• Tuber aestivum;
  • Tuber orchard;
  • host preference;
  • hazel;
  • hornbeam;
  • ITS sequencing


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

Truffles (Tuber spp.) and other ectomycorrhizal species form species-rich assemblages in the wild as well as in cultivated ecosystems. We aimed to investigate the ectomycorrhizal communities of hazels and hornbeams that are growing in a 24-year-old Tuber aestivum orchard. We demonstrated that the ectomycorrhizal communities included numerous species and were phylogenetically diverse. Twenty-nine ectomycorrhizal taxa were identified. Tuber aestivum ectomycorrhizae were abundant (9.3%), only those of Tricholoma scalpturatum were more so (21.4%), and were detected in both plant symbionts with a variation in distribution and abundance between the two different hosts. The Thelephoraceae family was the most diverse, being represented by 12 taxa. The overall observed diversity represented 85% of the potential one as determined by a jackknife estimation of richness and was significantly higher in hazel than in hornbeam. The ectomycorrhizal communities of hornbeam trees were closely related phylogenetically, whereas no clear distribution pattern was observed for the communities in hazel. Uniform site characteristics indicated that ectomycorrhizal relationships were host mediated, but not host specific. Despite the fact that different plant species hosted diverse ectomycorrhizal communities and that the abundance of T. aestivum differed among sites, no difference was detected in the production of fruiting bodies.


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

An ectomycorrhiza constitutes a mutualistic relationship between the mycelium of a fungus and the roots of a host plant (Smith & Read, 2008). All fungal species in the genus Tuber (Ascomycota, Tuberaceae) presumably develop ectomycorrhizae with woody angiosperms and gymnosperms, without a preference for a particular host taxonomic group (Agerer, 1987–2008; Norris et al., 1994; Hall et al., 2007). Tuber spp. produce subterranean edible ascocarps, which are highly sought after for their superb flavour and distinctive fragrance and are known as truffles.

Over the last few decades, the production of some truffle species, such as Tuber melanosporum, in Europe has declined rapidly, mainly as a result of overexploitation and the disruption of their natural habitat (Hall et al., 2001, 2003). To overcome this decline, seedlings that have been inoculated with the spores of truffles have been produced and cultivated in specialized orchards; spores from the most sought after species of truffle have been used in particular. To cultivate truffles, the mycorrhized plants are transplanted on a suitable field according to the ecological conditions that match the requirements of the specific host–fungus combination. This culture technique is now feasible for T. melanosporum, which is known commonly as black truffle (Bencivenga & Granetti, 1990), Tuber borchii (Zambonelli et al., 2002), and Tuber aestivum (Chevalier & Frochot, 1997), which is considered to be a synonym of Tuber uncinatum (Wedén et al., 2005), also known as summer truffle, autumn truffle, or Burgundy truffle. However, it is not yet possible to use this technique for Tuber magnatum (white truffle). This is mainly because it is difficult to obtain plants infected with T. magnatum that are not also contaminated with other ectomycorrhizal (ECM) species, such as Tuber maculatum, T. borchii, Sphaerosporella brunnea, and Pulvinula constellatio (Bertini et al., 2005). Tuber maculatum, T. borchii, and other Tuber spp. also seem to compete actively with T. magnatum in cultivated orchards in which this host–fungus combination is established (Donnini et al., 2000). In Italy, cultivated truffle orchards expanded rapidly in the 1980s when many nurseries became specialized in the production of mycorrhized plants (Granetti et al., 2005).

The cultivation of T. aestivum is drawing increasing interest from farmers for several reasons. (1) It is an economically important truffle, has a thriving market, and is sold throughout the world (Hall et al., 2007). (2) It is spread throughout the whole of Europe (between 37 and 57°N) and North Africa (Jeandroz et al., 2008). (3) It adapts easily to a broader range of climatic conditions and the physical and chemical characteristics of soil than T. melanosporum (Hall et al., 2007). (4) Cultivation of inoculated T. aestivum seedlings is feasible (Chevalier & Frochot, 1997). (5) Different from T. melanosporum and T. magnatum, T. aestivum shows two distinct growth and fruiting strategies, which occur both in closed (mature) forest stands and in completely open habitats, as reported by Bencivenga et al. (1995).

Several years after planting, the yield of all cultivated Tuber spp. is excellent in some orchards, but very poor or nonexistent in others. The causes of this phenomenon remain poorly understood. Some researchers have hypothesized that failure of yield could be due mainly to a poor choice of cultivation site, namely, unsuitable climatic and soil conditions (Garćιa Montero et al., 2008), and to the presence of few ectomycorrhizae on nursery-infected plants at the time of planting (Hall et al., 2003). Another hypothesis could be that strains with different mating types are not evenly distributed beneath productive soil patches as reported for T. melanosporum (Rubini et al., 2011a). Some other authors have focused their attention on the presence of competing indigenous ectomycorrhizal fungi. Despite the fact that recently molecular methods of identification have been used widely to support classical morphological techniques in the characterization of assemblages of ectomycorrhizal species (Gardes & Bruns, 1993; Horton & Bruns, 2001), knowledge of the diversity of truffle environment ectomycorrhizal fungi is generally poor. However, recent studies have shown that, in natural T. magnatum grounds, the most common ectomycorrhizal family is Thelephoraceae, which is represented in all cases by Tomentella (Murat et al., 2005). In contrast, T. magnatum ectomycorrhizae are very rare, which has also been reported by Bertini et al. (2005). Baciarelli Falini et al. (2006) described the diversity of ectomycorrhizal fungi in an unproductive 14-year-old T. melanosporum plantation. They found that T. melanosporum ectomycorrhizae were abundant and shared the habitat with other ectomycorrhizal fungi, such as T. borchii, T. aestivum, Tuber brumale, Scleroderma spp., and Cortinarius spp. In natural T. borchii grounds, Iotti et al. (2010) found that truffle ectomycorrhizae dominated the ectomycorrhizal population, although it also contained Thelephoraceae, Inocybaceae, Sebacinaceae, and Pyronemataceae. Pruett et al. (2008) studied a T. aestivum orchard that was established in Missouri with inoculated Quercus bicolor×Quercus robur seedlings. After lime application, trees were planted in a non-native field because T. aestivum is not present naturally in North-America. Two years after plantation, T. aestivum was still present together with T. maculatum, Tuber rufum, and Tomentella, Scleroderma, and Hebeloma spp.

Regardless of whether competition between fungi is one of the most important direct determinants of local ectomycorrhizal diversity (Bruns, 1995), it is essential to be able to detect species that are able to outcompete introduced truffles and to characterize their interactions in order to understand the conditions that promote truffle fructification and guide the making of decisions with respect to land management (Baciarelli Falini et al., 2006; Pruett et al., 2008). Understanding species diversity is also an important prerequisite for successful conservation efforts and orchard management when an introduced ectomycorrhizal community is being replaced slowly by a wild one. However, evidence that plants allocate resources to one particular ectomycorrhizal fungal partner in preference to another is very limited. Ishida et al. (2007), Tedersoo et al. (2008), and Morris et al. (2008) have demonstrated the existence of a strong, but not exclusive preference for a particular host. Comparison of the ectomycorrhizal fungi that are found among different host species within environments with the same conditions will help elucidate the importance of host preference in structuring the ectomycorrhizal community (Horton & Bruns, 1998; Dickie, 2007).

In the study reported herein, we analysed the diversity of ectomycorrhizal communities associated with hazel (Corylus avellana L.) and hornbeam (Ostrya carpinifolia Scop.) in a 24-year-old orchard that produces T. aestivum truffles. We aimed to achieve the following: (1) to verify and quantify the presence and distribution of T. aestivum and other Tuber spp. ectomycorrhizae; (2) to characterize other (non-Tuber) ectomycorrhizae present in the orchard; (3) to analyse the species composition and diversity of fungi associated with different plants using statistical techniques appropriate for studies of community ecology; and (4) to identify the importance of the effects of host species on the long-term structure of a community of ectomycorrhizal fungi.

Materials and methods

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

Description of the truffle orchard and collection of ectomycorrhiza samples

At the end of May 2009, samples of roots were collected in a truffle orchard that had been planted in 1985 and is located a few kilometres away from Spoleto, Central Italy (latitude 42°42′24.76″N; longitude 12°43′7.79″E). The site has an area of 5 ha, an average altitude of 350 m a.s.l., an E-SE aspect, and a slight slope. The following tree symbionts are found in this orchard: (1) Downy oak (Quercus pubescens Willd.), (2) European hazel (C. avellana L.), (3) European hornbeam (O. carpinifolia Scop.), and (4) Holm oak (Quercus ilex L.). The truffle plantation is a mature closed stand, with some hornbeams that are over 7 m in height. All species planted in the plantation are arranged with 5 m between the rows and 4 m between individuals within the rows.

Since 1998, the areas of the plantation that contain exclusively hornbeam have been monitored for the production of T. aestivum. Production started under a few hornbeam trees and then spread, year by year, to all hornbeam trees. To date, nearby hazel trees have also been found to produce fruiting bodies of T. aestivum. Given that truffles were produced in two different host species, two distinct types of plot, each with two replicates (to yield a total of four plots), were selected randomly within the orchard: plot A and plot B were located in the area with hazel and plot C and plot D were located in the area that contained hornbeam. Each plot covered approximately 600 m2 and contained 35 plants. It is necessary to select small plots that are occupied by a single host species to ensure homogeneous ecological conditions and to remove any possible effects of soil and climatic differences or other biotic or abiotic factors on ectomycorrhizal diversity (Bruns, 1995). Twenty-four years after the original inoculated seedlings had been planted, 10 trees per plot were selected randomly among the productive trees. A productive tree was defined as a tree that had produced at least one ascocarp per season in the previous 3 years. Trees that were located on the boundaries of the orchard were not sampled to reduce the edge effect. Three soil cores (6 cm diameter, 20 cm deep) were obtained 1 m from the base of each tree in three random directions. These three cores (per tree) were then combined to yield a single composite sample (Gehring et al., 1998; Avis et al., 2003). Ten combined samples were collected per plot. The collected samples were placed immediately in a cooler and kept at 4 °C until processed.

Morphological analysis

The samples were washed carefully with tap water over a 1-mm sieve and the roots that were present in the samples were floated in sterile glass Petri dishes that contained distilled water (Avis et al., 2003). The different types of ectomycorrhizae that were found in each sample were sorted according to morphotype and labelled using a serial number preceded by the initials UE (uncultured ectomycorrhiza). All vital ectomycorrhizal root tips that showed the same anatomical and morphological characteristics, as observed first under a stereomicroscope (Leica Leitz Wild MZ8) and then a light microscope (Leica Leitz DMRB), were considered to be distinct morphotypes. All morphotypes were described briefly (Agerer, 1987–2008; Zambonelli et al., 1993). From each root sample, up to three subsamples of root tips of the same morphotype were placed in 1.5-mL microfuge tubes with a drop of deionized water and stored at −80 °C for subsequent molecular analysis.

To determine the relative abundance of ectomycorrhizal fungi (percentage of all ectomycorrhizae), we counted each ectomycorrhiza as an individual (Bruns, 1995; Dickie et al., 2002). Nonmycorrhizal and nonvital tips were also recorded, stored separately, and included in the category of dried root tips. For each sample, roots were cut into 2-cm pieces, 10 of which were selected randomly, and a total of 400 tips were counted.

Nuclear ribosomal DNA (nrDNA)-internal transcribed spacer (ITS)-based identification of ectomycorrhizae

To remove all residual soil particles and other fungal structures (mycelia or spores), the tips of each morphotype were washed briefly by vortexing in a microfuge tube and spun for 2 min at 15 000 g. The ectomycorrhizae were analysed and manipulated under a stereomicroscope (× 20−40 magnification). For each morphotype, a single ectomycorrhiza was peeled from the surrounding hyphae using a fine needle and then a very small portion of the mantle was picked up and transferred directly into a PCR tube that contained 50 μL of PCR Mix, as reported by Iotti & Zambonelli (2006), with some modifications as reported below.

Primers ITS1 and ITS4 were used to amplify the ITS1, 5.8S, and ITS2 regions of the nrDNA (White et al., 1990). PCR reactions were performed in a 50-μL volume that contained the following: 1 × PCR buffer (Invitrogen), 200 μM dNTPs, 20 μg of bovine serum albumin, 10 pmol of the ITS1 and ITS4 primers, and 1.75 U of Taq DNA polymerase (Invitrogen). All reactions were performed using the following cycling parameters: 94 °C for 5 min, followed by 35 cycles of 94 °C for 30 s, 57 °C for 30 s, and 72 °C for 1 min. Aliquots of the PCR products (10 μL) were electrophoresed on a 1.5% (w/v) agarose gel and visualized by ethidium bromide staining. Amplified products were purified using an Illustra GFX PCR DNA kit (GE Healthcare) and sequenced. Sequencing reactions were performed using a Big Dye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems) and run on an ABI 3130xl DNA Analyzer.

Sequence identity searches were conducted using the blastn algorithm at the National Center for Biotechnology Information (NCBI) website ( using default settings (Altschul et al., 1997). After confirmation with blastn, all ITS sequences obtained that belonged to the genus Tomentella or Pseudotomentella were aligned with 37 additional congeneric ITS sequences that had been submitted previously to GenBank by Kõljalg et al. (2000) and Haug et al. (2004). The accession numbers of the reference sequences and provenance are indicated in Fig. 1. All sequence alignments, as well as phylogenetic and molecular evolutionary analyses, were carried out using mega version 4 software (Tamura et al., 2007). A neighbour-joining (NJ) tree was produced using the Jukes–Cantor model and support for clades was assessed using a bootstrap test (1000 replications), with values ≥55 indicated above the branches of the tree.


Figure 1.  Bar chart of the relative abundances of Tuber spp. for each sample collected in the truffle orchard.

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Statistical analysis

The diversity of the communities of ectomycorrhizal fungi was analysed for each plot and host using the following biodiversity indices: richness (S), evenness (E=exp H′/S), Shannon's H′ (H′=−Σni ln ni), and Simpson's D (D=1−Σ(ni/N)2). Once the presence of a normal distribution had been verified using the Shapiro–Wilk test, the values of the indices were used for multiple comparisons. Significant differences between the means of the indices (P<0.05) were detected by anova; Tukey's test was used to identify significant differences between plots and Student's t-test to identify significant differences between plant partners of ectomycorrhiza (hazel and hornbeam).

Differences among the relative abundances of ectomycorrhizal taxa were studied using the nonparametric Kruskal–Wallis test. To compare species richness, patterns of diversity, and to check the appropriateness of the sampling scheme, we constructed accumulation and rank-abundance curves. The species accumulation curve was obtained using 100 permutations with no randomization (Ugland et al., 2003; Colwell et al., 2004). Abundance and log10 of abundance values were used to generate the rank-abundance curves. First- and second-order jackknife estimators of richness were considered to test the potential richness of the surveyed areas.

Pairwise Pearson correlation tests (P=0.05) of all combinations of different plots and different host species were carried out using the data for total relative abundances (means: n=10 for A, B, C, and D; n=20 for hazel and hornbeam). A heatmap of pairwise Pearson correlation r-values (P=0.05) for all possible combinations of different root samples was plotted using the data for the total relative abundances in single counts. The dendrograms shown at the top and on the left side of the heatmap were generated using hierarchical cluster analysis (Warnes, 2009). A nonmetric multidimensional scaling (NMS) ordination technique (Bray–Curtis distance, 2 axes, 100 permutations) was used to compare communities of ectomycorrhizal fungi between different hosts and plots. For the nonparametric tests and NMS ordination, data were standardized with square-root transformation. Factor averages (centroids) of environmental variables were also plotted into the ordination. R2 was used as a goodness-of-fit statistic and the significance was tested with 1000 permutations. The names of the factor centroids were formed by combining the name of the factor and the name of the level. To characterize the influence of the host on the composition of ectomycorrhizal species and to assess whether the similarity within each group of samples (namely, those from each of the two hosts) was greater than that between the two groups of samples, an analysis of similarity (anosim) test was performed between the ectomycorrhizal communities of hazel and hornbeam (defined a priori) with 999 permutations for significance testing (Clarke, 1993).

All the analyses mentioned above were performed using r version 2.9 software (R Development Core Team, 2009). For the indices of diversity, species accumulation and rank-abundance curves, NMS ordination technique, and anosim test, the ‘vegan’ (Oksanen et al., 2009) and ‘biodiversityr’ (Kindt & Coe, 2005) packages were also used.


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

Identification and composition of ectomycorrhizal taxa

The morphological analysis of 9896 ectomycorrhizae allowed us to distinguish 29 different morphotypes, which were then subjected to ITS analyses. Fifty-three PCR products (at least one amplicon for each morphotype) were randomly chosen and sequenced. The ITS sequences generated allowed us to distinguish and classify 27 morphotypes; in particular, the analysis allowed us to assign 11 morphotypes to species, 11 to genus, and five to family level. For two of the 29 morphotypes, we could not obtain reliable sequence data (Table 1); we assigned these morphotypes to the Basidiomycetes class on the basis of the presence of clamp connections on the mycelial hyphae.

Table 1.   List of ectomycorrhizal (ECM) morphotypes found in the four plots studied (A, B, C, D) and similarity as determined by blast analysis of the ITS sequences obtained
NameMorphotypeAccession numberblast identifierMaximum identityAbbreviation
  1. Basidiomycete I and II were assigned to the Basidiomycetes class by morphological analysis alone because the ITS sequence was not determined (ND). blast identifier, accession number, and maximum identity are also reported. To avoid congestion in the ordination plots, a Cornell Ecology Programs (CEP) abbreviated name was given to each taxon.

UE1Tuber aestivumHM370452Tuber aestivum (EU753266)0.99Tubeaest
UE2Tuber sp.HM370453Tuber rufum (AY940646)0.99Tuberufu
UE3Tuber brumaleHM370454Tuber brumale (EU753268)0.99Tubebrum
UE4Tuber borchiiHM370455Tuber rapeodorum (EU784430)0.99Tuberape
UE5AD-typeHM370456Pyronemataceae (PAPM-Mycorrhiza) (EU822505)0.98PyroMyco
UE6AD-typeHM370457Pyronemataceae (ECMm7) (DQ402506)0.99PyroECMm
UE7UnidentifiedHM370458Tarzetta catinus (FM206478)0.99Tarzcati
UE8Tuber sp.HM370459Peziza michelii (DQ200839)0.98Pezimich
UE9Tomentella sp.HM370460Tomentella cf. sublilacina (AJ889982)1.00Tomesubl
UE10UnidentifiedHM370461Tomentella sp.1 (EU668198)0.99Tome1
UE11BasidiomyceteHM370462Tomentella sp.2 (EF411113)0.95Tome2
UE12BasidiomyceteHM370463Tomentella sp.3 (AJ879688)0.99Tome3
UE13BasidiomyceteHM370464Tomentella sp.4 (EU668215)0.96Tome4
UE14UnidentifiedHM370465Tomentella sp.5 (FJ897224)0.99Tome5
UE15UnidentifiedHM370466Tomentella sp.6 (AF430289)0.94Tome6
UE16UnidentifiedHM370467Pseudotomentella sp.1 (EU668196)0.99Pseu1
UE17Tomentella sp.HM370468Pseudotomentella sp.2 (EU668254)0.92Pseu2
UE18BasidiomyceteHM370469Thelephoraceae I (AJ893307)0.96ThelI
UE19BasidiomyceteHM370470Thelephoraceae II (FJ210762)0.90ThelII
UE20Tomentella sp.HM370471Thelephoraceae III (FJ210762)0.99ThelIII
UE21UnidentifiedHM370472Sebacina sp. (EU668222)0.99Sebasp
UE22UnidentifiedHM370473Sebacinaceae (AJ879667)0.99Sebacinc
UE23UnidentifiedHM370474Inocybe luteifolia (EU523569)0.95Inoclute
UE24UnidentifiedHM370475Inocybe cf. fuscidula (AM882842)0.98Inocfusc
UE25UnidentifiedHM370476Inocybaceae (EU523580)0.86Inocybac
UE26Tricholoma sp.HM370477Tricholoma scalpuratum (EU160596)1.00Tricscal
UE27UnidentifiedHM370478Hebeloma sp. (EF093151)0.98Hebesp
UE28Basidiomycete INDNDNDBasiI
UE29Basidiomycete IINDNDNDBasiII

With regard to taxonomy, eight out of the 29 morphotypes were Ascomycetes, whereas the rest belonged to the Basidiomycetes. Thelephoraceae was the most common family with 12 species (UE9-UE20), followed by Tuberaceae with four species (UE1–UE4), Pyronemataceae and Inocybaceae with three species each (UE5–UE7 and UE23–UE25, respectively), Sebacinaceae with two species (UE21, UE22), and Tricholomataceae, Pezizaceae, and Strophariaceae with a single species each (UE26, UE8, UE27) (Table 1). Regarding the relative abundance of ectomycorrhizal fungi, the Thelephoraceae family contributed the largest number of individuals, with 24.3% total relative abundance. This was followed by Tuberaceae with 21.7% total relative abundance; the Tricholomataceae family, which was surprisingly only represented by Tricholoma scalpturatum, with 21.4%; Pyronemataceae with 9.9%; Sebacinaceae with 8.4%; Strophariaceae with 3.9%; Inocybaceae with 2.7%; and Pezizaceae with 2.1% (Table 2).

Table 2.   Relative abundances of ectomycorrhizal (ECM) taxa and biodiversity indices (richness, evenness, and Shannon's and Simpson's indices) related to plot A, plot B, plot C, plot D, hazel and hornbeam host species, and total samples
ECM taxaPlot APlot BHazelPlot CPlot DHornbeamTotal samples
  • Values of the biodiversity indices (means ± SE, n=10) with different letters are significantly different at P<0.05 and df=3.36.

  • *

    Values (means ± SE, n=20) are significantly different at P<0.05 and df=1.38. The abundance of nonmycorrhizal and nonvital root tips was also calculated for the total tips counted and included in the category of dried root tips.

Tuber aestivum0.0860.0940.0910.0720.1250.0960.093
Tuber rufum0.0990.0480.0700.1400.0210.0850.076
Tuber brumale0.0100.0310.0220.0000.0130.0060.015
Tuber rapeodorum0.0310.0000.0130.0880.0190.0560.032
Pyronemataceae (PAPM-Mycorrhiza)0.1150.1230.1200.0080.0400.0230.077
Pyronemataceae (ECM7)0.0000.0000.0000.0250.0000.0130.006
Tarzetta catinus0.0290.0310.0300.0000.0000.0000.017
Peziza michelii0.0060.0610.0380.0000.0000.0000.021
Tomentella sublilacina0.0060.0000.0030.1420.0410.0950.044
Tomentella sp. 10.0690.0690.0690.0220.0000.0120.044
Tomentella sp. 20.0070.0170.0130.0000.0000.0000.007
Tomentella sp. 30.0000.0430.0250.0000.0000.0000.014
Tomentella sp. 40.0000.0350.0200.0000.0030.0010.012
Tomentella sp. 50.0570.0080.0290.0000.0000.0000.016
Tomentella sp. 60.1000.0300.0600.0000.0000.0000.033
Pseudotomentella sp. 10.0000.0660.0380.0000.0120.0060.024
Pseudotomentella sp. 20.0000.0090.0050.0000.0000.0000.003
Thelephoraceae I0.0400.0090.0220.0000.0000.0000.012
Thelephoraceae II0.0810.0000.0350.0000.0000.0000.019
Thelephoraceae III0.0000.0470.0270.0000.0000.0000.015
Sebacina sp.0.0430.0540.0490.0000.0000.0000.027
Inocybe luteifolia0.0000.0000.0000.0240.0000.0130.006
Inocybe cf. fuscidula0.0000.0000.0000.0290.0000.0160.007
Tricholoma scalpturatum0.0710.0870.0800.2600.5250.3830.214
Hebeloma sp.0.0290.0500.0410.0670.0000.0360.039
Basidiomycete I0.0360.0150.0240.0000.0200.0090.017
Basidiomycete II0.0660.0710.0690.0000.0000.0000.039
Dried root tips0.3810.2100.2930.4110.4920.4520.373
Richness4.400 ± 0.371 a4.800 ± 0.629 a4.600 ± 0.358*2.700 ± 0.153 b2.900 ± 0.180 b2.800 ± 0.117*3.700 ± 0.235
Evenness0.784 ± 0.038 a0.813 ± 0.034 a0.766 ± 0.0280.807 ± 0.051 a0.765 ± 0.046 a0.786 ± 0.0340.792 ± 0.021
Shannon's index1.186 ± 0.119 a1.274 ± 0.119 a1.184 ± 0.083*0.741 ± 0.083 b0.761 ± 0.087 b0.751 ± 0.058*0.991 ± 0.063
Simpson's index0.608 ± 0.056 ab0.672 ± 0.037 a0.618 ± 0.035*0.444 ± 0.053 b0.443 ± 0.053 b0.443 ± 0.037*0.542 ± 0.029

The distribution of Tuber spp. ectomycorrhizae among the 40 root samples collected for this study was of particular importance (Fig. 1). In general, T. aestivum was the most abundant species among the Tuberaceae family and the second most abundant species overall (9.3%; 923 out of 9896 ectomycorrhizae). Its ectomycorrhizae were found on 16 out of the total 40 samples (40%) and were always recovered together with other ectomycorrhizal taxa. Its relative abundance in individual samples varied considerably (from 0.03 to 0.90). No T. aestivum ectomycorrhizae were detected on the remaining 24 samples (60%). With respect to the ectomycorrhizal community in hazel, T. aestivum was the second most abundant species (9.1%) and its ectomycorrhizae were found in 30% of the samples (the relative abundance in individual samples ranged from 0.05 to 0.90). Within the community in hornbeam, T. aestivum was the third most abundant species (9.6%) and its ectomycorrhizae were found in 50% of the samples (the relative abundance in individual samples ranged from 0.03 to 0.60).

With regard to other truffle species, T. rufum was found in 10 samples from hazel and five samples from the hornbeam plots, whereas T. brumale was found in five samples from hazel and in only two samples from hornbeam. Tuber rapeodorum was found in four samples, two for each host species. The relative abundances of Tuber spp. in individual samples are detailed in Fig. 1.

Phylogenetic position of Tomentella and Pseudotomentella ectomycorrhizal fungi

The sequences of the Thelephoraceae family that were determined in this study showed a broad heterogeneity upon alignment, which suggested the presence of different species and underlined the high level of diversity present in the truffle orchard. The NJ tree revealed the relationships among the nine tomentelloid ectomycorrhizal fungi that were classified in the Tomentella and Pseudotomentella genera (Fig. 2): (1) Tomentella cf. sublilacina (UE9) clustered with the three samples of T. sublilacina with deviations of 0.023 ± 0.006 for T. sublilacina (Estonia AF272933), 0.023 ± 0.006 for T. sublilacina (Sweden AF272929), and 0.024 ± 0.006 for T. sublilacina (Norway AF272935); (2) Tomentella sp.3 (UE12) clustered with Tomentella ellisii with a deviation of 0.039 ± 0.008; (3) Tomentella sp.5 (UE14) clustered with Tomentella badia with a deviation of 0.002 ± 0.002; and (4) Tomentella sp.6 (UE15) clustered with Tomentella coerulea with a deviation of 0.060 ± 0.018. The Tomentella sp.1 (UE10) cluster could represent a basal lineage of Tomentella bryophila and Tomentella lapidum, given the intraspecific distances among clusters, which are approximately the same as those among several samples of a single morphological species (e.g. T. badia, Tomentella terrestris, and Pseudotomentella tristis).


Figure 2.  Phylogenetic relationships of Tomentella and Pseudotomentella genera. NJ analysis of nrDNA ITS partial sequences with the Jukes–Cantor substitution model combined with a bootstrap analysis from 1000 replicates (bootstrap values <55% not shown). For sequences retrieved from the NCBI database, the accession number and provenance are indicated when available; the sequences generated in this study are marked with a filled triangle (▴).

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For Pseudotomentella sp.1 (UE16) and Pseudotomentella sp.2 (UE17), the closest relationship was with P. tristis (Fig. 2). Pyronemataceae samples that were classified as AD-type morphotypes exhibited two different ITS sequences: blast analysis revealed that one morphotype (UE5) was similar to Quercirhiza quadratum (PAP-Mycorrhiza), which was described recently by Agueda et al. (2008), whereas the other morphotype (UE6) was similar to an uncultured ectomycorrhizal fungal isolate, ECMm7, reported by Baciarelli Falini et al. (2006). It is worth noting that UE6 did not show any similarity to any member of the Sarcosomataceae family, although such a similarity was reported by Baciarelli Falini et al. (2006) for ECMm7. As a consequence, our findings agree with new data from the same group, which were published by Rubini et al. (2011b). In the latter report, the authors concluded that Q. quadratum and AD-type ectomycorrhizae should be assigned to different species within the Trichophaea woolhopeia complex.

Analysis of the diversity of ectomycorrhizal communities

The first- and second-order jackknife estimators of the total richness of ectomycorrhizal communities were calculated to be 33.88 and 31.22, respectively. As such, the 29 ectomycorrhizal taxa that were identified represented 85.59% or 92.88% of the total estimated richness for the orchard (Table 3).

Table 3.   Observed richness, and first- and second-order jackknife estimators calculated per plot and for the two host species considered in this study (hazel and hornbeam)
HostObserved richness (per host)1st/2nd jackknife estimators (per host)PlotObserved richness (per plot)1st/2nd jackknife estimators (per plot)

The four diversity indices (Table 2) used in this study provided an important tool to summarize the total diversity of the ectomycorrhizal fungi. Our results showed that richness, together with Shannon's and Simpson's indices of the diversity of ectomycorrhizal communities, varied significantly among different host species and different plots, whereas evenness did not (t-tests and anova, P<0.05). In particular, the plots that contained hazel showed higher values of richness and Shannon's diversity index (4.600 ± 0.3584 and 1.184 ± 0.0829, respectively) than the hornbeam plots C and D (2.800 ± 0.1170 and 0.751 ± 0.0583, respectively).

By plotting the cumulative number of species collected against the amount of sampling undertaken, we generated species accumulation curves (Fig. 3). The curves obtained indicated that, using our applied sampling scheme, the observed richness was not a marked underestimation of the actual richness. In fact, as is clear from the figure, the curves approach the asymptote, which represents the actual total number of species; if >40 soil samples had been collected, very few new species would have been detected. The different curves show that the community of ectomycorrhizal fungi in hazel is richer in terms of taxa than that in hornbeam. Rank-abundance curves (Fig. 4a and b) also showed that the hazel community was more diverse than the hornbeam community and that only a few species of ectomycorrhizal fungi dominated in each host species.


Figure 3.  Accumulation curve for ectomycorrhizal species in hazel root samples (•), hornbeam root samples (▴), and total samples (inline image). The mean cumulative richness value in each site (± SE) was plotted after 100 permutations with no randomization.

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Figure 4.  Rank-abundance curves: (a) Abundance for ectomycorrhizal species in hazel root samples (•); hornbeam root samples (▴); and total samples (inline image). (b) Log10 of abundance of ectomycorrhizal species in plot A (○), plot B (▵), plot C (+), and plot D (×).

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Kruskal–Wallis tests (α=0.05) were performed between the two host species to make comparisons at the taxonomic levels of species, family, and class. The genus level was not considered for comparisons because we were unable to distinguish species within the same ectomycorrhizal taxa. Significant differences were only found with respect to Ascomycetes (P=0.026), Pyronemataceae (P=0.001), Thelephoraceae (P=0.004), and Tricholomataceae species (P=0.005), and with respect to comparisons of single species such as Pyronemataceae (PAPM-Mycorrhiza) (P=0.001), Tarzetta catinus (P=0.019), T. cf. sublilacina (P=0.034), Thelephoraceae sp.1 (P=0.038), T. scalpturatum (P=0.005), and Basidiomycete II (P=0.002).

Pearson tests were used to measure the strength of the hypothetical linear relationships between variables (plot, host, and single-sample ectomycorrhizal communities) and provided goodness of fit for the replicate plots (A vs. B and C vs. D). The tests showed that plots A did correlate with B and C did correlate with D, which confirmed the usefulness of the replicates (α=0.05; n=29); there were no significant correlations between the ectomycorrhizal communities of the two different host plants (Table 4). The pairwise Pearson linear relationship between samples is shown by the heatmap (Fig. 5). The Pearson correlation is +1 in the case of a perfect positive (increasing) linear relationship, −1 in the case of a perfect negative (decreasing) linear relationship, and some value between −1 and +1 in all other cases; the correlation indicates the degree of linear dependence between variables. Two main clades are visible in Fig. 5, which are also depicted by the hierarchical clustering dendrogram at the top and left side. Some ectomycorrhizal root samples from plot C (C5, C10, C4, C3, C1) and almost all samples from plot D (D1, D5, D4, D8, D6, D10, D3, D2) were correlated strongly with each other (dark squares in the heatmap). Most other samples showed no correlations, with a few random exceptions.

Table 4.   Correlation matrices of pairwise Pearson correlation r-values (α=0.05) computed using relative abundances
 Plot APlot BPlot CPlot D
Plot A1   
Plot B0.5221  
Plot C0.2180.1321 
Plot D0.2350.3210.7991
  1. Significant values are shown in bold .

  2. Comparisons are shown for different plots (means; n=10 for plots A, B, C, and D) and different host species (means; n=20 for hazel and hornbeam).


Figure 5.  Heatmap of the pairwise Pearson correlation r-value matrix (α=0.05) computed using relative abundance data for single root samples. A1–A10, B1–B10, C1–C10, and D1–D10 represent the 10 samples collected in the four different plots analysed in this study (plot A, plot B, plot C, and plot D, respectively). The colour key for the correlation values is shown on the top left of the plot: as the correlation value approaches 0, the relationship is weaker (light squares); otherwise, the closer the coefficient value is to 0.8, the stronger the correlation between the samples (dark squares). A dendrogram of hierarchical cluster analysis is also shown at the top and on the left.

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The NMS ordination of the ectomycorrhizal dataset showed and confirmed the segregation of communities of ectomycorrhizal fungi between the different host species along the two axes (Fig. 6). The spread of data is due to differences in several ectomycorrhizal taxa that were found at the levels of both host and plot, as reported in Table 2: T. catinus, Peziza michelii, Tomentella sp.1, Tomentella sp.2, Tomentella sp.3, Tomentella sp.5, Tomentella sp.6, Pseudotomentella sp.2, Thelephoraceae I, Thelephoraceae II, Thelephoraceae III, Sebacina sp., and Basidiomycete II were present exclusively in the roots of hazel trees, whereas others such as Pyronemataceae (ECMm7), Inocybe luteifolia, Inocybe cf. fuscidula, and Inocybaceae were found only in the roots of hornbeam trees. Other species, such as Pyronemataceae (PAPM-Mycorrhiza) and Tomentella sp.1, were found more frequently in hazel than in hornbeam, whereas T. rapeodorum, T. cf. sublilacina, and T. scalpturatum were more abundant in the latter (Table 2).


Figure 6.  NMS ordination graphs: (a) points related to hazel (○) and hornbeam (▵) root samples where environmental variable centroids for different host species are also shown; (b) points related to species (inline image) with CEP name abbreviations (as listed in Table 2); (c) merged figure.

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From the ordination plot (Fig. 6) we can conclude that the hornbeam samples showed a gathered behaviour and tended to group close to each other whereas hazel samples were more scattered; this confirmed the lack of similarity between the two host species communities that was observed in the correlation tests. The differences in the environmental factor averages for the two hosts, which were clearly separated in the ordination plot, showed a good level of significance (R2=0.178; P≤0.001). The result of the anosim test (R=0.272; P=0.001) provided evidence of the separation of the two groups and supported the data from the NMS.


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

For the best of our knowledge, this is the first work in which communities of ectomycorrhizal fungi between areas where T. aestivum ascocarps were produced under comparable ecological conditions, and that had a homogeneous record of ascocarp production and two common host species of the same age were compared. This investigation combined the analysis of morphotypes and of ITS sequences, and the use of statistical data on community ecology. The level of information in the literature about the ecology of ectomycorrhizal communities in truffle environments is low, especially with regard to studies that combine morphological and molecular tools (Murat et al., 2005; Baciarelli Falini et al., 2006; Iotti et al., 2010) rather than only morphological ones (Donnini & Bencivenga, 1995; Donnini et al., 1999; Garćιa Montero et al., 2008). However, such studies are essential to improve knowledge about truffle ecosystems and, as a consequence, to enhance the success of truffle cultivation.

From our data, we determined that T. aestivum exhibited behaviour similar to that of some other truffle species in that it dominated the ectomycorrhizal population. Its ectomycorrhizae were abundant (9.3%) and second only to those of T. scalpturatum (21.4%). This was consistent with the report by Iotti et al. (2010) for natural T. borchii grounds, where the ectomycorrhizal community was dominated by a few common species and a large number of rare ones. Baciarelli Falini et al. (2006) reported a similar situation for an orchard in which T. melanosporum was cultivated: truffle ectomycorrhizae were abundant and found on 44.3% of the samples. In contrast, it is interesting to note that the findings for the summer truffle were in contrast to those recorded by Murat et al. (2005) for T. magnatum in a natural truffle ground located in Northern Italy, where white truffle ectomycorrhizae were rare and identified in only 5% of the 335 tips analysed. In a similar study on T. magnatum, Bertini et al. (2005) obtained similar results. The difference in symbiotic behaviour between T. aestivum and T. magnatum is understandable, given that the two species are related distantly phylogenetically (Jeandroz et al., 2008). Anyway, we should take into account that our study refers to a truffle cultivated orchard where T. aestivum strains were intentionally introduced.

In our case study, T. aestivum was found to be a component of the ectomycorrhizal community in 40% of the trees sampled; this abundance was higher in the case of the root samples from hornbeam trees (50%) than for those from hazel (30%). In both of these plant symbionts, a large variation in the relative abundance was also identified.

When detected, ectomycorrhizae of the summer truffle always occurred alongside other ectomycorrhizal species, but its ectomycorrhizae were not detected in 60% of the samples, which were all obtained from productive trees. The absence of T. aestivum ectomycorrhizae in the samples from some plants that were still producing truffles might reflect the fact that the production of fruiting bodies of ectomycorrhizal fungi does not correlate with their distribution and abundance on a host root system (Horton & Bruns, 2001). The same situation was found for T. melanosporum (Rubini et al., 2011a), while the opposite situation was found for T. magnatum where ectomycorrhizae and mycelium in soil were present in a nonproductive area (Murat et al., 2005; Zampieri et al., 2009). However, the ecological niche of this species might also have a limited spatial distribution that cannot always be detected by random sampling.

The ectomycorrhizal species that were observed, which belonged to the Thelephoraceae, Tuberaceae, Pyronemataceae, Sebacinaceae, Inocybaceae, and Tricholomataceae families, are fairly common mycobionts in various communities of ectomycorrhizal fungi in plants (Kõljalg et al., 2000; Glen et al., 2002; Selosse et al., 2002; Avis et al., 2003; Urban et al., 2003) and are consistent with those reported previously for environments in which truffles are produced (Murat et al., 2005; Baciarelli Falini et al., 2006; Iotti & Zambonelli, 2006; Pruett et al., 2008; Iotti et al., 2010). Our data confirmed that tomentelloid fungi are widespread and an important component of orchards that contain Tuber spp. (De Miguel et al., 2002; Murat et al., 2005; Pruett et al., 2008; Iotti et al., 2010), as well as fungal communities in forest trees (Kõljalg et al., 2000; Avis et al., 2003; Jakucs & Erős Honti, 2008), and previously infected or noninfected seedlings after outplanting (Walker et al., 2005). However, we found that they were more abundant in hazel than in hornbeam trees.

In contrast, Russulaceae, which are very common in many ectomycorrhizal communities (Horton & Bruns, 2001; Avis et al., 2003), and Cenococcum geophylum, which is considered to be one of the most widespread ectomycorrhizal species (Iotti et al., 2010), were not detected. The absence of C. geophylum is probably due to the high pH and calcium content of the calcareous soils in which truffles grow (Hall et al., 2007); the presence of lime is known to influence the structure of ectomycorrhizal communities (Kjøller & Clemmensen, 2009; Rineau & Garbaye, 2009). Similarly, we did not record any species of Scleroderma and Cortinarius, which have been reported to be common in truffle grounds (Baciarelli Falini et al., 2006; Pruett et al., 2008). Pyronemataceae, Sebacinaceae, and Inocybaceae were common in the samples analysed, but their distributions were influenced markedly by the type of host plant (Table 2). Pyronemataceae (PAPM-Mycorrhiza) species that belonged to the AD-type morphotype were the most abundant in hazel trees; this morphotype is one of the most well-known competing fungi in both cultivated and natural truffle grounds (Baciarelli Falini et al., 2006; Rubini et al., 2011b). One of the two Sebacinaceae species was abundant, especially in hornbeam trees; interestingly, Sebacinaceae spp. have been reported to occur alongside T. magnatum in the root systems of host plants (Murat et al., 2005). Moreover, Inocybaceae spp. were only present in hornbeam trees, although at a low relative abundance.

If competition between ectomycorrhizal species is one of the most important factors that determine the structure of the ectomycorrhizal community (Bruns, 1995), indigenous congeneric species such as T. rufum, T. brumale, and T. rapeodorum, which were found in this study, could compete strongly and directly for natural resources with cultivated T. aestivum. In orchards, T. melanosporum is often replaced by T. aestivum and T. brumale (Garćιa Montero et al., 2008). The replacement of Tuber spp. by indigenous fungi often occurs in orchards and can lead to the formation of small assemblages that comprise numerous species, but have an unclear influence on truffle production (Donnini & Bencivenga, 1995; Baciarelli Falini et al., 2006).

The total observed richness of 29 ectomycorrhizal taxa (25 for hazel and 17 for hornbeam) for the area investigated in this study was similar to that found for other communities of truffle ectomycorrhizal fungi (Donnini et al., 1999; Murat et al., 2005; Baciarelli Falini et al., 2006; Iotti & Zambonelli, 2006; Pruett et al., 2008) or in small monocultural forests with different plant species in which different management practices were applied (Bruns, 1995; Taylor & Bruns, 1999; Korkama et al., 2006). In general, in these studies, the observed richness was lower than the potential ectomycorrhizal richness that was calculated for the study site owing to the finite sampling size, high diversity, and different abilities of ectomycorrhizal species to form ectomycorrhizae (Gardes & Bruns, 1996; Gehring et al., 1998; Hughes et al., 2001). In our case, the observed richness corresponded to 85% of the potential richness.

The statistical differences between ectomycorrhizal taxa, diversity indices, accumulation and rank-abundance curves, Pearson correlation tests between different plots, the NMS ordination, and the anosim test that were reported in this paper confirmed that the structure of the ectomycorrhizal community was differentiated between the two different species of host tree. Hornbeams showed lower levels of richness and ectomycorrhizal diversity than hazels. The ectomycorrhizal samples that were collected from beneath hornbeams showed strong relatedness to each other because of the similar pattern of ectomycorrhizal fungi and the homogeneous distribution of ectomycorrhizal species throughout the entire study area. However, for hazel, the diversity of the ectomycorrhizal species was higher and no clear distribution pattern was observed. There are several possible explanations for the recording of a higher number of ectomycorrhizal species for hazel than for hornbeam. (1) In the study area, the hazel trees were very close to the edge of the orchard and as a consequence it was likely that they would experience a type of edge effect, even though the trees at the boundaries of the orchard were discarded in the sampling phase (Dickie et al., 2002; Dickie & Reich, 2005; Hubert & Gehring, 2008). (2) Hazel trees are known to have a shallow root system, which makes them more accessible to possible sources of inoculum, fungal spores, and other propagules in the upper soil layer. (3) Hazel is a shrubby component of the natural Mediterranean environment and a pioneer species (Walker, 1975; Čušin & Dakskobler, 2006), which makes it more susceptible than other species to undergo ectomycorrhizal symbiosis under particular ecological conditions.

Host effects could be another important factor that determines the local structure of ectomycorrhizal diversity. In fact, in assemblages with numerous ectomycorrhizal species, different fungal species can access different sources of nutrients in the soil with different levels of efficiency (Jonsson et al., 2001; Lilleskov et al., 2002) and plants can allocate resources selectively to ‘preferred and ideal’ ectomycorrhizal associates (Dickie, 2007). Ishida et al. (2007) demonstrated that, in Japanese mixed forest, a preference for a certain host taxon over other taxa affects the ectomycorrhizal fungi community in forest ecosystems directly, and this preference is stronger with increased taxonomic distance and successional status of the host. Analogously, in a study in a wet sclerophyll forest in Tasmania, Tedersoo et al. (2008) identified ectomycorrhizal species that strongly preferred certain hosts, although they did not show a preference for a single species, and showed that these species can dominate the community underground. They found that the frequency of two-thirds of the most common ectomycorrhizal fungi was influenced by host species. Evidence of a preference for a particular host plant was also found by Iotti et al. (2010) in an area that contained natural, mixed-host T. borchii: the ectomycorrhizal community that was found in oak roots differed significantly from that found in pine. Morris et al. (2009) suggested that evergreen vs. deciduous habitats also influence the structure of the ectomycorrhizal community. The work of these authors on the ectomycorrhizal community in Californian woodlands showed that even congeneric oak species (Quercus douglasii and Quercus wislizeni) could create distinct ecological niches for ectomycorrhizal fungi.

In accordance with these previous findings, our data suggest the possibility that a host-preference effect exists and is involved in the partitioning of the ectomycorrhizal fungal community between hornbeam and hazel in the T. aestivum orchard under study. In particular, the anosim test provided significant evidence that there were two distinct communities, although there was some overlap, and confirmed the different spatial distributions of the samples that were shown by the NMS ordination plot. In terms of the scope of host preference, we confirmed that plant species of the same family (Fagaceae) that lived in the same area developed ectomycorrhizal relationships that were predominantly host mediated, but not host specific.

Truffles had been produced consistently by the analysed trees over the last 3 years. This suggested that truffle production was not affected by the observed differences between the ectomycorrhizal communities that were associated with hazel and hornbeam or by the presence or absence of a particular ectomycorrhizal species. Future studies on the complex interactions among host plants and other factors, such as soil properties and soil biota, at a smaller spatial scale are needed in order to understand the direct and indirect effects of host species on the structure of ectomycorrhizal communities better.

This in-depth and high-quality study of an ectomycorrhizal community in a mature and productive T. aestivum plantation, using both morphological and molecular approaches, provides considerable insight on truffle habitats. This knowledge is clearly useful for future planning, management, and long-term maintenance of truffle plantations. Commercially, it is important to understand the effects of different host trees on ectomycorrhizal diversity in order to identify the species that are best adapted for use in future plantations and to promote appropriate agronomical practices to increase truffle production.


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

We are grateful to the ‘Comunità Montana dei Monti Martani, Serano e Subasio’ and in particular to V. Campagnani, V. De Angelis, and M. Bartolomei for providing us with the opportunity to conduct this study on a truffle plantation and to operate freely in the field. We are indebted to Prof. F. Panella and Prof. F. Sarti of the Department of Applied Biology, University of Perugia, for invaluable help and advice regarding the statistical analysis of the data. Corrections and suggestions to improve the manuscript that were made by anonymous reviewers are appreciated greatly. T.G. was financed by the Ministry of Higher Education, Science, and Technology of the Republic of Slovenia through research project CRP-V4-0492.

Authors' contribution

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

G.M.N.B., L.R. and E.A. contributed equally to this work and must be considered first authors.


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