Biological Nitrogen Fixation☆

Biological Nitrogen Fixation☆ N Rascio and N La Rocca, University of Padua, Padua, Italy ã 2013 Elsevier Inc. All rights reserved.

Introduction Nitrogen-Fixing Organisms Nitrogenase and Nitrogen Fixation Ammonia Assimilation Diazothrophic Bacteria – Plant Symbioses Rhizobia–Legume Symbiosis Frankia-Dicotyledon Symbiosis Endophytic Diazotrophic Bacteria–Cereal Association Nitrogen Fixation in Free-Living Cyanobacteria Diazotrophic Cyanobacteria–Plant Symbioses Anabaeana–Azolla Symbiosis Cyanolichens A Swift Ecological Overview of Biological Nitrogen Fixation in Terrestrial and Aquatic Ecosystems References

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Introduction Nitrogen is a key element present in many biochemical compounds (such as nucleotide phosphates, amino acids, proteins, and nucleic acids) of living cells. Only oxygen, carbon, and hydrogen are more abundant in the cell. The entry of organic nitrogen in the food chains of natural ecosystems is essentially due to the activity of photoautotrophic organisms (cyanobacteria, algae, and terrestrial plants). These primary producers take up nitrogen from the environment mainly as nitrate, reduce it to ammonia, and then assimilate ammonia into organic compounds to form amino acids. However, this process of assimilatory reduction of nitrate is not the only change that nitrogen undergoes. In the biosphere nitrogen passes through many forms, ranging from the most reduced, NH3 (or NH4þ), to the most oxidized, NO3, in a biogeochemical cycle whose steps depend on both physical and biological events. The processes that involve living organisms (Figure 1) include the following:

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ammonification, carried out by saprophytic bacteria (e.g., Clostridium spp.) and fungi that detach NH3 from organic nitrogenous compounds and release this reduced form of inorganic nitrogen into the environment; nitrification, carried out by chemosynthetic bacteria that draw energy from oxidation of ammonia to nitrite (e.g., Nitrosomonas spp.) and of nitrite to nitrate (e.g., Nitrobacter spp.): NH4 þ þ 3=2 O2 ! NO2 þ H2 O þ 2Hþ DG0, ¼ 66 kcal mol1 NO2 þ 1=2 O2 ! NO3 DG0, ¼ 18 kcal mol1

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assimilatory reduction of nitrate to nitrite and then of nitrite to ammonia by nitrogen autotrophic organisms: NO3 þ NADðPÞH2 ! NO2 þ NADðPÞþ þ H2 O NO2 þ 6 reduced ferredoxins þ 8Hþ ! NH4 þ þ 6 oxidized ferredoxins þ 2 H2 O

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ammonia assimilation into aminoacids (see the pathway in the Ammonia Assimilation section) that leads to the recovery of organic nitrogen, thus closing the cycle.

Nonetheless, biological pathways of irreversible nitrogen loss also exist. One of them is carried out by facultative aerobic bacteria. These microorganisms (e.g., Pseudomonas spp. and Alcaligenes spp.) bring about a process of dissimilatory reduction of nitrate, called denitrification. Under anoxic conditions they activate anaerobic respiration by using nitrate and other oxidized forms of nitrogen instead of oxygen as final electron acceptors of the respiratory chain: 2 NO3 þ 4e þ 4Hþ ! 2 NO2 þ 2H2 O ☆ Change History: May 2013. N Rascio and N La Rocca updated keywords, introdution, references, further reading. Figure 1 legend, and cyanolichens, and A swift Ecological Overview of Biological Nitrogen Fixation in Terrestrial and Aquatic Ecosystems sections.

Reference Module in Earth Systems and Environmental Sciences

http://dx.doi.org/10.1016/B978-0-12-409548-9.00685-0

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Figure 1 A simplified scheme of the nitrogen cycle showing the steps carried out by living organisms: (1) ammonification; (2, 3) nitrification; (4, 5) assimilatory reduction of nitrate; (6) ammonia assimilation; (7) denitrification or anammox; and (8) nitrogen fixation.

2 NO2 þ 2e þ 4Hþ ! 2NO þ 2H2 O 2 NO þ 2e þ 2Hþ ! N2 O þ H2 O N2 O þ 2e þ 2Hþ ! N2 þ H2 O This leads to volatile nitrogen forms (e.g., N2O or N2), which are lost to the atmosphere. Another widespread process which contributes substantially to the loss of fixed nitrogen as N2 in different natural environments has recently been discovered. This process is the anaerobic ammonium oxidation (anammox) performed by chemoautotrophic bacteria (“anammox” bacteria, such as Brocadia anammoxidans and Scalindua spp.) that derive energy for growth from oxidation of NH4þ to N2 in complete absence of oxygen using nitrite as electron acceptor:
NH4 þ þ NO2 ! N2 þ 2H2 O DG0, ¼ 86 kcal mol1 In natural ecosystems, the recovery of nitrogen, necessary to satisfy the nutritional demands of the inhabiting organisms, occurs through biological nitrogen fixation (Figure 1). This event is of capital importance and consists in the reduction of molecular nitrogen (N2) to ammonia (NH3), providing the Earth’s ecosystems with about 200 milion tons N per year. It has been estimated that the 80–90% of the nitrogen available to plants in natural ecosystems originates from biological nitrogen fixation.

Nitrogen-Fixing Organisms Nitrogen constitutes almost 80% of the atmosphere, but is metabolically inaccessible to plants due to the exceptional stability of the triple covalent bond (N  N). The ability to catalyze enzymatic reduction of N2 to NH3 is limited to a variety of prokaryotes defined as nitrogen-fixing or diazotrophic microorganisms, which are widely distributed in all ecosystems as either free-living organisms or in symbiotic association with a number of different plants. These N2-fixing prokaryotes can be anaerobic, facultative aerobic, aerobic, photosynthetic, or nonphotosynthetic (Table 1). All carry out N2 reduction by an enzymatic complex termed nitrogenase.

Nitrogenase and Nitrogen Fixation The complex of nitrogenase (Figure 2) consists of two distinct enzymes: dinitrogenase reductase and dinitrogenase, neither of which has enzymatic activity by itself (Igarashi and Seefeldt, 2003). Dinitrogenase reductase is a dimeric (a2) Fe-protein of about 70 kDa with a 4Fe–4S cluster, which binds ATP and transfers electrons to dinitrogenase. The latter enzyme is a tetrameric (a2b2)

Biological Nitrogen Fixation

Table 1

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Some examples of organisms that carry out nitrogen fixation

N2-fixing prokaryotes

Genera

Aerobic bacteria

Azotobacter Azospirillum Klebsiella Bacillus Chromatium Chlorobium Clostridium Desulfovibrio Anabaena Nostoc Calotrix

Facultative bacteria Anaerobic bacteria photosynthetic Non-photosynthetic Cyanobacteria

Figure 2 The enzymatic complex of nitrogenase.

FeMo protein of about 220 kDa. It contains two Mo–Fe–S clusters and a variable number of Fe–S clusters and binds N2. Both these enzymes are very sensitive to oxygen, which rapidly inactivates them (Dixon and Wheeler, 1986). The molybdenum requiring nitrogenase (Mo-N2ase) is the long-studied enzyme complex found in all diazotrophs. However other two alternative nitrogenases have been identified in some free-living bacteria belonging to the genera Azotobacter and Rhodopseudomonas and in some cyanobacteria. These peculiar diazotrophic organisms can synthesize, in addition to Mo-Fe N2ase, a vanadium nitrogenase (V-N2ase) and a nitrogenase which requires only Fe (Fe-N2ase) (Newton, 2007). The alternative nitrogenases may serve as substitutive pathways for nitrogen fixation in molybdenum deficient conditions. They are structurally similar to the Mo-N2ase, except for the dinitrogenase components which are composed of a2b2d2 hexamers, they also operate in the same way, have the same requirements for activity and are reactive towards the same range of substrates. In the nitrogen reduction carried out by the Mo-N2ase (also referred to as conventional nitrogenase), the oxidized dinitrogenase reductase accepts an electron from a donor (reduced ferredoxin or reduced flavodoxin) and binds two molecules of adenosine 50 -triphosphate (ATP). This binding causes a conformational change of the Fe protein that lowers its redox potential (from 300 to 400 mV). The reduced Fe protein transfers the electron to the oxidized dinitroreductase with concomitant hydrolysis of both ATP molecules. Finally, the FeMo protein carries out the electron (and proton) transfer to the N2 bound to the MoFe cofactor. Since six electrons are required to reduce N2 to 2NH3, six sequential reduction events occur with the hydrolysis of 2ATP for each electron flowing through the nitrogenase. However, the nitrogenase also recognizes the protons (Hþ) in the cell, so that for each N2 reduced to 2NH3, two Hþ ions are reduced to H2, with the involvement of two additional electrons and the hydrolysis of another 4ATP. Thus, the overall reaction catalyzed by nitrogenase in the diazothrophic organisms is N2 þ 8e þ 8Hþ þ 16ATP ! 2NH3 þ H2 þ 16ADP þ 16Pi Under natural conditions the reduction of protons to hydrogen competes with that of nitrogen to ammonia for the electrons provided to nitrogenase by the donors. This lessens the efficiency of nitrogen fixation and leads to a waste of metabolic energy (ATP). Nevertheless, many nitrogen-fixing organisms have an uptake hydrogenase that reoxidizes H2 to 2Hþ and 2e The activity of this enzyme can greatly increase the efficiency of nitrogen fixation since it leads to ATP recovery by the flow of electrons through a respiratory transport chain, nitrogenase protection against the oxygen poisoning by reduction of O2 to H2O, and maintenance of nitrogenase activity by removal of H2 that inhibits the enzyme.

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For nitrogenase to function, low-potential electrons and energy (ATP) are needed. The most common source of electrons is ferredoxin (a small iron–sulfur protein). In phototrophic organisms the reduced ferredoxin can be derived from the photosynthetic electron flow, while in heterotrophic organisms the reduction occurs enzymatically through a pyruvate ferredoxin oxidoreductase that transfers electrons to oxidized ferredoxin from a-ketoacids such as pyruvate and a-ketoglutarate. An analogous reaction, carried out by a pyruvate flavodoxin oxidoreductase, produces reduced flavodoxin, a flavoprotein also used as an electron donor to nitrogenase. Some organisms may generate the required low-potential electrons by other alternative methods. The source of energy for reduction of N2 is ATP obtained from different metabolic pathways, according to the diazotrophic organism. In anaerobic phototrophic bacteria, the ATP comes from photosynthesis, while anaerobic heterotrophic organisms gain ATP essentially from fermentations that, due to the scarce oxidative efficiency, force them to consume large quantities of substrates. The aerobic organisms take advantage of the production of ATP through more efficient respiratory processes. Nevertheless, they still require mechanisms that keep oxygen away from the O2-sensitive nitrogenase. Some of these mechanisms are described in the following sections. Even if the nitrogen fixation is actually an exergonic process (DG0 ¼  15.2 kcal mol1), it has a high demand for energy due to the necessity to overcome unfavorable activation energies. The theoretical cost for reducing one molecule of N2 with the concomitant reduction of 2Hþ is 16ATP. However, under natural conditions the cost may be even higher due to the less than perfect efficiency of the process. This high energetic cost makes nitrogen fixation a strictly controlled process, through the modulation of the synthesis and activity of nitrogenase. All the other available forms of nitrogen (such as nitrate, nitrite, or amino acids) inactivate the enzymatic complex and inhibit the expression of the genes (nif genes) that code for the nitrogenase components (Helber et al., 1988; Cheng et al., 1999). The reaction products NH3 and H2 also cause strong inhibition. Thus, N2 fixation is an inductive process that diazotrophic organisms activate only in the absence of other more economic nitrogen sources.

Ammonia Assimilation The assimilation of NH3 obtained from N2 reduction occurs principally via the glutamine synthetase–glutamate synthase (GS-GOGAT) pathway (Nagatani et al., 1971). The first enzyme catalyzes the ATP-dependent assimilation of ammonia into glutamine using glutamate as a substrate: Glutamate þ NH3 þ ATP ! Glutamine þ ADP þ Pi The second enzyme catalyzes the reductive transfer of the amide group from glutamine to a-ketoglutarate, forming two molecules of glutamate. The reductants are NAD(P)H or reduced ferredoxin (Fdxred): Glutamine þ a  ketoglutarate þ NADðPÞH ðor Fdxred Þ ! 2glutamate þ NADðPÞþ ðor Fdx ox Þ One glutamate serves to keep the pathway going, whereas the other represents the gain in organic nitrogen.

Diazothrophic Bacteria – Plant Symbioses Nitrogen-fixing microorganisms have been found in roots or other organs of many species of plants with which they establish symbiotic associations. The diazotrophic partners can be aerobic bacteria or, in some cases, cyanobacteria. Symbiotic associations of current ecological importance for wide diffusion and the large nitrogen supply to the ecosystems are those between N2-fixing bacteria and roots of higher plants and, in particular, the rhizobia-legume and Frankia-dicotyledon symbioses. Another association of particular interest is that established by endophytic diazotrophic bacteria with cereals.

Rhizobia–Legume Symbiosis Most Leguminosae (about 90%) can establish a symbiotic association with aerobic diazotrophic Gram-negative bacteria commonly referred to as rhizobia. This symbiosis takes place in roots and brings about the formation of nodules in which N2 fixation occurs. The large contribution made by these symbioses to the nitrogen availability for agronomically important legumes is well known. Medicago sativa, for instance, can fix 300 kg N ha1 year1 and Vicia faba over 500 kg N ha1 year1. It has been calculated that soybean (Glycine max), which is the dominant crop legume, representing 68% of global production, can fix more than 16106 T N annually, corresponding to over 75% of the N fixed by crop legumes (Herridge et al., 2008). This makes biological nitrogen fixation a major component of sustainable agricultural systems, since it has the potential to greatly limit the use of chemical nitrogen fertilizers. Numerous species belonging to the family Leguminosae are also abundant in natural ecosystems, such as the forests of tropical regions (e.g., those in Brazil and Guyanas) (Koponen et al., 2003, Kreibich et al., 2006), where they can represent over 50% of all trees. Tropical forests often grow on substrates poor in mineral nutrients, and thus the continuous supply of nitrogen through biological N2 fixation acquires an essential role to maintain large nitrogen pools in these ecosystems.

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The rhizobia forming symbiosis with legume roots belong to five different genera: Rhizobium, Azorhizobium, Mesorhizobium, Sinorhizobium, and Bradyrhizobium. A given species of bacterium establishes symbiosis with one or few species of legumes (Table 2). This is due to the host-symbiont recognition occurring in the rhizosphere through the exchange of molecular signals. The first event of the root nodule formation is the chemotactic movement of the bacterium toward the root of the host plant in response to chemical attractants, usually specific flavonoids secreted by the root under nitrogen-starvation conditions. Each legume species produces its own particular cocktail of these compounds and this contributes to one of the distinctive characteristics of rhyzobiumlegume symbiosis, with a rhizobial species usually infecting a very limited host range (Marks, 2004). The compounds secreted by the roots induce the expression of host-specific bacterial genes (nod genes) coding for Nod factors (lipo-chito-oligosaccharides) that, in turn, induce plant responses and trigger the nodule developmental program (Spaink, 2000). Root infection starts with the bacterium-induced curling of a root hair, bacterium attachment to the hair surface, cell wall degradation, and formation of the infection thread. This is an internal tubular extension of the hair plasma membrane that carries out the proliferating rhizobia from the root surface into the root cortex. Concomitantly, some cortical cells undergo rapid divisions that give rise to the nodule primordium. When the branched infection thread reaches target cells within the developing nodule, its tip vesiculates releasing bacteria packaged in a membrane derived from the host cell plasmalemma. The rhizobia undergo some divisions but very soon they stop dividing and differentiate into diazotrophic bacteroids. Bacteroids and surrounding peribacteroid membrane form the symbiosome (Figure 3(b)), which is the site of N2 fixation. In the mature nodule (Figure 3(a)) specialized structures are developed around the infected tissue: an endodermis and a vascular system continuous with the root stele, and a layer

Table 2

Some examples of associations between rhizobia and legumes

Rhizobia

Host plants

Bradyrizobium japonicum Sinorhizobium meliloti Sinorhizobium fedii Azorhizobium caulinodans Rhizobium leguminosarum biovar. phaseoli Rhizobium leguminosarum biovar. trifolii Rhizobium leguminosarum biovar. viciae Mesorhizobium loti

Glycine, Vigna Medicago, Trigonella, Melilotus Glycine, Vigna Sesbania Phaseolus Trifolium Vicia, Pisum, Cicer Lotus, Lupinus, Anthillis

Figure 3 Schematic drawings of: (a) a determinate root nodule of a rhizobia-legume symbiosis and (b) a part of an infected cell with symbiosomes.

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Figure 4 Schematic drawing of an indeterminate root nodule of a rhizobia-legume symbiosis.

of cells hampering O2 diffusion to the root nodule interior. Some leguminous species such as soybean, peanut, and bean form spherical determinate nodules with a nonpersistent meristem (Figure 3(a)). Others, such as pea, clover, and alfalfa, form cylindrical indeterminate nodules with a persistent terminal meristem (Figure 4). The mature determinate nodule contains a homogenous central tissue with cells fully packed with nitrogen fixing bacteroids, whereas in the indeterminate nodule a gradient of developmental states occurs, due to the active meristem that continuously produces cells that become infected with bacteria (Ferguson et al., 2010). At maturity a meristematic zone, an invasion zone with infection threads, a N2-fixing zone with bacteroids and a senescent zone with degraded bacteroids can be distinguished along the nodule axis (Figure 4). Different mechanisms take place to obtain the microaerobic environment appropriate for maintaining ATP production in host cells and bacteroids and for preserving, at the same time, nitrogenase activity in N2-fixing tissue. The first hindrance to the entry of oxygen into the infected cells is the mechanical diffusion barrier in the nodule parenchyma. Moreover, leghemoglobin is synthesized in the cytoplasm of the host cells (Downie, 2005). This oxygen-binding protein plays a major role in delivering oxygen to the bacteroid surface and accounts for the characteristic pink color of N2-fixing tissue. Efficient bacteriod respiration also restricts oxygen penetration into the cytoplasm and provides nitrogenase with the ATP and reductants required. Finally, in most rhizobia the activity of an uptake hydrogenase is an additional help for protecting nitrogenase against the O2-poisoning (Ciccolella et al., 2010). Bacteroids do not have enzymes for the ammonia assimilation. For this reason, the NH3 obtained from N2 reduction is released into the root cell where the assimilation occurs via GS-GOGAT pathway (Figure 5). This leads to production of glutamine, glutamate, and, successively, of other nitrogenous transport compounds. Some of these organic compounds are returned to the bacteroids, but most are exported to the plant shoot via xylem. In order to sustain N2 fixation, the host plant must supply the bacteroids with a carbon source, which arrives to the root nodule via phloem as sucrose. However, this sugar is metabolized in the host cell and converted to C4 dicarboxylates, principally malate. The dicarboxylates, in fact, are transported across the peribacteroid membrane, becoming the primary carbon source for the N2-fixing organisms (White et al., 2007). Symbiotic nitrogen fixation is crucial to success of legumes, but the plant has to control the number of nodules it forms to balance the nitrogen gains with the developmental costs in order to avoid the severe energy drain that would be imposed by having to many nitrogen fixing nodules. The control of nodule number on the colonized root occurs via a complex systemic mechanism called “autoregulation of nodulation”, based on root-derived and shoot-derived signals (Kouchi et al., 2010). Furthermore legume regulates the nodule number in response to environmental nitrogen (nitrate or ammonia) availability to preferentially obtain this nutrient from sources energetically favourable relative to the cost of nodulation (Jeudy et al., 2009). In both determinate and indeterminate nodules bacteroids carry out an optimal N2-fixation for 4–5 weeks after infection. Beyond this period the bacteroid activity declines and a senescence process occurs in the N2-fixing zone, leading to degradation of the infected cells. Finally, bacteroids (or some undifferentiated bacteria) are released from decayed nodules to the soil where they can recolonize the rhizosphere resuming the free saprophytic lifestyle.

Frankia-Dicotyledon Symbiosis The aerobic Gram-positive actinomycetes belonging to the genus Frankia are diazotrophic bacteria that are capable of inducing formation of N2-fixing nodule lobes in roots of many dicotyledonous angiosperms. The plants nodulated by Frankia strains are

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Figure 5 A simplified diagram showing nitrogen fixation and ammonia assimilation in an infected cell of a legume root nodule. GOGAT, glutamate syntase; GS, glutamine syntetase; N2ase, nitrogenase.

Table 3

Association between Frankia and actinorhizal plants

Frankia phylogenetic groups

Plant families

Plant genera

Group I strains

Coriariaceae Datiscaceae Rosaceae Rhamnaceae Betulaceae Casuarinaceae Myricaceae Elaeagnaceae Rhamnaceae

Coriaria Datisca Cercocarpus, Chamaebatia, Dryas, Cowania, Purshia Ceanothus Alnus Casuarina, Allocasuarina, Ceuthostoma, Gymnostoma Myrica, Comptonia Elaeagnus, Hippophae¨, Shepherdia Calletia, Discaria, Kentrothamnus, Retanilla, Talguenea, Trevoa

Group II strains

Group III strainsa a

Group III strains are more promiscuous and can occasionally inhabit root nodules of Rosaceae, Coriariaceae and Ceanothus.

known as actinorhizal plants and include 8 families, 25 genera, and over 200 species, most of which are perennial woody shrubs or trees distributed in all landmasses except Antarctica. The actinorhizal plants share a predilection for marginally fertile soils and the majority are pioneers on nitrogen-poor sites. In addition, many actinorhizal species are able to tolerate environmental stresses such as heavy metals, high salinity, drought, cold, and extreme pH. They inhabit a variety of ecosystems, including coastal dunes, riparian zones, alpine communities, arctic tundra, glacial tills, and forests. Actinorhizal plants are especially important in high latitude regions, such as Scandinavia, Canada, Alaska, and New Zealand where Leguminosae are absent or rare while actinorhizal plants are abundant and capable of vigorous growth (Wall, 2000). Much of the new nitrogen entering these ecosystems comes from the actinorhizal symbioses that, on the whole, account for over 15% of the biologically fixed nitrogen worldwide. The filamentous frankiae, besides in symbiotic association with actinorhizal plants, can also occur as free-living diazotrophic organisms (Benson and Silvester, 1993). In pure culture, Frankia strains produce extensive hyphae and sporangia. In response to nitrogen deprivation, they also differentiate vesicles, named diazovesicles, which contain nitrogenase and are the site of N2 fixation. The diazovesicles are encapsulated by a series of laminated lipid layers that are rich in neutral lipids, glycolipids, and hopanoids. This envelope, whose thickness depends on the environmental O2 concentration, works as an oxygen-diffusion barrier, providing an anaerobic environment for nitrogenase to function inside vesicles (Berry et al., 1993). The Frankia strains that nodulated actinorhizal plants can be phylogenetically distinct in three groups (groups I, II, and III) that infect specific dicotyledon families (Table 3). Actinorhizal plants fall into families of three related orders: Rosales (Rosaceae, Eleagnaceae, Rhamnaceae), Fagales (Betulaceae, Casuarinaceae, Myricaceae) and Cucurbitales (Coriariaceae, Datiscaceae). Together with Fabales (legumes), they form a single “nitrogen-fixing clade” within the angiosperms (Soltis et al., 1995).

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Figure 6 Schematic drawing of a mature actinorhizal nodule lobe.

In the actinorhizal symbioses, root nodule formation begins with the host-symbiont recognition through the exchange of molecular signals, the knowledge of which is still limited. However, some findings suggest that the signaling mechanisms of Frankia-actinorhizal plants might be similar to those of rhizobia-legumes (Hocher et al., 2011b) and genomic analyses reinforce the hypothesis of a possible single origin for legumerhizobia and actinorhizal symbioses (Hocher et al., 2011a). Frankia strains can infect the host root by intracellular or intercellular mechanisms. Intracellular infection, such as that occurring in genera Myrica, Comptonia, Alnus, and Casuarina, starts with penetration of bacterial hyphae in a curled root hair. Afterward the hyphae move in cortical cells encapsulated with a layer of plant cell wall material surrounded by host plasmalemma. In intercellular infection, common in genera Elaeagnus, Ceanothus, and Cercocarpus, the bacterial hyphae penetrate between two adjacent rhizoderm cells and progress apoplastically through cortical cells encapsulated in a pectic matrix. Concomitantly, cell divisions induced in the root pericycle give rise to the nodule lobe primordium to which the hyphae move. The mature actinorhizal nodule lobe resembles a modified lateral root with an apical meristem but without a root cap. It shows a central stele with vascular tissues and has Frankia hyphae restricted to the cortical cells (Figure 6). In most actinorhizal symbioses, the N2-fixing activity of Frankia in infected cells is associated with differentiation of diazovesicles whose morphology is strictly controlled by the host plant. As in the free-living frankiae, these vesicles are surrounded by the multilayered lipid envelope and contain nitrogenase. However, in some symbioses (with plants of genera Myrica, Coriaria, Comptonia, and Casuarina), the Frankia hyphae proliferate without forming vesicles. The mature anatomy of a nodule lobe is reached at about 2 weeks after inoculation while the N2-fixation can be detected after three weeks (Huss-Danell, 1997). In infected cells of mature nodule lobes, some mechanisms take place to lower the oxygen tension near the site of the oxygenintolerant nitrogenase. The first diffusion resistance to oxygen is provided in diazovesicles by the multilayered envelope and a further reduction of the pO2 is obtained through their high respiration rate. In many nodule lobes devoid of diazovesicles, the infected cells contain high levels of hemoglobins that have homologous sequences to leghemoglobins and are believed to play the same role (Fleming et al., 1987). In these nodules, moreover, a low pO2 may be maintained by lignification of the host cell walls. Finally, the activity of uptake hydrogenases can also help to protect the nitrogenase against O2 in both hyphae and diazovesicles of the symbiotic frankiae (Leul et al., 2009). In free-living Frankia strains, as in the other free-living diazotrophs, the ammonia produced by N2 fixation is assimilated by the organism via the GS-GOGAT pathway. On the contrary, these enzymes are differently regulated in the symbiotic frankiae. In diazovesicles of root nodule lobes GS activity is very low and ammonia remains unassimilated (Alloisio et al., 2010). As in rhizobia–legume symbiosis, NH3 is released into the host cell where its assimilation gives rise to amino acids and other organic nitrogen compounds. Some are furnished to the bacterium, but most of them are transferred to the plant shoot. The scarcity or lack of GS activity in the diazotrophic symbiont also characterizes the rhizobia legumes as well as some cyanobacterial symbioses such as Anabaena–Azolla, showing a remarkable convergence of physiological strategies in the N2-fixing associations. The actinorhizal plant must provide photosynthates to the symbiotic bacterium. As in the rhizobia–legume symbiosis C4 dicarboxylates derived from sucrose metabolism occurring in the host cell are likely to be the carbon sources for Frankia strains in actinorhizal symbiosis. Furthermore, as it occurs in rhizobia–legume symbiosis, the actinorhizal plants can control infection by Frankia and regulate number and development of nodule lobes on roots by systemic autoregulatory processes (Wall et al., 2003).

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Endophytic Diazotrophic Bacteria–Cereal Association A recently discovered nitrogen-fixing association is that between grasses, such as sugar cane, maize, rice, wheat, sorghum, and other graminaceous species and endophytic diazotrophic bacteria that can colonize the plant interior without causing symptoms of disease. These bacteria enter the plants at root tips or at the emergence points of lateral roots and penetrate the root cortex, the stelar tissues, and the xylem vessels through which they may migrate toward the shoot (Cocking, 2003). Endophytic diazotrophic bacteria are generally restricted to intercellular spaces and especially to the xylem vessels where the low pO2 and the high bacterial respiration rate create the microaerobic conditions needed for nitrogenase activity. Some of these diazotrophic microorganisms, such as those belonging to the genera Acetobacter, Herbaspirillum, Azospirillum, and Azoarcus, are of extreme interest since they can significantly contribute to the nitrogen requirement of the graminaceous plants (Kennedy et al., 2004). Certain rice varieties, for instance, can obtain over 30% of their nitrogen from these endophytes (James et al., 2002) and some Brazilian sugarcane varieties up to 80%, with a total contribution of more than 170 kg N ha1 year1. (Urquiaga et al., 1992). Studies of these N2-fixing associations form a topical field of research whose aim is to explore the possibility of both enhancing the N2 fixation and extending this efficient system to other cereals. This would greatly reduce the use of nitrogen fertilizers with considerable economic benefits, and, above all, with enormous environmental advantages. Over two-thirds of arable lands, in fact, are dedicated to the growing of cereals, which provide 80% of the food for the world’s populations.

Nitrogen Fixation in Free-Living Cyanobacteria Among the free-living diazotrophs a prevailing interest is that addressed to cyanobacteria. This interest comes from the wide and abundant distribution of these microorganisms in all terrestrial and aquatic ecosystems as well as from their unique photosynthetic metabolism that makes the nitrogen fixation an apparently paradoxical event. Cyanobacteria, in fact, are the only prokaryotes that carry out oxygenic photosynthesis. A very great number of these microorganisms is able to both fix N2 under aerobic conditions and produce O2 by photosynthesis. Filamentous cyanobacteria resolve this oxygenic photosynthesis–diazotrophy paradox by segregating the oxygen-sensitive machinery for N2 fixation in specialized nonphotosynthetic cells named heterocysts, and by maintaining the oxygen evolving photosynthesis in the neighboring vegetative cells. Thus, the simultaneous operation of the two basically incompatible processes is made possible through their spatial separation. Nitrogen starvation leads to the appearance at regular intervals along the cyanobacterial filament of heterocysts which function as anaerobic factories for N2 fixation under external aerobic conditions. The ability to fix N2 ensues from changes that occur in vegetative cells that differentiate to heterocysts (Figure7).

Figure 7 Schematic drawing of a cyanobacterial heterocyst showing nitrogen fixation and metabolite exchange with the neighboring vegetative cell. Fdred, reduced ferredoxin; gluc6, glucose-6-phosphate; 6gluc, gluconate-6-phosphate; GOGAT, glutamate syntase; GS, glutamine syntetase; N2ase, nitrogenase; PSI, photosystem I; PSII, photosystem II; rib5P, ribulose-5-phosphate.

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First they build up a very thick cell wall with an innermost laminated glycolipid layer, whose function is to provide an O2 permeability barrier to avoid the inactivation of nitrogenase inside the cell (Nicolaisen et al., 2009). The connections between heterocyst and neighbor cell occur through thin cytoplasmic channels (microplasmodesmata), which traverse the septum separating the two cells and the plag (polar body) of cyanophycin filling the adjacent region (Mullineaux et al., 2008). In addition, the photosynthetic apparatus undergoes a deep reorganization during the heterocyst differentiation: phycobilisomes disappear and the oxygen-evolving photosystem II is totally dismantled, while photosystem I, which produces ATP through cyclic photophosphorylation, persists in thylakoid membranes (Wolk, 1996). The ATP necessary to fulfill the energy demand for nitrogenase activity, in fact, comes in heterocysts from cyclic photophosphorylation, while the reductant for the N2-fixing enzyme is furnished by neighboring photosynthetic cells probably as sucrose (Cumino et al., 2007). The sugar hydrolysis and glucose oxidation through the pentose phosphate pathway produces NADPH used for ferredoxin reduction. The heterocysts of free-living cyanobacteria contain high levels of glutamine synthetase (GS), but are deficient in GOGAT that, on the contrary, is active in vegetative cells. Thus, after N2 reduction, the NH3 assimilation in glutamine is carried out in the heterocyst while the successive reaction leading to glutamate synthesis occurs in the near vegetative cell into which glutamine moves through microplasmodesmata (Martin-Figueroa et al., 2000). A major role in protecting nitrogenase against O2 is played in heterocysts by the thick cell wall which prevents gas diffusion toward the cell. However, it is unlikely that this envelope provides a truly impermeable gas diffusion barrier, since this would also exclude nitrogen from the fixation site. Moreover, gases can reach the heterocyst cytoplasm through the junctions between them and the contiguous vegetative cells which produce O2 by photosynthesis. Thus, also cyanobacterial heterocysts, as the other aerobic diazotrophic organisms, need systems to remove oxygen that enters the cell. These include enhanced rate or respiration (Valladares et al., 2007), probable presence of hemoproteins in the cytoplasm peripheral region, and activity of an uptake hydrogenase (Tamagnini et al., 2007). A great ecological interest arose from the unexpected finding that unicellular and nonheterocystous filamentous cyanobacteria are also able to both fix nitrogen and carry out oxygenic photosynthesis. These cyanobacteria, which are very abundant in the phytoplanktonic populations of marine environments, are responsible for most of the photosynthetic organic carbon provided to the ecosystem (Karl et al., 2002), and they may also account for a high percentage of the nitrogen fixed biologically worldwide. The oxygenic photosynthesis–diazotrophy paradox is resolved by these nonheterocystous cyanobacteria through creative strategies enabling them to separate spatially or temporally the two incompatible processes. The marine filamentous cyanobacteria of genus Trichodesmium fix nitrogen during the light period in a subset of specialized nitrogenase-containing cells, named diazocytes, formed by cell division primarily confined to dark period. In addition, nitrogenase shows a diurnal pattern in which its activity is highest early in the day (Sandh et al., 2009). A number of unicellular diazotrophic cyanobacteria, such as those of genus Cyanothece, exhibit temporal separation of the two physiological processes that should necessarily occur in the same cell. They carry out only the oxygenic photosynthesis during the day and fix N2 only during the night, when the photosynthetic O2 production does not occur. This timing of N2 fixation is also related to the fact that the nitrogenase is active exclusively during the dark period. Interestingly, the daily oscillation of nitrogenase activity occurs according to an endogenous circadian rhythm (Toepel et al., 2008). The finding that the nitrogenase of nonheterocystous N2-fixing cyanobacteria possesses this kind of rhythmic activity was also of great scientific importance since it was the first clearly recognized circadian rhythm in prokaryotic organisms, which led to the backdrop of the former dogma that restricted the biological clocks to eukaryotes (Golden et al., 1997).

Diazotrophic Cyanobacteria–Plant Symbioses Many species of filamentous N2-fixing cyanobacteria, in the great majority of cases belonging to the genus Nostoc, can form symbiotic associations with a wide range of eukaryotic hosts, among which fungi (cyanolichens), microalgae (diatoms), corals, sponges and numerous plants. The cyanobacteria-plant symbioses include Bryophyta (liverworts, hornworts, and mosses), Pteridophyta (the genus Azolla), gymnosperms (Cycadaceae) and angiosperms (Gunneraceae) as hosts. The free-living cyanobacteria which can form these symbiotic associations share two major characteristics: they are able to differentiate heterocysts and to produce short motile filaments, known as hormogonia. Hormogonia, which provide a means of dispersal for otherwise immotile cyanobacteria, are the infective agents which enter the host plant tissues (Meeks and Elhai, 2002). The hormogonia, which are devoid of heterocysts, form in response to chemical signals (HIFs: hormogonia-inducing factors) released from the plant, that also produces chemoatractants to guide them into symbiotic cavities. Subsequent to the infection the host may release signals (HRFs: hormogonia-repressing factors) to prevent formation of further hormogonia and to shift the cyanobiont development towards heterocyst differentiation and N2 fixation (Rai et al., 2000). The frequency of heterocysts in filaments of the symbiotic cyanobacterium is several folds higher than in the same cyanobacterium in free-living state. However, only a low quantity of nitrogen fixed is retained by the cyanobiont, the remaining being transferred to the host plant as ammonia. Photosynthesis can be highly depresses in cyanobionts, relative to that in free-living strains, but the reduced CO2 fixation is compensated by sugars derived from the hosts (Meeks, 2003).

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Among the numerous cyanobacteria-plant symbioses, particular attention has been paid to that between the cyanobiont Anabaena azollae and the leaves of the aquatic fern Azolla. The interest for the Anabaena–Azolla association is mainly due to the potential use of Azolla as fertilizer in rice fields.

Anabaeana–Azolla Symbiosis Azolla is a small floating fern and is the only known pteridophyte that lives in symbiosis with a diazotrophic cyanobacterium. All the species of the genus harbour in their fronds a filamentous N2-fixing cyanobacterium until now referred as Anabaena azollae (Nostocaceae) (Papaefthimiou et al., 2008). The Azolla sporophyte (generally 0.5–7 cm in lenght but up to 40 cm in A. nilotica) consists of a multibranched rhizome generating, on the ventral surface, adventitious roots hanging down into the water to absorb nutrients directly. The rhizome bears small leaves (approximatively 1 mm in length) consisting of an aerial chlorophyllous dorsal lobe and a partially submerged colourless ventral lobe which is cup-shaped to provide buoyancy. Each dorsal lobe contains a specialized cavity housing the cyanobiont permanently. The mature cavity, ellipsoid in shape, has the interior surface covered by a mucilaginous layer delimited by an envelope, where 2000–5000 cyanobacterial cells are embedded and immobilized. Several trichomes, traditionally called hairs, protrude from the cavity surface into the mucilage layer and create an intimate contact between the symbiotic partners helping in the exchange of metabolites. Thus the leaf cavity can be considered as a natural microcosm with a self organization and an ecological defined structure. It behaves as both the physiological and dynamic interface unit of the symbiotic association where the main metabolic and energetic flows occur (Peters and Perkins, 1993; Rai et al., 2000a, 2000b). The diazotrophic cyanobacterium Anabaena azollae consists of unbranched filaments containing bead-like highly pigmented vegetative cells and lightly pigmented intercalary N2-fixing heterocysts. In the youngest leaves of the water fern the Anabaena filaments lack heterocysts while these gradually increase in frequency, relative to photosynthetic cells, reaching 30–40% of the cyanobacterial cells in Anabaena population of mature leaf cavities. This heterocyst frequency in Azolla-associated Anabaena (compared to 5–6% in other free-living species of Anabaena) is far higher than necessary to support the fixed N needs of the symbiotic cyanobacterium. Moreover, an overall decrease (up to 70%) in GS activity in the Azolla cyanobiont has also been shown, that limits the ammonia assimilation in heterocysts. Thus, some 50–90% of fixed nitrogen is delivered by Anabaena to the fern as ammonia. Translocation of fixed N from the symbiotic environment of mature cavities to other parts of the host plant occurs in the form of amino acids. Concomitantly with the differentiation of Anabaena into a higher proportion of nitrogen-fixing heterocysts, relative to the photosynthetic cells, a reduction of photosynthesis and CO2 fixation occurs in cyanobacterial filaments, which are supplied with sucrose by the host plant (Chapman and Margulis, 1998). The Anabaena–Azolla symbiosis is perpetual and hereditary and the symbiotic condition can be described as obligate for the cyanobiont whose free-living form is not found in nature. The symbiosis is maintained during all the life cycle of the pteridophyte throughout both sexual and asexual reproduction without requiring fresh infection from the environment. In contrast with other plant-cyanobacterial symbioses, Azolla hormogonium initiation factors (HIFs) are unknown. A colony of Anabaena is associated with each fern shoot apex and, as the plant grows, the cyanobacterial filaments are partitioned off into each new leaf. In the Azolla–Anabaena symbiosis, the cyanobiont growth is synchronized with that of the host plant. The growth rates of both partners are highest in the apical parts of the fern and decrease along the axis away from the apex. When growth of Azolla stops, cyanobiont cells cease to divide (Lechno-Yossef and Nierzwicki-Bauer, 2002). The water fern Azolla naturally occurs on lake surfaces, slow-moving rivers, canals, ponds, and ditches in warm-temperate to tropical climates, but its world distribution has been enlarged by humans. In fact, the Anabaena–Azolla association has been shown to be of major agronomic importance for its potentiality as a biofertilizer to substitute chemical nitrogen compounds. Azolla has been used as “green manure” in several countries to fertilize rice paddies and to increase rice yields (van Hove and Lejeune, 2002). Azolla–Anabaena is capable of fixing nitrogen at higher rates than legumes and is able to growth successfully in waterlogged habitats having low level of nitrogen. The Asians have recognized benefits of Azolla on rice cultivation first, since both rice crop and fern require similar environmental growing conditions. In rice fields, for instance, it can fix over 1 kg N ha1 day1, providing sufficient nitrogen to allow sustainable rice cultivation. Increase from 14% to 40% in grain yield of rice has been reported with Azolla used as dual crop. Moreover, Azolla has a high rate of multiplication, which helps in covering very rapidly the surface of water bodies where it is growing. Thus the thick mat produced helps to reduce the volatilization of ammonia in rice fields. The exploitation of this system in temperate environments is limited due to the Anabaena–Azolla sensitivity to low winter temperatures and to alternating day/night temperatures in spring before the rice sowing period (Pabby et al., 2003). Recently, however, Azolla strains have been selected as practicable biofertilizer even in the high latitude of the temperate rice areas, such as those of Northern Italy (Bocchi and Malgioglio, 2010). Besides its extensive use as a N-supplement in rice-based ecosystems, it has also been utilized in other crop cultivations such as taro, wheat, tomato, and banana. Furthermore, the Anabaena–Azolla association is also applied as controlling agent for weeds and mosquitos, due to its ability to cover water surfaces, and for improving water quality for its properties of removing excess quantities of nitrate and phosphorous.

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Cyanolichens Lichens are symbiotic associations between fungi (mycobionts, commonly Ascomycetes) and photosynthetic partners (photobionts) which can be green algae (commonly Trebouxia spp.) or diazotrophic cyanobacteria (commonly Nostoc spp.). These mycobiont-photobiont symbioses are regarded as mutualistic. The photoautotroph partners provide photosynthetic products and, in the case of cyanobacteria, also organic nitrogen compounds to the fungus. This latter, in turn, provides shelter for the photobionts by enclosing them within the thallus. In this way the photosynthetic organisms, protected from drying and strong sunlight, are allowed to growth in very harsh conditions. Lichens can tolerate the most extreme environments on Earth. They can live in hot deserts and arctic regions, on steril soils, bare rocks, wood debris and epiphytes on tree trunks, branches and leaves. Recently it has been reported that lichens are able to survive exposure to space conditions and even to adapt to the harsh Mars environment (de Vera, 2012; Raggio et al., 2011). Lichens can reproduce asexually through the dispersal of small fragments of thallus or through propagules (diaspores) tipically containing cells from the symbiotic partners. Bipartite lichens are symbiotic associations of a fungus with a single photosynthetic partner, which in so-called chlorolichens is a green alga while in lichens defined as cyanolichens is a diazotrophic cyanobacterium. In these latter associations the cyanobacterium may be localized in a distinct layer or dispersed through the thallus. As the sole photobiont, it is responsible for provision of both fixed carbon and fixed nitrogen to the mycobiont. The cyanobacterium maintains high photosynthetic levels and N2-fixing activity, with heterocyst frequency (7–8%) close to its free-living counterpart. In tripartite lichens the fungus is associated with both a green alga (phycobiont) and a cyanobacterium (cyanobiont). In these cyanolichens (also referred to as cephalolichens) the phycobiont is widely distributed through the thallus, while the cyanobiont is confined to special structures, named cephalodia. In cephalodia the cyanobiont becomes predominantly heterotrophic and is specialized for nitrogen fixation, as shown by the increase of heterocyst proportion over 30% of cyanobacterial cells. Thus, in these tripartite cyanolichens the fixed carbon is provided to the fungus by the phycobiont, while the cyanobiont supplies it with the fixed nitrogen.

A Swift Ecological Overview of Biological Nitrogen Fixation in Terrestrial and Aquatic Ecosystems Nitrogen is an essential element for all living organisms and the nitrogen supply is crucial for the ecology and biogeochemistry of terrestrial and aquatic ecosystems. A number of both physical (e.g., the dissolved nitrate exiting with percolating waters) and biological (e.g., denitrification and anammox) processes tend to restrict the biological nitrogen availability in the ecosystems. Apart from a minor contribution by lighting, the biological nitrogen fixation is the process which provides the largest source of new combined nitrogen to natural environments for replenishing the nitrogen losses. However, the fixation rates vary widely across the different ecosystems where symbiotic and free-living diazotrophic microorganisms can be differently involved in the process. Although the biological fixation is the major nitrogen input into terrestrial and aquatic ecosystems, currently there is only limited understanding about the factors that regulate this process in natural environments. In arid lands of the world, from arctic ecosystems (Stewart et al., 2011) to hot deserts of Southern Africa (Bu¨del et al., 2009), much of the nitrogen input via biological nitrogen fixation relies heavily on diazotrophic microorganisms, mostly cyanobacteria, living in biological soil crusts (BSCs). The BSCs, also referred to as cryptogamic crusts, are microbic communities (millimeters to centimeters thick) covering large portions of the dryland soils (more than 70% in untouched deserts of North America). BSCs are composed of fungi, cyanobacteria and microalgae, either as free-living organisms or as partners of lichen symbioses. Also symbiotic cyanobacteria-bryophyte associations can occur in well developed BSCs, which substantially contribute to available nitrogen increase in these extreme environments. Biological nitrogen fixation is limited in arctic and alpine tundra as well in enormous extended boreal forests of Eurasia and North America. The majority of nitrogen accretion in these environments occurs through cyanobacteria either free-living or symbiotic in cyanolichens or associated with bryophytes. The boreal forests lack significant quantities of plants forming root symbioses with diazotrophic bacteria (referred to as N2-fixing plants), with the exception of actinorhizal forms, among which species of Alnus and Ceanothus and of the coushion-forming dwarf shrubs Dryas. However, these fast-growing pioneer plants substantially contribute to nitrogen input only in early successional forests that develop after some form of disturbance, such as wildfire, windstorm or logging. In late successional and old-growth forests (more than 100–150 years since the last disturbance), instead, dominant sources of fixed nitrogen are communities of cyanobacteria (Nostoc spp.) associated with feather mosses (Pleurozium schrebery and Hylocomium splendens) that contribute approximatively 2 kg N ha1 year1 (DeLuca et al., 2002). The feather mosses, which account for as much as 95% of ground cover of the boreal forest floor, and their associated cyanobacteria may be the most broadly distributed N2-fixing association in Earth and are considered the primary source of nitrogen fixation in boreal ecosystems (Gundale et al., 2012) which are the second largest biome in the world. Many overstory and understory species of N2-fixing genera (e.g., Alnus, Robinia, Ceanotus, Myrica, Lupinus) can be present in temperate forests, accounting for high rates of nitrogen input in the environment (even more than 100 kg N ha-1 year1). However, like in boreal ones, these plants are prominent only in early successional stage of temperate forests. In mid and late successional forests the N2-fixer plants become rare, being totally absent in old-growth forests. Failing N2-fixing plants, the available nitrogen input into the ecosystem relies essentially on epiphytic cyanolichens (Antoine, 2004) and on nonsymbiotic N2 fixation that can account for over 1 kg N ha1 year1 and increases in magnitude toward mid, late and old-grown successional stages of forest. The

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nonsymbiotic N2 fixation is carried out by heterotrophic bacteria living on decomposing leaf litters or woody debris of forest floor, where reduced forms of carbon are available as energy sources for the energetically expensive microbial process (Pe´rez et al., 2010). Biological nitrogen fixation rates are very high in tropical rain forests that may fix more N2 than any other unmanaged ecosystem (even more than 200 kg N ha1 year1). Legumes are abundant in many of these forests, representing in some of them more than 50% of all trees. However, although these plants can be a major source of nitrogen input into the environment, their abundance may lead to overestimate the potential for N2-fixation. Not all leguminous trees, in fact, are able to establish an effective symbiosis with rhizobia or may not do so under natural conditions (Pons et al., 2007). High legume nodulation levels and N2 fixation rates occur in recently disturbed forests and in forests subjected to seasonal flooding, which have N-poor soils. Nodulation and N2 fixation, instead, are virtually absent from N-rich soils of undisturbed late successional and old-growth forests, despite the abundance of N2-fixing leguminous species. This is due to the ability of many symbiotic legumes to employ a strategy of facultative nitrogen fixation. These plants can induce N2 fixation in N-poor environments and down-regulate it to low or negligible levels in response to increased soil nitrogen availability (Barron et al., 2011). Therefore, the N2-fixing trees can act to rapidly redress any local nitrogen deficiencies that develop within tropical landscapes. Despite the lack of nitrogen fixation by tree in N-rich environmental conditions, substantial amounts of nitrogen enter the tropical forest ecosystem through several other diazotrophic organisms. Heterotrophic nonsymbiotic bacteria living on organic litter layers that cover the forest floor can bring about an input of available nitrogen up to 12 kg N ha1 year1. Canopy communities, dominated by autotrophic organisms, such as free-living cyanobacteria, cyanolichens and cyanobacteria-bryophyte associations can also contribute significant fluxes of nitrogen (up to 5 kg N ha1 year1) via biological fixation to the tropical rain forest ecosystems (Reed et al., 2008). Canopy cyanolichens and other epiphyte diazotrophic communities are a source of fixed nitrogen also in tropical dry forests, where the greatest contribution to available nitrogen input is given by actinorhizal tree species (e.g., those of genus Casuarina). Leguminous (e.g., Mimosa, Calliandra, Leucaena, Prosopis) and actinorhizal (e.g., Casuarina) genera are widespread in xeromorphic and arid woodlands from North and South America to West Africa and Ceanothus shrubs are common in European and American mediterranean shrublands. All these N2-fixing plants can greatly contribute to the nitrogen supply to their ecosystems (Cleveland et al., 1999). However, also in these environments an additional input of fixed nitrogen is provided by other symbiotic and nonsymbiotic diazotrophic communities. Legumes are not abundant in most grasslands and temperate savannas (from prairies of North America to pampas of South America and steppes of Eurasia) and contribute little available nitrogen to these environments that essentially rely on diazotrophic microorganisms of the soil (cyanobacteria and eterotrophic bacteria) as sources of fixed nitrogen. Conversely, N2fixing plants seem to be the largest source of available nitrogen in some tropical savannas. High proportions of legume trees, such as species of Acacia, are commonly present in tropical savannas that take up almost the half of the African continent, whereas Cycads (with symbiotic Nostoc spp. or Anabaena spp. in so called coralloid roots) form large populations in tropical savannas of Northern Australia. An estimate of biological nitrogen fixation, however, may be difficult in some of these ecosystems, due to the fact that they are often managed and contain N2-fixing forage plants (Cadish et al., 1994). Biological nitrogen fixation is an essential process in regulating biological productivity of freshwater and marine ecosystems that are most frequently under N-limited conditions also due to the worldwide presence of anammox bacteria whose activity accounts for up to 50% of the N2 gas released from these environments (Jetten et al., 2009). The major N2-fixers in the aquatic ecosystems are cyanobacteria. They are present as planktonic or benthic forms of nonheterocystous filamentous cyanobacteria, unicellular cyanobacteria and free-living and symbiotic heterocystous filamentous cyanobacteria. Recently, a significant role in N2 fixation has also been assigned to heterotrophic bacteria (Halm et al., 2012). Nitrogen fixation by planktonic cyanobacteria is rather low in oligotrophic lakes while the process may reach consistent levels in the mesotrophic and eutrophic ones. In these latter ecosystems the N2 fixation depends on some environmental factors including light intensity and phosphorus concentration. Phosphorus is often the limiting nutrient in lakes and diazotrophic cyanobacteria become abundant in planktonic communities only when phosphorus levels increase, leading to low nitrogen/phosphorus ratios. Benthic diazotrophic cyanobacteria also contribute the input of available nitrogen into lakes. They can develop extensive mats of filamentous species (such as Anabaena spp. and Oscillatoria spp.) on sediments, or they can occur as epilithic peryphyton forms (such as Nostoc spp., Calothrix spp., and many others). Extensive mats of diazotrophic cyanobacteria are also present in N-poor desert streams of western North America (Grimm and Petrone, 1997) which are characterized by warm temperature, high light, slow currents and ample supply of P. These environmental conditions support abundant cyanobacterial populations that can account for very high rates of N2 fixation (up to 150 mg N m2 day1). High current velocity, low light or high turbidity, instead, can limit growth of benthic and planktonic cyanobacteria and N2 fixation rate in flowing water systems, in most of which these diazotrophic microorganisms do not occur at all. Moreover, great turbulence, low availability of trace elements (in particular molybdenum) and also grazing by zooplankton (that breaks down diazotrophic filaments preventing the accumulation of enough photosynthetic cells to support the energetic requirement by heterocysts) are among the causes that negatively affect growth of cyanobacteria and N2 fixation in most temperate estuaries (Marino et al., 2006). N2-fixing heterotrophic bacteria have never been found in any of these freshwater ecosystems. Oceans are oligotrophic environments that make up 71% of the Earth’s surface. The South Pacific, which is the largest ocean system in the world, and the other tropical and subtropical oceanic gyres are regarded as biological deserts because of the extremely low availability of nutrients and minimum productivity. The only biological source of available nitrogen in all the oceanic

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ecosystems is the N2 fixation carried out by different types of cyanobacteria and heterotrophic bacteria (more than 100 million tons N per year). The most representative diazotrophs of North Atlantic Ocean, also abundant in the Arabian Sea and in the nearby Red Sea, are nonheterocystous filamentous cyanobacteria of genus Trichodesmium, which form large surface colonies at water temperature above 25  C. In North Pacific Ocean and in equatorial Pacific, instead, the dominant diazotrophs are small (<10 mm) unicellular coccoid cyanobacteria (e.g., Cyanoteche spp. and Crocosphaera spp.) together with heterotrophic bacteria (Halm et al., 2012). Heterocystous filamentous cyanobacteria (e.g., Nodularia spp. and Aphanizomenon spp.) are the N2 fixers living in the cooler brackish waters of the Baltic Sea. Although these free-living cyanobacteria are regarded as dominant diazotrophs in the oceanic environments, other N2-fixers also deserve consideration. Most notably, cyanobacteria-diatom symbioses capable of high N2 fixation rates, which are widely distributed through the warm oceans. Some genera of diatoms (e.g., Rhizosolenia, Hemiaulus and Guinardia), which are common members of the phytoplankton communities, form symbiotic associations with heterocystous filamentous cyanobacteria (e.g., Richelia intracellularis and Calothrix spp.) and represent a significant component of the nitrogen budget in these ecosystems (Foster et al., 2011). Planktonic diazotrophs are the only N2 fixers living in open oceans, while in coastal marine environments, such as salth marshers, intertidal zones and coral reefs, benthic filamentous cyanobacteria (e.g., heterocystous Calothrix and Anabaena and nonheterocystous Lyngbya and Oscillatoria genera) colonize sediments forming extensive mats that make substantial contributions to nitrogen supply to the ecosystem. The highest rates of N2 fixation in coastal environments are exhibited by the coral reef communities. Most of the fixed nitrogen entering the coral reef is provided by both free-living diazotrophs and benthic cyanobacteria that cover large areas of reef substratum. Additionally, several important members of the reef community, including sponges and corals, have also the capability to fix nitrogen through symbiotic associations with N2-fixing cyanobacteria and eterotrophic bacteria. Corals, in particular, are holobionts, with the coral animals that harbor a variety of microorganisms, including endosymbiotic dinoflagellates (Symbiodinium spp.), commonly referred as zooxanthellae, which provide over 95% of fixed carbon to the hosts. In addition, corals harbor endosymbiotic diazotrophic bacteria and cyanobacteria (e.g., Synechococcus and Prochlorococcus strains) that benefit with fixed nitrogen both zooxanthellae and coral hosts which possess enzymes enabling ammonium assimilation (Yellowlees et al., 2008). Interestingly, the symbiotic diazotrophic bacteria of many coral reefs, including the Great Barrier Reef, are closely related to bacterial species belonging to the order Rhizobiales (Lema et al., 2012). It is still unclear how the coral rhizobia might be protected against high concentrations of oxygen arising from the zooxanthella photosynthesis in host tissues. Possibly the diazotrophic communities are sheltered in oxygen-depleted coral microhabitats.

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Biological Nitrogen Fixation

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Further Reading Adams DG and Duggan PS (2008) Cyanobacteria-bryophyte symbioses. Journal of Experimental Botany 59: 1047–1058. Adams DG, Bergman B, Nierzwicki-Bauer SA, Rai AN, and Schu¨ßler A (2006) Cyanobacterial plant symbioses. In: Dworkin M, Falkow S, Rosengerg E, Schleifer K-H, and Stackebrandt E (eds.) The prokariotes: a handbook on the biology of bacteria. Symbiotic associations, biotechnology, applied microbiology 3rd edn., 1: pp. 331–363. New York: Springer. Benson DR, Brooks JM, Huang Y, et al. (2011) The Biology of Frankia sp. strains in post-genome era. Molecular Plant-Microbe Interactions 24: 1310–1316. Berry AM, Mendoza-Herrera A, Guo Y-Y, et al. (2011) New perspectives on nodule nitrogen assimilation in actinothizal symbioses. Functional Plant Biology 38: 645–652. Boys ESM, Hamiltion TL, and Peters JW (2011) An alternative path for the evolution of biological nitrogen fixation. Frontiers in Microbiology 2: 205. Church JG, Short CM, Jenkins BD, Karl DM, and Zehr JP (2005) Temporal patterns of nitrogenase gene (nifH) expression in the oligotrophic North Pacific Ocean. Applied and Environmental Microbiology 71: 5362–5370. Daalsgaard T, Thamdrup B, and Canfield DE (2005) Anaerobic ammonium oxidation (anammox) in the marine environment. Research in Microbiology 156: 457–464. Fiore CL, Jarret JK, Olson ND, and Lesser MP (2010) Nitrogen fixation and nitrogen transformations in marine symbioses. Trends in Microbiology 18: 455–463. Franche C, Lindstro¨m K, and Elmerich C (2009) Nitrogen-fixing bacteria associated with leguminous and non-leguminous plants. Plant and Soil 321: 35–59. Hedin LO, Brookshire ENJ, Menge DNL, and Barron AR (2009) The nitrogen paradox in tropical forest ecosystems. Annual Review of Ecology, Evolution, and Systematics 40: 613–635. Herridge DF, Peoples MB, and Boddey RM (2008) Global input of biological nitrogen fixation in agricultural systems. Plant and Soil 311: 1–18. Hirsch AM, Lum MR, and Downie JA (2001) What makes the rhizobia–legume symbiosis so special? Plant Physiology 127: 1484–1492. James EK (2000) Nitrogen fixation in endophytic and associative symbiosis. Field Crops Research 65: 197–209. Johnson CH, Mori T, and Xu Y (2008) A cyanobacterial circadian clockwork. Current Biology 18: R816–R825. Jones KM, Kobayashi H, Davies BW, Taga ME, and Walker GC (2007) How rhizobial symbionts invade plants: the Sinorhizobium-Medicago model. Nature Reviews Microbiology 5: 619–633. Kereszt A, Mergaert P, and Kondorosi E (2011) Bacteriod development in legume nodules: evolution of mutual benefit or of sacrifical victims? Molecular Plant-Microbe Interactions 24: 1300–1309. Kumar K, Mella-Herrera RA, and Golden JW (2009) Cyanobacterial heterocysts. Cold Spring Harbor Perspectives in Biology 2: a000315. Magnani GS, Didonet CM, Cruz LM, et al. (2010) Diversity of endophytic bacteria in brazilian sugarcane. Genetics and Molecular Research 9: 250–258. Markmann K and Parniske M (2009) Evolution of root endosymbiosis with bacteria: how novel are nodules? Trends in Plant Science 14: 77–86. Massena Reis V, Baldani JI, Divan Baldani VL, and Dobereiner J (2000) Biological dinitrogen fixation in gramineae and palm trees. Critical Reviews in Plant Sciences 19: 227–247. Murray JD (2011) Invasion by invitation: rhizobial infection in legumes. Molecular Plant-Microbe Interactions 24: 631–639. Oldroyd GED and Downie JA (2008) Coordinating nodule morphogenesis with rhizobial infection in legumes. Annual Review of Plant Biology 59: 519–546. Pawlowski K and Demchenko KN (2012) The diversity of actinorhizal symbiosis. Protoplasma 249: 967–979. Pawlowski K and Sirrenberg A (2003) Symbiosis between Frankia and actinorhizal plants: root nodules of non-legumes. Indian Journal of Experimental Biology 41: 1165–1183. Pawlowski K, Bogusz D, Ribeiro A, and Berry AM (2011) Progress on research on actinorhizal plants. Functional Plant Biology 38: 633–638. Perrine-Walker F, Gherbi H, Imanishi L, et al. (2011) Symbiotic signaling in actinorhizal symbioses. Current Protein and Peptide Science 12: 156–164. Reid DE, Ferguson BJ, Hayashi S, Lin Y-H, and Gresshoff PM (2011) Molecular mechanisms controlling legume autoregulation of nodulation. Annals of Botany 108: 789–795. Schumpp O and Deakin WJ (2010) How inefficient rhizobia prolong their existence within nodules. Trends in Plant Science 15: 189–195. Seefeldt LC, Hoffman BM, and Dean DR (2009) Mechanism of Mo-dependent nitrogenase. Annual Review of Biochemistry 78: 701–722. Sessitsch A, Hiwieson JG, Perret X, Antoun H, and Martinez-Romero E (2003) Advances in Rhizobium research. Critical Reviews in Plant Sciences 21: 323–378. Sohm JA, Webb EA, and Capone DG (2011) Emerging patterns of marine nitrogen fixation. Nature Reviews Microbiology 9: 499–508. Taule´ C, Mareque C, Barlocco C, et al. (2012) The contribution of nitrose fixation to sugarcane (Saccharum officinarum L.), and the identification and characterization of part of the associated diazotrophic bacterial community. Plant and Soil 356: 35–49. Vitousek PM, Cassman K, Cleveland C, et al. (2002) Towards an ecological understanding of biological nitrogen fixation. Biogeochemistry 57: 1–45. Wasson MF (1999) Global patterns of terrestrial biological nitrogen (N2) fixation in natural ecosystems. Global Biogeochemical Cycles 13: 623–645. Zhang CC, Laurent S, Sakr S, Peng L, and Be´du S (2006) Heterocyst differentiation and pattern formation in cyanobacteria: a chorus of signals. Molecular Microbiology 59: 367–375.