the ectomycorrhizal symbiosis: life in the real world

Mycologist, Volume 19, Part 3 August 2005. ©The British Mycological Society Printed in the United Kingdom.DOI: 10.1017/S0269915XO5003034

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Introduction

A recent volume entitled ‘Mycorrhizal Ecology’ (van derHeijden & Sanders, 2002) successfully broughttogether many current ideas on the function andecological significance of the mycorrhizal symbiosisand provides a good overview of the ectomycorrhizal(ECM) symbiosis. In this short article we focus primarilyon a small number of selected topics relating ECM fungito the biotic and abiotic environment in which theyoccur. After a short definition of the structural aspectsof the symbiosis, up-to-date summaries of the plantand fungal symbionts and their global distributions areprovided. The potential of ECM fungi to complete theirlifecycles as free-living organisms is then discussed.Finally, the question of where ECM fungi stockpileresources for sporocarp formation and the mechanismsevolved by the fungi to defend these stores are explored.

Ectomycorrhizas – a definition

The ECM symbiosis is typically formed between theterminal feeder roots of woody perennial plant speciesand a range of soil fungi (Smith and Read, 1997). Thefungi exchange soil-derived nutrients for carbohydrates

from the host plant. Nutrient uptake into the host isenhanced both as a consequence of the physicalgeometry of the fungal mycelium and by the ability ofthe fungi to mobilise N and P from organic substratesthrough the action of secreted catabolic enzymes(Leake & Read, 1997).

Within the root, the fungus ramifies between theouter cells forming a complex structure called theHartig net (Fig. 1), which provides a large surface areaof contact between the fungus and the host, allowingefficient transfer of metabolites. External to the root, amulti-layered, hyphal structure called the mantle orsheath develops (Fig. 1). Agerer (1987-2002) hasrecognised two main types of hyphal developmentwithin ECM mantles: pseudoparenchymatous - denselypacked, highly differentiated hyphal elements, andplectenchymatous - loosely interwoven hyphae, wheretheir linear nature is still evident. The hyphalarrangement within the mantle, particularly whenseen in plan view, has been used by Agerer and co-workers to characterise the mantles formed byindividual species as an aid to identification (Agerer1987-2002; Agerer et al., 1996 - 2004).

Many ECM fungi form mantles that are hydrophobic(Agerer, 1987-2002), implying that there is little direct

The ectomycorrhizal symbiosis: life in the real world

ANDY F. S. TAYLOR1 & IAN ALEXANDER2

1 Department of Forest Mycology and Pathology, Swedish University of Agricultural Sciences, PO Box 7026, Uppsala,Sweden, e-mail: [email protected] School of Biological Sciences, University of Aberdeen, AB24 3UU, Scotland. UK, e-mail: [email protected].

Ectomycorrhizas (ECM) are dual organs formed between the terminal feeder roots of many plant species and certain soil fungi. The species richness and taxonomic diversity of ECM symbionts is impressive: ca. 7-10,000 fungal and ca. 8,000 plant species may be capable of forming ECM. The latter are the dominant components offorest and woodland ecosystems over much of the earth’s surface. The obligate nature of the symbiosis for ECMfungi has been brought into question by reports that some species produce sporocarps under field conditions in theabsence of a host plant. We suggest that there is no unequivocal evidence to support this. The spread of tree rootsis often underestimated and small, overlooked hosts such as dwarf Salix spp or sedges may explain the appearanceof ECM sporocarps in vegetation apparently devoid of ECM hosts. Compared to plant material, the sporocarps ofECM fungi contain high concentrations of N and P. We show that it would take between 3 and 14 million mycorrhizal tips, or 1800 km of hyphae, to supply the N in one sporocarp of Boletus edulis. The mantle formed bythe fungus over the root tip is the likely site of storage for the N and P required for sporocarp production, and wediscuss the chemical and structural mechanisms developed on mantles by ECM fungi to defend this resourceagainst fungivory.

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exchange of solutes (uptake or exudation) with the soilsolution. Even in those mantles that are hydrophilic(e.g. many Lactarius species), there appears to be a tightcontrol over the movement of material through themantle (Ashford et al., 1988). These mantle features,coupled to the fact that the proportion of root tipscolonised is usually close to 100% (e.g. Taylor et al.,2000), mean that the host is effectively isolated fromthe soil environment. This isolation has importantimplications: any nutrients and water entering the rootmust first pass through the mantle, as must anymaterial leaving the root. Ectomycorrhizal fungitherefore occupy and, most probably control, theinterface between the soil environment and the hostplant. While the mantles may control the fluxes intoand out of the root, the mycelium extending out fromthe mantle surface (the extraradical or extramatricalmycelium) is considered to be the primary site fornutrient and water uptake.

The extramatrical mycelia produced by ECM fungivaries from a small number of hyphae growing out afew mm (e.g. many Russula spp, Fig. 2a), to highlydeveloped, extensive mycelial systems (e.g. Suillus spp,Cortinarius spp - Fig. 2b) that occupy large volumes ofsoil surrounding the colonised root tips (Agerer 1987-2002). The critical importance of this extramatricalmycelium in nutrient uptake has been emphasized inrecent years (Read & Perez-Moreno, 2003) and severalrecent studies have utilised molecular markers tolocalise the mycelium of ECM fungal species in differentsoil layers and substrates (Dickie et al., 2003; Guidot etal., 2003; Landeweert et al., 2003; Koide et al., 2004).In the quest to understand the roles of ECM fungi innature, we have progressed from counting sporocarps(e.g. Brandrud, 1995) to counting root tips (e.g. Tayloret al., 2000) to where we are today, localising mycelia.The next step will be to quantify the activity ofindividual species and/or individuals in situ. It is likelythat the large interspecific variation observed in therelationship between production of sporocarps andnumbers of associated mycorrhizas (see Gardes andBruns, 1996) will also hold between numbers ofmycorrhizas and the quantity and activity of theassociated mycelium (Koide et al., 2005).

The ectomycorrhizal mycobionts

Fungi capable of forming ectomycorrhizal associationsare believed to have evolved on a number ofoccasions from a diverse range of saprotrophic fungi(Hibbett et al., 2000). The great majority, ca. 95%, ofECM fungal species are homobasidiomycetes; theremaining species being ascomycetes (4.8%) and a few

zygomycetes within the genus Endogone (Molina et al., 1992). However, a recent study by Weiss et al. (2004) demonstrated that the importance ofheterobasidiomycetes within the Sebacinaceae asmycorrhizal formers has been underestimated.Members of this family have been strongly implicatedas the mycobionts in ECM (e.g. Urban et al., 2003),orchid (e.g. Taylor et al., 2003) and ericoid mycorrhizas(e.g. Allen et al., 2003). The recent description ofmycorrhizal associations in jungermannoid liverwortsalso seems to involve members of this family (Kottke etal., 2003). It seems likely that as the number ofmolecular studies on mycorrhizal symbionts increasesso will the taxonomic range of the mycobiontsidentified.

Within the homobasidiomycetes the ECM habit iswidespread, with seven of the twelve clades recentlyrecognised by Larsson et al. (2004) containing ECMtaxa. The basidiomes formed by ECM fungi areconsequently very diverse and include thin, crust-like(resupinate), coral-like (clavarioid), cantharelloid, andagaricoid as well as boletoid structures. The majority ofECM species are euagarics, and many of the mostfrequent and familiar sporocarps (e.g. Amanita spp, Fig.3) that appear in forests in the autumn are formed byECM taxa.

It is unusual for basidiomycete genera to displaymixed trophic status; usually all species within a‘mycorrhizal’ genus form mycorrhizas. However, whena genus is mycorrhizal it does not mean that the fungican form only ectomycorrhizas. A single fungal speciesmay be capable of forming ecto- and arbutoidmycorrhizas (Smith & Read, 1997) on different hostspecies (Horton et al., 1999). Typical ECM fungi canalso form orchid and monotropoid (Ericaceae)mycorrhizas (see Leake, this volume). At one time thegenus Paxillus was considered to be an exception as itwas thought to contain both ECM formers (P. involutusand P. rubicundulus) and saprotrophic species (e.g. P.atromentarius and P. panuoides). However, thesaprotrophic species have now been transferred to thegenus Tapinella (Sutara, 1992), a move supported bymolecular analyses (Kretzler and Bruns, 1999).

With the exception of some Tuber spp (see Murat etal., 2004), our knowledge of the ecology of ascomyceteECM fungi is very limited. Recent work (Vrålstad et al.,2000, 2002) suggests that the occurrence andimportance of the Helotiales as ECM fungi may havebeen underestimated. Villareal et al. (2004)demonstrated that an isolate from the Hymenoscyphusericae aggregate (Vrålstad et al, 2000) was capable ofsimultaneously forming ECM on Pinus sylvestris L. andericoid mycorrhizas with Vaccinium myrtillus L. It has


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forming ectomycorrhizas is relatively small. Around8000 spp., about 3%, of seed plants formectomycorrhizas (Meyer, 1973; see Smith & Read,1997, for a list of genera) but this minority of plantspecies is of enormous ecological and economicimportance, because they are the dominantcomponents of forest and woodland ecosystems overmuch of the earth’s surface. The great majority of ECMplants are woody perennials (Fitter & Moyersoen,1996). However, some sedges (Kobresia spp., see below),and herbaceous Polygonum spp. also formectomycorrhizas (Massicotte et al., 1998).

Conventional wisdom holds that ectomycorrhizalassociations are particularly characteristic of temperateand boreal forest, and that their occurrence elsewhereis patchy and of less ecological interest. It is certainlytrue that the forest dominants of the temperate andboreal zone (Fagaceae, Betulaceae, Salicaceae,Myrtaceae sf Leptospermoideae, Pinaceae) arehabitually ectomycorrhizal under natural conditions,and that the ECM habit shows particular adaptationsfor nutrient capture in temperate and boreal forests(Read & Perez-Moreno, 2003). However, much of rest ofthe land surface also supports vegetation with a strongECM component. Arctic and alpine habitats in thenorthern hemisphere are characterised by dwarf shrubcommunities of Dryas and Salix spp.: these are ECMplants that support species-rich communities ofmycobionts. Similarly, the winter-wet ecosystems of theMediterranean basin and California have a strongECM/arbutoid mycorrhizal component (Pinus, Cistus,Arbutus, Arctostaphylos).

However, it is in the tropics that the occurrence andimportance of ECM host species has been mostconsistently underestimated. Consider theDipterocarpaceae, of which all members formectomycorrhizas. This diverse family of over 500 spp. isthe source of most tropical hardwood timber and is ofimmense economic and ecological importance. Therange of dipterocarps extends from East Africa andMadagascar, through India, Bangladesh and Sri Lanka,to SE Asia, extending from S China in the north toPapua New Guinea in the south. There is one genus(Pakaraimea) in S. America. Dipterocarps make up 80%of the canopy trees and up to 40% of the understorey inSE Asian lowland and montane rain forest. They alsodominate the dry monsoonal forests of N. India, Burmaand Thailand. Dipterocarps are Gondwanan in origin(Ashton, 1982) and probably drifted away fromMadagascar/East Africa with the India/Seychelleslandmass around 88 million years ago, before radiatingthrough Asia. Recently the Sarcolaenaceae, anendemic family from Madagascar sharing a common

long been realised that ectomycorrhizal canopy treesand understorey ectomycorrhizal, or arbutoid mycorrhizal,shrubs might be linked by a common mycorrhizalnetwork (e.g. Kennedy et al., 2003). Villareal’s et al.(2004) findings extend this concept to the ericoidmycorrhizal understorey of the boreal forest. Thepotential of ECM fungi to form multiple and diverseassociations with different groups of overstorey andunderstorey hosts means that the common mycorrhizalnetwork in forests may be more inclusive thanpreviously thought (see below and Leake this volume).

Some groups or genera of ECM fungi appear to havebecome specialised to particular vegetation types.Cortinarius, the most species rich ECM genus, is prolificboth in terms of species-richness and sporocarpproduction in northern boreal regions (e.g. Brandrud,1995), but is conspicuously absent from tropicalregions (but see Peintner et al., 2003). By contrast,members of the Russulaceae (mainly Russula andLactarius species), while not uncommon in temperateand boreal regions, present a huge, largely undescribed,diversity in tropical regions where ECM hosts occur(South America – Henkel et al., 2002; Africa – Buyck etal., 1996; SE Asia – Lee et al., 2003). It is not clear whythis pattern exists. It may reflect historical hostbiogeography, or perhaps relate to the attributes ofhydrophobic (Cortinarius) versus hydrophilic(Russulaceae) mantles in different edaphic conditions(e.g. form and periodicity of nutrient fluxes, intensity ofbelow herbivory and disturbance).

The total number of ECM fungi is very unclear. Themost recent estimate (Molina et al., 1992) suggestedthat there were ca. 5500 species. This is likely to be aconsiderable underestimate. In recent years, moreintensive mycological explorations of tropical forests(e.g. Haug et al., 2005; Buyck et al., 1996), and of thehypogeous fungi associated with the eucalyptusvegetation in Australasia (Claridge, 2002), have revealedmany undescribed ECM. Since the early 1990s, the useof molecular markers to identify mycobionts directlyfrom ectomycorrhizas has greatly increased thenumber of known taxa (e.g. Weiss et al., 2004). Inaddition, a number of fungal groups previouslyconsidered to be saprotrophic have been found to beECM (e.g. tomentelloid fungi, Kõljalg et al., 2000). Insummary, an accurate estimate of the size of the globalcommunity of ECM fungi may not be known for sometime but it is likely to be ca. 7000 – 10,000 species.

The Phytobionts

In comparison to the huge diversity of plants formingarbuscular mycorrhizas, the number of host species

e

h

m

sd

s

2a 2b

4

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Fig 1 Transverse section through ectomycorrhizal tips of Didelotia africana . e - extramatrical mycelium, m – mantle, h- Hartignet. Fig 2 Ectomycorrhizal tips formed by a) Russula ochroleuca on Picea abies and b) Cortinarius cinnamomeus on Picea abies. Fig3 Sporocarps formed by the ectomycorrhizal fungus Amanita regalis. Fig 4 Sporocarps formed by the ectomycorrhizal fungusCortinarius agathosmus. Fig 5 Sporocarps of ectomycorrhizal fungi (a. Lactarius lanceolatus & b. Russula nana) growing in shortturf vegetation where the host plants (d – Dryas, s – Salix) are inconspicuous components of the ground vegetation. Fig 7 Densecovering of cystidia and setae on the mantle of an ectomycorrhiza formed by a Tomentella sp.


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ancestor with Dipterocarpaceae, has been found to beectomycorrhizal (Ducousso et al., 2004), suggestingthat the ECM habit in this clade was present more than88 million years ago. By the same line of reasoning, theconfirmation that Pakaraimea, the S American genus, isalso ectomycorrhizal sets ECM origin in the clade backto 130million years ago, i.e. prior to the separation of S America from Africa. The taxonomy and ecology of ECM fungi of dipterocarps is largely unexplored, yet they must have much to tell us about the evolution of the ECM habit and the ecology of tropicalforest ecosystems.

Ectomycorrhizas are also the norm in certaincaesalpinioid legume trees, which are importantcomponents of tropical rain forest and savannahwoodlands in Africa. The host genera in question (allnon-nitrogen fixing), are largely confined to the tribesDetarieae and, especially, Amherstieae in theCaesalpinioideae (Alexander, 1989). Genera such asTetraberlinia, Microberlinia and Julbernardia form grovesor extensive monodominant stands in the rain forests ofthe Guineo-Congolian basin (Newbery et al., 1988),while others such as Isoberlinia and Brachystegia covervast areas in the Zambezian miombo woodlands of Eastand South-Central Africa, and the Sudania savannahwoodlands of the sub-Sahara. These ecosystemssupport a rich ECM basidiomycete flora (Munyanziza &Oldeman, 1996; Verbeken & Buyck, 2002). One of thisgroup of ectomycorrhizal legumes (Intsia) extends intothe dipterocarp forests of SE Asia, while another(Dicymbe) forms monodominant stands on poor soilsthroughout the Guyana shield of S. America. This ‘S.American connection’ again raises tantalisingquestions about the mycorrhizal status of hosts, andthe radiation of ECM fungal clades at the breakup ofGondwanaland. That said, the penetration of ECM hostgenera into the neotropical rain forest of S. Americaseems much more restricted than that into thepalaeotropics of Africa and SE Asia. Elsewhere,ectomycorrhizas are also found on a few non-caesalpinioid legumes in Australasia, including severalAcacia spp. (Alexander, 1989)

Two temperate ECM host families, Fagaceae andPinaceae, have radiated into natural tropical forests.Tropical oaks (e.g. Lithocarpus, Castanopsis,Trigonobalanus) are found in Central America, andthroughout SE Asia where they co-exist with ECMdipterocarps. Southern beech (Nothofagus), althoughcentred on Chile and New Zealand, is found as far northas Papua New Guinea. Endemic ECM tropical pines arefound in Mexico and Central America (e.g. Pinusoocarpa, P. patula), in the Caribbean (e.g. P. caribea), andin SE Asia (e.g. P. khesiya, P. merkusii) as far as 2°S in

Sumatra. The ECM fungal flora of these natural tropicalpine forests is not well studied. However, several tropicalpines have been extensively used in plantationsthroughout the tropics outside their natural range, andoften in areas that have not previously supportedectomycorrhizal vegetation. Ironically, in theseplantations the mycobionts are either temperate speciesintroduced with inoculum from Europe, or theubiquitous ECM genus Pisolithus (Read, 1998).Eucalyptus spp are widespread throughout temperateand subtropical forest ecosystems in Australia wherethey support a bewildering diversity of, especiallyhypogeous, ectomycorrhizal fungi (Claridge, 2002).Like pines, this genus has been extensively plantedoutside its natural range in tropical countries, and likepines its mycobiont community appears depauperate inthese situations. Acacia mangium is another Australianspecies extensively used in tropical plantations.Apparently it too can form ectomycorrhizas, at leastwith Pisolithus sp. (Duponnois & Ba, 1999).

Facultative ectomycorrhizal fungi – fact or fiction

It has been claimed that certain ECM fungal species(e.g. some Hebeloma spp, Marmeisse et al., 1999) arefacultative symbionts i.e. they live as ectomycorrhizalsymbionts but then, when the opportunity or needarises, can also grow as saprotrophs. Most of theseclaims are based upon the observation of sporocarps ofECM fungi growing many metres away from the nearesthost plant or growing in vegetation apparently devoidof plants capable of forming ectomycorrhizas. However,the roots of many trees can easily extend up to 50metres from the stem base (Stone & Kalisz (1991); theroots are not restricted, as often suggested, to the extentof the crown (Smith 1964). The roots of shelterbelttrees on farms where there is minimal soil disturbance,or of isolated trees in parklands, extend for manymetres into the surrounding grassland. This couldeasily explain the occurrence of sporocarps of e.g.Xerocomus and Laccaria spp. that regularly appear infields many metres from the woodland margin (A.Taylor, pers. obs.). This also raises the vexed question –with which particular host tree is a sporocarpassociated? With the exception of fungi known to bespecific on a particular host spp, or where only a singlepotential host is present, it is usually impossible toidentify the host with certainty solely on the basis ofwhere the sporocarp is located. Recording the nearesttree species as the host could be very misleading(particularly when the host in question formsarbuscular mycorrhizas and not ectomycorrhizas!). Inmixed woodland containing large trees, all potential


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hosts within at least 20 m should be recorded. Aninnovative recent study by Saari et al. (2005), in a Scotspine forest, used molecular markers to identify both theECM fungi on root tips and the tree that had producedthe tips. They not only showed how many ECM fungalspecies could associate with a single tree but also thatthe roots from single trees extended for, at least, 18mfrom the trunk.

Furthermore, as already mentioned the ECM habit isnot restricted to large, obvious host plants (i.e. trees). Anumber of shrub-forming genera, in particular thedwarf Salix spp., form ECM with a range of fungi andare often very inconspicuous components of short turfvegetation (Fig. 5). The appearance of ECM sporocarpsin this vegetation type could be very misleading. Arecent, detailed study carried out on the Burren, W.Ireland demonstrated that Cortinarius (Dermocybe)cinnamomeus formed ECM-like structures with Carexflacca and C. pilulifera (Harrington & Mitchell, 2002).Such careful studies might explain old reports of ECMspecies appearing in grasslands (Wilkins & Patrick,1939).

The production of sporocarps by some ECM fungalspecies (e.g. Laccaria spp., Kropp & Mueller, 1999) in theabsence of a host plant in the laboratory on carbon richmedia has also been suggested as evidence for free-living ECM fungi. However, it is difficult to envisage anysituation under natural conditions where ECM fungicould exploit pools of readily available carbohydrateslarge enough for them to complete their life cycleswithout intense competition from other soilmicroorganisms. Fast growing fungi such asTrichoderma spp would easily outcompete any ECMfungus.

In summary, there is no unequivocal evidence thatany ECM fungus can complete its life cycles in theabsence of a host. In this context it is important to makethe distinction between those ECM fungi that haveconsiderable ability to degrade organic substrates(Leake and Read, 1997) and true saprotrophic fungithat depend entirely upon the catabolism of organicmatter for their carbon. If facultative ECM fungi didexist they would be at a considerable advantage overspecies restricted to a single trophic status (either ECMor saprotrophic) and it is difficult to see why such multi-trophic species should not outcompete their lessversatile relations.

The production of sporocarps by ECM fungi hasbeen closely linked to the supply of currentphotosynthate to the root systems (Lamhamedi et al.,1994). This dependency was elegantly demonstrated ina recent study carried out in a boreal forest in N.Sweden, where the flow of photosynthate from foliage

to roots was terminated by girdling the trees at breastheight (i.e. severing the phloem tissue) (Högberg et al.,2001). This resulted in a rapid (within two days) end tosporocarp production. Girdling was also associatedwith a 56% reduction in soil respiration. Most of thereduction in CO2 efflux was assumed to be because ofthe dependence of ECM fungal respiration on currentassimilate. These field data confirm earlier laboratoryobservations by Söderström and Read (1987), whodemonstrated a 50% reduction in respiration aftersevering ECM mycelium in microcosms.

Nutrient storage by ECM fungi

Sporocarp production represents a large sink not onlyfor carbon (C) but also for mineral nutrients. Thesporocarps of many ECM species are enriched innutrients, particularly nitrogen (N) and phosphorus(P), relative to the soil and the host plants (Vogt &Edmonds, 1980; Vogt et al., 1981, 1991). How do ECMfungi satisfy the nutrient demand of sporocarpproduction? It seems very unlikely given the highlycompetitive nature of nutrient acquisition that theycan rapidly acquire the necessary nutrients from thesurrounding soil. It is more likely that they mobilisestores of nutrients from within their mycelium. Manywood-inhabiting saprotrophic fungi producesporocarps without a net increase in fungal dry mass,meaning that they mobilise existing reserves within themycelium (Rayner & Boddy, 1988), much of which isprotected within the decomposing woody substrate.However, for ECM fungi this poses a problem – wherecan nutrients be safely stored away from grazingfungivores? Any fungal tissue accessible to fungivores isat risk of being consumed. For ECM fungi, grazingwould be particularly damaging where rhizomorphsare severed and large sectors of the mycelium becomedetached. The mycelium itself may therefore not be anefficient repository for nutrients. The storage potentialof the fungal mantle was recognised early in ECMresearch (Harley et al., 1958; Morrison, 1962). Anumber of studies have demonstrated seasonalchanges in the P and N contents of mantles ofectomycorrhizas and it has been suggested thatfluctuations were linked to the seasonal patterns ofnutrient demand by the host (e.g. Lussenhop & Fogel,1999; Genet et al., 2000)..

If we assume that the main store of N is the mantle,then it is possible to calculate roughly how manymycorrhizas would be required to provide the N for asingle sporocarp. Taking Boletus edulis as an example, asingle sporocarp can weigh ca. 11.5 g dw and contain6.5% N – i.e. 0.745g of N (pers. obs.; Taylor et al.,


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2003). A single ectomycorrhizal tip weighs ca. 30 µgdw (pers. obs.) of which the mantle is 10-40% (Harleyand McCready, 1952; Vogt et al., 1981; 1991). Högberget al. (1996) measured ca. 3.5% N in mantles ofunidentified ectomycorrhizas. Using this to calculatethe amount of N in a single ectomycorrhiza using the10-40% estimate of mantle weight gives a range of0.105 – 0.420 µg N. So 1.77 - 7.09 x 106 ectomycorrhizascontain the same amount of N as one sporocarp. Thiscomparison does not include N within the Hartig net:more importantly, it assumes that all of the N in the tipscould be mobilised for sporocarp formation. However,much of the N in fungal mycelia is in the chitin of thecell wall (Kerley & Read 1997), and therefore unavailablefor remobilisation. If we assume that only 50% of thefungal N in ectomycorrhizas is mobile, then theestimate doubles to 3.54 – 14.18 x 106 mycorrhizal roottips. How does this relate to the density of ectomycorrhizasin forests? Our studies of ectomycorrhizal communitiesin the boreal forest usually find 0.25 – 6 x 106

mycorrhizal root tips per square metre of forest floor(pers. obs.). So the N from all of the ectomycorrhizas in1-2 square metres of forest floor would be required toproduce a single sporocarp of B. edulis.

If we assume that the main site of storage is themycelium then it is also possible to estimate the amountof mycelium that would have to contribute N for theformation of a sporocarp. The N content of soilmycelium (ca. 1.5%) is lower than that in mantles andsporocarps (Vogt et al., 1981). Therefore, 50 g of soilmycelium would contain the same amount of N as asingle B. edulis sporocarp. How many metres ofmycelium would this be? The specific density of soilmycelium has been estimated at 1.1 g cm3 (Saito, 1955in Bååth & Söderström, 1979) and we can work out,using a hyphal diameter of 5 µm, that in 1 cm3 ofmycelium, there are 4,000m (4km) of hyphae.Therefore, 50 g of soil hyphae is the equivalent of181.81 km hyphae. The amount of mobile N in themycelium would probably be less than that in mantlesas the majority of the mycelium is vacuolated(Söderström, 1979). Assuming only 10-50% of the N isavailable, this gives an estimate of mycelial length of363-1818 km. The latter figure is the length ofSweden! Estimates of hyphal length in soils varyconsiderably with forest and soil type, and season(Bååth & Söderström, 1979). However, our estimatesare within the same range as those reported from 1-2square metres of some forest soils (Söderström, 1979).

The similarity between our estimates of the area offorest containing either sufficient mycorrhizal tips ormycelium to supply the N in a single sporocarp iscoincidental. It is very rare to find ECM communities

where all of the tips within a metre area are colonisedby a single ECM species. Usually a large number ofspecies coexist within even small (cm3) areas. Inaddition, estimates of mycelial length in soil, such asthat used above, include hyphae from mycorrhizal andsaprotrophic taxa. This would mean that we havesubstantially underestimated the amount of ECMhyphae necessary to supply N. Therefore, assumingthat ECM fungi are incapable of rapidly acquiring alarge pool of nutrients from the soil, these simplecalculations highlight an intriguing problem: fromwhere do ECM fungi mobilise the N and P (and othernutrients) necessary for sporocarp formation?

Physical and chemical defences

The earliest known fossil evidence of ECM structuresdates from ca. 50 million years (LePage et al., 1997),but molecular data suggests that their origin may bemuch earlier at ca. 180 million years (Berbee & Taylor,2001). Despite this antiquity, the ECM habit evolved insoils where there were already diverse communities ofsoil mites (Walter & Proctor, 1999); at least 11 speciesof mites are present in the Rhynie Chert, which datesfrom 380-400 million years (Shear & Kukalova-Peck,1990). Some of the modern descendent of these earlymites are fungivores and it seems possible thatfungivory might have been a major selection pressureon the evolution of ECM. Unlike arbuscularmycorrhizas, where essential fungal structures areprotected within the root, and where hyphal healingseems widespread (Giovannetti et al., 1999), in the ECMsymbiosis the mantle is exposed on the root surface, andhyphal healing has not been reported.

The mantle, which may be 20-60 µm thick (Fig. 1),is a storage site for nutrients acquired by the fungusfrom the soil (Smith & Read, 1997). Concentrations ofN and P in fungal tissue are 4 to 5 times higher thanthat of plant material (Vogt et al., 1981). Given thediffuse nature of most of the mycelium in the soil, themantle represents, to a fungivore, a small patch of highquality resource. In addition, in relation to the rest ofthe mycelium, the mantle is a relatively long-livedstructure. Estimates of the lifespan of individualmycorrhizal tips range from a few months to two years(Downes et al., 1992). There are no good estimates forthe turnover of ECM mycelia, but it is generallyassumed that it is more rapid than for mycorrhizal tips.

Many ECM fungi develop specialised cells on thesurface of the mantle (Fig. 6). These are usuallydifferentiated end cells of mantle hyphae that,particularly when the mycorrhizas are newly formed,may densely cover the whole surface of the mantle (Fig.


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7). It is possible to distinguish two types of cells basedon potential mode of action in defence. Thick-walled,pointed structures, which are often melanised, mayform a physical barrier to fungivory. There has been nodirect examination of the potential effects on grazing ofthe thick-walled, setae-like structures that many ECMfungi produce on their mantles, but it is tempting tothink that they do have a role to play. We know moreabout the thin-walled, swollen cells (cystidia) that arealso found on mantle surfaces. These have specialisedcellular contents and may function as reservoirs ofchemical deterrents. This latter group of cells areparticularly prevalent on mantles formed by membersof the Russulaceae (Agerer, 1986-2002; Eberhardt,2002). Even when no specialised cells are present, thechemical deterrents are still present within somesurface mantle cells or within lactifers or laticiferoushyphae that ramify through the mantle tissue. Thecells, both specialised and otherwise, contain thebiologically inactive precursor stearoylvelutinal that,upon injury to the cell, is converted in seconds tostrongly antibiotic and pungent sesquiterpenoiddialdehydes such as isovellerral (Mier et al., 1996).

Fig 6 The range of specialised cells observed on the mantlesurfaces of ectomycorrhizas (from R. Agerer, with permis-sion)

These substances have been shown to be active againsta range of potential arthropod grazers (Mier et al.,1996; Stadler and Sterner, 1998). The release of thesecompounds is seen on a large scale when sporocarps ofLactarius ‘milk’ in response to damage. In fact, the samemilking reaction can be seen from the mantles andrhizomorphs of Lactarius mycorrhizas.

The occurrence and appearance of cystidia andsetae on sporocarp surfaces has been extensively usedin fungal taxonomy (Largent et al., 1977). While adefence role can be envisaged for these structures in thedeveloping hymenium, their sparse presence in capcuticles is more difficult to explain. But we are perhapstrying to confer functions for these features at thewrong scale. Small ECM fungal structures (mantles,rhizomorphs, hyphae and sporocarp primordia) are allsubject to grazing pressure from microarthropods inthe soil. Small-scale defence mechanisms such as thoseoutlined above may be effective deterrents against smallgrazers. However, once sporocarps expand, slugs(Richter, 1980) and a diverse range of vertebrates (e.g.deer, Avila et al., 1999) become the major consumers.These large fungivores are unlikely to notice small-scalephysical defence mechanisms. The occurrence ofcystidia on large sporocarps may be coincidental – aconsequence of the need for protection at a small scale.

Many ECM species also produce ornamentedhyphae that are coated with crystals of calcium oxalateor other crystalline deposits, which could also providedefence against grazing (Brand, 1991). The ECMfungus Piloderma fallax (croceum) forms very conspicuousfans of bright yellow mycelium in boreal forests. Twofactors contribute to the apparent abundance of thisfungus. Firstly, the bright yellow mycelium makes itobvious and secondly, the mycelium appears to persistafter it is dead. An explanation for this persistence isthat the hyphae of P. fallax are coated with numerouscrystals, sometimes to the extent where the hyphalwalls are no longer visible beneath the deposits. It wouldappear that few soil organisms find the mycelium of P.fallax palatable. It is also common to find mycorrhizaltips formed by P. fallax that superficially appear viablebut a closer inspection of the tip reveals that the rootwithin the mantle is moribund and decayed. This is alsoa common feature of another ECM fungus, Cenococcumgeophilum, that forms black, heavily melanised mantlesthat also appear very unpalatable to fungivores.

Conclusions

We know that the ectomycorrhizal symbiosis iswidespread and an important component of manyecosystems. We also know a considerable amount


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about the taxonomy of the organisms involved. But,while the diversity of ECM fungi is impresive, it isimportant to place this species richness in relation tothe huge number of other soil inhabitating organisms.Many of these influence ECM fungi either directly viagrazing (fungivores) or indirectly via competition fornutrients (e.g. other microorganisms). We have muchto learn how the interactions between ECM fungi andorganisms at other trophic levels determine the activityand success of ECM fungi and their host plants.

ReferencesAgerer, R. (1987-2002). Colour Atlas of Ectomycorrhizae. 1st-

12th delivery. Einhorn Verlag, Schwäbisch Gmünd. Agerer, R. (2001). Exploration types of ectomycorrhizae.

A proposal to classify ectomycorrhizal mycelial systems according to their patterns of differentiation and putative ecological importance. Mycorrhiza 11: 107-114.

Agerer, R., Danielson, R. M., Egli S., Ingleby, K., Luoma D. &Treu R. (1996 – 2004). Descriptions of Ectomycorrhizae.Schwäbisch-Gmünd: Einhorn-Verlag.

Alexander, I. J. (1989). Systematics and ecology ofectomycorrhizal legumes. Advances in Legume Biology,Monographs in Systematic Botany 29, 607-624.

Allen, T. R., Millar, T., Berch, S. M. & Berbee, M. L. (2003).Culturing and direct DNA extraction find different fungifrom the same ericoid mycorrhizal roots. New Phytologist160: 255-272.

Ashford, A. E., Peterson, C. A., Carpenter, J. L., Cairney, J. W.G. & Allaway, W. G. (1988). Structure and permeability ofthe fungal sheath in the Pisonia mycorrhiza. Protoplasma147: 149-161.

Ashton, P. S. (1982). Dipterocarpaceae. In: Flora Malesiana,Series 1, Spermatophyta (edited by C. G. G. J. Van Steenis), pp.237-552. The Hague: Martinus-Niijhoff Publications.

Avila, R., Johanson, K. J. & Bergström, R. (1999). Model of theseasonal variations of fungi ingestion and 137Cs activityconcentrations in roe deer. Journal of EnvironmentalRadioactivity 46: 99-112.

Berbee, M. L. & Taylor, J. W. (2001). Fungal molecular evolution: gene trees and geologic time. In The Mycota VII.Part B. Systematics and Evolution. (edited by D. J.McLaughlin, R. G. McLaughlin, P. A. Lemke) pp. 229-245.Berlin: Springer-Verlag.

Brand, F. (1991). Ektomykorrhizen an Fagus sylvatica.Charakterisierung und Identifizierung, ökologischeKennzeichnung und unsterile Kultivierung. Libri Botanici,vol. 2, IHW-Verlag, Eching, Germany.

Brandrud, T. E (1995). The effects of experimental nitrogenaddition on the ectomycorrhizal fungus flora in an oligotrophic spruce forest at Gårdjön, Sweden. ForestEcology and Management 71: 111-122.

Buyck, B., Thoen, D & Watling, R. (1996). Ectomycorrhizalfungi of the Guinea Congo region. Proceedings of the RoyalSociety of Edinburgh 104B: 313-333.

Bååth, E. & Söderström, B. (1979). Fungal biomass and fungal immobilization of plant nutrients in Swedish coniferous forest soils. Revue D’Ecologie et de Biologie du Sol16: 477-489.

Claridge, A. W. (2002). Ecological role of hypogeous ectomycorrhizal fungi in Australia forests and woodlands.Plant and Soil 244: 291-305.

Dickie I. A., Xu, B. & Koide, R. T. (2003). Vertical niche differentiation of ectomycorrhizal hyphae in soil as shownby T-RFLP analysis. New Phytologist 156: 527-535.

Downes, G., Alexander, I. J. & Cairney, J. G. (1992). A light andelectron microscope study of ageing and death of spruceectomycorrhizas. New Phytologist 122: 141-152.

Ducousso, M., Bena, G., Bourgeois, C., Buyck, B., Eyssartier,G., Vincelette, M., Rabevohitra, R., Randrihasipara, L.,Dreyfus, B., & Prin, Y. (2004). The last common ancestor of Sarcolaenaceae and Asian dipterocarp treeswas ectomycorrhizal before the India-Madagascar separation, about 88 million years ago. Molecular Ecology13: 231-236.

Duponnois, R. & Ba, A. M. (1999). Growth stimulation ofAcacia mangium Willd by Pisolithus sp. in some Senegalesesoils. Forest Ecology and Management 119: 209-215.

Eberhardt, U. (2002). Molecular kinship analyses of theagaricoid Russulaceae: correspondence with mycorrhizalanatomy and sporocarp features in the genus Russula.Mycological Progress 1: 201-223.

Faber, J. H. (1991). Functional classification of soil fauna: anew approach. Oikos 61: 110-117.

Fitter, A. H. & Moyersoen, B. (1996). Evolutionary trends inroot-microbe symbioses. Philosophical Transactions of theRoyal Society of London. Series B-Biological Sciences 351:1367-1375.

Gardes, M. & Bruns, T. D. (1996). Community structure ofectomycorrhizal fungi in a Pinus muricata forest: above- andbelow-ground views. Canadian Journal of Botany 74: 1572-1583.

Giovannetti, M., Azzolini, D. & Citernesi, A. S. (1999).Anastomosis formation and nuclear and protoplasmicexchange in arbuscular mycorrhizal fungi Applied andEnvironmental Microbiology 65: 5571-5575.

Guidot, A., Debaud, J.-C., Effosse, A. & Marmeisse, R. (2003).Below-ground distribution and persistence of an ectomycorrhizal fungus. New Phytologist 161: 539-547.

Harley, J. L. & McCready, C. C. (1952). The uptake ofphosphorus by excised mycorrhizal roots of beech. III. Theeffect of the fungal sheath on the availability of phosphateto the core. New Phytologist 51: 343-348.

Harley, J. L., McCready, C. C. & Brierley, J. K. (1958). Theuptake of phosphorus by excised mycorrhizal roots ofbeech. VII. Translocation of phosphorus in mycorrhizalroots. New Phytologist 57: 353-362.

Harrington, T. J. & Mitchell, D. T. (2002). Colonisation of rootsystems of Carex flacca and C. pilulifera by Cortinarius(Dermocybe) cinnamomeus. Mycological Research 106: 452-459.

Haug, I., Weiss M., Homeier, J., Oberwinkler, F. & Kottke, I.(2005). Russulaceae and Thelephoraceae form ectomycorrhizas with members of the Nyctaginaceae(Caryophyllales) in the tropical mountain rain forest ofsouthern Ecuador. New Phytologist 165: 923-936.

Henkel, T. W., Terborgh, J. & Vilgalys, R. J. (2002).Ectomycorrhizal fungi and their leguminous hosts in thePakaraima Mountains of Guyana. Mycological Research106: 515-531.

Hibbett, D. S., Gilbert, L.-B. & Donohugue, M. J. (2000).Evolutionary instability of ectomycorrhizal symbioses inbasidiomycetes. Nature 407: 506-508

Horton, T., Bruns, T. D. & Parker, V. T. (1999). Ectomycorrhizalfungi associated with Arctostaphylos contribute toPseudotsuga menziesii establishment. Canadian Journal ofBotany 77: 93-102.


111

Högberg, P., Högbom, L., Schinkel, H., Högberg, M.,Johannisson, C., & Wallmark, H. (1996). 15N abundance ofsurface soils, roots and mycorrhizas in profiles of Europeanforest soils. Oecologia 108: 207-214.

Högberg, P., Nordgren, A., Buchmann, N., Taylor, A. F. S.,Ekblad, A., Högberg, M. N., Nyberg, G., Ottosson-Löfvenius,M. & Read, D. J. (2001). Large-scale forest girdling experiment demonstrates that current photosynthesis drives soil respiration. Nature 411: 789-792.

Kennedy, P. G., Izzo, A. D. & Bruns, T. D. (2003). There is highpotential for the formation of common mycorrhizal networks between understorey and canopy trees in a mixedevergreen forest. Journal of Ecology 91: 1071-1080.

Kerley, S. J. & Read, D. J. (1997). The biology of mycorrhiza inthe Ericaceae: XIX. Fungal mycelium as a nitrogen sourcefor the ericoid mycorrhizal fungus Hymenoscyphus ericaeand its host plants. New Phytologist 136, 691-701.

Koide, R., T., Xu, B., Sharda, J., Lekberg, Y. & Ostiguy, N.(2004). Evidence of species interactions within an ectomy-corrhizal fungal community. New Phytologist (on line).

Koide, R. T., Xu, B. & Sharda, J. (2005). Contrasting below-ground views of an ectomycorrhizal fungal community.New Phytologist (early on-line).

Kottke, I., Beiter, A., Weiss, M., Haug, I., Oberwinkler, F. &Nebel, M. (2003). Heterobasidiomycetes form symbioticassociations with hepatics: Jungermanniales have sebaci-noid mycobionts while Aneura pinguis (Metzgeriales) is asso-ciated with a Tulasnella species. Mycological Research 107:957-968.

Kretzler, A. M. & Bruns, T. D. (1999). Use of atp6 in fungalphylogenetics: an example from the Boletales. MolecularPhylogenetics and Evolution 13: 483-492.

Kropp, B. R. & Mueller, G. M. (1999). Laccaria. InEctomycorrhizal Fungi: Key genera in profile (edited by J.W.G.Cairney & S.M. Chambers), pp.65-88. Berlin: Springer.

Kõljalg, U., Dahlberg, A., Taylor, A. F. S., Larsson, E.,Hallenberg, N., Stenlid, J., Larsson, K. H., Fransson, P. M.,Kårén, O. & Jonsson., L. (2000). Diversity and abundance ofresupinate thelephoroid fungi as ectomycorrhizal sym-bionts in Swedish boreal forests. Molecular Ecology 9: 1985-1996.

Lamhamedi, M. S., Godbout, C. & Fortin, J. A. (1994).Dependence of Laccaria bicolor basidiomes development oncurrent photosynthesis of Pinus strobus seedlings. CanadianJournal of Forest Research 14: 412-415.

Landeweert, R., Leeflang., P., Kuyper, T. W., Hoffland E.,Rosling, A., Wernars, K. & Smit, E. (2003). Molecular iden-tification of ectomycorrhizal mycelium in soil horizons.Applied and Environmental Microbiology 69: 327-333.

Largent, D., Johnson, D. & Watling, R. (1977). How to identifymushrooms to genus III: Microscopic features. Eureka, CA:Mad River Press.

Larsson, K.-H., Larsson, E. & Kõljalg, U. (2004). High phylo-genetic diversity among corticoid homobasidiomycetes.Mycological Research 108: 983-1002.

Leake, J. R. & Read, D. J. (1997). Mycorrhizal fungi in terees-trial ecosystems. In: Wicklow D, Söderström B, eds. TheMycota IV environmental and microbial relationships. Berlin,Germany: Springer-Verlag, 281-301.

Lee, S. S., Watling, R. & Turnbull, E. (2003). Diversity of puta-tive ectomycorrhizal fungi in Pasoh Forest Reserve. Pasoh:Ecology of a lowland rain forest in Southeast Asia (eds T.Okuda, N. Manokaran, Y.Matsumoto, K. Niiyama, S. C.Thomas & P. S. Ashton), pp 149-159. Springer, Tokyo.

LePage, B. A., Currah, R., Stockey, R. & Rothwell, G. W.(1997). Fossil ectomycorrhiza in Eocene Pinus roots.American Journal of Botany 84: 410-412.

Lussenhop, J. & Fogel, R. (1999). Seasonal change in phosphorus content of Pinus strobus-Cenococcum geophilumectomycorrhizae. Mycologia 91: 742-746.

Marmeisse, R., Gryta, H., Jargeat, P., Fraissinet-Tachet, L.,Gay, G. & Debaud, J.-C. (1999). In Ectomycorrhizal Fungi:Key genera in profile (edited by J. W. G. Cairney & S. M.Chambers), pp.89-127. Berlin: Springer.

Massicotte, H. B., Melville, L. H., Peterson, R. L. & Luoma, D. L. (1998). Anatomical aspects of field ectomycorrhizas on Polygonum viviparum (Polygonaceae)and Kobresia bellardii (Cyperaceae). Mycorrhiza 7: 287-292.

McGonigle, T. P. (1995). The significance of grazing on fungiin nutrient cycling. Canadian Journal of Botany 73(Suppl.1):S1370-S1376.

Meyer, F. H. (1973). Distribution of ectomycorrhizae in nativeand man-made forests. In: Ectomycorrhizae (eds G. C.Marks, and T. T. Kozlowski). Academic Press, New York,USA. Pp. 79-105.

Mier, N., Canete, S., Klaebe, A., Chavant, L. & Fournier, D.(1996). Insecticidal properties of mushroom and toadstoolcarpophores. Phytochemistry 41: 1293-1299.

Molina, R., Massicotte, H. & Trappe, J. M. (1992). Specificityphenomenon in mycorrhizal symbiosis: community-ecological consequences and practical implications. In:Mycorrhizal Functioning (ed. M. F. Allen). Chapman andHall, London, UK. pp. 357-423.

Morrison, J. B. (1962). Absorption of phosphorus from soilsby mycorrhizal plants. New Phytologist 61: 10-20.

Munyanziza, E. & Oldeman, R. A. A. (1996). Miombo trees:Ecological strategies, silviculture and management. Ambio7: 454-458.

Murat, C., Díez, J., Luis, P., Delaruelle, C., Dupre, C., Chevalier,G., Bonfante, P. & Martin, F. (2004). Polymorphism at theribosomal DNA ITS and its relation to postglacial re-colo-nization routes of the Perigord truffle Tuber melanosporum.New Phytologist 164: 401-411.

Newbery, D. Mc., Alexander, I. J., Thomas, D. W. & Gartlan, J.S. (1988). Ectomycorrhizal legumes and soil phosphorus inKorup National Park, Cameroon. New Phytologist 109,433-450.

Peintner, U., Moser, M.M., Thomas, K.A. & Manimohan, P.(2003). First records of ectomycorrhizal Cortinariusspecies (Agaricales, Basidiomycetes) from tropical Indiaand their phylogenetic position based on rDNA ITSsequences. Mycological Research 107: 485-494.

Rayner, A. D. M. & Boddy, L. (1988). Fungal decomposition ofwood. Its biology and ecology. John Wiley & Sons. Chichester,New York. pp 587.

Read, D. J. (1998). The mycorrhizal status of Pinus. In Ecology& Biogeography of Pinus. Ed by D M Richardson. pp 324-340. Cambridge University Press.

Read, D. J. & Perez-Moreno, J. (2003). Mycorrhizas and nutri-ent cycling in ecosystems – a journey towards relevance?New Phytologist 157: 465-492.

Richter, K. O. (1980). Evolutionary aspects of mycophagy inAriolimax columbianus and other slugs. In Proceedings of theSeventh International Soil Zoology Colloquium of theInternational Society of Soil Science, (ed. D. L. Dindal). pp616-636. The Office of Pesticides and Toxic substances,E.P.A., Washington D.C.


112

Shaw, P. J. A. (1988). A consistent hierarchy in the fungalfeeding preferences of the collembola Onychiurus armatus.Pedobiologia 31: 179-187.

Shaw, P. J. A. (1992). Fungi, Fungivores, and Fungal FoodWebs. In. The fungal community. Its organisation and role inthe ecosystem. (eds. G. C. Carroll & D. T. Wicklow). MarcelDekker, Inc. New York, Basel, Hong Kong. pp. 295-310.

Shear, W. A. & Kukalová-Peck, J. (1990). The ecology ofPalaeozoic terrestrial arthropods: the fossil evidence.Canadian Journal of Zoology 68: 1807-1834

Smith, J. H. G. (1964). Root spread can be estimated fromcrown width of Douglas fir, Lodgepole pine, and otherBritish Columbia tree species. Forestry Chronicle 456-473.

Smith, S. E. & Read, D. J. (1997). Mycorrhizal Symbiosis.Academic Press, Harcourt Brace & Company, Publishers,San Diego. 605 pp.

Stadler, M. & Sterner, O. (1998). Production of bioactive secondary metabolites in the fruit bodies of macrofungi asa response to injury. Phytochemistry 49: 1013-1019.

Stone, E. L. & Kalisz, P. J. (1991). On the maximum extent oftree roots. Forest Ecology and Management. 46: 59-102.

Sutara, J. (1992). The genera Paxillus and Tapinella in CentralEurope. Ceska Mykologia 46: 50-56.

Söderström, B. (1979). Seasonal fluctuations of active fungalbiomass in the horizons of a podzolized pine forest soil incentral Sweden. Soil Biology and Biochemistry 11: 149-154.

Söderström, B. & Read, D. J. (1987). Respiratory activity ofintact and excised ectomycorrhizal mycelial systems growing in unsterilised soil. Soil Biology and Biochemistry19: 231-236

Taylor, A. F. S., Martin, F. & Read D. J. (2000). Fungal diversity in ectomycorrhizal communities of Norwayspruce (Picea abies [L.] Karst.) and beech (Fagus sylvatica L.)along north-south transects in Europe. In: Schulze E-D, ed.Carbon and Nitrogen Cycling in European Forest Ecosystems.Berlin Heidelberg, German: Springer Verlag, pp 343-365.

Taylor, A. F. S., Fransson, P. M., Högberg, P., Högberg, M. N. &Plamboeck A.-H. (2003). Species level patterns in 13C and15N abundance of ectomycorrhizal and saprotrophic fungalsporocarps. New Phytologist 159: 757-774.

Taylor, D. L., Bruns, T. D., Szaro, T. M. & Hodges, S. A. (2003).Divergence in mycorrhizal specialization within Hexalectrisspicata (Orchidaceae), a nonphotosynthetic desert orchid.American Journal of Botany 90: 1168-1179.

Urban, A., Weiss, M. & Bauer, R. (2003). Ectomycorrhizaeinvolving sebacinoid fungi and their analogs. Journal of theLinnaean Society of London, Botany 13: 31-42.

Van der Heijden, M. G. A. & Sanders, I. R. (2002). MycorrhizalEcology. Ecological Studies 157. Springer-Verlag BerlinHeidelberg New York. pp.469

Verbeken, A. & Buyck, B. (2002) Diversity and ecology oftropical ectomycorrhizal fungi in Africa. In TropicalMycology. Vol.1 Macromycetes (Edited by R. Watling, J.C.Frankland, M. Ainsworth, S. Isaac & C.H. Robinson) pp. 11-24. Wallingford, UK: CABI Publishing.

Villarreal-Ruiz, L., Anderson, I. C. & Alexander I. J. (2004).Interaction between an isolate from the Hymenoscyphusericae aggregate and roots of Pinus and Vaccinium. NewPhytologist 164: 183-192.

Vogt, K. A, & Edmonds, R. L. (1980) Patterns of nutrient concentration in basidiocarps in western Washington.Canadian Journal of Botany 58: 694-698.

Vogt, K. A, Edmonds, R. L. & Grier, C. C. (1981). Biomass and nutrient concentrations of sporocarps produced by mycorrhizal and decomposer fungi in Abies amabilis stands.Oecologia 50: 170-175

Vogt, K. A., Publicover, D. A. & Vogt, D. J. (1991). A critique ofthe role of ectomycorrhizas in forest ecology. Agriculture,Ecosystems and Management 35: 171-190.

Vrålstad, T., Fossheim, T. & Schumacher, T. (2000). Piceirhizabicolorata – the ectomycorrhizal expression of theHymenoscyphus ericae aggregate? New Phytologist 145:549-563

Vrålstad, T., Schumacher, T. & Taylor, A. F. S. (2002).Mycorrhizal synthesis between fungal strains of theHymenoscyphus ericae aggregate and potential ectomycor-rhizal and ericoid hosts. New Phytologist 153: 143-152.

Walter, D. E. & Proctor, H. C. (1999). Mites. Ecology, Evolutionand Behaviour. CABI Publishing, CABI International,Wallingford Oxon, UK. pp. 322

Weiss, M., Selosse, M.-A., Rexer, K.-H., Urban, A. &Oberwinkler, F. (2004). Sebacinales: a hitherto overlookedcosm of heterobasidiomycetes with a broad mycorrhizalpotential. Mycological Research 108: 1003-1010.

Wilkins, W. H. & Patrick, S. H. M. (1939). The ecology oflarger fungi III. Constancy and frequency of grasslandspecies with special reference to soil types. Annals ofApplied Biology 26: 25-46.

the ectomycorrhizal symbiosis: life in the real world

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