Introduction

The Burgundy truffle (Tuber aestivum Vittad.) is the highly priced fruit body of a hypogeous ascomycete fungus (Chevalier and Frochot 1989), which is forming ectomycorrhizae to promote plant-assimilate uptake for fungal growth and to enhance water and nutrition uptake of the host plant (Smith and Read 1997; Pennisi 2004). Besides the biological relevance for ecosystem functioning, the Burgundy truffle is an important economical factor in many southern European regions (Chevalier and Frochot 1989). T. aestivum mainly depends on mutual relationships with angiosperm hosts including Quercus spp., Fagus sylvatica L. and Corylus avellana L. (Chevalier and Frochot 1989; Smith and Read 1997). In contrast, Tuber sp. such as Tuber puberulum Berk. & Broome and Tuber borchii Vittad. prefer gymnosperm hosts (Bonito et al. 2010). One well-known exception is the frequent symbiotic association of T. aestivum to Pinus nigra Arnold in Austria (Urban and Pla 2008). Nevertheless, when focusing on ectomycorrhizal symbiosis, host specificity plays an important role but is not clearly defined for most wildlife associations (Bruns et al. 2002). Our traditional concept of truffle ecology is dominated by host specificity for deciduous trees (Hilszczanska et al. 2008), mostly occurring under warm climate conditions (Bonito et al. 2010). The ecological niche of Tuber spp. is generally assumed to be rather narrow (Büntgen et al. 2011), and the genus' true “specialists” such as Tuber magnatum Pico are restricted to small areas characterized by particularly suitable environmental parameters (Mello et al. 2004). Despite new truffle findings outside commonly accepted biogeographical boundaries (Stobbe et al. 2012), remain the microbiology, site ecology and host specificity of T. aestivum yet poorly understood (Bonito et al. 2010; Büntgen et al. 2011).

Here, we use two methodologically independent approaches to investigate whether an ectomycorrhizal symbiosis between T. aestivum and Picea abies in the wild exists, and if it can be obtained artificially in the greenhouse: (1) Morphological and molecular identification of ectomycorrhizae from pure wildlife P. abies stands. (2) Greenhouse inoculation of P. abies seedlings with T. aestivum, and subsequent morphological and molecular identification of their developed ectomycorrhizae. Results are discussed in the light of the potential cultivation of T. aestivum across central and eastern Europe.

Material and methods

Field observation

Two T. aestivum sites (I and II) in pure P. abies forests in southwest Germany, for which vegetation, geology, and climate is known (Table 1), were identified with trained truffle dogs (Bracco francese), and root samples were collected in November 2011, during the peak of truffle fruiting. Both sites have rendzic soils on Jurassic parent material with pH values (in H2O) between 7.26 and 7.96. In the understory of site I and II occurred a young growth of Quercus robur L. and F. sylvatica, respectively. The roots of these plants were included in the root samples and accordingly checked for T. aestivum ectomycorrhizae.

Table 1 Habitat characteristics of field sites I and II. Climatic data was obtained through DWD (Deutscher Wetterdienst; federal ministry of transport, building, and urban development) from the nearest weather stations. Potential host plants are in bold text. The lower part shows field study results with T. aestivum mycorrhization rates and presence of fruit bodies (ST-samples)
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Nine samples of topsoil containing fine roots (∼100 g each) were taken on a 2×2 m plot with a 1×1 m grid on each site. Additional samples were taken in the immediate periphery of the discovered fruit bodies and tagged ST (2 on site I and 5 on site II; Table 1). These samples were collected outside the grid points within the plots and 1 m around them. After gently washing off the soil with tap water, the fine roots were examined under a stereo microscope (Leica EZ4D, magnitude ×5–35) to detect the presence of T. aestivum ectomycorrhizae. Due to their tangly morphology, the root tips were counted roughly to provide ectomycorrhization rates in 5 % steps. Identification was performed with a light microscope (Zeiss AXIOPHOT, magnitude ×50–400) using the keys and illustrations by Agerer (1987–2006). Samples were taken from main roots adjacent to ectomycorrhized root tips to identify the associated host tree species by microscopic examination of thin-sections (magnitude ×50–400) (Schweingruber 1990). For molecular identification of fungal and plant partners, we randomly sampled 13 additional soil cores (0–25 cm) on site II, which were proceeded as indicated above. A total of 91 T. aestivum morphotype ectomycorrhizae were collected and subjected to DNA analyses.

Greenhouse experiment

Fifty P. abies plants were grown from seeds in a calcareous rendzic soil (pH 8.4) in 425 ml pots. The substrate was inoculated with 5 × 106 T. aestivum spores per plant according to the method described by Weden et al. (2009). As spore source, ripe fruit bodies with a dark brown gleba from southwest Germany were used. After 16 months, the fine root ectomycorrhization was analyzed (Fischer and Colinas 1996). To conduct the morphological examination, the root system was washed with water, cut into ∼2 cm pieces, and placed in a shallow dish with water over a 1 × 1 cm grid. At least 250 root tips per plant were counted in randomly chosen squares and categorized in “nonectomycorrhized” (N), “T. aestivum-ectomycorrhized” (T), and “other ectomycorrhizae” (K). The percentage of “T” and “K” was calculated, and descriptive statistics were performed with the software SPSS 19.0 (IBM). A total of 62 ectomycorrhized root tips from six plants were randomly sampled and preserved (frozen) for molecular identification.

Molecular identification

DNA extraction was performed with 91 ectomycorrhized root tips from site II and 62 root tips from the greenhouse experiment using NucleoSpin 96 PlantII DNA extraction kit (Macherey-Nagel GmbH), and following the protocol with the use of cell lysis buffer PL1 (CTAB) and an elution volume of 50 μl. The internal transcribed spacer (ITS) region of rDNA was amplified using the primer pairs Tu1sekvF/Tu2sekvR (Gryndler et al. 2011) and/or UNCI/UNCII (Mello et al. 2002) selective for T. aestivum. We used 2–5 μl of a 1:10 dilution of DNA (1–5 ng DNA) in a reaction volume of 25 μl. The REDTaq DNA polymerase and REDTaq PCR reaction buffer (1.1 mM MgCl2 final concentration) were used according to the protocol of SIGMA, with 200 μM of each dNTP, 0.2 μM of each primer, and 75 μg BSA (SIGMA, Cat no. B4287). The PCR was conducted in a Veriti Thermalcycler (Applied Biosystems) using an annealing temperature of 52 (Tu1sekvF/Tu2sekvR) and 59 °C (UNCI/UNCII) (Gryndler et al. 2011). PCR products were run on 1.0 % agarose gel and visualized with ethidium bromide. In all PCR reactions, a positive (DNA from a T. aestivum fruit body) and two negative controls (no DNA, DNA from Tuber melanosporum) were included. The presence of a PCR band was interpreted as a positive identification of T. aestivum in the respective sample. When no band was visible, we interpreted it as either the presence of a different ectomycorrhizal species at the root tip or as PCR failure of this sample.

To verify the host plant species of ectomycorrhizae sampled in the field, we amplified the plastid trnL intron in a second PCR reaction using DNA quantity and PCR ingredients as indicated above with primers and cycling parameters published by Brunner et al. (2001). Subsequently, restriction fragment length polymorphism (RFLP) analyses were performed, using the restriction enzyme Taq1 (Brunner et al. 2001). Based on the on-site tree species composition, RFLP analyses using Taq1 unambiguously distinguished P. abies from other species.

Results

Samples from site I and II revealed morphological evidence for T. aestivum ectomycorrhizae (Fig. 1b) in four out of eleven and six out of twelve samples, respectively (Table 1). Due to the presence of a distinctive feature combination of brown–red ectomycorrhiza color, angular mantle cells and wooly cystidia, morphological distinction of T. aestivum ectomycorrhizae from other ectomycorrhizae could be performed without difficulties. These features are illustrated in Fig. S 2 (supplementary material). Microscopic investigation of root thin sections identified P. abies as host tree according to the presence of characteristic resin canals in all cases (Fig. 1c). No T. aestivum ectomycorrhiza was detected in the root systems of the tree species occurring as young growth in the understory of the spruce stands. Details on sample numbers, ectomycorrhization data, and host tree species are provided in Table 1.

Fig. 1
figure 1

a Mycorrhization rate (mean, standard deviation, range) of 50 P. abies seedlings from the greenhouse experiment, b T. aestivum mycorrhiza on P. abies fine roots with characteristic appearance, c thin section of P. abies root with characteristic resin canals (circles) used for morphological fine root identification, and d the natural P. abies distribution overlapped with suitable soil pH (>7) for truffle habitats. The sample sites are marked red. The overlap covers an area of approximately 287.556 km2 (pH-map: FAO/IIASA/ISRIC/ISSCAS/JRC, 2012. Harmonized world soil database (version 1.2). FAO, Rome, Italy and IIASA, Luxemburg, Austria; P. abies distribution map: Euforgen, Bioversity International, Rome, Italy http://www.euforgen.org/distribution_maps.html)

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The molecular analyses of samples from site II confirmed the ectomycorrhizal association between P. abies and T. aestivum. The plant trnL intron was successfully amplified from 89 out of 91 ectomycorrhizae, and RFLP analyses identified P. abies as the host species in all samples. Of these 89 P. abies ectomycorrhizae, 72 % were assigned to T. aestivum. From the remaining 28 % of the P. abies ectomycorrhizae, the T. aestivum specific ITS PCR was negative.

The inoculation of P. abies with T. aestivum spores in the greenhouse experiment was positive on 27 out of the 50 seedlings. Morphological identification and quantification showed a mean ectomycorrhization rate of 19.6 % (range 0–69.4 %, STD 24.4 %) (Fig. 1a). As the other roots remained free of ectomycorrhizal colonization, no contaminating fungi were detected in the P. abies root systems. After 16 months, the plants had a mean height of 7.6 cm (minimum 3.8 cm/maximum 12.3 cm) and showed no signs of malnutrition or pathogens.

Seventy-four percent of the molecularly analyzed ectomycorrhizae from six P. abies seedlings were successfully amplified using T. aestivum specific primers and can therefore be assigned to T. aestivum. This broadly confirms the successful synthesis of P. abies/T. aestivum ectomycorrhizae.

Discussion

Our biological findings broaden the so far reported ecological niche of the Burgundy truffle, which was often restricted by the definition of host tree distribution and suitable climate (Hilszczanska et al. 2008). T. aestivum, however, can adapt to a much wider variety of habitats within calcareous soil conditions, instead of being limited to areas with characteristic deciduous hosts such as oak (Quercus spp.), beech (F. sylvatica), and hazel (C. avellana). P. abies inhabits a different distribution range, which generally extends to higher elevations and more continental settings (Ellenberg 1988). Nonetheless, T. aestivum studies from regions with prevalent P. abies habitats such as Poland, Czech Republic, and Slovakia do not refer to P. abies as a host plant (Hilszczanska et al. 2008; Miko et al. 2008; Streiblova et al. 2010), possibly due to a biased focus on habitats with deciduous vegetation. For a more comprehensive view on T. aestivum biology, a wider variety of calcareous habitats and a broader host tree range should be taken into consideration.

The natural distribution range of P. abies, characterized by cold climate and acidic soils, was expanded to calcareous soils for silvicultural reasons (Spiecker et al. 2004), which led to a vast overlap with T. aestivum distribution (Fig. 1d). Even if this was sometimes regarded as a mistake with unwanted ecological and economic consequences such as poor biodiversity, increased risk of pathogens, and drought stress (Spiecker et al. 2004), it revealed an unexpected adaptability to a wider range of symbiotic partners of the Burgundy truffle. By proving P. abies to be the host tree of T. aestivum on two productive truffle sites in the wild, our findings indicate the species' potential of supporting truffle production in commercially attractive amounts. Nevertheless, besides the presence of T. aestivum ectomycorrhizae, suitable habitat characteristics are probably the key factor for fruit body production.

The extended range of T. aestivum hosts will likely have beneficial implications for truffle cultivation, since P. abies roots remained free of contamination with other fungal species than T. aestivum in the greenhouse experiment. A reason might be the poor ability of P. abies-specific ectomycorrhizal fungi to adapt to the high pH substrate (Lehto 1994). Picea abies might therefore be a suitable and trouble-free task for producing T. aestivum seedlings in nurseries, which frequently have to deal with contamination (Hall et al. 2003). However, production methods have to be refined in further studies to ensure better success, before a commercial use is advisable. Particularly in eastern European regions, where P. abies and T. aestivum distributions overlap, truffle cultivation with P. abies could be a promising opportunity. Furthermore, mixed stands on truffle orchards would reduce the risk of pathogens and promote high biodiversity.