SEARCH

SEARCH BY CITATION

Keywords:

  • truffles;
  • Tuber melanosporum ;
  • ectomycorrhiza;
  • ITS ;
  • haplotype;
  • microbial community

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Conclusions
  8. Acknowledgements
  9. Authors' contribution
  10. References
  11. Supporting Information

Truffles are hypogeous ectomycorrhizal (EM) fungi belonging to the genus Tuber. Although outplanting of truffle-inoculated host plants has enabled the realization of productive orchards, truffle cultivation is not yet standardized. Therefore, monitoring the distribution of fungal species in different truffle fields may help us to elucidate the factors that shape microbial communities and influence the propagation and fruiting of Tuber spp. In this study, we compared the fungal biodiversity in cultivated and natural Tuber melanosporum truffle fields located in Central Italy. To this end, ectomycorrhizas (ECM) and soil samples were molecularly analyzed, and an inventory of the fungi associated with Quercus pubescens plants colonized by T. melanosporum, Tuber aestivum or Tuber brumale was compiled. T. melanosporum and T. aestivum were dominant on the cultivated plants, and the number of EM species was markedly lower in the cultivated sites than in the natural sites. However, in the same site, EM biodiversity was higher in T. brumale-colonized plants than in T. melanosporum-colonized plants. These results suggest that different Tuber spp. may have different competitive effects on the other mycobionts. Additionally, in keeping with our previous findings, we found that the number of T. melanosporum genotypes recovered from the soil samples was higher than that of the underlying ECM.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Conclusions
  8. Acknowledgements
  9. Authors' contribution
  10. References
  11. Supporting Information

Tuber spp. are symbiotic ascomycetes that must establish a mutualistic interaction with the roots of their host plants for vegetative propagation and fruiting body formation. Because of the exceptional flavor of its fruit bodies, Tuber melanosporum, also known as the Périgord Black truffle, is one of the most appreciated truffle species worldwide. Reliable nursery procedures of root inoculation to produce host plants colonized with T. melanosporum were established in 1970s and have enabled large-scale cultivation programs not only in Europe, where this fungus is endemic, but also in Australia, Chile, New Zealand, and the USA. These programs are aimed at counteracting the declining spontaneous production of this fungus, satisfying the increasing worldwide demand and producing truffles for out-of-season Northern Hemisphere markets (Garland, 1999; Hall et al., 2003). However, not all black truffle orchards are productive and, even in productive plantations, some plants never produce ascocarps. Thus, black truffle cultivation cannot yet be considered a fully standardized and reliable agronomic procedure.

Recently, our understanding of the reproductive system of this species has increased considerably (Riccioni et al., 2008; Martin et al., 2010; Rubini et al., 2011c), and it is expected that the acquisition of basic information on the truffle life cycle will positively impact truffle field management in the near future. However, ecological factors controlling both the vegetative and reproductive spread of these fungi remain largely undeciphered. Therefore, studies aimed at assessing fungal biodiversity and dynamics in natural and cultivated truffle grounds will help us to shed light on the underground microbial network that may play a pivotal role in controlling truffle-host plant colonization, hyphal growth and fructification.

Several studies have been performed on the biodiversity of ectomycorrhizas (ECMs) in different truffle grounds (Chevalier et al., 1982; Granetti & Angelini, 1992; Donnini & Bencivenga, 1995; Granetti & Baciarelli Falini, 1997; Etayo & De Miguel, 1999; Ferrara et al., 1999; De Miguel & Sáez, 2005; Garcia-Barreda & Reyna, 2011). In most of these studies, which generally have focused on cultivated orchards, the different ectomycorrhizal (EM) species have been typed using morphological methods. Accurate species sorting, however, may be difficult if based only on morphotyping. Indeed, species-specific morphological traits used to distinguish among ECMs of different species are few and highly influenced by environmental conditions, age and the host plant species (Egger, 1995). The development of molecular markers has provided mycologists with new tools to evaluate EM fungal biodiversity. Markers based on the internal transcribed spacer (ITS) region of the rDNA locus are now broadly used to study ECMs in various environments (Horton & Bruns, 2001). For example, molecular typing has been recently used to study the EM communities in T. melanosporum and Tuber aestivum orchards (Baciarelli Falini et al., 2006; Pruett et al., 2008; Benucci et al., 2011) and in natural Tuber magnatum and Tuber borchii truffle grounds (Murat et al., 2005; Bertini et al., 2006; Iotti et al., 2010). Napoli et al. (2010) compared the fungal communities in natural and cultivated T. melanosporum truffle grounds within and outside the ‘brulé’, that is, the ‘burnt’ zone next to truffle-colonized plants characterized by the scarcity or complete absence of understory vegetation (Fasolo-Bonfante & Fontana, 1971; Pacioni, 1991) and reported that inside the brulé, T. melanosporum reduced the richness of other EM species. Indeed, the competition of fungal species may play a significant role in determining the structure of EM communities (Kennedy & Bruns, 2005; Kennedy, 2010). Therefore, the analysis of EM communities associated with both natural and cultivated productive truffle grounds may help to deepen our insight into the ecology of T. melanosporum and to ultimately annotate those EM species that might antagonistically interact with truffles for niche and host plant colonization.

To this end, we performed an analysis of ECMs and soil fungi in both cultivated and natural T. melanosporum truffle grounds to achieve the following goals: (1) to compare the biodiversity of EM species in natural and cultivated T. melanosporum truffle grounds; (2) to identify potential fungal competitors with respect to T. melanosporum; and (3) to gain preliminary insights into T. melanosporum strain biodiversity in different truffle grounds via ITS haplotyping. Additionally, the presence, within the T. melanosporum-cultivated sites, of plants originally inoculated with T. aestivum and of plants colonized by Tuber brumale provided us with an opportunity to compare the fungal biodiversity that exists on and next to the host plants that sustain different Tuber spp. on their roots.

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Conclusions
  8. Acknowledgements
  9. Authors' contribution
  10. References
  11. Supporting Information

Sites description

The sites under investigation were two natural and two man-made T. melanosporum truffle grounds located in Central Italy. The natural truffle grounds were located near Leonessa (Rieti), in the surrounding areas of Vallunga (VNM; 42°34′57.91″N, 12°59′53.88″E) and Villa Gizzi (VGNM; 42°36′31.49″N, 12°57′57.49″E). These sites are at an altitude of approximately 900 m. The predominant tree is the downy oak (Quercus pubescens Willd.), with a patchy distribution (10–15 m of distance between trees). The productive trees were selected with the help of local truffle pickers, and all of the trees showed the characteristic brulé. The first cultivated plantation is located only 200 m from the natural truffle ground of Vallunga (42°35′10.23″N; 13°0′5.20″E). It was established in 1998 using Q. pubescens plants inoculated with T. melanosporum (VCM) in the eastern part and T. aestivum in the western part (VCA) of the truffle ground. This truffle plantation has been considered as unproductive because neither T. melanosporum nor T. aestivum truffles were ever harvested by the owner. The second cultivated plantation is located in Montemartano (42°47′26.49″N, 12°35′24.64″E) near Spoleto (Central Italy). It was established in 1995 with Q. pubescens plants inoculated with T. melanosporum (MCM). Most of the trees have produced Tmelanosporum truffles since 2002, but some trees only produced the competing species T. brumale (MCB).

The natural and cultivated truffle grounds were selected to have similar pedo-ecological characteristics All truffle grounds present more than 30% of stones. Granulometric fractions content on average consist of 34.5 ± 3.7% clay, 42.5 ± 2.2% silt, and 23 ± 2.5% sand. The average content of organic matter is 3.92 ± 0.5% and pH is approximately 8.

ECM and soil sampling

Root samples were collected in July and October 2007 and January and April 2008. In each truffle ground, three oak plants were selected, and two samples were collected from each plant from opposite directions (north-south) and then pooled. In total, 12 root samples for each truffle ground (VCM, MCM, VNM, and VGNM) were analyzed. Twelve additional root samples from three oak plants producing T. brumale in the plantation of Montemartano (MCB) and from three oak plants inoculated with T. aestivum in the plantation of Vallunga (VCA) were sampled and analyzed.

Root samples were collected at a depth of approximately 15–20 cm using a soil corer, and the samples were stored at 4 °C until processed. Each root sample was soaked in water and sieved to separate the root fragments and ECMs from the soil.

In July 2007 and January 2008, two soil samples were collected from the brulé areas of VCM, MCM, VNM, and VGNM trees after the removal of the first 3-cm layer. For each of the four truffle ground, the soil samples (~ 3 g from each sample) were pooled, stored at −20 °C and used for DNA isolation.

Morphological analysis of ECMs

Each root sample was carefully examined to identify all EM morphotypes, according to Agerer (1986, 1991). The morphology and color of the ECMs were evaluated with a dissecting microscope on freshly isolated root tips. The features of the mantle surface and emanating elements were examined on ECMs mounted in 50% glycerol using a light microscope. The different EM morphotypes were collected and stored in 95% ethanol. Single ECMs were also stored at −80 °C and later used for molecular analysis.

Molecular identification of ECMs

Genomic DNA was isolated from single ECMs as described by Paolocci et al. (1999). The ITS region of the rDNA was amplified by PCR using the ITS1/ITS4 primer pair (White et al., 1990) according to Paolocci et al. (1999). The PCR products were purified using a Jet-Quick spin column (Genomed) and directly sequenced using a BigDye terminator sequencing kit (Applied Biosystems) according to the instructions of the supplier. Sequencing reactions were run on an ABI 3130 Genetic Analyzer (Applied Biosystems) and the resulting sequences were deposited in GenBank under the following accession no. JF926843JF927155.

Isolation and amplification of DNA from soil organisms

DNA was isolated from 0.25 g of soil using the PowerSoil DNA Isolation Kit (Mo Bio) according to the manufacturer's instructions. Isolated DNA was visualized by electrophoresis on an agarose gel containing ethidium bromide and quantified by comparisons with DNA mass ladders (Invitrogen).

Soil DNA was analyzed by PCR amplification of the ITS region using the ITS1F/ITS4 primers pair (White et al., 1990; Gardes & Bruns, 1993). PCR amplifications were carried out in a Gene Amp 9700 PCR system (Applied Biosystems) with the following cycling parameters: an initial denaturation step at 95 °C for 3 min; 25 cycles consisting of 30 s at 95 °C, 30 s at 55 °C, and 45 s at 72 °C, with a final extension of 7 min at 72 °C. PCRs were performed in a 50-μL reaction mixture containing 200 μM of each dNTP, 10 pmol of each primer, 4 mM MgCl2, 10 mM Tris–HCl (pH 9.0), 50 mM KCl, 2.5 units of Taq polymerase (GE healthcare), and 10–20 ng of target DNA. All of the PCR experiments included a negative control (no DNA template).

The obtained PCR products were used to generate ITS clone libraries. For this purpose, approximately 20 ng of amplified DNA was ligated into the pGEM-T Easy (Promega) plasmid and used to transform competent E. coli cells following standard protocols (Sambrook et al., 1989). In total, four soil ITS clone libraries, one for each T. melanosporum truffle ground, were produced.

Individual colonies were picked and boiled in 20 μL of sterile water, and the DNA was PCR amplified with ITS1F/ITS4 primers. Each PCR product (5 μL) was run on a 1.5% agarose gel and visualized with ethidium bromide. The amplicons showing different sizes were purified using the JetQuick PCR purification Kit (Genomed) and sequenced. In total, from 40 to 60 clones were sequenced from each library.

The presence of T. melanosporum DNA in the soil samples was also verified using the species-specific ITS primers pair, ITSml/ITS4lng, according to Paolocci et al. (1999).

Sequence analysis

Sequence assembly and editing were performed using the Bioedit software (Hall, 1999). Operational taxonomic units (OTUs) were defined using Sequencher (Genecodes Corporation), considering the same OTU sequences with ≥ 97% similarity and without taking into account the differences in sequence length (Arnold et al., 2007). For each OTU, the longest ITS sequence was selected and analyzed for the similarity with other sequences in the GenBank database using a blast search.

Data analysis

The diversity of EM communities in each truffle ground was evaluated using different estimators based on the abundance and frequency of ECMs. The OTU frequency and abundance were calculated for each tree and for each truffle ground. Because the ECMs with a similar morphotype from a given root sample always exhibited an identical ITS sequence (see Results), they were considered to belong to the same OTU. The relative frequency of each OTU was calculated as follows: inline image, where Xi is the number of root or soil samples on which a given OTU was detected (Paulus et al., 2006).

The relative abundance was defined as the number of ECMs of a given OTU divided by the total number of ECMs detected in each tree or in each truffle ground. The abundance was evaluated from the root samples collected in October 2007 by performing a subsampling of roots, according to Richard et al. (2005). More specifically, from each plant, 20 distinct aggregates of ECMs were randomly selected, and all of the ECMs of each morphotype, for a total of 3292, were counted.

Species richness (S), consisting of the total number of OTUs in a given truffle ground, Simpson's diversity index (D) and Shannon–Weaver information index (H) (Shannon, 1948) were calculated using Estimates v. 8.2.0 (Colwell, 2005). Bray–Curtis and Jaccard dissimilarity indices and hierarchical clustering (HCLUST, using the average linkage option) were calculated using R 2.9 software (R Development Core Team, 2009) with the ‘vegan’ package v. 11.1-4 (Oksanen et al., 2010). Correlation tests were performed using Xlstat v. 7.5.2.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Conclusions
  8. Acknowledgements
  9. Authors' contribution
  10. References
  11. Supporting Information

Identification and comparison of EM fungi in the root samples from natural and cultivated grounds

The ECMs were identified by a combination of morphological and molecular analyses. Among all of the samples, 28 morphotypes were detected, and their description is provided in the Supporting information, Table S1.

For each plant, all of the different EM morphotypes were collected, and the ITS region was PCR amplified and sequenced. As two or more single, randomly peaked ECMs for each morphotype per plant always showed identical ITS sequences (or sequences with similarity levels higher than 97%; data not shown), ECMs with similar morphotypes were assigned to the same OTU.

In total, considering all truffle grounds and seasonal sampling, 128 ECMs were molecularly analyzed. Grouping of the ITS sequences on the basis of 97% nucleotide similarity revealed 43 OTUs (Tables 1 and 2). The blast analyses revealed that 51% (22 OTUs) were basidiomycetes and 35% (15 OTUs) were EM ascomycetes. In a few samples (OTUs 66, 67, and 77), PCR-amplified saprophytic or pathogenic fungi that were likely associated with the EM fungus on the root tips. OTUs 5 and 65 were root endophytic fungi, and OTU 68 was ascribed to glomeromycetes (Table 1).

Table 1. OTUs detected and molecular identification
OTUSourceaEcologybPhylumcMorphotypeblastn tentative identification
SpeciesAccession no.Query Length (bp)Query Coverage (%)Identity (%)
  1. Morphotype descriptions are reported in Table S1.

  2. a

    R = root, S = soil.

  3. b

    EM = ectomycorrhizal, AM = arbuscular mycorrhizal, E = endophyte, SA = saprobic, PA = plant pathogen.

  4. c

    B = Basidiomycota, A = Ascomycota, G = Glomeromycota, Z = Zygomycota.

1S + REMBT1Sistotrema muscicola (Corticiaceae) AJ606041 7137385
2S + REMAT2Tuber melanosporum (Tuberaceae)U8935962010099
3S + REMBT3Russula brevipes (Russulaceae) FJ845429 70810096
4S + REMBT4Tricholoma scalpturatum (Tricholomataceae) EU160589 7279499
5REAThysanorea papuana (Herpotrichiellaceae) EU041814 6089091
6SUncultured fungus FJ197879 7719395
7SSAAMicrodochium bolleyi (Xylariales) GU566298 6796697
8SSAAPenicillium sp. (Trichocomaceae) FJ376592 5868781
9SSAAPenicillium sp. (Trichocomaceae) FJ379806 5926486
10S + REMAT5Cenococcum geophilum (Dothideomycetes) EU427331 59210098
11S EMBHymenogaster citrinus (Hymenogastraceae) EU784360 6979898
12S EMBUncultured ectomycorrhiza (Hymenogastraceae) AY634136 70210099
13SEMAUncultured ectomycorrhiza (Sordariomycetes) FJ554320 6908393
14SSAZUncultured zygomycete (Mortierellaceae) EF027378 6949796
15SSABConocybe sp. (Bolbitiaceae) AY194553 6486488
16SAMGGlomus etunicatum (Glomeraceae) AY236328 6169995
17SAMGUncultured Glomus (Glomeraceae) GQ388467 5759798
18SEMBUncultured Tomentella (Thelephoraceae) AF184746 66610094
19SEMAPhaeangium lefebvrei (Pyronemataceae) AF387653 6559890
20SEAExophiala sp. (Herpotrichiellaceae) GQ302685 6759497
21SSABLycoperdon rimulatum (Lycoperdaceae) EU833664 3339498
22SPAAFusarium oxysporum (Hypocreales) GU566205 5839999
23SSAZUncultured zygomycete EU490026 6789995
24SAUncultured Ascomycota HM239907 6179998
25SEMBHygrocybe persistens (Hygrophoraceae) FM208872 6908894
26SEMBHygrocybe conica (Hygrophoraceae) EU784300 70310092
27SPAATruncatella angustata (Amphisphaeriaceae) GU566260 62410099
28SE/PAARhizopycnis vagum (Ascomycota) AF022786 5809399
29SE/PAAFusarium sp. (Hypocreales) HQ130713 5766697
30SSA/PAAUncultured Hypocreales (Hypocreales) GU0556802 4929881
31SSAAUncultured Sordariales (Sordariales) DQ182442 5607587
32S + REMAT6Trichophaea woolhopeia (Pyronemataceae) DQ200835 5959199
33SEMATrichophaea woolhopeia (Pyronemataceae) DQ200835 5989196
34REMAT7Tuber rufum (Tuberaceae) FM205636 64595100
35SSAALophodermium minor (Rhytismataceae) AY100665 440100100
36SSABCryptococcus aerius (Filobasidiales) AF444376 6809499
37SAUncultured Ascomycete AY970123 6279190
38SSAZMortierella sp. (Mortierellaceae) GQ302682 6849498
39S + REMAT8Tuber aestivum (Tuberaceae)AY226040.7109798
40REMAT9Hydnobolites cerebriformis (Pezizaceae) EU784271 5879991
41SEMBSebacina incrustans (Sebacinaceae) AF490395 6289994
42SUncultured soil fungus DQ421205 6796696
43SUncultured soil fungus DQ421190 7389999
44SBUncultured Basidiomycota HM240166 6919997
45REMBT10Uncultured Cortinarius (Cortinariaceae) EU668287 5989999
46SSA/PAAUncultured Hypocreales (Hypocreales) GU055550 53210083
47S + REMBT10Hebeloma leucosarx (Cortinariaceae) AB211268 7009997
48REMBT10Hebeloma cf. oculatum (Cortinariaceae) DQ974696 40399100
49REMBT11Scleroderma cepa (Sclerodermataceae) EU784412 7279899
50REMAT7Tuber rufum (Tuberaceae) AF106892 7049285
51REMAT12Peziza cf. succosa (Pezizaceae) EU819417 60410096
52REMAT13Tuber brumale f. moschatum (Tuberaceae) AF001010 803100100
53REMAT14Uncultured Genea (Pyronemataceae) EU668290 7546083
54REMBT15Uncultured Tomentella (Thelephoraceae) FJ581421 58110093
55REMBT16Tomentella sp. (Telephoraceae) AJ879642 6339099
56REMBT15Tomentella sp. (Telephoraceae) DQ974780 64310095
57S + REMBT16Uncultured Tomentella (Thelephoraceae) EU668199 67610096
58REMBT16Tomentella sp. (Thelephoraceae) AJ534912 68210044
59S EMBUncultured Tomentella (Thelephoraceae) EU625888 7059993
60REMBT15Tomentella sp. (Thelephoraceae) EU668208 66110096
61REMBT15Thelephora terrestris (Thelephoraceae) EU819444 6859992
62REMBT16Uncultured ECM (Tomentella) (Thelephoraceae) EF218826 65810095
63REMBT16Uncultured ECM (Tomentella) (Thelephoraceae) AY748876 6779998
64REMBT17Uncultured Tomentella (Thelephoraceae) GQ469534 543100100
65RET18Fungal endophyte AF373058 3008796
66RSA/PAAT19Uncultured Nectriaceae DQ182425 7117999
67RSAABisporella citrina (Helotiales) GQ411507 5779590
68RAMGT20Glomus sp. (Glomeraceae) AY174691 42510093
69REMAT14Geopora arenicola (Pyronemataceae) FM206450 45210094
70REMBT21Uncultured Inocybe (Cortinariaceae) FJ210736 559100100
71S + REMAT22Tuber maculatum (Tuberaceae) FM205511 6968899
72S + REMBT23Uncultured Sebacina (Sebacinaceae) EU668260 64610099
73REMBT24Clavulina cinerea (Clavulinaceae) AJ889937 27810096
74S + REMBT25Uncultured Tomentella (Thelephoraceae) FJ378804 7549395
75REMAT14Geopora sp. (Pyronemataceae) FM206471 5539998
76REMAT26Tarzetta catinus (Pyronemataceae) DQ200833 5719991
77RPAAT27Cadophora luteo-olivacea (Helotiales) DQ404348 59810097
78REMAT24Uncultured ectomycorrhiza (Clavulinaceae) GQ254856 40010099
79S + REMBT28Inocybe umbrinella (Cortinariaceae) FJ904165 7299894
80SEMBInocybe umbrinella (Cortinariaceae) FJ904165 7259899
81SEMBUncultured ectomycorrhiza (Sclerodermataceae) EF644144 7779099
82SEMBUncultured Scleroderma (Sclerodermataceae) FJ197962 6806697
83SSAAPodospora didyma (Lasiosphaeriaceae) AY999127 5849789
Table 2. Distribution of ECM taxa in the plant roots
OTUVCAVCMMCMMCBVNMVGNM
P1P2P3P4P5P6P7P8P9P10P11P12P13P14P15P16P17P18
  1. Sample collections as follows: July 2007 (a); October 2007 (b); January 2008 (c); and April 2008 (d). P1 to P18 indicate the sampled plants.

1            d d   
2    abababcdabcdabcdcd d  abcdabcdad
3            ab  ad 
4             abd   ab
5        c         
10           bdabacdbbdbabcd
32 c    bd  acaabcc    b
34       b         b
39abcdabcdabcdabcdbcdbcd   ab     b c
40            a     
45                b 
47            a a   
48            d     
49         abc       d
50               d cd
51            cbb a 
52         bcdbcdb      
53                b 
54      bb          
55           bc      
56          bc    d d
57          d    b bcd
58         bab       
60           cc  bcac 
61        b   b b bb
62            a     
63             b    
64         bd   a    
65                 ac
66        a    d    
67             b    
68                 ac
69        b         
70          b       
71           b      
72               d  
73               a  
74    bb     b   d  
75             d    
76               d  
77             d d  
78                 c
79               d  

A very low number of EM species was detected in the cultivated T. melanosporum truffle grounds, irrespective of the sampling season (Table 2). As such, in VCM and MCM plants, 3 and 8 species, respectively, were detected. Such a low number of EM species was also detected in VCA plants, where only one species in addition to T. aestivum was identified. Interestingly, T. aestivum turned out to be the dominant species not only on VCA but also on VCM plants (Fig. 1a and b, Tables S2 and S3), conversely, T. melanosporum was the dominant species in MCM plants, being the most abundant and the most frequent (Table 2, Fig. 1c). A higher number of OTUs (15) was detected on plants producing T. brumale (MCB) from the same truffle ground. In this case, the most frequent species was T. brumale (OTU 52), followed by Trichophaea woolhopeia (OTU 32), some basidiomycetes (OTUs 49 and 58) and Tmelanosporum (OTU 2), whereas the most abundant were Tomentella spp. (OTU 74 and 64), followed by Inocybe sp. (OTU 70), T. brumale, T. puberulum (OTUs 52 and 71) and some Telephoraceae (OTUs 55, 56; Fig. 1d).

image

Figure 1. Dominance diversity curves for each truffle ground. The lines and bars indicate the OTU relative frequency and abundance, respectively. The OTU numbers are reported as in Table 1.

Download figure to PowerPoint

In the natural truffle grounds, the fungal diversity at root level was higher than in the cultivated sites: 19 and 25 species were recorded in VNM and VGNM, respectively (Table 2). In the natural truffle ground of Vallunga, the EM community was dominated by Cenococcum geophilum (OTU 10), followed by T. melanosporum. These two species were also the most abundant, together with a Thelephora sp. (OTU 61) (Fig. 1e).

Similarly, in the natural truffle ground of Villa Gizzi, the most frequent species detected were Cenococcum geophilum and T. melanosporum. Cenococcum geophilum was also one of the most abundant, together with some Telephoraceae (OTUs 60, 57, and 61) and T. rufum (OTU 34) but not T. melanosporum (Fig. 1f).

Diversity indices based on abundance and frequency confirmed that the two natural sites presented a higher level of diversity than the cultivated ones. The only exception was the T. brumale-producing plants from Montemartano, which showed a diversity of ECMs similar to that found in the two natural sites (Table 3).

Table 3. ECM diversity of the studied sites
Diversity indexCultivated sitesNatural sites
VCAVCMMCMMCBVNMVGNM
  1. S = Species richness; H = Shannon index; D = Simpson index

  2. a

    Calculated from frequency data of all of the samples.

  3. b

    Calculated from abundance data of the samples collected in October 2007.

S238151925
Ha0.270.91.492.52.742.98
1/Db1.172.132.819.9912.6315.57
Hb00.512.061.651.84
1/Db11.331.96.154.554.88

The number of EM species was lower in plants showing a high abundance of T. melanosporum or T. aestivum mycorrhizas: the Spearman's test indicated a significant negative correlation between the relative abundance (r2 = −0.890, P < 0.0001, α = 0.05) or the relative frequency (r2 = −0.879, P < 0.0001, α = 0.05) of T. melanosporum T. aestivum ECMs and the number of total EM species (species richness) detected on each plant (Fig. 2).

image

Figure 2. Correlation between the relative abundance (a) or relative frequency (b) of Tuber melanosporum Tuber aestivum ECMs and the number of species per tree.

Download figure to PowerPoint

Similarity indices based on incidence (Jaccard) and frequency (Bray–Curtis) data and cluster analysis (Table S4, Fig. S1) indicated close relationships between VCM and VCA, whereas the cultivated sites of Montemartano MCM and MCB were more closely related to the two natural sites (VNM and VGNM). Similar results were obtained using Sørensen and Morisita indices (data not shown).

Cenococcum geophilum (OTU 10) ECMs were shared by all of the natural plants and were detected on the plant 12 from MCB but not on the other cultivated plants. Similarly, OTUs 61 and 51 were present in most of the natural plants but were not or were rarely detected in the cultivated sites.

Characterization and comparison of fungal species from soil clone libraries

From the analysis of ITS clone libraries from the four T. melanosporum truffle grounds, 53 fungal taxa were found and, among them, 26 were EM fungi (18 basidiomycetes and eight ascomycetes), half of which also found at the root level (Table 1).

Non-EM fungal species (21 OTUs), most of which were amplified from the VCQM truffle ground (14 OTUs), consisted of mitosporic ascomycetes, basidiomycetes, zygomycetes, and glomeromycota. Furthermore, six OTUs were of undetermined classification (Tables 1 and 4).

Table 4. Distribution of fungi from soil clone libraries in the sampling sites
EcologyOTUMCMVCMVNMVGNM
  1. Sample collections as follows: July 2007 (a) and January 2008 (c).

EM1a ac 
2ac a 
3a aa
4  ac
10a aca
11a aac
12  cc
13  c 
18  a 
19   c
25   a
26  a 
32c   
33 c  
39 ac  
41  a 
47a   
57   c
59 c  
71  c 
72  cc
74  c 
79  cc
80  c 
81   c
82   c
Non-EM7  c 
8 a  
9 a a
14 a  
15 a  
16   c
17   c
20 c  
21  c 
22 c  
23 c  
27 c  
28 c  
29 c  
30 a  
31 a  
35a   
36 c  
38c   
46  a 
83 c  
Undetermined6 a  
24 c  
37 a  
42  c 
43  c 
44  c 

The cultivated truffle grounds showed a lower number of EM species than natural sites. More specifically, in soil samples from VCM and MCM, only three and seven OTUs, respectively, relative to EM fungi were found (Table 4). In the soil samples from VCM site, two EM species were found in addition to T. melanosporum (see the analysis with species-specific primers reported below) and T. aestivum: T. woolhopeia (OTU 33) and a Tomentella sp. (OTU 59) (Table 4, Fig. 3a). In the soil samples from the MCM site, besides the two most abundant species, that is, T. melanosporum and T. woolhopeia (OTUs 2 and 32) present at root level, five additional EM species were identified as soil free-living mycelia (OTUs 1, 3, 10, 11, and 47) (Table 4, Fig. 3b).

image

Figure 3. Venn diagram showing the number of EM species detected in the root (R) and soil (S) samples.

Download figure to PowerPoint

In the soil from the two natural truffle grounds (VNM and VGNM), 16 and 12 OTUs, respectively, relative to EM species were detected. On VNM and VGNM truffle grounds, only five (OTUs 1, 2, 3, 4, and 10) and six (OTUs 3, 4, 10, 57, 72, and 79) species, respectively, were shared between soil and root samples (Table 4, Fig. 3c and d). In both cases, these OTUs were among the most abundant and frequent species detected at the root level (Fig. 1e and f).

As shown in Table 4, some of the soil EM fungi were shared between the truffle grounds. Five (OTUs 1, 2, 3, 10, and 11) and three (OTUs 3, 10, and 11) species from MCM were also present in the VNM and in VGNM natural sites, respectively, and the two natural sites shared seven EM species (OTUs 3, 4, 10, 11, 12, 72, and 79).

In contrast, VCM site did not share any EM species with the other sites in addition to T. melanosporum.

Non-EM fungi were detected predominantly from the VCM site (14 OTUs), and only OTU 9 was shared between two sites.

Distribution and genetic variability of T. melanosporum

Tuber melanosporum ECMs were found in both the natural and cultivated sites. T. melanosporum ECMs were also found on two of the three T. brumale-producing plants sampled in the Montemartano (MCB) truffle ground but not on plants originally inoculated with T. aestivum in the truffle ground of Vallunga. Analyses of soil clone libraries confirmed the presence of T. melanosporum in the natural truffle ground of Vallunga and in the truffle plantation of Montemartano but not in the cultivated plantation of Vallunga and in the natural truffle ground of Villa Gizzi. However, the presence of this fungus in the soil of all of these truffle grounds was detected using T. melanosporum ITS species-specific primers. Using the sequences obtained from both the ECMs and soil clone libraries, the ITS haplotypes I, II, and IV, which were previously described by Murat et al. (2004) and Riccioni et al. (2008), were detected. More specifically, haplotypes I, II and IV were detected in the soil clone libraries from MCM, whereas ITS haplotypes I and IV were detected in the VNM soil samples. The analysis of T. melanosporum ECMs from VNM, VCM, MCB, and MCM revealed the presence of ITS haplotype I only.

Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Conclusions
  8. Acknowledgements
  9. Authors' contribution
  10. References
  11. Supporting Information

The analysis of the community ecology underneath truffle grounds has the potential to deepen our understanding of microbial determinants that might actively control the vegetative spreading of Tuber spp. By performing a parallel morphotyping and molecular typing of ECMs and a molecular typing of soil fungi associated to T. melanosporum fields here we show that T. melanosporum natural truffle grounds are more species-rich than the cultivated ones and that under similar ecological conditions the fungal community is likely shaped by species-specific compounds secreted by Tuber ECMs.

Fungal biodiversity differs between natural and cultivated truffle grounds

Our extensive morphological and molecular survey of the fungal communities from four Tmelanosporum grounds allowed the molecular identification of more than 80 OTUs, 50 of which were EM species. For the fungal species detected as ECMs, a preliminary morphological description is also provided. A reduction in the number of EM species on plants from cultivated truffle grounds compared with those from natural sites was detected; in the two cultivated sites, species diversity was significantly lower on plants highly colonized by T. melanosporum and T. aestivum.

A low number of EM species in cultivated T. melanosporum plants was previously reported. Baciarelli Falini et al. (2006) detected only one or two morphotypes in plants highly colonized by T. melanosporum. A recent study conducted on a T. aestivum-cultivated orchard found that fungal EM communities vary with the host species at root level (Benucci et al., 2011). As such, the community of EM fungi on hazel plants colonized by T. aestivum was found to be richer in terms of taxa than T. aestivum-colonized hornbeam plants grown in the same environment. However, whether this effect depends on the abundance of T. aestivum ECMs on host plant roots was not investigated. The present study shows that the number of EM species is negatively correlated with the abundance of T. melanosporum and T. aestivum ECMs. Napoli et al. (2010), reported a lower diversity of fungi in the soils sampled within than outside the brulé formed by T. melanosporum-colonized plants, but whether the fungal communities differ both quantitatively and qualitatively between brulé from natural and cultivated sites was not addressed. The ITS-based screening, which was here performed to identify, in a cost-affordable manner, as many fungal variants in the soils as possible, proves that the decrease in richness of EM species on roots of plants from cultivated vs. natural T. melanosporum truffle grounds is reflected by a parallel decrease in soil fungi. At the light that nurseries take care that contaminating fungi are excluded from the plants produced, so that greenhouse-inoculated plants have generally a high percentage of truffle ECMs at the time of outplanting, it can be argued that, once inoculated, some if not all Tuber species negatively affect fungal competitors from spreading on and next to their host plants. The competitive advantage of inoculated ECM species over soil-resident fungi and of early colonists over later arrivers is a well-documented phenomenon known as ‘priority effects’ (Kennedy et al., 2009). Yet, we note that T. aestivum and T. melanosporum on Q. pubescens both formed a brulé on both cultivated and natural sites. Indeed, volatile organic compounds (VOCs) and methanolic extracts from different truffle species have recently been shown to exert allelopathic activity (Pacioni, 1991; Angelini et al., 2010; Splivallo et al., 2011). It is conceivable that these metabolites may also participate in controlling processes involved in plant–microorganism or microorganism–microorganism interactions, thus negatively affecting the growth of putative fungal competitors for host plant and/or niche colonization. Therefore, it is conceivable that the priority effect conferred by the artificial mycorrhization and allelopathy may act synergistically (Kennedy, 2010). The alternative explanation that, in cultivated truffle grounds, agronomic practices might interfere with the spreading and colonization of indigenous EM species seems less likely. In the cultivated plantation of Montemartano, plants producing T. brumale exhibited a richness of ECMs higher than that of the plants producing T. melanosporum suggesting that the anthropogenic impact is not the cause of the inferior number of EM taxa found on and next to T. melanosporum-inoculated plants. Indeed, in contrast to T. melanosporum-colonized plants, those colonized by T. brumale do not form a brulé. Thus, halting of the invasion of grasses and competitor fungi may be an intrinsic attribute of only a few truffle species whose magnitude can vary from species to species and according to the environment. On this context, Vallunga orchard is a case study in that T. aestivum was the dominant species on T. aestivum-inoculated plants, as expected, but also on plants originally inoculated with T. melanosporum. This is consistent with previous observation showing that T. melanosporum is often replaced by T. aestivum but also by T. brumale under open field conditions (Garcìa Montero et al., 2008).

Furthermore, the parallel screening of ECMs and soil fungi from both natural and cultivated, productive and unproductive, grounds here carried out provided us this intriguing hints concerning the composition of EM communities in different T. melanosporum sites: indeed, in the cultivated sites of Montemartano (MCM and MCB), the associated fungal communities were more similar to those of the natural sites (VNM and VGNM) than to those of the cultivated sites of Vallunga (VCM, VCA), despite that these sites are geographically closer to VNM and VGNM. The cultivated site of Montemartano is productive, whereas in the cultivated site of Vallunga the production of T. aestivum or T. melanosporum was never achieved. In VCM and VCA, the number of EM fungi is markedly lower than in the other sites (Tables 2 and 4). Altogether it can be argued that the association with non-Tuber EM species is not per se a factor sufficient to perturb truffle production on both naturally or artificially inoculated truffle plants.

Soil fungal biodiversity within and outside the brulé was studied in French T. melanosporum truffle grounds by denaturing gradient gel electrophoresis (DGGE) (Napoli et al., 2010) and by pyrosequencing of the ITS (Mello et al., 2011). The comparison of our results with those from Napoli et al. (2010) highlights the presence of 21 OTUs shared between French and Italian sites, many of these (11) related to fungi detected inside the brulé (Table S5). Similarly, Mello et al. (2011) reported the abundant presence of Hypocreales, Basidiomycetes such as Cryptococcus, Hymenogaster as well as members of Thelephoraceae, also detected in our samples. Differently from what found in French T. melanosporum sites, our screening highlighted the presence either as root resident or as soil free-living mycelia of OTUs relative to the T. woolhopeia complex as well as to C. geophilum. Notably, T. woolhopeia species complex forms the AD-type ECMs described by Giraud (1988) (Rubini et al., 2011a). Indeed, AD-type ECMs have been frequently found in both T. melanosporum and T. aestivum truffle orchards (Giraud, 1988; De Miguel & Sáez, 2005; Baciarelli Falini et al., 2006; Suz et al., 2010; Benucci et al., 2011). The presence of T. woolhopeia in both cultivated and natural truffle grounds under investigation lends further support to the idea that fungi belonging to this species complex are competitors of T. melanosporum and T. aestivum (Benucci et al., 2011; Rubini et al., 2011a). A similar conclusion could be drawn for C. geophilum, which is considered a cosmopolitan EM species (LoBuglio et al., 1996). Cenococcum geophilum was in fact one of the dominant species in the two natural truffle grounds and this is in line with previous studies reporting its presence on both cultivated and natural T. melanosporum grounds (Baciarelli Falini et al., 2008; González-Armada et al., 2010).

We finally note that some dominant EM species such as T. melanosporum and T. aestivum in cultivated sites or C. geophilum in natural sites were not detected across all the seasonal samplings in all the sampled hosts. Indeed, when the distribution of a specific EM taxon on the plant roots is not sufficiently uniform, it is more than conceivable that its presence could be underestimated. Along the same reasoning, the fluctuating presence of most EM species as soil free-living mycelia was an expected result, as we have previously shown that the presence of T. melanosporum free-living mycelia was dependent on the season (Rubini et al., 2011b). Overall, the life cycle and seasonal effects appear to control the spreading of EM as well as non-EM fungal species as soil free-living mycelia.

Thus far, only a few studies on the competitive interactions involving Tuber species, such as T. melanosporum vs. Tbrumale (Mamoun & Olivier, 1993) or T. aestivum vs. other EM fungi (Pruett et al., 2008), have been performed. Indeed, dedicated studies are needed to ascertain the extent of the competitive interaction between Tuber and other EM species. The isolation of mycelia from most of the ECMs identified in this study is expected to help establish in vitro dual cultures to assess whether co-colonizing fungi exert any competing effects on the growth of T. melanosporum mycelia.

Tuber melanosporum strain distribution by ITS haplotyping

The data presented here show that ITS haplotype I was the most common T. melanosporum haplotype. This result agrees with those reported in Murat et al. (2004) and Riccioni et al. (2008) who showed that haplotype I is the most widespread T. melanosporum haplotype.

More specifically, all of the T. melanosporum ECMs analyzed showed ITS haplotype I. Conversely, in addition to haplotype I, haplotype IV and haplotypes II and IV were detected in the soil samples collected next to the VNM and MCM plants, respectively. That ECMs with a single ITS haplotype are surrounded by soil living mycelia with different haplotypes from each other in productive truffle sites (VNM and MCM) is an evidence that fits nicely with heterothallism in T. melanosporum and the model that ECM strains act as maternal partner and the soil strains as paternal one in the fertilizing process (Rubini et al., 2011b). Along the same line, present ITS-based data are in keeping with the model which suggests that single host plants and even nearby plants in a given site are colonized by the same black truffle strain that likely represents a genet (Rubini et al., 2011b).

The findings that in some soil samples (VCM and VGNM), T. melanosporum was detected only by means of T. melanosporum-specific ITS primers and that different strains were detected within a single truffle ground prompt us to argue the following: (1) the presence and distribution of soil free-living T. melanosporum mycelia can be below the amplification detection limit (using universal ITS primers) in complex and species-rich environmental samples, and (2) the development of species-specific and highly polymorphic markers, such as SSR loci, provides a new tool to analyze and compare the T. melanosporum biodiversity at both the ECM and soil levels in natural and cultivated as well as productive and nonproductive black truffle grounds. Thus, SSR- and mating-type-based genotyping of the ECMs and soil samples from the truffle sites investigated in this study will be performed in the future to assess and compare further the spatial and temporal distribution of T. melanosporum strain biodiversity under different ecological niches.

Conclusions

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Conclusions
  8. Acknowledgements
  9. Authors' contribution
  10. References
  11. Supporting Information

This study, based on a molecular inventory at both the root and soil levels of fungal biodiversity, provides an insight into the distribution and seasonal variation of the fungal community in natural and cultivated T. melanosporum sites and advances our understanding of the factors that influence fungal assemblage on truffle sites. For the first time, we provide evidence that fungal communities underneath cultivated and natural T. melanosporum plants can diverge and, under identical environmental conditions (i.e. host plant species and microhabitats), the composition of soil and EM mycobionts depends highly on the truffle species colonizing the root system.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Conclusions
  8. Acknowledgements
  9. Authors' contribution
  10. References
  11. Supporting Information

This study was partially funded by Regione Lazio (PRAL 2003–2005) and the Italian Ministry for the Environment. The authors thank Prof. A. Rambelli, Dr S. Arcioni, Mr M. Guaragno and Dr V. Passeri. The authors are also grateful to Mr D. Manna and Mr R. Paiella, the owners of the cultivated truffle grounds and Mr D. Bigioni, Mr S. Palla and ‘Consorzio Stedi’ for the their help in the identification of the natural truffle grounds.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Conclusions
  8. Acknowledgements
  9. Authors' contribution
  10. References
  11. Supporting Information
  • Agerer R (1986) Colour Atlas of Ectomycorrhizae. Einhorn, Schwäbisch-Gmünd.
  • Agerer R (1991) Characterization of ectomycorrhiza. Techniques for the Study of Mycorrhiza (Norris JR, Read DJ & Varma AK, eds), pp. 2573. Academic Press, London, UK.
  • Angelini P, Donnini D, Pagiotti R, Tirillini B, Granetti B & Venanzoni R (2010) Biological activities of methanolic extract from Tuber aestivum, T. borchii and T. brumale f. moschatum. Osterr Z Pilzkd 19: 281290.
  • Arnold AE, Henk DA, Eells R, Lutzoni F & Vilgalys R (2007) Diversity and phylogenic affinities of foliar fungal endophytes in loblolly pine inferred by culturing and environmental PCR. Mycologia 99: 185206.
  • Baciarelli Falini L, Rubini A, Riccioni C & Paolocci F (2006) Morphological and molecular analyses of ectomycorrhizal diversity in a man-made T. melanosporum plantation: description of novel truffle-like morphotypes. Mycorrhiza 16: 475484.
  • Baciarelli Falini L, Bencivenga M, Donnini D & Di Massimo G (2008) Risultati delle ricerche nelle tartufaie della regione Umbria. Atti 3° Congresso Internazionale di Spoleto sul tartufo (Comunità Montana dei Monti Martani, Serano e Subasio, eds), 25–28 November 2008, Spoleto, Italia, pp. 577589.
  • Benucci GMN, Raggi L, Alberini E, Grebenc T, Bencivenga M, Falcinelli M & Di Massimo G (2011) Ectomycorrhizal communities in a productive Tuber aestivum Vittad. Orchard: composition, host influence and species replacement. FEMS Microbiol Ecol 76: 170184.
  • Bertini L, Rossi I, Zambonelli A, Amicucci A, Sacchi A, Cecchini M, Gregori G & Stocchi V (2006) Molecular identification of Tuber magnatum ectomycorrhizae in the field. Microbiol Res 161: 5964.
  • Chevalier G, Giraud M & Bardet MC (1982) Interaction entre le mycorrhizes de Tuber melanosporum et celles d'autres champignons ectomycorrhiziennes en sols favorable al la truffe. Le Colloques de l'INRA 13: 313321.
  • Colwell RK (2005) EstimateS: Statistical Estimation of Species Richness and Shared Species from Samples, Version 8.2. User's Guide and application published online at: http://viceroy.eeb.uconn.edu/estimates.
  • De Miguel AM & Sáez R (2005) Algunas micorrizas competidoras de plantaciones truferas. Publ Biol Univ Navarra Ser Bot 16: 118.
  • Donnini D & Bencivenga M (1995) Micorrize inquinanti frequenti nelle piante tartufigene. Inquinanti in campo. Micol Ital 20: 185207.
  • Egger KN (1995) Molecular analysis of ectomycorrhizal fungal communities. Can J Bot 73: S1415S1422.
  • Etayo ML & De Miguel AM (1999) Effect of mulching on Tuber melanosporum Vitt. mycorrhizae vs. other competing mycorrhizae in a cultivated truffle bed. Abstract of the 5th international congress on the science and cultivation of truffle and other edible hypogeous mushrooms, 4–6 March 1999, Aix-en-Provence, France, pp. 378381.
  • Fasolo-Bonfante P & Fontana A (1971) Studi sull'ecologia del Tuber melanosporum i) dimostrazioni di un effetto fitotossico. Allionia 17: 4754.
  • Ferrara AM, Palenzona M & Lo Bue G (1999) Osservazioni preliminari sull'incidenza di alcune pratiche colturali in una tartufaia impiantata a Tuber aestivum Vitt. Proceedings of the 5th international congress on the science and cultivation of truffle and other edible hypogeous mushrooms, 4–6 March 1999, Aix-en-Provence, France, pp. 382386.
  • Garcìa Montero LG, Díaz P, Martín-Fernández S & Casermeiro MA (2008) Soil factors that favour the production of Tuber melanosporum carpophores over other truffle species: a multivariate statistical approach. Acta Agr Scand B-SP 58: 322329.
  • Garcia-Barreda S & Reyna S (2011) Below-ground ectomycorrhizal community in natural Tuber melanosporum truffle ground and dynamics after canopy opening. Mycorrhiza. DOI: 10.1007/s00572-011-0410-2.
  • Gardes M & Bruns TD (1993) ITS primers with enhanced specificity for basidiomycetes – application to the identification of mycorrhizae and rusts. Mol Ecol 2: 113118.
  • Garland F (1999) Growing Tuber melanosporum under adverse acid soil conditions in the United States of America. Proceedings of the 5th international congress on the science and cultivation of truffle and other edible hypogeous mushrooms, 4–6 March 1999, Aix-en-Provence, France, pp. 393.
  • Giraud M (1988) Prélévement et analyse de mycorhizes. La Truffe. CTIFL, Paris. Bull FNTP 10: 4963.
  • González-Armada B, De Miguel AM & Cavero RY (2010) Ectomycorrhizae and vascular plants growing in brulés as indicators of below and above ground microecology of black truffle production areas in Navarra (Northern Spain). Biodivers Conserv 19: 38613891.
  • Granetti B & Angelini P (1992) Competizione tra alcuni funghi micorrizici e T. melanosporum Vitt. in una tartufaia coltivata. Micol Vegetazione Mediterr 7: 173188.
  • Granetti B & Baciarelli Falini L (1997) Competizione tra le micorrize di T. melanosporum Vitt. e quelle di altri funghi in una tartufaia coltivata a Quercus ilex L. Micol Ital 1: 4549.
  • Hall TA (1999) BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp 41: 9598.
  • Hall IR, Yun W & Amicucci A (2003) Cultivation of edible ectomycorrhizal mushrooms. Trends Biotechnol 21: 433438.
  • Horton TR & Bruns TD (2001) The molecular revolution in ectomycorrhizal ecology: peeking into the black-box. Mol Ecol 10: 18551871.
  • Iotti M, Lancellotti E, Hall I & Zambonelli A (2010) The ectomycorrhizal community in natural Tuber borchii grounds. FEMS Microbiol Ecol 72: 250260.
  • Kennedy P (2010) Ectomycorrhizal fungi and interspecific competition: species interactions, community structure, coexistence mechanisms, and future research. New Phytol 187: 895910.
  • Kennedy P & Bruns TD (2005) Priority effects determine the outcome of ectomycorrhizal competition between two Rizopogon species colonizing Pinus muricata seedlings. New Phytol 166: 631638.
  • Kennedy PG, Peay KG & Bruns TD (2009) Root tip competition among ectomycorrhizal fungi: are priority effects a rule or an exception? Ecology 90: 20982017.
  • LoBuglio KF, Berbee ML & Taylor JW (1996) Phylogenetic origins of the asexual mycorrhizal symbiont Cenococcum geophilum Fr. and other mycorrhizal fungi among the ascomycetes. Mol Phylogenet Evol 6: 287294.
  • Mamoun M & Olivier JM (1993) Competition between Tuber melanosporum and other ectomycorrhizal fungi under two irrigation regimes. I. Competition with Tuber brumale. Plant Soil 149: 211218.
  • Martin F, Kohler A, Murat C et al.2010) Périgord black truffle genome uncovers evolutionary origins and mechanisms of symbiosis. Nature 464: 10331038.
  • Mello A, Napoli C, Murat C, Morin E, Marceddu G & Bonfante P (2011) ITS-1 versus ITS-2 pyrosequencing: a comparison of fungal populations in truffle grounds. Mycologia 103: 11841193.
  • Murat C, Díez J, Luis P, Delaruelle C, Dupré C, Chevalier G, Bonfante P & Martin F (2004) Polymorphism at the ribosomal DNA ITS and its relation to postglacial re-colonization routes of the Perigord truffle Tuber melanosporum. New Phytol 164: 401411.
  • Murat C, Vizzini A, Bonfante P & Mello A (2005) Morphological and molecular typing of the below-ground fungal community in a natural Tuber magnatum truffle-ground. FEMS Microbiol Lett 245: 307313.
  • Napoli C, Mello A, Borra A, Vizzini A, Sourzat P & Bonfante P (2010) Tuber melanosporum, when dominant, affect fungal dynamics in truffle grounds. New Phytol 185: 237247.
  • Oksanen J, Blanchet FG, Kindt R, Legendre P, O'Hara RB, Simpson GL, Solymos P, Stevens MHH & Wagner H (2010) Vegan: Community Ecology Package. R package. version 1.17-4. Available at http://CRAN.R-project.org/package=vegan.
  • Pacioni G (1991) Effects of Tuber metabolites on the rhizospheric environment. Mycol Res 95: 13551358.
  • Paolocci F, Rubini A, Granetti B & Arcioni S (1999) Rapid molecular approach for a reliable identification of Tuber spp. ectomycorrhizae. FEMS Microbiol Ecol 28: 2330.
  • Paulus BC, Kanowski J, Gadek PA & Hyde KD (2006) Diversity and distribution of saprobic microfungi in leaf litter of an Australian tropical rainforest. Mycol Res 110: 14411454.
  • Pruett G, Bruhn J & Mihail J (2008) Temporal dynamics of ectomycorrhizal community composition on root systems of oak seedlings infected with Burgundy truffle. Mycol Res 112: 13441345.
  • R Development Core Team (2009) R: A Language and Environment for Statistical Computing. Foundation for Statistical Computing, Vienna, Austria. Available at http://www.R-project.org.
  • Riccioni C, Belfiori B, Rubini A, Passeri V, Arcioni S & Paolocci F (2008) Tuber melanosporum outcrosses: analysis of the genetic diversity within and among its natural populations under this new scenario. New Phytol 180: 466478.
  • Richard F, Millot S, Gardes M & Selosse MA (2005) Diversity and specificity of ectomycorrhizal fungi retrieved from an old-growth Mediterranean forest dominated by Quercus ilex. New Phytol 166: 10111023.
  • Rubini A, Belfiori B, Passeri V, Baciarelli Falini L, Arcioni S, Riccioni C & Paolocci F (2011a) The AD-type ectomycorrhizas, one of the most common morphotypes present in truffle fields, result from fungi belonging to the Trichophaea woolhopeia species complex. Mycorrhiza 21: 1725.
  • Rubini A, Belfiori B, Riccioni C, Arcioni S, Martin F & Paolocci F (2011b) Tuber melanosporum: mating type distribution in a natural plantation and dynamics of strains of different mating types on the roots of nursery-inoculated host plants. New Phytol 189: 723735.
  • Rubini A, Belfiori B, Riccioni C, Tisserant E, Arcioni S, Martin F & Paolocci F (2011c) Isolation and characterization of MAT genes in the symbiotic ascomycete Tuber melanosporum. New Phytol 189: 710722.
  • Sambrook J, Fritsch EF & Maniatis T (1989) Molecular Cloning. A Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
  • Shannon CE (1948) A mathematical theory of communication. Bell Syst Tech J 27: 379423.
  • Splivallo R, Ottonello S, Mello A & Karlowsky P (2011) Truffle volatiles: from chemical ecology to aroma biosynthesis. New Phytol 189: 688699.
  • Suz LM, Martín M, Fisher CR, Bonet J & Colinas C (2010) Can NPK fertilizers enhance seedling growth and mycorrhizal status of T. melanosporum-inoculated Quercus ilex seedlings? Mycorrhiza 20: 349360.
  • White TJ, Bruns T, Lee S & Taylor J (1990) Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. PCR Protocols. A Guide to Methods and Applications (Gelfand MA, Sininski DM & White TJ, eds), pp. 315322. Academic Press, San Diego, CA.

Supporting Information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Conclusions
  8. Acknowledgements
  9. Authors' contribution
  10. References
  11. Supporting Information
FilenameFormatSizeDescription
fem1379-sup-0001-TableS1-S5-FigureS1.docWord document425KFig. S1. Average linkage clustering of truffle grounds based on Jaccard (a) and Bray-Curtis (b) dissimilarities. Table S1. Morphological characteristics of ECMs. Table S2. Relative frequency (%) of ECM morphotypes per plant and per site. Table S3. Relative Abundance (%) of ECM morphotypes per plant and per site (samples collected in October 2007). Table S4. Similarity values among the six sites. Table S5. OTUs shared between the sites investigated in this study and those investigated by Napoli et al. (2010).

Please note: Wiley Blackwell is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.