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9Biotic Influences

9A. Symbiotic Associations

1. Introduction

Symbiosis is the ‘‘living together’’ of two or moreorganisms. In its broadest sense, symbiotic associa-tions include parasitic and commensal as well asmutually beneficial partnerships. As is common inthe ecophysiological literature, however, we use theterm symbiosis in a narrow sense to refer tomutually beneficial associations between higherplants and microorganisms. Mutual benefits maynot always be easy to determine, particularly forthe microsymbiont. In this chapter benefits for themacrosymbiont (‘‘host’’) are often expressed interms of ability to accumulate biomass. In an ecolo-gical context, benefits in terms of ‘‘fitness’’ may bemore relevant but are rarely documented. In themutually beneficial associations discussed in thischapter, nutrients or specific products of the part-ners are shared between two or three partners; themacrosymbiont and the microsymbiont(s). Parasiticassociations between higher plants are dealt with inChapter 9D; parasitic associations between micro-organisms and higher plants are discussed briefly inthis chapter, and more elaborately in Chapter 9C oneffects of microbial pathogens.

In Chapter 6 on mineral nutrition, we discussednumerous special mechanisms that allow somehigher plants to acquire sparingly soluble nutrientsfrom soils (e.g., excretion of carboxylates and phyto-siderophores). We also pointed out (Sects. 2.2.5 and2.2.6 of Chapter 6 on mineral nutrition) that somespecies are quite capable of growing on soils where

P is sparingly available, without having a large capa-city to excrete protons or carboxylates. How do theseplants manage to grow? It is also obvious that specialmechanisms to take up nutrients (e.g., N) are of littleuse, if the N is simply not there. Such plants musthave alternative ways to acquire N.

This chapter discusses associations betweenhigher plants and microorganisms that are of vitalimportance for the acquisition of nutrients. Suchsymbiotic associations play a major role in environ-ments where the supply of P, N, or immobile cationslimits plant growth. In the rhizosphere (or else-where in the plant’s immediate surroundings),mycorrhiza-forming fungi and N2-fixing bacteriaor cyanobacteria may form symbiotic associations.For those species that are capable of such symbioses,it tends to be profitable for both the higher plant(macrosymbiont) and the microorganism (micro-symbiont). Indeed, it is so profitable for themacrosymbiont that some plants are associatedwith more than one microsymbiont at the same time.

2. Mycorrhizas

Vast majority of higher plant species can form sym-biotic associations with mycorrhizal fungi. Mycor-rhizas are the structures arising from the associationof roots and fungi; except for nonmycorrhizal species(Sect. 2.2), roots in soil should be considered inconjunction with their mycorrhizal symbionts. Thereare four main types of mycorrhizas (Sect. 2.1; Smith &

H. Lambers et al., Plant Physiological Ecology, Second edition, DOI: 10.1007/978-0-387-78341-3_9,� Springer ScienceþBusiness Media, LLC 2008

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Read 2008). The most ancient type dates back tothe early Devonian, some 400 million years ago(Nicholson 1975, Cairney 2000). The first bryophyte-like land plants had endophytic associationsresembling arbuscular mycorrhizas (AM), evenbefore roots evolved. The ectomycorrhizal symbiosishas evolved repeatedly over the last 130—180 millionyears (Martin et al. 2001). Like root hairs (Sects. 2.2.1and 2.2.5 of Chapter 6 on mineral nutrition), themycorrhizal associations enhance the symbioticplant’s below-ground absorbing surface. For somemycorrhizas (Table 1), this is the primary mechanismby which mycorrhizal plants are able to acquirescarcely available, poorly mobile nutrients, especiallyP (Sect. 2.1). For other mycorrhizas, additionalmechanisms, such as excretion of hydrolytic enzymesand carboxylates, may also play a role (Sect. 2.1).

The mycorrhizal associations generally enhanceplant growth, especially when P or other immobilenutrients limit plant growth; they may also bebeneficial when water is in short supply and suppressinfection by parasitic plants (Sect. 2.1 of Chapter 9D onparasitic associations). As such, mycorrhizas are ofgreat ecological and agronomic significance. Whenthe nutrient supply is high, however, they are a poten-tial carbon drain on the plant, providing less benefit inreturn. Plants, however, may have mechanisms to

suppress the symbiotic association at a high supplyof P (Sect. 2.3.1).

Some species never form a mycorrhizal associa-tion, even when P is in short supply. Some of these(e.g., Proteaceae; Sect. 2.2.5.2 of Chapter 6 onmineral nutrition) perform well when P is severelylimiting; other nonmycorrhizal plants may even beharmed by mycorrhizal fungi (Sect. 7 of Chapter 9E).On the other hand, some nonmycorrhizal plantsseverely inhibit the growth of mycorrhizal hyphae.Before dealing with these complex interactions(Sect. 2.2), we discuss some general aspects ofmycorrhizal associations.

2.1 Mycorrhizal Structures: Are TheyBeneficial for Plant Growth?

Mycorrhizas occur in 82% of all angiosperm speciesinvestigated to date; all gymnosperms are mycor-rhizal (Brundrett 2002). A mycorrhizal associationconsists of three vital parts:

1. The root2. The fungal structures in close association with

the root3. The external mycelium growing in the soil

TABLE 1. The length of mycorrhizal hyphae per unit colonized root length asmeasured for a number of plant species, infected with different arbuscularmycorrhiza-forming fungal species.

Fungus Host Hyphal length (m cm–1 root)

Glomus mosseae Allium cepa (onion) 0.79–2.5Glomus mosseae Allium cepa 0.71Glomus macrocarpum Allium cepa 0.71Glomus microcarpum Allium cepa 0.71Glomus sp. Trifolium sp. (clover) 1.29Glomus sp. Lolium sp. (ryegrass) 1.36Glomus fasciculatum Trifolium sp. 2.50Glomus tenue Trifolium sp. 14.20Gigaspora calospora Allium cepa 0.71Gigaspora calospora Trifolium sp. 12.30Acaulospora laevis Trifolium sp. 10.55

Source: Various authors, as cited in Smith & Gianinazzi-Pearson (1988).

FIGURE 1. (A) Schematic structure of an arbuscularmycorrhiza (AM). (B) Arbuscules (A) and a vesicle (V)of Glomus sp. colonized on the root of Tagetes patula(marigold). (C) Arbuscules (A) and intercellular hyphae(IH) of Glomus etunicatum isolated from the root ofTagetes patula after enzymatic digestion. (D) Detail of

the intraradical hyphae of Glomus mosseae in a root ofTagetes patula (marigold), after enzymatic digestion ofthe root, showing fine branches and trunks of an arbus-cule (Ezawa et al. 1995) (courtesy T. Ezawa, Faculty ofHorticulture, Nagoya University, Japan).

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Mycorrhizas 405

Mycorrhizas are classified in different types, butsome are very similar (Brundrett 2002). Arbuscularmycorrhizas (AMs) are the most widespread.A large fraction of the fungal tissue is within rootcortical cells, outside their plasma membrane(Fig. 1). They frequently occur on herbaceous plants,but are also found on trees, especially in tropical

forests. Ericoid mycorrhizas in the Ericaceae andorchid mycorrhizas in the Orchidaceae have some-what different structures and functions (Fig. 2).Alnus (alder), Cupressus (cypress), Eucalyptus (euca-lypt), Fraxinus (ash), Populus (poplar), and Salix(willow) are genera that have both AMs andectomycorrhizas (ECMs) (Fig. 3; Brundrett 2002).

FIGURE 2. (A) Orchidaceous mycorrhizal association in astem of Pterostylis sanguinea (greenhood orchid). (B)Transverse section of a stem of Caladenia arenicola(carousel spider orchid) showing intracellular fungalcoils (courtesy A.L. Batty and M.C. Brundrett, The Uni-versity of Western Australia, Australia). (C) Ericoidmycorrhizal association of Woollsia pungens, showingepidermal cells colonized by coils of an ericoid

mycorrhizal fungus (stained blue, arrowed) (courtesyS. Chambers and J.W.G. Cairney, Centre for Plant andFood Science, University of Western Sydney, Australia;copyright Elsevier Science, Ltd.). Ericoid mycorrhizalassociation of Leucopogon verticillatus (courtesy M.C.Brundrett, The University of Western Australia,Australia).

FIGURE 3. (A) Schematic representation of an ectomycor-rhiza, showing the fungal mantle around the root and thehyphae in the cortex, which form the Hartig net. (B–D)Ectomycorrhizal association between Pinus resinosa(red pine) and an unknown fungal species. A highermagnification of Pinus resinosa and Pisolithus tinctoriusas the mycobiont, showing thickened branched rootlets,

covered in a fungal mantle, and external hyphae. Thehighest magnification is of a longitudinal section of aPinus resinosa–Pisolithus tinctorius mycorrhizal rootshowing mantle hyphae on the root surface and Hartignet hyphae surrounding epidermal and cortical cells(courtesy R.L. Peterson, University of Guelph, Canada).


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Less than 200 fungal species form AM; they arethe most widespread mycorrhizal association(Nicholson 1975, Brundrett 2002). AMs are classifiedin six genera within the Glomeromycota, with Glo-mus being the largest genus. These fungi are consid-ered to be ‘‘primitive’’ because they have relativelysimple spores and they associate with a wide rangeof plant species. They are not capable of growingwithout a plant host. The AMs have been namedafter the arbuscules (which are tree-like structuresthat occur inside root cortical cells; Fig. 1). Althoughthe arbuscule can fill most of the cell space, it doesnot compromise the integrity of the plant plasmamembrane, because cortical cells envelop the arbus-cule in a specialized host membrane, the periarbus-cular membrane (Javot et al. 2007). The roots of 80%of all surveyed plant species and 92% of all familiescan be infected with AM-forming fungi (Wang &Qiu 2006). Even species that are typically ectomy-corrhizal form AM associations in the absence ofectomycorrhizal inoculum.

In ericoid and orchid mycorrhizas, like in AMs, alarge fraction of the fungal tissues is within the rootcortical cells. In ectomycorrhizas, however, mostfungal tissue is outside the root. This symbioticassociation is frequently found between trees (Dipter-ocarpaceae: 98%; Pinaceae: 95%; Fagaceae: 94%;Myrtaceae: 90%; Salicaceae: 83%; Betulaceae: 70%;Fabaceae: 16%) and more than 5000 species thatbelong to the Basidiomycota (agarics, bolets) or Asco-mycota (truffles) (Barker et al. 1998, Martin et al. 2001).Fossil ectomycorrhizas have been found amongplant remains of Pinus (pine) from the Middle Eocene40 million years ago (LePage et al. 1997). Althoughectomycorrhizas occur mostly in woody angios-perms and Pinaceae, they have also been found insome monocotyledons and in ferns. Ectomycorrhizasindependently evolved many times through parallelevolution. Co-evolution between plant and fungalpartners in ECM has probably contributed to diver-sification of both plant hosts and fungal symbionts(Wang & Qiu 2006).

2.1.1 The Infection Process

Root exudates from AM host plants enhance, but arenot required for spore germination, whereas exu-dates from nonhost plants, e.g., Lupinus (lupin) orBrassica (cabbage) species, do not stimulate germi-nation. Roots of host plants also release a signal orsignals that stimulate(s) the directional growth ofthe AM fungus toward them. CO2 may be onesuch signal (Becard et al. 2004), but there are others,analogous to the signal molecules (flavonoids)

involved in the legume—rhizobium recognitioninteractions (Sect. 3.3; Scervino et al. 2005, Catfordet al. 2006). In fact, flavonoids have been implicatedin the recognition between AM hosts and fungi(Harrison 2005, Hause & Fester 2005). However,root exudates of Zea mays (corn) mutants deficientin chalcone synthase, which is an enzyme involvedin flavonoid synthesis, show similar AM coloniza-tion as those of the wild-type (Becard et al. 1995).This suggests that flavonoids are not the key com-ponents in the AM fungus—host regonition (Bueeet al. 2000, Harrison 2005). In one of the first stagesof AM host recognition, the hyphae of AM fungishow extensive branching in the vicinity of hostroots in response to signaling molecules (Fig. 4).Root exudates contain a branching factor identifiedas a strigolactone, 5-deoxy-strigol (Akiyama et al.2005, Paszkowski 2006). Strigolactones are a groupof sesquiterpene lactones, previously isolated asseed-germination stimulants for the parasiticweeds Striga and Orobanche (Sect. 2.1 of Chapter9D on parasitic associations). Strigolactones induceextensive hyphal branching in germinating sporesof the AM fungus Gigaspora margarita at very lowconcentrations (Bouwmeester et al. 2007). They alsoplay a role in monocotyledonous species [e.g., Sor-ghum bicolor (millet)], interacting with AM fungi atconcentrations as low as 10—13 M. Within 1 hour ofexposure, the density of mitochondria in the fungalcells increases and their shape and movementchanges dramatically which is associated with arapid increase of mitochondrial density and respira-tion (Besserer et al. 2006). Isolation and identifica-tion of plant symbiotic signals open up new waysfor studying the molecular basis of plant—AMfungus interactions. This discovery also provides aclear answer to a long-standing question on theevolutionary origin of the release from host rootsof molecules that stimulate seed germination inparasitic plants (Sect. 2.1 of Chapter 9D on parasiticassociations; Akiyama & Hayashi 2006). Similar sig-naling between host and fungus also plays a role inmycorrhizal associations other than AM, but weknow much less about this (Martin et al. 2001).

During the establishment of AM, fungal hyphaethat grow from spores in the soil or from adjacentplant roots contact the root surface, where they dif-ferentiate to form an appressorium in response tosignals released from the host roots, and initiate theinternal colonization phase (Genre & Bonfante2005). Appressoria form only on epidermal cells,and generally not on the roots of nonhost plants, afurther indication of recognition signals (Harrison2005). Penetration of the root occurs via the appres-soria, and the fungus frequently enters by forcing its


way between two epidermal cells. Alternatively, ahypha may penetrate the cell wall of an epidermalor root-hair cell and grow through the cell as a resultof localized production of hydrolytic enzymes bythe fungus.

Once inside the root, the fungus produces inter-cellular hyphae, coils, and arbuscules. The functionof the arbuscules is most likely to increase the surfacearea of membranes over which exchange of metabo-lites occurs and so to enhance active transportbetween the plasma membrane of the host and thehyphae of the fungus. The invaginated membranesare highly specialized; they contain mycorrhiza-inducible Pi transporters (Rausch et al. 2001, Karan-dashov & Bucher 2005) and H+-pumping ATPases(Ferrol et al. 2002, Requena et al. 2003). Arbusculesare short lived and usually degenerate within a weekor two. Thus, progression of colonization requirescontinuous arbuscule formation as the fungusspreads in the roots (Gadkar et al. 2001). The hyphaeproliferate both in the cortex and in the soil. Vesicles,in which lipids are stored, are sometimes formed at alater stage, either between or within cells. The AMfungus does not penetrate into the endodermis, stele,or meristems; the fungus usually colonizes roots

where the endodermis does not have a completesuberin barrier yet (Brundrett 2002).

The structure of orchid mycorrhizas have alsobeen studied intensively; as with AM, there isextensive intracellular growth with fungi formingintracellular fungal coils, rather than arbuscules(Fig. 2). The fungi forming the mycorrhizas are Basi-diomycota and many belong to the genus Rhizocto-nia. As soon as they have germinated, the orchidseedlings, which have very few reserves, dependon organic matter in the soil or from other hostplants which is supplied via the mycorrhizalfungus. Rhizoctonia species may form associationswith both orchids and conifers. The orchids aretherefore not saprophytic, but mycoheterotrophic(i.e., parasitic on the fungus) (Leake 2004); the asso-ciation between host and fungus does not appear tobe mutually beneficial. Even orchids that have theability to photosynthesize may form ECM with for-est trees, and their stable N- and C-isotope signa-tures indicate a dependence on ECM. This wouldexplain the success of orchids in low-light environ-ments (Bidartondo et al. 2004). In those orchids thatremain nonphotosynthetic during their entire lifecycle [e.g., the Western Australian fully

FIGURE 4. The complete life cycle of arbuscular mycor-rhizal fungi, involving recognition, communication, andestablishment of symbiosis between fungus and host.

The pre-germination stages may be stimulated by theplant root exudates, but may also occur in its absence(after Gadkar et al. 2001).

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subterranean Rhizanthella gardneri (Batty et al.2004)], the fungus continues to play this role. In allorchids, including those that are green (photosyn-thetic) as adults, the fungi also absorb mineral nutri-ents from soil (like AM, see below) (Cameron et al.2006).

In ericoid mycorrhizas, a large number of infectionpoints are found: up to 200 per mm root in Calluna, asopposed to 2—10 per mm in Festuca ovina (sheep’sfescue), infected by an AM fungus. Up to 80% of thevolume of these mycorrhizas may be fungal tissue (notincluding the external mycelium). The fungi infectingEricaceae are Ascomycota (e.g., Hymenoscyphus ericae)(Cairney & Ashford 2002).

Spores of ectomycorrhizal fungi in the rhizospheremay germinate to form a monokaryotic mycelium.This fuses with another hypha, forming a dikaryoticmycelium, which can then colonize the root, forminga mantle of fungal hyphae that enclose the root. Thehyphae usually penetrate intercellularly into the cor-tex, where they form the Hartig net (Fig. 3; Massicotteet al. 1999). The hyphae always remain apoplastic andcan colonize the epidermal (angiosperms) and thecortical (gymnosperms) layers. As hyphae contactthe root surface, roots may respond by increasingtheir diameter and switching from apical growth toprecocious branching (Peterson & Bonfante 1994).Fungal biomass constitutes about 40% of ectomycor-rhizas. Numerous fungal species have the capacity toform ectomycorrhizas. Most of these belong to theBasidiomycota and Ascomycota, and they are oftenspecies that we are familiar with as toadstools. Someof these are edible (e.g., Boletus, truffles), whereasothers are highly toxic (e.g., Amanita).

In contrast to infection by pathogenic fungi, colo-nization with mycorrhizal fungi never causes diseasesymptoms. In the presence of mycorrhizal fungi inthe rhizosphere, flavonoids accumulate in the rootsof Medicago sativa (alfalfa) host roots, similar to, butmuch weaker than, the response to pathogenic fungalattack (Sect. 3 of Chapter 9C on effects of microbialpathogens). AM fungi initiate a transient hostdefense response in the early stages of colonization,followed by suppression to levels well below those ofnoncolonized plants. The production of defense-related gene products is restricted to arbusculatedcells; intercellular hyphae and vesicles elicit no suchdefense response (Harrison & Dixon 1994, Harrison1999, Hause & Fester 2005). The rate and location offungal growth within the root may be controlledthrough activation of plant defense mechanisms.

Some mutants of Pisum sativum (pea) and otherlegumes are characterized by aborted mycorrhizalinfections, after formation of appressoria (Duc et al.1989, Shirtliffe & Vessey 1996). The genes appear to

be linked with genes that control nodulation byrhizobia (Sect. 3.3) which may point to a tight controlof two carbon-consuming and potentially compet-ing symbioses (Sect. 2.6).

2.1.2 Mycorrhizal Responsiveness

In soils with low P availability, plants vary widely inthe extent to which their growth responds to rootcolonization by mycorrhizal fungi (Johnson et al.1997). Mycorrhizal dependency is the ratio of thedry mass of mycorrhizal plants to that of nonmycor-rhizal plants. Species that depend less on AM fungifor their nutrient acquisition are generally colonizedto a lesser extent in the field than more AM-dependentones. This suggests that species that have root systemswith low dependency on mycorrhizas also havemechanisms to suppress mycorrhizal colonization(Sect. 2.3.1). Mycorrhizal dependency of a plantspecies varies with the AM fungal species involvedin the symbiosis (Van der Heijden et al. 1998a). Ingrassland communities, dominant species tend tohave a greater mycorrhizal dependency than subordi-nate species, so that suppression of the AM symbiosisenhances plant species diversity (Hartnett & Wilson2002). In other herbaceous communities, mycorrhizasenhance plant species diversity by increasing theestablishment and abundance of subordinate speciesrelative to the community dominants, and plant diver-sity may be positively correlated with the speciesdiversity of mycorrhizal fungi (Van der Heijden et al.1998b, O’Connor et al. 2002).

Plants that show little responsiveness to AM fungiin terms of growth, may, in fact, acquire significantamounts of P via the fungus (Smith et al. 2003, 2004, Liet al. 2006). Using a compartmented pot system and33P-labeled Pi (Fig. 5A), the contribution of the mycor-rhizal uptake pathway to total plant Pi uptake can beestimated. The hyphal compartment is capped with25 mm nylon mesh, which allows hyphae to penetrate,but excludes roots. Unlabeled P can be absorbeddirectly by roots or via the mycorrhizal pathway.Compared with noninoculated plants without addi-tional P, Linum usitatissimum (flax) grows better, but todifferent extents, depending on the AM fungi tested(Gigaspora rosea, Glomus caledonium, or Glomus intrara-dices). Medicago truncatula (medic) responds positivelyto the two Glomus species in terms of dry weightproduction, but shows a small growth depressionwith Gigaspora rosea, compared with nonmycorrhizalplants. Solanum lycopersicum (tomato) does notrespond positively to any of the fungi. P uptake alsovaries among the different plant—fungus combina-tions, and mycorrhizal phosphorus dependencies


FIGURE 5. (A) Diagrammatic repre-sentation (not to scale) of a com-partmented pot design to assess Puptake by mycorrhizas and roots.The main root + hyphae compart-ment is a nondraining pot contain-ing a mixture of sand and soil. Formycorrhizal treatments, this mixincludes inoculum of three appro-priate fungi: Gigaspora rosea,Glomus caledonium, and Glomusintraradices. Nonmycorrhizal treat-ments receive no inoculum. Thehyphal compartment is a small plas-tic tube containing the same soil +sand mix, but without inoculum; it iscapped with 25 mm nylon mesh,which allows hyphae (shown inred) but not roots (shown in black)to grow into the hyphae compart-ment. The soil in the hyphae com-partment is well mixed with33P-labeled orthophosphate of highspecific activity. (B–D) Mycorrhizaleffects on (A) growth, (B) total Puptake, and (C) specific activities of33P, in Linum usitatissimum (flax,blue bars), Medicago truncatula(medic, yellow bars), and Solanumlycopersicum (tomato, red bars).Mycorrhizal dependencies forgrowth and P-uptake are calculatedas: 100 (value for mycorrhizal plant– mean value for nonmycorrhizalplants)/value for mycorrhizal plant.In the bottom panel, the dotted hor-izontal line indicates the predictedspecific activity of 33P in the plants if100% of P is derived via the mycor-rhizal pathway. This percentage iscalculated using values for specificactivities of 33P in the plants andbicarbonate-extractable P in thehyphae compartment and the totalP available in the pots and in thehyphae compartment. It assumesthat the densities of hyphae (metersper gram of soil) are the same inthe hyphae compartment and inthe hyphae + roots compartment(after Smith et al. 2003). CopyrightAmerican Society of Plant Biologists.

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(Fig. 5B, middle) are similar to mycorrhizal growthdependencies (Fig. 5B, top). By supplying33 P in acompartment to which only the fungal hyphae hasaccess (Fig. 5A), it is possible to show that the mycor-rhizal pathway differs in its contribution to totalP uptake, depending on fungal and plant species. Ptransfer via the mycorrhizal pathway is extremelyhigh in five out of the nine individual plant—funguscombinations, although this is not correlated withmycorrhizal P dependency. With Glomus intraradicesas the fungal partner, all of the P is delivered via themycorrhizal pathway to all tested plants (Fig. 5B,bottom). These findings indicate that mycorrhizasmay be an important pathway of Pi uptake, evenin plants that lack a positive change in growth orP status as a result of AM colonization. It wouldbe interesting to learn more about the conditionsthat cause down-regulation of Pi transportersresponsible for Pi uptake from the root environ-ment (Sect. 2.2.2 of Chapter 6 on mineral nutrition)when AM fungi infect the roots (Karandashov &Bucher 2005).

Mycorrhizal responsiveness is generally assessedusing single species in a pot experiment, whichpoorly reflects the real world. When paired with anear-isogenic nonmycorrhizal genotype, even Sola-num lycopersicum (tomato) shows a positive growthresponse (Cavagnaro et al. 2004), when this is not thecase when tested singly (Fig. 5B, top). This aspect isfurther discussed in Sect. 2.2.

Crop cultivars, e.g., of Zea mays (corn) that havebeen developed for high-input systems in Europehave not lost their ability to be colonized, and maybe more responsive to inoculation by Glomus intrara-dices than those suited for low-input African systems.However, specific adaptations that allow nonmycor-rhizal plants developed for low-input systems to

perform well in low-P soils may limit their ability torespond to higher nutrient supply rates and mycor-rhizal infection. High-input cultivars may have traitsthat are useful for low-input cropping systems wheremycorrhizal symbioses are established (Wright et al.2005). A high mycorrhizal responsiveness is com-monly associated with lack of well-developed roothairs and coarse fibrous roots (Sect. 2.2.1 of Chapter6 on mineral nutrition; Collier et al. 2003).

2.2 Nonmycorrhizal Species and TheirInteractions with Mycorrhizal Species

Although mycorrhizal associations are very com-mon, some species cannot be colonized, or only mar-ginally so (Brundrett 2002). These nonmycorrhizalspecies can be broadly categorized as either ruderalspecies that inhabit relatively fertile sites or speciesthat occur on severely P-impoverished soils. Thenonmycorrhizal ruderals include many that belongto Brassicaceae, Caryophyllaceae, Chenopodiaceae,and Urticaceae. The species from severely P-impoverished habitats include Cyperaceae, Protea-ceae, and Restionaceae, as well as carnivorous(Chapter 9F) and parasitic species (Chapter 9D). The‘‘scavenging’’ strategy of mycorrhizal species, whichaccess P that is in the soil solution, but too far awayfrom roots or inside soil pores that are too small forroots to enter, does not work on severely P-impover-ished soils. The little amount of P that is present inthese soils is predominantly sorbed to soil particles.Cluster roots, which release large amounts of carbox-ylates in an exudative burst (Sect. 2.2.5.2 of Chapter 6on mineral nutrition) effectively ‘‘mine’’ P from thesesoils (Fig. 6). In younger landscapes, nonmycorrhizalspecies with cluster roots tend to occur on either

FIGURE 6. Changes in total soil P and N andin plant nutrient-acquisition strategies(green) as dependent on soil age. ‘‘Poorlydeveloped, very young soils’’ refers to soilsthat result from, e.g., recent volcanic erup-tions or glaciation; ‘‘ancient, highly weath-ered soils’’ refers to soils that have beenabove sea level and not been glaciated formillions of years. Whilst never becomingdominant in severely P-impoverishedsoils, some mycorrhizal species do co-occur with nonmycorrhizal, cluster-bearing species. The width of the trianglesreferring to the different ecological strate-gies provides a measure of the abundanceof these strategies as dependent on soil age(modified after Lambers et al. 2008). Copy-right Elsevier Science, Ltd.


calcareous soils, where the availability of P is low dueto precipitation as calcium phosphates, or on acidssoils, where P precipitates as iron or aluminum phos-phates (Fig. 6.1 in Sect. 2.1 of Chapter 6 on mineralnutrition; Lambers et al. 2006, 2008).

Even within typical nonmycorrhizal genera,mycorrhizal infection has been observed in somespecies (Muthukumar et al. 2004, Boulet & Lambers2005). It is interesting, as discussed in Sect. 2.2.5.2 ofChapter 6 on mineral nutrition, that many of thenonmycorrhizal species from severely P-impoverishedsoils have ‘‘cluster roots’’ (Cyperaceae, Proteaceae, andRestionaceae). Other nonmycorrhizal species includecarnivorous species, e.g., Drosera (sundew), andhemiparasitic species, e.g., Nuytsia floribunda (WesternAustralian Christmas tree).

The mechanisms that prevent colonization innonmycorrhizal species are not yet fully under-stood. In some species, the exudation of fungi-toxiccompounds, such as glucosinolates in Brassicaceae(Koide & Schreiner 1992) or a chitin-binding agglu-tinin in Urtica dioica (stinging nettle) (Vierheiliget al. 1996), may prevent infection. More impor-tantly, the correct chemical cues necessary for devel-opment after spores have germinated (Sect. 2.2.1)may be lacking (Harrison 2005).

Mycorrhizal fungi may enhance growth ofmycorrhizal plants, at least at a low P supply. Insome nonmycorrhizal species, however, the oppo-site is found (Sanders & Koide 1994). In these spe-cies, the mycorrhizal fungus may cause localizedlesions on the roots around aborted penetrationpoints (Allen et al. 1989), inhibit root-hair elonga-tion, probably via its exudates, and reduce stomatalconductance (Allen & Allen 1984). This mechanismmay well explain why nonmycorrhizal speciesshow poor growth in a community dominated bymycorrhizal species, unless the P level is increased(Francis & Read 1994).

2.3 Phosphate Relations

Like root hairs, the external mycelium of mycorrhi-zas increases the roots’ absorptive surface. In fact,the effective root length of the mycorrhizal associa-tions may increase 100-fold or more per unit rootlength (Table 1).

2.3.1 Mechanisms That Account for EnhancedPhosphate Absorption by Mycorrhizal Plants

Arbuscular mycorrhizal associations enhance Puptake and growth most strongly, when the

availability of P in the soil is fairly low (Sect. 2.2.5.2of Chapter 6 on mineral nutrition; Bolan et al. 1987,Thingstrup et al. 1998). Experiments using differentsoil compartments and different labeled sources ofPi show that AM plants do not have access to differ-ent chemical pools of Pi in soil (Fig. 7). They arecapable, however, of acquiring P outside the deple-tion zone that surrounds the root, because of thewidely ramified hyphae. These hyphae allow Ptransport over as much as 10 cm from the rootsurface and at rates that far exceed diffusion in soil(Fig. 8). In addition, they may access smaller soilpores and compete effectively with other microor-ganisms (Joner & Jakobsen 1995); some AM mayaccess organic P, but the ecological significance ofthis remains to be established (Joner et al. 2000a,b,Koide & Kabir 2000). Ectomycorrhizal hyphae canextend even greater distances, possibly severalmeters. Ectomycorrhizal and ericoid mycorrhizalroots also have access to additional chemical poolsof P; they may release phosphatases, which enhancethe availability of organic P, or exude carboxylates,which increase the availability of sparingly solubleP (Landeweert et al. 2001, Van Leerdam et al. 2001,Van Hees et al. 2006). Hymenoscyphus ericae, whichforms mycorrhizas with a number of host plantsbelonging to the Ericaceae, also produces extracel-lular enzymes that may allow the fungus to decom-pose components of plant cell walls which facilitatesaccess to mineral nutrients that are sequestered inthese walls (Leake & Read 1989, Read 1996, Cairney& Burke 1998). Laccaria bicolor, an ectomycorrhizalfungus, is capable of paralyzing, killing, and digest-ing springtails. A significant portion of the organicN that is subsequently hydrolyzed ends up in itsmacrosymbiont, Pinus strobus (eastern white pine)(Klironomos & Hart 2001). Springtails selectivelyfeed on fungi, including mycorrhizal fungi. Shouldthis phenomenon prove to be widespread, forest—nutrient cycling may turn out to be more complicatedand tightly linked than is currently recognized. Inter-estingly, springtails prefer nonmycorrhizal specieswhen given a choice; their fecundity is reducedwhen fed a diet of less palatable mycorrhizal fungi(Klironomos et al. 1999).

Ectomycorrhizas and ericoid mycorrhizas fre-quently occur in organic soils, whereas AMs aremore typical of mineral soils. Weatherable mineralsunder many European coniferous forests contain adense network of tubular micropores with a dia-meter of 3—10 mm that are formed by carboxylatesexuded by mycorrhizal or saprotrophic fungi. Con-centrations of succinate, oxalate, formate, citrate,and malate may be in the micromolar to millimolarrange. These exuded carboxylates enhance mineral

Mycorrhizas 413

FIGURE 8. Schematic presentation of components of thenutrient acquisition from the soil by arbuscular mycor-rhizal roots (right). The thickness of the circle indicatesthe importance of AM in acquiring this nutrient (? indi-cates lack of definitive information). Additional compo-nents in ectomycorrhizal roots are also shown (left).Note that all mycorrhizas enhance the availability of

soil nutrients by enlarging the soil volume that isexploited, and that this is most relevant for those nutri-ents that are least mobile (e.g., phosphate). Ectomycor-rhizas excrete hydrolytic enzymes, which allow them touse organic forms of both P and N, and chelating organicacids, which allows the use of poorly soluble forms ofphosphate (after Marschner & Dell 1994).

FIGURE 7. Diagram showing thedesign of the rhizoboxes used toassess which chemical forms ofP can be accessed by arbuscularmycorrhizal hyphae. The 32P-labeled P source is added tothe hyphal compartment only.The 32P content is expressed asa proportion of the total P con-tent of the shoots of Trifoliumpratense (red clover) (Yao et al.2001).


weathering by forming complexes with Al and dis-solve Ca-rich rock (Wallander 2000). In this wayabout 107 hyphal tips are eating their way throughsand grains at any one time, forming 150 km ofmicropores per m3 of soil per year. Ectomycorrhizalhyphae of the species Suillus granulatus and Pilodermacroceum are thought to transport the released miner-als directly to their hosts, thereby bypassing competi-tion for nutrient uptake by other organisms. Thismechanism by which mycorrhizal hyphae bypassthe soil to reach minerals might help explain whyforest productivity has not decreased, despite recentexcessive soil acidification (Jongmans et al. 1997).

The external AM mycelium consists of both large‘‘runner’’ hyphae and finer hyphae with a role innutrient absorption. As in roots, Pi uptake by thefine mycorrhizal hyphae occurs via active transport,against an electrochemical potential gradient, with aproton-cotransport mechanism (Sect. 2.2.2 of Chap-ter 6 on mineral nutrition). Once absorbed by theexternal hyphae, P is rapidly transported intovacuoles where most of it is polymerized into inor-ganic polyphosphate (poly-P), a linear polymer ofthree to thousands of inorganic phosphate mole-cules, connected by high-energy phosphate bonds(Ezawa et al. 2002). This reaction is catalyzed bypolyphosphate kinase, which is induced whenexcess Pi is absorbed. Poly-P accumulated in thevacuole is translocated from the external hyphae tothe hyphae inside the roots, possibly via cytoplas-mic streaming. This transport of P and that of otherimmobile ions through the external hyphae to theplant is relatively rapid, bypassing the very slowdiffusion of these ions in soil. Once the poly-P hasarrived near the plant cells, it is degraded, presum-ably catalyzed by phosphatases. The mechanismsaccounting for poly-P production at one end of thehyphae and breakdown at the other are unknown. Itmight involve sensing a gradient in plant-derivedcarbohydrates. Transfer of P and other nutrientsfrom fungus to plant is a two-step process over themembranes of the two symbionts, probably invol-ving passive efflux from the fungus and activeuptake by the plant (Javot et al. 2007). Some of theplant Pi-transporter genes involved in this processare mycorrhiza-specific and differ genetically fromthose discussed in Sect. 2.2.2 of Chapter 6 on mineralnutrition (Karandashov & Bucher 2005). Thesemycorrhiza-inducible Pi-transporter genes are up-regulated in roots that are colonized by AM fungiand expressed at a very low level in noncolonizedparts of the same root system. These findings showthat in species that form AM associations with mem-bers of the Glomeromycota Pi transporters haveevolved that are involved in scavenging P from theapoplast between intracellular AM structures and

root cortical cells (Rausch et al. 2001, Glassop et al.2005).

2.3.2 Suppression of Colonization at HighPhosphate Availability

The AM fungus colonizes roots to a greater extent inlow-P soils than in soils that are more fertile (Fig. 9;Smith & Read 2008). To some extent this greater

FIGURE 9. Response of Casuarina cunninghamiana(sheoak) seedlings to inoculation with an arbuscularmycorrhizal fungus (Glomus sp.) over a range of P sup-plies in sand culture. (A) Shoot dry weight; (B) mycor-rhizal colonization of roots; (C) occurrence of clusterroots. Filled symbols: inoculated with Glomus; opensymbols uninoculated (redrawn after Reddell et al.1997, Australian Journal of Botany 45: 41–51, CopyrightCSIRO, Australia).

Mycorrhizas 415

frequency may be associated with a decreased rateof root elongation so that the colonization by thefungus keeps up with the growth of the root.Systemic effects of P are also important, however.An analysis of Solanum tuberosum (potato) grownwith a divided root system of which only one half isinoculated with the AM fungus Glomus intraradicesshows that when high P levels are applied to thenoncolonized part of the root system, the formationof both arbuscules and vesicles is suppressed in thecolonized portion of the root, despite the presence ofmore internal hyphae. Moreover, a high Pi supplymay lead to down-regulation of the expression levelof both the mycorrhiza-inducible and other Pi trans-porters (Rausch et al. 2001, Glassop et al. 2005).

Once a root is infected by an AM fungus, furtherinfection of the roots tends to be suppressed(Vierheilig et al. 2000); this phenomenon is calledautoregulation. A decreased availability of carbo-hydrates or an improved P status of the plant hasbeen offered as an explanation for this suppressionof further colonization. To test this, plants ofHordeum vulgare (barley) were grown with a dividedroot system, where one half is inoculated with anAM fungus. After extensive root colonization, theother half is inoculated, but colonization is sup-pressed (Fig. 10). In such plants some of the Pacquired by the colonized root part ends up in theroots that are not colonized, as a result of transportvia the xylem to the shoot, followed by export viathe phloem from the shoot. Because the biomass andthe P concentration of both root halves are the same,neither a shortage of carbohydrates nor elevated Pconcentrations explain the suppression (Fig. 10). Asystemic suppression, i.e., a response triggered bymycorrhizal colonization followed by signaling tothe rest of the plant, is the most likely explanationfor the autoregulatory effect of prior mycorrhizationon subsequent colonization. This involves signalingmolecules such as strigolactones (Sect. 2.1.1), whoserelease from roots of plants that are hosts for AMfungi is promoted by P deficiency (Yoneyama et al.2007a). By contrast, in roots of nonmycorrhizalspecies, P deficiency does not affect exudation(Yoneyama et al. 2007b). Interestingly, the systemicautoregulatory effect not only suppresses mycorrhi-zation in Hordeum vulgare (barley), but also reducesroot infection by the take-all disease caused by thefungus Gaeumannomyces graminis var. tritici. Thiseffect is found when barley plants show a highdegree of mycorrhizal root colonization, whereas alow mycorrhizal root colonization has no effect ontake-all (Khaosaad et al. 2007).

2.4 Effects on Nitrogen Nutrition

Unlike AM, some ectomycorrhizas may utilizeorganic N, including proteins. This has been docu-mented in situ, as revealed by a comparison of 15Nfractionation in plants with and without ectomycor-rhiza. Ectomycorrhizal plants may have a 1.0—2.5‰

more positive �15N value than do plants infectedwith arbuscular mycorrhiza (Table 2), demonstrat-ing that different N sources are used (either differentcompounds or different regions in the soil). A studyon a Tanzanian woodland (Table 2) provides noevidence for a difference in isotope composition indifferent soil layers. Fractionation of the heavy Nisotope (15N) occurs during mineralization andnitrification; therefore, the organic N becomesenriched with 15N. The data in Table 2 provideevidence that the ectomycorrhizal plants use a sig-nificant amount of N from a pool that was notdecomposed and nitrified (i.e., organic N). In borealforest and arctic tundra, ectomycorrhizal plant spe-cies also have distinctive 15N signatures, with 15Nconcentrations that are higher than those of specieswith ericoid mycorrhizas, but lower than those ofAM or nonmycorrhizal species (Schulze et al. 1995,Nadelhoffer et al. 1996). In these studies either root-ing depth or the form of N that is utilized could havecontributed to the different 15N signatures.

Ericoid mycorrhizas, like ectomycorrhizas, canuse quite complex organic sources of N and P(Fig. 8; Read 1996). This ability may contribute tothe dominance by Ericaceae of many cold and wetsoils, where rates of decomposition and mineraliza-tion are low (Read & Perez-Moreno 2003). AMs areat the other extreme of the continuum of mycorrhi-zal associations. They cannot access organic P,because of a very low capacity to release phospha-tases into the soil (Joner et al. 2000a). Their predo-minant significance lies in the acquisition ofsparingly available inorganic nutrients, especiallyP. AMs are relatively unimportant for acquisitionof N, if this is available as NO3

—, but they do enhanceN acquisition when mineral N is present as the lessmobile NH4

+ (Johansen et al. 1994, Tanaka & Yano2005). In the extraradical hyphae, ammonium isassimilated into arginine, and then transportedtoward the arbuscules, where arginine is brokendown, followed by transfer of NH4

+ to the hostcells (Govindarajulu et al. 2005, Jin et al. 2005, Chalotet al. 2006). AM may enhance the uptake of NO3

—

from dry soils, when mass flow and diffusion arelimited, but not in wet soils (Tobar et al. 1994). Ecto-mycorrhizas are thought to be intermediate between


the ericoid and arbuscular mycorrhiza in terms ofaccessing organic N (Lambers et al. 2008).

2.5 Effects on the Acquisition of Water

Arbuscular mycorrhizal plants may have anenhanced capacity to acquire water from the rootenvironment (Auge 2001). Several hypotheses havebeen put forward to explain this increased capacity.One suggests an indirect effect via the improved Pstatus of the plant which increases the hydraulic

conductance of the roots or affects the plant’s hor-mone metabolism and stomatal conductance. Lactucasativa (lettuce) plants colonized by the AM fungiGlomus coronatum, Glomus intraradices, Glomus claroi-deum, and Glomus mosseae deplete soil water to agreater extent than uninoculated control plants orplants colonized by Glomus constrictum or Glomusgeosporum. The differences in soil-moisture depletioncan be ascribed to the activity of AM fungi, but fungidiffer in their effectiveness to enhance plant wateruptake from soil, probably related to the amount ofexternal mycelium produced by each AM fungus

FIGURE 10. Diagram showing theexperimental design of the rhizo-boxes used to assess the systemiceffect of prior infection of Hor-deum vulgare (barley) by arbuscu-lar mycorrhizal fungi(compartment B) on subsequentinfection (in compartment C).Infection takes place from roots ofPhaseolus vulgaris (common bean)in adjacent compartments (A or D)(modified after Vierheilig et al.2000).

Mycorrhizas 417

and to the frequency of root colonization in terms oflive and active fungal structures (Marulanda et al.2003). The multihyphal strands of ectomycorrhizasare thought to have a particularly high capacity totransport water. Alternatively, the improved plantwater status may be an effect on effects of mycorrhi-zas on soil structure (Bearden & Petersen 2000).

2.6 Carbon Costs of the MycorrhizalSymbiosis

Mycorrhizas provide nutritional benefits, which mayenhance resource allocation to leaves (Baas &Lambers 1988, Grimoldi et al. 2005) and photosyn-thetic carbon gain of the host (Douds et al. 1988,Wright et al. 1998a), but they also incur carbon costsfor the host. The amount of carbon that is exudedfrom intact roots into the apoplast and normally endsup in the rhizosphere is not sufficient to satisfy thedemand of the microsymbiont of the mycorrhizalassociation. It is possible that passive efflux of carbonis increased in mycorrhizal plants; alternatively, thehost’s active carbon-uptake system may be inhibited.So far there is no molecular information on transportproteins that account for carbon efflux from or carbonre-uptake in mycorrhizal plants (Bago et al. 2000).

Costs associated with the AM symbiosis havebeen estimated in various ways (e.g., by comparingplants with and without the mycorrhizal symbiontat the same growth rate). This can be achieved byproviding more P to the nonmycorrhizal plant, com-pared with the supply to the mycorrhizal plant. Thecarbon use for growth and respiration by the roots of

both types of plants can then be used to quantifycosts of the mycorrhizal symbiosis (Snellgrove et al.1982, Grimoldi et al. 2006). The problem with thismethod is that it assumes steady-state rates of Pacquisition and carbon consumption, whereas infact these may vary following active root coloniza-tion. A variation of this approach is to grow nonmy-corrhizal plants at a range of P supplies, so that aP—response curve can be constructed with which tocompare the mycorrhizal plants (Rousseau & Reid1991, Eissenstat et al. 1993).

An alternative approach to quantify the costs ofthe mycorrhizal symbiosis has been to grow plantswith a divided root system. That is, part of the rootsystem is grown in one pot, and the remaining part ina separate pot. One part of the divided root is theninoculated with a mycorrhizal fungus, while theother is not and remains nonmycorrhizal. The shootis then given 14CO2 to assimilate in photosynthesisand the partitioning of the label over the two rootparts is measured (Table 3; Koch & Johnson 1984,Douds et al. 1988). It is also possible to calculatecarbon costs of the mycorrhizal symbiosis by measur-ing the flow of 14C-labeled assimilates into soil andexternal hyphae (Jakobsen & Rosendahl 1990).

The estimates of the carbon costs of the AM sym-biosis vary between 4 and 20% of the carbon fixed inphotosynthesis (Lambers et al. 2002). Only a minorpart (15%) of the increased rate of root respiration isassociated with an increased rate of ion uptake by themycorrhizal roots. The major part (83%) is explainedby the respiratory metabolism of the fungus and/orother effects of the fungus on the roots’ metabolism(Baas et al. 1989). Construction costs of fibrous roots

TABLE 2. 15N abundance of leaf samples collected in different years in Tanzania.*

d15N

Species Symbiotic status 1980 1981 1984

Brachystegia boehmii EC 1.64 1.32 1.23B. microphylla EC 1.53 1.51 1.73Julbernardia globiflora EC 2.81 1.63 1.60Pterocarpus angolensis AMþNO �0.81 �0.87 �0.93Diplorynchus condylocarpon AM � �0.36 �0.60Xeroderris stuhlmannii AMþNO � 0.01 0.62Dichrostachys cinerea AMþNO � 0.45 �0.38

Source: Hogberg (1990).* EC ¼ ectomycorrhizal; AM ¼ arbuscular mycorrhizal; NO ¼ nodulated. The experimentssummarized here were actually carried out with the aim to determine the extent ofsymbiotic N2 fixation of the nodulated plants. Since nodulated plants have access todinitrogen from the atmosphere, they are expected to have d15N‰ values closer to atmo-spheric N2 than do plants that do not fix dinitrogen. The data presented here stress thatcontrol plants need to be sampled to allow a proper comparison. This table shows that thechoice of the control plants is highly critical (see also Sect. 3 of this chapter).


are also higher for mycorrhizal than for nonmycor-rhizal roots, because of their higher fatty acid con-centration (Sect. 5.2.1 of Chapter 2B on plantrespiration; Peng et al. 1993). The costs for the ECMsymbiosis are probably higher, but there is little infor-mation available (Hobbie 2006).

In addition to a higher carbon expenditure, mycor-rhizal plants also tend to have a higher rate of photo-synthesis per plant, partly due to higher rates ofphotosynthesis per unit leaf area and partly due totheir greater leaf area (Wright et al. 1998a, b). Thehigher rate of CO2 assimilation is most pronouncedwhen the soil water potential is low (Sanchez-Diazet al. 1990). When P and water are limiting for growth,therefore, benefits outweigh the costs, and mycorrhizalplants usually grow faster, despite the large carbonsink of the symbiotic system. The relatively high costsof the mycorrhizal association, however, may help toexplain why mycorrhizal plants sometimes grow lessthan their nonmycorrhizal counterparts (Thompsonet al. 1986, Fredeen & Terry 1988), especially when asecond microsymbiont (rhizobium) plays a role(Fig. 11). Under drought, however, mycorrhizal plantsmay still show more benefit from an association withrhizobium than do nonmycorrhizal control plants(Penas et al. 1988).

2.7 Agricultural and EcologicalPerspectives

From an ecological point of view, information on themycorrhizal status of plants in a community is mostimportant. In a mixed community nonmycorrhizalspecies may profit most from fertilization with Pbecause the mycorrhizal association is often sup-pressed at a higher P supply, and not necessarilybecause the growth of nonmycorrhizal species ismore severely P-limited (Sect. 2.3). Suppression ofthe mycorrhizas might then reduce the harmful

effect of the mycorrhizal fungus on nonmycorrhizalspecies (Sect. 2.2). We should therefore be cautiousin interpreting the effects of P fertilization on thegrowth of certain plants in a community.

Close proximity between the roots of a seedlingand those of an established, infected plant may speedup AM infection, but, for unknown reasons, this isnot always the case (Newman et al. 1992). AMs mayhave profound effects on interactions between plantsin a community, as discussed in Sect. 7 of Chapter 9Eon interactions among plants.

Mycorrhizas can obviously never enhancegrowth and productivity of crop plants in theabsence of any P. Mycorrhizal associations,

TABLE 3. Comparison of accumulated 14C and fresh mass in mycorrhizal andnonmycorrhizal halves of root system of two citrus cultivars.*

14C recovered from below-ground tissue dpm g–1 Fresh mass mg plant–1

Species þ � þ �

Sour orange 66.4 33.6 1580 1240NSCarrizo citrange 67.7 32.3 1990 1520NS

Source: Koch & Johnson (1984).*þ and � denote mycorrhizal and nonmycorrhizal plants, respectively; NS indicates thatthere was no significant difference.

FIGURE 11. The relative host response to mycorrhizalinfection as dependent on the supply of hydroxyapatite[Ca10(PO4)6(OH)2]. Phaseolus vulgaris (common bean)plants are infected with Rhizobium phaseoli, a nitrogen-fixing bacteria. Half of the plants are also infected withthe mycorrhizal fungus Glomus fasciculatum. The dif-ference in mass of the parts of the mycorrhizal plantsrelative to the nonmycorrhizal control plants is calcu-lated as percentage of the difference. Negative valuesindicate that the shoot or nodule mass is less in themycorrhizal plants (after Bethlenfalvay et al. 1982).Copyright American Society of Plant Biologists.

Mycorrhizas 419

however, do have great potential in improving cropproduction when P or other immobile nutrients arein short supply. Introduction of spores of the bestmicrosymbiont and breeding for genotypes with a

more efficient mycorrhizal symbiosis are tools thatcan be used to enhance food production in countrieswhere immobile nutrients restrict crop production.As such, mycorrhizas allow good crop growth and

FIGURE 12. (Top) Planting arrangement ina split-root design. (Bottom) N content ofCasuarina cunninghamiana and Eucalyp-tus maculata seedlings (blue bars fordonors and green bars for receivers) asaffected by mycorrhizas, nodulation, Nsource, and identity of the N-donor orN-receiver. Plants were fertilized with14N for 5 months and with 15N (Ndonor only) for 1 month before harvest.Abbreviations: C, Casuarina cunning-hamiana; E, Eucalyptus maculata; M,mycorrhizal status; F, nodulation status(He et al. 2004). Copyright Trustees ofThe New Phytologist.


may reduce nutrient losses to the surroundingenvironment (Tisdall 1994).

Arbuscular and ectomycorrhizal mycorrhizalfungi may infect many plants at the same time,even plants of different species, potentially providinga pathway for transport of carbon or nutrientsbetween roots of different plants. Two-way N trans-fers mediated by a common mycorrhizal networkhave been examined by growing plants in two-cham-bered pots separated by nylon mesh, and supplying15NH4

+ or 15NO3— ; the mesh effectively excludes root

contact, but allows movement of water-soluble sub-stances and penetration of fungal hyphae (Johansen& Jensen 1996). Using this method, it can be demon-strated that nitrogenous compounds are transportedbidirectionally between N2-fixing Casuarina cunning-hamiana trees (sheoak) and Eucalyptus maculata trees(spotted gum), connected via the ectomycorrhizalfungus Pisolithus sp. (Fig. 12), especially in the nodu-lated, ectomycorrhizal treatment. About twice asmuch N moves from Eucalyptus maculata towardCasuarina cunninghamiana as in the opposite direc-tion, irrespective of the source of N (15NH4

+ or15NO3

—), resulting in increased growth of Casuarinacunninghamiana due to interspecific N transfer. Sincethere is virtually no N transfer in the nonmycorrhizaltreatment, but significant N transfer in the mycorrhi-zal treatment, N transfer between the two tree speciesis obviously mediated by ectomycorrhizal fungi. Themuch higher N transfer between nodulated mycor-rhizal plants indicates that mycorrhizas and Frankiatogether enhance bidirectional N fluxes between N2-fixing Casuarina cunninghamiana and non-N2-fixingmycorrhizal Eucalyptus maculata, contradicting theview that N flows from N2-fixing to non-N2-fixingplants. However, the fraction of N derived fromtransfer is similar for both species, because the Nconcentrations are higher in the N2-fixing Casuarinacunninghamiana (He et al. 2004, 2005). It remains to beestablished how significant bidirectional transport ofnutrients or carbon is in the field, especially via AMnetworks (Pfeffer et al. 2004). Using stable carbonisotopes to assess carbon transfer via AM, providesno evidence for carbon transfer from Festuca idahoen-sis (Idaho fescue) to the exotic invasive Centaureamaculosa (spotted knapweed). However, Centaureamaculosa exploits its mycorrhizal symbiosis moreeffectively than the native grassland species, prob-ably due to the luxury consumption of P throughefficient utilization of extra-radical hyphae. Exploita-tion of AM networks may be a mechanism bywhich invasive weeds outcompete their neighbors(Zabinskey et al. 2002). Transport of carbon viamycorrhizal hyphae is obviously also important inmycoheterotrophic plants, which depends on

carbon supplied by a photosynthetic plant towhich they are connected via mycorrhizal hyphae(Sect. 2.1.1; Selosse et al. 2006). However, putativelyautotrophic orchids also receive significant amountsof carbon from their fungal associates (Gebauer &Meyer 2003).

Benefits of N nutrition to non-N2-fixing plants byneighboring N2-fixing plants are well documented(Sect. 3.6). Differences in net N transfer with differentnodulation/mycorrhizal combinations could haveimportant ecological implications, both for nutrientcycling and for the structure and function of naturalor agricultural plant communities, particularly withrespect to those plants that can potentially construct acommon mycorrhizal network to transfer nutrients(He et al. 2005). AM colonization can result inresource equalization or sharing, thus reducingdominance of aggressive species and promoting coex-istence and biodiversity (Van der Heijden et al. 1998a).In tallgrass prairie, Fischer Walter et al. (1996) docu-mented transfer of (labeled) P over distances of up to0.5 m. Transfer of label, however, does not imply a nettransfer of P. Although tracer experiments do showthat interplant transfer does occur, the amount oftransfer of 32P between mycorrhizal plants of Loliumperenne and Plantago lanceolata appears to be much tooslow to be ecologically significant (Eissenstat 1990).

3. Associations with Nitrogen-Fixing Organisms

Nitrogen is a major limiting nutrient for the growth ofmany plants in many environments in young land-scapes (Sect. 2.1 of Chapter 6 on mineral nutrition).Terrestrial N is subject to rapid turnover, and is even-tually lost as nitrogen gas into the atmosphere ordeposited in marine sediments. Its maintenance,therefore, requires a continuous reduction of atmo-spheric N2. Biological reduction of dinitrogen gas toammonia can be performed only by some prokar-yotes and is a highly O2-sensitive process. The mostefficient N2-fixing microorganisms establish asymbiosis with higher plants, in which the energyfor N2 fixation and the O2-protection system are pro-vided by the plant partner (Mylona et al. 1995).

Symbiotic associations with microorganisms thatfix atmospheric N2 may be of major importance for asymbiotic plant’s N acquisition, especially in envir-onments where N is severely limiting to plantgrowth. As such, the symbiosis is also of agronomicand environmental importance, because it reducesthe need for costly fertilizers and greenhousegas emissions associated with their production.

Associations with Nitrogen-Fixing Organisms 421

A nonsymbiotic association [e.g., with Azospirillum inthe rhizosphere of tropical grasses or Gluconacetobac-ter diazotrophicus in the apoplast of the stems of Sac-charum officinarum (sugarcane)] is sometimes found.Contrary to the strictly symbiotic systems, no specialmorphological structures are induced.

Symbiotic N2-fixing systems require a carboninput from the host that is far greater than the car-bon requirements for the acquisition of N in thecombined form (e.g., NO3

—, NH4+, or amino acids).

Are there mechanisms to suppress the symbiosiswhen there is plenty of combined N around? Howdoes a plant discriminate between a symbiotic guest

and a pathogenic microorganism? Finally, what isthe significance of nonsymbiotic N2 fixation forplants? To answer these ecological questions wewill first provide a basic understanding of somephysiological aspects of this symbiotic association.

3.1 Symbiotic N2Fixation Is Restrictedto a Fairly Limited Number of Plant Species

Because of its overwhelming economic importance,the most widely studied associations between N2-fixing microorganisms and vascular plants are those

FIGURE 13. N2-fixing symbio-tic systems. (Top, left)Legume–rhizobium symbiosison the South African Chamae-crista mimosoides (fishbonedwarf cassia) (photo H. Lam-bers). (Bottom, left) Symbio-tic structure (rhizothamnia)between the Western Austra-lian Allocasuarina humilisand an Actinobacteria (Fran-kia) (courtesy M.W. Shane,School of Plant Biology, TheUniversity of Western Austra-lia, Australia). (Right) Sym-biotic structure betweenMacrozamia riedlii and cya-nobacteria (courtesy M.W.Shane, School of Plant Biol-ogy, The University of Wes-tern Australia, Crawley,Australia).


that involve a symbiosis between bacteria of thefamily Rhizobiaceae (genera: Allorhizobium, Azorhi-zobium, Bradyrhizobium, Rhizobium, Mesorhizobium,and Sinorhizobium, collectively known as rhizobia;the numbers of genera are increasing steadily, andthey include both alpha and beta Proteobacteria;Sprent & James 2007) and more than 3000 speciesof Fabaceae (Geurts & Bisseling 2002, Vessey et al.2005). Parasponia is the only nonlegume speciesknown to have a symbiotic association with rhizo-bium [Bradyrhizobium and Rhizobium (Vessey et al.2005)]. Invariably, root nodules are formed (Figs. 13and 14), with the exception of Azorhizobium andBradyrhizobium, which induces nodules on bothstems and roots (of Sesbania rostrata and Aeschyno-mene species, respectively). The Fabaceae familycomprises three subfamilies; the less specializedsubfamily Caesalpinioideae includes far morenonnodulating species than do the other twosubfamilies (Van Rhijn & Vanderleyden 1995). Thesymbiosis between rhizobia and legume crops is ofenormous agronomic importance, especially wherefertilizer inputs are low.

There are also nonlegume species capable offorming a symbiotic association with N2-fixingorganisms, other than rhizobia. First, there is the

actinorhizal symbiosis between soil bacteria(Frankia, Actinobacteria) and more than 200 speciesfrom eight nonlegume families of angiosperms [e.g.,Alnus (alder), Hippophae (sea buckthorn), Myrica(myrtle), Elaeagnus (silverberry), Dryas (mountainavens), and Casuarina (sheoak)]. In all these sym-bioses root nodules are formed (Figs. 13 and 14;Akkermans & Hirsch 1997, Vessey et al. 2005). Sec-ond, there are symbioses between Cyanobacteria(Nostoc, Anabaena) and plant species of the generaAzolla, Macrozamia, and Gunnera (Vessey et al. 2005).Special morphological structures are sometimesformed on the roots (e.g., the coralloid roots inMacrozamia species) (Figs. 13 and 9.A.14; Pate et al.1988). The endosymbiont only fixes N2, not CO2,although cyanobacteria are photosynthetically activewhen free-living (Lindblad et al. 1991). Nostoc in theGunnera—Nostoc symbiosis maintains its photosystemII (PS II) units, but their photochemical efficiency ismuch reduced (Black & Osborne 2004). In the sym-biosis between fungi of the genus Collema and cyano-bacteria (Nostoc), the cyanobacteria arephotosynthetically active; this symbiosis occurs inlichens.

Table 4 gives an overview of major symbioticassociations between plants and microorganisms

FIGURE 14. Diagrammatic representation of longitu-dinal sections through (A) an indeterminate legumenodule, (B) a determinate legume nodule, (C) anactinorhizal nodule, and (D) a lobe of a symbioticcoralloid root cluster. The red colored regions repre-sent the infected zones. The dark, thick lines repre-sent vascular tissues. Outer cortical (OC) tissue,inner cortical (IC) tissue, and meristems (m, blue)are indicated. In the indeterminate legume nodule(B) and the actinorhizal nodule (C), the zones ofinfection (1), N2 fixation (2), and senescence (3) areindicated (Vessey et al. 2005).


capable of fixing atmospheric N2. It shows thatN2-fixing organisms can be very significant for theinput of N into natural and agricultural systems.

3.2 Host–Guest Specificity in theLegume–Rhizobium Symbiosis

The associations between legumes and rhizobiahave been studied most elaborately. Many arehighly specific. For example, Rhizobium meliloti willinfect Medicago truncatula (medic), Melilotus alba(honey clover), and Trigonella coerulea (fenugreek),but not Trifolium repens (white clover) or Glycine max(soybean). Bradyrhizobium japonicum will nodulateGlycine max (soybean), but not Pisum sativum (pea)and Medicago truncatula (medic). Other rhizobia(e.g., Rhizobium strain NGR234) may infect up to100 host species, from different genera, includingParasponia andersonii, a nonlegume, but this is excep-tional. What determines the specificity and whydoes this specificity vary among different rhizobia?To answer these questions, we first discuss theinfection process in more detail.

3.3 The Infection Process in theLegume–Rhizobium Association

Nodule formation in legumes such as Pisum sativum(pea) and Glycine max (soybean) is preceded by the

release of specific phenolic compounds (flavo-noids: flavones, flavanones, or isoflavones) andbetaines from the legume roots (Phillips et al.1994). The same or similar flavonoids are inducedas antibiotics (phytoalexins) upon infection bypathogenic microorganisms (Sect. 3 of Chapter 9Con effects of microbial pathogens). Subtle differ-ences between host plant species, of which we areonly just beginning to understand the details, deter-mine if an interaction between a bacterium and aplant results in symbiosis or pathogenesis. The fla-vonoids bind with a bacterial gene product, andthen interact with a specific promoter in the genomeof rhizobium. This promoter is associated with thegenes responsible for inducing nodulation (thenodulation, or nod genes). As detailed in the follow-ing sections, the products of these genes, the Nodfactors, induce root-hair curling on the plant andcortical cell divisions, which are among the earliest,microscopically observable events in the nodulationof most legume species. Nod factors are not requiredfor symbiosis in some legumes, e.g., Aeschynomenespecies forming a symbiosis with photosyntheticBradyrhizobium species (Giraud et al. 2007). Rhizobiamay also enter through cracks in the epidermis,associated with lateral-root formation, or wounds;for 25% of all legumes this is the only way of entry(Sprent 2007).

The actinorhizal symbiosis between plantspecies like Alnus glutinosa (black alder) and the Acti-nobacteria Frankia also involves the release of specificcompounds (flavonols) that enhance the level of

TABLE 4. Symbiotic associations between plants and microorganisms capable of fixing atmospheric N2.*

Plant type Genus Microorganism LocationAmount of N2 fixed (kg N ha–1

season–1)

Fabaceae Pisum Rhizobium Root nodules 10–350Glycine Bradyrhizobium Root nodules 15–250Medicago Sinorhizobium Root nodules 440–790Sesbania Azorhizobium Stem nodules 7–324

MesorhizobiumUlmaceae Parasponia Bradyrhizobium Root nodules 20–70Betulaceae Alnus Frankia Root nodules 15–300Casuarinaceae Casuarina (Actinobacteria) Root nodules 9–440Eleagnaceae Eleagnus (Actinobacteria) Root nodules ndRosaceae Rubus (Actinobacteria) Root nodules ndPteridophytes Azolla Anabaena Heterocysts in cavities of dorsal

leaf lobes40–120

Cycads Ceratozamia Nostoc Coralloid roots 19–60Lichens Collema Nostoc Interspersed between fungal

hyphaend

Source: Kwon & Beevers (1992), Gault et al. (1995), Peoples et al. (1995), Vance (2002).* Only a limited number of species are listed, just to provide an example; nd is not determined.


nodulation, but their exact role in the process isunknown (Van Ghelue et al. 1997, Vessey et al. 2005).Very little is known about the chemical nature ofattractants from hosts to Cyanobacteria (Vessey et al.2005).

3.3.1 The Role of Flavonoids

To some extent specificity between the host andrhizobium is determined by the type of flavonoidsreleased by the host and by the sensitivity of therhizobium promoter for a given type of flavonoid(Table 5). Rhizobium species that form symbioseswith a broad range of plant species respond to awider range of flavonoids than those that are morespecific, but, if the flavonoid concentration isincreased, a response may occur, even in thosemore specific rhizobia. In addition, nonlegumesmay also exude flavonoids, and several legumesexude flavonoids that also activate the promoter ofrhizobium species that are unable to establish asymbiosis. Other factors must, therefore, also con-tribute to specificity (Spaink 1995). What ultimateeffects do the flavonoids have in rhizobium?

To study the effect of flavonoids on rhizobium anappropriate assay is required that is less elaboratethan measuring root-hair curling (Maxwell et al.1989). A construct has been made, coupling thegene from Escherichia coli that encodes b-galactosi-dase to the promoter of the nod genes. The activity ofthe enzyme b-galactosidase is then measured in asimple spectrophotometric assay. In this way, therelative effect of various flavonoids can be assessedby determining the activity of b-galactosidase,

rather than the extent of root-hair curling. In thefollowing section we examine the kind of productsproduced by the bacterial nod genes.

3.3.2 Rhizobial nod Genes

There are three types of rhizobial nod genes (Fig.15). First, all rhizobia have a nod gene that is tran-scribed constitutively and probably confers somehost specificity. The product of this gene bindswith flavonoids produced by the host plant to acti-vate the common nod genes, which are found in allrhizobium species and lead to the production of abacterial lipochitooligosaccharide. The flavonoidsalso activate the transcription of host-specific nodgenes, which encode enzymes that ‘‘decorate’’ thislipochitooligosaccharide. The ‘‘decorated’’ lipochi-tooligosaccharides are known as the Nod factors.The lipid component of the Nod factor allowspenetration through membranes. Different sidegroups are added to the backbone of this molecule,and this confers the specificity of a certain rhizo-bium (Fig. 15). Rhizobial species with a broad spe-cificity produce many different Nod factors, asopposed to ones with a narrow host range. Thatis, the structure of the lipochitooligosaccharidedetermines if it will be recognized as a symbiontor as a pathogen by a potential host plant. Becausethe Nod factor is effective at concentrations as lowas 10—12 M, a plant receptor must be involved, andevidence for this has recently been obtained(Radutoiu et al. 2003, Oldroyd et al. 2005). Rhizo-bial Nod factors trigger nod gene transcription inplant epidermal cells within 1 hour of exposure;

TABLE 5. Comparisons of indeterminate and determinate nodules.

Parameter Indeterminate Determinate

Nodule initiation Inner cortex Outer cortexCell infection through Infection threads Infection threads and cell divisionMeristem Persistent (months) Nonpersistent (days)Bacteroid size Larger than bacteria Variable, although usually not too much larger than

bacteriaPeribacteroid membrane One bacteroid per symbiosome Several bacteroids per symbiosomeN2 fixation products

transportedAmides usually Ureides usually

Infected cells Vacuolate NonvacuolateGeographical origin Temperate Tropical to subtropicalGenera Medicago, Trifolium, Pisum,

LupinusIsoflavones

nod Gene inducers Flavones, isoflavones Glycine, Phaseolus, VignaPloidy level of bacteroids Polyploid DiploidViability of bacteroids Nonviable upon release in soil Viable upon release in soil

Source: Vance (2002), Vessey et al. (2005), Mergaert et al. (2006).


maximum expression is in the differentiatingregion of the root between the growing root tipand the zone of root-hair emergence, where thenodulation process is initiated. An increase of thecytosolic Ca concentration, which originates from

intracellular sources and from the apoplast,appears to play a role in the signal-transductionpathway that starts with the perception of the Nodfactor and leads to nod gene expression (Pingretet al. 1998).

FIGURE 15. Symbiotic signaling between legume plantsand rhizobia. (1) Flavonoids are exuded by the legumeroots. (2) The flavonoids bind to the gene product of aconstitutively expressed nodulation (nod ) gene. (3) Afterthis binding, common nod genes are activated. (4) Thisleads to the production of a lipochitooligosaccharide. (5)

The flavonoids also activate the transcription of specificnod genes. (6) The products of the specific nod genes leadto modification of the lipochitooligosaccharides and theformation of nodulation (nod) factors, which confer spe-cificity. (7) The Nod factors are recognized by receptorson the surface of the legume-host’s roots.


Much less is known about plant factors deter-mining if a rhizobium will recognize a plant as anappropriate host. Flavonoids offer only a partialexplanation. When rhizobial genes that confer hostspecificity are transferred to a rhizobium strain witha different specificity, these genes alter the bacterialacidic polysaccharide structure and in situ bindingto the host’s root hairs. Furthermore, introducingthe genes encoding specific root-hair proteins (lec-tins) into Trifolium repens (white clover) allowsnodulation of clover roots by a Rhizobium strain,which is usually specific for Pisum sativum (pea). Ithas therefore been thought that the host—Rhizobiumspecificity involves the interaction of the root hairlectins with specific carbohydrates on the bacterialsurface (Diaz et al. 1989).

3.3.3 Entry of the Bacteria

After the release of flavonoids by the host and therelease of the Nod factors by rhizobium, the bacteriamultiply rapidly in the rhizosphere. The bacteriamay adhere to root hairs and affect those that havejust stopped growing. Younger and older root hairsare not affected. Alternatively, entry may occurthrough cracks in the epidermis or wounds. This isthe only way of entry for 25% of all legumes (Sprent2007) as well as for entry into nonlegumes, e.g.,Oryza sativa (rice), Triticum aestivum (wheat), andZea mays (corn) (Webster et al. 1997). However,upon entry into nonlegumes, with the exception ofParasponia species, no nodules are formed.

When rhizobia adhere to root hairs, the cell wall ofthe affected root hair is then partly hydrolyzed atthe tip. In this process the root hairs curl, attachingthe bacteria to the root hair. On those locations on theroot hairs that have become deformed due to the pre-sence of rhizobia, the cell wall is degraded, allowingthe bacteria to enter. An infection thread is formed byinvagination of the cell wall. This thread consists ofcell-wall components similar to those that form thenormal root-hair cell wall. The infection threadgrows down the root hair at a rate of 7—10 mm h—1

and provides a conduit for bacteria to reach the rootcortex. The tip of the transcellular infection threadappears to be open; sealing of the thread tip resultsin abortion of the infection thread. The formation ofthe infection thread may well be analogous to theenlargement of epidermal cell walls, in response to apathogen’s attempted penetration (Vance 2002).

Only 1—5% of all root hairs become infected, andonly 20% of these infections result in nodules. Whyare most of the infections unsuccessful? This is likelyto be due to the production of chitinases by the

plant. These enzymes hydrolyze the lipochitooligo-saccharide Nod factor (Mellor & Collinge 1995).Legumes contain different chitinases. In an earlystage of infection the host produces a chitinase thatbreaks down the Nod factors of Rhizobium speciesthat are not suitable guests. In this way they preventthe entry of bacteria that cannot form a symbiosis.Chitinases are therefore another factor that confershost—guest specificity. At a later stage, differentchitinases are produced that are effective againstthe Nod factor of ‘‘homologous’’ rhizobium bacteria,i.e., suitable guests. This is probably a mechanism tocontrol the entry of rhizobium and to prevent morenodules being formed than can be supported by thehost. In addition, the breakdown of Nod factorsprevents entering bacteria from being erroneouslyrecognized as pathogens. Rhizobium strains thatoverproduce the Nod factor do indeed lead to adefense response. Not only is the Nod factor brokendown by plant chitinase activity, but expression ofthe bacterial nod genes is also suppressed at a laterstage of infection. Plant phenolics may play a role inthis suppression. If the Rhizobium fails to recognizethe plant-derived suppressor molecule, the bacteriamay be recognized as a pathogen and further devel-opment stops. This offers another possibility forhost—guest specificity. The train of events thatlikely plays a role in the early stages of infectionand the role of chitinases is outlined in Fig. 16.

If the infection is successful, then specific genesare activated in the cortex and pericycle whichallows the formation of an infection thread throughwhich the bacteria enter (Vessey et al. 2005). Celldivisions start in the inner cortex (indeterminatenodules) or outer cortex (determinate nodules),opposite protoxylem poles, so that a new meristemis formed due to the presence of the rhizobia. Thismeristem gives rise to the root nodule. The infectionthread grows inward, and, finally, the bacteria aretaken up into the cytoplasm of the parenchyma cellsof the center of the developing nodule. Inside theinfected host plant cell, the bacteria continue todivide for some time, now differentiating into bac-teroids, which have a diminished ability to grow onlaboratory culture media; they may be greatlyenlarged with various shapes. In most legumes,bacteroids are enclosed within a peribacteroidmembrane, to form a symbiosome (White et al.2007). Most symbiosomes are of a similar volume,but in some nodules, each symbiosome contains asingle, enlarged pleomorphic bacteroid, whereasthere may be several (up to 20) smaller, rod-shapedbacteroids in others. Symbiosomes with a singlebacteroid are more typical of elongate, cylindricalnodules of the so-called indeterminate class, as


found on Trifolium repens (white clover) or Pisumsativum (pea) (Table 5). These nodules have a persis-tent meristem. Bacteroids in indeterminate nodulesare polyploid, and have lost their ability to divide;hence they are nonviable when released fromnodules (Mergaert et al. 2006). Symbiosomes withseveral bacteroids are common in spherical, deter-minate nodules (with no meristem), such as those ofVigna unguiculata (cowpea) or Glycine max (soy-bean). Bacteroids in determinate nodules arediploid, like free-living rhizobia, and can divide insoil upon release from nodules. The ploidy level ofthe bacteroids is controlled by the host (Mergaertet al. 2006). A few legumes have nodules in whichthere are no symbiosomes, the bacteria beingretained within multiply branched infectionthreads. Mature nodules of the determinate andindeterminate types are strikingly different, buttheir initiation is rather similar (Sprent 2007).

3.3.4 Final Stages of the Establishmentof the Symbiosis

Each infected cell may contain many hundreds ofsymbiosomes. The symbiosome membrane (peri-bacteroid membrane) originates from an invagina-tion and endocytosis of the plasma membrane of theinfected cortical cells. This membrane acts as a selec-tive permeability barrier to metabolite exchangebetween the bacteroids and the cytosol of theinfected cells. Interspersed between the infectedcells of many nodules are smaller, uninfectedcells, which occupy about 20% of the total volumeof the central zone of the nodules of Glycine max(soybean). Plasmodesmata connect uninfectedwith infected cells and with other uninfected cellsin the central zone of the nodule. These plasmodes-mata allow for the massive transport of carbon fromthe uninfected cells to the infected ones and of

FIGURE 16. Tentative scheme to account forevents that determine the establishment of afunctional symbiosis (effective nodulation)between rhizobium and a host legume. ‘‘Elici-tation’’ is a combination of the elicitor activityof a specific bacterial Nod factor and the sen-sitivity of the plant to that elicitor (Mellor &Collinge 1995). Reproduced with kind permis-sion of Oxford University Press.


nitrogenous compounds in the reverse direction(Brown et al. 1995). Both infected and uninfectedcells contain numerous plastids and mitochondria.Uninfected and infected cells in nodules have dif-ferent metabolic roles in symbiotic N2 fixation (Day& Copeland 1991). The central tissue of somenodules, however, contains no uninfected cells.

In Glycine max (soybean) nodules, an outer layerof cortical cells surrounds an endodermal cell layer,which in turn encloses several layers of subcorticalcells. The central zone of the nodules containsseveral thousand infected cells. The nodule isconnected with the vascular tissue in the stele, dueto the proliferation of cells from the pericycle.

The pattern of gene expression in the host cellsthat are part of the nodules is altered by the presenceof the bacteria, resulting in the synthesis of manydifferent proteins, known as nodulins. Some ofthese nodulins have been characterized biochemi-cally, including the O2 carrier leghemoglobin andnodule-specific forms of the enzymes uricase,glutamine synthetase, and sucrose synthase.

3.4 Nitrogenase Activity and Synthesisof Organic Nitrogen

Biological reduction of N2 to NH3 is catalyzed bynitrogenase, in a highly O2-sensitive process. ThisO2 sensitivity accounts for the pink color of thenodule tissue which is due to the presence of leghe-moglobin in the cytoplasm of the infected legumecells or hemoglobin in nodules of species livingsymbiotically with Frankia (Gualtieri & Bisseling2000). This heme-protein may comprise 35% of thetotal nodule soluble protein. Leghemoglobin isrelated to the myoglobin of mammalian muscle; itsprotein component is encoded by the DNA of theplant. The enzymes responsible for the synthesis ofthe O2-binding heme-group of the protein in legumenodules are encoded both in rhizobium and inthe host [e.g., Pisum sativum (pea)]. Because theenzymes are up-regulated in the infected host cellsin root nodules that synthesize the other componentof leghemoglobin, the heme moiety is probablysynthesized by the macrosymbiont (Santana et al.1998). Leghemoglobin plays a role in the O2 supplyto the bacteroid. It has a high affinity for O2 and arelatively fast O2-dissociation rate, which ensuressufficiently rapid O2 supply for the highly activerespiratory processes in the plant and bacteroid com-partment while maintaining a low concentration offree O2 (between 3 and 30 nM). The latter is veryimportant because nitrogenase, which is the enzymeresponsible for the fixation of atmospheric N2 to

NH3, is rapidly damaged by free O2 (Arredondo-Peter et al. 1998, Gualtieri & Bisseling 2000).

To control the O2 supply and O2 concentration toand within infected cells, nodule permeability toO2 diffusion varies within seconds to hours inresponse to changes in carbohydrate supply via thephloem, adenylate demand, and O2 status. This per-meability control is associated with the reversibleflow of water into or out of intercellular spaces.When nodulated Glycine max (soybean) plants areexposed to treatments that decrease the nodules’ O2

permeability, the Kþ concentration in the nodulecortex increases relative to that in the central zoneof the nodules. On the other hand, treatments thatincrease O2 permeability have the opposite effect.The energy-dependent coupled movement of ionsand water into and out of infected cells offers apossible mechanism for diffusion barrier control inlegume nodules (Wei & Layzell 2006).

The bacteroids contain nitrogenase, which is anenzyme complex that consists of two proteins. One,nitrogenase-reductase, is an Fe—S-protein thataccepts electrons, via an intermediate electron car-rier, from NADPH and then binds ATP. At the sametime, the other subunit (an Fe—Mo-protein) bindsN2. Reduction of this N2 occurs if the two subunitshave formed an active complex. A minimum of 12,and possibly as many as 16 ATP are required per N2;therefore the overall equation is:

N2 þ 8 Hþ þ 8 e� þ 16 ATP

! 2 NH3 þH2 þ 16ADPþ 16 Pi

(1)

Most of the N2 fixed by the bacteroids is releasedas NH3 to the peribacteroid space and then as NH4

þ,via a voltage-driven channel across the peribacter-oid membrane, to the cytosol of the nodule cells(Fig. 17; Mouritzen & Rosendahl 1997). Alterna-tively, NH3 may be converted into alanine,and then exported (Fig. 17; White et al. 2007).A nodule-specific glutamine synthetase is expressedin the cytosol of infected cells. Glutamine 2-oxoglu-tarate aminotransferase (GOGAT) then catalyses theformation of two molecules of glutamate from onemolecule of glutamine. The major N-containing pro-ducts exported via the xylem are the amides aspar-agine and glutamine in such species as Pisumsativum (pea), Medicago sativa (alfalfa), and Trifoliumrepens (white clover). These products are typical fornodules that are elongate-cylindrical with indeter-minate apical meristematic activity (Fig. 18, Table 5).In Phaseolus vulgaris (common bean) and Glycine max(soybean), the products are predominantly ureides:allantoin and allantoic acid (Fig. 18, Table 5).Nodules exporting these compounds are spherical


FIGURE 17. (Top) Electron micrograph of a nodule ofGlycine max (soybean), infected with Bradyrhizobiumjaponicum. Three infected cells and one uninfected cellare shown, with two air spaces in between. Note thatthe uninfected cell is much smaller than the infectedones, and that the bacteroids are grouped as ‘‘symbio-somes’’, surrounded by a peribacteroid membrane(courtesy D. Price, Research School of BiologicalSciences, Australian National University, Canberra,Australia). (Bottom) Scheme of N2-fixation and NH3

production in bacteroids. NH3, being an unchargedmolecule, can cross the bacteroid membrane andarrives in the peribacteroid space, where it picks up aproton and becomes NH4

+. Subsequently, NH4+ leaves

the symbiosome via a monovalent cation channel (1).Alternatively, NH3 may react with pyruvate, producingalanine. An ATPase in the peribacteroid membrane (2)creates a membrane potential (positive inside of thesymbiosome), which drives the passive uptake of nega-tively charged organic acids (predominantly malate andsuccinate) into the symbiosome (3). Alternatively,organic acids may arrive symplastically from neighbor-ing uninfected cells. An electron-transport chain in thebacteroid membrane (etc.) creates a proton-motiveforce, which drives the uptake of malate (and succinate)into the bacteroids via a H+-co-transport mechanism (4)[Whitehead et al. (1995), White et al. (2007)].


with determinate internal meristematic activity(Vessey et al. 2005).

The ureides released to the xylem of plants withdeterminate nodules are products that are onlyfound when the symbiotic plants are fixing N2

(Peoples et al. 1996). Hence the concentration ofthese compounds in the xylem sap, relative to thetotal amount of N transported in the xylem, has beenused to estimate the proportion of N derived fromN2 fixation, as opposed to the assimilation of com-bined N (Peoples et al. 1996). The amides released tothe xylem of plants with indeterminate nodules arealso found when these plants grow nonsymbioti-cally. In fact, they are not even typical for legumes.Hence, they cannot be used as ‘‘markers’’ for sym-biotic N2 fixation.

3.5 Carbon and Energy Metabolismof the Nodules

Carbohydrates are supplied via the phloem to thenodules, where they are rapidly converted in the

plant compartment to dicarboxylic acids (malate,succinate), predominantly in the uninfected cells inthe nodules (Lodwig & Poole 2003, White et al.2007). Malate and succinate are the major substratesfor the bacteroids (Fig. 17). How do the infected cellsprevent a large part of the organic acids from beingoxidized via the nonphosphorylating alternativepath in a situation where the demand for organicacids of the bacteroids is very large (Sect. 2.3 ofChapter 2B on plant respiration)? Mitochondriafrom nodules have very little alternative path capa-city; the little capacity they have is restricted to theuninfected cortical cells, rather than to the infectedones (Table 6). There is, therefore, no risk of oxidiz-ing the organic acids destined for the bacteroids.

Apart from N2, H+ is also reduced by nitrogen-ase [see Equation (1) in Sect. 3.4], leading to theproduction of H2. Most rhizobia, however, containhydrogenase, which is an enzyme that consumesH2, using it as an electron donor. The characteristicof nitrogenase to reduce acetylene (ethyne) to ethy-lene (ethene) is frequently used to assay nitrogen-ase activity in vivo. Because the assay itself,

FIGURE 18. Major nitrogen transportproducts from legume nodules. TheC:N ratio of ureides is 1:1, whereasthat of amides is 2:1 (asparagine) or2.5:1 (glutamine).

TABLE 6. Cyanide-resistant, SHAM-sensitive respiration in infected anduninfected cells isolated from Glycine max (soybean).*

O2 consumption [nmol mg–1 (protein) min–1]

Cells from rootnodules Control

KCN-resistantrespiration

KCN resistantrespiration (%)

Infected 60 0 0Uninfected 45 22 49

Source: Kearns et al. (1992).* Measurements were made on isolated mitochondria from different tissues, as wellas on infected and uninfected cells from root nodules.


however, interferes with the process of N2 fixation,it should only be considered as a qualitative to semi-quantitative indicator for the occurrence of nitro-genase activity, rather than as a good quantitativemeasure for its actual activity (Hunt & Layzell1993, Vessey 1994).

3.6 Quantification of N2 Fixation In Situ

The contribution of symbiotic N2 fixation to the totalaccumulation of N in the above-ground biomass ofa crop or a plant community can be determinedby applying 15N-labeled inorganic N (15NO3

—

or 15 NH4þ) separately to N2-fixing plants and refer-

ence plants (i.e., nonfixing species or mutants). Forexample, it can be given to a plant community thatconsists of both N2-fixing species (e.g., clover) andother species (e.g., grasses) (15N is a nonradioactiveisotope of N). The grasses have a 15N/14N ratio,which is used as a reference. N2 fixation in the N2-fixing clovers will ‘‘dilute’’ their 15N concentration.The extent of the dilution is used to calculate thecontribution of fixation to the total amount of N thataccumulates in the clover plants. The contribution ofN2 fixation to the total amount of N that accumu-lates in the plant may amount to 75 and 86% inTrifolium repens (white clover) and Trifolium pratense(strawberry clover). Transfer of N is most likely viarelease of ammonium and amino acids from legumeroots (Paynel et al. 2001), but under some conditionsN transfer is not detected (McNeill & Wood 1990).The contribution depends on the amount of inor-ganic combined N that the plants receive from soiland fertilizer, and also varies with the developmentalstage of the plant and the time of the year (Fig. 19).The overwhelming importance of symbiotic N2

fixation in some agricultural systems is illustratedin Table 7.

Sometimes the natural abundance of 15N inthe soil is used to quantify N2 fixation (Table 2 inSect. 2.4). Instead of adding 15N-labeled inorganiccombined N, the natural abundance of N in the soilcan be used. The natural abundance of soil N islikely to differ from that of N2 in the atmosphere,due to discrimination against the heavy isotope invarious biological processes (e.g., nitrification anddenitrification). N in the soil is therefore ‘‘enriched’’with the heavy isotope, relative to N2 in the air. Toapply this technique in situ, reliable control plantshave to be used. As discussed in Sect. 2 (Table 2), thisis not always easy, if it is at all possible (Handleyet al. 1993).

The 15N technique referred to earlier has alsobeen used to demonstrate a significant transfer of

FIGURE 19. The contribution of symbiotic nitrogen fixationto the total N accumulation in the above-ground biomassof Trifolium repens (white clover) (top) and Trifolium pra-tense (strawberry clover) (bottom). The white clover plantsgrew in a mixed culture with Lolium perenne (perennialryegrass) and the strawberry clover with Lolium multi-florum (Italian ryegrass). Circles: no N fertilizer; triangles:30 kg N ha–1 per cut; open and closed symbols refer todifferent years (Boller & Nosberger 1987).

TABLE 7. Symbiotic N2 fixation by some legume crops1

and native species in their natural environment inBrazil2.

Species

N2 fixedkg ha–1

per season

Plant N absorbedfrom atmosphere

(%)

Medicago sativa 440–780 65–96Glycine max 120 53Lotus corniculatus 92 55Lupinus angustifolius 170 65Medicago sativa 180 70Phaseolus vulgaris 65 40Pisum sativum 72 35Trifolium pratense 170 59Vicia faba 151 ndVigna angularis 80 70Chamaecrista

speciesnd 66–79

Mimosoid legumes nd 42–63Papilionoid legumes nd 68–79

Source: 1Gault et al. (1995), Vance (2002); 2Sprent et al.(1996).


N from symbiotic plants to neighboring grasses (upto 52 kg N ha—1 year—1; on average a value of 17 kgha—1 year—1 is found). Conditions favoring N2 fixa-tion by the legume, such as a high irradiance, afavorable temperature, long days, and a relativelyhigh Pi supply, enhance the transfer of N from thelegume to the nonfixing neighbors. This transfer isto some extent the result of the uptake of nitrogen-ous compounds released after decomposition ofparts of the legumes. Some of it is also due, however,to the exudation of nitrogenous compounds by thelegumes, followed by absorption by the nonfixingplants. Some transfer of N may occur throughmycorrhizal hyphae (Sect. 2.3.1).

The maximum yield of Lolium rigidum (annualryegrass) is reached at a much lower supply of Pi

than that of Trifolium subterraneum (subclover), atleast when both species are grown in monoculture(Bolan et al. 1987). This demand for a higher P isfairly common for nodulated legumes, although it isby no means universal (Koide et al. 1988, Sprent1999). The high demand for P of many crop legumesmay reflect their adaptation to soils with a highavailability of Pi, and it points out that a benefitfrom such legumes can only be expected whenthese have access to sufficient Pi. Apart from Pi,also Mo and S have to be available to the legume toallow symbiotic N2 fixation.

What is the role of N2-fixing species in morebiodiverse grasslands? To address this question,test plants (‘‘phytometers’’) can be planted in plotsunder investigation, to sample their above-groundbiomass at a later stage (both N concentration andnatural abundance of 15N: �15N). Phytometers in arecent study belonged to four ‘‘plant functionalgroups’’ (Chapter 9E): Festuca pratensis (meadowfescue), Plantago lanceolata (snake plantain), Knautiaarvensis (field scabious), and Trifolium pratense(strawberry clover). Significantly lower �15N valuesand higher N concentrations and N contents were

found in all phytometer species growing withlegumes, indicating a facilitative role for legumesin these natural grassland ecosystems. The magni-tude of the positive interactions depends on theexact phytometer species, but increased N uptakein communities containing legumes is found in allthree nonlegume phytometer species, with a subse-quent strong increase in biomass in the grass Festucapratensis across all diversity levels, and a lesser bio-mass gain in Plantago lanceolata and Knautia arvensis.In contrast, the legume phytometer species Trifoliumpratense is negatively affected when other legumesare present in their host communities across all diver-sity levels (Temperton et al. 2007).

3.7 Ecological Aspects of theNonsymbiotic Association with N2-FixingMicroorganisms

Next to the truly symbiotic associations that lead toN2 fixation, as discussed in Sect. 3.3, a somewhatlooser association between Azospirillum and higherplants (especially grasses) has been investigated.Inoculation of the soil in which Zea mays (corn)plants are grown with Azospirillum bacteria signifi-cantly enhances the yield of the corn plants, espe-cially when the N supply is relatively low (Table 8).It is by no means certain, however, that this is adirect result of the fixation of N2 by the Azospirillumbacteria. These organisms are more likely toenhance the growth of higher plants in a differentmanner, such as the production of phytohormones(Dobbelaere et al. 1999). In a comparison of threecultivars of Triticum aestivum (wheat), grown withAzospirillum brasiliense, most N2 is fixed in the rhizo-sphere of Al-resistant cultivars. Because these culti-vars also exude more dicarboxylic acids (Sect. 3.1 ofChapter 6 on mineral nutrition), it has been sug-gested that fixation is enhanced by the excretion of

TABLE 8. The effect of inoculation with Azospirillum brasiliense on the productionof Zea mays (corn) plants as dependent on the N supply.*

Shoot dry mass (g) Root dry mass (g)

N supply(g L�1) Inoculated Control Inoculated Control

Relative increment oftotal plant mass (%)

0 0.49 0.32 0.36 0.27 440.04 0.97 0.66 0.76 0.53 450.08 1.84 1.23 0.97 0.86 340.16 2.93 2.52 1.96 1.70 16

Source: Cohen et al. (1980).* N was supplied as NH4NO3.


these organic molecules (Christiansen-Weniger et al.1992). Microorganisms might promote plant growthin many other ways, such as suppression of patho-genic organisms and the production of vitamins.

In some areas in Brazil, Saccharum officinarum(sugarcane) has been grown continuously for morethan a century without any nitrogenous fertilizer.Although it had long been suspected that substan-tial N2 fixation occurs in such systems, none of theN2-fixing bacteria isolated from the rhizosphere ofSaccharum officinarum occur in large enough num-bers to account for the high rates of N2 fixationfound in these crops. An acid-tolerant N2-fixingbacterium (Glucoacetobacter diazotrophicus and arange of others) has been identified in the intercel-lular spaces of sugarcane stem parenchyma (Donget al. 1994, James et al. 1994). These spaces are filledwith a solution that contains 12% sucrose (pH 5.5).Glucoacetobacter diazotrophicus has most unusualgrowth requirements; it shows optimal growthwith 10% sucrose and pH 5.5. It will grow in amedium with 10% sucrose and rapidly acidifies itssurroundings by the formation of acetic acid. It hasbeen isolated from sugarcane tissues, but was notfound in the soil between rows of sugarcane or ingrasses from the same location. The apoplastic fluidoccupies approximately 3% of the stem volume,which is equivalent to 3 tons of fluid per hectare ofthe sugarcane crop. It has been suggested that thisamount suffices to make the sugarcane independentof N fertilizers.

Enterobacter agglomerans, Herbaspirillum seropedi-cae, and Klebsiella terrigena are also believed to be

able to fix atmospheric N2 in the apoplast of plantsthat have high apoplastic sugar concentrations.Some of these (e.g., Herbaspirillum seropedicae) arepathogens on certain grass species which restrictstheir potential use as inoculants in agriculture(Palus et al. 1996, Triplett 1996). To date, with theexception of sugarcane, little success has beenattained in elucidating which endophyte is reallyresponsible for the observed biological N2 fixation,and in what site, or sites, within plants the N2 fixa-tion mainly occurs. Until such important questionsare answered, further developments or extension ofthis novel N2-fixing system to other economicallyimportant nonlegumes (e.g., cereals) will be ser-iously hindered (Boddey et al. 2003).

3.8 Carbon Costs of theLegume–Rhizobium Symbiosis

Because all the organic acids required for symbioticN2 fixation by rhizobium and for maintenance ofthe root nodules come from the plant (Fig. 17), thereare costs involved in this symbiotic system for thehigher plant. These costs exceed those required forassimilation of NO3

— or NH4+. They have been esti-

mated for an association of Trifolium repens (whiteclover) and rhizobium, in which the clover plantsare totally dependent on the microsymbiont fortheir supply of N (Fig. 20). In this symbiotic system,the N2 fixation is briefly interrupted by decreasingthe O2 concentration that surrounds the plants, butkept sufficiently high to fully maintain aerobic

FIGURE 20. Respiration and acetylene reduction (mea-sured as ethylene production) in roots of Trifolium repens(white clover) at either high or low O2 concentration.The low O2 concentration was sufficiently high to main-tain aerobic metabolism of the plant cells, but virtuallycompletely abolished the activity of the N2-fixing activity

of the bacteroids. The decline in respiration after addingacetylene reflects the sensitivity of the nodules to‘‘manipulations’’. It also shows that this technique cannotbe used to give reliable quantitative estimates of the rateof N2 fixation (Ryle et al. 1985). Reproduced with kindpermission of Oxford University Press.


metabolism of the plant. It is sufficiently low, how-ever, to completely block the respiration and N2

fixation by the bacteroids. By relating the decreasein respiration upon blocking the N2 fixation to theactivity of N2 fixation as determined from the accu-mulation in the clover plants, carbon costs per unitfixed N are calculated. The costs for N2 fixationamount to approximately 25% of all the carbonfixed in photosynthesis per day. This proportionis rather high when compared with the figures forN acquisition by nonsymbiotic plants given inTable 2 of Chapter 2B on respiration: 4—13% whenplants are grown at an optimum nutrient supply.At a limiting nutrient supply, the percentage islikely to be more similar to that of the costs of N2

fixation.Because of the high carbon costs of symbiotic N2

fixation, it has been suggested that elevated atmo-spheric CO2 concentrations will stimulate thisprocess. However, a meta-analysis shows that sym-biotic N2 fixation is stimulated by elevated atmo-spheric [CO2] only when sufficient soil nutrients,other than N, are available (Van Groenigen et al.2006). Short-term experiments frequently show apositive effect of elevated atmospheric [CO2] onsymbiotic N2 fixation, but in the long run, a reducedavailability of, in particular, Mo, a key compo-nent of nitrogenase, leads to rates of N2 fixationsimilar to those under ambient [CO2] (Hungateet al. 2004) .

3.9 Suppression of the Legume–RhizobiumSymbiosis at Low pH and in the Presenceof a Large Supply of Combined Nitrogen

Once rhizobia have successfully infected legumeroots and nodules have been formed, further infec-tion is suppressed. This is called autoregulation ofroot nodule formation, a phenomenon we encoun-tered before in the establishment of a mycorrhizalsymbiosis (Sect. 2.3.1). Using plants grown with adivided root system, it can be shown that autoregu-lation depends on systemic signals (Van Brusselet al. 2002). Infection of one root of Vicia sativa (com-mon vetch) with Rhizobium leguminosarum bacteriainhibits nodulation of a spatially separated root,when this root is inoculated 2 days later with thesame bacteria. The mechanism by which nodulationis autoregulated is related to that by which com-bined N inhibits nodulation (see below). Genesthat are involved in Nod-factor signaling may betargets for mechanisms that suppress nodulation(Limpens & Bisseling 2003).

At low pH, nodule formation on legumes tendsto be inhibited. Because fixation of N2 lowers soil pH(Sect. 3.1 of Chapter 6 on mineral nutrition), contin-ued use of legumes in agriculture requires regularliming. Why is nodulation impaired at a low soilpH? The assay system described in Sect. 3.3.1 hasbeen used to establish that a relatively acid or alka-line, as opposed to a neutral pH of the soil, leadsto less effective root exudates. This offers an expla-nation for the common observation of a poor infec-tion of legumes by rhizobium in acid soils. Survivalof rhizobia is also lower in soils with a low pH, butsome degree of adaptation of rhizobia strains hasbeen observed.

N2 fixation is an energetically expensive process, ascompared with the assimilation of NO3

— or NH4+. Remi-

niscent of the effect of Pi on the formation of the mycor-rhizal symbiosis, NO3

— often inhibits the infection oflegumes by rhizobia, but this is not invariably found(Sprent 1999). When Medicago sativa (alfalfa) plants aregrown under N-limiting conditions, the expression ofthe genes involved in flavonoid biosynthesis and theproduction of root flavonoids are enhanced. This mayaccount for greater infection by Rhizobium meliloti underconditions when N is in short supply, as opposed tosuppression of nodulation in the presence of high NO3

—

concentrations (Coronado et al. 1995).

TABLE 9. Apparent nitrogenase activity and the O2-limitation coefficient, 2 days after addition of NO3

–

to the root environment of nodulated 21-day-oldplants of Pisum sativum (pea).*

[NO3](mM)

Apparent nitrogenaseactivity [nmol H2 g�1

(nodule dry mass) s�1]O2 limitationcoefficient

0 45 0.895 38 0.6410 22 0.4515 24 0.49

Source: Kaiser et al. (1997).* The apparent nitrogenase activity was measured as the rate ofH2 evolution. As explained in Sect. 3.4, nitrogenase activityleads to the production of H2. There is normally no net evolu-tion of H2, because rhizobia have a hydrogenase (i.e., anenzyme that uses H2 as an electron donor). In the presentexperiment, a rhizobium strain was used that lacks hydroge-nase so that the evolution of H2 could be measured. The O2

limitation coefficient is calculated as the ratio between totalnitrogenase activity (H2 evolution in the absence of N2) andpotential nitrogenase activity (H2 evolution in the absence ofN2 at an optimum concentration of O2).


NO3— also inhibits the process of fixation itself

(Table 9). Several mechanisms have been proposedto account for this inhibition (Hunt & Layzell 1993):

1. Competition for carbohydrates between nitro-genase and nitrate reductase, located in leaves,roots, or nodules

2. Inhibitory effects of NO2—, the product of nitrate

reductase. NO2— may inhibit nitrogenase directly,

by irreversibly binding to the enzyme, or indir-ectly, by forming a bond with leghemoglobin, sothat it can no longer function in O2 transport

3. A decrease of the partial pressure of O2 in thenodule, due to a decrease in the conductance forgas transport in the pathway between the outsideair and the infected cells

4. Feedback inhibition of nodule metabolism bynitrogenous compounds that arrive via the phloem

There is evidence for a decrease in the conductancefor O2 transport to the infected cells which leads to amore severe limitation of nitrogenase activity by O2

(Table 9). It is unlikely, however, that this is the onlymechanism that accounts for NO3

— inhibition ofnodule activity. Rather, all four mechanisms probablyoccur at one stage or another in some species.

Several mutants of a number of legumes havebeen produced, of which neither infection nor N2

fixation itself is inhibited by nitrate. These mutants

are expected to enhance input of N through thelegume—rhizobium symbiosis in agriculturalsystems.

4. Endosymbionts

Many plants are infected by fungal endophytes(family Clavicipitaceae, Ascomycota) that live theirentire life cycle within a plant (Bacon & De Battista1991). The fungi form nonpathogenic and usuallyintercellular associations in living plant tissue. Theendophytes are often transmitted through the plantseed, particularly in grasses and sedges, but seedsmay lose their endophytes upon prolonged storage.Infection through germinating spores is an alterna-tive way to enter the macrosymbiont. The associa-tion between higher plants and endosymbiotic fungihas been well studied in grasses in which the fungimay produce alkaloids in the tissue of their hosts,many of which have a neurotoxic effect, and hencemake the infected plants poisonous to domesticmammals and increase their resistance to insectherbivores (Table 10).

In some species, plant growth and seed produc-tion can be increased by infection with the endo-phyte. The symbiotic associations between grasses

TABLE 10. Antiherbivore effects of fungal endophytes that infect grasses.

AnimalHost genus

grassFungal endophyte

genus Comments

MammalsCattle, horses Festuca Acremonium Reduced mass gain, gangrene, spontaneous abortionCattle, sheep, deer Lolium Acremonium Reduced mass gain, tremors, staggers, deathCattle, goats Andropogon Balansia Reduced milk production, deathCattle Paspalum Myriogenospora Reduced mass gain, tremors gangrene

InsectsFall armyworm Cenchrus Balansia Avoidance, reduced survival, reduced growth, increased

development timeCyperus BalansiaFestuca AcremoniumLolium AcremoniumPaspalum MyriogenosporaStipa Atkinsonella

Aphids Festuca Acremonium AvoidanceBillbugs Lolium Acremonium Reduced feeding and ovipositionCrickets Lolium Acremonium Complete mortalityCutworms Dactylis Epichloe Reduced survival and mass gainFlour beetles Lolium Acremonium Reduced population growthSod webworms Lolium Acremonium Reduced feeding and ovipositionStem weevils Lolium Acremonium Reduced feeding and oviposition

Source: Clay (1988).Note: The examples are representative but not exhaustive.


and fungal endophytes may be an association inwhich the fungi derive carbohydrates from theirhost and defend their host against herbivory,thereby defending their own resources. Similar tothe effect that mycorrhizal fungi have on interac-tions among mycorrhizal and nonmycorrhizalplants (Sect. 2.2), fungal endophytes may also influ-ence competitive interactions between plants. Forexample, grass plants infected with fungal endo-symbionts are less nutritious (Clay et al. 1993) andpreferred less than the uninfected plants of the samespecies. The presence of endophytes may also affectcompetition among grasses in interaction with her-bivory (Clay et al. 1993) and suppress the fungaltake-all disease in Triticum aestivum (wheat)(Dewan & Sivasithamparam 1988).

The presence or absence of fungal endophytes isnot a specific trait of a plant species; rather, itdepends on environmental conditions in an as-yet-unclear manner. For example, in WesternAustralian heaths (Ericaceae), the number of fungalassociates is considerably smaller on a mesicwetland site when compared with a dryland habitat,even when comparing the endophytes associatedwith the same plant species (Lysinema ciliatum).This appears to reflect the response of different fun-gal endophytes to water stress (Hutton et al. 1996).

Bacteria may also act as endosymbionts. Someplant-growth-promoting endosymbiotic bacteria havealready been discussed in Sect. 3.7: Gluconacetobacterdiazotrophicus, which fixes N2 in the tissues of Saccharumofficinarum (sugarcane). Common endophytic bacteriafrom healthy tubers of Solanum tuberosum (potato)belong to six genera (Pseudomonas, Bacillus, Xantho-monas, Agrobacterium, Actinomyces, and Acinetobacter).As discussed in Sect. 3 of Chapter 9C on effects ofmicrobial pathogens, many bacterial endophytesmake the host plant more resistant to pathogen attack(induced resistance) or they enhance growth. Thereare also endophytic bacteria, however, that are plant-growth-neutral or plant-growth-retarding (Sturz 1995).

5. Plant Life AmongMicrosymbionts

At one stage in the history of plant ecophysiology itmay have seemed most appropriate to discuss themineral nutrition and performance of plants devoidof their microsymbionts. If we wish to unravel basicprinciples of plant mineral nutrition (e.g., the natureof a NO3

— transporter or the function of a micronu-trient), then this remains a valid approach. If the aimis to understand plant functioning in a real

environment, whether a natural ecosystem or anagricultural field, however, then we cannot ignorethe existence and overwhelming importance of themicrosymbionts that interact with higher plants inan intricate manner. This is most certainly true formycorrhizal fungi, which affect both mycorrhizaland nonmycorrhizal species, although in a very dif-ferent manner.

Plant—microbe interactions do not always receivethe attention they deserve. Interactions with N2-fixing symbionts have been the target of plant phy-siological research for a long time. Why buy N if youcan grow your own? In recent years there has beenan enormous development in the understanding ofsignaling between rhizobia and legumes. Similarsignaling processes probably exist between otherN2-fixing microsymbionts and their nonlegumehosts, and major progress has been made on signal-ing between mycorrhizal fungi and their macrosym-biotic partners.

Endophytes other than the ‘‘classic’’ mycorrhizalfungi and N2-fixing microorganisms include the fas-cinating N2-fixing microorganisms in the apoplast ofSaccharum officinarum (sugarcane) and toxin-producingendophytes in grasses. We are only just beginning tounderstand the agronomic and ecological significanceof these endophytes. Another question that remains tobe answered is how symbiotic microorganisms areallowed entry into the plant when we know that plantshave a wide array of defense mechanisms to keepmicroorganisms at bay.

In this chapter we have showcased one of manyareas in plant physiological ecology where ‘‘estab-lished’’ terms like ecology and molecular plantphysiology have become obsolete. We can onlyfurther our basic understanding of interactionsbetween plants and their microsymbionts if we abol-ish barriers that hinder the developments in thisfield. Many applications of a basic understandingof symbiotic associations between plants and micro-organisms are to be expected.

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