Kumar 2017

Bot. Rev.
https://doi.org/10.1007/s12229-017-9196-z
REVIEW
Studies on Ectomycorrhiza: An Appraisal
Jitender Kumar 1 & N. S. Atri 1,2

1
Department of Botany, Punjabi University, Patiala -147002, India
2
Author for Correspondence; e-mail: narinderatri04@gmail.com
# The New York Botanical Garden 2017
Abstract Ectomycorrhizal (ECM) fungi are obligate symbionts of dominant vascular

plants, liverworts and hornworts. There are reports of about 20,000 to 25,000 ECM
fungi that promote plant growth by facilitating enhanced water and nutrient absorption,
and provide tolerance to environmental stresses. These below-ground fungi play a key
role in terrestrial ecosystems as they regulate plant diversity, nutrient and carbon cycles,
and influence soil structure and ecosystem multifunctionality. Because ECM fungi are
obligate root symbionts, host plant can have a strong effect on ECM species richness
and community composition. The biogeographic pattern and detailed functioning and
regulation of these mycorrhizosphere processes are still poorly understood and require
detailed study. More recent researches have placed emphasis on a wider, multifunc-
tional perspective, including the effects of ectomycorrhizal symbiosis on plant and
microbial communities, and on ecosystem processes. Over the years the main focus in
ECM research has been on the study of diversity and specificity of ECM strains, the
role of ECM in regeneration of degraded ecosystem, the growth and establishment of
seedlings through nutrient acquisition and the mediation of plant responses to various
types of stress. In this review, recent progresses in ectomycorrhizal biology are
presented, especially the potential role of ECM symbioses in resistance or tolerance
to various biotic and abiotic stresses, and in maintinance of plant diversity for proper
ecosystem functioning.
Keywords Ectomycorrhizal symbiosis . Ecosystem . Evolution . ECM diversity .

Afforestation
Introduction
It was Albert Bernard Frank (1885), a forest pathologist, who for the first time
introduced the term mycorrhiza. In Greek language Bmykes^ refers to Fungus and
Brhiza^ refers to Root. Since Frank’s description of mycorrhizal association in 1880’s
(Frank, 1885), a lot of work has been generated by different investigators as a
consequence of which it is estimated that 86% of terrestrial plant species are benefited
as they acquire their mineral nutrients via mycorrhizal roots (Brundrett, 2009). In fact
the mutualistic phenomenon of symbiosis has been reported to result from the co-
evolution between plants and fungi and became essential and obligatory for terrestrial
J. Kumar, N.S. Atri
plant nutrition (Allen, 2007; Alvez et al., 2010; Brundrett, 2002; Bücking & Heyser,
2003; Trappe, 1977).
Of the seven types of mycorrhizae described (arbuscular, arbutoid, ectomycorrhiza,
ectendo, ericoid, monotropoid, and orchidaceous), both arbuscular (AM) and
ectomycorrhizae (ECM) are reported to be the most abundant and widespread in forest
communities (Smith & Read, 2008; Taylor & Alexander, 2005). The number of ECM
fungal species is estimated between 20,000 - 25000, and the number of plants, mainly
trees and woody shrubs of tropical and temperate forests are estimated to 6000 species
(Brundrett, 2009; Rinaldi et al., 2008; Roy-Bolduc et al., 2016). Forest trees belonging
to Pinaceae, Fagaceae, Betulaceae, Nothophagaceae, Fabaceae, Gnetaceae,
Leptospermoidae of Myrtaceae, Dipterocarpaceae and Amhersteae of Casalpiniaceae,
which form dominant component in boreal, temperate and tropical rain forests, are
reported to harbour a great diversity of ECM fungi. ECM fungi that associates with
these ecologically and economically important trees appears to differ greatly in their
ecology and dispersal abilities and could be subject to different selective forces. Recent
studies suggest that typical biogeographic patterns and distance decay of similarity
resulting from dispersal limitation are usually evident in ECM fungi, but these patterns
do not always match those observed in plants and animals (Bahram et al., 2013;
Tedersoo et al., 2012). A number of liverworts and lycophytes are also reported to
form fungal associations that physically resemble ectomycorrhizas (Bidartondo et al.,
2003; Horn et al., 2013). The ectomycorrhiza-like associations of these liverwort-
fungus and Lycophytes-fungus associations are integral parts of terrestrial ecosystems,
but have rarely been studied.
In different forest ecosystems, ECM fungi have been reported to play an important
role in seedling survival, establishment and growth (Sebastiana et al., 2017; Smith &
Read, 2008; Tedersoo et al., 2010a). Studies have shown that the ECM fungi mostly
belong to higher Basidiomycetes, Ascomycetes and a very few Zygomycetes.
Researches have confirmed that ECM fungi play a key role in terrestrial ecosystems
as drivers of global carbon and nutrient cycles. ECM plant provides photosynthetically
fixed carbon and habitat for the fungi, while mycobionts provide dissolved and
organically bound nutrients mainly nitrogen (N) and phosphorus (P) to their hosts
(Simard et al., 1997; Simard & Durall, 2004; Tedersoo et al., 2010a). ECM symbiosis
differ from other mutualistic plant fungi interactions by the presence of a mantle,
formed by fungal colonisation of short feeder roots, and a Hartig net representing an
intercellular hyphal penetration between epidermal or cortical cells (Agerer, 1999;
Smith & Read, 2008; Grelet et al., 2009). Hartig net is the place of massive bidirec-
tional exchanges of nutrients between the fungus and host plant. ECM symbionts are
normally reported to colonise soils where nutrients are bound in organic compound.
The mycelium of ECM fungi promotes the release of complexed nutrients through the
excretion of extracellular enzymes and acids (Finlay, 2008).
Mycorrhizal taxonomy and molecular techniques have uncovered the unexpected
diversity and functioning of mycorrhizal associations and their temporal and spatial
dynamics in boreal, temperate, tropical and subtropical ecosystems. The development
of –omics (transcriptomics, metagenomics) have now revolutionised our knowledge of
ECM functioning, diversity and biogeograpy. At the ecosystem levels, more and more
clues have revealed the role of mycorrhizal fungi in evolution, plant growth, soil
structure and responses to environmental changes and global carbon and nutrient
cycles. The present review aims at summarizing our knowledge on ECM biology,
evolution, global diversity, specificity, ecology and potential role of their association in
ecosystem sustainance.
Evolutionary History of ECM:
The origin of ECM association is reported to be approximately 125 million year old.
Despite being widespread they are associated with only 3% of vascular plants families
(Smith & Read, 2008). The oldest ECM root fossil was reported to be Pinaceae which
was recorded about 50 million year ago (mya). In this regard Berbee and Taylor (1993)
has emphasized that ECM origin may date from the time of mushroom evolution (130
mya) on earth, although this view seems to lack evidence as the fossilised plant material
of Pinaceae, an ECM family known since the Triassic (200 mya; Hibbett et al., 1997),
suggests that ECM association may have appeared before mushroom forming fungi.
Molecular evidences suggest that ECM taxa, belonging to class Agaricomycetes and
some members of order Pezizales originated around 200 and 150 mya, respectively
(Berbee & Taylor, 2001, 2010). In this regard largest numbers of fungal lineages are
reported in mushrooms belonging to orders Pezizales, Agaricales, Helotiales, Boletales
and Cantherellales (Tedersoo et al., 2010a, 2012). This phylogenetic diversity shows
that ECM symbiosis has arisen several times independently (Floudas et al., 2012;
Hibbett et al., 2000; Tedersoo et al., 2010a, 2012). The examination of available
ECM fungi Laccaria bicolor and Tuber melanosporum genomes (Martin et al., 2008,
2010) and new genomic data documented by the Mycorrhizal Genomics Initiative
Consortium (MGI) concurs with the hypothesis that mycorrhizal symbiosis in nature
has originated with a loss of lignocellulose- degrading genes compared with
saprotrophic ancestors (Eastwood et al., 2011; Martin & Selosse, 2008; Plett &
Martin, 2011; van der Heijden et al., 2015; Wolfe et al., 2012). The loss of lignocel-
lulose degrading enzymes has been reported to be responsible for the dependence of
ECM fungi on their host plant for harvesting fixed carbon. Molecular clock analysis on
the reconciled tree suggested that ECM fungi evolved far latter than the appearance of
the last common ancestor of brown and white rot fungi about 300 mya (Marcel et al.,
2015). These results supported the long standing hypothesis that ECM fungi evolved
polyphyletically from multiple saprophytic species. The regular updates of the ECM
lineages and genera can be found at the UNITE homepage http://unite.ut.ee/EcM_
lineages (Abarenkov et al., 2010).
Diversity and Specificity of ECM
ECM are reported to be common in temperate and boreal ecosystems and in large
forested areas of tropical and subtropical regions (Corrales et al., 2016; Diédhiou et al.,
2014; Henkel et al., 2011; Sharma et al., 2009; Tedersoo et al., 2012). In the temperate
and boreal ecosystems most studies have been focused on ECM of Picea and Pinus. In
the BColour Atlas of Ectomycorrhiza^ by Agerer (1987-2012) most morphotypes
documented are from coniferous trees. A large number of fungal species, mostly
belonging to Basidiomycota and Ascomycota associate with conifers. Species of
J. Kumar, N.S. Atri
Amhinema, Boletus, Hebeloma, Laccaria, Paxillus, Phialophora, Russula, Lactarius,

Suillus and Thelephora are the commonest associates of conifer roots (Bent et al., 2011;
Gao & Yang, 2010; Garcia et al., 2016; Obase et al., 2009) and the Cenococcum
geophilum often dominate the community (Horton & Bruns, 1998; Koide et al., 2008;
Obase et al., 2009; Taniguchi et al., 2007). These fungi often coexist and for example
100 different ECM fungal species have been detected on the same tree species locally
(Roy-Bolduc et al., 2016). In a field survey of Swedish boreal forest soil, 60,000 to 1.2
million ECM have been reported in one square meter of forest soil in which 95% of
plant roots are reported to form ECM association. While investigating the ECM
associates of Pinus radiata over 2 years in New Zealand, Walbert et al. (2010)
documented eighteen species fruiting above ground and nineteen ECM species
fruiting below ground. Similarly 20 fungal taxa were reported to form ECM
association with Pinus muricata by Gardes and Bruns (1996) and 27 taxa with P.
thumbergii by Obase et al. (2009). In all these studies Cenococcum geophilum and
species of Clavulinaceae, Russulaceae and Thelephoraceae are reported to be the main
members of ECM fungal communities.
In Europe, Cenococcum geophilum and members of other genera including
Lactarius, Russula, Tomentella, Cortinarius, Laccaria and Paxillus were found to be
most diverse in these ecosystems (Rudawska et al., 2016). Most temperate and boreal
tree species of Europian mountains including Abies alba (Silver fir) have been docu-
mented to develop obligate mutualism with Lactarius, Russula, Tomentella, Laccaria
and Cortinarius, which plays significant role in the survival and growth of trees
(Ważny, 2014). Moreover in a Japanese conifer forest, Matsuda et al. (2009) docu-
mented the members of family Clavulinaceae, Russulaceae, Thelephoraceae and the
genus Trichophaea as ECM associates of Pinus thunbergii. Similarly, Kranabetter
et al. (2009) documented 63 ECM taxa, including a dark septate fungus, four
species of Piloderma, 7 of Tomentella and Psudotomentella, 8 of Russula, 27 of
Cortinarius, 1 of Tricholoma and several unknown fungi from boreal forest of
British Columbia (Canada).
In India, ECM diversity of different conifer species are being investigated from
North West Himalayas since 1979 by Lakhanpal and his associates (Bhatt &
Lakhanpal, 1990; Lakhanpal & Kumar, 1984). In South India, Mohan et al. (1993)
and Natarajan et al. (2005) has done substantial work on ECM association of different
fungi with coniferous trees. In a survey conducted in North-Western Himalyas for
about 12 year by Lakhanpal and his associats, 72 species belonging to 15 fungal genera
of mushrooms and toadstools were observed to form ECM association with Abies
pindrow, Betula utilis, Cedrus deodara, Picea smithiana, Pinus roxburghii, P.
wallichiana, Rhododendron arboreum, Quercus incana and Q. semicarpifolia. Beside
these records, ECM fungi forming close association with coniferous and
angiospermic trees have also been reported by various workers from India (Atri
& Saini, 1986; Atri et al., 1997; Bhatt & Lakhanpal, 1990; Kumar & Atri, 2016;
Pande et al., 2004; Sharma et al., 2008a, 2008b, 2009; Sharma et al., 2016;
Watling & Abraham, 1992). The lack of trained mycorrhizal taxonomists in
India and even in Asian continent has been a major limiting factor in the
generation of knowledge on ECM symbiosis from this region.
In China, The ECM community composition of Castanopsis fargesii (Fagaceae) was
investigated by Wang et al. (2011) in a subtropical evergreen broad-leaved forest and
Zhang et al. (2013) studied the ECM fungal communities of Quercus liaotungensis
(Fagaceae), along local slopes in the temperate oak forests on the Loess Plateau.
Members of Thelephoraceae (Tomentella spp.), Clavulinaceae (Clavulina spp.) and
Russulaceae (Lactarius spp.) are reported to be the most species-rich and abundant
ECM fungi in these regions. Similarly, Wang et al. (2017b) investigated ECM com-
munities associated with Quercus liaotungensis, from five typical habitats, and docu-
mented some of the dominant ECM lineages in these communities (/tomentella-
thelephora, /cenococcum, /russula-lactarius, and /inocybe). The ECM fungal richness
and diversity was reported to be positively correlated with soil organic matter and
elevation. Amongest the various lineages documented, /tomentella-thelephora, /russula-
lactarius, and /cenococcum are the most dominant lineages associated with Fagaceae in
a wide range of northern China (Smith et al., 2007; Jumpponen et al., 2010; Wang
et al., 2017b).
ECM Fungal diversity of wet and dry sclerophyll Australian temperate eucalypt
(Eucalyptus delegatensis, a Myrtaceae) forests was investigated by Horton et al. (2017).
They hypothesised that ECM fungal community richness and composition would differ
between forest types. Cortinariaceae represented the dominant family irrespective of
forest type. Similarly, Waseem et al. (2017) investigated ECM fungal diversity of
Tristaniopsis (Myrtaceae) tree species growing under contrasting soil conditions in
the natural ecosystems of New Caledonia, about 1200 km east from Australia. They
also documented Cortinarius to be the most dominant genus followed by Pisolithus
and Russula.
Only recently, tropical rain forests in South East Asia (Peay et al., 2010; Riviere
et al., 2007; Yuwa-Amornpitak et al., 2006), Africa (Bâ et al., 2012; Diédhiou et al.,
2010; Riviere et al., 2007; Suvi et al., 2010) South America (Geml et al., 2014; Morris
et al., 2008; Smith et al., 2011, 2013; Tedersoo et al., 2010b) and southern Ecuador
(Haug et al., 2005; Kottke et al., 2008, 2013) have been studied for below-ground ECM
fungal diversity. However, very little is known from the tropical forests that cover
relatively large forested areas in the world (Alexander & Selosse, 2009). ECM plants in
tropical forests mostly belong to families Dipterocarpaceae, Fabaceae, Juglandaceae,
Betulaceae, and Fagaceae (Henkel, 2003; Henkel et al., 2011; Morris et al., 2008).
Tropical Fagaceae are still most studied in comparision Dipterocarpaceae in this regard.
Castanopsis fargessii (Fagaceae) in subtropical evergreen broad leaved forest host 3
ECM fungi belonging to Ascomycetes and 14 belonging to Basidiomycetes especially,
Russulaceous and Thelephoroid members (Wang et al., 2011). Morris et al. (2008)
characterised diversity and richness of ECM communities in Quercus crassifolia in a
tropical forest in Southern Mexico and documented Russulaceae, Cortinariaceae,
Inocybaceae and Thelephoraceae as dominant ECM fungal families. Similar results
were documented by Hynes et al. (2009) in California and Lancelloti and Franceschini
(2013) in North–West Sardinia, while studying the ECM community in a declining
Fagaceae stand. Dipterocarpaceae in South – East Asia comprises 470 tree species
including dominant tree Shorea robusta, which is the only dominant tree species of
tropical South – East Asia including India (Maury-Lechon & Curtet, 1998). Despite
having great bio-geographic significance Shorea robusta ECM fungi have received little
attention. However, Shorea robusta associate with diverse fungi such as species of
Russula, Boletus, Agaricus, Amanita, Lactarius, Lactifluus, Cortinarius, Laccaria,
Pisolithus, Scleroderma, Suillus, Strobilomyces and Cantharellus (Kumar & Atri, 2016;
J. Kumar, N.S. Atri
Natarajan et al., 2005; Pyasi et al., 2011; Tapwal et al., 2013). Moreover dipterocarp forest
revealed 17 phylogenetic lineages spread over 69 species primarily belonging to the
/russula-lactarius, /tomentella-thelephora, /sordariales, /sebacina and /cantharellus line-
ages. As in temperate forests, these lineages were the most species-rich and Cenococcum
geophilum was found to be the most frequent fungal taxon (Dickie, 2007; Matsuda et al.,
2008, 2009; Phosri et al., 2012; Smith et al., 2013).
In tropical Africa, The /russula–lactarius and /tomentella– thelephora lineages dom-
inated ECM fungal flora on caesalpionioid legumes, Dipterocarpaceae, Sarcolaenaceae,
Phyllantaceae, Asterpeiaceae, Sapotaceae, Papilionoideae, Gnetaceae and Proteaceae.
Most studies, in Africa indicated that ECM is mainly found on caesalpinoid legume tree
species that play a major role in forestry and agroforestry (Bâ et al., 2012). As in the
temperate region, Russulales (Russula and Lactarius) showed the largest number of
described species in West Africa. Some of the species of Russula harvested from West
African region have also been described from East and Central Africa (Bâ et al., 2012).
Bâ et al. (2011) investigated 195 fungal taxa from West Africa, with dominant
thelephoroid taxa. Tedersoo et al. (2011) identified 18 phylogenetic lineages from
Zambia, Gabon, Madagascar and Cameroon, with some shared species. The work of
Tedersoo et al. (2011) confirmed the previous studies of ECM fungi from West Africa
(Riviere et al., 2007; Diédhiou et al., 2010) and the Seychelles (Tedersoo et al., 2007),
from where the dominance of species of the /russula–lactarius, /tomentella– thelephora,
/boletus and /pisolithus–scleroderma lineages have been reported. The remarkable
dominance of the /russula–lactarius and /tomentella–thelephora lineages is reminiscent
of temperate and other tropical forests (Horton & Bruns, 1998; Peay et al., 2010). Other
ECM fungal lineages such as /sebacina, /sordariales, /marcelleina-peziza gerardii and/
elaphomyces were absent or rarely encountered from African tropical habitats (Riviere
et al., 2007; Tedersoo et al., 2007; Diédhiou et al., 2010; Jairus et al., 2011), whereas
Cenococcum geophilum, one of the most widespread or dominant ECM fungi in
Holartic communities was absent (Bâ et al., 2012). In African tropical forests plant
species diversity is much higher than in temperate forests, and also the ECM fungi
associated with tropical trees could be very diverse and similar to that observed in
temperate forests; so further efforts should be made to assess the genetic and functional
ECM diversity of Africa.
ECM fungi are phylogenetically highly diverse in South America, but tropics are
very much overlooked in this regard (Roy et al., 2017). The orders Pezizales,
Agaricales, Boletales, Helotiales and Cantharellales include the largest number of
fungal lineages of host families Fabaceae, Fagaceae, Dipterocarpaceae,
Nyctaginaceae and Polygonaceae. ECM fungal diversity of Dicymbe corymbosa,
Dicymbe altsonii, and Aldina insignis trees in Guiana Shield are reported to show high
species diversity and richness in the ⁄russula-lactarius, ⁄boletus and ⁄tomentella-
thelephora lineages (Henkel et al., 2011; Smith et al., 2011), which was on a par with
dipterocarp forests of Borneo and many ECM-rich forests from boreal and temperate
zones (Peay et al., 2010). The community structure of ECM in Nothofagus forests of
South America are reported to be relatively diverse, mostly involving members of the
/cortinarius, /inocybe,/tomentella-thelephora, /clavulina and /tricholoma lineages
(Tedersoo et al., 2010a, 2010b, 2010c; Nouhra et al., 2013; Geml et al., 2014). The/
russula-lactarius lineage was relatively poor in this region, whereas the /suillus-
rhizopogon, /boletus and /pisolithus-scleroderma lineages were not recovered, which
is a unique distribution at the global scale (Tedersoo et al., 2012). Similarly, Moyersoen
and Weiß (2014) and Smith et al. (2013) examined the ECM fungal community
associated with Pakaraimaea dipterocarpacea in Venezuela and Guyana respectively,
and reported thirteen ECM fungal lineages (/amanita, /boletus, /cantharellus-craterellus,
/clavulina, /coltricia, /cortinarius, /hydnum, /inocybe, /russula-lactarius, /sebacina,
/tomentella-thelephora, /elaphomyces and /hysterangium). The diversity of ECM fungi
associated with Pakaraimaea dipterocarpacea is reported to be similar to what has
been documented with dipterocarps in Southeast Asia and Africa (Peay et al., 2010;
Tedersoo et al., 2011).
The different ECM communities on different plant species support similar ecosys-
tem functions in the soil (Wang et al., 2017a), but all these studies on ECM diversity
point out that Russulaceae show a great diversity in tropical forest ecosystems, and are
among the commonest ECM family (Maba et al., 2014, 2015; Malysheva et al., 2016;
Peay et al., 2010; Riviere et al., 2007). Smith et al. (2011) reported that, only the ⁄
russula-lactarius lineage is more diverse in tropical than in temperate habitats and by
contrast, the ⁄inocybe lineage is more diverse in the temperate zones. The dominance of
Russulaceae in low nutrient soil has been linked with its unique role in nutrient uptake
from the soil (Alexander, 2006).
ECM fungi are reported to be more or less host plant specific. On the basis of
specificity to the host, Molina and Trappe (1982) classified ECM fungi into three
groups, fungi with wide ECM host potential, fungi with limited host potential and fungi
with narrow host potential, that only form ECM with a specific host species. Most
ECM fungal species associate with a broad host range, or at least several species of the
same genus (Dickie, 2007). This potential has been attributed to the wood-wide-web of
fungal mycelium, where one fungus is reported to be connected with several plants to
stabilise forest ecosystem (Dickie, 2007; Diédhiou et al., 2010; Peay et al., 2015). Only
few ECM fungi are specialized on a single plant species (Bruns et al., 2002; Tedersoo
et al., 2008, 2010c). Reciprocally, only few ECM plants associate with a low number of
ECM fungi, such as the species of genus Alnus (Pritsch et al., 1997; Roy et al., 2013).
Most studies indicate that large numbers of ECM fungi are associated with
multihost. Peay et al. (2015) studied 13 genera of Dipterocarpaceae and observed
non significant differences in ECM communities of different genera. Similarly, Roy-
Bolduc et al. (2016) also observed non significant differences in ECM community of
four different tree species of family Pinaceae, indicating that at family level ECM host-
fungus interaction is well conserved. Some earlier studies, however, in contrast reported
host identity as the main determinant of ECM fungal community structure and com-
position (Ishida et al., 2007; Morris et al., 2008; Tedersoo et al., 2014). Due to the lack
of knowledge of complex interactions between host plants and microbial communities,
the extent of this host effect has been documented to remain incompletely understood
(Peay et al., 2015; Roy-Bolduc et al., 2016). In the northern hemisphere heath plants
(Ericaceae) and some groups of liverworts and lycophytes form mycorrhizal associa-
tions with the same group of fungi (Chambers et al., 1999; Horn et al., 2013). Several
myco-heterotrophic vascular plants have been shown to be epiparasitic upon
neighbouring photosynthetic plants through shared ECM fungal symbionts (Cullings
et al., 1996; Horn et al., 2013; Leake, 2004). Although photosynthetic plants are
generalists in their compatibility with fungal partners, the epiparasites examined so
far are reported to display exceptional specificity towards narrow groups of closely
J. Kumar, N.S. Atri
related fungi (Bidartondo et al., 2003). Leafy liverworts genera (Jungermanniales)

predominantly associate with the members of Sebaciana species and thalloid liverworts
associate nearly exclusively with Tulasnella species (Bidartondo & Duckett, 2010;
Pressel et al., 2010). For example, Cryptothallus (liverworts) associates with a narrow
clade of Tulasnella and that these same fungi have the ability to form ECM on Betula
and Pinus (Bidartondo & Duckett, 2010). Horn et al. (2013) reported that Sebacinales
Fungi colonise the Diphasiastrum alpinum (Lycopodiaceae) gametophytes as well as
adjacent Ericaceae plants simultaneously; indicating mycoheterotrophic gametophyte
to be epiparasitic on Ericaceae.
Recently, more and more studies have emerged on ECM diversity in plantations or
associated with invasive plants. Outside their native range, tree species are reported to
have relatively species-poor ECM communities in comparison to those growing in their
natural environment (Dickie et al., 2010; Nuñez et al., 2009; Tedersoo et al., 2007;
Walbert et al., 2010). This has been primarily attributed to low number or lack of
specific symbionts (Bahram et al., 2013; O’Hanlon, 2012; Ważny, 2014). More
generalist fungi are reported to survive through the invasion of several Pinaceae in
the southern hemisphere including Argentina, Brazil and New Zealand (Alberton et al.,
2014; Hayward et al., 2015; Hynson et al., 2013; Moeller et al., 2015). Moreover ECM
fungi with low host specificity tend to be more successful in colonizing new hosts in
the invaded ranges than those with high specificity (Wolfe & Pringle, 2012). For
example Amanita phalloids becomes widespread throughout North America, showed
host shift from pine plantation in which it was introduced, to pines and oaks in the
surrounding native forests (Pringle et al., 2009; Wolfe et al., 2010). In contrast high host
specificity ECM fungi can not grow beyond its host range and can not colonise new
host. For example, /suillus-rhizopogon lineage is specific to Pinaceae and even indi-
vidual species of ECM fungi to different genera of Pinaceae, and Laccaria which is
generalist shows additional host jumps which is responsible for diversification shifts
and dispersal events associated with its ECM ecology and dispersal throughout the
southern and northern hemisphere (Wilson et al., 2017). Thus ECM fungi of /suillus-
rhizopogon lineage is unsuccessful in colonizing new hosts in the invaded ranges than
those with low specificity lineages of ECM (Tedersoo et al., 2010c). So host preference
or specificity is the most important determinant of ECM fungal community composi-
tion in most studies ranging from local to global scale (Ishida et al., 2007; Tedersoo
et al., 2008, 2012; Bahram et al., 2013).
Factors Affecting ECM Diversity and Composition in Terrestrial

Ecosystem
It is well recognised that ectomycorrhizal fungal communities are diverse and species
rich, containing a few dominant species and some rare species (Horton & Bruns, 1998;
Izzo et al., 2005). ECM fungal diversity plays an important role in influencing the
ecosystem functioning, as well as plant diversity (Johnson et al., 2012; Kottke et al.,
2013; van der Heijden et al., 1998). In turn environmental and biological factors such as
host plant diversity and competition are also reported to influence fungal community
dynamics (van der Heijden et al., 1998; Peay et al., 2010). There are number of factors
including heterogenous distribution of nutrients, pH, elevation, soil moisture,
disturbance, coexistence of different host plants and competition between fungi and
other soil microbes which have been reported to play a significant role in this regard
(Bruns, 1995; Diédhiou et al., 2014; Druebert et al., 2009; Gao et al., 2014; Kennedy &
Peay, 2007; Kranabetter et al., 2009; Lambers et al., 2010; Malysheva et al., 2016).
Nutrient status of soil is reported to be the most important factor affecting the ECM
diversity and richness (Erlandson et al., 2016; Garcia et al., 2016; Taniguchi et al.,
2008; Tedersoo et al., 2011). Erlandson et al. (2016) studied the influence of soil
environment on ECM fungal communities across hydrologic gradient in temperate
North America and documented that ECM diversity is influenced by soil nutrient status
primarily P and N. Similar observations have also been made by Garcia et al. (2016) on
ECM communities of ponderosa pine (Pinus ponderosa) and lodgepole pine (Pinus
contorta) in south-central Oregon. Higher nutrient status of soil has been reported to
negatively influence the ECM diversity and richness (Corrales et al., 2016; Cox et al.,
2010; Lilleskov et al., 2002; Parrent et al., 2006). Erlandson et al. (2016) documented
that ECM fungi richness and diversity is negatively correlated with phosphorus
availability. Similar observations have been made by Corrales et al. (2016) while
working with the Oreomunnea adult saplings and seedlings across site differing in soil
fertility. In the characterization of the ECM fungal community associated with
Oreomunnea, it has been documented that, infection frequency of ECM fungi is lower
in more fertile soils, which is consistent with the general view that benefits of ECM
fungi depend on soil conditions (Treseder, 2004). So in high-fertility sites phylogenetic
diversity of ECM fungi and ECM colonization rate has been reported to decrease.
The decrease in pH level in the soil up to certain extent has been reported to
positively affect ECM diversity (Barrow, 1984; Benucci et al., 2016; Kluber et al.,
2010; Wang et al., 2017a). Marx et al. (1984) while investigating the effect of soil pH
and Pisolithus tinctorius on pecan seedlings (Carya illinoensis) documented that the
percentage of roots infected by ECM increased from 22% to 44% as soil pH decreased
from 6.5 to 5.5. Plant dry weight, N and K content has also been reported to increase
with decreasing pH.
Elevated atmospheric CO2 concentration has also been reported to affect ECM
diversity and richness by increasing carbon allocation to ECM fungi by their tree host
(Andrew & Lilleskov, 2009; Druebert et al., 2009; Parrent et al., 2006; Rouhier &
Read, 1999). It is also reported that elevated CO2 more strongly and positively
influences production and composition of ECM sporocarps which suggest that
ECM sporocarps are most sensitive to a reduction in C supply (Andrew &
Lilleskov, 2014). With the available techniques in hand the elevated CO2 concen-
tration and future climate change will definitely have a strong influence on ECM
diversity (Cairney, 2012; Corcobado et al., 2015) and this should be seen as
priority area for future research.
Light is also another important abiotic factor that has been reported to influence
ECM diversity and richness.Various studies have shown that plant biomass decreases
with the reduction of light (Tester et al., 1986; Welander & Ottosson, 1998). Recently it
has been shown that under low light condition, Pinus sylvestris seedlings inoculated
with the ECM fungus Suillus bovinus decreased P acquisition and P transfer to the host
plant (Bücking & Heyser, 2003; Nehls et al., 2010). While working on this aspect,
Lambers et al. (1998) reported that shaded ECM plants showed lower root biomass in
comparison to plants grown under optimum light conditions. This is primarily because
J. Kumar, N.S. Atri
under low light condition plants are reported to allocate proportionally less carbon to
the roots which result in decrease in the ECM fungal diversity and richness.
It has been reported that in contrast to light, host damage, clear cutting, low soil
moisture, and elevation may also negatively affect ECM fungal abundance and diver-
sity (Lilleskov et al., 2002; Perez-Moreno & Read, 2000; Trocha et al., 2016;
Kyaschenko et al., 2017). Trocha et al. (2016), while evaluating the effect of biotic
(interspecific competition) and abiotic (organ loss or damage, light shortage) stresses on
ECM root tip colonization, diversity, seedling biomass, and nitrogen content of leaf,
reported that for both light demanding (Pinus) and shade-tolerant (Fagus) species, the
amount of light had a more pronounced effect on ECM colonization and diversity than
did juvenile damage. Wang et al. (2017a) reported that, in Austrian Alps species
richness and diversity of ECM fungi decreased with increasing elevation and decreas-
ing soil moisture. ECM diversity and species richness has also been reported to be
affected by successional stages in the ecosystem. With the ecosystem development, the
communities of ECM fungi are reported to become more diverse with early-
successional species and additional species or late successional species which appear
in the later stages of ecosystem development (Dickie et al., 2009; Kyaschenko et al.,
2017; Muhlmann et al., 2008; Nara, 2006; Peay et al., 2011, 2012).
All these factors shape ECM diversity locally, and interestingly, at a larger scale,
some other factors explain ECM distribution, such as the host, soil quality and volume
(Tedersoo et al., 2012, 2014). The host remain the main factor shaping ECM commu-
nities, which confirms that ECM fungi preferences are conserved at a larger scale
(Bahram et al., 2013; Tedersoo et al., 2012). For example, Tedersoo et al. (2014) have
compared Alnus ECM fungal communities across large scales (Europe, Asia, South
America and North America), and found that soil chemistry explained only a small
amount of variability. Intrageneric phylogenetic relations among Alnus spp. indicate
that closely related hosts generally exhibit more similar fungal communities largely
independent of geographical distance and environmental variables at global scale.
Finally, the distribution of ECM diversity centered in the temperate zone, contrasts
with the latitudinal gradient of diversity observed for plants and insects. This trend
shows that ECM biogeography may also modulate their species richness locally. More
and more studies have investigated the biogeography of ECM genera (Kennedy et al.,
2012; Looney et al., 2016; Matheny et al., 2009), and such studies have highlighted that
tropical zone could host diverse ECM taxa, or even their center of origin (for
Inocybaceae, see Matheny et al., 2009). Most ECM fungi have relatively restricted
ranges. Kõljalg et al. (2013) reported that approximately 80% of molecularly defined
species level operational taxonomic units were restricted to a single continent. Tedersoo
et al. (2014) also found that, at a global scale, biogeographic regions contained unique
species and the differences between geographic regions are consistent with the effec-
tiveness of oceans and mountains as dispersal barriers, which play a major role in the
current structure of ECM fungal communities. A major aim of these biogeographical
studies is to identify predictable drivers of ECM fungal composition, so there is a need
to work more on ECM biogeography. It is reported that ECM fungi follow biogeo-
graphical rules of microbes, such as island biogeography (Bahram et al., 2013; Peay
et al., 2012, 2015) and relationships with altitude (Bahram et al., 2013; Tedersoo et al.,
2012). Bahram et al. (2013) try to find out the spatial patterning and the underlying
mechanisms driving these patterns across different ecosystems at the local and global
scales. They examined the distance decay of similarity– diminishing similarity with
increasing geographical distance. The distance-decay relationship reflects the rate of
ECM species turnover (i.e. beta diversity) with increasing geographical distance and
enables the prediction of gamma diversity (global richness) based on alpha diversity
(local richness; Bahram et al., 2013). Distance from the equator and host density was
reported to be the main determinants of the extent of distance decay in ECM fungal
communities. Tropical ECM fungal communities are reported to be exhibit stronger
distance-decay patterns compared to non-tropical communities, suggesting a relatively
greater spatial aggregation of ECM fungi diversity and richness in tropical ecosystems.
At the global scale, Bahram et al. (2013) reported that lineage-level ECM community
similarity decayed faster with latitude than with longitude, suggesting that climate has
an important effect on distance decay of ECM fungal communities at the global scale,
directly or indirectly by influencing host-plant distribution and soil processes. Despite
considerable progress in our understanding of alpha diversity and community compo-
sition of ECM fungi, little is known about spatial structure of ECM fungal communities
in different ecosystems and the relative roles of niche processes in creating these
patterns of diversity.
Morphoanatomical and Molecular Studies on ECM
Despite recent advances in the use of molecular techniques, there are still many
advantages associated to classical methods for studying ECM fungal diversity.
Tracing mycelial connections between fruit bodies and ECM is still the most reliable
way of assessing the trophic status of fungi in the field. For the recognition of fungal
relationship and type of mycorrhizal association is advantageous over molecular
method (Rinaldi et al., 2008). Some time morphoanatomical based taxonomy is not
well supported by molecular taxonomy. To overcome such discrepancy, combined
approach of morphoanatomical and molecular characterization of ECM in combination
with phylogeny was applied (Mrak et al., 2017). Before molecular revolution, most
studies on ECM roots were focused on morphoanatomical characters. Detailed struc-
ture of the ECM mantle was for the first time given by Foster and Marks (1966). Zak
(1973) documented that mantle surface can range from thin to profuse and texture may
vary from smooth, cottony, velvety and warty to granular. The mantle can differ in
organisation, colour, texture, thickness and presence or absence of cystidia on mantle
surface, and Hartig net, depending on the host and ECM fungus identity (Agerer, 1986;
Smith & Read, 2008; Tedersoo et al., 2010a). The presence or absence of rhizomorphs
and mycelial growth pattern have also been used to classify fungi by exploration type,
which is an important aspect to understand their ecological function (Agerer, 2001),
and the concept has since been widely used in the studies of ECM ecology.
Before the contribution of Agerer and his co-investigators (Agerer, 1986, 2006;
Agerer & Rambold, 2004-2016) the method for describing the ECM association varied
greatly. Agerer through series of publications started to publish descriptions for iden-
tification and characterisation of ECM which has now become standard and is being
widely used throughout the world. Computer based software has also been developed
to follow uniform pattern for characterisation and determination of ECM (Agerer &
Rambold, 2004-2016). Agerer (1986) described and identified the mycorrhizae of
J. Kumar, N.S. Atri
Spruce with Lactarius deterrimus, L. picinus, Russula ochroleuca and R. xerampelina

by tracing hyphal connections between the fruiting bodies and the ECM roots and also
observed that morphological and anatomical characteristics of ECM are conserved at
the genus and species level. Yamada et al. (2001) worked out ECM either synthesized
in vitro or produced in natural conditions in association with Pinus densiflora seedlings
and reported that mycorrhizal morphology and anatomy within a species/isolate may
vary with substrate and other external environmental conditions. The similarity in
hyphal features of sporophores and the mantle, sometimes help in conformation of
organic connections. For example, Kumar and Atri (2016) and Leonardi et al. (2016)
reported similar laticifers in mantle and sporocarp while investigating the ECM formed
by Lactifluus. Indeed, observations on ECM roots are scarce and more observations in
Neotropics are still required to correct sampling bias, and to answer more specific
puzzels and challenges (Roy et al., 2017).
Most studies available till date have focused on ECM of coniferous trees such as
Picea and Pinus (Agerer & Rambold, 2004-2016). Roman et al. (2005) compiled 1244
descriptions of ECM published in 479 papers till 2005. Most ECM described by them
were collected from Europe (862), Germany (331) followed by Italy (177). A signif-
icant number of ECM was also reported to be collected from USA (96) and Canada
(175). Gymnosperms are reported to be the most common tree associates in as many as
510 descriptions with Pinaceae being the commonest host. Among angiosperms, only
members of Fagaceae family are reported to be the ECM hosts in 339 descriptions with
Quercus (188) being the commonest host followed by Fagus (116). Other angiosperms
are poorly represented in comparison (Fig. 1). In South America, most studies on ECM
descriptions deal with Fagaceae, Fabaceae, Nyctaginaceae, and Polygonaceae family
(Becerra & Zak, 2011). However, in Africa, most ECM described belongs to
caesalpionioid legumes, Dipterocarpaceae, Sarcolaenaceae, Asterpeiaceae,
Sapotaceae, Gnetaceae, and Proteaceae host (Bâ et al., 2012). The available data
indicate that most ECM studies have been undertaken in the temperate and boreal
forests. The ecologically and economically most important ECM trees dominate
woodland and forest communities in boreal, Mediterranean, and temperate forests of
the Northern Hemisphere and parts of South America, seasonal savanna and rain forest
habitats in Africa, India and Indo-Malay as well as temperate rain forest and seasonal
woodland communities of Australia. ECM-dominated habitats in Southeast Asia,
Africa, Australia and to some extent South America, remain undersampled relative to
the north temperate regions (Fig. 2). In comparision, ECM diversity of tropical and sub
tropical plants is least investigated, which require exploration in this regard to supple-
ment the present figures.
Earlier studies on ECM fungal communities were only based on morphological and
anatomical observations. However, these investigations proved to be time consuming
and the connection between feeder root and ectomycorrhizal fungi was sometime hard
to trace. The application of molecular techniques has dramatically changed the situa-
tion. Along with ECM morphoanatomical study it is now possible to investigate
accurately both fungi and plant (Tedersoo et al., 2006). The ECM species that dominate
in a given forest soil are not the abundant fruiting ones, indicating fruiting -body based
fungal inventories are not sufficient to describe ECM fungal communities (Zoll et al.,
2016). In recent years, use of molecular methods has provided new insights into the
below-ground fungal community and a more precise approach to fungal diversity
Fig. 1 Pie chart showing the proportion of ECM plant families studied so far
studies.The Genetic analysis of ECM symbiosis is largely being done using PCR
techniques. Internal transcribed spaces (ITS region) of rDNA are reported to be the
most frequently used sequences for the identification of ECM fungi (Avis, 2012;
Bahram et al., 2011; Benucci et al., 2016; Healy et al., 2013; Horton et al., 2017).
The DNA analysis of ECM also relied on the use of restriction fragment length
polymorphisms to evaluate the unique types of strains (Dahlberg, 2001; DeBellis
et al., 2006; Gehring et al., 2006; Gryta et al., 2006; Molinier et al., 2016; Wang &
Guo, 2010). The techniques were often improved to avoid sequencing contaminants: a
direct PCR analysis of ECM mantle can be more accurate and precise than amplifying
the whole root tip (Lotti & Zambonelli, 2006). The development of late oligo-array
based transcript profile now allow to investigate the molecular mechanisms within
ECM root tip, such as N and P exchanges (Willmann et al., 2014; Zhang et al., 2010;
Zheng et al., 2016), hormone cross-talks (Plett et al., 2014) and belowground ECM
fungal communities interactions (Taniguchi et al., 2007). Comparative proteome anal-
ysis between mycorrhizal and nonmycorrhizal plants is also used to understand the
mechanism behind nutrient exchange (Sebastiana et al., 2017).
Several scientists have pioneered the use of molecular markers for studies into the
ecology of ECM fungi and contributed to the recent advances in mycorrhizal research
where DNA sequencing and phylogenetic analyses have been used to identify species,
and trace the phylogenetic position of known and unknown ECM fungi directly from
ECM root tips (Avis, 2012; Bahram et al., 2011; Benucci et al., 2016; Healy et al., 2013;
Horton et al., 2017; Morris et al., 2008; Smith & Peay, 2014; Tedersoo et al., 2014).
J. Kumar, N.S. Atri
Fig. 2 Global map showing location of the ECM diversity and morphoanatomical study sites (Triangles)
Through DNA-sequencing and phylogenetic analyses, it has become clear that many of
the unidentified taxa on the roots belonged to species which had previously not been
known to form ECM. The data provided by DNA sequences have provided the tools
necessary for mapping of microbial distributions, and have advanced the development
of ECM fungal biogeography in a number of critical dimensions (Tedersoo et al., 2014).
Recent advances in DNA sequencing have greatly progressed the field of ECM ecology
and allowed for the study of complex communities in unprecedented detail. Next
generation sequencing (NGS) can reveal powerful insights into the diversity and
richness of cryptic ECM species. NGS is reported to be advantageous compared with
the traditional Sanger method because of its much higher data throughput and much
lower cost. It works best when species are mixed together, and generates thousands to
millions of sequences at low cost, where traditional methods do not work well (Smith &
Peay, 2014; Zoll et al., 2016). The traditional methods are adequate for sequencing ITS
from fruit bodies, pure cultures, and even individual ECM root tips, but not as suitable
when samples include multiple species (forest soil sample). NGS have made outstanding
contributions for our understanding of ECM fungal diversity, ecology and biogeography
(Smith & Peay, 2014; Peay & Matheny, 2017; Roy et al., 2017).
Molecular and proteomics techniques in ECM provide opportunities for visualizing
how organisms interact in the soil and to assess the ECM community structure. The use
of fluorescent markers, that can reveal the location of the nucleic acid (DNA) of
specific organisms, can provide important additional information on microbial-plant-
soil interactions. With the advances in sequencing technology it becomes possible to
further explore mycorrhizal networks and interactions with other organisms, including
interactions with bacteria colonizing the ECM or even endosymbiotic bacteria living
inside mycorrhizal hyphae (Deveau et al., 2016).
Potential Role of Ectomcorrhiza to Sustain Terrestrial Ecosystems
ECM associations bring several advantages to host plants. Potential role of ECM in
plant growth and development have been evaluated by number of investigators includ-
ing Aroca et al., 2009; Camilo-Alves et al., 2013; Danielsen et al., 2013; Diagne et al.,
2013; Jourand et al., 2014; Kayama & Yamanaka, 2014; Luo et al., 2011, 2014; Maurel
& Plassard, 2011; Mohan et al., 2015; Tapwal et al., 2015 Xu et al., 2016 and Zong
et al., 2015.They enhance water absorption, nutrient uptake, reduce the need of external
fertilizer, improve plant resistance against pathogens, improve seedling growth, surviv-
al and establishment, can protect against heavy metal stress and other pollutants.
However, ECM fungal species show differences in host compatibility and host growth.
The host populations have diverged in community composition of their ECM fungi,
and have also diverged genetically in several traits related to interactions of seedlings
with particular ECM fungi, growth, and biomass allocation. Even some ECM symbi-
onts are reported to decrease the growth of plant. Patterns of genetic variation among
host plant populations for compatibility with ECM fungi has been reported to differ for
the species of ECM fungi, suggesting that host plant can evolve differently in their
compatibility and function with different symbiont species (Hoeksema et al., 2012).
Role in Growth and Establishment of Seedlings
The idea of inoculating ECM fungi on seedlings in plant nurseries was developed by
Fortin (1966). Vozzo and Hacskaylo (1971) while working on ECM in United States
experimentally demonstrated that field survival and growth of tree seedlings with
specific potantial ECM enhance the performance of seedlings and contribute to the
proper functioning of forest ecosystems. Numerous investigators demonstrated the
impact of ECM fungi in enhancing the survival rate and early growth performance of
various plant species (Danielsen et al., 2013; Kayama & Yamanaka, 2014; Kumla et al.,
2016; Menkis et al., 2012; Rincón et al., 2007; Tapwal et al., 2015).
Several techniques were developed to inoculate nursery seedlings with selected
ECM fungi. Mostly spore, solid substrate and liquid mycelia slurry inoculi are being
used in normal practice (Marx et al., 1991; Molina & Trappe, 1982, 1994; Wan et al.,
2016). Out of these solid substrate inoculums (vermiculite- peat or wheat grains) of
ECM fungi are reported to be as effective as liquid mycelia slurry and more effective as
compared to spore slurry inoculums (Quoreshi et al., 2008; Quoreshi & Khasa, 2008).
However, soil obtained from natural forests or established plantations was also directly
used. One problem with this type of inoculant is that large amounts of soil are required
to inoculate nursery plants, and another important problem is the risk of introducing
plant pathogens and weeds. Moreover, there is no precise information on the introduced
fungal species (Castellano & Molina, 1989).
Potential isolates of ECM fungi are selected based on their efficiency and compat-
ibility which is usually measured by evaluating parameters such as the height of plant,
the diameter of the stem, the overall fresh, the dry mass and the nutrient content of the
inoculated plant, especially phosphorus (Desai et al., 2014; Jourand et al., 2014; Marx
et al., 1991). Turjaman et al. (2005) reported a greater survival of Shorea pinanga,
when seedlings were inoculated with Pisolithus arhizus and Scleroderma sp. in com-
parison to control seedlings. Artificially inoculated ECM fungi are also reported to
J. Kumar, N.S. Atri
enhance growth and nutrition of the seedlings both under nursery conditions and in the
field after outplanting (Browning & Whitney, 1993; Kropp & Langlois, 1990; Quoreshi
et al., 2008; Villeneuve et al., 1991). The establishment of Diphasiastrum
(Lycopodiaceae) gametophytes by their ECM fungal partners are reported to develop
improved conservation strategies for this genus, which is endangered throughout
Central Europe (Grulich, 2012; Horn et al., 2013).
In some ECM association even bacteria are reported to play a mediatory role in
mycorrhizal symbiosis. In plant-fungus-bacterial association mycorrhiza helper bacteria
(MHB) are reported to have positive influence in enhancing the efficiency and intensity
of the ectomycorrhizal symbiosis (Brulé et al., 2001; Cumming et al., 2015; Deveau
et al., 2016; Dominguez et al., 2012; Izumi et al., 2006; Kurth et al., 2013; Mediavilla
et al., 2016; Shakya et al., 2013; Zhang et al., 2010). Mediavilla et al. (2016) undertook
in- vitro synthesis of ECM between Boletus edulis and Cistus ladanifer to test the
effects of fungal culture and co-inoculation with the MHB Pseudomonas fluorescens.
This co-inoculation with a MHB doubled the plant mycorrhization and plant growth
levels as compared to plant colonized with Boletus edulis alone. Similarly significant
increase in colonization of Pinus halepensis root by Tuber melanosporum in the
presence of Pseudomonas fluorescens has been reported by Dominguez et al. (2012).
Pseudomonas fluorescens greatly improve the symbiotic relationship of ECM by
increasing the percentage of mycorrhizal short roots to total short roots (Duponnois
& Garbaye, 1991; Duponnois, 2006), the stimulation of mycelial growth, the reduction
of environmental stress on the mycelium and solublisation of nutrients (Brulé et al.,
2001; Frey-Klett et al., 2007; Kurth et al., 2013).
Role in Regeneration of Forest Ecosystems
It is well established that artificial ECM inoculation has a great potential in the
restoration of natural ecosystems, degraded and disturbed sites (Bent et al., 2011;
Bois et al., 2005; Danielson & Visser, 1989; Dickie et al., 2013; Marx et al., 1991;
Miller & Jastrow, 1992; Rincón et al., 2006; van der Heijden & Horton, 2009).
Successful revegetation of severely disturbed mine lands in various parts of the world
has been accomplished successfully by using biological tools. The inoculation of
nursery seedlings with appropriate ECM fungi is reported to be the most environment
friendly approach, particularly, for disturbed and degraded ecosystem. This inoculation
of nursery seedlings with appropriate ECM fungi is known to promote uptake of
nutrients and water, protection against various stresses, increased resistance
against some pathogens and enhanced seedling regeneration and performance
(Bois et al., 2005; Quoreshi et al., 2008;). Without a host, however, the amount
and diversity of ECM fungal inoculums has been reported to decrease rapidly
(Dahlberg, 2002; Jones et al., 2003).
Role of ECM in Improving Soil Fertility
It is well known that soil microorganisms influence the growth and development of
plant communities through increasing soil nutrient availability and mediating plant co-
existence (Bâ et al., 2002; Bonfante & Genre, 2010; Pritsch & Garbaye, 2011; van der
Heijden et al., 2008; van der Putten et al., 2013). It is a well established fact that in
ECM fungi improves the nutritional status of their host plant by supplying macro and
micronutrients (Table. 1) while plant provides increased allocation of carbohydrates to
the fungi in order to sustain the symbiosis (Sebastiana et al., 2017). Hatch (1936) was
amongst the pioneers to document increased N and P content in mycorrhizal white pine
(Pinus strobus) in comparison to non mycorrhizal plant. ECM fungi are reported to
produce ectoenzymes which help to facilitate the release of organic N and P into the
soil which are otherwise unavailable to the plants (Desai et al., 2014; Ma et al., 2013;
Quoreshi & Khasa, 2008).
Mycorrhizal symbionts are reported to improve the host plant nutrient uptake,
especially P uptake quite efficiently (Bucher, 2007; de Campos et al., 2013; Smith &
Read, 2008; Stonor et al., 2014). Smith and Smith (2012) and Pedersen et al. (2013)
documented that high affinity phosphate transporters of mycorrhizal fungi play an
important role in absorbing P from outside of P depletion zone. Similar observation was
made by Facelli et al. (2014). From ECM fungi, the PT genes in case of Boletus edulis
(Wang et al., 2014), Hebeloma cylindrosporum (Tatry et al., 2009), Laccaria bicolor
(Martin et al., 2008) and Tuber melanosporum (Martin et al., 2010) have been
characterized. So far at least 34 PT genes of 11 ECM fungal species, including
Amanita muscaria, Laccaria amethystina, Paxillus involutus, Paxillus rubicundulus,
Piloderma croceum, Pisolithus microcarpus, Pisolithus tinctorius, Scleroderma
citrinum, Sebacina vermifera, Suillus luteus, and Tulasnella calospora are reported
(Casieri et al., 2013).
ECM fungi contribute to increased P demand of trees when they increase their root
colonisation level, mostly during active growth period of plant because at that time
requirenment of nutrient has been reported to increase many fold (Cairney, 2011;
Szuba, 2015). While working on Pinus tabulaeformis during active growth period in
Northern China, Zhang et al. (2010) isolated two phosphate transporter genes, RlPT and
LbPT from Rhizopogon luteolus and Leucocortinarius bulbiger, respectively. Further
investigations have revealed that RlPT and LbPT are significantly up-regulated at lower
P level, and enhance P uptake and transport (Zheng et al., 2016).
Importance of ECM fungi in the N nutrition of trees and in forest N cycling process
has been emphasised in recent years (Averill et al., 2014; Avis et al., 2008; Cairney,
2011; Danielsen et al., 2013; Obase et al., 2009; Taylor et al., 2004; Willmann et al.,
2007, 2014; Wu et al., 2003). ECM fungi are reported to use different nitrogen sources
such as nitrate, NH4+, urea (Guidot et al., 2005; Morel et al., 2008), di- and tripeptides
(Benjdia et al., 2006), and protein (Guidot et al., 2005). Mycorrhiza formation strongly
increase expression of fungal high-affinity NH4+ transporters (Couturier et al., 2007;
Javelle et al., 2001, 2003; Kuster et al., 2007; Willmann et al., 2007).
The breakdown and mobilization of nitrogen from complex organic matter, as well
as the nitrogen transporter genes transcription, are strongly activated by carbon avail-
ability (Rineau et al., 2013). The inhibition of mycorrhization when mineral supply is
sufficient and its subsequent reversal when mineral supply is insufficient has been
reported (Kemppainen et al., 2009), revealing the crucial role of the nitrogen in
establishing ECM symbiosis. Javelle et al. (2003) reported that genes that are typically
induced by nitrogen starvation are suppressed by high nitrogen availability. The
silencing of LbNrt—the nitrate-transporter–encoding gene of the fungus substantially
decreases growth and symbiotic interaction with plant (Kemppainen & Pardo, 2013),
illustrating the important role of fungi during the establishment of ECM.
Table 1 ECM fungi, associated host plant and their role in nutrient uptake
Sr.No Fungi Host Nutrient Refrences
1 Hebeloma crustuliniforme, Betula pendula N Abuzinadah & Read, 1986

Amanita muscaria,
Paxillus involutus
2 Trichoderma harzianum, Laccaria laccata Pinus wallichiana N, P, K Ahangar et al., 2012
3 Suillus varigatus, Pinus sylvestris P, K, Mg, S Ahonen-Jonnarth et al., 2000
Rhizopogon roseolus
4 Hebeloma longicaudum, Laccaria bicolor Pinus sylvestris N, P, K, Mg Ahonen-Jonnarth et al., 2003
5 Paxillus involutus, Picea abies P, Ca Andersson et al., 1996
Betula pendula
6 Paxillus involutus, Pinus spp. N Arnebrant, 1994
Suillus bovines
7 Pisolithus tinctorius Eucalyptus pilularis P Ashford et al., 1999
8 Suillus bovinus, Schima wallichii P Bendangmenla & Ajungla, 2014
Boletus edulis,
Scleroderma citrinum
9 Hebeloma cylindrosporum Pinus pinaster K Benito & Gonzalez-Guerrero, 2014
10 Descolea maculata, Eucalyptus diversicolor P Bougher et al., 1990
Pisolithus tinctorius,
Paxillus involutus
11 Paxillus involutus Picea abies P Brandes et al., 1998
12 Suillus bovinus Populus tremela P Bücking & Heyser, 2001
P. alba
13 Suillus bovines Pinus sylvestris P Bücking & Heyser, 2003
14 Laccaria lateritia Eucalyptus globules P Chen et al., 2000
15 Paxillus involutus, Pinus sylvestris P Colpaert et al., 1999
J. Kumar, N.S. Atri
Table 1 (continued)
Suillus luteus,
Suillus bovinus,
Thelephora terrestris
16 Laccaria bicolor, Pinus rigida P Cumming, 1996
Paxillus involutus
17 Pisolithus tinctorius, Rhizopogan vulgaris, Suillus Pinus wallichiana N, P Dar et al., 2007
granulates, Laccaria laccata,
Hebeloma crustuliniforme,
18 Laccaria bicolor Populus tremuloides P Desai et al., 2014
19 Pisolithus albus, Scleroderma dictyosporum, Acacia mangium N Diagne et al., 2013
S. verrucosum,
Scleroderma sp.
20 Tuber melanosporum Quercus ilex, P Domínguez Núñez et al., 2006
Quercus faginea.
21 Tuber melanosporum Quercus petraea, Quercus faginea, P, N, K, Ca, Mg Domínguez Núñez et al., 2008
Pinus halepensis
22 Rhizopogon roseolus, Pinus sylvestris N Finlay et al., 1988
Suillus bovinus,
Paxillus involutus
23 Thelephora terrestris Pinus contorta N Finlay & Söderström, 1992
24 Amanita muscaria, Elaphomyces antracinus, Pisolithus Eucalyptus urophylla N, P, K Gandini et al., 2015
microcarpus, Scleroderma areolatum
25 Hebeloma cylindrosporum Pinus pinaster P, K Garcia et al., 2014
26 Thelephora terrestris Pinus sylvestris N Hilszczańska et al., 2008
27 Pisolithus arhizus Pinus sylvestris N Högberg et al., 2003
Table 1 (continued)
28 Paxillus involutus Picea abies Mg Jentschke et al., 2000

29 Laccaria bicolor, Thelephora terrestris Eucalyptus coccifera P Jones et al., 1998
30 Wilcoxina sp., Cenococcum sp., Amphinema byssoides Picea engelmannii N Jones et al., 2009
31 Pisolithus albus Acacia spirorbis, Eucalyptus globules N, P Jourand et al., 2014
32 Scleroderma verrucosum Quercus acutissima P, K, Mg, Ca Jung & Tamai, 2013
33 Rhizopogon occidentalis, Pinus muricata N Kennedy & Peay, 2007
R. salebrosus,
R. vulgaris,
Tomentella sublilacina
34 Pisolithus albus Eucalyptus tereticornis Ca, K Khosla & Reddy, 2008
35 Paxillus involutus Populus deltoids P, Ca, Mg Khosla et al., 2009
36 Pisolithus sp. Pinus pinaster N, P, K, Ca, Mg Lamhamedi et al., 1992
37 Laccaria laccata Fagus sylvatica K, Mg Leyval & Berthelin, 1989
38 Pisolithus tinctorius Pinus caribaea var. hondurensis N, P Marx et al., 1985
39 Hebeloma cylindrosporum Pinus pinaster N Müller et al., 2007
40 Suillus tomentosus Pinus contorta N Paul et al., 2007
41 Laccaria laccata Picea mariana N Quoreshi & Timmer, 2000
42 Melanogaster ambiguous, Pisolithus tinctorius, Rhizopogon Pinus pinea N, P Rincón et al., 2005
luteolus, Rhizopogon roseolus, Scleroderma verrucosum
43 Pisolithus tinctorius, Pinus taeda P Rousseau et al., 1994
Cenococcum geophilum
44 Hebeloma crustuliniforme Populus tremuloides N Siemens et al., 2011
45 Hebeloma cylindrosporum Pinus pinaster P Tatry et al., 2009
46 Amanita rubescens, Pinus sylvestris N Taylor et al., 2004
J. Kumar, N.S. Atri
Table 1 (continued)
Lactarius deterrimus
47 Pisolithus arhizus, Scleroderma columnare Shorea seminis N, P Turjaman et al., 2006
48 Suillus variegates Pinus sylvestris P Wallander, 2000
49 Pisolithus tinctorius Pinus resinosa N Wu et al., 2003
50 Pisolithus sp., Pinus densiflora, K, P, Ca, Mg Zong et al., 2015
Cenococcum geophilum, Laccaria laccata Quercus variabilis
Table 2 ECM fungi and their plant associate mediating heavy metal stress
Sr.No. Fungi Host Heavy metal Refrences
1 Suillus luteus Pinus sylvestris Zn Adriaensen et al., 2004

2 S. luteus P. sylvestris Cu Adriaensen et al., 2005
3 S. bovines P. sylvestris Zn Adriaensen et al., 2006
4 S. varigatus, P. sylvestris Al, Cu, Ni Ahonen-Jonnarth et al., 2000
Rhizopogon roseolus
5 Hebeloma longicaudum, Laccaria bicolor P. sylvestris Al Ahonen-Jonnarth et al., 2003
6 Pisolithus tinctorius Eucalyptus grandis Mn Canton et al., 2016
7 Pisolithus sp., Pinus densiflora Cu Chen et al., 2015
Cenococcum geophilum
8 Paxillus involutus Pinus sylvestris Zn Colpaert & Van Assche, 1993
9 Amanita muscaria, Cenococcum geophilum, Laccaria laccata, Pinus spp. Hg Crane et al., 2010
Piloderma bicolor, Pisolithus tinctorius, Suillus decipiens,
10 Pisolithus tinctorius, Pinus rigida Al Cumming & Weinstein, 1990
Hebeloma crustuliniforme
11 Hebeloma crustuliniforme Picea abies Cd, Zn Frey et al., 2000
12 Pisolithus tinctorius Populous alba X glandulosa Cd Han et al., 2011
13 Paxillus involutus Picea abies Al Hentschel et al., 1993
14 Laccaria bicolor, Picea abies Cd Jentschke et al., 1999
Paxillus involutus
15 Pisolithus albus Eucalyptus globules Ni Jourand et al., 2010
16 Pisolithus albus Acacia spirorbis, Eucalyptus globules Jourand et al., 2014
17 Paxillus involutus Populus deltoids Al Khosla et al., 2009
18 Suillus luteus Pinus sylvestris Cd Krznaric et al., 2009
J. Kumar, N.S. Atri
Table 2 (continued)
Sr.No. Fungi Host Heavy metal Refrences
19 Suillus luteus Pinus sylvestris Zn Krznaric et al., 2010

20 Astraeus hygrometricus, Quercus glauca, Al Kayama & Yamanaka, 2014
Scleroderma citrinum Q. saliciana, Castanopsis cuspidate
21 Pxillus involutus Populus x canascens Cd Ma et al., 2013
22 Pisolithus tinctorius Pinus taeda Al Moyer-Henry et al., 2005
23 Pisolithus tinctorius Pinus strobes Al Schier & McQuattie, 1995
24 Cadophora finlandica Salix sp. Cd, Zn Utmazian et al., 2007
25 Suillus bovines, Pinus sylvestris Cu Van Tichelen et al., 2001

Thelephora terrestris
J. Kumar, N.S. Atri
It is well known fact that fungi exude several organic acids that contribute to the
release of some other nutrients such as potassium (K), calcium (Ca) and magnesium
(Mg) which become available to the plants (Benito & Gonzalez-Guerrero, 2014;
Kayama & Yamanaka, 2014; Plassard & Dell, 2010). So as to obtain C in the form
of sugars, ECM fungi provide physical, physiological and biochemical access to
nutrients in soil that the host tree would otherwise be unable to access. Similarly soil
bacteria which ubiquitously colonize these ECM roots and rhizospheres are document-
ed to play an important role along with ECM fungi in influencing tree productivity by
providing limiting nutrients (N, P, K, Ca, Mg) to the host plant (Frey-Klett et al., 2007;
Navarro-Ródenas et al., 2016). Moreover ECM roots are reported to select bacteria
with higher mineralising potential (Calvaruso et al., 2007; Kataoka et al., 2009; Kluber
et al., 2010; Uroz et al., 2007).
Many non–photosynthetic gametophyte and vascular plants, such as some liverworts
and lycophyte obtain all of their carbon from fungi (Bidartondo & Duckett, 2010). The
liverwort Cryptothallus mirabilis is epiparasitic and is specialized on Tulasnella species
that form ECM with surrounding trees. By using microcosm experiments it was
observed that the interaction with Tulasnella is necessary for growth of Cryptothallus
(Bidartondo et al., 2003). Diphasiastrum alpinum (Lycopodiaceae) gametophytes
colonised by ECM fungi and obtain all of their carbon and nutrition from Ericaceae
plant through ECM fungi (Horn et al., 2013).
ECM Role in Amelioration of Heavy Metal Stress:
There are number of heavy metals like Zn, Fe, Cu, and Mn which are otherwise
essential micronutrients for plant growth and a wide variety of cellular processes but
are reported to become toxic above threshold limit (Adeleke et al., 2012; Chen et al.,
2015; Ott et al., 2002). Other heavy metals in soil like Pb, Cd, As, Se, Cr, and Al are
biologically non essential and toxic to plant. Despite their toxic effect on plant growth,
heavy metals also influence the uptake and concentration of essential elements such as
P and N (Krznaric et al., 2009; Luo et al., 2014). It has been suggested several times
that microorganism exhibit higher tolerance to metal toxicity in comparison to
plants. At least some ECM fungi possess the ability to grow in and possibly
decompose such compounds and also impart protective effect to plants against
heavy metals via prevention of translocation of heavy metals into the host
(Table 2). Till date most studies have indicated that ECM plants accumulate less
metal inside their tissue and grow better than non- mycorrhizal plants, when
exposed to heavy metal stress (Adriaensen et al., 2004, 2005, 2006; Jourand
et al., 2010; Kayama & Yamanaka, 2014; Luo et al., 2014).
Fungal organic acids are reported to play an important role in binding heavy metals
thereby preventing their translocation in the host plant (Ahonen-Jonnarth et al., 2000;
Fomina et al., 2006). Hentschel et al. (1993) emphasized the importance of Lactarius
thiogalus, L. rufus, and Paxillus involutus in association with Picea abies for
mycoremediation of Al. Krupa and Kozdrój (2004) documented that the fruiting body
of fungi accumulated several times higher contents of Cd and Pb compared to those found
in the soil and five fold higher concentration of metals in mycorrhizal roots compared to
plants, indicating the importance of mycorrhizal fungi in forming an efficient biological
barrier for checking the movement of heavy metals into the host tissues.
Ahonen-Jonnarth and Finlay (2001) reported that Laccaria bicolor is a potential

ECM associate of Pinus sylvestris which prevent the uptake of Ni and Cd into
P. sylvestris in polluted soil. Paxillus involutus and Suillus bovinus associated with
Pinus sylvestris are also reported to prevent the uptake of Zn in contaminated soil
(Adriaensen et al., 2004; Fomina et al., 2006).The impact of association of Astraeus
hygrometricus and Scleroderma citrinum on the seedlings of Quercus glauca, Q.
saliciana and Castanopsis cuspidata in Al polluted soil was investigated by Kayama
and Yamanaka (2014). In the leaves of seedlings of C. cuspidata inoculated with
Astraeus hygrometricus and Scleroderma citrinum the amount of Al has been reported
to be higher in comparision to those of Quercus glauca and Q. saliciana indicating the
role of ECM fungi in protection of Q. glauca and Q. saliciana from heavy metal stress.
The cellular mechanisms involved in detoxification of heavy metals by mycorrhizal
fungi include biosorption of metals to fungal cell wall, chelation of metal ion in the
cytosol by compounds such as Glutathione and Metallothioneins, metal exclusion
mechanisms in metal-tolerant ECM fungi and the compartmentation of metals in the
vacuole, where metal ions are probably complexed in a chemically inactive form
(Colpaert et al., 2011; Daghino et al., 2016; Krpata et al., 2009; Luo et al., 2014).
Thus metal tolerant fungal isolates are reported to successfully colonize heavy metal
polluted soils and act as a better filter than non-tolerant ecotypes because the former are
known to more strongly prevent metal transfer to their host (Colpaert et al., 2005, 2011;
Luo et al., 2014)
Role of ECM in Disease Resistance to Plants:
It has been documented by several authors that mycorrhizal fungi improve disease
resistance of their host plant primarily by direct competition, enhanced or altered plant
growth, nutrition and morphology, induced resistance and development of antagonist
microbiota. Direct competition or inhibition is reported to be due to production and
release of antibiotics and physical sheathing by mantle of ECM (Blom et al., 2009;
Branzanti et al., 1999; Corcobado et al., 2015; Duchesne et al., 1988). Japanese red
pine (Pinus densiflora) inoculated with ECM fungi Suillus lubens and Rhizopogon
rubescens has been reported to result in improved growth of seedlings and decreased
seedlings mortality caused by pinewood nematode (Kikuchi et al., 1991). Similar
observation was made by Nakashima et al. (2016) while working on Japanese black
pine (Pinus thunbergii). Number of pathogenesis related (PR) proteins and enzymes
that help in defense are reported to be produced in ECM infected plants by number of
investigators (Guenoune et al., 2001; Guillon et al., 2002; Pfabel et al., 2012).
Pisolithus tinctorius, Laccaria laccata, Leucopaxillus cerealis and Suillus luteus are
reported to show antagonistic effect against pathogenic fungus Phytophthora
cinnamomi (Marx, 1972). Shrestha et al. (2005) investigated the antagonistic activity
of ECM fungi Pisolithus and Scleroderma against plant pathogen such as Pythium sp.,
Rhizoctonia solani, Fusarium sp., Agrobacterium tumifaciens, Klebsiella sp.,
Staphylococcus aureus, Shigella dysetriae and Escherichia coli. Mohan et al. (2015)
investigated the pathogenic effect of ECM fungi including Alnicola sp., Laccaria
fraterna, Lycoperdon perlatum, Pisolithus albus, Russula parazurea, Scleroderma
citrinum, Suillus brevipus, and S. subluteus against pathogenic fungus Alternaria
solani, Botrytis sp., Fusarium oxysporum, Phytophthora sp., Pythium sp.,
J. Kumar, N.S. Atri
Rhizoctonia solani, Sclerotium rolfsii, and Subramanispora vesiculosa. The antagonis-

tic effect was reported to be maximum in the ECM fungus Suillus brevipes (60.31%)
followed by S. subluteus (49.46%). Out of the pathogenic fungi employed for inves-
tigation, growth of Phytophthora sp. was reported to be highly inhibited by all 8
different ECM fungi.
Camilo-Alves et al. (2013) documented Phytophthora cinnamomi to be the main
biotic factor in Quercus ilex decline. This invasive pathogen has been reported to be
responsible for multiple fine root infections and tree death due to impairment of Q. ilex
water uptake and photosynthesis (Corcobado et al., 2013a, 2013b). ECM fungi have been
shown to protect trees from P. cinnamomi infection along with supporting their survival
and growth in comparison to non mycorrhizal seedlings (Azul et al., 2014; Branzanti
et al., 1999; Corcobado et al., 2015). Thus ECM fungi can also be used as fungicide in
nursery plantations for better growth, survival and establishment of seedlings.
Role of ECM in Drought and Salt Tolerance:
Symbiosis between plants and mycorrhizal fungi, have also been reported to enhance
drought and salt tolerance of their host plant (Breda et al., 2006; Luo et al., 2011, 2014;
Richard et al., 2011; Xu et al., 2016). Water has been documented as vital regulator of
plant growth, and mycorrhizal fungal colonisation of root tips has been reported to
significantly impact root water uptake by altering both apoplastic and symplastic
pathways (Lehto & Zwiazek, 2011; Smith & Read, 2008) and formation of more
lateral roots (Felten et al., 2009). Under drought stress, mycorrhizal symbiosis has
been documented to possess a remarkable capacity to alter hydraulic properties of plant
roots by altering symplastic pathways and by their impact on plant aquaporins (AQPs)
(Aroca et al., 2009; Dietz et al., 2011; Maurel & Plassard, 2011; Nehls & Dietz, 2014;
Xu et al., 2015). As reported by Xu et al. (2015) in case of Picea glauca seedlings
colonized by Laccaria bicolor, the fungal aquaporin (JQ585595) have significant
impact on root hydraulics under water stress.
Symbiosis between plants and ECM fungi has been documented to help plants to
cope with salt stress (Ishida et al., 2009; Luo et al., 2011, 2014; Richard et al., 2011).
While working with Poplar, Langenfeld-Heyser et al. (2007) documented the positive
effect of ECM on the host in increasing plant biomass and decreasing Na+
accumulation in poplar leaves. Similarly while working on roots under salt stress, Li
et al. (2012) reported that there is ECM mediated remodeling of ion flux which helps to
maintain K+ /Na+ homeostasis by increasing the release of Ca2+. ECM has also been
reported to change the plant phytohormone balance during salt stress. The presence of
Paxillus involutus as ECM partner has been associated with increased level of abscisic
acid and salicylic acid and decreased level of jasmonic acid and auxin in poplar roots in
comparison to non-mycorrhizal roots under water stress (Luo et al., 2009; Szuba, 2015).
ECM in Carbon Cycling and Decomposition of Organic Matter:
The ‘mycorrhizal decomposition theory’ of Frank seems to have been largely ignored
until the mid-1980s, when main emphasis was laid on the abilities of ectomycorrhizal
fungi to retrieve nitrogen (N) and phosphorus from plant litter. In such situations ECM
fungi are reported to act as facultative saprotrophs and benefit from organic matter
decomposition primarily through increased nitrogen mobilization rather than through

release of metabolic carbon (Lindahl & Tunlid, 2015; Hupperts et al., 2017). This view
was supported by the observation of Rineau et al. (2013) that organic matter transfor-
mation by Paxillus involutus in pure culture was stimulated rather than repressed by
glucose additions. In the study of Bodeker et al. (2014), organic matter degrading
enzyme activity has been reported to decline after the addition of ammonium because
organic N mobilization was suppressed in the presence of more easily accessible N.
Facultative saprotrophism has been reported to help mycorrhizal fungi in survival in the
absence of host plants or when host plants fail to supply enough carbon (Buee et al.,
2005; Courty et al., 2007). Hupperts et al. (2017) proposed two competing models to
explain carbon mobilization by ectomycorrhizal fungi. ‘Saprotrophy model’, where
decreased allocation of carbon may induce saprotrophic behaviour in ectomycorrhizal
fungi, resulting in the decomposition of organic matter to mobilize carbon and second
‘nutrient acquisition model’, where decomposition may instead be driven by the
acquisition of nutrients locked within soil organic matter compounds.
Hofrichter et al. (1999) demonstrated oxidation of synthetic lignin to CO2 in a cell-
free in vitro system. Martin et al. (2008) reported limited capacity of ECM fungi to
decompose plant litter. Some of the mycorrhizal mushrooms including Laccaria bicolor
(Martin et al., 2008), Tuber melanosporum (Martin et al., 2010) and Amanita species
(Wolfe et al., 2012) are reported to have lost most of their enzymes degrading plant cell
wall over a period of time. At the contrary, in some ECM fungi including Cortinarius
glaucopus the retention of a large number of Class II peroxidase genes has been reported
(Bodeker et al., 2014), which plays an important role in lignin degradation (Sinsabaugh,
2010). Furthermore, the potential ability to degrade lignin is more common in low-
biomass ectomycorrhizas as compared to high-biomass ectomycorrhizas (Hupperts
et al., 2017). Thus, Cortinarius and some other ECM species are documented to have
retained the capacity of their Agaricales ancestors, such as white-rot fungi (Matheny
et al., 2006; Shah et al., 2016), able to enzymatically oxidize and decompose phenolic
macromolecules, and especialiy the lignin. Indeed, peroxidase activity in soils has been
found to positively correlate with the species richness and relative abundance of ECM
fungi (Phillips et al., 2014; Talbot et al., 2013).
Hupperts et al. (2017) tested whether phenology-induced shifts in carbon reserves of
fine roots of aspen (Populus tremuloides) affect potential activity of four carbon-
compound d egrading enzymes, β -g luc uronidase, β-glucosidase, N -
acetylglucosaminidase and laccase, by ectomycorrhizal fungi. Ectomycorrhizal roots
from mature aspen were collected and analysed during tree dormancy, leaf flush, full
leaf expansion and leaf abscission and observed that extracellular enzyme activity to be
highest when root carbon reserves were lowest and root carbon reserves were positively
correlated with invertase of plant, suggesting that phenology may affect carbon allo-
cation to ECM fungi.
Conclusions
Ectomycorrhizal fungi represent an important component of soil microbiota and

associate with roots of several and abundant tree species. ECM associations are formed
by a restricted group of higher plant families such as Pinaceae, Betulaceae, Fabaceae,
J. Kumar, N.S. Atri
Dipterocarpaceae, Fagaceae and Myrtaceae with fungal symbionts belonging to the

Basidiomycota and Ascomycota. There are about 80 phylogenetic lineages of ECM
fungal species that are reported to have independently evolved from saprotrophic
ancestors (Tedersoo & Smith, 2013). ECM association is the result of coevolution
between plant and fungi which show mycorrhiza to be essential in terrestrial plant
nutrition and play a significant role in survival and growth of associated trees. It is now
established fact that mycorrhizal inoculation is beneficial for reclamation of variety of
disturbed sites and had great potential in restoration of natural ecosystem. Most studies
on ECM have been carried out in temperate and boreal forests and the associated fungi
mostly belong to Basidiomycota, especially Russuleceae.
Earlier studies on ECM fungal communities were carried out using
morphoanatomical characters, however presently molecular techniques are being
employed which along with morphoanatomical features helps to precisely understand
the ECM communities structure and diversity. Present day molecular tools alongwith
standard PCR analysis and transcriptome analysis such as oligoarray based tran-
scriptome profiling are being used so as to understand the molecular mechanisms of
ECM influence on host plant including N and P acquisition and carbohydrate transport.
Investigation done so far in this regard clearly suggest that plants inoculated with ECM
fungi have greater biomass and survival rate than those without associated mycorrhizal
partner, or those growing only with fertilizers. NGS has changed the face of microbial
ecology, and have gained unprecedented insight into the community dynamics and
biogeography of cryptic ECM species.
Despite recent advances in understanding community ecology of ECM fungi, little is
known about spatial distribution patterns of ECM fungal communities from local to
global scales. However, using population genetics and phylogenetic tools based on a
group of closely related species or within biological species, spatial and phylogenetic
scale of ECM fungal biogeography needs to be further explored. There are many areas
which remain to be explored on ECM interactions and their dynamic. Accurate
mycological data in man-made, native, or disturbed forests are required to better
manage plant and fungi biodiversity. We need to work on the complete ECM
microbiome so as to unearth information of all fungi and bacteria associated with plant
roots and magnitude of infection amongst them. Increased taxonomic efforts especially
in tropical and subtropical forests are required in this regard where not much informa-
tion is available. The development and identification of new strains able to increase
forest productivity is yet another area which requires attention. Along with molecular
investigation of ECM and host species, scope of application of proteomics and
metabolomics needs to be investigated with a view to elucidate the biochemistry
underlying this ECM related symbiosis. Coevolutionary processes between plants
and mycorrhizal fungi are also still poorly understood along with the effect of ECM
responses to expected future climate change and its consequences: these are some of the
areas which need to be thoroughly investigated.
Acknowledgements We are grateful to Mélanie Roy and another anonymous reviewer for their useful
comments on the manuscript. Thanks are due to Council of Scientific and Industrial Research (CSIR), New
Delhi, India for financial assistance under CSIR-JRF fellowship scheme to the first author. To University
Grants Commission we are indebted for giving liberal grants to the department under SAP programme and to
Department of Biotechnology, Government of India, for grant under IPLS programme.
Literature Cited
Abarenkov, K., R.H. Nilsson, K.H. Larsson, I.J. Alexander, U. Eberhardt, S. Erland et al. 2010. The
UNITE database for molecular identification of fungi-recent updates and future perspectives. New
Phytologist 186: 281–285. https://doi.org/10.1111/j.1469-8137.2009.03160.x
Abuzinadah, R.A., & D.J. Read. 1986. The role of proteins in the nitrogen nutrition of ectomycorrhizal
plants I. Utilization of peptides and proteins by ectomycorrhizal fungi. New Phytologist 103: 481-493.
https://doi.org/10.1111/j.1469-8137.1986.tb02886.x
Adeleke, R.A., T.E. Cloete, A. Bertrand & D.P. Khasa. 2012. Iron ore weathering potentials of
ectomycorrhizal plants. Mycorrhiza 22: 535–544. https://doi.org/10.1007/s00572-012-0431-5
Adriaensen, K., D. van der Lelie, A. Van Laere, J. Vangronsveld & J.V. Colpaert. 2004. A zinc-adapted
fungus protects pines from zinc stress. New Phytologist 161:549–555. https://doi.org/10.1046/j.1469-
8137.2003.00941.x
———, T. Vrålstad, J. P. Noben, J. Vangronsveld & J. V. Colpaert. 2005. Copper-adapted Suillus luteus, a
symbiotic solution for pines colonizing Cu mine spoils. Applied and Environmental Microbiology 71:
7279–7284. https://doi.org/10.1128/AEM.71.11.7279-7284.2005
———, J. Vangronsveld, & J.V. Colpaert. 2006. Zinc tolerant Suillus bovinus improves growth of Zn-
exposed Pinus sylvestris seedlings. Mycorrhiza 16: 553–558. https://doi.org/10.1007/s00572-006-0072-7
Agerer, R. 1986. Studies on ectomycorrhizae III. Mycorrhizae formed by four fungi in the genera Lactarius
and Russula on spruce. Mycotaxon 27: 1-59.
———. 1987-2012. Colour atlas of ectomycorrhizae. 1st – 15th ed.-Einhorn-Verlag. Schwäbisch Gmünd,
Germany.
———. 1999. Anatomical charecterstics of identified ectomycorrhizas: an attempt towards a natural classi-
fication. In: A.Verma, B. Hock (Eds.), mycorrhiza. Springer Verlag, Berlin, pp. 685-734.
———. 2001. Exploration types of ectomycorrhizae- A proposal to classify ectomycorrhizal mycelial
systems according to their paterns of differentiation and putative ecological importance.
Mycorrhiza 11: 107-114.
——— . 2006. Fungal relationships and structural identity of their ectomycorrhizae. Mycological Progress 5:
67-107. https://doi.org/10.1007/s11557-006-0505-x
——— & G. Rambold. 2004-2016. DEEMY-An Information System for Characterization and Determination
of Ectomycorrhizae. www.deemy.de - München, Germany.
Ahangar, M.A., G.H. Dar & Z.A. Bhat. 2012. Growth response and nutrient uptake of blue pine (Pinus
wallichiana) seedlings inoculated with rhizosphere microorganisms under temperate nursery conditions.
Annals of Forest Research 55(2): 217-227.
Ahonen-Jonnarth, U. & R.D. Finlay. 2001. Effects of elevated nickel and cadmium concentrations on
growth and nutrient uptake of mycorrhizal and non-mycorrhizal Pinus sylvestris seedlings. Plant Soil 236:
129-138. https://doi.org/10.1023/A:1012764902915
———, P.A.W. van Hees, U. Lundström & R.D. Finlay. 2000. Production of organic acids by mycorrhizal
and non-mycorrhizal Pinus sylvestris exposed to elevated concentrations of aluminium and heavy metals.
New Phytologist 146: 557-567. https://doi.org/10.1046/j.1469-8137.2000.00653.x
———, A. Göransson & R.D. Finlay. 2003. Growth and nutrient uptake of ectomycorrhizal Pinus sylvestris
seedlings in a natural substrate treated with elevated Al concentrations. Tree Physiology 23: 157–167.
https://doi.org/10.1093/treephys/23.3.157
Alberton, O., D. Aguiar, R.M.T. Gimenes & R. Carrenho. 2014. Meta-analysis for responses of eucalyptus
and pine inoculated with ectomycorrhizal fungi in Brazil. Journal of Food Agriculture and Environment
12: 1159–1163.
Alexander, I.J. 2006. Ectomycorrhizas – out of Africa? New Phytologist 172: 589–591.
——— & M.A. Selosse. 2009. Mycorrhizas in tropical forests: a neglected research imperative. New
Phytologist 182 (1): 14-16. https://doi.org/10.1111/j.1469-8137.2009.02798.x
Allen, M.F. 2007. Mycorrhizal fungi: highways for water and nutrients in arid soils. Vadose Zone Journal 6:
291–297. https://doi.org/10.2136/vzj2006.0068
Alvez, L., V.L. Oliveira & G.N.S. Filho. 2010. Utilization of rocks and ectomycorrhizal fungi to promote
growth of eucalypt. Brazilian Journal of Microbiology 41: 676–684. https://doi.org/10.1590/s1517-
83822010000300018
Andersson, S., P. Jensén & B. Söderström. 1996. Effects of mycorrhizal colonization by Paxillus involutus
on uptake of Ca and P by Picea abies and Betula pendula grown in unlimed and limed peat. New
Phytologist 133: 695-704. https://doi.org/10.1111/j.1469-8137.1996.tb01938.x
J. Kumar, N.S. Atri
Andrew, C. & E.A. Lilleskov. 2009. Productivity and community structure of ectomycorrhizal fungal
sporocarps under increased atmospheric CO2 and O3. Ecology Letters 12(8): 813–822. https://doi.
org/10.1111/j.1461-0248.2009.01334.x
——— & ———. 2014. Elevated CO2 and O3 effects on ectomycorrhizal fungal root tip communities in
consideration of a post-agricultural soil nutrient gradient legacy. Mycorrhiza 24: 581–593. https://doi.
org/10.1007/s00572-014-0577-4
Arnebrant, K. 1994. Nitrogen amendments reduce the growth of extramatrical ectomycorrhizal mycelium.
Mycorrhiza 5: 7–15. https://doi.org/10.1007/BF00204014
Aroca, R., A. Bago, M. Sutka, J.A. Paz, C. Cano, G. Amodeo et al. 2009. Expression analysis of the first
arbuscular mycorrhizal fungi aquaporin described reveals concerted gene expression between salt-
stressed and nonstressed mycelium. Molecular Plant-Microbe Interaction 22(9): 1169-1178. https://doi.
org/10.1094/MPMI-22-9-1169
Ashford, A.E., P.A. Vesk, D.A. Orlovich, A.L. Markovina & W.G. Allaway. 1999. Dispersed
polyphosphate in fungal vacuoles in Eucalyptus pilularis/Pisolithus tinctorius ectomycorrhizas. Fungal
Genetics and Biology 28: 21-33. https://doi.org/10.1006/fgbi.1999.1140.
Atri, N.S. & S.S. Saini. 1986. Further contribution on the studies of North-West Himalyan Russulaceae.
Geobios new reports 5: 100-105.
———, ——— & M.K. Saini. 1997. Studies on genus Russula Pers. from North Western Himalayas.
Mushroom Research 6(1): 1-6.
Averill, C., B.L. Turner & A.C. Finzi. 2014. Mycorrhiza-mediated competition between plants and
decomposers drives soil carbon storage. Nature 505: 543–545. https://doi.org/10.1038/nature12901
Avis, P.G. 2012. Ectomycorrhizal iconoclasts: the ITSrDNA diversity and nitrophilic tendencies of fetid
Russula. Mycologia 104 (5): 998–1007. https://doi.org/10.3852/11-399
———, G.M. Mueller & J. Lussenhop. 2008. Ectomycorrhizal fungal communities in two North American
oak forests respond to nitrogen addition. New Phytologist 179: 472–483. https://doi.org/10.1111/j.1469-
8137.2008.02491.x
Azul, A.M., J. Nunes, I. Ferreira, A.S. Coelho, P. Veríssimo, J. Trovão, A. Campos, P. Castro & H.
Freitas. 2014. Valuing native ectomycorrhizal fungi as a mediterranean forestry component for sustain-
able and innovative solutions. Botany 92: 161–171. https://doi.org/10.1139/cjb-2013-0170
Bâ, A.M., K.B. Sanon & R. Duponnois. 2002. Influence of ectomycorrhizal inoculation Afzelia quanzensis
Welw. seedlings in a nutrient-deficient soil. Forest Ecology and Management 161:215–219. https://doi.
org/10.1016/S0378-1127(01)00484-4
———, R. Duponnois, M. Diabaté & B. Dreyfus. 2011. Les champignons ectomycorhiziens des arbres
forestiers en Afrique de l’Ouest. Institute of Research for Development, Montpellier, pp. 252.
———, ———, R. B. Moyersoen & A.G. Diédhiou. 2012. Ectomycorrhizal symbiosis of tropical African
trees. Mycorrhiza 22:1–29. https://doi.org/10.1007/s00572-011-0415-x
Bahram, M., S. Põlme, U. Kõljalg & L. Tedersoo. 2011. A single European aspen (Populus tremula) tree
individual may potentially harbor dozens of Cenococcum geophilum ITS genotypes and hundreds of
species of ectomycorrhizal fungi. FEMS Microbiology Ecology 75: 313–320. https://doi.org/10.1111
/j.1574-6941.2010.01000.x
———, U Kõljalg, P. Kohout, S. Mirshahvaladi & L. Tedersoo. 2013. Ectomycorrhizal fungi of exotic
pine plantations in relation to native host trees in Iran: evidence of host range expansion by local
symbionts to distantly related host taxa. Mycorrhiza 23: 11–19. https://doi.org/10.1007/s00572-012-
0445-z
Barrow, N.J. 1984. Modelling the effects of pH on phosphate sorption by soils. Journal of Soil Science 35(2):
283–297. https://doi.org/10.1111/j.1365-2389.1984.tb00283.x
Becerra, A.G. & M.R. Zak. 2011. The ectomycorrhizal symbiosis in South America: Morphology,
Colonisation and Diversity. In: M. Rai & A. Verma (Eds.), Diversity and Biotechnology of
ectomycorrhizae, springer-verlag Berlin Heidelberg, pp. 19-41.
Bendangmenla & T. Ajungla. 2014. Field performance of Schima wallichii (dc.) korth. seedlings inoculated
with ectomycorrhizal fungi. International Journal of Recent Scientific Research 5(4): 970-973.
Benito, B. & M. Gonzalez-Guerrero. 2014. Unravelling potassium nutrition in ectomycorrhizal associations.
New Phytologist 201: 707–709. https://doi.org/10.1111/nph.12659
Benjdia, M., E. Rikirsch, T. Müller, M. Morel, C. Corratgé, S. Zimmermann, M. Chalot, W.B. Frommer
& D. Wipf. 2006. Peptide uptake in the ectomycorrhizal fungus Hebeloma cylindrosporum: character-
ization of two di- and tripeptide transporters (HcPTR2A and B). New Phytologist 170: 401–410.
https://doi.org/10.1111/j.1469-8137.2006.01672.x
Bent, E., P. Kiekel, R. Brenton & D.L. Taylor. 2011. Root-associated ectomycorrhizal fungi shared by
various boreal forest seedlings naturally regenerating after a fire in interior Alaska and correlation of
different fungi with host growth responses. Applied and Environmental Microbiology 77: 3351–3359.
https://doi.org/10.1128/AEM.02575-10
Benucci, G.M.N., C. Lefevre & G. Bonito. 2016. Characterizing root-associated fungal communities and
soils of Douglas-fir (Pseudotsuga menziesii) stands that naturally produce Oregon white truffles (Tuber
oregonense and Tuber gibbosum). Mycorrhiza 26: 367–376. https://doi.org/10.1007/s00572-015-0677-9
Berbee, M.L. & J.W. Taylor. 1993. Dating the evolutionary radiations of the true fungi. Canadian Journal of
Botany 71: 1114–1127. https://doi.org/10.1139/b93-131
——— & ———. 2001. Fungal molecular evolution: gene trees and geologic time, in: D. McLaughlin, E.
McLaughlin, & P. Lemke (Eds.), The Mycota. Springer-Verlag, Berlin, pp. 229–245.
——— & ———. 2010. Dating the molecular clock in fungi – how close are we? fungal biology reviews 24:
1–16. https://doi.org/10.1016/j.fbr.2010.03.001
Bhatt, R.P. & T.N. Lakhanpal. 1990. Fleshy fungi of North Western Himalayas V. Indian Phytopathology
43: 156- 164.
Bidartondo, M.I. & J.G. Duckett. 2010. Conservative ecological and evolutionary patterns in liverwort-
fungal symbioses. Proceedings of the Royal Society B. 277: 485–492. https://doi.org/10.1098
/rspb.2009.1458
———, T.D. Bruns, M. Weiss, C. Sérgio & D.J. Read. 2003. Specialized cheating of the ectomycorrhizal
symbiosis by an epiparasitic liverwort. Proceedings of the Royal Society B. 270: 835–842. https://doi.
org/10.1098/rspb.2002.2299
Blom, J.M., A. Vannini, A.M. Vettraino, M.D. Hale & D.L. Godbold. 2009. Ectomycorrhizal community
structure in a healthy and a Phytophthora-infected chestnut (Castanea sativa Mill.) stand in central Italy.
Mycorrhiza 20: 25–38. https://doi.org/10.1007/s00572-009-0256-z
Bodeker, I.T.M., K.E. Clemmensen, W. de Boer, F. Martin, A. Olson & B.D. Lindahl. 2014.
Ectomycorrhizal Cortinarius species participate in enzymatic oxidation of humus in northern forest
ecosystems. New Phytologist 203: 245–256. https://doi.org/10.1111/nph.12791
Bois, G., Y. Piché, M.Y.P. Fung & D.P. Khasa. 2005. Mycorrhizal inoculum potentials of pure reclamation
materials and revegetated tailing sands from the Canadian oil sand industry. Mycorrhiza 15: 149–158.
https://doi.org/10.1007/s00572-004-0315-4
Bonfante, P. & A. Genre. 2010. Mechanisms underlying beneficial plant–fungus interactions in mycorrhizal
symbiosis. Nature Communication 1: 48. https://doi.org/10.1038/ncomms1046
Bougher, N.L., T.S. Grove & N. Malajczuk. 1990. Growth and phosphorus acquisition of Karri (Eucalyptus
diversicolor F. Muell.) seedlings inoculated with ectomycorrhizal fungi in relation to phosphorus supply.
New Phytologist 114: 77-85. https://doi.org/10.1111/j.1469-8137.199 0.tb 00376.x
Brandes, B., D.L. Godbold, A.J. Kuhn & G. Jentschke. 1998. Nitrogen and phosphorus acquisition by the
mycelium of the ectomycorrhizal fungus Paxillus involutus and its effect on host nutrition. New
Branzanti, M.B., E. Rocca & A. Pisi. 1999. Effect of ectomycorrhizal fungi on chestnut ink disease.
Mycorrhiza 9: 103-109. https://doi.org/10.1007/s005720050007
Breda, N., R. Huc, A. Granier & E. Dreyer. 2006. Temperate forest trees and stands under severe drought: a
review of ecophysiological responses, adaptation processes and long-term consequences. Annals of Forest
Science 6: 625–644. https://doi.org/10.1051/forest:2006042
Browning, M.H.R. & R.D. Whitney. 1993. BInfection of containerized jack pine and black spruce by
Laccaria species and Thelephora terrestris and seedlings survival and growth after out planting^.
Canadian Journal of Forest Research 23: 330-333. https://doi.org/10.1139/x93-046
Brulé, C., P. Frey-Klett, J.C. Pierrat, S. Courrier, F. Gerard, M.C. Lemoine, J.L. Rousselet, G. Sommer
& J. Garbaye. 2001. Survival in the soil of the ectomycorrhizal fungus Laccaria bicolor and the effects
of a mycorrhiza helper Pseudomonas fluorescens. Soil Biology and Biochemistry 33: 1683-1694.
https://doi.org/10.1016/S0038-0717(01)00090-6
Brundrett, M.C. 2002. Coevolution of roots and mycorrhizas of land plants. New Phytologist 154: 275-304.
https://doi.org/10.1046/j.1469-8137.2002.00397.x
———. 2009. Mycorrhizal associations and other means of nutrition of vascular plants: understanding the
global diversity of host plants by resolving conflicting information and developing reliable means of
diagnosis. Plant Soil 320: 37–77. https://doi.org/10.1007/s11104-008-9877-9
Bruns, T.D. 1995. Thoughts on the processes that maintain local species diversity of ectomycorrhizal fungi.
Plant Soil 170: 63–73. https://doi.org/10.1007/BF02183055
———, M.I. Bidartondo & D.L. Taylor. 2002. Host specificity in ectomycorrhizal communities: what do
the exceptions tell us? Integrative and Comparative Biology 42: 352–359. https://doi.org/10.1093
/icb/42.2.352
J. Kumar, N.S. Atri
Bucher, M. 2007. Functional biology of plant phosphate uptake at root and mycorrhiza interfaces. New
Bücking, H. & W. Heyser. 2001. Microrautoradiographic localization of phosphate and carbohydrate in
mycorrhizal roots of Populus tremela x Populus alba and the implications for transfer processes in
ectomycorrhizal associations. Tree Physiology 21:101–107.
——— & ———. 2003. Uptake and transfer of nutrients in ectomycorrhizal Associations: Interactions
between Photosynthesis and Phosphate Nutrition. Mycorrhiza 13(2): 59-68. https://doi.org/10.1007
/s00572-002-0196-3
Buee, M., D. Vairelles & J. Garbaye. 2005. Year-round monitoring of diversity and potential metabolic
activity of the ectomycorrhizal community in a beech (Fagus silvatica) forest subjected to two thinning
regimes. Mycorrhiza 15: 235– 245. https://doi.org/10.1007/s00572-004-0313-6
Cairney, J.W.G. 2011. Ectomycorrhizal fungi: the symbiotic route to the root for phosphorus in forest soils.
Plant Soil 344: 51–71. https://doi.org/10.1007/s11104-011-0731-0
———. 2012. Extramatrical mycelia of ectomycorrhizal fungi as moderators of carbon dynamics in forest
soil. Soil Biology and Biochemistry 47: 198–208. https://doi.org/10.1016/j.soilbio.2011.12.029
Calvaruso, C., M.P. Turpault, E. Leclerc & P. Frey-Klett. 2007. Impact of ectomycorrhizosphere on the
functional diversity of soil bacterial and fungal communities from a forest stand in relation to nutrient
mobilization processes. Microbial Ecology 54: 567–577. https://doi.org/10.1007/ s002 48-007-9260-z
Camilo-Alves, C.S.P., M.I.E. Clara & N.M.C.A. Ribeiro. 2013. Decline of Mediterranean oak trees and its
association with Phytophthora cinnamomi: a review. European Journal of Forest Research 132: 411–432.
https://doi.org/10.1007/s10342-013-0688-z
Canton, G.C., A.A. Bertolazi, A.J. Cogo, F.J. Eutriópio, J. Melo, S.B. de Souza et al. 2016. Biochemical
and ecophysiological responses to manganese stress by ectomycorrhizal fungus Pisolithus tinctorius and
in association with Eucalyptus grandis. Mycorrhiza 26: 475–487. https://doi.org/10.1007/s00572-016-
0686-3
Casieri, L., N.A. Lahmidi, J. Doidy, C. Veneault-Fourrey, A. Migeon, L. Bonneau, P.E. Courty, K.
Garcia, M. Charbonnier & A. Delteil. 2013. Biotrophic transportome in mutualistic plant–fungal
interactions. Mycorrhiza 23: 597–625. https://doi.org/10.1007/s00572-013-0496-9
Castellano, M.A. & R. Molina. 1989. Mycorrhizae. In: T.D. Landis, R.W. Tinus, S.E. McDonald, J.P. Barnett
(Eds.), The Biological Components: Nursery Pests and Mycorrhizae – The Container Tree Nursery
Manual, US Department of Agriculture, Forest Service, Washington, DC, USA, pp. 101–167.
Chambers, S.M., P.G. Williams, R.D. Seppelt, J.W.G. Cairney. 1999. Molecular identification of
Hymenoscyphus sp. from rhizoids of the leafy liverwort Cephaloziella exiliflora in Australia and
Antarctica. Mycological Research 103:286–288. https://doi.org/10.1017/S0953756298007217
Chen, Y.L., M.C. Brundrett & B. Dell. 2000. Effects of ectomycorrhizas and vesicular-arbuscular mycor-
rhizas, alone and in competition, on root colonization and growth of Eucalyptus globulus and E.
urophylla. New Phytologist 146: 545–556. https://doi.org/10.1046/j.1469-8137.2000.00663.x
———, K. Nara, Z. Wen, L. Shi, Y. Xia, Z. Shen & C. Lian. 2015. Growth and photosynthetic responses of
ectomycorrhizal pine seedlings exposed to elevated Cu in soils. Mycorrhiza 25: 561–571. https://doi.
org/10.1007/s00572-015-0629-4
Colpaert, J.V. & J.A. Van Assche. 1993. The effects of cadmium on ectomycorrhizal Pinus sylvestris L. New
Phytologist 123: 325-333. https://doi.org/10.1111/j.1469-8137.1993.tb03742.x
———, K.K. Van Tichelen, J.A. Van Assche & A. Van Laere. 1999. Short-term phosphorus uptake rates in
mycorrhizal and non-mycorrhizal roots of intact Pinus sylvestris seedlings. New Phytologist 143: 589–
597. https://doi.org/10.1046/j.1469-8137.1999.0047.x
———, K. Adriaensen, L.A.H. Muller, M. Lambaerts, C. Faes, R. Carleer & J. Vangronsveld. 2005.
Element profiles and growth in Zn-sensitive and Zn-resistant Suilloid fungi. Mycorrhiza 15: 628–634.
https://doi.org/10.1007/s00572-005-0009-6
———, J.H.L. Wevers, E. Krznaric & K. Adriaensen. 2011. How metal-tolerant ecotypes of ecto-
mycorrhizal fungi protect plants from heavy metal pollution. Annals of Forest Science 68: 17–24.
https://doi.org/10.1007/s13595-010-0003-9
Corcobado, T., E. Cubera, G. Moreno, & A. Solla. 2013a. Quercus ilex forests are influenced by annual
variations in water table, soil water deficit and fine root loss caused by Phytophthora cinnamomi.
Agriculture and Forest Meteorology 169: 92–99. https://doi.org/10.1016/j.agrformet.2012.09.017
———, A. Solla, M.A. Madeira & G. Moreno. 2013b. Combined effects of soil properties and
Phytophthora cinnamomi infections on Quercus ilex decline. Plant Soil 373: 403–413. https://doi.
org/10.1007/s11104-013-1804-z
———, G. Moreno, A.M. Azul & A. Solla. 2015. Seasonal variations of ectomycorrhizal commu-
nities in declining Quercus ilex forests: interactions with topography, tree health status and
Phytophthora cinnamomi infections. Forestry 88: 257 –266. https://doi.org/10.1093/ forestry/

cpu056
Corrales, A., A.E. Arnold, A. Ferrer, B.L. Turner & J.W. Dalling. 2016. Variationin ectomycorrhizal
fungal communities associated with Oreomunnea mexicana (Juglandaceae) in a Neotropical montane
forest. Mycorrhiza 26: 1–17. https://doi.org/10.1007/s00572-015-0641-8
Courty, P.E., N. Breda & J. Garbaye. 2007. Relation between oak tree phenology and the secretion of
organic matter degrading enzymes by Lactarius quietus ectomycorrhizas before and during bud break.
Soil Biology and Biochemistry 39: 1655–1663. https://doi.org/10.1016/j.soilbio.2007.01.017
Couturier, C., B. Montanini, F. Martin, A. Brun, D. Blaudez & M. Chalot. 2007. The expanded family of
ammonium transporters in the perennial poplar plant. New Phytologist 174: 137–150. https://doi.
org/10.1111/j.1469-8137.2007.01992.x
Cox, F., N. Barsoum, E.A. Lilleskov & M.I. Bidartondo. 2010. Nitrogen availability is a primary
determinant of conifer mycorrhizas across complex environmental gradients. Ecology Letter 13: 1103–
1113. https://doi.org/10.1111/j.1461-0248.2010.01494.x
Crane, S., T. Barkay & J. Dighton. 2010. Growth responses to and accumulation of mercury by
ectomycorrhizal fungi. Fungal Biology 114: 873-880. https://doi.org/10.1016/j.funbio.2010.08.004.
Cullings, K.W., T.M. Szaro & T.D. Bruns. 1996. Evolution of extreme specialization within a lineage of
ectomycorrhizal epiparasites. Nature 379, 63–67. https://doi.org/10.1038/379063a0
Cumming, J.R. & L.H. Weinstein. 1990. Nitrogen source effects on Al toxicity in nonmycorrhizal and
myconhizal pitch pine (Pinus rigida) seedlings Growth and nutrition. Canadian Journal of Botany 68:
264-2652. https://doi.org/10.1139/b90-334
——— . 1996. Phosphate-limitation physiology in ectomycorrhizal pitch pine (Pinus rigida) seedlings. Tree
Physiology 16: 977- 983. https://doi.org/10.1093/treephys/16.11-12.977
———, C. Zawaski, S. Desai & F.R. Collart. 2015. Phosphorus disequilibrium in the tripartite plant
ectomycorrhiza-plant growth promoting rhizobacterial association. Journal of Soil Science and Plant
Nutrition 15 (2): 464-485. https://doi.org/10.4067/S0718-95162015005000040
Daghino, S., E. Martino & S. Perotto. 2016. Model systems to unravel the molecular mechanisms of heavy
metal tolerance in the ericoid mycorrhizal symbiosis. Mycorrhiza 26: 263–274. https://doi.org/10.1007
/s00572-015-0675-y
Dahlberg, A. 2001. Community ecology of ectomycorrhizal fungi: an advancing interdisciplinary field. New
———. 2002. Effects of fire on ectomycorrhizal fungi in Fennoscandian boreal forests. Silva Fennica 36: 69–
80. https://doi.org/10.14214/sf.551
Danielsen, L., G. Lohaus, A. Sirrenberg, P. Karlovsky, C. Bastien, G. Pilate & A. Polle. 2013.
Ectomycorrhizal colonization and diversity in relation to tree biomass and nutrition in a plantation of
transgenic poplars with modified lignin biosynthesis. PLoS ONE 8(3): e59207. https://doi.org/10.1371
/journal.pone.0059207
Danielson, R.M. & S. Visser. 1989. Host response to inoculation and behaviour of induced and indigenous
ectomycorrhizal fungi of jack pine grown on oil-sands tailings. Canadian Journal of Forest Research 19:
1412–1421.
Dar, G.H., M.A. Beig & N.A. Ganai. 2007. Effect of source and inoculum load of ectomycorrhizae on the
growth and biomass of containerized kail pine (Pinus wallichiana) seedlings. Applied Biological
Research 9:19-28.
de Campos, M.C., S.J. Pearse, R.S. Oliveira, & H. Lambers. 2013. Viminaria juncea does not vary its
shoot phosphorus concentration and only marginally decreases its mycorrhizal colonization and cluster-
root dry weight under a wide range of phosphorus supplies. Annals of Botany 111: 801–809. https://doi.
org/10.1093/aob/mct035
DeBellis, T., G. Kernaghan, R. Bradley & P. Widden. 2006. Relationships between stand composition and
ectomycorrhizal community structure in boreal mixed-wood forests. Microbial Ecology 52:114-126.
https://doi.org/10.1007/s00248-006-9038-8
Desai, S., D. Naik & J.R. Cumming. 2014. The influence of phosphorus availability and Laccaria bicolor
symbiosis on phosphate acquisition, antioxidant enzyme activity, and rhizospheric carbon flux in Populus
tremuloides. Mycorrhiza 24: 369–382. https://doi.org/10.1007/s00572-013-0548-1
Deveau, A., S. Antony-Babu, F. Le Tacon, C. Robin, P. Frey-Klett & S. Uroz. 2016. Temporal changes of
bacterial communities in the Tuber melanosporum ectomycorhizosphere during ascocarp development.
Mycorrhiza 26: 389–399. https://doi.org/10.1007/s00572-015-0679-7
Diagne, N., J. Thioulouse, H. Sanguin, Y. Prin, T. Krasova-Wade & S. Sylla et al. 2013. Ectomycorrhizal
diversity enhances growth and nitrogen fixation of Acacia mangium seedlings. Soil Biology and
Biochemistry 57: 468–476. https://doi.org/10.1016/j.soilbio. 2012.08.030
J. Kumar, N.S. Atri
Dickie, I.A. 2007. Host preference, niches and fungal diversity. New Phytologist 174: 230–233. https://doi.
org/10.1111/j.1469-8137.2007.02055.x
———, S.J. Richardson & S.K. Wiser. 2009. Ectomycorrhizal fungal communities in two temperate
Nothofagus rainforests respond to changes in soil chemistry after small-scale timber harvesting.
Canadian Journal of Forest Research 39: 1069–1079. https://doi.org/10.1139/X09-036
———, N. Bolstridge, J.A. Cooper & D.A. Peltzer. 2010. Co-invasion by Pinus and its mycorrhizal fungi.
New Phytologist 187: 475–484. https://doi.org/10.1111/j.1469-8137.2010.03277.x
———, L.B. Martínez-García, N. Koele, G-A. Grelet, J.M. Tylianakis, D.A. Peltzer & S.J. Richardson.
2013. Mycorrhizas and mycorrhizal fungal communities throughout ecosystem development. Plant Soil
367: 11–39. https://doi.org/10.1007/s11104-013-1609-0
Diédhiou, A.G., M.A. Selosse, A. Galiana, M. Diabate, B. Dreyfus, A.M. Ba, S. Miana de Faria, & G.
Bena. 2010. Multihost ectomycorrhizal fungi are predominant in a Guinean tropical rainforest and shared
between canopy trees and seedlings. Environmental Microbiology 12: 2219–2232. https://doi.org/10.1111
/j.1462-2920.2010.02183.x
———, H. Christelle, M. Ebenye, M.A. Selosse, N. Onguene & A.M. Bâ. 2014. Diversity and community
structure of ectomycorrhizal fungi in mixed and monodominant African tropical rainforest. In: A.M. Bâ,
K.L. McGuire, A.G. Diédhiou (Eds.), Ectomycorrhizal symbioses in tropical and neotropical forests.
CRC Press, pp 1–18.
Dietz, S., J. von Bülow, E. Beitz & U. Nehls. 2011. The aquaporin gene family of the ectomycorrhizal fungus
Laccaria bicolor: lessons for symbiotic functions. New Phytologist 190: 927–940. https://doi.org/10.1111
/j.1469-8137.2011.03651.x
Domínguez Núñez, J.A., J. Selva Serrano, J.A. Rodríguez Barreal & J.A. Saiz de Omeñaca González.
2006. The influence of mycorrhization with Tuber melanosporum in the afforestation of a Mediterranean
site with Quercus ilex and Quercus faginea. Forest Ecology and Management 231: 226–233. https://doi.
org/10.1016/j.foreco.2006.05.052
———, R.G. Planelles, J.A. Rodriguez Barreal & J.A. Saiz de Omenaca Gonzalez. 2008. The effect of
Tuber melanosporum Vitt. mycorrhization on growth, nutrition, and water relations of Quercus petraea
Liebl., Quercus faginea Lamk., and Pinus halepensis Mill. seedlings. New Forest 2: 159–171. https://doi.
org/10.1007/s11056-007-9069-0
———, A. Martin, A. Anriquez & A. Albanesi. 2012. The combined effects of Pseudomonas fluorescens
and Tuber melanosporum on the quality of Pinus halepensis seedlings. Mycorrhiza 22: 429–436.
https://doi.org/10.1007/s00572-01 1-0420-0
Druebert, C., C. Lang, K. Valtanen & A. Polle. 2009. Beech carbon productivity as driver of
ectomycorrhizal abundance and diversity. Plant Cell and Environment 32(8): 992-1003. https://doi.
org/10.1111/j.1365-3040.2009.01983.x
Duchesne, L.C., R.L. Peterson & B.E. Ellis. 1988. Pine root exudate stimulates antibiotic synthesis by the
ectomycorrhizal fungi Paxillus involutus. New Phytologist 108: 471-476. https://doi.org/10.1111/j.14 69-
8137.1988.tb04188.x
Duponnois, R. 2006. Bacteria helping mycorrhiza development. In: K.G. Mukerji & J. Manoharachary (Eds.),
Soil Biology. Springer- Verlag, Berlin, Germany.
——— & J. Garbaye. 1991. Mycorrhization helper bacteria associated with the Douglas fir-Laccaria laccata
symbiosis: effects in aseptic and in glasshouse conditions. Annals of Science 48: 239–251. https://doi.
org/10.1051/forest:19910301
Eastwood, D.C., D. Floudas, M. Binder, A. Majcherczyk, P. Schneider, A. Aerts, F.O. Asiegbu,
S.E. Baker, K. Barry, M. Bendiksby et al. 2011. The plant cell wall-decomposing machinery
underlies the functional diversity of forest fungi. Science 333: 762–765. https://doi.org/10.1126
/science.1205411
Erlandson, R.S., J.A. Savage, J.M. Cavender-Bares & K.G. Peay. 2016. Soil moisture and chemistry
influence diversity of ectomycorrhizal fungal communities associating with willow along an hydrologic
gradient. FEMS Microbiology Ecology 92(1). https://doi.org/10.1093/femsec/fiv148
Facelli, E., T. Duan, S.E. Smith, H.M. Christophersen, J.M. Facelli & F.A. Smith. 2014. Opening the
black box: outcomes of interactions between arbuscular mycorrhizal (AM) and non-host genotypes of
Medicago depend on fungal identity, interplay between P uptake pathways and external P supply. Plant
Cell and Environment 37: 1382–1392. https://doi.org/10.1111/pce.12237
Felten, J., A. Kohler, E. Morin, R.P. Bhalerao, K. Palme, F. Martin, F.A. Ditengou & V. Legué. 2009.
The ectomycorrhizal fungus Laccaria bicolor stimulates lateral root formation in poplar and Arabidopsis
through auxin transport and signaling. Plant Physiology 151: 1991–2005. https://doi.org/10.1104
/pp.109.147231
Finlay, R.D. 2008. Ecological aspects of mycorrhizal symbiosis: with special emphasis on the functional
diversity of interactions involving the extraradical mycelium. Journal of Experimental Botany 59: 1115–
1128. https://doi.org/10.1093/jxb/ern059
——— & B. Söderström. 1992. Mycorrhiza and carbon flow to the soil. In: Allen, M.F. (ed.), Mycorrhizal
functioning: an integrative plant-fungal process. Chapman and Hall, Routledge, pp 134–162.
———, H. Ek, G. Odham & B. Soderstrom. 1988. Mycelial uptake, translocation and assimilation of
nitrogen from 15N labelled ammonium by Pinus sylvestris plants infected with four different
ectomycorrhizal fungi. New Phytologist 110: 59–66. https://doi.org/10.1111/j.1469-8137.1988.tb00237.x
Floudas, D., M. Binder, R. Riley, K. Barry, R.A. Blanchette, B. Henrissat, A.T. Martinez, R. Otillar, J.W.
Spatafora, J.S. Yadav et al. 2012. The Paleozoic origin of enzymatic lignin decompositionnre con-
structed from 31fungal genomes. Science 336: 1715–1719. https://doi.org/10.1126/science.1221748
Fomina, M., J.M. Charnock, S. Hillier, I.J. Alexander, & G.M. Gadd. 2006. Zinc phosphate transforma-
tions by the Paxillus involutus/pine ectomycorrhizal association. Microbial Ecology 52(2): 322-333.
https://doi.org/10.1007/s00248-006-9004-5
Fortin, J.A. 1966. Synthesis of mycorrhizae on explants of the root hypocotyls of Pinus sylvestris L.
Canadian Journal of Botany 44: 1087-1092. https://doi.org/10.1139/b66-116
Foster, R.C. & G.C. Marks. 1966. The fine structure of the mycorrhizas of Pinus radiata. Australian Journal
of Biological Sciences 19(6): 1027-1038. https://doi.org/10.1071/BI9661027
Frank, B. 1885. Ueber die auf Wurzelsymbiose beruhende Ernährung gewiser Bäume durch unterirdishe
Pilze. Berichte der Deutschen Botanischen Gesellschaft 3: 128–145.
Frey, B., K. Zierold & I. Brunner. 2000. Extracellular complexation of Cd in the Hartig net and cytosolic Zn
sequestration in the fungal mantle of Picea abies–Hebeloma crustuliniforme ectomycorrhizas. Plant Cell
and Environment 23(11): 1257–1265. https://doi.org/10.1046/j. 1365-3040.2000.00637.x
Frey-Klett, P., J. Garbaye & M. Tarkka. 2007. The mycorrhiza helper bacteria revisited. New Phytologist
176: 22–36. https://doi.org/10.1111/j.1469-8137.2007.02191.x
Gandini, A.M.M., P.H. Grazziotti, M.J. Rossi, D.C.F.S. Grazziotti, E.M.M. Gandini, E. de Barros Silva
& C. Ragonezi. 2015. Growth and nutrition of eucalypt rooted cuttings promoted by ectomycorrhizal
fungi in commercial nurseries. The Revista Brasileira de Ciência do Solo 39: 1554-1565. https://doi.
org/10.1590/01000683rbcs20150075
Gao, Q. & Z.L. Yang. 2010. Ectomycorrhizal fungi associated with two species of Kobresia in an alpine
meadow in the eastern Himalaya. Mycorrhiza 20 (4): 281–287. https://doi.org/10.1007/s00572-009-
0287-5
Gao, C., N.N. Shi, Y.X. Liu, Y. Zheng, Q. Ding, X.C. Mi, K.P. Ma, T. Wubet, F. Buscot & L.D. Guo.
2014. Host plant richness explains diversity of ectomycorrhizal fungi: Response to the comment of
Tedersoo et al. (2014). Molecular Ecology 23(5): 996-999. https://doi.org/10.1111/mec.12659
Garcia, K., A. Delteil, G. Conejero, A. Becquer, C. Plassard, H. Sentenac & S. Zimmermann. 2014.
Potassium nutrition of ectomycorrhizal Pinus pinaster: overexpression of the Hebeloma cylindrosporum
HcTrk1 transporter affects the translocation of both K+ and phosphorus in the host plant. New Phytologist
201: 951–960. https://doi.org/10.1111/nph.12603
Garcia, M., O. Jane, E. Smith, E. Daniel, L. Luoma, L. Melanie & D. Jones. 2016. Ectomycorrhizal
communities of ponderosa pine and lodgepole pine in the south-central Oregon pumice zone. Mycorrhiza
26: 275–286. https://doi.org/10.1007/s00572-015-0668-x
Gardes, M. & T.D. Bruns. 1996. Community structure of ectomycorrhizal fungi in a Pinus
muricata forest: above and below ground views. Canadian Journal of Botany 74: 1572–1583.
https://doi.org/10.1139/b96-190
Gehring, C.A., R.C. Mueller & T.G. Whitham. 2006. Environmental and genetic effects on the formation of
ectomycorrhizal and arbuscular mycorrhizal associations in cottonwoods. Oecologia 149: 158–164.
https://doi.org/10.1007/s00442-006-0437-9
Geml, J., N. Pastor, L. Fernandez, S. Pacheco, T.A. Semenova, A.G. Becerra, C.Y. Wicaksono, E.R.
Nouhra. 2014. Large-scale fungal diversity assessment in the Andean Yungas forests reveals strong
community turnover among forest types along an altitudinal gradient. Molecular Ecology 23: 2452–2472.
https://doi.org/10.1111/mec.12765
Grelet, G.-A., D. Johnson, E. Paterson, I.C. Anderson & I.J. Alexander. 2009. Reciprocal carbon and
nitrogen transfer between an ericaceous dwarf shrub and fungi isolated from Piceirhiza bicolorata
ectomycorrhizas. New Phytologist 182: 359-366. https://doi.org/10.1111/j.14698137.2009.0281 3.x
Grulich, V. 2012. Red List of vascular plants of the Czech Republic: 3rd edition. Preslia 84: 631–645.
Gryta, H., F. Carriconde, J.Y. Charcosset, P. Jargeat, & M. Gardes. 2006. Population dynamics of the
ectomycorrhizal fungal species Tricholoma populinum and Tricholoma scalpturatum associated with
J. Kumar, N.S. Atri
black poplar under differing environmental conditions. Environmental Microbiology 8(5): 773-786.
https://doi.org/10.1111/j.1462-2920.2005.00957.x
Guenoune, D., S. Galili, D.A. Phillips, H. Volpin, I. Chet, Y. Okon & Kapulnik. 2001. The defense
response elicited by the pathogen Rhizoctonia solani is suppressed by colonization of the AM-fungus
Glomus intraradices. Plant Science 160: 925–932. https://doi.org/10.1016/S0168-9452(01) 00329-6
Guidot, A., M.C. Verner, J.C. Debaud & R. Marmeisse. 2005. Intraspecific variation in use of different
organic nitrogen sources by the ectomycorrhizal fungus Hebeloma cylindrosporum. Mycorrhiza 15(3):
167-177. https://doi.org/10.1007/s00572-004-0318-1
Guillon, C., M. St-Arnaud, C. Hamel & S.H. Jabaji-Hare. 2002. Differential and systemic alteration of
defence-related gene transcript levels in mycorrhizal bean plants infected with Rhizoctonia solani.
Canadian Journal of Botany 80: 305-315. https://doi.org/10.1139/b02-015
Han, S.H., D.H. Kim, & J-C. Lee. 2011. Effects of the ectomycorrhizal fungus Pisolithus tinctorius and Cd
on physiological properties and Cd uptake by hybrid poplar Populus alba x glandulosa. Journal of
Ecology and Enviornment 34: 393–400. https://doi.org/10.5141/JEFB. 2011. 041
Hatch. 1936. The role of mycorrhiza in afforestation. Journal of Forest Research 34: 22-29.
Haug, I., M. Wei, J. Homeier, R. Oberwinkler & I. Kottke. 2005. Russulaceae and Thelephoraceae form
ectomycorrhizas with members of the Nyctaginaceae (Caryophyllales) in the tropical mountain rainforest
of southern Ecuador. New Phytologist 165: 923–936 https://doi.org/10.1111/j.1469-8137.2004.01284.x
Hayward, J., T.R. Horton & M.A. Nuńĩz. 2015. Ectomycorrhizal fungal communities coinvading with
Pinaceae host plants in Argentina: Gringos bajo el bosque. New Phytologist 208(2): 497-506 https://doi.
org/10.1111/nph.13453
Healy, R.A., M.E. Smith, G.M. Bonito, D.H. Pfister, Z.W. Ge, G.G. Guevara, G. Williams, K. Stafford,
L. Kumar, T. Lee, C. Hobart, J. Trappe, R. Vilgalys & D.J. McLaughlin. 2013. High diversity and
widespread occurrence of mitotic spore mats in ectomycorrhizal Pezizales. Molecular Ecology 22: 1717–
1732. https://doi.org/10.1111/mec.12135
Henkel, T.W. 2003. Monodominance in the ectomycorrhizal Dicymbe corymbosa (Caesalpiniaceae) from
Guyana. Journal of Tropical Ecology 19: 417– 437. https://doi.org/10.1017/S0266467403003468
———, M.C. Aime, M.M.L. Chin, S.L. Miller, R. Vilgalys, & M.E. Smith. 2011. Ectomycorrhizal fungal
sporocarp diversity and discovery of new taxa in Dicymbe monodominant forests of the Guiana Shield.
Biodiversity Conservation 21: 2195–2220. https://doi.org/10.1007/s10531-011-0166-1
Hentschel, E., D.L. Godbold, P. Marschner, H. Schlegel & G. Jentschke. 1993. The effect of Paxillus
involutus Fr. on aluminium sensitivity of Norway spruce seedlings. Tree Physiology 12: 379–390.
https://doi.org/10.1093/treephys/12.4.379
Hibbett, D.S., D. Grimaldi & M.J. Donoghue. 1997. Fossil mushrooms from Miocene and Cretaceous
ambers and the evolution of homobasidiomycetes. American Journal of Botany 84: 981–991. https://doi.
org/10.2307/2446289
———, L.B. Gilbert & M.J. Donaghue. 2000. Evolutionary instability of ectomycorrhizal symbioses in
basidiomycetes. Nature 407: 506–508. https://doi.org/10.1038/35035065
Hilszczańska, D., M. Małecka & Z. Sierota. 2008. Changes in nitrogen level and mycorrhizal structure of
Scots pine seedlings inoculated with Thelephora terrestris. Annals of Forest Science 65: 409. https://doi.
org/10.1051/forest:2008020
Hoeksema, J.D., J.V. Hernandez, D.L. Rogers, L.L. Mendoza & J.N. Thompson. 2012. Geographic
divergence in a species-rich symbiosis: interactions between Monterey pines and ectomycorrhizal fungi.
Ecology 93: 2274–2285. https://doi.org/10.1890/11-1715.1
Hofrichter, M., K. Vares, M. Kalsi, S. Galkin, K. Scheibner, W. Fritsche & A. Hatakka. 1999. Production
of manganese peroxidase and organic acids and mineralization of 14C-labelled lignin (14C-DHP) during
solid-state fermentation of wheat straw with the white rot fungus Nematoloma frowardii. Applied and
Environmental Microbiology 65: 1864–1870.
Högberg, M.N., E. Bååth, A. Nordgren, K. Arnebrant & P. Högberg. 2003. Contrasting effects of nitrogen
availability on plant carbon supply to mycorrhizal fungi and saprotrophs – a hypothesis based on field
observations in boreal forests. New Phytologist 160: 225–238. https://doi.org/10.1046/j.1469-
8137.2003.00867.x
Horn K, T. Frank, M. Unterseher, M. Schnittler & L. Beenken. 2013. Morphological and molecular
analyses of fungal endophytes of achlorophyllous gametophytes of Diphasiastrum alpinum
(Lycopodiaceae). American Journal of Botany 100: 2158–2174. https://doi.org/10.3732/ajb.1300011
Horton, T.R. & T.D. Bruns. 1998. Multiple-host fungi are the most frequent and abundant ectomycorrhizal
types in a mixed stand of Douglas fir (Pseudotsuga menziesii) and bishop pine (Pinus muricata). New
Horton, B.M., M. Glen, N.J. Davidson, D.A. Ratkowsky, D.C. Close, T.J. Wardlaw & C. Mohammed.
2017. An assessment of ectomycorrhizal fungal communities in Tasmanian temperate high-altitude
Eucalyptus delegatensis forest reveals a dominance of the Cortinariaceae. Mycorrhiza 27: 67-74.
https://doi.org/10.1007/s00572-016-0725-0
Hupperts, S.F., J. Karst, K. Pritsch & S.M. Landhäusser. 2017. Host phenology and potential
saprotrophism of ectomycorrhizal fungi in the boreal forest. Functional Ecology 31: 116-126
https://doi.org/10.1111/1365-2435.12695
Hynes, M.M., M.E. Smith, R.J. Zasoski & C.S. Bledsoe. 2009. A molecular survey of ectomycorrhizal
hyphae in a California Quercus-Pinus woodland. Mycorrhiza. https://doi.org/10.1007/s00572-009-028/-y
Hynson, N.A., V.S.F.T. Merckx, B.A. Perry & K.K. Treseder. 2013. Identities and distributions of the co-
invading ectomycorrhizal fungal symbionts of exotic pines in the Hawaiian Islands. Biology Invasions 15:
2373–2385. https://doi.org/10.1007/s10530-013-0458-3
Ishida T.A., K. Nara & T. Hogetsu. 2007. Host effects on ectomycorrhizal fungal communities: insight from
eight host species in mixed conifer–broadleaf forests. New Phytologist 174: 430–440. https://doi.
org/10.1111/j.1469-8137.2007.02016.x
———, ———, S. Ma, T. Takano & S. Liu. 2009. Ectomycorrhizal fungal community in alkaline-saline
soil in northeastern China. Mycorrhiza 19(5): 329-335. https://doi.org/10.1007/s00572-008-0219-9
Izumi, H., I.C. Anderson, I.J. Alexander, K. Killham & E.R. Moore. 2006. Endobacteria in some
ectomycorrhiza of Scots pine (Pinus sylvestris). FEMS Microbiology Ecology 56: 34–43. https://doi.
org/10.1111/j.1574-6941.2005.00048.x
Izzo, A., J. Agbowo & T.D. Bruns. 2005. Detection of plot level changes in ectomycorrhizal communities
across years in an old-growth mixed-conifer forest. New Phytologist 166: 619– 625. https://doi.
org/10.1111/j.1469-8137.2005.01354.x
Jairus, T., R. Mpumba, S. Chinoya & L. Tedersoo. 2011. Invasion potential and host shifts of Australian
and African ectomycorrhizal fungi in mixed eucalypt plantations. New Phytologist 192:179–187.
https://doi.org/10.1111/j.1469-8137.2011.03775.x
Javelle, A., B.R. Rodríguez-Pastrana, B. Botton, B. André, A.M. Marini, A. Brun & M. Chalot. 2001.
Molecular characterization of two ammonium transporters from the ectomycorrhizal fungus Hebeloma
cylindrosporum. FEBS Letters 505(3): 393-398. https://doi.org/10.1016/S0014-5793(01)02802-2
———, M. Morel, B.R. Rodríguez-Pastrana, B. Botton, B. André, A.M. Marini, A. Brun & M. Chalot.
2003. Molecular characterization, function and regulation of ammonium transporters (Amt) and
ammonium-metabolizing enzymes (GS, NADP-GDH) in the ectomycorrhizal fungus Hebeloma
cylindrosporum. Molecular Microbiology 47(2): 411-430. https://doi.org/10.1046/j.1365-
2958.2003.03303.x
Jentschke, G., S. Winter, & D.L. Godbold. 1999. Ectomycorrhizas and cadmium toxicity in Norway spruce
seedlings. Tree Physiology 19: 23–30. https://doi.org/10.1093/treephys/19.1.23
———, B. Brandes, A.J. Kuhn, W.H. Schröder, J.S. Becker & D.L. Godbold. 2000. The mycorrhizal
fungus Paxillus involutus transports magnesium to Norway spruce seedlings. Evidence from stable
isotope labeling. Plant Soil 220: 243–246. https://doi.org/10.1023/A: 1004727331860
Johnson, D., F. Martin, J.W.G. Cairney & I.C. Anderson. 2012. The importance of individuals: intraspe-
cific diversity on mycorrhizal plants and fungi in ecosystems. New Phytologist 194: 614–628. https://doi.
org/10.1111/j.1469-8137.2012.04087.x
Jones, D.L., P.G. Dennis, A.G. Owen & P.A.W. van Hees. 2003. Organic acid behavior in soils- miscon-
ceptions and knowledge gaps. Plant Soil 248(1): 31–41. https://doi.org/10.1023/A: 1022304332313
Jones, M.D., D.M. Durall & P.B. Tinker. 1998. A comparison of arbuscular and ectomycorrhizal Eucalyptus
coccifera: growth response, phosphorus uptake efficiency and external hyphal production. New
———, F. Grenon, H. Peat, M. Fitzgerald, L. Holt, L.J. Philip & R.L. Bradley. 2009. Differences in 15N
uptake amongst seedlings colonized by three pioneer ectomycorrhizal fungi in the field. Fungal Ecology
2: 110-120. https://doi.org/10.1016/j.funeco.2009.02.002
Jourand, P., M. Ducousso, R. Reid, C. Majorel, C. Richert, J. Riss & M. Lebrun. 2010. Nickel-tolerant
ectomycorrhizal Pisolithus albus ultramafic ecotype isolated from nickel mines in New Caledonia
strongly enhance growth of the host plant Eucalyptus globulus at toxic nickel concentrations. Tree
Physiology 30: 1311–1319. https://doi.org/10.1093/treephys/tpq070
———, L. Hannibal, C. Majorel, S. Mengant, M. Ducousso & M. Lebrun. 2014. Ectomycorrhizal
Pisolithus albus inoculation of Acacia spirorbis and Eucalyptus globules grown in ultramafic top soil
enhances plant growth and mineral nutrition while limits metal uptake. Journal of Plant Physiology 171:
164–172. https://doi.org/10.1016/j.jplph.2013.10.011
J. Kumar, N.S. Atri
Jumpponen A, K.L. Jones, J. David Mattox, C. Yaege. 2010. Massively parallel 454-sequencing of fungal
communities in Quercus spp. ectomycorrhizas indicates seasonal dynamics in urban and rural sites.
Molecular Ecology 19:41–53. https://doi.org/10.1111/j.1365-294 X.2009.04483.x
Jung, N.C. & Y. Tamai. 2013. Polyphosphate (phytate) formation in Quercus acutissima-Scleroderma
verrucosum ectomycorrhizae supplied with phosphate. Journal of Plant Interaction 8(4): 291-303.
https://doi.org/10.1080/17429145.2013.816789
Kataoka, R., T. Taniguchi & K. Futai. 2009. Fungal selectivity of two mycorrhiza helper bacteria on five
mycorrhizal fungi associated with Pinus thunbergii. World Journal of Microbiology and Biotechnology
25: 1815–1819. https://doi.org/10.1007/s11274-009-0082-7
Kayama, M. & T. Yamanaka. 2014. Growth characteristics of ectomycorrhizal seedlings of Quercus glauca,
Quercus salicina, and Castanopsis cuspidata planted on acidic soil. Trees 28: 569–583. https://doi.
org/10.1007/s00468-013-0973-y
Kemppainen, M.J. & A.G. Pardo. 2013. LbNrt RNA silencing in the mycorrhizal symbiont Laccaria
bicolor reveals a nitrate-independent regulatory role for a eukaryotic NRT2-type nitrate transporter.
Environmental Microbiology Reports 5(3): 353-366. https://doi.org/10.1111/1758-2229.12029
———, S. Duplessis, F. Martin & A.G. Pardo. 2009. RNA silencing in the model mycorrhizal fungus
Laccaria bicolor: gene knock-down of nitrate reductase results in inhibition of symbiosis with Populus.
Environmental Microbiology 11(7): 1878-1896. https://doi.org/10.1111/j. 1462-2920.2009.01912.x
Kennedy, P.G. & K.G. Peay. 2007. Different soil moisture conditions change the outcome of the ecto-
mycorrhizal symbiosis between Rhizopogon species and Pinus muricata. Plant Soil 291: 155–165.
https://doi.org/10.1007/s11104-006-9183-3.
———, P.B. Matheny, K.M. Ryberg, T.W. Henkel, J.K. Uehling & M.E. Smith. 2012. Scaling up:
examining the macroecology of ectomycorrhizal fungi. Molecular Ecology 21: 4151–4154.
Khosla, B. & M.S. Reddy. 2008. Response of ectomycorrhizal fungi on the growth and mineral nutrition of
Eucalyptus seedlings in bauxite mined soil. American-Eurasian Journal of Agricultural and
Environmental Science 3: 123-126.
———, H. Kaur & M.S. Reddy. 2009. Influence of ectomycorrhizal colonization on the growth and mineral
nutrition of Populus deltoides under Aluminum toxicity. Journal of Plant Interactions 4(2): 93-99.
https://doi.org/10.1080/17429140802382953
Kikuchi, J., N. Tsuno & K. Futai. 1991. The effect of mycorrhizae as a resistance factor of pine trees to the
pinewood menatode. Journal of Japanese Forest Society 73: 216–218.
Kluber, L.A., K.M. Tinnesand, B.A. Caldwell, S.M. Dunham, R.R. Yarwood, P.J. Bottomley & D.D.
Myrold. 2010. Ectomycorrhizal mats alter forest soil biogeochemistry. Soil Biology and Biochemistry
42: 1607–1613. https://doi.org/10.1016/j.soilbio.2010.06.001
Koide, R.T., J.N. Sharda, J.R. Herr & G.M. Malcolm. 2008. Ectomycorrhizal fungi and the
biotrophy–saprotrophy continuum. New Phytologist 178: 230–233. https://doi.org/10.1111
/j.1469-8137.2008.02401.x
Kõljalg, U., R.H. Nilsson, K. Abarenkov, L. Tedersoo, A.F.S. Taylor, E. Larsson, et al. 2013. Towards a
unified paradigm for sequence-based identification of Fungi. Molecular Ecology 22: 5271-5277.
https://doi.org/10.1111/mec.12481
Kottke, I., A. Beck, I. Haug, S. Setaro & J.P. Suárez. 2008. Mycorrhizal fungi and plant diversity in tropical
mountain rain forest of southern Ecuador. In: S.R. Gradstein, J. Homeier & D. Gansert, (Eds.), The
Tropical Mountain Forest: Patterns and Processes in a Biodiversity Hotspot. Biodiversity and Ecology
Series, Universitätsverlag Göttingen, vol. 2 pp. 7-67.
———, S. Setaro, I. Haug, P. Herrera, D. Cruz, A. Fries, J. Gawlik, J. Homeier, F.A. Werner, A.
Gerique & J.P. Suárez. 2013. Mycorrhiza networks promote biodiversity and stabilize the tropical
mountain rain forest ecosystem: Perspectives for understanding complex communities In: J.Bendix, E.
Beck, I. Kottke, F. Makeschin & R. Mosandl (Eds.), Ecosystem Services, Biodiversity and Environmental
Change in a Tropical Mountain Ecosystem of South Ecuador, Ecological Studies. Springer-Verlag Berlin
Heidelberg, pp.187-203.
Kranabetter, J.M., D.M. Durall & W.H. MacKenzie. 2009. Diversity and species distribution of
ectomycorrhizal fungi along productivity gradients of a southern boreal forest. Mycorrhiza 19: 99–111.
https://doi.org/10.1007/s00572-008-0208-z
Kropp, B.R. & C-G. Langlois. 1990. Ectomycorrhizae in reforestation. Canadian Journal of Forest Research
20(4): 438-451. https://doi.org/10.1139/x90-061
Krpata, D., W. Fitz, U. Peintner, I. Langer & P. Schweiger. 2009. Bioconcentration of zinc and cadmium in
ectomycorrhizal fungi and associated aspen trees as affected by level of pollution. Environmental
Pollution 157: 280–286. https://doi.org/10.1016/j.envpol.2008.06.038
Krupa, P. & J. Kozdrój. 2004. Accumulation of Heavy Metals by Ectomycorrhizal Fungi Colonizing Birch
Trees Growing in an Industrial Desert Soil. World Journal of Microbiology and Biotechnology 20(4):
427-430. https://doi.org/10.1023/B:WIBI.0000033067.64061.f3
Krznaric, E., N. Verbruggen, J.H.L. Wevers, R. Carleer, J. Vangronsveld & J.V. Colpaert. 2009. "Cd-
tolerant Suillus luteus: a fungal insurance for pines exposed to Cd.". Environmental Pollution 157: 1581–
1588. https://doi.org/10.1016/j.envpol.2008.12.030
———, J.H.L. Wevers, C. Cloquet, J. Vangronsveld, F. Vanhaecke & J.V. Colpaert. 2010. Zn pollution
counteracts Cd toxicity in metaltolerant ectomycorrhizal fungi and their host plant, Pinus sylvestris.
Environmental Microbiology 12: 2133–2141. https://doi.org/10.1111/j.1462-2920.2009.02082.x
Kumar, J. & N.S. Atri. 2016. Characterisation of ectomycorrhiza of Russula and Lactifluus (Russulaceae)
associated with Shorea robusta from Indian Shiwaliks. Nova Hedwigia 103(3-4): 501-513. https://doi.
org/10.1127/nova_hedwigia/2016/0368
Kumla, J., E.A. Hobbie, N. Suwannarach & S. Lumyong. 2016. The ectomycorrhizal status of a tropical
black bolete, Phlebopus portentosus, assessed using mycorrhizal synthesis and isotopic analysis.
Kurth, F., K. Zeitler, L. Feldhahn, T.R. Neu, T. Weber, V. Krištůfek, T. Wubet, S. Herrmann, F. Buscot
& M. Tarkka. 2013. Detection and quantification of a mycorrhization helper bacterium and a mycor-
rhizal fungus in plant-soil microcosms at different levels of complexity. BMC Microbiology 13: 205.
https://doi.org/10.1186/1471-2180-13-205
Kuster, H., A. Becker, C. Firnhaber, N. Hohnjec, K. Manthey, A.M. Perlick, T. Bekel, M. Dondrup, K.
Henckel, A. Goesmann, F. Meyer et al. 2007. Development of bioinformatic tools to support EST-
sequencing, in silico- and microarray-based transcriptome profiling in mycorrhizal symbioses.
Phytochemistry 38: 19-32. https://doi.org/10.1016/j.phytochem.2006.09.026
Kyaschenko, J., K. E. Clemmensen, A. Hagenbo, E. Karltunand & B. D. Lindahl. 2017. Shift in fungal
communities and associated enzyme activities along an age gradient of managed Pinus sylvestris stands.
The ISME Journal 11: 863-874. https://doi.org/10.1038/ismej.2016.184
Lakhanpal, T.N. & S. Kumar. 1984. Studies on mycorrhiza and mycorrhizosphere of Picea smithiana.
Journal of Tree Science 3: 5–9.
Lambers, H., F.S. Chapin & T.L. Pons. 1998. Plant physiological ecology. Springer, New York.
———, M.C. Brundrett, J.A. Raven & S.D. Hopper. 2010. Plant mineral nutrition in ancient landscapes:
high plant species diversity on infertile soils is linked to functional diversity for nutritional strategies. Plant
Soil 334: 11–31. https://doi.org/10.1007/s11104-010-0444-9
Lamhamedi, M.S., P.Y. Mernier & J.A. Fortin. 1992. Growth, nutrition and response to water stress of
Pinus pinaster inoculated with ten dikaryotic strains of Pisolithus sp. Tree Physiology 10: 153-167
Lancelloti, E. & A. Franceschini. 2013. Studies on the ectomycorrhizal community in a declining Quercus
suber L. Stand. Mycorrhiza 23: 533-542. https://doi.org/10.1007/s00572-013-0493-2
Langenfeld-Heyser, R., J. Gao, T. Ducic, Ph. Tachd, C.F. Lu, E. Fritz, A. Gafur & A. Polle. 2007.
Paxillus involutus mycorrhiza attenuate NaCl-stress responses in the salt-sensitive hybrid poplar Populus
× canescens. Mycorrhiza 17: 121-131. https://doi.org/10.1007/s00572-006-0084-3
Leake, J.R. 2004. Myco-heterotroph/epiparasitic plant interactions with ectomycorrhizal and arbuscular
mycorrhizal fungi. Current Opinion in Plant Biology 7: 422 – 428.
Lehto, T. & J.J. Zwiazek. 2011. Ectomycorrhizas and water relations of trees: a review. Mycorrhiza 21(2):
71–90. https://doi.org/10.1007/s00572-010-0348-9
Leonardi, M., O. Comandini & A.C. Rinaldi. 2016. Peering into the Mediterranean black box: Lactifluus
rugatus ectomycorrhizas on Cistus. IMA FUNGUS 7(2): 275–284. https://doi.org/10.5598/ imafungus.
2016.07.02.07
Leyval, C. & J. Berthelin. 1989. Interactions between Laccaria laccata, Agrobacterium radiobacter and
beech roots: influence on P, K, Mg and Fe mobilization from minerals and plant growth. Plant Soil 117:
103-120. https://doi.org/10.1007/BF02206262
Li, J., S. Bao, Y. Zhang, X. Ma, M. Mishra-Knyrim, J. Sun, G. Sa, X. Shen, A. Polle & S. Chen. 2012.
Paxillus involutus strains MAJ and NAU mediate K(+)/Na(+) homeostasis in ectomycorrhizal Populus x
canescens under sodium chloride stress. Plant Physiology 159: 1771–1786, https://doi.org/10.1104
/pp.112.195370
Lilleskov, E.A., T.J. Fahey, T.R. Horton & G.M. Lovett. 2002. Belowground ectomycorrhizal fungal
community change over a nitrogen deposition gradient in Alaska. Ecology 83:104-115. https://doi.
org/10.2307/2680124
Lindahl, B.D. & A. Tunlid. 2015. Ectomycorrhizal fungi – potential organic matter decomposers, yet not
saprophytes. New Phytologist 205: 1443–1447. https://doi.org/10.1111/nph.13201
J. Kumar, N.S. Atri
Looney, B.P., M. Ryberg, F. Hampe, M. Sánchez-García & P.B. Matheny. 2016. Into and out of the
tropics: global diversification patterns in a hyper-diverse clade of ectomycorrhizal fungi. Molecular
Ecology 25: 630–647. https://doi.org/10.1111/mec.13506
Lotti, M. & A. Zambonelli. 2006. A quick and precise technique for identifying ectomycorrhizas by PCR.
Mycological Research 110: 60–65. https://doi.org/10.1016/j.mycres.2005.09.010
Luo, Z.B., D. Janz, X. Jiang, C. Göbel, H. Wildhagen, Y. Tan, H. Rennenberg, I. Feussner & A. Polle.
2009. Upgrading root physiology for stress tolerance by ectomycorrhizas: insights from metabolite and
transcriptional profiling into reprogramming for stress anticipation. Plant Physiology 151: 1902–1917.
https://doi.org/10.1104/pp.109.143735
———, K. Li, Y. Gai, C. Gobel, H. Wildhagen, X.N. Jiang, I. Feussner, H. Rennenberg & A. Polle. 2011.
The ectomycorrhizal fungus (Paxillus involutus) modulates leaf physiology of poplar towards improved
salt tolerance. Environmental and Experimental Botany 72: 304–311. https://doi.org/10.1016/j.
envexpbot.2011.04.008
———, Ch. Wua, Ch. Zhang, H. Lic, U. Lipkad, Polleda & A. College. 2014. The role of ectomycorrhizas
in heavy metal stress tolerance of host plants. Environmental and Experimental Botany 108: 47–62.
https://doi.org/10.1016/j.envexpbot.2013.10.018
Ma, Y., J. He, C. Ma, J. Luo, H. Li, T. Liu, A. Polle, C. Peng & Z.B. Luo. 2013. Ectomycorrhizas with
Paxillus involutus enhance cadmium uptake and tolerance in Populus canescens. Plant Cell Environment
37: 627–642. https://doi.org/10.1111/pce.12183
Maba, D.L., A.K. Guelly, N.S. Yorou, A. DE Kesel, A. Verbeken & R. Agerer. 2014. The genus Lactarius
s. str. (Basidiomycota, Russulales) in Togo (West Africa): phylogeny and new species described. IMA
Fungus 5: 39-49. https://doi.org/10.5598/imafungus.2014.05.01.05
———, A.K. Guelly, N.S. Yorou, A. Verbeken & R. Agerer. 2015. Phylogenetic and microscopic studies in
the genus Lactifluus (Basidiomycota, Russulales) in West Africa, including the description of four new
species. IMA Fungus 6 (1): 39-49. https://doi.org/10.5598/ima fungus.2015 .06.01.02
Malysheva, E.F., V.F. Malysheva, A.E. Kovalenko, E.A. Pimenova, M.N. Gromyko, S.N. Bondarchuk &
E.Y. Voronina. 2016. Below-Ground Ectomycorrhizal Community Structure in the Postfire Successional
Pinus koraiensis Forests in the Central Sikhote-Alin (the Russian Far East). Botanica Pacifica- Journal of
plant science and conservation 5(1): 19–31. 10.17581/bp.2016.05102
Marcel, G., A. van der Heijden, F.M. Martin, M.A. Selosse & I.R. Sanders. 2015. Mycorrhizal ecology
and evolution: the past, the present, and the future. New Phytologist 205: 1406–1423. https://doi.
org/10.1111/nph.13288
Martin, F. & M.A. Selosse. 2008. The Laccaria genome: a symbiont blueprint decoded. New Phytologist
180: 296–310. https://doi.org/10.1111/j.1469-8137.2008.02613.x
———, A. Aerts, D. Ahren, A. Brun, E.G.J. Danchin, F. Duchaussoy, J. Gibon, A. Kohler E. Lindquist,
V. Pereda et al. 2008. The genome of Laccaria bicolor provides insights into mycorrhizal symbiosis.
Nature 452: 88–92. https://doi.org/10.1038/nature06556
———, A. Kohler, C. Murat, R. Balestrini, P.M. Coutinho, O. Jaillon, B. Montanini, E. Morin, B. Noel,
R. Percudani et al. 2010. Perigord black truffle genome uncovers evolutionary origins and mechanisms
of symbiosis. Nature 464: 1033–1038. https://doi.org/10.1038/nature08867
Marx, D.H. 1972. Ectomycorrhizae as biological deterrents to pathogenic root infections. Annual Review of
Phytopathology 10: 429-454. https://doi.org/10.1146/annurev.py.10.090172.002241
———, K. Jarl, J.L. Ruehle & W. Bell. 1984. Development of Pisolithus tinctorius ectomycorrhizae on pine
seedlings using basidiospore encapsulated seeds. Forest Science 30: 897–907.
———, A. Hedin & S.F.P. Toe. 1985. Field performance of Pinus caribaea var. Hondurensis seedlings with
specific ectomycorrhizae and fertilizer after three years on a savana site Liberia. Forest Ecology and
Management 13(12): 1-25. https://doi.org/10.1016/0378-1127(85)90002-7
———, J.L. Ruehle & C.E. Cordell. 1991. Methods for Studying Nursery and Field Response of Trees to
Specific Ectomycorrhiza. In: J.R. Norris, D.J. Read & A.K. Varma (Eds.), Methods in Microbiology,
Academic Press, London, UK. pp. 383–411.
Matheny, P.B., J.M. Curtis, V. Hofstetter, M.C. Aime, J.M. Moncalvo, Z.W. Ge, Z.L. Yang, J.C. Slot,
J.F. Ammirati, T.J. Baroni et al. 2006. Major clades of Agaricales: a multilocus phylogenetic overview.
Mycologia 98: 982–995. https://doi.org/10.3852/mycologia.98.6.982
———, M.C. Aime, N.L. Bougher, B. Buyck, D.E. Desjardin et al. 2009. Out of the Palaeotropics?
Historical biogeography and diversification of the cosmopolitan ectomycorrhizal mushroom family
Inocybaceae. Journal of Biogeography 36: 577–592. https://doi.org/10.1111/j.1365-2699.2008.02055.x
Matsuda, Y., N. Hayakawa & S. Ito. 2008. Local and microscale distributions of Cenococcum geophilum in
soils of coastal pine forests. Fungal Ecology 2: 31–35. https://doi.org/10.1016/ j.funeco.2008.10.002
———, Y. Noguchi & S. Ito. 2009. Ectomycorrhizal fungal community of naturally regenerated Pinus
thunbergii seedlings in a coastal pine forest. Journal of Forest Research 14: 335–341. https://doi.
org/10.1007/s10310-009-0140-x.
Maurel, C. & C. Plassard. 2011. Aquaporins: for more than water at the plant–fungus interface? New
Phytologist 190(4): 815–817. https://doi.org/10.1111/j.1469-8137.2011.03731.x
Maury-Lechon, G. & L. Curtet. 1998. Biogeography and Evolutionary Systematics of Dipterocarpaceae. In:
A Review of Dipterocarps: Taxonomy, ecology and silviculture Center for International Forestry Research
Bogor, Indonesia ISBN 979-8764-20-X.
Mediavilla, O., J. Olaizola, L. Santos-del-Blanco, J.A. Oria-de-Rueda & P. Martín-Pinto. 2016.
Mycorrhization between Cistus ladanifer L. and Boletus edulis Bull is enhanced by the mycorrhiza
helper bacteria Pseudomonas fluorescens Migula. Mycorrhiza 26: 161–168. https://doi.org/10.1007
/s00572-015-0657-0
Menkis, A., D. Burokiene, T. Gaitnieks, A. Uotila, H. Johannesson, A. Rosling, R. Finlay, J. Stenlid & R.
Vasaitis. 2012. Occurrence and impact of the root-rot biocontrol agent Phlebiopsis gigantea on soil
fungal communities in Picea abies forests of northern Europe. FEMS Microbiology Ecology 81: 438-445.
https://doi.org/10.1111/j.1574-6941.2012.01366.x
Miller, R.M. & J.D. Jastrow. 1992. The application of VA mycorrhizae to ecosystem restoration and
reclamation. In: M. Allen (Ed.), Mycorrhizal functioning. Chapman & Hall, New York.
Moeller, H.V., I.A. Dickie, D.A. Peltzer & T. Fukami. 2015. Mycorrhizal coinvasion and novel interactions
depend on neighborhood context. Ecology 96: 2336–2347. https://doi.org/10.1890/14-2361.1
Mohan, V., K. Natarajan & K. Ingleby. 1993. Anatomical studies on ectomycorrhizas. I. The
ectomycorrhizas produced by Thelephora terrestris on Pinus patula. Mycorrhiza 3: 39–42. https://doi.
org/10.1007/BF00213466
———, R. Nivea, S. Menon. 2015. Evaluation of Ectomycorrhizal Fungi as Potential Bio-control Agents
against Selected Plant Pathogenic Fungi. Journal of Academia and Industrial Research (JAIR) 3 (9): 408-
412.
Molina, R. & J.M. Trappe. 1982. Patterns of ectomycorrhizal host specificity and potential among Pacific
North – West Conifer and fungi. Forest Science 28: 423-458.
——— & ———. 1994. Biology of the ectomycorrhizal genus: Rhizopogon I. Host associations, host-
specificity and pure culture syntheses. New Phytologist 126: 653–675. https://doi.org/10.1111/j.1469-
8137.1994.tb02961.x
Molinier, V., C. Murat, M. Peter, A. Gollotte, H. De la Varga, B. Meier, S. Egli, B. Belfiori, F. Paolocci &
D. Wipf. 2016. SSR-based identification of genetic groups within European populations of Tuber
aestivum Vittad. Mycorrhiza 26: 99–110. https://doi.org/10.1007/s00572-015-0649-0
Morel, M., C. Jacob, M. Fitz, D. Wipf, M. Chalot & A. Brun. 2008. Characterization and regulation of
PiDur3, a permease involved in the acquisition of urea by the ectomycorrhizal fungus Paxillus involutus.
Fungal Genetics and Biology 45: 912–921. https://doi.org/10.1016/j.fgb.2008.01.002
Morris, M.H., M.E. Smith, D.M. Rizzo, M. Rejmanek & C.S. Bledsoe. 2008. Contrasting ectomycorrhizal
fungal communities on the roots of co-occurring oaks (Quercus spp.) in a California woodland. New
Phytologist 178: 167–176. https://doi.org/10.1111/j.14698137.2007.02348.x
Moyer-Henry, K., I. Silva, J. Macfall, E. Johannes, N. Allen, B. Goldfarb & T. Rufty. 2005.
Accumulation and localization of aluminium in root tips of loblolly pine seedlings and the associated
ectomycorrhiza Pisolithus tinctorius. Plant Cell Environment 28: 111–120. https://doi.org/10.1111/j.1365-
3040.2004.01240.x
Moyersoen, B. & M. Weiß. 2014. New Neotropical Sebacinales Species from a Pakaraimaea
dipterocarpacea Forest in the Guayana Region, Southern Venezuela: Structural Diversity and
Phylogeography. PLOS ONE 9(8): e107078. https://doi.org/10.1371/journal.pone.0107078
Mrak, T., K. Kühdorf, T. Grebenc, I. Štraus, B. Münzenberger & H. Kraigher. 2017. Scleroderma
areolatum ectomycorrhiza on Fagus sylvatica L. Mycorrhiza 27: 283-293. https://doi.org/10.1007
/s00572-016-0748-6
Muhlmann, O., M. Bacher & U. Peintner. 2008. Polygonum viviparum mycobionts on an alpine primary
successional glacier forefront. Mycorrhiza 18: 87–95. https://doi.org/10.1007/s00572-007-0156-z
Müller, T., M. Avolio, M. Olivi, M. Benjdia, E. Rikirsch, A. Kasaras et al. 2007. Nitrogen transport in the
ectomycorrhiza association: the Hebeloma cylindrosporum-Pinus pinaster model. Phytochemistry 68:
41–51. https://doi.org/10.1016/j.phytochem.2006.09.021
Nakashima, H., N. Eguchi, T. Uesugi, N. Yamashita & Y. Matsuda. 2016. Effect of ectomycorrhizal
composition on survival and growth of Pinus thunbergii seedlings varying in resistance to the pine wilt
nematode. Trees 30: 475–481. https://doi.org/10.1007/s00468-015-1217-0
J. Kumar, N.S. Atri
Nara, K. 2006. Ectomycorrhizal networks and seedling establishment during early primary succession. New
Natarajan, K., G. Senthilrasu, V. Kumaresan & T. Riviera. 2005. Diversity in Ectomycorrhizal fungi of a
dipterocarp forest in Western Ghats. Current Science 88 (12): 1893-1895.
Navarro-Ródenas, A., L.M. Berná, C. Lozano-Carrillo, A. Andrino & A. Morte. 2016. Beneficial native
bacteria improve survival and mycorrhization of desert truffle mycorrhizal plants in nursery conditions.
Mycorrhiza 26(7): 769-779. https://doi.org/10.1007/s00572-016-0711-6
Nehls, U., Dietz, S. 2014. Fungal aquaporins: cellular functions and ecophysiological perspectives. Fungal
Aquaporins: Cellular Functions and Ecophysiological Perspectives. Applied Microbiology and
Biotechnology 98(21): 8835-8851. https://doi.org/10.1007/s00253-014-6049-0
———, F. Göhringer, S. Wittulsky & S. Dietz. 2010. Fungal carbohydrate support in the ectomycorrhizal
symbiosis: a review. Plant Biology 12(2): 292-301. https://doi.org/10.1111/j.1 438-8677.2009.00312.x
Nouhra, E., C. Urcelay, S. Longo & L. Tedersoo. 2013. Ectomycorrhizal fungal communities associated to
Nothofagus species in Northern Patagonia. Mycorrhiza. https://doi.org/10.1007/s00572-013-0490-2
Nuñez, M.A., T.R. Horton & D. Simberloff. 2009. Lack of belowground mutualisms hinders Pinaceae
invasions. Ecology 90(9): 2352–2359. https://doi.org/10.1890/08-2139.1
O’Hanlon, R. 2012. Below-ground ectomycorrhizal communities: the effect of small scale spatial and short
term temporal variation. Symbiosis 57: 57–71. https://doi.org/10.1007/s13199-012-0179-x
Obase, K., J.Y. Cha, J.K. Lee, S.Y. Lee, J.H. Lee & K.W. Chun. 2009. Ectomycorrhizal fungal commu-
nities associated with Pinus thunbergii in the eastern coastal pine forests of Korea. Mycorrhiza 20(1): 39-
49. https://doi.org/10.1007/s00572-009-0262-1
Ott, T., E. Fritz, A. Polle & A. Schützendübel. 2002. Characterization of antioxidative systems in the
ectomycorrhiza-building basidiomycete Paxillus involutus (Bartsch) Fr. and its reaction to cadmium.
FEMS Microbiology Ecology 42: 359–366. https://doi.org/10.1111/j.1574-6941.2002.tb01025.x
Pande, V., U.T. Palni & S.P. Singh. 2004. Species diversity of ectomycorrhizal fungi associated with
temperate forest of Western Himalaya: a preliminary assessment. Current Science 86: 1619-1623.
Parrent, J.L., W.F. Morris & R. Vilgalys. 2006. CO2-enrichment and nutrient availability alter
ectomycorrhizal fungal communities. Ecology 87: 2278–2287. https://doi.org/10.1890/0012-9658(2006)
87[2278:CA NAAE] 2.0.CO;2
Paul, L.R., B.K. Chapman & C.P. Chanway. 2007. Nitrogen fixation associated with Suillus tomentosus
tuberculate ectomycorrhizae on Pinus contorta var. latifolia. Annals of Botany 99: 1101–1109. https://doi.
org/10.1093/aob/mcm061
Peay, K.G. & P.B. Matheny. 2017. Biogeography of ectomycorrhizal fungi. In: F. Martin (eds.), The
Molecular Mycorrhizal Symbiosis, John Wiley & Sons, pp. 341-361.
———, P.G. Kennedy, S.J. Davies, S. Tan & T.D. Bruns. 2010. Potential link between plant and fungal
distributions in a dipterocarp rainforest: community and phylogenetic structure of tropical
ectomycorrhizal fungi across a plant and soil ecotone. New Phytologist 185: 529–542. https://doi.
org/10.1111/j.1469-8137.2009.03075.x
———, ——— & T.D. Bruns. 2011. Rethinking ectomycorrhizal succession: are root density and hyphal
exploration types drivers of spatial and temporal zonation? Fungal Ecology 4: 233–240. https://doi.
org/10.1016/j.funeco.2010.09.010
———, M.G. Schubert, N.H. Nguyen & T.D. Bruns. 2012. Measuring ectomycorrhizal fungal dispersal:
macroecological patterns driven by microscopic propagules. Molecular Ecology 21: 4122–4136.
https://doi.org/10.1111/j.1365-294X.2012.05666.x
———, S.E. Russo, K.L. McGuire, Z. Lim, J.P. Chan, S. Tan & S.J. Davies. 2015. Lack of host specificity
leads to independent assortment of dipterocarps and ectomycorrhizal fungi across a soil fertility gradient.
Ecology Letter 18(8): 807–816 https://doi.org/10.1111/ele.12459
Pedersen, B.P., H. Kumar, A.B. Waight, A.J. Risenmay, Z. Roe-Zurz, B.H. Chau, A. Schlessinger, M.
Bonomi, W. Harries & A. Sali. 2013. Crystal structure of a eukaryotic phosphate transporter. Nature
496: 533– 536. https://doi.org/10.1038/nature12042
Perez-Moreno, J. & D.J. Read. 2000. Mobilization and transfer of nutrients from litter to tree seedlings via
the vegetative mycelium of ectomycorrhizal plants. New Phytologist 145: 301–309, https://doi.
org/10.1046/j.1469-8137.2000.00569.x
Pfabel, C., K.U. Eckhardt, C. Baum, C. Struck, P. Frey & M. Weih. 2012. Impact of ectomycorrhizal
colonization and rust infection on the secondary metabolism of poplar (Populus trichocarpa deltoides).
Tree Physiology 32: 1357–1364. https://doi.org/10.1093/treephys/tps093
Phillips, L.A., V. Ward & M.D. Jones. 2014. Ectomycorrhizal fungi contribute to soil organic matter cycling
in sub-boreal forests. ISME Journal 8: 699–713. https://doi.org/10.1038/ismej.2013.195
Phosri, C., S. Põlme, A.F.S. Taylor, U. Kõljalg, N. Suwannasai & L. Tedersoo. 2012. Diversity and
community composition of ectomycorrhizal fungi in a dry deciduous dipterocarp forest in Thailand.
Biodiversity and Conservation 21: 2287–2298. https://doi.org/10.1007/s10531-012-0250-1
Plassard, C. & B. Dell. 2010. Phosphorus nutrition of mycorrhizal trees. Tree Physiology 30: 1129–1139.
https://doi.org/10.1093/treephys/tpq063
Plett, J.M. & F. Martin. 2011. Blurred boundaries: lifestyle lessons from ectomycorrhizal fungal genomes.
Trends in Genetics 27: 14–22. https://doi.org/10.1016/j.tig.2010.10.005
———, Y. Daguerre, S. Wittulsky, A. Vayssières, A. Deveau, S.J. Melton, A. Kohler, J.L. Morrell-
Falvey, A. Brun, C. Veneault-Fourrey & F. Martin. 2014. Effector MiSSP7 of the mutualistic fungus
Laccaria bicolor stabilizes the Populus JAZ6 protein and represses jasmonic acid (JA) responsive genes.
Proceedings of National Academy of Sciences USA. 111(22): 8299-8304. https://doi.org/10.1073
/pnas.1322671111
Pressel, S., M.I. Bidartondo, R. Ligrone, J.G. Duckett. 2010. Fungal symbioses in bryophytes: New
insights in the twenty first century. Phytotaxa 9: 238-253. 10.11646/phytotaxa.9.1.13
Pringle, A., R.I. Adams, H.B. Cross & T.D. Bruns. 2009. The ectomycorrhizal fungus Amanita phalloides
was introduced and is expanding its range on the west coast of North America. Molecular Ecology 18(5):
817–833. https://doi.org/10.1111/j.1365-294X.2008.04030.x
Pritsch, K., & J. Garbaye. 2011. Enzyme secretion by ECM fungi and exploitation of mineral nutrients from
soil organic matter. Annals of Forest Science 68: 25–32. https://doi.org/10.1007/s13595-010-0004-8
———, H. Boyle, J.C. Munch & F. Buscot. 1997. Characterization and identification of black alder
ectomycorrhizas by PCR/RFLP analyses of the rDNA internal transcribed spacer. New Phytologist
137: 357–369. https://doi.org/10.1046/j.1469-8137.1997.00806.x
Pyasi, A., K.K. Soni & R.K. Verma. 2011. Dominant occurrence of ectomycorrhizal colonizer Astraeus
hygrometricus of sal (Shorea robusta) in forest of Jharsuguda Orissa. Journal of Mycology and Plant
Pathology 41(2): 222-225.
Quoreshi, A.M. & D.P. Khasa. 2008. Effectiveness of mycorrhizal inoculation in the nursery on root
colonization, growth, and nutrient uptake of aspen and balsam poplar. Biomass Bioenergy 32: 381–
391. https://doi.org/10.1016/j.biombioe.2007.10.010
———, Y. Piché & D.P. Khasa. 2008. Field performance of conifer and hardwood species five years after
nursery inoculation in the Canadian Prairie Provinces. New Forests 35: 235–253. https://doi.org/10.1007
/s11056-007-9074-3
Quoreshi, M. & V.R. Timmer. 2000. Growth, nutrient dynamics, and ectomycorrhizal development of
container-grown Picea mariana seedlings in response to exponential nutrient loading. Canadian Journal
of Forest Research 30: 191–201. https://doi.org/10.1139/x99-208
Richard, F., M. Roy, O. Shahin, C. Sthultz, M. Duchemin, R. Joffre & M.A. Selosse. 2011.
Ectomycorrhizal communities in a Mediterranean forest ecosystem dominated by Quercus ilex: seasonal
dynamics and response to drought in the surface organic horizon. Annals of Forest Science 68: 57–68.
https://doi.org/10.1007/s13595-010-0007-5
Rinaldi, A.C., O. Comadini & T.W. Kuyper. 2008. Ectomycorrhizal fungal diversity: separating the wheat
from the chaff. Fungal Diversity 33:1–45.
Rincón, A., J. Parladé & J. Pera. 2005. Effects of ectomycorrhizal inoculation and the type of substrate on
mycorrhization, growth and nutrition of containerised Pinus pinea L. seedlings produced in a commercial
nursery. Annals of Forest Science 62: 1–6. https://doi.org/10.1051/forest:2005087
———, B. Ruiz-Diez, M. Fernandez-Pascual, A. Probanza, J.M. Pozuelo & M.R. de Felipe. 2006.
Afforestation of degraded soils with Pinus halepensis Mill.: effects of inoculation with selected micro-
organisms and soil amendment on plant growth, rhizospheric microbial activity and ectomycorrhizal
formation. Applied Soil Ecology 34:42–51. https://doi.org/10.1016/ j.apsoil.2005. 12.004
———, M.R. de Felipe & M. Fernández-Pascual. 2007. Inoculation of Pinus halepensis Mill. with selected
ectomycorrhizal fungi improves seedling establishment 2 years after planting in a degraded gypsum soil.
Mycorrhiza 18(1): 23–32. https://doi.org/10.1007/s00572-007-0149-y
Rineau, F., F. Shah, M. Smits, P. Persson, T. Johansson, R. Carleer, C. Troein & A. Tunlid. 2013. Carbon
availability triggers the decomposition of plant litter and assimilation of nitrogen by an ectomycorrhizal
fungus. ISME Journal 7: 2010– 2022. https://doi.org/10.1038/ismej.2013.91
Riviere, T., A.G. Diedhiou, M. Diabate, G. Senthilarasu, K. Natarajan, A. Verbeken, B. Buyck, B.
Dreyfus, G. Bena & A.M. Bâ. 2007. Genetic diversity of ectomycorrhizal basidiomycetes from African
and Indian tropical forests. Mycorrhiza 17: 415–428. https://doi.org/10.1007/s00572-007-0117-6
Roman, M.D., V. Claveria & A.M.D. Miguel. 2005. A revision of the descriptions of ectomycorrhizas
published since 1961. Mycological Research 109 (10): 1063-1104. https://doi.org/10.1017
/S0953756205003564
J. Kumar, N.S. Atri
Rouhier, H. & D.J. Read. 1999. Plant and fungal responses to elevated atmospheric CO2 in mycorrhizal
seedlings of Betula pendula. Environmental Experimental Botany 42: 231–241. https://doi.org/10.1016/
S0098-8472(99)00039-8
Rousseau, J.V.D., D.M. Sylvia & A.J. Fox. 1994. Contribution of ectomycorrhiza to the potential nutrient-
absorbing surface of pine. New Phytologist 128: 639–644. https://doi.org/10.1111/j.1469-8137.1994.
tb04028.x
Roy, M., J. Rochet, S. Manzi, P. Jargeat, H. Gryta, P.A. Moreau et al. 2013. What determines Alnus
associated ectomycorrhizal community diversity and specificity? A comparison of host and habitat effects
at a regional scale. New Phytologist 198: 1228–1238. https://doi.org/10.1111/ nph.12212
———, A.Vasco-Palacios, J. Geml, B. Buyck, L. Delgat et al. 2017. The (re)discovery of ectomycorrhizal
symbioses in Neotropical ecosystems sketched in Florianópolis. New Phytologist 214: 920–923.
Roy-Bolduc, A., E. Laliberté & M. Hijri. 2016. High richness of ectomycorrhizal fungi and low host
specificity in costal sand dune ecosystem revealed by network analysis. Ecology Evolution 6(1): 349-362.
https://doi.org/10.1002/ece3.1881
Rudawska, M., M. Pietras, I. Smutek, P. Strzeliński & T. Leski. 2016. Ectomycorrhizal fungal assem-
blages of Abies alba Mill. outside its native range in Poland. Mycorrhiza 26: 57–65. https://doi.
org/10.1007/s00572-015-0646-3
Schier, G. & C. McQuattie. 1995. Effect of aluminium on the growth, anatomy, and nutrient content of
ectomycorrhizal and nonmycorrhizal eastern white pine seedlings. Canadian Journal of Forest Research
25: 1252– 1262. https://doi.org/10.1139/x95-138
Sebastiana, M., J. Martins, A. Figueiredo, F. Monteiro, J. Sardans, J. Peñuelas, A. Silva, P. Roepstorff,
M.S. Pais & A.V. Coelho. 2017. Oak protein profile alterations upon root colonization by an
ectomycorrhizal fungus. Mycorrhiza 27: 109-128. https://doi.org/10.1007/s00572-016-0734-z
Shah, F., C. Nicolàs, J. Bentzer, M. Ellström, M. Smits, F. Rineau, B. Canbäck, D. Floudas, R. Carleer,
G. Lackner, J. Braesel, D. Hoffmeister, B. Henrissat, D. Ahrén, T. Johansson, D.S. Hibbett, F.
Martin, P. Persson & A. Tunlid. 2016. Ectomycorrhizal fungi decompose soil organic matter using
oxidative mechanisms adapted from saprotrophic ancestors. New Phytologist 209: 1705–1719.
https://doi.org/10.1111/nph.13722
Shakya, M., N. Gottel, H. Castro, Z.K. Yang, L. Gunter, J. Labbe, W. Muchero, G. Bonito, R. Vilgalys,
G. Tuskan et al. 2013. A multifactor analysis of fungal and bacterial community structure in the root
microbiome of mature Populus deltoides trees. PLoS ONE 8(10): e76382. https://doi.org/10.1371/journal.
pone.0076382
Sharma R, R.C. Rajak, A.K. Pandey. 2008a. Some ectomycorrhizal mushrooms of Central India-I. Russula.
Journal of Mycopathological Research 46(2): 201-212.
———, ———, ———. 2008b. Some ectomycorrhizal mushrooms of Central India-II. Lactarius. Journal of
Mycopathological Research 47(1): 43-47.
———, ———, ———. 2009: Ectomycorrhizal mushrooms in Indian tropical forests. Biodiversity 10 (1):
25-30. https://doi.org/10.1080/14888386.2009.9712634
Sharma, S., M.K. Saini & N.S. Atri. 2016. Some new records of Russulaceous mushrooms from North West
Himalayas. Kavaka 46: 5-13.
Shrestha, V.G., K. Shrestha & H. Wallander. 2005. Antagonistic study of ectomycorrhizal fungi isolated
from Baluwa forest (Central Nepal) against pathogenic fungi and bacteria. Science World 3: 44-56.
Siemens, J.A., M. Calvo-Polanco & J.J Zwiazek. 2011. Hebeloma crustuliniforme facilitates ammonium
and nitrate assimilation in trembling aspen (Populus tremuloides) seedlings. Tree Physiology 31: 1238–
1250. https://doi.org/10.1093/treephys/tpr104
Simard, S.W., & D.M. Durall. 2004. Mycorrhizal networks: a review of their extent, function, and
importance. Canadian Journal of Botany 82: 1140–1165. https://doi.org/10.1139/b04-116
———, A.P. David, M.D. Jones, D.D. Myrold, D.M. Durall & R. Molina. 1997. Net transfer of carbon
between ectomycorrhizal tree species in the field. Nature 388: 579–582.
Sinsabaugh, R.L. 2010. Phenol oxidase, peroxidase and organic matter dynamics of soil. Soil Biology and
Biochemistry 42: 391–404. https://doi.org/10.1016/j.soilbio.2009.10.014
Smith, D. & K.G. Peay. 2014. Sequence depth, not PCR replication, improves ecological inference from Next
Generation DNA Sequencing. PLoS One 92, e90234. https://doi.org/10.1371/journal.pone.0090234
Smith, S.E. & D.J. Read. 2008. Mycorrhizal symbiosis 3rd edition. Academic, New York.
——— & F.A. Smith. 2012. Fresh perspectives on the roles of arbuscular mycorrhizal fungi in plant nutrition
and growth. Mycologia 104: 1– 13. https://doi.org/10.3852/11-229
Smith, M.E, G.W. Douhan, D.M. Rizzo. 2007. Ectomycorrhizal community structure in xeric Quercus
woodland based on rDNA sequence analysis of sporocarps and pooled roots. New Phytologist 174:847–
863. https://doi.org/10.1111/j.1469-8137.2007.02040.x
———, T.W. Henkel, M.C. Aime, A.K. Fremier & R. Vilgalys. 2011. Ectomycorrhizal fungal diversity and
community structure on three co-occurring leguminous canopy tree species in a Neotropical rainforest.
———, ———, J.K. Uehling, A.K. Fremier, H.D. Clarke & R. Vilgalys. 2013. The ectomycorrhizal
fungal community in a neotropical forest dominated by the endemic dipterocarp Pakaraimea
dipterocarpacea. PLoS ONE 8: e55160. https://doi.org/10.1371/ journal. Po ne.0055160
Stonor, R.N., S.E. Smith, M. Manjarrez, E. Facelli & F.A. Smith. 2014. Mycorrhizal responses in wheat:
shading decreases growth but does not lower the contribution of the fungal phosphate uptake pathway.
Mycorrhiza 24(6): 465-472. https://doi.org/10.1007/s00572-014-0556-9
Suvi, T., L. Tedersoo, K. Abarenkov, J. Gerlach, K. Beaver & U. Kõljalg. 2010. Mycorrhizal symbionts of
Pisonia grandis and P. sechellarum in Seychelles: identification of mycorrhizal fungi and description of
new Tomentella species. Mycologia 102: 522–533. https://doi.org/10.3852/09-147
Szuba, A. 2015. Ectomycorrhiza of Populus. Forest Ecology and Management 347: 156-169. https://doi.
org/10.1016/ j.foreco.2015.03.012
Talbot, J.M., T.D. Bruns, D.P. Smith, S. Branco, S.I. Glassman, S. Erlandson, R. Vilgalys & K.G. Peay.
2013. Independent roles of ectomycorrhizal and saprophytic communities in soil organic matter decom-
position. Soil Biology and Biochemistry 57: 282–291. https://doi.org/10.1016/j.soilbio.20 12.10.004
Taniguchi, T., N. Kanzaki, S. Tamai, N. Yamanaka & K. Futai. 2007. Does ectomycorrhizal fungal
community structure vary along a Japanese black pine (Pinus thunbergii) to black locust (Robinia
pseudoacacia) gradient. New Phytologist 173: 322–334. https://doi.org/10.1111/ j.1469-
8137.2006.01910.x
———, R. Kataoka, & K. Futai. 2008. Plant growth and nutrition in pine seedlings (Pinus thunbergia)
seedlings and dehydrogenase and phosphatase activity of ectomycorrhizal root tips inoculated with seven
individual ectomycorrhizal fungal species at high and low nitrogen conditions. Soil Biology and
Biochemistry 40: 1235–1243. https://doi.org/10.1016/j.soilbio.2007.12.017
Tapwal, A., R. Kumar & S. Pandey. 2013: Diversity and frequency of macrofungi associated with wet ever
green tropical forest in Assam, India. Biodiversitas 14(2): 73-78. 10.13057/biodiv/ d140204
———, ——— D. Borah. 2015. Effect of mycorrhizal inoculations on the growth of Shorea robusta
seedlings. Nusantara Bioscience 7: 1-5. 10.13057/nusbiosci/n070101
Tatry, M.V., E.E. Kassis, R. Lambilliotte, C. Corratgé, I. Van Aarle, L.K. Amenc, R. Alary, S.
Zimmermann, H. Sentenac & C. Plassard. 2009. Two differentially regulated phosphate transporters
from the symbiotic fungus Hebeloma cylindrosporum and phosphorus acquisition by ectomycorrhizal
Pinus pinaster. Plant Journal 57: 1092–1102. https://doi.org/10.1111/j.1365-313X .2008.03749.x
Taylor, A.F.S. & I.J. Alexander. 2005. The ectomycorrhizal symbiosis: life in the real world. Mycologist 19:
102–112. https://doi.org/10.1017/S0269-915X(05)00303-4
———, G. Gebauer & D.J. Read. 2004. Uptake of nitrogen and carbon from double-labelled 15N and 13C
glycine by mycorrhizal pine seedlings. New Phytologist 164: 383–388. https://doi.org/10.1111/j.1469-
8137.2004.01164.x
Tedersoo, L. & M.E. Smith. 2013. Lineages of ectomycorrhizal fungi revisited: foraging strategies and novel
lineages revealed by sequences from belowground. Fungal Biology Reviews 27: 83–99. https://doi.
org/10.1016/j.fbr.2013.09.001
———, T. Suvi, E. Larsson & U. Kõljalg. 2006. Diversity and community structure of ectomy-corrhizal
fungi in a wooded meadow. Mycological Research 110: 734–748. https://doi.org/10.1016/j.mycres.20
06.04.007
———, ———, K. Beaver & U. Kõljalg. 2007. Ectomycorrhizal fungi of the Seychelles: diversity patterns
and host shifts from the native Vateriopsis seychellarum (Dipterocarpaceae) and Intsia bijuga
(Caesalpiniaceae) to the introduced Eucalyptus robusta (Myrtaceae), but not Pinus caribea (Pinaceae).
———, T. Jairus, B.M. Horton, K. Abarenkov, T. Suvi, I. Saar & U. Kõljalg. 2008. Strong host
preference of ectomycorrhizal fungi in a Tasmanian wet sclerophyll forest as revealed by DNA barcoding
and taxon- specific primers. New Phytologist 180: 479–490. https://doi.org/10.1111/j.1469-
8137.2008.02561.x
———, T.W. May & M.E. Smith. 2010a. Ectomycorrhizal lifestyle in fungi: global diversity, distribution,
and evolution of phylogenetic lineages. Mycorrhiza 20: 217–263. https://doi.org/10.1007/s00572-009-
0274-x
———, R.H. Nilsson, K. Abarenkov, T. Jairus, A. Sadam, I. Saar, M. Bahram, E. Bechem, G. Chuyong,
& U. Kõljalg. 2010b. 454 Pyrosequencing and Sanger sequencing of tropical mycorrhizal fungi provide
similar results but reveal substantial methodological biases. New Phytologist 188: 291–301. https://doi.
org/10.1111/j.1469-8137.2010.03373.x
J. Kumar, N.S. Atri
———, A. Sadam, M. Zambrano, R. Valencia & M. Bahram. 2010c. Low diversity and high host
preference of ectomycorrhizal fungi in Western Amazonia, a neotropical biodiversity hotspot. ISME
Journal 4: 465–471. https://doi.org/10.1038/ismej.2009.131
———, L., M. Bahram, T. Jairus, E. Bechem, S. Chinoya, R. Mpumba, M. Leal, E. Randrianjohany, S.
Razafi- mandimbison, A. Sadam, T. Naadel & U. Kõljalg. 2011. Spatial structure and the effects of
host and soil environments on communities of ectomycorrhizal fungi in wooded savannas and rain forests
of Continental Africa and Madagascar. Molecular Ecology 20: 3071–3080. https://doi.org/10.1111
/j.1365-294X.2011.05145.x
———, ———, M. Toots, A.G. Diédhiou, T.L. Henkel, R. Kjoller, M.H. Morris, K. Nara, E. Nouhara,
K. Peay, S. Polme, M. Ryberg, M.E. Smith & U. Kõljalg. 2012. Towards global patterns in the
diversity and community structure of ectomycorrhizal fungi. Molecular Ecology 21: 4160–4170.
https://doi.org/10.1111/j.1365-294X.2012.05602.x
———, ———, S. Põlme, U. Kõljalg, N.S. Yorou, R. Wijesundera, L.V. Ruiz, A.M. Vasco-Palacios, P.
Quang Thu, A. Suija et al. 2014. Global diversity and geography of soil fungi. Science 346(6213):
1256688. https://doi.org/10.1126/science.1256688
Tester, M., S.E. Smith, F.A. Smith & N.A. Walker. 1986. Effects of photon irradiance on the growth of
shoots and roots, on the rate of initiation of mycorrhizal infection and on the growth of infection units in
Trifolium subterraneum L. New Phytologist 103: 375–390. https://doi.org/10.1111/j.1469-8137.1986.
tb00623.x
Trappe, J.M. 1977. Selection of fungi for ectomycorrhizal inoculation in nurseries. Annual Review of
Phytopathology 15: 203-222. https://doi.org/10.1146/annurev.py.15.090177.001223
Treseder, K.K. 2004. A meta-analysis of mycorrhizal responses to nitrogen, phosphorus and
atmospheric CO2 in field studies. New Phytologist 164: 347–355. https://doi.org/10.1111
/j.1469-8137.2004.01159.x
Trocha, L.K., E. Weiser & P. Robakowski. 2016. Interactive effects of juvenile defoliation, light conditions,
and interspecific competition on growth and ectomycorrhizal colonization of Fagus sylvatica and Pinus
sylvestris seedlings. Mycorrhiza 26: 47–56. https://doi.org/10.1007/s00 572-011-0387-x
Turjaman, M., Y. Tamai, H. Segah, S.H. Limin, J.Y. Cha & M.O.K. Tawaraya. 2005. Inoculation with the
ectomycorrhizal fungi Pisolithus arhizus and Scleroderma sp. improves early growth of Shorea pinanga
nursery seedlings. New Forest 30: 67–73. https://doi.org/10.1007/s11056-004-1954-1
———, ———, ———, ———, M.K. Osaki & K. Tawaraya. 2006. Increase in early growth and nutrient
uptake of Shorea seminis seedlings inoculated with two ectomycorrhizal fungi. Journal of Tropical Forest
Science 18 (4): 243-249.
Uroz, S., C. Calvaruso, M.P. Turpault, J.C. Pierrat, C. Mustin & P. Frey-Klett. 2007. Effect of the
mycorrhizosphere on the genotypic and metabolic diversity of the soil bacterial communities involved in
mineral weathering in a forest soil. Applied and Environmental Microbiology 73:3019–3027. https://doi.
org/10.1128/AEM.00121-07
Utmazian, M.N.D.S., P. Schweiger, P. Sommer, M. Gorfer, J. Strauss & W.W. Wenzel. 2007. Influence of
Cadophora finlandica and other microbial treatments on cadmium and zinc uptake in willows grown on
polluted soil. Plant, Soil and Environment 53: 158–166.
van der Heijden, M.G.A. & T.R. Horton. 2009. Socialism in soil? The importance of mycorrhizal fungal
networks for facilitation in natural ecosystems. Journal of Ecology 97: 1139–1150. https://doi.org/10.1111
/j.1365-2745.2009.01570.x
———, J.N. Klironomos, M. Ursic, P. Moutoglis, R. Streitwolf-Engel, T. Boiler, A. Wiemken & I.R.
Sanders. 1998. Mycorrhizal fungal diversity determines plant biodiversity, ecosystem variability and
productivity. Nature 396: 69-72. https://doi.org/10.1038/23932
———, R.D. Bardgett & N.M. van Straalen. 2008. The unseen majority: soil microbes as drivers of plant
diversity and productivity in terrestrial ecosystems. Ecology Letters 11: 296–310. https://doi.org/10.1111
/j.1461-0248.2007.01139.x
———, F.M. Martin, M.A. Selosse & I.R. Sanders. 2015. Mycorrhizal ecology and evolution: the past, the
present, and the future. New Phytologist 205(4): 1406–1423. https://doi.org/10.1111/nph.13288
van der Putten, W.H., R.D. Bardgett, J.D. Bever, T.M. Bezemer, B.B. Casper, T. Fukami, P. Kardol, J.N.
Klironomos, A. Kulmatiski, J.A. Schweitzer, K.N. Suding, T.F.J. Van de Voorde & D.A. Wardle.
2013. Plant–soil feedbacks: the past, the present and future challenges. Journal of Ecology 101: 265–276.
https://doi.org/10.1111/1365-2745.12054
Van Tichelen, K.K., T. Vanstraelen & J.V. Colpaert. 2001. Ectomycorrhizal protection of Pinus
sylvestris against copper toxicity. New Phytologist 150: 203–213. https://doi.org/10.1046/j.1469-
8137.2001.00081.x
Villeneuve, N., F. Le Tacon & D. Bouchard. 1991. Survival of inoculated Laccaria bicolor in competition
with native ectomycorrhizal fungi and effects on the growth of outplanted Douglas fir seedlings. Plant
Soil 135:95-107. https://doi.org/10.1007/BF00014782
Vozzo, J.A. & E. Hacskaylo. 1971. Inoculation of Pinus caribaea with ectomycorrhizal fungi in Puerto Rico.
Forest Science 17: 239-245.
Walbert, K., T.D. Ramsfield, H.J. Ridgway & E.E. Jones. 2010. Ectomycorrhizal species associated with
Pinus radiata in New Zealand including novel associations determined by molecular analysis.
Wallander, H. 2000. Uptake of P from apatite by Pinus sylvestris seedlings colonized by different
ectomycorrhizal fungi. Plant Soil 218: 249–256. https://doi.org/10.1023/A:1014936217105
Wan, S.P., F-Q. Yu, L. Tang, R. Wang, Y. Wang, P-G. Liu, X-H. Wang & Y. Zheng. 2016.
Ectomycorrhizae of Tuber huidongense and T. liyuanum with Castanea mollissima and Pinus armandii.
Wang, Q. & L-D. Guo. 2010. Ectomycorrhizal community composition of Pinus tabulaeformis assessed by
ITS-RFLP and ITS sequences. Botany 88: 590–595. https://doi.org/10.1139/B10-023
———, C. Gao & L-D. Guo. 2011. Ectomycorrhizae associated with Castanopsis fargesii (Fagaceae) in a
subtropical forest China. Mycological Progress 10: 323–332. https://doi.org/10.1007/s 11557-010-0705-2
Wang, J., T. Li, X. Wu, & Z. Zhao. 2014. Molecular cloning and functional analysis of a H+-dependent
phosphate transporter gene from the ectomycorrhizal fungus Boletus edulis in southwest China. Fungal
Biology 118: 453–461. https://doi.org/10.1016/j.funbio.2014.03.003
Wang, X., J. Liu, D. Long, Q. Han & J. Huang. 2017a. The ectomycorrhizal fungal communities associated
with Quercus liaotungensis in different habitats across northern China. Mycorrhiza. https://doi.
org/10.1007/s00572-017-0762-3
Wang, L., B. Otgonsuren & D. L. Godbold. 2017b. Mycorrhizas and soil ecosystem function of co-existing
woody vegetation islands at the alpine tree line. Plant Soil 411:467–481. https://doi.org/10.1007/s11104-
016-3047-2
Waseem, M., M. Ducousso, Y. Prin, Q. Domergue, L. Hannibal, C. Majorel, P. Jourand & A. Galiana.
2017. Ectomycorrhizal fungal diversity associated with endemic Tristaniopsis spp. (Myrtaceae) in
ultramafic and volcano-sedimentary soils in New Caledonia. Mycorrhiza. https://doi.org/10.1007
/s00572-017-0761-4
Watling, R. & S.P. Abraham. 1992. Ectomycorrhizal fungi of Kashmir forests. Mycorrhiza 2: 81-87.
https://doi.org/10.1007/BF00203254
Ważny, R. 2014. Ectomycorrhizal communities associated with silver fir seedlings (Abies alba Mill.) differ
largely in mature silver fir stands and in Scots pine forecrops. Annals of Forest Science 71: 801–810.
https://doi.org/10.1007/ s13595-014-0378-0
Welander, N.T. & B. Ottosson. 1998. The influence of shading on growth and morphology in seedlings of
Quercus robur L. and Fagus sylvatica L. Forest Ecology and Management 107: 117–126. https://doi.
org/10.1016/S0378-1127(97)00326-5
Willmann, A., M. Weiss & U. Nehls. 2007. Ectomycorrhiza-mediated repression of the high-affinity
ammonium importer gene AmAMT2 in Amanita muscaria. Current Genetics 51: 71–78. https://doi.
org/10.1007/s00294-006-0106-x
———, S. Thomfohrde, R. Haensch & Nehls. 2014. The poplar NRT2 gene family of high affinity nitrate
importers: Impact of nitrogen nutrition and ectomycorrhiza formation. Environmental and Experimental
Botany 108: 79–88.
Wilson, A.W., K. Hosaka & G.M. Mueller. 2017. Evolution of ectomycorrhizas as a driver of diversification
and biogeographic patterns in the model mycorrhizal mushroom genus Laccaria. New Phytologist 213:
1862–1873. https://doi.org/10.1111/nph.14270
Wolfe, B.E. & A. Pringle. 2012. Geographically structured host specificity is caused by the range expansions
and host shifts of a symbiotic fungus. ISME Journal 6: 745–755. https://doi.org/10.1038/ismej.2011.155
———, F. Richard, H.B. Cross & A. Pringle. 2010. Distribution and abundance of the introduced
ectomycorrhizal fungus Amanita phalloides in North America. New Phytologist 185: 803–816.
https://doi.org/10.1111/j.1469-8137.2009.03097.x
———, R.E. Tulloss & A. Pringle. 2012. The irreversible loss of a decomposition pathway marks the single
origin of an ectomycorrhizal symbiosis. PLoS ONE 7: e39597. https://doi.org/10.1371/ journal.
pone.0039597
Wu, T., J.N. Sharda & R.T. Koide. 2003. Exploring interactions between saprotrophic microbes and
ectomycorrhizal fungi using a protein-tannin complex as an N source by red pine (Pinus resinosa).
J. Kumar, N.S. Atri
Xu, H., M. Kemppainen, W. El Kayal, S.H. Lee, A.G. Pardo, J.E.K. Cooke et al. 2015. Overexpression of
Laccaria bicolor aquaporin JQ585595 alters root water transport properties in ectomycorrhizal white
spruce (Picea glauca) seedlings. New Phytologist 205: 757–770. https://doi.org/10.1111/nph.13098
———, A. Navarro-Ródenas, J.E.K. Cooke & J.J. Zwiazek. 2016. Transcript profiling of aquaporins
during basidiocarp development in Laccaria bicolor ectomycorrhizal with Picea glauca. Mycorrhiza 26:
19–31. https://doi.org/10.1007/s00572-015-0643-6
Yamada, A., T. Ogura & M. Ohmasa. 2001. Cultivation of mushrooms of edible ectomycorrhizal fungi
associated with Pinus densiflora by in vitro mycorrhizal synthesis. II. Morphology of mycorrhizas in
open-pot soil. Mycorrhiza 11: 67–81. https://doi.org/10.1007/s005720000093
Yuwa-Amornpitak, T., T. Vichitsoonthonkul, M. Tanticharoen, S. Cheevadhanarak & S.
Ratchadawong. 2006. Diversity of ectomycorrhizal fungi on Dipterocarpaceae in Thailand. Journal of
Biological Sciences 6: 1059–1064. https://doi.org/10.3923/jbs.2006.1059.1064
Zak, B. 1973. Classification of ectomycorrhizae. In: G.C. Marks & T.T. Kozlowski (Eds.), Ectomycorrhizae,
Acadmic press, New York, pp. 43-78.
Zhang, H-H., M. Tang, H. Chen & C-L. Zheng. 2010. Effects of inoculation with ectomycorrhizal fungi on
microbial biomass and bacterial functional diversity in the rhizosphere of Pinus tabulaeformis seedlings.
European Journal of Soil Biology 46: 55–61. https://doi.org/10.1016/j.ejsobi.2009.10.005
Zhang, J., T. Taniguchi, R. Tateno, M. Xu, S. Du, G-B. Liu & N. Yamanaka. 2013. Ectomycorrhizal
fungal communities of Quercus liaotungensis along local slopes in the temperate oak forests on the Loess
Plateau, China. Ecological Research 28:297–305. https://doi.org/10.1007/s11284- 012-1017-6
Zheng, R., J. Wang, M. Liu, G. Duan, X. Gao, S. Bai & Y. Han. 2016. Molecular cloning and functional
analysis of two phosphate transporter genes from Rhizopogon luteolus and Leucocortinarius bulbiger,
two ectomycorrhizal fungi of Pinus tabulaeformis. Mycorrhiza 26(7): 633-644. https://doi.org/10.1007
/s00572-016-0702-7
Zoll, J., E. Snelders, P.E. Verweij & W.J.G. Melchers. 2016. Next-Generation Sequencing in the Mycology
Lab. Current Fungal Infection Reports 10: 37–42. https://doi.org/10.1007/s12281-016-0253-6
Zong, K., J. Huang, K. Nara, Y. Chen, Z. Shen & C. Lian. 2015. Inoculation of ectomycorrhizal fungi
contributes to the survival of tree seedlings in a copper mine tailing. Journal of Forest Research 20(6):
493–500. https://doi.org/10.1007/s10310-015-0506-1

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Bot. Rev.

Studies on Ectomycorrhiza: An Appraisal

Jitender Kumar 1 & N. S. Atri 1,2

# The New York Botanical Garden 2017

Abstract Ectomycorrhizal (ECM) fungi are obligate symbionts of dominant vascular

Keywords Ectomycorrhizal symbiosis . Ecosystem . Evolution . ECM diversity .

Evolutionary History of ECM:

Diversity and Specificity of ECM

Amhinema, Boletus, Hebeloma, Laccaria, Paxillus, Phialophora, Russula, Lactarius,

related fungi (Bidartondo et al., 2003). Leafy liverworts genera (Jungermanniales)

Factors Affecting ECM Diversity and Composition in Terrestrial

Morphoanatomical and Molecular Studies on ECM

Spruce with Lactarius deterrimus, L. picinus, Russula ochroleuca and R. xerampelina

Potential Role of Ectomcorrhiza to Sustain Terrestrial Ecosystems

Role in Growth and Establishment of Seedlings

Role in Regeneration of Forest Ecosystems

Role of ECM in Improving Soil Fertility

Sr.No Fungi Host Nutrient Refrences

1 Hebeloma crustuliniforme, Betula pendula N Abuzinadah & Read, 1986

Sr.No Fungi Host Nutrient Refrences

Sr.No Fungi Host Nutrient Refrences

28 Paxillus involutus Picea abies Mg Jentschke et al., 2000

Sr.No Fungi Host Nutrient Refrences

Sr.No. Fungi Host Heavy metal Refrences

1 Suillus luteus Pinus sylvestris Zn Adriaensen et al., 2004

Sr.No. Fungi Host Heavy metal Refrences

19 Suillus luteus Pinus sylvestris Zn Krznaric et al., 2010

25 Suillus bovines, Pinus sylvestris Cu Van Tichelen et al., 2001

ECM Role in Amelioration of Heavy Metal Stress:

Ahonen-Jonnarth and Finlay (2001) reported that Laccaria bicolor is a potential

Role of ECM in Disease Resistance to Plants:

Rhizoctonia solani, Sclerotium rolfsii, and Subramanispora vesiculosa. The antagonis-

Role of ECM in Drought and Salt Tolerance:

ECM in Carbon Cycling and Decomposition of Organic Matter:

decomposition primarily through increased nitrogen mobilization rather than through

Ectomycorrhizal fungi represent an important component of soil microbiota and

Dipterocarpaceae, Fagaceae and Myrtaceae with fungal symbionts belonging to the

Phytophthora cinnamomi infections. Forestry 88: 257 –266. https://doi.org/10.1093/ forestry/

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