar cane and other graminaceous plants by endophytic diazotrophs. Critical Reviews in Plant Sciences, Vol.17 pp.77–119
  55. 55. Jayasinghearachchi, H.S. & Seneviratne, G. (2004). Can mushrooms fix atmospheric nitrogen? J. Biosci., Vol.29, pp.293–296
  56. 56. Jones, D.L.; Hodge, A. & Kuzyakov, Y. (2004). Plant and mycorrhizal regulation of rhizodeposition. New Phytologist, Vol.163, pp.459 –480
  57. 57. Leaf, G.; Gardner, I.C. & Bond, G. (1958). Observations on the composition and metabolism of the nitrogen-fixing root nodules of Alnus. Journal of Experimental Botany, Vol.9, pp.320-31
  58. 58. Lindow, S.E. & Brandl, M.T. (2003). Microbiology of the phyllosphere. Appl. Environ., Vol.69, pp.1875-1883
  59. 59. Loper, J.E.; Haack, C. & Schroth, M.N. (1985). Population dynamics of soil Pseudomonads in the rhizosphere of potato (Solanum tuberosum L.). Applied and Environmental Microbiology, Vol.49, pp.416-422
  60. 60. Ferrario, M.E.; Villafane, V.; Helbling, W. & Holm-Hansen, O. (1995). The occurrence of the symbiont Richelia in Rhizosolenia and Hemiaulus in the North Pacific. Revista Brasileira de Biologia, Vol. 55, pp.439–443
  61. 61. Malik, K.A.; Bilal, R.; Mehnaz, S.; Rasul, G.; Mirza, M.S. & Ali, S. (1997). Association of nitrogen-fixing, plant-growth-promoting rhizobacteria (PGPR) with kallar grass and rice. Plant and Soil, Vol.194, pp.37–44
  62. 62. Meeks, J.C. (1998). Symbiosis between nitrogen-fixing cyanobacteria and plants. Bioscience, Vol.48, pp.266-276
  63. 63. Mendes, R.; Pizzirani-Kleiner, A.A.; Araujo, W.L.; Raaijmakerset, J.M. (2007). Diversity of cultivated endophytic bacteria from sugarcane: genetic and biochemical characterization of Burkholderia cepacia complex isolates. Appl Environ Microbiol., Vol.73, pp.7259–7267. doi: 10.1128/AEM.01222-07
  64. 64. Mia , M.A.B. & Shamsuddin, Z.H. (2010). Nitrogen fixation and transportation by rhizobacteria: a scenario of rice and banana. International Journal of Botany, Vol.6, pp.235-242
  65. 65. Momose, A.; Ohtake, N.; Sueyoshi, K..; Sato, T.; Nakanishi, Y.; Akao, S. & Ohyama, T. (2009). Nitrogen fixation and translocation in young sugarcan (Saccharum officinarum L.) plants associated with endophytic nitrogen-fixing bacteria. Microbes Environ., Vol.24, pp.224-230
  66. 66. Montañez, A.; Abreu, C.; Gill, P.R.; Hardarson, G. & Sicardi, M. (2009). Biological nitrogen fixation in maize (Zea mays L.) by 15N isotope-dilution and identification of associated culturable diazotrophs. Biol Fertil Soils Vol.45, pp.253–263. doi:10.1007/s00374-008-0322-2
  67. 67. Myrold, D.D. & Bottomley, P.J. (2007). Biological N inputs. In Soil microbiology, ecology and biochemistry. Elsevier, Burlington, MA, pp. 365-388
  68. 68. Newton, W.E. 2007. Physiology, biochemistry, and molecular biology of nitrogen fixation. In Biology of the Nitrogen Cycle. Elsevier, Amsterdam pp. 109-129
  69. 69. Nguyen, C. (2003). Rhizodeposition of organic C by plants: mechanisms and controls. Agronomie Vol.23, pp.375 –396
  70. 70. Oldroyd, G.E.D.; Murray, J.D.; Poole, P.S. & Downie, J.A. (2011). The rules of engagement in the legume-rhizobial symbiosis. Annu. Rev. Genet., Vol.45, pp.119–44
  71. 71. Postgate, J.R. (1998). Nitrogen fixation. Cambridge University Press, Cambridge, UK
  72. 72. Preisig, H.R. & Hibberd, D.J. (1984). Virus-like particles and endophytic bacteria in Paraphysomonas and Chromophysomonas (Chrysophyceae). Nord J Bot., Vol.4, pp.279–285
  73. 73. Reed, H.E. & Martiny, J.B.H. (2007). Testing the functional significance of microbial composition in natural communities. FEMS Microbiol. Ecol., Vol.62, pp.161-170
  74. 74. Reed, S.C.; Cleveland, C.C. & Townsend., A.R. (2008). Tree species control rates of free-living nitrogen fixation in a tropical rain forest. Ecology Vol.89, pp.2924-2934
  75. 75. Reed., S.C.; Cleveland, C.C. & Townsend, A.R. (2011). Functional ecology of free-living nitrogen fixation: a contemporary perspective. Annu. Rev. Ecol. Evol. Syst., Vol.42, pp.489–512
  76. 76. Reinhold, B.; Hurek, T.; Gillis, M.; Hoste, B.; Vancanneyt, M.; Kersters, K. & De Ley, J. (1993). Azoarcus gen.nov. nitrogen-fixing proteobacteria associated with roots of kallar grass (Leptochloa fusca L. Kunth), and description of two species, Azoarcus indigens sp. nov. and Azoarcus communis sp. nov. International Journal of Systematic Bacteriology, Vol.43, pp.574–584
  77. 77. Reinhold-Hurek, B. & Hurek, T. (1998a). Interactions of gramineous plants with Azoarcus spp. and other diazotrophs: identification, localization, and perspectives to study their function. Critical Reviews in Plant Sciences, Vol.17, pp.29–54
  78. 78. Reinhold-Hurek, B. & Hurek, T. (1998b). Life in grasses: diazotrophic endophytes. Trends in Microbiology, Vol.6, pp.139–144
  79. 79. Richardson, A.E.; Barea, J.M.; NcNeill, A.M. & Prigent-Combaret, C. (2009). Acquisition of phosphorus and nitrogen in the rhizosphere and plant growth promotion by microorganisms. Plant Soil., Vol.321, pp.305-339
  80. 80. Rosenblueth, M. & Martinez-Romero, E. (2006). Bacterial endophytes and their interactions with hosts. MPMI, Vol.19, pp. 827-837
  81. 81. Ruschel, A.; Henis, Y. & Salati, E. (1975). Nitrogen 15N tracing of N2-fixation with soil grown sugarcane seedlings. Soil Biol. Biochem., Vol.5, pp.83–89
  82. 82. Santi, C.; Bogusz, D. & Franche, C. (2013). Biological nitrogen fixation in non-legume plants. Annals of Botany, doi:10.1093/aob/mct048
  83. 83. Schell, D.M. & Alexander, V. (1973). Nitrogen fixation in Arctic coastal tundra in relation to vegetation and micro-Relief. Arct., Vol.26, pp.130–137
  84. 84. Schultz, B. & Boyle, C. (2005). The endophyte continuum. Mycological Research, Vol.109, pp.661-686
  85. 85. Sevilla, M.; Burris, R. H.; Gunapala, N. & Kennedy, C. (2001). Comparison of benefit to sugarcane plant growth and 15N incorporation following inoculation of sterile plants with Acetobacter diazotrophicus wild-type and Nif- mutant strains. Mol. Plant-Microbe Interact., Vol.14, pp.358–366
  86. 86. Shi, S.; O’Callaghan, M.; Jones, E.E.; Richardson, A.E.; Walter, C.; Stewart, A. & Condron, L. (2012). Investigation of organic anions in tree root exudates and rhizosphere microbial communities using in situ and destructive sampling techniques. Plant Soil., Vol.359, pp.149-163
  87. 87. Shishido, M. & Chanway, C.P. (1999). Spruce growth response specificity after treatment with plant growth-promoting pseudomonads. Can. J. Bot., Vol.77, pp.22-31
  88. 88. Shrestha, R.K. & Ladha, J.K. (1996). Genotypic variation in promotion of rice nitrogen fixation as determined by nitrogen 15N dilution. Soil Sci. Soc. Am. J., Vol.60, pp.1815-1821
  89. 89. Sinclair, J.B. & Cerkauskas, R.F. (1996). Latent infection vs. endophytic colonization by fungi. In: Redlin SC, Carris LM (eds) Endophytic fungi in grasses and woody plants. APS, St Paul, MN, pp 3-30
  90. 90. Solheim, B. & Zielke, M. (2002). Associations between cyanobacteria and mosses. In: Rai AN, Bergman B, Rasmussen (eds) Cyanobacteria in symbiosis. Dordrecht: Kluwer Academic Publishers, 37–152
  91. 91. Stephan, M.P.; Oliveira, M.; Teixeira, K.R.S.; Martinez-Drets, G. & Döbereiner, J. (1991). Physiology and dinitrogen fixation of Acetobacter diazotrophicus. FEMS Microbiol. Lett., Vol.77, pp.67–72
  92. 92. Talley, S.N.; Talley, B.J. & Rains, D.W. (1977). Nitrogen fixation by azolla in rice fields. In Genetic Engineering for Nitrogen Fixation (ed) Hollander, A. pp 259-282. New York: Plenum Press
  93. 93. Taulé, C.; Mareque, C.; Barlocco, C.; Hackembruch, F.; Reis, V.M.; Sicardi, M. & Battistoni, F. (2012). The contribution of nitrogen fixation to sugarcane (Saccharum officinarum L.), and the identification and characterization of part of the associated diazotrophic bacterial community. Plant Soil., Vol.356, pp.35–49
  94. 94. Trevet, I.W. & Hollis, J.P. (1948). Bacteria in storage organs of healthy plants. Phytopathology, Vol.38, pp.960-967
  95. 95. Udvarte, M. & Poole, P.S. (2013). Transport and metabolism in legume-Rhizobia symbioses. Annu. Rev. Plant Biol., Vol.64, pp.781–805
  96. 96. Urquiaga S.; Cruz K.H.S. & Boddey R.M. (1992). Contribution of nitrogen fixation to sugar cane: nitrogen-15 and nitrogen-balance estimates. Soil Science Society of America Journal, Vol.56, pp.105-114
  97. 97. van der Lelie, D.; Taghavi, S.; Monchy, S. et al. (2009). Poplar and its bacterial endophytes: coexistence and harmony. Critical reviews in Plant Sciences, Vol.28, pp.346-358
  98. 98. Wall, L.G. & Berry, A.M. (2008). “Early interactions, infection and nodulation in actinorhizal symbiosis” in Nitrogen-Fixing Actinorhizal Symbioses, K. Pawlowski and W. E. Newton (Eds), pp. 147–166, Springer, Dordrecht, The Netherlands
  99. 99. Warembourg, F.R. (1997). The 'rhizosphere effect': a plant strategy for plants to exploit and colonize nutrient-limited habitats. Bocconea, Vol.7, pp.187-194
  100. 100. Weetman, G.F.; Fournier, R.M. & Schnorbus, E. (1988). Lodgepole pine fertilization screening trials: four-year growth response following initial predictions. Soil Sci Soc Am J., Vol.52, pp.833–839
  101. 101. Wilson, D. (1995). Endophyte: the evolution of a term, and clarification of its use and definition. Oikos, Vol.73, pp.274-276
  102. 102. Wolk, C.P.; Ernst, A. & Elhai, J. (1994). Heterocyst metabolism and development. The Molecular Biology of Cyanobacteria, Vol.1, pp.769-823
  103. 103. Yanni, Y.G. (1992). The effect of cyanobacteria and azolla on the performance of rice under different levels of fertilizer nitrogen. World Journal of Microbiology and Biotechnology, Vol.8, pp.132-136
  104. 104. You, C.B.; Song, W.; Wang, H.X.; Li, J.P.; Lin, M. & Hai, W.L. (1991). Association of Alcaligenes faecalis with wetland rice. Plant and Soil, Vol.137, pp.81-85

Written By

C.P. Chanway, R. Anand and H. Yang

Submitted: 16 October 2013 Published: 29 January 2014