Annals of Botany 116: 423–435, 2015
doi:10.1093/aob/mcv085, available online at www.aob.oxfordjournals.org
PART OF A HIGHLIGHT ON ORCHID BIOLOGY
The importance of associations with saprotrophic non-Rhizoctonia fungi among
fully mycoheterotrophic orchids is currently under-estimated: novel
evidence from sub-tropical Asia
Yung-I Lee1,2, Chih-Kai Yang3,4 and Gerhard Gebauer5,*
1
Biology Department, National Museum of Natural Science, No 1, Kuan-Chien Rd, Taichung, Taiwan, 2Department of Life
Sciences, National Chung Hsing University, Taichung 40227, Taiwan, 3The Experimental Forest, College of Bio-Resources
and Agriculture, National Taiwan University, 12 Chienshan Rd., Sec. 1, Chushan Township, Nantou 55750, Taiwan,
4
Department of Life Science, National Taiwan Normal University, 88 Tingchow Rd., Sec. 4, Taipei 11677, Taiwan
and 5Laboratory of Isotope Biogeochemistry, Bayreuth Center of Ecology and Environmental
Research (BayCEER), University of Bayreuth, D-95440 Bayreuth, Germany
* For correspondence. E-mail [email protected]
Received: 18 December 2014 Returned for revision: 4 February 2015 Accepted: 27 April 2015 Published electronically: 25 June 2015
Background and Aims Most fully mycoheterotrophic (MH) orchids investigated to date are mycorrhizal with
fungi that simultaneously form ectomycorrhizas with forest trees. Only a few MH orchids are currently known to be
mycorrhizal with saprotrophic, mostly wood-decomposing, fungi instead of ectomycorrhizal fungi. This study provides evidence that the importance of associations between MH orchids and saprotrophic non-Rhizoctonia fungi is
currently under-estimated.
Methods Using microscopic techniques and molecular approaches, mycorrhizal fungi were localized and identified for seven MH orchid species from four genera and two subfamilies, Vanilloideae and Epidendroideae, growing
in four humid and warm sub-tropical forests in Taiwan. Carbon and nitrogen stable isotope natural abundances of
MH orchids and autotrophic reference plants were used in order to elucidate the nutritional resources utilized by the
orchids.
Key Results Six out of the seven MH orchid species were mycorrhizal with either wood- or litter-decaying saprotrophic fungi. Only one orchid species was associated with ectomycorrhizal fungi. Stable isotope abundance
patterns showed significant distinctions between orchids mycorrhizal with the three groups of fungal hosts.
Conclusions Mycoheterotrophic orchids utilizing saprotrophic non-Rhizoctonia fungi as a carbon and nutrient
source are clearly more frequent than hitherto assumed. On the basis of this kind of nutrition, orchids can thrive in
deeply shaded, light-limiting forest understoreys even without support from ectomycorrhizal fungi. Sub-tropical
East Asia appears to be a hotspot for orchids mycorrhizal with saprotrophic non-Rhizoctonia fungi.
Key words: Orchids, Orchidaceae, mycoheterotrophy, mycorrhiza, ectomycorrhiza, Vanilloideae, Epidendroideae,
Gastrodia, stable isotopes, carbon, nitrogen, saprotrophic fungi.
INTRODUCTION
In nature, orchids are known to begin their life cycle as mycoheterotrophs (Rasmussen, 1995; Leake, 2004). Because of the
rudimentary embryo and the lack of endosperm in seeds, the
germination of orchid seeds is dependent on the formation of a
mycorrhizal association, which supplies young seedlings, i.e.
protocorms, with all carbon (C) and mineral nutrients until the
seedlings develop green leaves and become putatively autotrophic (Leake, 1994; Merckx, 2013). Mycorrhizal partners of the
majority of these adult green orchids are widely distributed
fungi of the polyphyletic Rhizoctonia group, including
Tulasnella, Ceratobasidium, Thanatephorus and Sebacina clade
B (Dearnaley et al., 2012). In contrast to the putatively autotrophic nutritional mode of chlorophyllous orchids, a few orchids
remain achlorophyllous and depend on their mycorrhizal partners for C and mineral nutrient supplies throughout their entire
life cycle. These achlorophyllous orchids are known as mycoheterotrophic (MH) plants (Leake, 1994; Merckx, 2013).
In temperate forests, MH orchids usually associate with narrow clades of ectomycorrhizal (ECM) fungi and obtain photosynthates from neighbouring trees through underground
mycorrhizal networks (Taylor and Bruns, 1997; Hynson et al.,
2013). In tropical and sub-tropical forests, a few MH orchids
have been reported to associate with saprotrophic (SAP) nonRhizoctonia fungi and obtain nutrients through the ability of the
fungi to cause wood or litter decay. For example, Gastrodia
spp. have been shown to associate with the litter- and/or wooddecomposing fungi Armillaria, Mycena, Resinicium and
Campanella or Marasmius (Kusano, 1911; Kikuchi et al.,
2008; Martos et al., 2009; Ogura-Tsujita et al., 2009;
Dearnaley and Bougoure 2010), Epipogium roseum associates
with a litter-decomposing species of Coprinaceae in culture
conditions (Yamato et al., 2005), Wullschlaegelia aphylla associates with litter-decaying species of Gymnopus and Mycena
(Martos et al., 2009), Eulophia zollingeri associates with another litter decomposer, Psathyrella cf. candolleana (OguraTsujita and Yukawa 2008), and Erythrorchis spp. associate
C The Author 2015. Published by Oxford University Press on behalf of the Annals of Botany Company.
V
All rights reserved. For Permissions, please email: [email protected]
424
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
with a wide range of wood-rotting fungi of Hymenochaetaceae
and Polyporaceae (Umata, 1995, 1997a; Dearnaley, 2007). In
addition, some MH orchids in tropical and sub-tropical forests
also associate with ECM fungi, but lack mycorrhizal specificity
as in temperate forests (Roy et al., 2009). A recent molecular
approach indicates that Lecanorchis associates with diverse
ECM fungi, e.g. Lactarius, Russula, Atheliaceae and Sebacina
clade A (Okayama et al., 2012).
The analysis of 13C and 15N (nitrogen) isotope natural abundances has been extensively used to elucidate the nutritional resources utilized by organisms in ecosystems. Along food chains
most organisms have isotope values similar to their food resources (Fry, 2006). ECM fungi are significantly enriched in
13
C and 15N as compared with autotrophic plants (Gebauer and
Dietrich, 1993; Gleixner et al., 1993), and consequently, MH
plants associated with ECM fungi have isotope signatures close
to those of ECM fungi (Gebauer and Meyer, 2003; Trudell
et al., 2003). As compared with ECM fungi, wood-decaying
SAP fungi are even more enriched in 13C, but less enriched in
15
N, and, therefore, MH plants associated with wood-decaying
SAP fungi should have isotope signatures similar to this group
of SAP fungi. Unfortunately, little information about the isotopic composition of MH plants associated with non-Rhizoctonia
SAP fungi is available to date (Ogura-Tsujita et al., 2009;
Martos et al., 2009; Dearnaley and Bougoure, 2010; Sommer
et al., 2012; Hynson et al., 2013). Furthermore, the currently
available knowledge provides no clue about whether MH plants
associated with wood-decaying or litter-decaying fungi are different in terms of their isotopic composition. Such a distinction
should be expected from the different isotopic composition of
the substrates on which these fungi live (Gebauer and Schulze,
1991; Cernusak et al., 2009).
A great diversity of MH orchids (>120 species) occurs in
tropical and sub-tropical Asia. In Taiwan, among the approx.
400 native orchids, >50 fully MH orchids in 15 genera have
been recorded (Su, 2000). The Xitou Experimental Forest located in central Taiwan has long humid seasons with warm
temperatures that obviously favour the growth of MH orchids.
According to the report by Yang et al. (2010), nine fully MH
orchids, including representatives of the genera Cyrtosia,
Galeola and Lecanorchis (subfamily Vanilloideae) and
Gastrodia and Epipogium (subfamily Epidendroideae) occur in
this misty forest with abundant litter and dead wood. Among
these MH orchids in the Xitou Experimental Forest, three
Gastrodia spp., G. appendiculata, G. fontinalis and G. nantoensis, occur sympatrically in a bamboo forest, whereas in another
bamboo forest, two vanilloid orchids, C. javanica and L. thalassica, grow sympatrically.
The richness of MH orchids in the sub-tropical forests in central Taiwan allows us to test the following questions. (1) What
are their mycorrhizal partners? Current knowledge about the
identity of mycorrhizal fungi in the vanilloid genera Cyrtosia
and Erythrorchis, close relatives of Galeola, are primarily
based on only in vitro isolations (Hamada, 1939; Umata, 1995,
1997a). Here we identify the fungal associates of seven MH orchids using molecular methods. (2) Gastrodia spp. occur
mainly in Asia, Africa and Australia. How much diversity is
there in mycorrhizal partners over the range of Gastrodia spp.?
We compare the fungal composition in mycorrhizas of sympatric and allopatric species. (3) Although the mycorrhizal partners
of Cyrtosia, Galeola and Lecanorchis of subfamily
Vanilloideae have already been investigated, their nutritional
resources are still not clear. Cyrtosia and Galeola appear to associate with SAP fungi, whereas Lecanorchis presumably associates with ECM fungi. However, Lyophyllum shimeji, an ECM
fungus, could stimulate seed germination in vitro of
Erythrorchis, a close relative genus of Galeola (Umata, 1997b),
suggesting the possible recruitment of an ECM mycorrhizal
partner in the natural environment. In this study we analyse for
the first time the C and N stable isotope abundances of three
vanilloid orchids and four Gastrodia spp. to reveal their nutritional resources, either ECM fungi or wood-decaying or litterdecaying non-Rhizoctonia SAP fungi.
MATERIALS AND METHODS
Sample collection and locations
Specimens of seven fully MH orchids (Figs 1 and 2) were sampled from four sites in Central Taiwan from 2011 to 2012
(Supplementary Data Table S1). The four sites are located
approx. 500–3000 m from each other in the Xitou Experimental
Forest (College of Bio-resources and Agriculture, National
Taiwan University), Nantou County, Taiwan at 1000 m above
sea level. The climate is sub-tropically moist, with a mean annual temperature of 166 C and a mean annual precipitation of
2600 mm. Site A (23 690 2900 N, 120 790 1200 E) consists of a
broadleaf forest on organic soil (pH 37) dominated by Phoebe
formosana and Machilus japonica (Lauraceae) with some
understorey plants (see Table S2). The target plant at this site
was the MH orchid Galeola falconeri. Site B (23 400 4400 N,
120 450 5300 E) consists of a dense bamboo (Phyllostachys
edulis) forest mixed with Cryptomeria japonica trees on organic soil (pH 44) with only few understorey plants (see Table
S2). The MH orchids Cyrtosia javanica and Lecanorchis thalassica grow sympatrically at this site. Site C (23 400 2600 N,
120 470 4500 E) consists of a coniferous forest on organic soil
(pH 40) dominated by Cryptomeria japonica with some understorey plants (see Table S2). The target orchid at this site was
the MH Gastrodia flabilabella. Site D (23 400 4100 N,
120 470 4300 E) consists of a dense bamboo (Phyllostachys
edulis) forest on organic soil (pH 44) with only few understorey plants (see Table S2). The MH orchids Gastrodia appendiculata, G. fontinalis and G. nantoensis co-occur at this site.
Voucher specimens of G. falconeri (Yung-I Lee 201225),
C. javanica (Yung-I Lee 201223), L. thalassica (Yung-I Lee
201224), G. appendiculata (Yung-I Lee 201122), G. fontinalis.
(Yung-I Lee 201217), G. nantoensis (Yung-I Lee 201121) and
G. flabilabella (Yung-I Lee 201117) have been deposited in the
herbarium of the National Museum of Nature and Science,
Taichung, Taiwan. Light climate data of the four sites were
measured with a LM-8000 Lux meter (Lutron Electronic
Enterprise Co., Ltd, Taipei, Taiwan) at 20 cm from ground level
at three different points in each site. Site A received a mean of
3670 lux (¼ 6 %), site B a mean of 980 lux (¼ 2 %), site C a
mean of 3050 lux (¼ 5 %) and site D a mean of 950 lux (¼
2 %), whereas outside of the forests at the same time a mean of
57 800 lux (¼ 100 %) was measured.
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
425
B
A
C
FIG. 1. Flower morphology of vanilloid species. (A) Galeola falconeri, scale bar ¼ 5 cm; (B) Cyrtosia javanica, scale bar ¼ 1 cm; (C) Lecanorchis thalassica, scale
bar ¼ 1 cm.
Microscopy
Mycorrhizal roots were collected and fixed in 25 % glutaraldehyde and 16 % paraformaldehyde buffered with 005 M
phosphate buffer, overnight at 4 C. After fixation, the samples
were dehydrated using an ethanol series, and embedded in
Technovit 7100 (Kulzer & Co., Wehrheim, Germany). Sections
of 3 mm thickness were obtained using Ralph knives on a
Reichert-Jung 2040 Autocut rotary microtome. Sections were
stained with 005 % (w/v) toluidine blue O (TBO) in benzoate
buffer for general histology (Yeung, 1984). The sections were
examined and the images were captured using a digital camera
attached to a microscope (Axioskop 2, Carl Zeiss AG, Jena,
Germany).
Molecular identification of mycorrhizal fungi
After checking for fungal colonization by free-hand sections
under the microscope, mycorrhizal roots were washed in water
and kept at –80 C until use. DNA was extracted from each
sample by using a DNeasy Plant Mini Kit (Qiagen, Hilden,
Germany). The internal transcribed spacer (ITS) region of the
fungal nuclear rRNA gene was amplified with the primer combinations ITS1F/ITS4 or ITS1F/ITS4B (White et al., 1990;
Gardes and Bruns, 1993). The large subunit (LSU) nuclear ribosomal DNA (nrDNA) sequences were amplified using primer
combinations LR0R/LR5 (Moncalvo et al., 2000) or LR0R/
LR3 (Vilgalys and Hester, 1990). PCR amplification and sequencing were carried out as described by Ogura-Tsujita et al.
(2009). PCR products that were difficult to sequence directly
were cloned using the pGEM-T Vector System II (Promega,
Madison,
WI,
USA).
Sequences
were
identified
(Supplementary Data Table S3) using a BLAST search against
the NCBI sequence database (National Center for
Biotechnology Information, GenBank) to find the closest sequence matches in the database. For phylogenetic analysis,
LSU marasmioid sequences from GenBank were added to the
analysis by referring to Moncalvo et al. (2000, 2002), Wilson
and Desjardin (2005), Matheny et al. (2006), Martos et al.
(2009) and Ogura-Tsujita et al. (2009), and sequences of
Cyphella digitalis, Nia vibrissa and Henningsomyces candidus
were used as outgroup taxa. LSU sequences of Polyporales
from GenBank were added to the analysis by referring to Justo
and Hibbett (2011) and Binder et al. (2013), and sequences of
Coltriciella oblectabilis was used as outgroup taxa. ITS sequences of Russula from GenBank were added to the analysis
by referring to Okayama et al. (2012), and sequences of
Arcangeliella camphorata and Lactarius quietus were used as
outgroup taxa. DNA sequences were aligned using
CLUSTALX (Thompson et al., 1997), followed by manual adjustment. Phylogenetic relationships were analysed by a modelbased Bayesian approach using MrBayes 32.1 (Ronquist and
Huelsenbeck, 2003). The ‘best-fit’ model of evolution was selected under the Akaike information criterion test (Akaike,
1974) as implemented in MrModeltest 22 (Nylander, 2004).
The general time reversal plus invariant rates and a gamma
distribution (GTR þ I þ C) was selected for the analyses. Two
separate runs of four Monte Carlo Markov chains (MCMCs;
Yang and Rannala, 1997) were performed for 10 000 000 generations until the mean deviation of split frequency dropped below 001, and a tree was sampled every 1000th generation.
Trees from the first 25 % of generations were discarded using
the ‘burn-in’ command, and the remaining trees were used to
calculate a 50 % majority-rule consensus topology and to determine the posterior probability (PP) for individual branches.
The alignment data sets were further analysed by maximum
426
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
A
C
B
D
FIG. 2. Flower morphology of Gastrodia species. (A) Gastrodia appendiculata, scale bar ¼ 1 cm; (B) Gastrodia fontinalis, scale bar ¼ 1 cm; (C) Gastrodia flabilabella, scale bar ¼ 1 cm; (D) Gastrodia nantoensis, scale bar ¼ 1 cm.
parsimony (MP) using PAUP* version 40b10 (Swofford,
2002). Support for groups was evaluated using the bootstrap
method (Felsenstein, 1985) with 1000 replicates. The trees
obtained in these analyses were drawn with the TreeGraph 2
software (Stover and Muller, 2010).
Stable isotope abundance analysis
Five 1 m2 plots were selected at each site; each plot included
fully MH orchids and four to five autotrophic reference plant
species. Flower stalks of seven fully MH orchids, leaves of four
to five autotrophic reference plants and soil samples from the
organic layer were taken from each of the five plots at each
site. The reference plants collected at the respective sites are
listed in Supplementary Data Table S2. In addition, on site B,
fruit bodies of a litter-decaying SAP fungus (Marasmius sp.)
were found and collected in five replicates.
Samples were dried at 105 C, ground to a fine powder and
stored in a desiccator with silica gel until analysed. Relative N
and C isotope abundances of the samples were measured using
a dual-element analysis mode with an elemental analyser coupled to a continuous flow isotope ratio mass spectrometer as described in Bidartondo et al. (2004). Measured abundances are
denoted as d values that were calculated according to the given
equation d15N or d13C ¼ (Rsample/Rstandard 1) 1000 [%],
where Rsample and Rstandard are the ratios of heavy isotope to
light isotope of the samples and the respective standard.
Standard gases (N2 and CO2) were calibrated with respect to international standards by using the reference substances N1 and
N2 for N isotopes and ANU sucrose, NBS 18 and NBS 19 for
C isotopes, provided by the International Atomic Energy
Agency (Vienna, Austria). Reproducibility and accuracy of the
isotope abundance measurements were routinely controlled by
measuring the test substance acetanilide (Gebauer and Schulze,
1991). At least six test substances with varying sample weight
were routinely analysed in each batch of 50 samples. The maximum variation of d13C and d15N within and between batches
was always <02 %. To compare isotope abundances of orchids and reference plants from different sites, the data were
normalized. Enrichment factors (e) were calculated per plot:
e ¼ dS dREF, with S as a single d13C or d15N value of an adult
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
orchid and REF as the mean value of non-orchid reference
plants from the respective plot (Preiss and Gebauer, 2008). The
original d13C and d15N values of orchids, the respective reference plants, fungi and soil samples are available in
Supplementary Data Table S2. Total N concentrations in leaf,
stem, fungus and soil samples were calculated from sample
weights and peak areas using a six-point calibration curve per
sample run based on acetanilide measurements (Gebauer and
Schulze, 1991). Acetanilide has a constant N concentration of
1036 %. Enrichment factors and total N concentrations of orchids and reference plants were tested for normal distribution.
Enrichment factors e13C and e15N were not normally distributed and therefore these data were tested for statistical differences using the Kruskal–Wallis non-parametric test followed
by a post-hoc Mann–Whitney U-test with an adjusted significance level according to Holm (1979). The autotrophic reference plants were treated as one group after confirming
insignificant differences among the data of each species. Total
N concentrations were normally distributed and thus the
Student’s t-test was used to test total N concentrations in orchids and reference plants for statistical differences.
427
EX
OC
H
MC
A
EX
H
OC
MC
B
RESULTS
EX
Histological studies
OC
In the three vanilloid study species, the below-ground structure
of Galeola falconeri had a long rhizome with a few thick roots.
Cyrtosia javanica had a short rhizome with a few thick roots.
Lecanorchis thalassica had a slender rhizome with a number of
thick roots (Supplementary Data Fig. S1). The exodermal cells
of roots in the three vanilloid orchids were characterized by the
thickened outer and lateral walls. Colonization by fungal hyphae in the middle cortex cells could be observed (Fig. 3), and
their exodermal and outer cortex layers were occasionally colonized (Fig. 3B).
The below-ground structures of Gastrodia appendiculata,
G. fontinalis and G. nantoensis were similar, having thick rhizomes with a few slim roots (Supplementary Data Fig. S1E).
The epidermal cells remained intact or became collapsed without fungal colonization. The outer and inner cortex cells were
usually uncolonized, whereas the middle cortex cells were filled
with fungal hyphae (Fig. 4A, B, D). Gastrodia flabilabella had
a tuberous rhizome with several coralloid roots (Supplementary
Data Fig. S1D). As observed in the other three Gastrodia spp.,
the epidermal, outer and inner cortical layers were rarely colonized. The middle cortex cells were heavily colonized by fungal
hyphae. It is worth noting that several papillae-like cell wall
thickenings could be observed at the adjoining walls between
the outer and middle cortex cells (Fig. 4A, C, D).
Molecular identification of mycorrhizal fungi
Two of the three MH vanilloid study species were associated
with SAP non-Rhizoctonia fungi known to be wood-decaying.
The ITS sequences obtained from seven G. falconeri individuals (22 samples) and those obtained from five C. javanica individuals (20 samples) had a high DNA sequence homology
with species of Meripilaceae (order Polyporales) by BLAST
H
MC
C
IC
FIG. 3. Histology of mycorrhizas in three vanilloid species. In their epidermal
cells, the outer and lateral walls become thickened. (A) Light micrograph showing a transverse section of a root of G. falconeri. Fungal colonization in the middle cortex cells could be observed. (B) Light micrograph showing a transverse
section of a root of C. javanica. A few cells in the exodermal, outer and inner
cortex cells are filled with fungal hyphae. (C) Light micrograph showing a transverse section of a root of L. thalassica. Fungal colonization is mainly found in
the middle cortex cells. EX, exodermal cells; H, fungal hyphae; IC, inner cortex;
MC, middle cortex; OC, outer cortex. Scale bar ¼ 100 mm.
analysis. Thus, wood-decaying Meripilaceae have to be considered as the exclusive fungal associates of G. falconeri and
C. javanica (Fig. 5). In contrast, for the third MH vanilloid orchid L. thalassica, the ITS sequences obtained from seven individuals (24 samples) demonstrate a high DNA sequence
homology with species of the ECM fungus Russula (Fig. 6).
For all Gastrodia spp. studied, associations with SAP nonRhizoctonia fungi were found. For the three Gastrodia spp.
growing sympatrically in a bamboo forest (G. appendiculata,
15 samples of five individuals; G. fontinalis, 22 samples of six
individuals; G. nantoensis, 20 samples of six individuals), the
ITS sequences demonstrate a high DNA sequence homology
with Mycena spp. The ITS sequences of Mycena obtained from
the roots of G. appendiculata, G. fontinalis and G. nantoensis
(57 root samples) are grouped into three types (Supplementary
428
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
E
E
OC
OC
H
MC
MC
IC
H
A
C
E
E
OC
H
OC
MC
MC
H
IC
IC
B
D
FIG. 4. Histology of mycorrhizas in four Gastrodia species. Their epidermal, outer and inner cortical layers are rarely colonized. (A) Light micrograph showing a
transverse section of a root of G. appendiculata. The middle cortex cells are heavily colonized by fungal hyphae. The thickened papillae-like cell walls could be observed at the adjoining walls between the outer and middle cortex cells (arrows). (B) Light micrograph showing a transverse section of a root of G. fontinalis. (C)
Light micrograph showing a transverse section of a root of G. flabilabella. (D) Light micrograph showing a transverse section of a root of G. nantoensis. A few papillae-like cell walls occur at the adjoining walls between the outer and middle cortex cells (arrows). E, epidermal cells; H, fungal hyphae; IC, inner cortex; MC, middle
cortex; OC, outer cortex. Scale bar ¼ 50 mm.
Data Table S4). Type I was only detected in G. fontinalis, and
type II was only detected in G. appendiculata. Type III was detected in both G. appendiculata and G. nantoensis. In addition,
the mycorrhizal roots of G. fontinalis (eight samples) were also
colonized by Gymnopus spp. For G. flabilabella (21 samples of
seven individuals), collected in a coniferous forest, the generated ITS sequences indicate homology with species of the fungal genus Hydropus (Fig. 7).
Stable isotope natural abundance and total N concentrations
Comparisons of enrichment factors e13C (H ¼ 77; d.f. ¼ 7;
P < 0001) and e15N (H ¼ 77; d.f. ¼ 7; P < 0001) among the
seven orchid species and the set of reference plants revealed
highly significant differences in our data set. All study orchids
were enriched by 79 6 02 % (G. appendiculata) to 121 6 05
% (G. flabilabella) in 13C and by 47 6 06 % (G. flabilabella)
to 88 6 27 % (L. thalassica) in 15N in comparison with autotrophic reference plants growing at the same sites (Fig. 8).
Based on post-hoc tests, this enrichment in 13C and 15N was
highly significant for all orchid species (in all cases U ¼ 0;
P < 0001).
The orchids themselves fall into three distinct groups. The
ECM-associated L. thalassica was significantly more enriched
in 15N than all other orchids associated with SAP fungi (U ¼ 0
to U ¼ 3; P < 0001 to P ¼ 0003; mean difference 37 %) and
significantly less enriched in 13C than the group composed of
G. falconeri, C. javanica and G. flabilabella (U ¼ 0; P < 0001;
mean difference 30 %). However, L. thalassica was not significantly distinguishable in its 13C enrichment from the orchid
group composed of G. nantoensis, G. appendiculata and G. fontinalis (U ¼ 25; P ¼ 0295). The group composed of G. falconeri, C. javanica and G. flabilabella was more enriched in 13C
than the group composed of G. nantoensis, G. appendiculata
and G. fontinalis (U ¼ 0; P < 0001; mean difference 35 %).
However, both of these groups were not significantly different
in their 15N enrichments (U ¼ 107; P ¼ 0836).
The investigated litter-decaying SAP fungus Marasmius sp.
was enriched by 94 6 05 % in 13C and by 43 6 02 % in 15N
compared with autotrophic reference plants (Fig. 8) and thus
had similar enrichments in 13C and 15N to the orchid group
composed of G. nantoensis, G. appendiculata and G. fontinalis.
Orchid flower stems had a slightly higher mean total N concentration (288 6 050 mmol g d. wt–1; n ¼ 35) than reference
plant leaves (267 6 067 mmol g d. wt–1; n ¼ 90;
Supplementary Data Table S2). However, this slight difference
was not significant (t ¼ 1671; P ¼ 0097; d.f. ¼ 123).
DISCUSSION
Fungal colonization in the roots
In three MH vanilloid orchids, mycorrhizal colonization of
wood-decaying and ECM fungi was mainly observed in the
cortical layers of the old roots, a finding similar to reports for
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
87/1·0
Mycobiont of Cyrtosia javanica (KP238184)
Mycobiont of Cyrtosia javanica (KP238182)
–/0·63
57/0·99
429
58/0·99
84/1·0
Polyporales sp. (AB470242)
Meripilus giganteus (AF287874)
Physisporinus vitreus (JQ031129)
Mycobiont of Galeola falconeri (KP238181)
Rigidoporus vinctus (AY333794)
74/1·0
Rigidoporus microporus (AY333795)
Podoscypha elegans (JN649356)
100/1·0
Podoscypha sp. (JN649365)
90/1·0
71/0·78
Trametes aff. maxima (JN164802)
Polyporus brumalis (AF347108)
Hypochnicium michelii (JN939579)
100/1·0
90/1·0
71/0·99
Hypochnicium eichleri (AJ406508)
Hypochnicium erikssonii (DQ677508)
Hypochnicium lyndoniqe (JX124704)
–/0·53
60/0·54
Mycoacia cf. columellifera (JN710572)
Nigroporus vinosus (JN710576)
Steccherinum cf. murashkinskyi (JN710586)
Antrodiella foliaceodentata (JN710515)
Climacocystis borealis (JN710527)
Coltriciella oblectabilis (KC155387)
0·1
FIG. 5. Phylogenetic relationships of the mycorrhizal fungi of G. falconeri and C. javanica based on the Bayesian analysis of LSU ribosomal DNA sequences of
Polyporales available in GenBank (Justo and Hibbett, 2011; Binder et al., 2013). GenBank accession numbers are shown in parentheses. The values above branches
are bootstrap percentages and Bayesian posterior probabilities (>50 %), respectively. ‘–’ indicates that the node was not supported in MP analysis.
mycorrhizal colonizations in chlorophyllous Vanilla spp. by
Porras-Alfaro and Bayman (2007). The outermost layers of old
roots in MH vanilloid orchids are characterized by thickened
exodermal layers, whereas the epidermal cells are no longer
alive. Colonization of fungal hyphae was also found in the
thickened exodermal layers (Fig. 3B), suggesting their roles for
maintenance of nutrient uptake by older roots (Esnault et al.,
1994).
In the Gastrodia spp. investigated, fungal colonization was
restricted to a few cortical layers of root systems, but was not
commonly observed in rhizomes. In the colonized cortical
layers, the papillae-like cell wall thickenings were abundant at
the adjoining walls between the outer and middle cortex cells,
corresponding to pathways for fungal hyphae (Fig. 4A, C, D).
The presence of papillae-like cell wall thickenings could be potentially underdeveloped structures of wall ingrowths in specialized transfer cells as described in symbiotic associations by
Pate and Gunning (1972), suggesting a specific nutrient transport network in the mycorrhiza (Martos et al., 2009).
Mycorrhizal partners
Subfamily Vanilloideae contain a number of non-photosynthetic genera (40 % of the 15 genera of vanilloid orchids), e.g.
Cyrtosia, Erythrorchis, Galeola, Pseudovanilla and
Lecanorchis (Cameron, 2009). Chlorophyllous Vanilla spp.
have been shown to associate with a wide range of Rhizoctonia
fungi, including Ceratobasidium, Thanatephorus and
Tulasnella (Porras-Alfaro and Bayman, 2007), whereas the MH
vanilloid taxa Cyrtosia and Erythrorchis mainly associate with
wood-decaying fungi, such as Armillaria and species of
Hymenochaetaceae and Polyporaceae (Umata, 1995, 1997a;
Cha and Igarashi, 1996; Dearnaley, 2007). In this study, G. falconeri and C. javanica were identified to associate exclusively
with wood-decaying fungi of Meripilaceae (order Polyporales).
According to the findings of Okayama et al. (2012),
Lecanorchis spp. in Japan associate with a broad range of ECM
fungi, including Lactarius, Russula, Atheliaceae and Sebacina,
with Lactarius and Russula dominating. In this study, Russula
was identified as the preferred mycorrhizal partner of L. thalassica. In phylogenetic analyses, MH vanilloid orchids, i.e.
Cyrtosia, Erythrorchis, Galeola, Pseudovanilla and
Lecanorchis, are closely related to chlorophyllous Vanilla spp.
in tribe Vanilleae (Cameron, 2009). All these results together
suggest that the nutritional shift from autotrophy to mycoheterotrophy in vanilloid orchids correlates with shifts in fungal
partners from Rhizoctonia fungi to wood-decaying nonRhizoctonia fungi or to ECM fungi. A shift towards either a
wood-decaying fungus or an ECM fungus can even happen for
sympatrically growing closely related species, as in our case for
C. javanica and L. thalassica. Liebel et al. (2015) recently argued that this shift in fungal partners is essential for the MH
mode of nutrition. The annual C and N flux from Rhizoctonia
fungi to their orchid partners is rather low (Stöckel et al.,
2014). This low matter flux is obviously sufficient to support
growth of the tiny initially MH orchid protocorms, but appears
to be insufficient to support growth of adult MH orchids.
430
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
–/0·75
68/0·99
79/1·0
96/1·0
83/0·98
100/1·0
98/1·0
56/0·99
95/1·0
–/0·59
–/0·99
94/1·0
97/1·0
80/0·99
–/0·81
–/0·77
–/0·98
100/1·0
62/0·77
100/1·0
51/0·98
98/1·0
Russula brunneoviolacea (AM113956)
Russula cuprea (AY061667)
100/1·0
Mycobiont of L. kiusiana var. kiusiana KK-06M Mugi (AB597680)
Mycobiont of L. kiusiana var. kiusiana KK-07M Mugi (AB597681)
Russula azurea (AY061660)
Russula lilacea (AY061731)
Mycobiont of L. japonica var. hokuriluensis JH-13M Jindai (AB597651)
Russula romellii (AY061714)
Mycobiont of L. japonica var. japonica JJ-08M Hachioji (AB597661)
Russula rosea (AY061715)
Mycobiont of L. japonica var. hokuriluensis JH-04M Joetsu (AB597642)
Russula peckii (EU598174)
Mycobiont of L. thalassica (KP238187)
Russula fellea (AF418616)
Russula ochroleuca (EU350580)
Russula raoultii (AF418621)
Russula gracillima (AY061678)
Mycobiont of L. nigricans NI-02M Shizuoka (AB597695)
60/0·89
61/0·96
–/0·54
–/0·52
–/0·66
100/1·0
–/0·82
99/1·0
71/0·91
100/1·0
–/0·96
–/0·68
64/0·97
86/1·0
71/0·67
61/0·51
–/0·67
100/1·0
–/1·0
100/0·52
91/0·88
96/1·0
93/1·0 63/0·89
100/1·0
–/0·53
53/0·99
100/1·0
–/0·93
100/1·0
100/1·0
86/1·0
100/1·0
89/1·0
74/0·99
100/1·0
56/0·61
100/1·0
100/1·0
76/0·97
100/1.0
Lactarius quietus (AF096982)
Arcangeliella camphorata (EU644702)
Russula postiana (AF230898)
Mycobiont of L. virella VI-01M Kumanoe (AB597714)
Russula roseipes (AY061716)
Mycobiont of L. nigricans NI-11M Kiyosumi (AB597704)
Russula turci (EF530935)
Russula solaris (AF418627)
Russula firmula (AF418631)
Russula rubra (AY061717)
Mycobiont of L. japonica var. japonica JJ-10M Hachioji (AB597663)
Mycobiont of L. japonica var. japonica JJ-10M Hachioji (AB597664)
Mycobiont of L. japonica var. japonica JJ-10M Nichinan (AB597665)
Mycobiont of L. japonica var. japonica JJ-13M Nichinan (AB597666)
Russula claroflava (AY061665)
Russula occidentalis (AY534206)
Russula integra (AY061683)
Russula decolorans (DQ367913)
Russula xerampelina (AF418632)
Russula favrei (EF530944)
Russula pascua (AF061705)
Russula aeruginea (EU819421)
Mycobiont of L. japonica var. japonica JJ-14M Kiyosumi (AB597667)
Mycobiont of L. kiusiana var. kiusiana KK-08M Mugi (AB597682)
Mycobiont of L. trachycaula TR-02M Shishikui (AB597706)
Mycobiont of L. trachycaula TR-05M Shishikui (AB597709)
Russula nitida (AY061696)
Russula sphagnaphila (AY061719)
Russula paludosa (AJ971402)
Russula xerampelina var. xerampelina (AJ971402)
Russula mairei (AM113959)
Mycobiont of L. nigricans NI-10M Kiyosumi (AB597703)
Russula emetica (AY061673)
Russula betularum (EU598183)
Russula raoultii (AY061712)
Russula cremoricolor (DQ974755)
Mycobiont of L. nigricans NI-04M Shizuoka (AB597697)
Mycobiont of L. nigricans NI-05M Shizuoka (AB597698)
Russula vesca (AM113965)
Russula vesca (AY606965)
Mycobiont of L. japonica var. japonica JJ-09M Hachioji (AB597662)
Mycobiont of L. japonica var. hokurikuensis JH-11M Keta (AB597649)
Russula heterophylla (DQ422006)
Russula parazurea (DQ422007)
Russula ilicis (AY061682)
Mycobiont of L. japonica var. hokurikuensis JH-10M Kanazawa (AB597648)
Mycobiont of L. trachycaula TR-09M Amami (AB597713)
Russula pallescens (DQ421987)
Russula pallidoapora (AY061701)
Russula littoralis (AY061702)
Mycobiont of L. japonica var. kiiensis JK-02M Tajimi (AB597671)
Russula laurocerasi (AY061735)
Russula illota (DQ422024)
Russula nigricans (EF534352)
Mycobiont of L. flavicans var. flavicans FA-01M Mugi Oshima (AB597630)
Russula nigricans (EU597075)
Mycobiont of L. trachycaula TR-06M Setouchi (AB597710)
Russula compacta (EU598172)
Mycobiont of L. trachycaula TR-06M Shishikui (AB597705)
0·1
FIG. 6. Phylogenetic relationships of the mycorrhizal fungi of Lecanorchis thalassica based on the Bayesian analysis of partial ITS ribosomal DNA sequences of
Russulaceae available in GenBank (with the position of mycorrhizal fungi found in Lecanorchis by Okayama et al., 2012). GenBank accession numbers are shown
in parentheses. The values above branches are bootstrap percentages and Bayesian posterior probabilities (>50 %), respectively. ‘–’ indicates that the node was not
supported in MP analysis.
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
100/0·74
100/0·87
63/0·55
100/0·58
95/1·0
100/0·95
100/0·89
–/0·57
84/1
75/1·0
100/0·76
91/1·0
51/0·51
100/0·99
100/1·0
100/0·97
–/0·89
–/0·51
–/0·52
100/0·99
94/0·68
74/0·81
94/1·0
–/0·56
52/0·96
61/0·97
64/0·95
–/0·57
99/1·0
88/0·78
100/1·0
58/1·0
93/1·0
–/0·67
75/0·92
55/0·59
64/0·97
–/0·9
–/0·72
99/1·0
100/1·0
91/0·65
53/1·0
–/0·62
–/0·77
–/0·91
–/0·6
–/0·99
99/1·0
–/0·97
–/0·92
80/0·88
82/1·0
99/1·0
–/0·98
60/0·74
81/1·0
55/1·0
71/1
56/0·99
–/0·99
86/1·0
–/0·97
–/0·67
66/0·98
–/0·51
0·1
431
Mycena polygramma (DQ071707)
Mycena polygramma (AY207243)
Mycena leaiana (AF261411)
Mycena galericulata (AF042636)
Mycena crocata (AY207241)
Mycena aurantiomarginata (AY207246)
Mycena niveipes (AY207242)
Mycena leptocephala (AY207253)
Mycena renati (AY207256)
Mycena citrinomarginata (AY207248)
Mycena olivaceomarginata (AY207255)
Mycena galopus (AY207250)
Mycena sanguinolenta (AY207257)
Mycena haematopus (AY207252)
Mycena maculata (AY207254)
Mycena insignis (AF261413)
Mycena calavicularis (AF042637)
Mycena tintinnabulum (AY207258)
Mycena epipterygia (AY207249)
Cruentomycena viscidocruenta (AF261414)
Dictyopanus pusillus (AF261425)
Dictyopanus sp. (AF261425)
Panellus stypticus (AF261427)
Resinomycena acadiensis (AF042638)
Resinomycena rhododendri (AF261415)
Poromycena sp. (AF261429)
Mycobiont of Gastrodia confusa (AB454415)
Mycena plumbea (DQ470813)
Mycena galericulata (AY647216)
Mycena galericulata (AY207251)
Mycena rubromarginata (AY207245)
Mycena zephirus (AY207259)
Poromycena sp. (AF261421)
Fiboboletus gracilis (AF261422)
Mycenoporella griseipora (AF261428)
Poromycena manipularis (AF261423)
Mycena aff. pura (AY261409)
Mycena aff. pura (AF261410)
Mycena rutilanthiformis (AF042606)
Mycena aff. pura (AY207244)
Mycobiont of Gastrodia confusa (AB454414)
Mycobiont of Gastrodia appendiculata (KP238191)
Mycobiont of Gastrodia nantornsis (KP238188)
Mycobiont of Gastrodia fontinalis (KP238190)
Mycobiont of Gastrodia confusa (AB454416)
Mycena amicta (DQ457692)
Mycena aff. pura (DQ457688)
Mycena aff. pura (DQ457689)
Mycena aff. pura (DQ457690)
Cotobrusia calostomoides (AF261424)
Mycena capillaripes (AY207247)
Tricholoma matsutake (U62964)
Entoloma prunuloides (AY700180)
Catathelasma ventricosum (DQ089012)
Lyophyllum decastes (AF042583)
Mycobiont of Gastrodia flabilabella (KP238192)
Hydropus cf scabripes (DQ411536)
Hydropus marginellus (DQ457674)
Megacollybia platyphylla (AY635778)
Gymnopus aff moseri (AY639409)
Mycobiont of Gastrodia similis (FJ179468)
Gymnopus aff menehune (AY639408)
Gymnopus indoctus (AY639418)
Mycobiont of Wullschlaegelia aphylla (FJ179476)
Marasmiellus synodicus (AY639435)
Gymnopus nonnullus var attenuatus (AY639426)
Gymnopus melanopus (AY639422)
Mycobiont of Gastrodia fontinalis (KP238189)
Gymnopus brunneigracilis (AY639412)
Gymnopus gibbosus (AY639415)
Gymnopus termiticola (AY639430)
Mycobiont of Wullschlaegelia aphylla (FJ179475)
Marasmius androsaceus (AF261585)
Gymnopus fusioes (AY256710)
Gymnopus bicolor (AY639411)
Gymnopus sepiiconicus (AY639427)
Gymnopus spissus (AY639428)
Rhodocollybia maculata (AY639880)
Rhodocollybia butyracea (EU486454)
Marasmiellus opacus (AF261330)
Marasmius scorodonius (AF261332)
Marasmius alliaceus (AY635776)
Marasmius alliaceus (AY639436)
Marasmius copelandii (AY639438)
Marasmiellus palmivorus (AY639434)
Marasmius oreades (DQ156126)
Marasmius rotula (DQ457686)
Henningsomyces candidus (AF287864)
Nia vibrissa (AF334750)
Cyphella digitalis (AY635771)
FIG. 7. Phylogenetic relationships of the mycorrhizal fungi of Gastrodia appendiculata, G. fontinalis, G. flabilabella and G. nantoensis based on the Bayesian analysis of LSU ribosomal DNA sequences of marasmioid available in GenBank (Moncalvo et al., 2000, 2002; Wilson and Desjardin, 2005; Matheny et al., 2006; Martos
et al., 2009; Ogura-Tsujita et al., 2009). The values above branches are bootstrap percentages and Bayesian posterior probabilities (>50 %), respectively. ‘–’ indicates that the node was not supported in MP analysis.
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
432
Enrichment factor e 15N (%o)
12
Lecanorchis thalassica
10
Galeola falconeri
8
G. nantoensis
G. fontinalis
6
4
G. appendiculata
Marasmius sp.
2
G. flabilabella
Cyrtosia javanica
0
–2
Ref
–2
0
2
4
6
8
10
12
Enrichment factor e 13C (%o)
FIG. 8. Mean enrichment factors e13C and e15N 6 1 s.d. as calculated for six MH
orchid species associated with SAP non-Rhizoctonia fungi (triangles), one
mycoheterotrophic orchid species associated with a fungus forming ECM (filled
square), one litter-decaying SAP fungus (open circle) (n ¼ 5 each) and of photosynthetic reference plants (Ref, n ¼ 90, see Supplementary Data Table S2) collected together with each of the orchids at four sites in the Xitou Experimental
Forest, Nantou County, Taiwan. Please note that the orchids associated with
SAP non-Rhizoctonia fungi fall into two significantly distinct groups, Galeola
falconeri, Cyrtosia javanica, Gastrodia flabilabella (filled triangles) and
Gastrodia fontinalis, G. nantoensis and G. appendiculata (open triangles). Both
of these groups are significantly different from Lecanorchis thalassica, the orchid associated with an ECM fungus. Mean e values of reference plants are zero
by definition. The box represents 6 1 s.d. for the reference plants.
Despite the fairly high number of fully MH species in subfamily Vanilloideae, no partially MH species have been reported so
far to occur among green species of Vanilloideae.
The four Gastrodia spp. investigated here also fall into two
groups with respect to their mycorrhizal partners. Mycena dominates as fungal host in the three species growing sympatrically
in a bamboo forest (G. appendiculata, G. fontinalis and G. nantoensis). Furthermore, Gymnopus was detected in the mycorrhizal roots of G. fontinalis. It is worth mentioning that the three
Gastrodia species produce their fruit bodies at different times
in the year. Gastrodia fontinalis produces fruit bodies in spring
and G. nantoensis and G. appendiculata appear successively in
October. Note that G. fontinalis associates with type I Mycena
and Gymnopus, G. nantoensis only associates with type III
Mycena, and G. appendiculata recruits Mycena types II and III.
Thus, the results demonstrate a preference towards different lineages of Mycena by three sympatric Gastrodia spp. In G. confusa from Japan, an association with various types of Mycena
was found in the same and in different populations, suggesting
an additional geographical mosaic of mycorrhizal specificity
(Ogura-Tsujita et al., 2009).
The fourth Gastrodia sp., growing in a coniferous forest
(G. flabilabella), has Hydropus as a mycorrhizal partner.
Unlike the other three Gastrodia spp., G. flabilabella forms its
own clade (Hsu, 2008) and mainly occurs in coniferous forests.
The change of mycorrhizal fungal associations may reflect not
only the independent evolution of Gastrodia lineages, but also
a switch in forest types as their habitats. Previous studies have
shown that Gastrodia spp. associate with litter- and wooddecomposing SAP basidiomycetes of the genera Armillaria,
Mycena, Resinicium and Campanella or Marasmius (Kusano,
1911; Martos et al., 2009; Ogura-Tsujita et al., 2009; Dearnaley
and Bougoure, 2010). Here we provide molecular evidence that
Gymnopus and Hydropus are additional mycorrhizal partners of
Gastrodia spp., and this broadens the diversity of SAP basidiomycetes associating with Gastrodia. Gastrodia is one of the
largest MH genera, consisting of >50 fully MH species, distributed in Asia, Australia and Africa (Hsu, 2008), and the recruitment of various basidiomycetes as mycorrhizal partners needs
further investigation. All fungal hosts so far known of all
Gastrodia spp. so far investigated are either litter- or wooddecaying non-Rhizoctonia SAP fungi. A better knowledge
about the preferred substrates (litter or wood) of the respective
fungal hosts would be desirable for the future in order to allow
an even more comprehensive use of information available from
stable isotope natural abundance (see below).
Carbon and nitrogen sources and total nitrogen concentrations
All seven of the MH study orchids turned out to be significantly enriched in the heavy isotopes 13C and 15N in comparison with autotrophic plants growing in identical habitats. This
finding confirms that fungi also known to be enriched in 13C
and 15N (Gebauer and Dietrich, 1993; Gleixner et al., 1993)
serve as their C and N source. Though leafless, some putative
MH orchids have green flowering stems and thus a low photosynthetic activity. One leafless orchid with frequently green
flowering stems and green seed capsules is Corallorhiza trifida.
This species is associated with an ECM fungus and is less enriched in 13C and 15N than fully MH orchids also associated
with ECM fungi (Zimmer et al., 2008). Thus, C. trifida has
been classified as being partially MH. All study orchids of this
investigation provided no indications for chlorophyllous flowering stems and had isotope signatures typical of various types of
fully MH plants. According to their pattern in 13C and 15N enrichment, the MH orchids fall into three groups. (1) The ECMassociated L. thalassica is the species most enriched in 15N.
This pattern is in agreement with current knowledge (Hynson
et al., 2013) and can be traced back to the fact that ECM fungi
are more enriched in 15N than sympatrically growing wood- or
litter-decaying SAP fungi (Gebauer and Taylor, 1999). (2) The
two orchid species associated with wood-decaying
Meripilaceae, Galeola falconeri and Cyrtosia javanica, belong
to the group most enriched in 13C. The specifically high 13C enrichment of this group of MH orchids is expected due to the
fact that wood is more enriched in 13C than leaf tissue (Gebauer
and Schulze, 1991; Cernusak et al., 2009) and confirms previous findings of similar patterns in MH orchids associated with
wood-decaying fungi (Ogura-Tsujita et al., 2009; Martos et al.,
2009; Hynson et al., 2013). The third MH orchid belonging to
this group is the Hydropus-associated Gastrodia flabilabella,
suggesting that this Hydropus sp. should be a wood-decaying
fungus. (3) The third MH orchid cluster characterized by a significantly lower 15N enrichment than (1) and a significantly
lower 13C enrichment than (2) is composed of the three
Gastrodia spp. growing sympatrically in a bamboo forest,
G. appendiculata, G. fontinalis and G. nantoensis. The isotopic
pattern of this cluster of MH orchids provides strong evidence
that their mycorrhizal associates, Mycena of three different
types and Gymnopus, are all litter-decaying fungi. This
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
evidence is further strengthened by the rather similar isotopic pattern found for the fruit bodies of the litter-decaying
fungus Marasmius sp. This is the first report we are aware
of that indicates a significantly different isotopic pattern
for MH orchids associated with either wood-decaying or
litter-decaying fungi. The only hint towards an isotopic
distinction between MH orchids associated with wood- or
litter-decaying fungi was previously given by Martos et al.
(2009), who investigated the isotopic compositions of
the MH orchids Gastrodia similis associated with wooddecaying
Resinicium
from
La
Réunion
and
Wullschlaegelia aphylla mycorrhizal orchids with litterdecaying Gymnopus and Mycena from Guadeloupe.
Unfortunately, comparisons of the data from the study of
Martos et al. with each other and with our investigation based
on normalized enrichment factors is not possible due to a lack
of stable isotope data for forest ground plants growing in micro-environments identical to the MH orchids.
Total N concentrations in MH orchids have been reported to
be significantly higher than in the majority of co-occurring autotrophic plants (Gebauer and Meyer, 2003; Liebel et al., 2010;
Liebel and Gebauer, 2011; Sommer et al., 2012). The most
likely reason for these unusually high total N concentrations in
MH orchid tissues is their N gain through the fungal source.
Fungi are known to have considerably higher total N concentrations in their tissue than autotrophic plants growing in the same
environment (Gebauer and Dietrich, 1993; Gebauer and Taylor,
1999). Total N concentrations found in the flower stems of the
MH orchid species investigated here (288 6 050 mmol g
d. wt–1) are rather similar to the total N concentrations reported
for other fully or partially MH orchids (Gebauer and Meyer,
2003; Liebel et al., 2010; Liebel and Gebauer, 2011; Sommer
et al., 2012). The reference plant leaf total N concentrations investigated here (267 6 067 mmol g d. wt–1) were only slightly
lower than N concentrations in the flower stems of the MH orchids and statistically not distinguishable. Total N concentrations in the leaves of the understorey plants investigated here
from four different forests in Taiwan are about twice as high as
leaf total N concentrations in autotrophic forest ground plants
from temperate Central Europe (Gebauer et al., 1988; Gebauer
and Meyer, 2003) and Mediterranean Italy (Liebel et al., 2010),
and three times higher than leaf total N concentrations in autotrophic forest ground plants from severely N-limited regions,
such as SW Australia (Sommer et al., 2012) or boreal Norway
(Liebel and Gebauer, 2011). The most likely reason for the
high total N concentrations in the leaves of autotrophic forest
ground plants in Taiwan is a high mineral N availability due to
high soil N mineralization rates. Thus, N limitation as a driver
for the switch from autotrophy towards mycoheterotrophy is
unlikely for the MH orchids investigated here. More likely as a
driving factor for the development of mycoheterotrophy is light
limitation for photosynthesis on the forest ground. As reported
for other habitats of MH plants (Bidartondo et al., 2004;
Zimmer et al., 2007; Preiss et al., 2010) only 2–6 % of incoming light reached the ground of our study forests and thus severely limited the photosynthetic capacity of forest ground
vegetation.
In conclusion, our data provide further evidence for the importance of associations with non-Rhizoctonia SAP fungi
among fully MH orchids from sub-tropical Asia. Abundant
433
fallen litter and wood in the warm and humid forests of Taiwan
obviously supplies ideal substrates for the continuous growth of
SAP fungi. For the MH orchids studied here, as seeds germinate, they can construct efficient mycorrhizal interactions preferably with wood- or litter-decaying SAP fungal partners, but
sympatrically also with ECM fungi, in order to thrive in deeply
shaded forest understoreys with low light conditions.
SUPPLEMENTARY DATA
Supplementary data are available online at www.aob.oxfordjournals.org and consist of the following. Table S1: voucher
and sampling sites of seven mycoheterotrophic orchids in Xitou
Experimental Forest used in molecular identification of mycorrhizal fungi. Table S2: mean d15N and d13C values, mean enrichment factors e15N and e13C, and mean total N
concentrations of all plant and fungal samples in this study.
Table S3: putative taxonomic identity of the fungi detected in
this study. Table S4: ITS sequence types in Mycena from mycorrhizal roots of three sympatric Gastrodia spp. in a bamboo
forest. Figure S1: morphology of rhizomes and roots of mycoheterotrophic orchids in this study.
ACKNOWLEDGEMENTS
The authors thank Christine Tiroch (BayCEER–Laboratory of
Isotope Biogeochemistry, University of Bayreuth) for skilful
technical assistance with stable isotope abundance measurements. The authors also thank Yu-Hsiu Cho (National
Museum of Natural Science) for molecular identifications of
fungal partners, and Sheng-Kun Yu for providing the photo of
Lecanorchis thalassica. This work is a contribution to the
German Research Foundation Project GE 565/7-2. This work
was also supported by the funding from National Museum of
Natural Science, Taiwan to Y.-I.L.
LITERATURE CITED
Akaike H. 1974. A new look at the statistical model identification. IEEE
Transactions on Automatic Control 19: 716–723.
Bidartondo MI, Burghardt B, Gebauer G, Bruns TD, Read DJ. 2004.
Changing partners in the dark: isotopic and molecular evidence of ectomycorrhizal liaisons between forest orchids and trees. Proceedings of the Royal
Society B: Biological Sciences 271: 1799–1806.
Binder M, Justo A, Riley R, et al. 2013. Phylogenetic and phylogenomic overview of the Polyporales. Mycologia 105: 1350–1373.
Cameron KM. 2009. On the value of nuclear and mitochondrial gene sequences
for reconstructing the phylogeny of vanilloid orchids (Vanilloideae,
Orchidaceae). Annals of Botany 104: 377–385.
Cernusak LA, Tcherkez G, Keitel C, et al. 2009. Why are non-photosynthetic
tissues generally 13C enriched compared with leaves in C3 plants? Review
and synthesis of current hypotheses. Functional Plant Biology 36: 199–213.
Cha JY, Igarashi T. 1996. Armillaria jezoensis, a new symbiont of Galeola septentrionalis (Orchidaceae) in Hokkaido. Mycoscience 37: 21–24.
Dearnaley JDW. 2007. Further advances in orchid mycorrhizal research.
Mycorrhiza 17: 475–486.
Dearnaley JDW, Bougoure JJ. 2010. Isotopic and molecular evidence for saprotrophic Marasmiaceae mycobionts in rhizomes of Gastrodia sesamoides.
Fungal Ecology 3: 288–294.
Dearnaley JDW, Martos F, Selosse M-A. 2012. Orchid mycorrhizas:
molecular ecology, physiology, evolution and conservation aspects. In: B
Hock, ed. Fungal associations. The mycota IX, 2nd edn. Berlin: Springer
Verlag, 207–230.
434
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
Esnault AL, Masuhara G, McGee PA. 1994. Involvement of exodermal passage cells in mycorrhizal infection of some orchids. Mycological Research
98: 672–676.
Felsenstein J. 1985. Confidence limits on phylogenies: an approach using the
bootstrap. Evolution 39: 783–791.
Fry B. 2006. Stable isotope ecology. New York: Springer.
Gardes M, Bruns TD. 1993. ITS primers with enhanced specificity for basidiomycetes: application to the identification of mycorrhizae and rusts.
Molecular Ecology 2: 113–118.
Gebauer G, Dietrich P. 1993. Nitrogen isotope ratios in different compartments
of a mixed stand of spruce, larch and beech trees and of understorey vegetation including fungi. Isotopenpraxis 29: 35–44.
Gebauer G, Meyer M. 2003. 15N and 13C natural abundance of autotrophic and
myco-heterotrophic orchids provides insight into nitrogen and carbon gain
from fungal association. New Phytologist 160: 209–223.
Gebauer G, Schulze ED. 1991. Carbon and nitrogen isotope ratios in different
compartments of a healthy and a declining Picea abies forest in the
Fichtelgebirge, NE Bavaria. Oecologia 87: 198–207.
Gebauer G, Taylor AFS. 1999. 15N natural abundance in fruit bodies of different functional groups of fungi in relation to substrate utilization. New
Phytologist 142: 93–101.
Gebauer G, Rehder H, Wollenweber B. 1988. Nitrate, nitrate reduction and organic nitrogen in plants from different ecological and taxonomic groups of
Central Europe. Oecologia 75: 371–385.
Gleixner G, Danier HJ, Werner RA, Schmidt HL. 1993. Correlations between
the 13C content of primary and secondary plant products in different cell
compartments and that in decomposing basidiomycetes. Plant Physiology
102: 1287–1290.
Hamada M. 1939. Studien über die Mykorrhiza von Galeola septentrionalis
Reichb. F. – ein neuer Fall der Mykorrhiza-Bildung durch intraradicale
Rhizomorpha. Japanese Journal of Botany 10: 151–211.
Holm S. 1979. A simple sequentially rejective multiple test procedure.
Scandinavian Journal of Statistics 6: 65–70.
Hsu TC. 2008. Taxonomy of Gastrodia (Orchidaceae) in Taiwan. Master thesis,
National Taiwan University, Taipei.
Hynson NA, Madsen TP, Selosse M-A, et al. 2013. The physiological ecology
of mycoheterotrophy. In: VSFT Merckx, ed. Mycoheterotrophy: the biology
of plants living on fungi. Berlin: Springer Verlag, 297–342.
Justo A, Hibbett DS. 2011. Phylogenetic classification of Trametes
(Basidiomycota, Polyporales) based on a five-marker dataset. Taxon 60:
1567–1583.
Kikuchi G, Higuchi M, Morota T, Nagasawa E, Suzuki A. 2008. Fungal symbiont and cultivation test of Gastrodia elata Blume (Orchidaceae). Journal
of Japanese Botany 83: 88–95.
Kusano S. 1911. Gastrodia elata and its symbiotic association with Armillaria
mellea. Journal of the College of Agriculture Imperial University of Tokyo
4: 1–65.
Leake JR. 1994. The biology of myco-heterotrophic (‘saprophytic’) plants. New
Phytologist 127: 171–216.
Leake JR. 2004. Myco-heterotroph/epiparasitic plant interactions with ectomycorrhizal and arbuscular mycorrhizal fungi. Current Opinions in Plant
Biology 7: 422–428.
Liebel HT, Gebauer G. 2011. Stable isotope signatures confirm carbon and nitrogen gain through ectomycorrhizas in the ghost orchid Epipogium aphyllum Swartz. Plant Biology 13: 270–275.
Liebel HT, Bidartondo MI, Preiss K, et al. 2010. C and N stable isotope
signatures reveal constraints to nutritional modes in orchids
from the Mediterranean and Macaronesia. American Journal of Botany 97:
903–912
Liebel HT, Bidartondo MI, Gebauer G. 2015. Are carbon and nitrogen exchange between fungi and the orchid Goodyera repens affected by irradiance? Annals of Botany 115: 251–261.
Martos F, Dulormne M, Pailler T, et al. 2009. Independent recruitment of saprotrophic fungi as mycorrhizal partners by tropical achlorophyllous orchids.
New Phytologist 184: 668–681.
Matheny PB, Curtis JM, Hofstetter V, et al. 2006. Major clades of Agaricales:
a multilocus phylogenetic overview. Mycologia 98: 982–995.
Merckx VSFT. 2013. Mycoheterotrophy: an introduction. In: VSFT Merckx, ed.
Mycoheterotrophy: the biology of plants living on fungi. Berlin: Springer
Verlag, 1–17.
Moncalvo JM, Lutzoni FM, Rehner SA, Johnson J, Vilgalys R. 2000.
Phylogenetic relationships of agaric fungi based on nuclear large subunit
ribosomal DNA sequences. Systematic Biology 49: 278–305.
Moncalvo JM, Vilgalys R, Redhead SA, et al. 2002. One hundred and
seventeen clades of euagarics. Molecular Phylogenetics and Evolution 23:
357–400.
Nylander JAA. 2004. MrModeltest 2.2. Program distributed by the author.
Evolutionary Biology Centre, Uppsala University.
Ogura-Tsujita Y, Yukawa T. 2008. High mycorrhizal specificity in a widespread mycoheterotrophic plant, Eulophia zollingeri (Orchidaceae).
American Journal of Botany 95: 93–97.
Ogura-Tsujita Y, Gebauer G, Hashimoto T, Umata H, Yukawa T. 2009.
Evidence for novel and specialized mycorrhizal parasitism: the orchid
Gastrodia confusa gains carbon from saprotrophic Mycena. Proceedings of
the Royal Society B: Biological Sciences 276: 761–767.
Okayama M, Yamato M, Yagame T, Iwase K. 2012. Mycorrhizal diversity
and specificity in Lecanorchis (Orchidaceae). Mycorrhiza 22: 545–553.
Pate JS, Gunning BES. 1972. Transfer cells. Annual Review of Plant
Physiology 23: 173–196.
Porras-Alfaro A, Bayman P. 2007. Mycorrhizal fungi of Vanilla: diversity, specificity and effects on seed germination and plant growth. Mycologia 99:
510–525.
Preiss K, Gebauer G. 2008. A methodological approach to improve estimates
of nutrient gains by partially myco-heterotrophic plants. Isotopes in
Environmental and Health Studies 44: 393–401.
Preiss K, Adam IKU, Gebauer G. 2010. Irradiance governs exploitation
of fungi: fine-tuning of carbon gain by two partially myco-heterotrophic
orchids. Proceedings of the Royal Society B: Biological Sciences 277:
1333–1336.
Rasmussen HN. 1995. Terrestrial orchids – from seed to mycotrophic plant.
Cambridge: Cambridge University Press.
Ronquist F, Huelsenbeck JP. 2003. MRBAYES 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19: 1572–1574.
Roy M, Whatthana S, Stier A, Richard F, Vessabutr S, Selosse M-A. 2009.
Two mycoheterotrophic orchids from Thailand tropical dipterocarpacean
forests associate with a broad diversity of ectomycorrhizal fungi. BMC
Biology 7: 51.
Sommer J, Pausch J, Brundrett MC, Dixon KW, Bidartondo MI, Gebauer
G. 2012. Limited carbon and mineral nutrient gain from mycorrhizal fungi
by adult Australian orchids. American Journal of Botany 99: 1133–1145.
Stöckel M, Těšitelová T, Jersáková J, Bidartondo MI, Gebauer G. 2014.
Carbon and nitrogen gain during the growth of orchid seedlings in nature.
New Phytologist 202: 606–615.
Stover BC, Muller KF. 2010. TreeGraph 2: combining and visualizing evidence
from different phylogenetic analyses. BMC Bioinformatics 11: 7.
Su HJ. 2000. Orchidaceae. In TC Huang, ed. Flora of Taiwan, 2nd edn, Vol. 5.
Editorial Committee of the Flora of Taiwan, Department of Botany,
National Taiwan University, Taipei, Taiwan, 729–1086.
Swofford DL. 2002. PAUP*: phylogenetic analysis using parsimony (*and other
methods), version 4.0b10 for Macintosh. Sunderland, MA: Sinauer
Associates.
Taylor DL, Bruns TD. 1997. Independent, specialized invasions of ectomycorrhizal mutualism by two non-photosynthetic orchids. Proceedings of the
National Academy of Sciences of the USA 94: 4510–4515.
Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG. 1997.
The Clustal X windows interface. Flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Research
24: 4876–4882.
Trudell SA, Rygiewicz PT, Edmonds RL. 2003. Nitrogen and carbon stable
isotope abundances support the myco-heterotrophic nature and host-specificity of certain achlorophyllous plants. New Phytologist 160: 391–401.
Umata H. 1995. Seed germination of Galeola altissima, an achlorophyllous orchid, with Aphyllophorales fungi. Mycoscience 36: 369–372.
Umata H. 1997a. Formation of endomycorrhizas by an achlorophyllous orchid,
Erythrorchis ochobiensis, and Auricularia polytricha. Mycoscience 38:
335–339.
Umata H. 1997b. In vitro germination of Erythrorchis ochobiensis
(Orchidaceae) in the presence of Lyophyllum shimeji, an ectomycorrhizal
fungus. Mycoscience 38: 355–357.
Vilgalys R, Hester M. 1990. Rapid genetic identification and mapping of enzymatically amplified ribosomal DNA from several Cryptococcus species.
Journal of Bacteriology 172: 4238–4246.
White TJ, Bruns TD, Lee S, Taylor JW. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: MA Innis,
DH Gelfand, JJ Sninsky, TJ White, eds. PCR protocols: a guide to methods
and application. San Diego: Academic Press, 315–322.
Lee et al. — Associations with saprotrophic fungi among fully mycoheterotrophic orchids
Wilson AW, Desjardin DE. 2005. Phylogenetic relationships in the gymnopoid
and marasmioid fungi (Basidiomycetes, euagarics clade). Mycologia 97:
667–679.
Yamato M, Yagame T, Suzuki A, Iwase K. 2005. Isolation and identification
of mycorrhizal fungi associating with an achlorophyllous plant, Epipogium
roseum (Orchidaceae). Mycoscience 46: 73–77.
Yang CK, Lee YI, Chen YF, Chung NJ, Wang YN, Chung KF. 2010. Study
on species diversity and conservation of wild orchids at Xitou Tract
Experimental Forest of National Taiwan University. Journal of the
Experimental Forest of National Taiwan University 24: 123–136. (in Chinese)
Yang Z, Rannala B. 1997. Bayesian phylogenetic inference using DNA sequences: a Markov chain Monte Carlo method. Molecular Biology and
Evolution 14: 717–724.
435
Yeung EC. 1984. Histological and histochemical staining procedures. In: IK
Vasil, ed. Cell culture and somatic cell genetics of plants. Orlando:
Academic Press, 689–697.
Zimmer K, Hynson NA, Gebauer G, Allen EB, Allen MF, Read DJ. 2007.
Wide geographical and ecological distribution of nitrogen and carbon gains
from fungi in pyroloids and monotropoids (Ericaceae) and in orchids. New
Phytologist 175: 166–175.
Zimmer K, Meyer C, Gebauer G. 2008. The ectomycorrhizal specialist orchid
Corallorhiza trifida is a partial myco-heterotroph. New Phytologist 178:
395–400.