For germination and establishment, orchids depend on carbon (C) and nutrients supplied by mycorrhizal fungi. As adults, the majority of orchids then appear to become autotrophic. To compare the proportional C and nitrogen (N) gain from fungi in mycoheterotrophic seedlings and in adults, here we examined in the field C and N stable isotope compositions in seedlings and adults of orchids associated with ectomycorrhizal and saprotrophic fungi.
Using a new highly sensitive approach, we measured the isotope compositions of seedlings and adults of four orchid species belonging to different functional groups: fully and partially mycoheterotrophic orchids associated with narrow or broad sets of ectomycorrhizal fungi, and two adult putatively autotrophic orchids associated exclusively with saprotrophic fungi.
Seedlings of orchids associated with ectomycorrhizal fungi were enriched in 13C and 15N similarly to fully mycoheterotrophic adults. Seedlings of saprotroph-associated orchids were also enriched in 13C and 15N, but unexpectedly their enrichment was significantly lower, making them hardly distinguishable from their respective adult stages and neighbouring autotrophic plants.
We conclude that partial mycoheterotrophy among saprotroph-associated orchids cannot be identified unequivocally based on C and N isotope compositions alone. Thus, partial mycoheterotrophy may be much more widely distributed among orchids than hitherto assumed.
The production of tiny dust-like seeds is a common feature of orchids; the biomass of orchid seeds varies from 0.3 to 24 μg dry weight (Arditti & Ghani, 2000). As a consequence of their lack of endosperm (Rasmussen, 2002) and storage of only marginal amounts of carbon (C) reserves within the embryo (Manning & van Staden, 1987), orchid seeds are dependent for their germination and establishment on C and nutrients supplied by fungal partners. This kind of C and nutrient gain from fungi in the early seedling (protocorm) stage is called initial mycoheterotrophy (Leake, 1994; Merckx, 2013). There is broad agreement that protocorms of terrestrial orchids are often extremely slow to develop in nature; it may take years until seedlings produce for the first time a photosynthetic leaf above the soil surface (Rasmussen & Whigham, 1993, 1998; Leake, 1994; Smith & Read, 2008). Quantitative information about growth in this initially mycoheterotrophic stage and about nutrient gains from fungal symbionts under natural conditions, however, is almost entirely missing.
As adults, the majority of orchid species produce green leaves and become putatively autotrophic. However, among the > 20 000 orchid species there are c. 200 that remain nonphotosynthetic and entirely dependent on fungal C, and probably nutrients, throughout their entire life cycle. These orchids are called full mycoheterotrophs (Leake, 1994; Merckx, 2013). In addition, some species of green orchids obtain C both from their own photosynthesis and from their fungal symbionts (Gebauer & Meyer, 2003). These orchids are called partial mycoheterotrophs (Gebauer & Meyer, 2003; Merckx, 2013). Partial mycoheterotrophy is a particular case of plant mixotrophy.
Over a century ago, a group of abundant and easily cultivable, asexual, saprotrophic fungi was identified as major mycorrhizal symbionts of orchids (Bernard, 1909). This group of fungi was called ‘rhizoctonia’. Based on molecular systematics it is now clear that rhizoctonia fungi are a phylogenetically heterogenous assemblage (Dearnaley et al., 2012). More recently, mostly using molecular techniques, other groups of fungi have been identified as capable of forming orchid mycorrhizas. Among fully and partially mycoheterotrophic orchids, association with hardly cultivable basidiomycetes and ascomycetes that simultaneously form ectomycorrhizas with forest trees is widely distributed. Furthermore, for some fully mycoheterotrophic orchids, associations with nonrhizoctonia saprotrophic wood-decomposer or litter-decaying fungi have been reported (Hynson & Bruns, 2010).
The different functional groups of fungi forming orchid mycorrhizas have access to different soil-derived and/or plant-derived C compounds and nutrients, which leads to characteristic stable isotope abundance patterns in fungal tissues (Gebauer & Dietrich, 1993; Gleixner et al., 1993; Gebauer & Taylor, 1999; Kohzu et al., 1999; Taylor et al., 2003) and consequently also in orchids that gain C and nutrients from these various fungi. Early stable isotope investigations showed that adult fully mycoheterotrophic orchids are significantly enriched in the heavy C and nitrogen (N) isotopes 13C and 15N in comparison to neighbouring autotrophic plants (Gebauer & Meyer, 2003; Trudell et al., 2003), and that adult partially mycoheterotrophic orchids are positioned between fully mycoheterotrophic orchids and autotrophic plants (Gebauer & Meyer, 2003). Based on these early studies and a series of subsequent investigations, we are now able to distinguish through their characteristic C and N stable isotope abundance patterns the following adult orchid nutritional strategies: fully mycoheterotrophic associated with ectomycorrhizal fungi; fully mycoheterotrophic associated with wood-decomposer fungi; partially mycoheterotrophic associated with ectomycorrhizal fungi; and putatively autotrophic associated with rhizoctonia (Hynson et al., 2013a).
Hitherto missing is information about the C and N isotope compositions of orchids in their initially mycoheterotrophic stage. Here we compare, to our knowledge for the first time, stable isotope natural abundance data of mycoheterotrophic protocorms and adults of four orchid species symbiotic with fungi belonging to three different functional groups: an adult fully mycoheterotrophic orchid associated with a single ectomycorrhizal fungal lineage (Neottia nidus-avis (L.) L. C. Rich.); an adult partially mycoheterotrophic orchid associated with a set of ectomycorrhizal fungi (Epipactis helleborine (L.) Crantz); and two adult putatively autotrophic orchid species associated with rhizoctonia fungi (Serapias parviflora Parl. and Pseudorchis albida (L.) A. & D. Löve). We hypothesized that in all cases protocorms mirror the isotope composition of C and N gained from the fungal source and, thus, are ideally suited to help identify potential C and N gains in adults even of those orchid species that become photosynthetic. In addition to C and N isotope compositions, we identified the fungi associated with orchid protocorms, and investigated relationships between protocorm biomass, total C and N pool sizes and protocorm total N concentrations.
Materials and Methods
Study sites, seed germination and sample collection
The orchid seeds used for germination originated from ripe capsules collected at natural sites in Sardinia (N. nidus-avis, E. helleborine and S. parviflora) and the Czech Republic (P. albida). The mean biomass per seed ranged from 0.6 μgDW (S. parviflora) to 5.4 μgDW (N. nidus-avis). Following the seed packet method of Rasmussen & Whigham (1993), c. 100 seeds were transferred into nylon net pockets of 50-μm mesh (Plastok, Birkenhead, UK) or 42-μm mesh (Silk and Progress Ltd, Brněnec, Czech Republic) and sealed with a photographic slide frame. The seeds were not pretreated by cold stratification or chemical scarification. In October (P. albida) and December (N. nidus-avis and E. helleborine) 2007 or January 2008 (S. parviflora), 120 (P. albida), 300 (E. helleborine and S. parviflora) or 400 (N. nidus-avis) of these seed packets were buried vertically 1 cm below the soil surface at natural sites where the respective orchid species occur. For in situ seed germination the following four sites were chosen.
Sardinia, 39.9°N, 9.5°E: evergreen forest dominated by Quercus ilex for germination of N. nidus-avis.
Continental Italy, 44.4°N, 8.2°E: mixed deciduous forest dominated by Fagus sylvatica and Corylus avellana for germination of E. helleborine.
Tenerife, 28.3°N, 16.8°W: grassland with Erica arborea shrubs for germination of S. parviflora.
Czech Republic, 49.1°N, 13.6°E: Nardus grassland with scattered Vaccinium shrubs and Picea abies trees for germination of P. albida.
In April (S. parviflora) or May (N. nidus-avis and E. helleborine) 2009, that is, after c. 1 yr of burial, or in July 2011 (P. albida), that is, after nearly 4 yr of burial, seed packets were excavated and stored moist at 4°C for a maximum of 1 wk until processing. In addition, leaf or stem samples of flowering adults of the respective orchid species (n =5) and leaf samples of three or four autotrophic nonorchid reference plant species per site were collected following the sampling protocol described by Gebauer & Meyer (2003). In two cases (sites 1 and 3), C and N isotope abundance and C and N concentration data for the respective orchid adults and reference plants were available from a previous investigation (Liebel et al., 2010) and in those cases we did not re-sample orchid adults and reference plants. At site 2 no adult leaves of E. helleborine could be found at the time of seed packet collection.
Thirteen plant species served as autotrophic nonorchid reference plants (n =5): site 1, Cyclamen repandum S. et S., Mycelis muralis (L.) Dumort. and Quercus ilex L.; site 2, Calystegia sepium (L.) R. Br., Hedera helix L. and Primula vulgaris Huds.; site 3, Avena barbata Pot. ex Link, Erica arborea L. and Plantago lagopus L.; site 4, Hieracium laevigatum Willd., Melampyrum pratense L., Nardus stricta L. and Vaccinium vitis-idaea L.
About half of the originally buried seed packets were used for protocorm preparation. Germination success was assessed using a dissecting microscope. Thirty-four packets contained a total of 259 germinated individuals. The numbers of packets and protocorms were distributed among the orchid species unequally: N. nidus-avis, 46 protocorms in 11 packets; E. helleborine, 196 protocorms in 15 packets; S. parviflora, five protocorms in two packets; and P. albida, 12 protocorms in six packets. One protocorm per packet was placed in 300 μl of lysis buffer (cetrimethyl ammonium bromide (CTAB)) for fungal DNA analysis. The majority of E. helleborine protocorms were visually much smaller than all other protocorms. For this reason, 177 E. helleborine protocorms were pooled in seven batch samples for biomass determination and isotope abundance analysis (see Supporting Information Table S1). All other protocorms, in total 48, were treated individually. Protocorms were dried at 105°C to constant weight for the determination of biomass, C and N concentrations and stable isotope abundance. The dry weight of the protocorms was determined using a micro balance (CPA 2 P; Sartorius AG, Göttingen, Germany) with repeatability from 0.001 to 0.002 mg at a weighing range from 0.5 to 1 mg. Biomass data for E. helleborine protocorms from batch samples are mean values for four to 36 individuals (see Table S1).
Analysis of C and N concentrations and stable isotope abundances
Leaf and stem plus scale-like leaf samples (only the latter for N. nidus-avis) of adult orchids and reference plants were oven-dried at 105°C, ground to a fine powder in a ball mill (Schwingmühle MM2; Retsch, Haan, Germany), and stored in a desiccator until further analysis. From 2.5 to 3.5 mg of this ground material was used for the analysis of relative N and C isotope abundances in a dual element analysis mode with an elemental analyser (1108; Carlo Erba, Milano, Italy) coupled to a delta S gas-isotope ratio mass spectrometer (Finnigan MAT, Bremen, Germany) through a ConFlo III interface (Thermo Fisher Scientific, Bremen, Germany). Isotope abundances are denoted as δ values according to the following equation: δ13C or δ15N = (Rsample/Rstandard − 1) × 1000 (‰), where R is the ratio of heavy to light isotopes of the sample or the respective standard. Standard gases were calibrated with respect to international standards using the reference substances N1 and N2 for N isotopes and ANU sucrose and NBS 19 for C isotopes, all provided by the International Atomic Energy Agency (IAEA) (Vienna, Austria). The reproducibility and accuracy of the isotope abundance measurements were routinely controlled by measuring the test substance acetanilide (Gebauer & Schulze, 1991). At least six test substances with varying sample weight were routinely analysed within each batch of 50 samples. The maximum variation of δ13C and δ15N both within and between batches was always below 0.2 ‰.
Few protocorms had individually sufficient biomass for the above-described analytical procedure. Therefore, a different analytical approach was developed. The two approaches were basically cross-checked by using identical IAEA standards for calibration and the identical test substance acetanilide for quality control. We coupled an elemental analyser (NA 1500; Carlo Erba) with a more sensitive gas-isotope ratio mass spectrometer (Delta V Plus; Thermo Fisher Scientific) through a ConFlo III interface (Thermo Fisher Scientific). We furthermore checked the accuracy and reproducibility of the δ13C and δ15N measurements in the critical range below 1 V peak height using acetanilide. Even in the range from 1 V to 50 mV peak height, the δ13C of acetanilide did not vary with sample amount, and only the maximum variation increased from 0.2 to 0.3‰. Therefore, no further correction of the δ13C data was required. For the measurement of δ15N in the peak height range below 1 V signal intensity, a blank correction had to be performed to account for minor contamination with atmospheric N2. The blank corrected measurement of δ15N also revealed no dependence on sample amount down to peak areas of 25 V s at m/z 28 (equals c. 450 mV peak height). However, at peak areas below 25 V s at m/z 28 there was a drift in δ15N with decreasing sample amount (Fig. 1). In order to correct for this, we calculated a logarithmic regression curve (δ15Nuncor = 0.9305 × loge(aream/z28) − 1.8307; F1,23 = 50.2; r²adj = 0.67; P <0.001), where δ15Nuncor is the uncorrected δ15N value of the sample and aream/z28 is the peak area at m/z 28. For every δ15Nuncor value, the difference (Diff) from the reference value of the test substance (acetanilide with a δ15N of 0.895‰) was calculated and subtracted from the δ15Nuncor value. After this correction, the maximum variation of δ15N was 0.4‰. This analytical approach allowed the δ13C and δ15N measurement of individual protocorms down to biomasses of c. 0.030 mgDW with satisfactory accuracy and reproducibility (see Table S1). Epipactis helleborine protocorms with mostly far lower biomass were pooled and analysed together (see ‘Protocorm preparation’ above).
Total C and N concentrations in leaf, stem and protocorm samples were calculated from sample weights and peak areas using a six-point calibration curve per sample run based on acetanilide measurements (Gebauer & Schulze, 1991). Acetanilide has a constant C concentration of 71.09% and N concentration of 10.36%.
Fungal DNA analysis
A subset of 34 protocorms from different packets (see ‘Protocorm preparation’ above) were sampled, placed in CTAB and stored at −32°C. Genomic DNA of P. albida was extracted as described in Gardes & Bruns (1993). For the three other orchid species the same method was used, with the only modification being the use of a GeneClean III Kit (Q-BioGene, Carlsbad, CA, USA) to bind and purify the DNA. Following the methods described in Bidartondo & Duckett (2010) the nuclear ribosomal internal transcribed spacer (ITS) region was amplified with the fungal-specific primers ITS1F/ITS4 and ITS1/ITS4-tul. Fungal ITSs from P. albida protocorms were amplified using ITS1OF/ITS4OF following the thermal cycling profile in Taylor & McCormick (2008) but with the annealing temperature decreased to 55°C. Most PCR products showed a single band and were purified with ExoSap-IT (Affymetrix, High Wycombe, UK) and sequenced bidirectionally with an ABI3730 Genetic Analyzer using the BigDye 3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, CA, USA) and absolute ethanol/EDTA precipitation. Double-banded PCR products in two P. albida protocorms were excised separately from the gel, purified using the JetQuick Gel Extraction Spin Kit (Genomed, Löhne, Germany), and resequenced. After checking electrophoretograms using Sequencher 4.5 (Gene Codes, Ann Arbor, MI, USA) all sequences were compared to GenBank using BLAST (http://www.ncbi.nlm.nih.gov) to ascertain taxonomic affinity. All unique DNA sequences have been submitted to GenBank (KF386021, KF386022 and KF850619–KF850623).
Calculations and statistics
To compare isotope abundances of the adult orchids and protocorms from different sites, the data were normalized. Enrichment factors (ε) were calculated per site: ε = δS − δREF, where δS is a single δ13C or δ15N value for an adult orchid or a protocorm and δREF is the mean value for nonorchid reference plants from the respective site (Preiss & Gebauer, 2008). The original δ13C and δ15N values of orchid protocorms and adults and of the respective reference plants are available in Table S1. The following data were analysed for the significance of differences: total N concentrations in protocorms, orchid adults and respective references for each of the four orchid species; ε13C and ε15N values in protocorms, adults and reference plants from the protocorm sampling sites and adults and reference plants from the literature for each of the four orchid species; ε13C and ε15N values in protocorms of the four orchid species; ε13C and ε15N values in initially mycoheterotrophic protocorms of the four orchid species in comparison to adults of the fully mycoheterotrophic N. nidus-avis.
A one-way ANOVA and a subsequent Tukey test were used, if data were normally distributed and variances homogenous (only in the case of ε13C for the comparison of protocorms). In all other cases a Kruskal–Wallis nonparametric test and a sequential Bonferroni-corrected (Holm, 1979) Mann–Whitney U-test for post hoc comparison were used.
Relationships between ε13C or ε15N and protocorm biomass were tested for significance using a linear correlation analysis. Statistical analyses were performed with SigmaPlot 11.0 (Systat Software Inc., San José, CA, USA).
PCR amplification and sequencing succeeded in all protocorms of N. nidus-avis, S. parviflora and P. albida, and in c. 80% of E. helleborine protocorms. The molecular identification of fungi in the orchid protocorms confirmed for N. nidus-avis a highly specialized association with Sebacina of the ectomycorrhizal clade A (Table 1). Epipactis helleborine protocorms were associated with the ectomycorrhizal fungi Tomentella, Tuber and Sebacina clade A and representatives of the Pyronemataceae. In protocorms of S. parviflora and P. albida either Tulasnella alone or Tulasnella and Sebacina of clade B, probably belonging to the saprotrophic rhizoctonia, were detected.
Table 1. Mycorrhizal fungal associates detected in orchid protocorms
Numbers in brackets indicate the number of protocorms in which the respective fungus was detected. n, number of investigated protocorms. Obligate ectomycorrhizal fungi are shown in bold. Taxa that contain some ectomycorrhizal lineages are denoted by an asterisk.
Sebacinaclade A (11)
Pyronemataceae* (5), Sebacinaclade A (3), Tomentella (1), Tuber (1)
Tulasnella* (6), Sebacina clade B* (2)
During 1 yr in the soil the germinated seedlings developed heterogeneously (Figs 2, 3). They reached a maximum biomass of 170 (E. helleborine), 360 (N. nidus-avis) or 500 μgDW (S. parviflora). After 4 yr in the soil the biomass of P. albida protocorms ranged between 170 μgDW and 3.8 mgDW (Figs 2, 3). Thus, the contribution of the initial seed biomass (0.6–5.4 μgDW; see the 'Materials and Methods' section) to the protocorm biomass was considerable only for the tiniest protocorms (Fig. 3). The differences in protocorm biomass between species were also not related to the initial seed biomass, as the species with the tiniest seeds (S. parviflora) produced by no means the tiniest protocorms. The total C and N contents in the protocorms of all four orchid species increased in a mostly linear manner with increasing biomass (Fig. 3). The amount of C and N that the orchid protocorms accumulated ranged after 1 yr in the soil between 0.2 and 18 μmol of C and between 20 and 800 nmol of N. For P. albida after 4 yr in the soil, C and N gains were between 9 and 119 μmol of C and between 1 and 13.3 μmol of N. The total N concentrations in the protocorms of the four orchid species were even higher than those of adult orchids and twofold to almost fourfold higher than N concentrations in reference plants (Table 2).
Table 2. Mean total nitrogen (N) concentrations (± 1 SD) in protocorms (n =3–35; see Fig. 3) and leaves or stems of adults (n =5) of a fully mycoheterotrophic (Neottia nidus-avis), a partially mycoheterotrophic (Epipactis helleborine) and two in the adult stage putatively autotrophic (Serapias parviflora and Pseudorchis albida) orchids and of co-occurring reference plants (three or four species in five replicates each) collected at one forest site in Sardinia (N. nidus-avis), one forest site in continental Italy (E. helleborine) or at open grassland sites in Tenerife (S. parviflora) and the Czech Republic (P. albida)
Protocorm N conc. (mmol gDW−1)
Adult N conc. (mmol gDW−1)
Reference plant N conc. (mmol gDW−1)
A Kruskal–Wallis test indicates significance of differences in total N concentrations between protocorms, adults and reference plants (H =38.7; P <0.001). Different letters indicate significant differences in total N concentrations for each of the four orchid species (U between 0 and 4, in all cases P <0.01). nd, not determined.
4.8 ± 1.0a
2.6 ± 0.1b
1.8 ± 0.7c
3.4 ± 1.0a
1.4 ± 0.5b
2.2 ± 0.8a
1.6 ± 0.2a
1.0 ± 0.1b
4.2 ± 1.7a
1.8 ± 0.2b
1.1 ± 0.3c
Comparisons of ε13C revealed significant differences: among protocorms, adults and reference plants of each of the four orchid species; among protocorms within each the four orchid species; among protocorms of the four orchid species and adults of the fully mycoheterotrophic N. nidus-avis (Table 3). Post hoc tests showed that all protocorms were significantly enriched in 13C (in all cases U =0; P <0.01) in comparison to autotrophic reference plants collected at the respective protocorm sampling sites (Fig. 4a). The relative enrichment factor ε13C of orchid protocorms in comparison to autotrophic reference plants was independent of the protocorm biomass; that is, no significant correlation between ε13C and protocorm biomass was found (r2adj between 0.39 (S. parviflora; df = 1) and 0.06 (E. helleborine; df = 9); P >0.05 for all orchid species). However, the relative enrichment in 13C was distinctive for ectomycorrhizal fungi-associated (N. nidus-avis and E. helleborine) and rhizoctonia-associated (S. parviflora and P. albida) protocorms. The mean ± SD 13C enrichments of N. nidus-avis and E. helleborine protocorms were 7.6 ± 0.6‰ (n =35) and 7.3 ± 0.7‰ (n =11), respectively, and did not differ significantly from each other (Q =2.44; P =0.321). They were also not different from those of adults of the fully mycoheterotrophic N. nidus-avis collected at one of the protocorm sampling sites (U =47; P =0.102 and U =17; P =0.257, respectively). By contrast, initially mycoheterotrophic protocorms of E. helleborine were significantly more enriched in 13C than partially mycoheterotrophic E. helleborine adults (U =0; P <0.001). The mean 13C enrichments of mycoheterotrophic protocorms of S. parviflora and P. albida were 4.2 ± 0.3‰ (n =3) and 4.9 ± 0.3‰ (n =6), respectively, with no significant difference from each other (Q =2.39; P =0.34). However, they were significantly less enriched in 13C than protocorms of the two orchid species associated with ectomycorrhizal fungi (Q between 11.51 and 15.13; in all cases P <0.001). While protocorms of P. albida were significantly more enriched in 13C than P. albida adults (U =0; P =0.004), the 13C enrichment of S. parviflora protocorms was not sufficient to distinguish them significantly from S. parviflora adults either collected at the protocorm sampling site (U =0; P =0.036; not significant after sequential Bonferroni correction (SBC)) or taken from the literature (U =0; P =0.014; not significant after SBC). Adults of S. parviflora and P. albida were not significantly different from autotrophic reference plants in their 13C enrichment after SBC (U =45; P =0.008 and U =789; P =0.045 for S. parviflora from the protocorm sampling site and from literature data, respectively; U =103; P =0.178 for P. albida).
Table 3. Results of statistical tests for differences in the enrichment factors ε13C or ε15N in the following groups of independent variables: (1) protocorms, adults and reference plants from the protocorm sampling sites and adults and reference plants from the literature for each of the four orchid species, (2) protocorms of the four orchid species, and (3) protocorms of the four orchid species and adults of the fully mycoheterotrophic Neottia nidus-avis
One-way ANOVA was used, if data were normally distributed and variances homogenous (only in the case of ε13C for the comparison of protocorms). In all other cases a Kruskal–Wallis nonparametric test was used.
In contrast to 13C, the 15N enrichment of protocorms changed significantly with biomass for two of the four orchid species (Fig. 4b). For N. nidus avis and for E. helleborine a linear increase in ε15N with growing biomass was found (ε15N = 18.87 × biomass + 5.15; r2adj = 0.56; df = 33; P <0.001 for N. nidus avis; ε15N = 16.50 × biomass + 9.22; r2adj = 0.65; df = 9; P <0.01 for E. helleborine). Comparisons between ε15N in protocorms, adults and reference plants revealed significant differences for three of the four orchid species (Table 3). Only in the case of P. albida were no significant relationships found (Table 3). Comparisons among initially mycoheterotrophic protocorms of the four orchid species and between protocorms and fully mycoheterotrophic adults of N. nidus-avis indicated significant differences (Table 3). Epipactis helleborine protocorms reached the highest 15N enrichment. In comparison to autotrophic reference plants from the protocorm sampling sites, E. helleborine protocorms were enriched by 10.5 ± 2.5‰ in 15N (U =0; P <0.001; n =11). The 15N enrichment of E. helleborine protocorms was similar to that of E. helleborine adults in the literature (ε15N = 13.1 ± 4.4‰; n =18; U =56; P =0.056). The mean 15N enrichment of N. nidus-avis protocorms was 6.5 ± 1.0‰ and thus was significantly lower than that found for E. helleborine protocorms (U =13; P <0.001; n =35). Nonetheless, N. nidus-avis protocorms were significantly enriched in 15N compared with autotrophic reference plants from the protocorm sampling site (U =0; P <0.001). Interestingly, N. nidus-avis protocorms were significantly less enriched in 15N than adults from the protocorm sampling site (ε15N = 13.9 ± 1.7‰; n =5; U =0; P <0.001) and adult data from the literature (ε15N = 11.0 ± 3.1‰; n =36; U =128; P <0.001). The mean protocorm 15N enrichments of the two orchid species associated with rhizoctonias were 1.8 ± 0.6‰ (S. parviflora; n =3) and 1.4 ± 1.3‰ (P. albida; n =6) and thus were significantly lower than the 15N enrichments in protocorms of E. helleborine (U =0; P =0.013 and U =0; P <0.001, respectively) and N. nidus-avis (U =0; P =0.005 and U =0; P =0.001, respectively) and not significantly different from each other (U =21; P =0.714). Serapias parviflora and P. albida protocorms were significantly different in 15N enrichment neither from autotrophic reference plants collected at the protocorm sampling sites (for S. parviflora, U =17; P =0.017; not significant after SBC; for P. albida, see Table 3) nor from adults of the respective species (U =2; P =0.143 and U =3; P =0.052 for S. parviflora from the protocorm sampling site and from literature data, respectively; for P. albida, see Table 3). Adults of S. parviflora had mean ε15N values of 2.8 ± 0.9‰ (data from protocorm sampling site) or 3.2 ± 1.1‰ (data from the literature) and were significantly enriched in 15N in comparison to autotrophic reference plants (U =11; P <0.001 for protocorm sampling sites and U =41; P <0.001 for literature data). By contrast, adults of P. albida had a mean ε15N value of 0.5 ± 1.2‰ at the protocorm sampling site and were not significantly different from autotrophic reference plants (Table 3).
The fungi identified here in orchid protocorms are identical to the fungi already known to form mycorrhizas with adults of the four investigated orchid species (McKendrick et al., 2002; Selosse et al., 2002; Bidartondo et al., 2004; Ogura-Tsujita & Yukawa, 2008; Liebel et al., 2010; Těšitelová et al., 2012; Kohout et al., 2013). With this finding, an essential prerequisite for our initial hypothesis was fulfilled, namely, C and N transferred from fungi to protocorms and potentially to adults of each of the four investigated orchid species originated from the same fungal clades and from, plausibly, identical fungal isotope compositions. Although for some Tulasnella and Sebacina clade B strains an ectomycorrhizal association has been documented (Bidartondo et al., 2003; Hynson et al., 2013b), there is good evidence that the Tulasnella and Sebacina clade B fungi found here in protocorms of the meadow-dwelling orchids S. parviflora and P. albida belong to the saprotrophic rhizoctonia.
In agreement with the first part of our hypothesis, initially mycoheterotrophic protocorms of the two orchid species associated with ectomycorrhizal fungi and fully mycoheterotrophic adults of N. nidus-avis were enriched in 13C and 15N in comparison to autotrophic reference plants, matching the typical 13C and 15N enrichment of fruit bodies of ectomycorrhizal fungi (Trudell et al., 2003) and thus confirming that ectomycorrhizal fungi serve as their exclusive C and N source. The position of E. helleborine adult 13C enrichment between that of fully mycoheterotrophic E. helleborine protocorms and that of autotrophic reference plants indicates a complementary photosynthetic C gain of this orchid at adulthood, and thus confirms the second part of our hypothesis. Simultaneously, E. helleborine adults and protocorms had similar 15N enrichments, which suggests exclusive N gain from the fungal source in both mycoheterotrophic protocorms and partially mycoheterotrophic adults. The even higher 15N enrichment found for the adults of the fully mycoheterotrophic N. nidus-avis in comparison to the protocorms of this species may be related to the observed increase in protocorm 15N enrichment with growing biomass.
The isotope composition of protocorms and adults of the two rhizoctonia-associated orchid species provides a less clear answer to the question of whether our hypothesis is supported. The C and N isotope compositions found here in the protocorms of rhizoctonia-associated orchids indicate that organic matter in saprotrophic fungi of the rhizoctonia group and derived from these fungi by orchid protocorms is significantly less enriched in 13C and 15N than organic matter in ectomycorrhizal fungi and gained by their associated orchid protocorms. This discovery has far-reaching consequences because it means that partially mycoheterotrophic nutrition among adults of the rhizoctonia-associated orchids cannot be identified unequivocally based on their C and/or N isotope composition alone. At least for N, this finding is in agreement with significantly lower 15N enrichments found for fully mycoheterotrophic orchids associated with saprotrophic wood-decomposer fungi than for fully mycoheterotrophic ectomycorrhizal fungi-associated orchids (Martos et al., 2009; Ogura-Tsujita et al., 2009). With our current knowledge we cannot unequivocally trace back the reason for the lower 13C and 15N enrichment of rhizoctonia-associated protocorms compared with ectomycorrhizal fungi-associated protocorms. The most likely reason for the different 13C and 15N enrichments is the different isotopic signal of the fungal C and N sources. Gebauer & Taylor (1999) have already shown that saprotrophic wood-decomposer fungi are depleted in 15N in comparison to ectomycorrhizal fungi, because bulk N from wood is depleted in 15N compared with humus bulk N. Unfortunately, for rhizoctonia fungi per se there are no data on isotope compositions available yet, because these fungi produce no fruit bodies easily accessible for isotope abundance analysis, and laboratory-cultured fungal tissue may not mirror their natural isotopic signal. We can only speculate that C and N compounds gained by saprotrophic rhizoctonia fungi from the soil and delivered to their associated orchid protocorms, and potentially to orchid adults, are less enriched in 13C and 15N than C and N compounds delivered by ectomycorrhizal fungi to their associated orchid protocorms and adults.
There are indications available from the literature on stable isotope abundance analyses for some rhizoctonia-associated orchid species that point towards gain of N from fungi apparently without significant C gain (Gebauer & Meyer, 2003; Liebel et al., 2010; Girlanda et al., 2011; Sommer et al., 2012). N gain from the fungal source without any C gain could only be based on exclusive fungus-to-plant transfer of inorganic N compounds (e.g. ammonium) or a cycling of organic N compounds, where an organic N compound (e.g. glutamate) gained from the fungus is deaminated in plant cells and then a C skeleton (e.g. 2-oxoglutarate) is transferred back to the fungus. However, current knowledge indicates that amino acids serve as preferred N compounds for a one-way fungus-to-plant transfer (Chalot et al., 2002; Smith & Read, 2008). Furthermore, it is known that fungal pelotons are degraded within orchid root cells (Burgeff, 1932), thus potentially leading also to fungus-to-plant transfer of organic N compounds. Gain of organic N from fungi, however, also means gain of fungal C, because organic N compounds are inevitably based on C skeletons. Thus, the occurrence of partial mycoheterotrophy has to be expected at least for some rhizoctonia-associated orchids. Hynson et al. (2013a) called this kind of organic N gain without apparent organic C gain ‘cryptic partial mycoheterotrophy’. Novel approaches required to elucidate this phenomenon might include laboratory tracer experiments (Cameron et al., 2008; Bougoure et al., 2010) and field studies of stable isotope natural abundance of elements other than C and N.
The significantly lower 13C and 15N enrichment in protocorms of rhizoctonia-associated orchids than in protocorms of orchids associated with ectomycorrhizal fungi also has consequences for the two-source isotopic mixing model approach. This model has been used as a proxy to calculate the proportional C and/or N gain from the fungal source by partially mycoheterotrophic plants (Gebauer & Meyer, 2003; Bidartondo et al., 2004; Tedersoo et al., 2007; Zimmer et al., 2007, 2008; Hynson et al., 2013a). The model requires as endpoints the isotopic composition of the two sources utilized by a partially mycoheterotrophic plant and assumes that these sources are mixed in a linear manner in the target plant. The mixing model furthermore assumes that the isotopic compositions of the photosynthetic C source and N gained by autotrophic plants as one of the two endpoints are represented by the isotopic composition of fully autotrophic reference plants living in close spatial proximity to the partially mycoheterotrophic target plant (= 0% C or N gain of fungal origin). The other endpoint is represented by fully mycoheterotrophic plants associated with ectomycorrhizal fungi (= 100% C or N gain of fungal origin). To improve this approach, specifically with respect to orchids associated with rhizoctonias, we suggest considering as the upper endpoint the isotopic composition of initially mycoheterotrophic protocorms instead of the mean values of fully mycoheterotrophic adults dependent on ectomycorrhizal fungi. As a consequence, more data on isotope signatures of protocorms of a wider spectrum of orchid species are now required.
The only quantitative data currently available on growth of orchid protocorms in nature report size and volume over time (McKendrick et al., 2000, 2002). Here we showed that the biomass of the investigated orchid protocorms after 1 yr in the soil reached a maximum of 0.5 mgDW and rose linearly to a maximum of 3.8 mgDW after 4 yr. This biomass production is a result of annual C gains from fungi in the range of only a few μmol and N gains as low as a few hundred nmol.
The unusually high total N concentrations found in the protocorms are striking. Total N concentrations of up to almost 5 mmol N gDW−1 (= 70 mg N gDW−1 = 7%) in orchid protocorms are far above those of nonorchid plant tissues. They are about twice as high as total N concentrations in leaves of legumes (Gebauer et al., 1988). As a result of their symbiosis with N2-fixing bacteria, legumes have already above-average total N concentrations in their tissues. However, total N concentrations in orchid protocorms match total N concentrations typically found in fungal tissue (Gebauer & Dietrich, 1993; Gebauer & Taylor, 1999). Fungal hyphae and pelotons in protocorms contribute (in unknown proportions) to protocorm biomass. This fungal contribution may explain to a certain extent the unusually high total N concentrations found in protocorms; nevertheless, the N compounds transferred from fungi to plant tissues in protocorms are expected to match also the N concentrations of typical fungal bulk tissues. It is already known for partially mycoheterotrophic orchids that the total N concentration in their tissue increases with increasing reliance on the fungal source (Stöckel et al., 2011). The chemical nature and function of the unusually high total N concentrations in orchid tissues are unknown. Metabolomic investigations are required to elucidate the major N compounds responsible for the high total N concentrations in orchid adults as well as in orchid protocorms.
The authors thank Isolde Baumann, Iris Schmiedinger and Christine Tiroch (BayCEER – Laboratory of Isotope Biogeochemistry, University of Bayreuth) for skillful technical assistance with stable isotope abundance measurements; Cesario Giotta, Rossana Segreto, Heiko Liebel and Rüdiger Otto for help with the field work; student research assistants and Nadja Schilloks for their help with the seed packets. Permissions granted by the respective authorities in Italy, Spain and the Czech Republic to collect tissues of protected orchids are gratefully acknowledged. Valuable comments by three anonymous reviewers on an earlier version of this manuscript are highly appreciated. The work was supported by the German Research Foundation (DFG, project GE 565/7-1) and the Czech Science Foundation (project no. 31-P505/10/0786).