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•Rhizanthella gardneri is a rare and fully subterranean orchid that is presumably obligately mycoheterotrophic. R. gardneri is thought to be linked via a common mycorrhizal fungus to co-occurring autotrophic shrubs, but there is no experimental evidence to support this supposition.
•We used compartmentalized microcosms to investigate the R. gardneri tripartite relationship. 13CO2 was applied to foliage of Melaleuca scalena plants and [13C-15N]glycine was fed to the common mycorrhizal fungus, and both sources traced to R. gardneri plants.
•In our microcosm trial, up to 5% of carbon (C) fed as 13CO2 to the autotrophic shrub was transferred to R. gardneri. R. gardneri also readily acquired soil C and nitrogen (N), where up to 6.2% of C and 22.5% of N fed as labelled glycine to soil was transferred via the fungus to R. gardneri after 240 h.
•Our study confirms that R. gardneri is mycoheterotrophic and acquires nutrients via mycorrhizal fungus connections from an ectomycorrhizal autotrophic shrub and directly from the soil via the same fungus. This connection with a specific fungus is key to explaining why R. gardneri occurs exclusively under certain Melaleuca species at a very limited number of sites in Western Australia.
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The rare underground orchid Rhizanthella gardneri is an obligately mycoheterotrophic orchid that lacks the ability to photosynthesize. Even during flowering, R. gardneri is entirely subterranean, a feature unique amongst known mycoheterotrophic orchids (Leake, 1994). R. gardneri also lacks roots and so has limited independent access to nutrient resources (Dixon & Pate, 1984; Dixon et al., 1990; Bougoure et al., 2008). Consequently, it has been presumed that R. gardneri, like many obligately mycoheterotrophic orchids, is dependent on a tripartite relationship with an orchid mycorrhizal fungus that also forms ectomycorrhizas with Melaleuca shrubs from the Melaleuca uncinata complex (Warcup, 1985; Mursidawati, 2004; Bougoure et al., 2008, 2009). Attempts at growing R. gardneri ex situ have only been successful in the presence of both its autotrophic and fungal partners (putatively a Ceratobasidium sp; Warcup, 1991; Bougoure et al., 2009). The orchid has also only ever been found growing within 1 m of the base of Melaleuca shrubs under natural conditions. While the relationships among these three species appear highly specialized, there has been no clear evidence of carbon (C) transfers to the orchid from the Melaleuca. Similarly, we know very little about how mycoheterotrophic orchids such as R. gardneri acquire nitrogen (N).
The autotrophic host is most likely to be the predominant C source in the tripartite relationships formed by mycoheterotrophic plants, particularly when those plants remain nongreen for their entire life (Leake, 1994, 2005). However, in the R. gardneri tripartite relationship, the autotrophic host may not be the only provider of C, as the mycorrhizal fungus linking the autotroph to the orchid belongs to the Ceratobasidiales, a group of known saprotrophs and pathogens (Warcup, 1991; Roberts, 1999; Bougoure et al., 2009). Little is known about the ability of an individual fungal species to simultaneously obtain C by both saprotrophic and mycorrhizal means (Hibbett et al., 2000; Wiemken & Boller, 2002). Indeed, there is generally clear taxonomic and functional separation between fungi considered saprophytic or mycorrhizal, and the latter usually fail to persist in ecosystems without a host plant (Molina et al., 1992; Brundrett, 2004). Consequently, the R. gardneri–Ceratobasidium–Melaleuca relationship may also access C and nutrients from soil organic matter.
Mycorrhizal fungi are known to transfer organic C and N as amino acids into their host plants, including green-leaved orchids (Cameron et al., 2006). However, the possibility of cotransport of C and N from fungus to orchid as well as to the autotrophic partner has not been demonstrated experimentally in any tripartite relationship. While mutualism in orchid mycorrhizas has been demonstrated in green-leaved orchids (Cameron et al., 2006), we would expect that the transfer of C from the fungal symbiont to R. gardneri would be unidirectional, as these plants never become autotrophic. Similarly, we would also expect that the orchid is dependent on the fungus for N acquisition from soil organic matter, consistent with most other orchid species (Leake, 2004).
Our objective was to determine to what extent a tripartite relationship with both a specialist fungus and autotrophic host facilitates C and N transfers to R. gardneri. We used isotopically labelled tracers, 13CO2 and double-labelled [13C-15N]glycine, to assess the direction of C and N transfers between autotrophic and mycoheterotrophic plants via fungal connections. We hypothesized that photosynthetically derived carbohydrates are transferred from Melaleuca scalena (a shrub species that R. gardneri occurs under) to R. gardneri via a connecting mycorrhizal fungus. We also hypothesized that the fungus that simultaneously forms mycorrhizas with R. gardneri and M. scalena would be able to access soil-derived nutrients saprophytically and subsequently transfer both C and N to R. gardneri and N to M. scalena.
Materials and Methods
Source and preparation of M. scalena seeds, R. gardneri seeds and mycorrhizal fungal inoculum
Rhizanthella gardneri (Rogers) is extremely rare and highly threatened, limited to two distinct populations of < 100 known individuals (Bougoure et al., 2008). Consequently, we were severely restricted in the amount of seed and thus replication that could be used for this experiment. Material for the microcosm experiment was collected in November 2005 from the northern population of R. gardneri near Babakin, c. 320 km east of Perth, in southwestern Australia (Bougoure et al., 2008). Seeds of both M. scalena and R. gardneri, used in the microcosm trials described later, were sterilized by immersion in 70% ethanol (v/v; 1 min), 5% H2O2 (v/v; 5 min) and 1% NaCl (v/v; 5 min) with thorough Milli-Q water rinses after each sterilant immersion.
The mycorrhizal fungus of R. gardneri used in all microcosms was isolate RGBAB1, putatively identified as a Ceratobasidium sp. belonging to the Ceratobasidiales (Bougoure et al., 2009). The isolate was cultured in liquid MMN medium (Marx, 1969) and made into an inoculum in the form of calcium alginate gel beads prepared using a modification of the method described by Da Silva Rodrigues et al. (1999). Briefly, under sterile conditions using sterile solutions, large clumps of actively growing fungal hyphae were rinsed in Milli-Q water, added to a sterile 500 ml bottle along with 4-mm-diameter glass beads and a small amount of Milli-Q water and shaken vigorously to break the hyphae into small fragments. The macerated fungal hyphae solution was mixed with 2% w/v sodium alginate (alginic acid sodium salt from brown algae; Sigma-Aldrich) solution, in a 1 : 1 ratio. This mycelial slurry was added to a 0.05 M CaCl2 solution drop-wise, using a 20 ml sterile syringe, and left for 45 min to set. Each drop formed a gel bead, c. 3 mm in diameter, containing a small amount of living fungal hyphae. The alginate beads were rinsed in sterile Milli-Q water and stored in sealed sterile vials at 4°C. A few beads were added immediately after synthesis, and also 1 wk before their use in pot trials, to fresh liquid MMN medium (Marx, 1969) to test if the inoculum was viable and not contaminated.
Microcosm experimental design
Our microcosm was a concentric tricompartment system that separated M. scalena roots from R. gardneri plants so that any connection between the two plant species was via a connecting mycorrhizal fungus (Fig. 1). Individual microcosms were created using large glass jars (1 l) wrapped in aluminium foil (Fig. 1). Each microcosm was divided into three compartments separated by a 42-mm-diameter polyvinyl chloride (PVC) pipe outside a 21-mm-diameter PVC pipe – both capped at one end, perforated with 8 mm holes and covered with 32 μm aperture nylon mesh (Sefar, Freibach Thal, Switzerland). All components were thoroughly cleaned before being autoclaved or gamma-irradiated at 25 kGy (Steritech, Melbourne, Victoria, Australia). Microcosms were then filled with loamy-sandy soil collected from the top 15 cm of soil from the Babakin site, beneath the canopy of a randomly selected M. scalena plant near to where R. gardneri plants had previously been recorded. Soil was dried at 60°C for 48 h, sieved < 2 mm, and then weighed into 1400 g lots for individual microcosms and gamma-irradiated at 25 kGy (Steritech).
Fifty-five replicate microcosms were constructed under sterile conditions. Half of the sterilized soil (700 g) was added to microcosms with internal dividers in position before five to 10 alginate beads of fungal inoculum were added to each of the three compartments. Two sealed 32-μm-aperture nylon pouches, each containing six to 10 seeds of R. gardneri, were added to the innermost compartment (Fig. 1) and the remaining 700 g of sterilized soil was added to fill all compartments of the microcosm. Between five and 10 M. scalena seeds were added to the outermost compartment and buried 3 mm below the soil surface. Microcosms were watered to c. 10% w/w moisture using c. 150 ml Milli-Q water then sealed using upside-down sunbags (Sigma-Aldrich), which provide a ventilated sterile growing environment for plants. Microcosms were placed in a controlled-environment room receiving 16 h of light per day with day : night temperatures of 18 : 23°C. These 55 microcosms were used for isotope labelling experiments. A further 24 microcosms were created for the purpose of observing R. gardneri germination so that experimental microcosms were not being continually disturbed.
Melaleuca scalena germination was observed after 7 d with all but one M. scalena plant per microcosm removed after 1 month. All microcosms were weighed fortnightly and watered to maintain 10% soil moisture content. Microcosms were created in December 2005 and checked for R. gardneri germination every few months in spare microcosms. In January 2007 the 55 large microcosms were checked for R. gardneri germination by carefully removing and opening nylon R. gardneri seed pouches. R. gardneri seeds and/or seedlings were carefully returned to their relevant microcosms and left for a month to allow for fungal reattachment or recolonization (Warcup, 1985).
Microcosm experiments to examine C and N transfers between R. gardneri, M. scalena and their connecting mycorrhizal fungus
Isotopically labelled tracers were added to microcosms in February 2007, 60 wk after establishment. We first sought to demonstrate the transfer of photosynthetically fixed C from the Melaleuca to R. gardneri via the fungus. Sunbags were removed from 22 (nine with and 12 without germinated R. gardneri) microcosms to expose M. scalena plants. A transparent 10 l Tedlar gasbag (SKC Inc., Eighty Four, PA, USA) was placed over M. scalena foliage and tightly sealed around the stem using tape and silicon to minimize gas escape. Bags were checked for major leaks before all air was removed. Labelled 13CO2 (180 ml 13C, 99%– Cambridge Isotope Laboratory, Andover, MA, USA) was injected at atmospheric pressure (c. 131 mg of 13C) into the gasbag via the tap-controlled hose connection using a 60 ml syringe. The gasbag remained on the M. scalena plant for 12 h during which time the microcosms were subject to the same daytime growing conditions described earlier. Microcosms were then taken out of the controlled-environment chamber and gasbags were removed from the plants so that no residual 13CO2 gas would become available to other replicates in the experiment. Sunbags were then replaced over microcosms, which were returned to the same controlled-environment chamber but kept separate from control plants so that any reabsorption of respired 13C-rich CO2 would be minimal.
A second labelling experiment assessed the saprotrophic potential of the mycorrhizal fungus and its subsequent capacity to transfer soil-derived organic C and N to either R. gardneri or M. scalena. Glycine was chosen to represent an organic C and N source because it is the smallest amino acid and it has been used in similar studies with success (Jennings, 1995; Cameron et al., 2006). Double-labelled glycine (5 ml of 2 mM containing 137.3 μg 15N and 235.4 μg 13C; U-13C2, 98%; 15N, 98%; Cambridge Isotope Laboratories) was added drop-wise to only the fungal (middle) compartment of 12 microcosms, four of which had germinated R. gardneri and eight of which had no germinated R. gardneri (Fig. 1). [13C-15N]glycine-labelled microcosms were separated from other microcosms but remained in the controlled-environment room, as described earlier, until sampled. Two control microcosms containing no mycorrhizal fungus, M. scalena or R. gardneri had glycine added as described earlier and were used to confirm that no movement of the added glycine occurred across compartment barriers (i.e. to confirm that M. scalena or R. gardneri could not independently access the added glycine).
Microcosms to which 13CO2 or [13C-15N]glycine had been added were sampled at 0, 6, 12, 24, 48, 96, 120 or 240 h postlabelling with three replicate microcosms sampled at each time. At each sampling, R. gardneri plants (when present) and a soil sample were carefully removed from the middle compartment of microcosms. Foliage was also removed from M. scalena plants and divided into three age categories (new, intermediate and old – easily determined as M. scalena foliage formed in consistent flushes). M. scalena root systems were then carefully removed from the soil and rinsed clean in water. Three replicate samples, each of between five and 10 ectomycorrhizal or nonectomycorrhizal root tips, were collected from single lateral roots of each M. scalena under a dissecting microscope. All samples were dried at 45°C for 5 d after which M. scalena leaf and soil samples were ball-milled (Retsch Ball Mill; Retsch, Haan, Germany) to a fine powder.
Isotopic analysis of δ15N and δ13C
All samples were analysed for total N and C content (%), and δ15N and δ13C isotope signatures, using an automated nitrogen carbon analyser-mass spectrometer consisting of a 20 : 20 mass spectrometer connected with an ANCA–NT system (Europa Scientific Ltd, Crewe, UK) in the West Australian biogeochemistry bentre at the University of Western Australia. All samples were standardized against a secondary reference of radish collegate (3.167% N, δ15N 5.71‰; 41.51% C, δ13C −28.61‰) that was in turn standardized against primary analytical standards (IAEA, Vienna, Austria). Accuracy was measured at 0.07‰ (SD), while precision was measured at 0.03‰ (SD), according to the stipulations for reporting analytical error in stable isotope analysis outlined by Jardine & Cunjak (2005).
Quantifying C movement from Melaleuca to Rhizanthella
The percentage of 13CO2 that was photosynthetically assimilated by M. scalena foliage and then transferred to R. gardneri plants was estimated for individual microcosms using the following equation:
where %CRGA is the percentage of C acquired by R. gardneri derived from the 13C taken up by M. scalena from the 12 h 13CO2 pulse; RGATMlab is the atom percentage of 13C in R. gardneri from labelled samples; RGATMcon is the atom percentage of 13C in R. gardneri samples at natural abundance; %CRG is the C percentage of R. gardneri sample; BMRG is the dry weight of R. gardneri (mg); MSATMlab is the atom percentage of 13C in M. scalena foliage samples directly after the 12 h pulse period; MSATMcon is the atom percentage of 13C in M. scalena samples at natural abundance; %CMS is the C percentage of M. scalena foliage samples; and BMMS is the dry weight of M. scalena foliage samples (mg).
13C and 15N enrichment of R. gardneri tissue and M. scalena foliage derived from double-labelled [13C-15N]glycine were calculated individually for each microcosm using the following equation:
where %ESAMA is the percentage of element (C or N) acquired by the sample (R. gardneri or M. scalena foliage) derived from added [13C-15N]glycine; SAMATMlab is the atom percentage of element (13C or 15N) in the sample (R. gardneri or M. scalena foliage) from labelled microcosms; SAMATMcon is the atom percentage of element (13C or 15N) in the sample (R. gardneri or M. scalena foliage) at natural abundance; %ESAM is the element percentage (C or N) of the sample (R. gardneri or M. scalena foliage); BMSAM is the dry weight (mg) of the sample (R. gardneri or M. scalena foliage); and Eadd is the amount (mg) of element (13C or 15N) added to microcosms as [13C-15N]glycine
Differences in 13C and 15N isotopic composition between sample types and sampling times were analysed using the software Primer (version 6, Clarke & Gorley, 2006) with PERMANOVA (Anderson et al., 2008). Permutational analysis of variance (PERMANOVA) was performed on square-root-transformed data to provide a robust comparison of treatments that was unaffected by data heterogeneity or variations in replication of individual treatments.
Formation of M. scalena ectomycorrhizas and R. gardneri germination
Of the 55 large microcosms created, ectomycorrhiza formation between the M. scalena seedling and fungal inoculant was substantial in 35 (> 75% root tips ectomycorrhizal), weaker in 16 (25–75% root tips ectomycorrhizal) and negligible in four. There was no obvious greater growth of M. scalena in microcosms where mycorrhizal formation was most substantial. Growth of M. scalena individuals was very slow and plants were only c. 28 cm in height after 60 wk in the microcosms. There was no evidence of contamination by foreign ectomycorrhizal fungi in any of the microcosms in sunbags even after 60 wk growth.
Rhizanthella gardneri seeds did not germinate until c. 50 wk after microcosm establishment. When the large microcosms were examined 56 wk post-creation, 38% had signs of R. gardneri growth at varying stages, ranging from imbibed seeds with broken seed coats to developing protocorms of 15 mm diameter (Fig. 1). When germination was observed, often all R. gardneri seeds in that microcosm had germinated and they tended to be associated with extensive ectomycorrhiza formation.
13C labelling of the R. gardneri tripartite system derived from 13CO2 pulsing
Analysis of M. scalena leaves before and after pulsing with 13CO2 showed that that the 13C assimilation in the leaves was highest directly after the 12 h pulse and gradually reduced over the time course (Fig. 2). Differences in 13C enrichment between sample types and over sample times were significant (P =0.001). Enrichment was greatest in new foliage compared with old foliage, which is consistent with incorporation into metabolically active leaves. Differences in the timing of maximum isotopic enrichment between sample types, clearly visible as peaks in Fig. 2, are also confirmed by a significant sample type × time interaction in the PERMANOVA analysis (P =0.001).
In microcosms not containing R. gardneri plants, foliage of M. scalena was more enriched over the time course compared to microcosms with R. gardneri plants even though 13C abundance of the samples taken directly after the 13CO2 pulse were similar (δ13C 3092 ± 288‰, n = 6). M. scalena roots showed 13C enrichment of both ectomycorrhizal and nonmycorrhizal roots within 48 h after the addition of 13CO2 (Fig. 2). Enrichment was greater in nonmycorrhizal roots than in mycorrhizal roots in microcosms with or without R. gardneri present. There was slight 13C enrichment (c. 2‰) of soil in the fungus-only compartment of the microcosms at 96 h after the 13CO2 pulse (Fig. 2). Specific data for all microcosms are presented in the Supporting Information, Table S1.
15N and 13C labelling of the R. gardneri tripartite system derived from added [13C-15N]glycine
15N enrichment of R. gardneri tissue only became evident 120 h after the labelled glycine was added to the microcosms (Fig. 3). 15N enrichment of ectomycorrhizal and nonmycorrhizal roots was variable among microcosm replicates with and without R. gardneri present (Fig. 3). Generally, ectomycorrhizal roots showed greater 15N enrichment in the earlier sampling times, 24–48 h after addition of labelled glycine compared with nonmycorrhizal roots. Foliage of M. scalena was enriched in 15N at 120 h after labelled glycine addition in microcosms without R. gardneri and 240 h after labelled glycine addition in microcosms with R. gardneri (Fig. 3). Enrichment was most obvious in the youngest foliage in both cases. Soil from the fungus-only compartment, where the labelled glycine was added, remained 15N-enriched through the entire time course in all microcosms. However, soils (fungal only compartment) from microcosms containing R. gardneri showed a dramatic decrease in 15N enrichment at 240 h after labelling compared with earlier time points. This depletion was not observed in the microcosms without the orchid (see Table S3).
13C enrichment of R. gardneri tissue became obvious c. 120 h after addition of [13C-15N]glycine (Fig. 3). By contrast, ectomycorrhizal roots of M. scalena showed little variation in 13C across the time course, in microcosms with or without R. gardneri. However, nonmycorrhizal roots of M. scalena became slightly 13C-enriched over the sampling time course, particularly in microcosms without R. gardneri (Fig. 3). M. scalena foliage was slightly but variably enriched in 13C for most samples, especially in the youngest foliage (Fig. 3). Soil from the fungus-only compartment (where [13C-15N]glycine was added) remained 13C-enriched through the entire time course in all microcosms, although this decreased with time (Fig. 3). Specific data for all microcosms are presented in Table S2.
Quantification of C and N transfer to R. gardneri
Immediately after the 12 h pulse, M. scalena foliage dry weight had a 13C excess of 16.1 μg mg−1 (SE 1.3, n =3). In one microcosm sampled 120 h after 13CO2 labelling, 5.03% (or 2.6 μg mg−1R. gardneri dry weight) of the added 13C was present in total R. gardneri biomass of that microcosm. In two other replicates sampled at the same time, 0.22% (or 1.65 μg mg−1R. gardneri dry weight) and 0.28% (or 0.2 μg mg−1R. gardneri dry weight) of the added 13C was present in total R. gardneri biomass in those microcosms.
For 13C derived from [13C-15N]glycine, up to 10.8% of the 13C was present in R. gardneri tissue (total of all plants in individual microcosm sampled 240 h after labelling). Two replicate microcosms 120 h after labelled glycine addition had 1.13 and 1.39% of added 13C present in R. gardneri tissue (total of all plants in the individual microcosm). Transfer of 15N in the same three microcosms showed that 8.5% of the 15N added in the glycine was transferred to R. gardneri protocorms in the microcosm sampled 240 h after labelling, and in the two microcosms sampled 120 h after glycine labelling, 3.48 and 4.13% of the 15N were transferred to R. gardneri protocorms (results of all R. gardneri plants in the individual microcosms combined). However, when excess 13C and 15N derived from the dual-labelled glycine was calculated in relation to the amount of R. gardneri biomass in each microcosm, 13C and 15N transferred in each of the same three microcosms were more uniform, with c. 0.2 μg 13C and 0.3–1 μg 15N per mg R. gardneri dry weight biomass. Transfer of 15N to M. scalena in two replicate microcosms sampled 120 h after double-labelled glycine addition was equivalent to 0.001 and 0.002%, respectively, of the excess 15N added to individual microcosms. The microcosm sampled 240 h after glycine addition showed a significant increase in excess 15N content of the total foliage biomass of M. scalena, which represented 3.26% of the 15N added.
Our study clearly demonstrates for the first time that R. gardneri can obtain nutrients from an associating mycorrhizal fungus, which in turn derives C from both an autotrophic shrub (via ectomycorrhizas) and, interestingly, also from the soil organic substrates via saprophytic activity. Furthermore, we have confirmed a mutualistic (i.e. nutrient-sharing) mycorrhizal relationship between M. scalena and a species of Ceratobasidium, a fungal genus not generally implicated in ectomycorrhizal formation (Roberts, 1999; Bougoure et al., 2009). When autotrophic M. scalena plants were supplied with enriched CO2, 13C enrichment of R. gardneri tissue, which contained fungal pelotons, was evident at 96 h post-labelling, with increased concentrations observed 24 h later. The number of microcosms where R. gardneri germinated limited the duration of our experiment and it is not clear if further enrichment would have occurred after the 120 h final time point of the experiment. Thus, transfer into the orchid tissue may take longer than the duration of the time course in this study. It is likely that the observed 13C enrichment in R. gardneri was still confined to the fungal peloton hyphae within orchid cortical cells. While portions of plant with and without peloton presence could not be analysed separately because of the limited number and size of R. gardneri plants used in this study, we might expect that some fungus–orchid transfer would have occurred during the sampling period. However, little is known of the specific nutrient transfer mechanisms across orchid mycorrhizas (Rasmussen, 2002; Smith & Read, 2008), specifically whether C is transferred from fungus to orchid before or only as a result of peloton lysis.
Our results greatly strengthen the hypothesis that some mycoheterotrophic plants rely heavily on their mycorrhizal fungi’s ability to simultaneously form ectomycorrhizas with autotrophic hosts (Leake, 1994, 2004, 2005; McKendrick et al., 2000a,b). Although Bougoure et al. (2009) demonstrated the ability of R. gardneri’s mycorrhizal fungus to form both orchid mycorrhizas and ectomycorrhizas, until now there was no direct evidence to suggest that nutrients could be transferred from the autotrophic shrub to the fungus and then to the orchid. We found R. gardneri acquired 5.03% of the 13C added as 13CO2 to the Melaleuca in one microcosm (after 120 h), but in two other replicates only 0.2% of the added 13C was transferred to R. gardneri. Similar studies by McKendrick et al. (2000b) also found that the mycoheterotrophic orchid, Corallorhiza trifida, acquired c. 0.3% of enriched 14C fixed by an autotrophic shrub from a mycorrhizal fungus that linked the two plants. While further investigation would be required to determine whether R. gardneri would acquire more 13C given more time after labelling, it is likely that biomass, particularly peloton biomass in orchid tissue, will determine how much C is allocated to the orchid. Our results for both R. gardneri, together with those previously published for C. trifida (McKendrick et al., 2000b), demonstrate that these mycoheterotrophic orchids may represent < 6% C drain on their autotrophic host. While C transfers to the orchid may be slightly greater if we assume that the orchids respire some of the transferred C before analysis, it is still unlikely that > 10% of the C fixed by the host shrub would be transferred to the orchid. When extrapolating microcosm data to the natural situation, we would expect that R. gardneri’s C drain on its autotrophic host would be even less given the large ratio of shrub to orchid biomass (these shrubs normally grow 2–3 m tall). We were unable to accurately quantify C transfer from M. scalena to its associated ectomycorrhizal fungus because of difficulties isolating fungi from other components of the microcosms. However, we would expect an order of magnitude difference between C allocated from autotrophic M. scalena to the fungus, compared with that ending up in R. gardneri, based on suggestions by Hobbie (2006) that C allocation to the ectomycorrhiza-forming fungus can amount to up to a quarter of total C taken up by its autotrophic plant host (depending on nutrient availability and host tree biomass). Therefore, R. gardneri’s C drain on its mycorrhizal fungus is quite significant if we assume that much of the C acquired via ectomycorrhizas is respired, as has been suggested for other ectomycorrhizal systems (Colpaert et al., 1996; Högberg et al., 2001).
Saprobic capacity of the fungus and contribution to C and N transfer to R. gardneri
By labelling with [13C-15N]glycine, we showed that R. gardneri is able to receive fungal C transfers not only from the Melaleuca shrub but also directly from the soil. Fungi have the ability to absorb most amino acids intact, including glycine (Jennings, 1995; Taylor et al., 2004), and the acquisition of the 13C derived from the added labelled glycine may thus be a by-product of the fungi’s N acquisition strategy. However, the ability of the R. gardneri mycorrhizal fungus to compete with other soil microorganisms (in situ) for access to soil C, and therefore the significance of this mechanism for the orchid, remains unresolved. While ectomycorrhizal fungi in boreal habitats compete poorly with saprotrophic fungi for access to more readily degradable soil C sources (Lindahl et al., 2007), the Ceratobasidiales fungus that forms mycorrhizas with R. gardneri cannot be assumed to behave in the same way, as all other investigated species in the group are parasites or saprotrophs (Roberts, 1999). Further research with different C sources that are characteristic of soil organic matter in its natural environment and which are likely predominantly derived from Melaleuca litter (Bougoure et al., 2008) is required to elucidate the physiological potential of the Ceratobasidiales fungus.
Our results suggest that glycine was not transferred within the fungus as an intact amino acid and that some portion of the excess 13C was being lost, probably via fungal respiration. In microcosms fed with double-labelled glycine, R. gardneri tissue had higher 13C : 15N ratios than M. scalena foliage but both were less than that of the [13C-15N]glycine. For example, R. gardneri acquired only one atom of 13C for every five atoms of 15N derived from glycine acquired by its mycorrhizal fungus (sampled 120 and 240 h after labelled glycine added). Cameron et al. (2008) recently reported that up to 15% of C added to microcosms with a green-leaved orchid (Goodyera repens) and mycorrhizal fungus (C. cornigerum) was lost via fungal respiration. If we assume consistencies in physiological functioning across mycorrhizal species, fungal respiration in our system may account for the lower C : N, derived from added [13C-15N]glycine, in R. gardneri tissue. Furthermore, respired 13C from the fungus may contribute to the slight 13C enrichment of M. scalena foliage in some of the dual-labelled glycine microcosms.
It was expected that the nonphotosynthetic R. gardneri would have a reduced demand for N, and therefore low N concentration, because most N found in other plants is associated with chlorophyll and photosynthetic processes (Marschner, 1995; Lambers et al., 2008). However, this was not the case; significant amounts of 15N were transferred to R. gardneri and its tissue had higher N concentration than M. scalena foliage. Gebauer & Meyer (2003) also found high N concentrations (2.13–2.87 mmol g−1) in a number of mycoheterotrophic orchids compared with surrounding autotrophic plants (1.44–1.77 mmol g−1). The high N concentrations in mycoheterotrophic orchids suggest that nutrient transfer from the orchid mycorrhiza may predominantly occur via peloton lysis and not across active mycorrhizal interfaces (Rasmussen, 2002; Peterson & Massicotte, 2004). Presumably the fungus has high concentrations of N-rich proteins and chitin, which the orchid can acquire after peloton lysis. The role of fungal peloton lysis in orchid mycorrhizal nutrient transfer is further supported by the similarity of isotopic natural abundance signatures, especially for N and C, between mycoheterotrophic orchids and their fungal partners (Hobbie et al., 2001; Gebauer & Meyer, 2003; Trudell et al., 2003), suggesting bulk flow of nutrients from fungus to orchid. However, this process has yet to be demonstrated experimentally.
The results presented here show that R. gardneri can obtain both C and N from its mycorrhizal fungus, which in turn can obtain N from a simple amino acid source and C from both amino acid applied to soil and a second autotrophic host via ectomycorrhizas. Furthermore, insights as to the magnitude of C and N transferred within the R. gardneri tripartite system have been achieved. The tripartite system is not simply two plants connected by a ‘pipeline’ (fungus), and results in this study demonstrate that the organisms involved in this relationship should be considered as parts of two distinct systems: a fungus obtaining a steady consistent carbohydrate source and the orchid exploiting or parasitizing a mycorrhizal fungus for all its nutritional needs. This relationship is likely to be critical to the ability of the fungus to obtain ample carbohydrates and nutrients to promote R. gardneri germination and continued growth. The importance of the tripartite relationship was inadvertently demonstrated in this study given that we found R. gardneri germination only occurred in microcosms with a high degree of Melaleuca ectomycorrhiza formation. Variable Melaleuca mycorrhization amongst replicate microcosms was unexpected but suggests that the R. gardneri–fungus–Melaleuca relationship is sensitive to subtle environmental or genetic differences. Understanding fundamental aspects of how R. gardneri and its mycorrhizal fungus have adapted to their environment, and specifically how they utilize available nutrients are key first steps in developing conservation and restoration strategies for this critically endangered orchid (see Bougoure et al., 2008). In the future, conservation efforts should focus on all three organisms, including how they interact with each other and their environment.
Financial assistance for this project was received from the Australian Research Council (ARC Linkage Project 0454276). We are also grateful to King’s Park and Botanic Gardens, WA, Australia for supplying some of the R. gardneri seeds used in this study. We are grateful to two anonymous reviewers and Prof. Ian Alexander for their comments on the manuscript.