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OPEN
ORIGINAL ARTICLE
Functional analysis of liverworts in dual symbiosiswith
Glomeromycota and Mucoromycotina fungiunder a simulated Palaeozoic
CO2 decline
Katie J Field1, William R Rimington2,3,4, Martin I
Bidartondo2,3, Kate E Allinson5,David J Beerling5, Duncan D
Cameron5, Jeffrey G Duckett4, Jonathan R Leake5 andSilvia
Pressel41School of Biology, Faculty of Biological Sciences,
University of Leeds, Leeds, UK; 2Department of LifeSciences,
Imperial College London, London, UK; 3Jodrell Laboratory, Royal
Botanic Gardens, Kew, UK;4Department of Life Sciences, Natural
History Museum, London, UK and 5Department of Animal and
PlantSciences, Western Bank, University of Sheffield, Sheffield,
UK
Most land plants form mutualistic associations with arbuscular
mycorrhizal fungi of theGlomeromycota, but recent studies have
found that ancient plant lineages form mutualisms
withMucoromycotina fungi. Simultaneous associations with both
fungal lineages have now been found insome plants, necessitating
studies to understand the functional and evolutionary significance
ofthese tripartite associations for the first time. We investigate
the physiology and cytology of dualfungal symbioses in the
early-diverging liverworts Allisonia and Neohodgsonia at modern
andPalaeozoic-like elevated atmospheric CO2 concentrations under
which they are thought to haveevolved. We found enhanced carbon
cost to liverworts with simultaneous Mucoromycotina
andGlomeromycota associations, greater nutrient gain compared with
those symbiotic with only onefungal group in previous experiments
and contrasting responses to atmospheric CO2 amongliverwort–fungal
symbioses. In liverwort–Mucoromycotina symbioses, there is
increased P-for-C andN-for-C exchange efficiency at 440 p.p.m.
compared with 1500 p.p.m. CO2. In liverwort–Glomeromy-cota
symbioses, P-for-C exchange is lower at ambient CO2 compared with
elevated CO2. Nocharacteristic cytologies of dual symbiosis were
identified. We provide evidence of a distinctphysiological niche
for plant symbioses with Mucoromycotina fungi, giving novel insight
into whydual symbioses with Mucoromycotina and Glomeromycota fungi
persist to the present day.The ISME Journal (2016) 10, 1514–1526;
doi:10.1038/ismej.2015.204; published online 27 November 2015
Introduction
Symbioses with soil fungi have existed since plantsfirst began
to colonize the Earth’s land masses(Redecker et al., 2000; Redecker
and Raab, 2006;Smith and Read, 2008) and are thought to have
playeda key role in establishing terrestrial ecosystems(Pirozynski
and Malloch, 1975; Malloch et al., 1980).There are numerous lines
of supporting evidence forthis view, including plant and fungal
fossils(Stubblefield et al., 1987; Remy et al., 1994; Tayloret al.,
1995) and molecular data (Simon et al., 1993;Redecker et al., 2000;
Redecker and Raab, 2006).Recent studies of ultrastructure (Pressel
et al., 2010)and plant–fungal physiology of early-diverging
extant
land plant lineages (Field et al., 2012, 2015a) providednew
insights into the structure–function relationshipsof non-vascular
plants and their symbiotic fungi. Untilrecently, the fungal
associates of the earliest branchingplant lineages have been
assumed to be members of thearbuscular mycorrhiza-forming clade of
fungi, theobligately biotrophic Glomeromycota that lack
sapro-trophic capabilities.
Application of universal DNA primers, enablingdetection of fungi
beside Glomeromycota, togetherwith detailed physiological and
cytological observa-tions, have now established that the
earliestbranching lineage of extant liverworts, the
Haplo-mitriopsida (Heinrichs et al., 2005, 2007; Crandall-Stotler
et al., 2009; Wikström et al., 2009), often formmutualistic
mycorrhiza-like associations exclusivelywith Mucoromycotina fungi
(Bidartondo et al., 2011;Field et al., 2015a). This partially
saprotrophicfungal lineage is basal or sister to the
Glomeromycota(James et al., 2006; Lin et al., 2014), raising
thehypothesis that plant–Mucoromycotina associations
Correspondence: KJ Field, School of Biology, Faculty of
BiologicalSciences, University of Leeds, Miall Building, Leeds LS2
9JT, UK.E-mail: [email protected] 23 March 2015; revised
8 October 2015; accepted 12October 2015; published online 27
November 2015
The ISME Journal (2016) 10, 1514–1526© 2016 International
Society for Microbial Ecology All rights reserved 1751-7362/16
www.nature.com/ismej
http://dx.doi.org/10.1038/ismej.2015.204mailto:[email protected]://www.nature.com/ismej
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represent the ancestral mycorrhizal type for landplants and that
these were replaced by the strictlybiotrophic Glomeromycota as
plants evolved andsoil organic matter accumulated (Bidartondo et
al.,2011). Although some early branching clades of landplant taxa
have been found to associate exclusivelywith one or other of these
fungal groups, representa-tives of nearly all extant early
branching clades ofland plants examined thus far host both
fungallineages, sometimes simultaneously (Desirò et al.,2013;
Rimington et al., 2015, and a recent report forHaplomitrium
mnioides by Yamamoto et al., 2015)(Figure 1a). These discoveries
point to more versatileand shifting evolutionary scenarios in early
plant–fungus symbioses than hitherto assumed (Field et al.,2015b),
suggesting that the ability to engage insimultaneous partnerships
with both Mucoromyco-tina and Glomeromycota fungi may be a basal
trait(Desirò et al., 2013; Rimington et al., 2015).
The latest evidence (Field et al., 2015a) shows
thatliverwort–Mucoromycotina symbioses functionallydiffer from
those between liverworts and Glomer-omycota fungi (Field et al.,
2012) in their ability tomaintain efficiency of carbon-for-nutrient
exchangebetween partners across atmospheric CO2 concentra-tions
(a[CO2]). The conditions in these studiessimulate the 90% a[CO2]
drop coincident with thediversification of terrestrial ecosystems
through thePalaeozoic (Berner, 2006; Franks et al., 2014).Although
the symbiotic functional efficiency ofliverwort–Glomeromycota
associations was severelycompromised by a simulated Palaeozoic fall
ina[CO2], that of Haplomitriopsida liverwort–Mucor-omycotina
partnerships was unaffected or increasedunder the modern-day a[CO2]
scenario. These find-ings parallel those in
Glomeromycota-associatedsporophytes of some vascular plants, which
alsoincreased in functional efficiency under lowera[CO2] (Field et
al., 2012). Therefore, the hypothesisthat Mucoromycotina fungi,
switching from sapro-trophy to facultative biotrophy, facilitated
the evolu-tion and diversification of early land plants under ahigh
a[CO2] and were among the first fungi to formmutualistic symbioses
with plants is strengthened(Bidartondo et al., 2011). It remains an
open questionas to why dual
Mucoromycotina/Glomeromycotaplant–fungus partnerships today are
often restrictedto early-branching lineages of land plants (Figure
1a)and thus ‘lost out’ to Glomeromycota-specific ones
inlater-branching plant lineages, such as the angios-perms (Field
et al., 2015b).
We investigated the functionality and detailedcytology of the
dual fungal associations in wild-collected Neohodgsonia mirabilis,
the sister taxon toall other complex thalloid liverworts
harbouringmycorrhiza-like associations, and Allisonia cockay-nei in
the earliest divergent clade of simple thalloidliverworts (Forrest
et al., 2006; Crandall-Stotler et al.,2008, 2009; Villarreal et
al., 2015) (Figure 1).Using molecular methods, we found that
bothliverworts hosted simultaneous Glomeromycota
and Mucoromycotina fungal partners (see Results).We used a
combination of isotope tracers under amodern ambient a[CO2] of 440
p.p.m. and a simu-lated Palaeozoic (c. 410–390Ma) atmosphere of1500
p.p.m. [CO2] (Franks et al., 2014).
We aimed to answer the following questions;
(1) Is there an enhanced carbon cost to liverwortsassociated
simultaneously with Mucoromyco-tina and Glomeromycota fungi
compared withthose harbouring single fungal symbionts?
(2) Do plants with dual colonization by Mucoro-mycotina and
Glomeromycota fungi benefitfrom enhanced nutrient gain in
comparison tothose harbouring single fungal associations?
(3) Are the costs decreased and benefits increasedby elevated
a[CO2] for liverworts maintainingdual symbioses with both
Mucoromycotina andGlomeromycota fungi?
(4) Are there any characteristic cytological signa-tures of dual
fungal symbiosis as opposed tosingle fungal associations?
Haplomitriopsida
Blasiales
Sphaerocarpales
Neohodgsonia
LunulariaPreissia
Marchantia
Allisonia
Pellia
Pallavicinaceae
Pleurozia
Aneura
Porellales
Cyanobacteria
Jungermanniales
Pel
lidae
Mar
chan
tiops
ida
None
Mucoromycotina
Glomeromycota
Basidiomycota
Ascomycota
Met
zger
idae
or Fungal associates
Figure 1 Liverwort phylogeny and species used in the
presentstudy. (a) Liverwort phylogeny (following Wikström et al.,
2009)showing key nodes alongside commonly associated
fungalsymbionts (James et al., 2006; Pressel et al., 2008;
Bidartondoand Duckett, 2010; Humphreys et al., 2010; Pressel et
al., 2010;Bidartondo et al., 2011; Field et al., 2012; Desirò et
al., 2013).Plants of (b) Allisonia cockaynei and (c) Neohodgsonia
mirabilisphotographed in the field (photo credits: KJ Field andJG
Duckett).
Dual fungal symbioses in thalloid liverwortsKJ Field et al
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Materials and methods
Plant material and growth conditionsThe liverworts Neohodgsonia
mirabilis (Perss.)Perss. and Allisonia cockaynei (Steph.) RM
Schust.were collected from the South Island of New Zealandin April
2012, and vouchers were deposited in theNatural History Museum,
London. We planted theliverworts directly into pots (120mm diameter
× 100mm depth) soon after collection. Native soil sur-rounding
liverwort rhizoids was left intact to act as anatural inoculum, and
pots were carefully weededregularly to remove any other plant
species.
Based on the methods of Johnson et al. (2001), weinserted three
mesh-windowed cylindrical cores(Supplementary Figure S1) into each
experimentalpot. The mesh covering the cores was fine enough
toexclude liverwort rhizoids but allows the ingrowth offungal
hyphae. Two of the cores were filled with ahomogeneous mixture of
acid-washed silica sand(89% core volume), native soil gathered from
aroundthe rhizoids of wild plants (10% core volume) andfine-ground
tertiary basalt (1% core volume) to act asfungal bait (Field et
al., 2012). The third core was filledwith glass wool and enabled
below-ground respirationsampling throughout the 14C-labelling
period.
We maintained plants in controlled environmentchambers (BDR16,
Conviron, Winnipeg, MB, Canada)with settings chosen according to
those of the plant’snatural environment (see Supplementary
Information).Each species was grown at either 440p.p.m.a[CO2]
(n=10) or at a simulated early-Palaeozoica[CO2] concentration of
1500p.p.m. (n=10) (Berner,2006, Franks et al., 2014). a[CO2] was
monitoredusing CARBOCAP GMP343 CO2 sensors (Vaisala,Birmingham, UK)
and maintained through addition ofgaseous CO2. Cabinet settings and
contents werealternated every 2 weeks, and we regularly rotated
allpots within cabinets. Plants were acclimated to cham-ber/growth
regimes for 12 weeks to allow establishmentof mycelial networks
within pots.
Molecular identification of fungal associatesWild Neohodgsonia
and Allisonia thalli were pre-pared for molecular analysis within 1
day ofcollection and immediately following our isotopelabelling
experiments at the end of the growth periodat different a[CO2]. We
dissected both plant speciesin the same way to leave the central
part of thethallus and rhizoidal ridge (2–3mm2) where
fungalcolonization is the highest. The DNA extraction,amplification
and sequencing were performed as perthe methods of Gardes and Bruns
(1993), Desirò et al.(2013) and Field et al. (2015a) (see
SupplementaryInformation). Sequence identity was inferred fromtheir
most closely related BLAST hits (Altschulet al., 1997). Bayesian
inference was used to confirmthe fungal identity of samples shown
to be Glomer-omycota or Mucoromycotina by BLAST. Sequenceswere
aligned with reference DNA sequences fromGenBank (Benson et al.,
2005) using MUSCLE
alignment algorithms (Edgar, 2004) within MEGAv. 5.1 (Tamura et
al., 2011). We tested evolutionarymodels in MEGA and selected HKY85
(nst = 2) withinvgamma rates for Bayesian analysis using
MrBayes(Huelsenbeck and Ronquist, 2001).
Quantification of fluxes of C, 33P and 15N betweenliverworts and
fungiAfter the 12-week acclimation period, we introduced100 μl of
an aqueous mixture of 33P-labelled ortho-phosphate (specific
activity 148 GBqmmol −1, total111 ng 33P added) and 15N-ammonium
chloride(1mgml− 1) into one of the soil-filled mesh cores ineach
pot and 100 μl distilled water into the controlcore via the
installed capillary tubes. Cores in whichisotope tracers were
introduced were left static inhalf of the pots to preserve direct
hyphal connectionswith the liverworts. In the remaining half,
labelledcores were rotated through 90°, severing the
hyphalconnections between the plants and core soilimmediately prior
to addition of isotopes and everyother day thereafter
(Supplementary Figure S2).
We sealed the top of all soil cores with lanolin andcaps 21 days
after addition of the isotope tracers. Glasswool-filled cores were
sealed with a rubber septum(SubaSeal, Sigma). We then sealed each
pot into a 3-l,gas-tight labelling chamber and added 2ml 10%
lacticacid to 15 μl Na14CO3 (specific activity 2.04 TBq-mmol−1) in
a cuvette within the chamber prior toillumination at 0700 hours.
This resulted in the releaseof a 1.1-MBq pulse of 14CO2 gas. Pots
were maintainedunder growth chamber conditions, and 1ml
oflabelling-chamber headspace gas was sampled after1 h and every 4
h thereafter. Below-ground gas wassampled via the glass-wool filled
core after 1 h andevery 2 h thereafter to monitor below-ground
respira-tion and 14C flux for around 17 h (see
SupplementaryInformation for further details).
Plant harvest and sample analysesPlant and soil materials were
separated, freeze-dried,weighed and homogenized. In all, 10–30mg
ofhomogenized samples were digested in 1ml ofconcentrated H2SO4.
These were heated to 365 °Cfor 15min, and 100 μl H2O2 was added to
eachsample when cool. Samples were reheated to 365 °C,and each
clear digest solution was diluted to 10mlwith distilled water. Two
ml of each diluted digestwere then added to 10ml of the
scintillation cocktailEmulsify-safe (Perkin Elmer, Beaconsfield,
UK) andquantified through liquid scintillation. 33P trans-ferred to
the plant via fungal mycelium was thencalculated as detailed in
Supplementary Information(Cameron et al., 2007).
15N abundance was determined using IsotopeRatio Mass
Spectrometry (IRMS). Between 2 and5mg of freeze-dried, homogenized
plant tissue wasweighed out into 6× 4mm2 tin capsules (Sercon
Ltd,Crewe, UK) and analysed using a continuous flowIRMS (PDZ 2020
IRMS, Sercon Ltd). Air was used as
Dual fungal symbioses in thalloid liverwortsKJ Field et al
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the reference standard, and the IRMS detector wasregularly
calibrated to commercially availablereference gases.
14C activity was quantified through sample oxida-tion and liquid
scintillation. Approximately 10–100mg of freeze-dried sample was
placed inCombusto-cones (Perkin Elmer) before oxidation(Model 307
Packard Sample Oxidiser Isotech, Ches-terfield, UK). CO2 released
through oxidation wastrapped in 10ml Carbosorb prior to mixing
with10ml Permafluor. Total carbon (12C+14C) fixed by theplant and
transferred to the fungal network wascalculated as a function of
the total volume and CO2content of the labelling chamber and the
proportionof the supplied 14CO2 label fixed by the plants.
Thedifference in carbon between the static and rotatedcores is
taken as equivalent to the total C transferredfrom plant to
symbiotic fungus within the soil core,noting that a small
proportion will be lost throughsoil microbial respiration. The
total carbon budgetfor each experimental pot was calculated
usingequations from Cameron et al. (2006), which aredetailed in
Supplementary Information.
Data from Allisonia and Neohodgsonia are com-pared in the
discussion to published and unpub-lished data from Haplomitrium and
Treubiaassociated exclusively with Mucoromycotina fungiobtained
from experiments using identical condi-tions within the same
controlled environmentgrowth chambers (see Field et al., 2015a).
Data arealso presented alongside previously published datafor
Preissia and Marchantia associated only withGlomeromycota fungi
from experiments using near-identical experimental conditions
within the samecontrolled environment growth chambers (Fieldet al.,
2012). In these experiments, pots were filledwith soil from dune
slacks at Aberfraw, Anglesey,UK (Grid Reference: SH 397 648) but
were otherwiseidentical to those of all our other experiments.
Ultrastructural analysesWe processed plants that were
wild-collected andfrom experiments where they were grown at
twoa[CO2] for transmission and scanning electron
microscopy as described previously (Duckett et al.,2006). For
transmission electron microscopy, thalliwere fixed in 3%
glutaraldehyde, 1% fresh formal-dehyde and 0.75% tannic acid in
0.05 M Na-cacodylate buffer, pH 7, for 3 h at room
temperature.After rinses in 0.1 M buffer, samples were postfixedin
buffered (0.1 M, pH 6.8) 1% osmium tetroxideovernight at 4 °C,
dehydrated in an ethanol seriesand embedded in TAAB low viscosity
resin viaethanol. Thin sections were cut with a diamondknife,
stained with methanolic uranyl acetate for15min and in Reynolds’
lead citrate for 10min andobserved with a Hitachi H-7100
transmission elec-tron microscope (Hitachi High-Technologies
Europe,Maidenhead, UK) at 100 kV. For scanning electronmicroscopy,
we fixed thalli in 3% glutaraldehyde,dehydrated through an ethanol
series, critical-pointdried using CO2 as transfusion fluid, sputter
coatedwith 390 nm palladium-gold and viewed themusing a FEI Quanta
scanning electron microscope(FEI, Hillsboro, OR, USA).
StatisticsEffects of plant species, a[CO2] and the
interactionbetween these factors on the C, 33P and 15N
fluxesbetween plants and fungi from this and previousstudies (Field
et al., 2012, 2015a) were tested usinganalysis of variance with
additional post-hoc Tukey'stests where indicated. Data were checked
for homo-geneity of variance and normality. Where assumptionsfor
analysis of variance were not met, data weretransformed using log10
or arcsine-square-root asindicated in Table 1. Different letters in
the figuresdenote statistical difference (Po0.05) in all the
figures.All statistics were carried out using the
statisticalsoftware package R 3.1.2 (R Core Team, 2012).
ResultsMolecular identification of fungiMolecular analyses of
fungal partners (n=6) showedthat Allisonia and Neohodgsonia plants
freshlycollected from the field and after our isotope tracing
Table 1 Summary of differences in mycorrhizal functionality (F
ratio from ANOVA) between Neohodgsonia, Alisonia,
Haplomitrium,Treubia, Preissia and Marchantia at elevated a[CO2]
(1500 p.p.m.) and ambient a[CO2] (440 p.p.m.)
df Plant species CO2 treatment Species×CO2
Biomass (g) 1, 30 16.276*** 18.911*** 1.937Fungal carbon in
cores (ng)a 1, 54 14.042*** 31.334*** 5.087***Percentage of carbon
allocationb 1, 54 5.756*** 13.900*** 3.278*Total 33P uptake (ng)b
1, 30 4.498** 5.714* 6.483***[33P] in plant tissue (ng g−1) 1, 36
6.259*** 3.142 3.857*Total 15N uptake (ng) 1, 20 1.889 0.953
0.822[15N] in plant tissue (ng g− 1)a 1, 20 0.235 1.147
1.30433P-for-C efficiency (ng ng−1) 1, 36 46.220*** 0.885
31.747***15N-for-C efficiency (ng ng− 1) 1, 20 0.413 13.523**
1.913
Abbreviations: ANOVA, analysis of variance; p.p.m., parts per
million. *Po0.05, **Po0.01, ***Po0.001; post-hoc Tukey's test.aData
have been log10 transformed to meet the assumptions for ANOVA.bData
have been arcsine-square-root transformed to meet the assumptions
for ANOVA.
Dual fungal symbioses in thalloid liverwortsKJ Field et al
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experiments are colonized by both Mucoromycotinaand
Glomeromycota fungi (Supplementary FigureS3). The Mucoromycotina
fungi identified here werethe same as those found previously in
wild popula-tions of both species (Bidartondo et al.,
2011)belonging to groups I and H (sensu, Desirò et al.,2013) in
Neohodgsonia and Allisonia, respectively.The Glomeromycota fungal
associates were exclu-sively Glomerales in Allisonia while
Neohodgsoniaharboured members of both Glomerales and
Archae-osporales. Sequences are deposited in
GenBank(KR779272-KR7792784).
Plant biomassOverall, there was a consistent trend of
liverwortsachieving greater biomass when grown at a[CO2] of1500
p.p.m. compared with 440 p.p.m. a[CO2](Figure 2). We found greater
biomass of bothAllisonia (41%) and Neohodgsonia (45%) grown at1500
p.p.m. a[CO2] compared with those grown ata[CO2] of 440 p.p.m.
Liverwort-to-fungus carbon transferBoth Allisonia and
Neohodgsonia allocated aroundfour times more photosynthate to their
fungalsymbionts under the simulated Palaeozoic a[CO2](Figure 3a)
compared with the lower [CO2](Figure 3a). In terms of total carbon
transferred fromplants to fungal partners (Figure 3b), each
liverwortspecies transferred more carbon to their fungalsymbionts
at 1500 p.p.m. a[CO2] than at 440 p.p.m.a[CO2], this difference
being significant in Allisoniaand Neohodgsonia. As such, the dual
fungal sym-bioses of Neohodgsonia and Allisonia have a greatertotal
carbon ‘cost’ at both a[CO2] than any of the
other Glomeromycota– or Mucoromycotina–liver-wort symbioses
(Figure 3b).
Fungal transfer of 33P and 15N to host liverwortsAllisonia and
Neohodgsonia acquire 78% and 67%more 33P, respectively, at 440
p.p.m. compared with1500 p.p.m. a[CO2], also reflected in plant
tissue [33P](Figures 3c and d). When grown at the 1500
p.p.m.a[CO2], the liverworts with dual fungal symbiontsshowed
reduced total 33P uptake (Figure 3c), result-ing in greatly reduced
33P concentrations in theirtissues (Figure 3d).
The total uptake and assimilation of 15N is reducedby 11% in
Allisonia and 57% in Neohodgsonia at1500 p.p.m. a[CO2] compared
with 440 p.p.m. a[CO2](Figure 3e). In terms of tissue
concentration, the sametrend is amplified with [15N] being far
greater by 250%and 119% in Allisonia and Neohodgsonia,
respec-tively, at 440 p.p.m. a[CO2] compared with whenplants are
grown at 1500 p.p.m. a[CO2] (Figure 3f).
Nutrient-for-carbon exchange efficiency33P-for-C exchange
efficiency in Allisonia was 413times greater at 440 p.p.m. a[CO2]
than it was at1500 p.p.m. a[CO2] (Figure 4a). The same pattern
wastrue in Neohodgsonia, with three times greater 33P-for-C
exchange efficiency at the lower a[CO2](Figure 4a). 15N-for-C
exchange was an order ofmagnitude greater in both Allisonia and
Neohodgso-nia at the lower a[CO2] compared with the elevateda[CO2]
(Figure 4b).
Cytology of colonizationThe cytology of dual colonization by
Mucoromycotinaand Glomeromycota fungi in wild plants of
Neohodg-sonia and Allisonia is described here for the first time.As
our detailed electron microscopic analyses revealedno major
differences, only a minor one in Allisonia(detailed below), between
wild and experimental plantsgrown at contrasting a[CO2] (440 and
1500p.p.m.), theresults are presented together unless otherwise
stated.
Neohodgsonia mirabilisFungal colonization occupies the central
thallusmidrib, extending all the way from the rhizoid-bearing
ventral surface, the point of fungal entry (seeSupplementary
Information), to just below the largedorsal air chambers (Figure
5a). Fungal structurescomprise numerous arbuscules at various
stages ofdevelopment, from young (Figure 5b) to collapsedand large
vesicles occupying a significant proportionof the host cell (Figure
5c). Healthy (Figure 5d) anddegenerated arbuscules (Figure 5e),
large livinghyphae, vesicles and active host cytoplasm are
mostoften present in the same host cell. Fungal trunkhyphae and
arbuscular hyphae are surrounded bythe host plasma membrane, and
the cytoplasm of thehost cells comprises numerous Golgi bodies,
Haplomitrium
Treubia
Preissia
Marchantia
Allisonia
Neohodgsonia
0
5
10
15
20
Bio
mas
s (n
g)
1,500 ppm a[CO2] 440 ppm a[CO2]
Mucoromycotina Glomeromycota Dual associations
a
a
abab abc
bcbc bc
c
ab
Figure 2 Mean total plant biomass (dry) at the end of
experi-mental period in five liverwort species (Field et al., 2012,
2015a) atboth 1500 p.p.m. a[CO2] (black bars) and 440 p.p.m. a[CO2]
(whitebars). Error bars show s.e.m. (n=4 for all species);
different lettersdenote statistical difference where Po0.05
(Tukey's post hoc).
Dual fungal symbioses in thalloid liverwortsKJ Field et al
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mitochondria, plastids and microbodies (Figures 5dand e). The
latter have a well-developed thylakoidsystem but are largely devoid
of starch deposits(Figure 5d). Fungal hyphae are aseptate and
oftencontain multiple mitochondrial stacks, each com-prising of 5–6
mitochondria (Figure 5f).
Allisonia cockayneiFungal entry is via the rhizoids
(Supplementary FigureS4) with the fungal zone occupying the central
regionof the thallus, generally the first 10 cell layers from
therhizoid-bearing ventral side with approximately 1/3 ofthe
thallus midrib remaining free of fungal structures(Figure 6a).
These comprise large hyphae, arbuscules(Figure 6b) and prominent
vesicles (Figure 6c). Hostcells are characterised by active
cytoplasm, including
numerous mitochondria and plastids in close associa-tion with
the fungus (Figure 6d). Colonizing hyphaetraverse the walls of
adjacent host cells and have athick layer of fibrillar material in
between the funguscell wall and the host plasma membrane
thatsurrounds them (Figure 6d) while arbuscular hyphaeare
characterized by thin cell walls (Figure 6e). Theseare often
collapsed while the colonizing hyphae andhost cytoplasm surrounding
them persist (Figure 6f).Whereas the plastids of wild plants and
those grown at440 p.p.m. a[CO2] contain little or no starch
deposits(Figure 6g), those of plants grown at 1500 p.p.m.
a[CO2]have prominent starch grains (Figure 6h). The largecolonizing
hyphae of wild and experimental plantsgrown under contrasting
a[CO2] regimes are allcharacterized by plasmodesmata-like channels
in thefibrillar material that surrounds them (Figure 6i).
1
10
100
1000
10000
Fung
al c
arbo
n in
cor
es (n
g)
Mucoromycotina Glomeromycota Dual association
ab b b
ab
d
c c
dde de
e
0.01
0.1
1
% C
allo
catio
n to
fung
us in
cor
e Mucoromycotina Glomeromycota Dual colonisationa
ababc
abcb
bc
cdd d
cdcd bc
0
5
10
15
20
25
Plan
t tis
sue
[33 P
] (n
g g-
1 )
Mucoromycotina Glomeromycota Dual association
a
ab
a
bbbbab
b
cc ccc
0.001
0.01
0.1
1
10
100
1000
Plan
t tis
sue
33P
(ng)
Mucoromycotina Glomeromycota Dual association
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Treubia
Preissia
Marchantia
Allisonia
Neohodgsonia
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Mucoromycotina Glomeromycota Dual colonisation
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ababab
1,500 ppm a[CO2] 440 ppm a[CO2]
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Neohodgsonia
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Figure 3 Carbon-for-nutrient exchange between liverworts and
their fungal partners. (a) Percentage allocation of plant-derived
carbon tofungi within soil cores, (b) total measured plant-fixed
carbon transferred to fungi in soil for liverworts with different
fungal associations(Mucoromycotina-only, Glomeromycota-only and
dual fungal associations; Field et al., 2012, 2015a); (c) total
plant tissue 33P content (ng) and(d) tissue concentration (ng g−1)
of fungal-acquired 33P in six liverwort species with different
fungal associations under 1500p.p.m. (blackbars) a[CO2] and 440
p.p.m. (white bars) a[CO2] (Field et al., 2012, 2015a); (e) total
tissue 15N content (ng) and (f) concentration (ng g−1)
offungal-acquired 15N in four liverwort species with different
fungal associations (Field et al., 2015a) at both 1500 p.p.m.
(black bars) and 440 p.p.m. (white bars) a[CO2]. In all panels,
error bars show ±s.e.m. Different letters represent where Po0.05
(analysis of variance, Tukey's posthoc; see Table 1). In panels (a)
and (b), n=6 for all species apart from Marchantia, where n=4;
(c–e) n=4, where data are available.
Dual fungal symbioses in thalloid liverwortsKJ Field et al
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Discussion
The currently emerging paradigm considers the Mucor-omycotina
symbiosis with plants to have evolved priorto the emergence of
plant–Glomeromycota fungalsymbioses (Bidartondo et al., 2011).
Moreover, untilvery recently it has been assumed that early
diverginglineages of plants associate with only Glomeromycota(Wang
and Qui, 2006). In direct contrast to this, ourwork shows that
basal liverwort lineages (Figure 1a)form simultaneous mutualistic
symbioses with bothMucoromycotina and Glomeromycota fungi.
Thisraises novel questions regarding mycorrhizal evolution;given
the global radiation and dominance of glomer-omycotean symbioses,
why have associations withMucoromycotina fungi persisted? We can
now beginto answer this question with the present demonstrationthat
dual associations are significantly more efficient atmodern day
atmospheric CO2 compared with Palaeo-zoic CO2, whereas single
fungal group partnerships areeither unaffected by a[CO2]
(Mucoromycotina fungi) orare less efficient under modern day a[CO2]
(Glomer-omycota fungi). This trade-off provides a
physiologicalniche facilitating the persistence of plant
symbioseswith Mucoromycotina fungi, singly and in dualpartnerships
with Glomeromycota fungi to thepresent day.
Physiological costs and benefitsOur experiments reveal that
Neohodgsonia andAllisonia with dual Glomeromycota and
Mucoromy-cotina fungal associations allocated greater percen-tages
and total amounts of photosynthate to theirfungal partners at 1500
p.p.m. a[CO2] than at 440 p.p.m. a[CO2] (Figures 3a and b). Our
previous studiesshow that in terms of percentage of carbon
alloca-tion, Mucoromycotina partners of Treubia receiveseven times
greater percentage allocation of plant-fixed carbon at 1500 p.p.m.
a[CO2] compared with at440 p.p.m. a[CO2]. There is little
difference inpercentage of C allocation in Haplomitrium whilein
Marchantia and Preissia the percentage of Callocation to
Glomeromycota fungi is 1.9 and 1.2times greater, respectively. This
likely resulted in thegreater biomasses recorded in all liverworts
atelevated a[CO2] (Figure 2).
In all of the combinations of liverwort–fungalsymbioses examined
thus far, partnerships in whichthere is a Mucoromycotina fungal
symbiont (that is, inHaplomitrium, Treubia, Allisonia and
Neohodgsonia)display increased 33P-for-C and 15N-for-C
exchangeefficiency at 440 p.p.m. a[CO2] compared with at1500 p.p.m.
a[CO2] (Figure 4). In liverwort–Glomer-omycota symbioses, the
opposite trend is apparent,with 33P-for-C being several orders of
magnitudelower in both Marchantia and Preissia at 440
p.p.m.compared with at 1500 p.p.m. a[CO2] (Figure 4a).
Decreased fungal-acquired nutrient uptake inliverworts with
Mucoromycotina fungal partners(either single or dual colonizations)
at elevated a[CO2] seems at first counter-intuitive,
particularlygiven their larger biomass (Figure 2) and
increasedphotosynthate allocation to fungal partners(Figures 3a and
b) in those conditions. However, itis possible that the plants in
our experimental potsexperienced nutrient limitation (for P, N or
both).This seems likely considering the lack of plant-available
nutrients in the surrounding sand and itslimited accessibility
within the soil cores. As such,when a[CO2] is at 1500 p.p.m., the
liverworts likelyproduced excess photosynthates that they mighthave
been unable to utilize for growth or reproduc-tion owing to
nutrient limitation. As liverworts arestructurally simple plants,
with no vasculature orspecialized storage organs to provide
transport andstorage of excess carbohydrates (Kenrick and
Crane,1997), surplus sugars must be either stored asinsoluble
starch granules within the thallus(observed here in Allisonia;
Figure 6h), supplieddirectly to fungal partner(s) (see Figures 3a
and b), orbe released into the surrounding soil as exudates.
It is likely that the greater C allocation we observedfrom
liverworts to Mucoromycotina fungal partners inour experiments
allows increased hyphal prolifera-tion and fungal sporulation.
Given that these pro-cesses are demanding in terms of energy
andresources (Denison and Kiers, 2011), the funguswould have
greater N and P requirements andtherefore may assimilate more of
the nutrients
0.0001
0.001
0.01
0.1
133
P-fo
r-C
effi
cien
cy (
ng n
g-1 )
Mucoromycotina Glomeromycota Dual associations
1,500 ppm a[CO2] 440 ppm a[CO2]
a aaa a
cc
c c
d
c
a
Haplomitrium
Treubia
Preissia
Marchantia
Allisonia
Neohodgsonia
0.0001
0.001
0.01
0.1
1
15N
-for-
C e
ffici
ency
(ng
ng-
1 ) Mucoromycotina Glomeromycota Dual associations
N/A
a a
bb
abab
abab
Figure 4 Nutrient-for-carbon exchange efficiencies
betweenliverworts and their fungal partners. (a) 33P-for-carbon and
(b)15N-for-carbon efficiency for different liverwort species
withdifferent fungal associations under both 1500 p.p.m. (black
bars)and 440 p.p.m. (white bars) a[CO2] (Field et al., 2012,
2015a,b).Error bars show s.e.m. (n=4 for all species). Different
lettersindicate where Po0.05 (analysis of variance, Tukey's post
hoc).
Dual fungal symbioses in thalloid liverwortsKJ Field et al
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Figure 5 Cytology of Neohodgsonia mirabilis grown at 440 and
1500 p.p.m. a[CO2]. Scanning (a–c) and transmission (d–f)
electronmicrographs (TEM). Both the distribution and cytology of
the association remained the same between a[CO2] treatments and are
illustratedhere in plants grown at 440 p.p.m. a[CO2]. (a) Fungal
colonization zone extending from the rhizoid (R) bearing ventral
surface of the thallusto just below the dorsal air chambers (AC).
(b, c) Young arbuscules (b) and collapsed ones (c) (*) adjacent to
a large vesicle (arrowed). (d)fungal hyphae surrounded by active
host cytoplasm. Note the plastids (P) with well-developed thylakoid
systems but largely devoid ofstarch. (e) Degenerated arbuscular
hyphae (*) surrounded by healthy host cytoplasm. (f) Fungal hyphae
typically contain multiplemitochondrial stacks (M). Scale bars: (a)
200 μm; (c) 50 μm; (b) 20 μm; (d) 3 μm; (e, f) 1 μm.
Dual fungal symbioses in thalloid liverwortsKJ Field et al
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Figure 6 Cytology of Allisonia cockaynei grown at 440 and 1500
p.p.m. a[CO2]. Scanning (a–c) and transmission (e–i) electron
micrographs(TEM). There was no change in the overall distribution
of fungal colonization and in the cytology of the fungus between
[CO2] treatments, bothillustrated here in plants grown at 440
p.p.m. a[CO2] except for panels (h and i). (a) Fungal colonization
zone (arrowed) occupying the first 10cell layers from the
rhizoid-bearing ventral surface. (b) Collapsed arbuscules (arrowed)
and (c) large vesicle (arrowed). (d) Host cell with activecytoplasm
in close association with fungal hyphae (H). Note the colonizing
hypha (CH) traversing the host cell wall. N, nucleus; OB, oil body.
(e)colonizing hypha with thick layer of fibrillar material (*) in
between the fungus cell wall and the host plasma membrane (arrowed)
and thin-walled arbuscular hyphae (AH) in close proximity to
plastids (P). (g) Arbuscular hyphae in close association with
starch-free plastids. (h) Inplants grown at 1500 p.p.m. a[CO2]
plastids have prominent starch deposits. (i) Plasmodesmata-like
channels are present in the fibrillar materialthat surrounds the
colonizing hyphae. Scale bars: (a) 200μm; (b, c) 20μm; (d–i)
3μm.
Dual fungal symbioses in thalloid liverwortsKJ Field et al
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acquired from its surroundings, rather than surrenderthem in
return for plant carbohydrate. This mayprovide a mechanism to
explain our observations ofreduced fungal-acquired nutrient uptake
inMucoromycotina-exclusive and dual
Mucoromyco-tina/Glomeromycota-partnered liverworts at
elevateda[CO2], even with enhanced C allocation to fungalpartners
(Figures 4 and 5). It is also possible that thereare further
non-nutritional benefits for liverworts insymbiosis with
Mucoromycotina fungi that have notbeen explored here, such as
enhanced disease and/orherbivore resistance (Cameron et al.,
2013).
In contrast, the liverworts partnered exclusivelywith obligately
biotrophic Glomeromycota fungi inprevious experiments (that is,
Marchantia and Pre-issia in Field et al., 2012) operated a more
linearexchange of nutrients-for carbon. In this scenario,more
photosynthate is supplied to the fungal myce-lium at elevated
a[CO2], which in turn supplies more33P to the host plant. At 440
p.p.m. a[CO2], the plantdoes not maintain the same supply of
photosynthatesto the fungus, and so the fungus does not return
asmuch nutrient to its host. This pattern of
‘tit-for-tat’reciprocity in plant–Glomeromycota symbiosis
haspreviously been demonstrated in various vascularplant species,
both in root-organ culture systems(Kiers et al., 2011) and in
whole-plant experiments(Hammer et al., 2011; Fellbaum et al.,
2014). Here wedemonstrate that this model does not apply in ourcase
of a plant symbioses involving more than onefungal partner and
involving Mucoromycotina fungi.
It is possible that by allocating excess photo-synthates
directly to Mucoromycotina fungal part-ners, rather than releasing
them as C-rich plantexudates, the liverworts avoid providing
excesscarbohydrate resources to surrounding saprotrophicmicrobes.
This may help to reduce nutrient immobi-lization by free-living
saprotrophic microorganismsand damage or toxicity caused by
potential microbialpathogens (Otten et al., 2004). These
potentialbenefits to the plants may contribute to the main-tenance
of Mucoromycotina fungal partnerships evenin plants that can form
symbiotic associations withGlomeromycota fungi and may explain why
thesehave not been lost entirely from extant plants
throughevolutionary time (Rimington et al., 2015). If
excessphotosynthates are released as exudates from theliverworts,
they are likely to enhance nutrientimmobilization and increase
their nutrient limitation.
Cytological characteristicsOur investigation reveals that the
cytology of fungalcolonization in both Neohodgsonia and Allisonia
istypical of mycorrhizal associations involving Glo-meromycota
fungi; in both it comprises prominentvesicles and well-developed,
short-lived arbusculesand/or fine hyphae surrounded by active
hostcytoplasm. The last feature is also typical of theintracellular
phase in Mucoromycotina associations(Desirò et al., 2013;
Strullu-Derrien et al., 2014;
Rimington et al., 2015; Field et al., 2015a). However,the key
feature of intracellular colonization
inHaplomitriopsida–Mucoromycotina symbiosis—hyphal coils with
terminal swellings (‘lumps’)(Carafa et al., 2003; Duckett et al.,
2006)—seems tobe unique and has not been observed in any
otherliverwort–fungus partnerships, including those inNeohodgsonia
and Allisonia.
Another diagnostic feature of Mucoromycotinacolonization,
intercellular fungal proliferation withthe production of
thick-walled spores in mucilagi-nous spaces, does occur across
plant lineages,including hornworts and lycopods, but
neitherNeohodgsonia nor Allisonia develop mucilage-filled
schizogenous intercellular spaces in theirthalli. It is
unsurprising therefore that in these twospecies we did not observe
any of the majorcytological differences between ambient and
ele-vated a[CO2]-grown plants reported in the Haplomi-triopsida
(Field et al., 2015a) as the latter wereexclusively associated with
the intercellular phase offungal colonization. The single minor
cytologicaldifference observed between wild and experimentalplants
grown at contrasting a[CO2], and restricted toAllisonia, was the
presence of far more starchgranules within thalli of this species
when grownat high a[CO2] (Figure 6h).
The only cytological features that may potentiallybe indicative
of fungal identity in these dualsymbioses are the fine/arbuscular
and trunk/coloniz-ing hyphal diameters (Strullu-Derrien et al.,
2014). InMucoromycotina–liverwort symbiosis, the finehyphae range
from 0.5 to 1.0 μm and the largerclasses are 3–4 μm (Haplomitrium,
Treubia), but inGlomeromycota–liverwort symbiosis
(Marchantia,Preissia, Pellia) the corresponding dimensions arefrom
1 to 3 μm and from 4 to 8 μm. Measurements ofNeohodgsonia and
Allisonia reveal that the finehyphae range from 0.6 to 1.2 μm, that
is, mostly inthe Mucoromycotina range, whereas the trunkhyphae,
ranging from 3 to 8 μm, are more typical ofGlomeromycota. In
contrast, vesicles are consistentlydiagnostic of Glomeromycota.
Thus, although theidentification of the two different fungi in
Neohodg-sonia and Allisonia largely rests with the
molecularevidence, there are indications from cytology for
thepresence of both Mucoromycotina and Glomeromy-cota fungi that
could be further explored bycytochemical and cytogenetic
techniques. Conse-quently, regarding the large number of
previousstudies, particularly in early-diverging plantlineages, in
which electron microscopy hasbeen used to describe mycorrhizal
associations as‘glomeromycotean’, our findings suggest that
crypticMucoromycotina associations may sometimes alsobe occurring
simultaneously.
In vitro isolation and resynthesis experiments withliverworts
known to engage in dual symbioses andwhereby either of the two
mycobionts is reintro-duced in the host plant will help to
determinecytological similarities and/or differences between
Dual fungal symbioses in thalloid liverwortsKJ Field et al
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the two fungal symbionts. Fluorescence in situhybridization may
allow localization of Glomeromy-cota and Mucoromycotina fungi
co-existing in thesame host plant to establish which structures
belongto which fungus. In the meantime, it is essential thatfungal
identification is carried out using appropri-ately inclusive
molecular techniques in any mycor-rhizal or mycorrhizal-like
symbiosis.
Wider perspectivesOur findings indicate that under a modern
near-ambient a[CO2], liverworts in partnership withMucoromycotina,
either in single or dual associa-tions alongside Glomeromycota
fungi, benefit fromgreater nutrient gain for carbon outlay than
liver-worts that maintain mutualistic symbioses with onlya
Glomeromycota fungal symbiont. From an evolu-tionary perspective,
the relative increases in nutrientexchange efficiency of plants
harbouring bothtypes of symbiont at lower a[CO2] may at
leastpartially explain why declining atmospheric a[CO2]over the
course of the Palaeozoic would havefavoured the retention of both
functional types ofsymbiosis. However, it is important to note
thatplants living in 1500 p.p.m. a[CO2] were likely toexperience
other abiotic factors that changed asplants evolved, including soil
mineralogy andnutrient supply.
The question remains whether Mucoromycotinafungal symbioses
resemble an ancestral conditionthat gave way to dual (for example,
Neohodgsoniaand Allisonia) and then solely Glomeromycotasymbiosis
(for example, Marchantia, Preissia, Con-ocephalum) or whether
co-evolution of plant andfungal symbioses have been more dynamic
thanpreviously thought (Field et al., 2015b). Indeed, theliverwort
phylogeny (Figure 1a) is associated withrepeated losses and
re-acquisitions of the same ordifferent fungal symbionts. That
liverwort cladessupporting dual fungal partnerships have
fungus-free sister groups, for example, the Sphaerocarpalesand
Blasiales (Pressel et al., 2010), points to shiftingfungal
associations during liverwort evolution.Exclusive
plant–Mucoromycotina fungal symbiosisbeing a basal trait is only
supported by theseassociations being present in liverworts of
theHaplomitriopsida (Bidartondo et al., 2011), the sistergroup to
all other liverworts (Forrest et al., 2006;Crandall-Stotler et al.,
2008, 2009), with liverwortsthemselves being the earliest diverging
land plantlineage (Alaba et al., 2014; Cox et al., 2014; Qiu et
al.,1998, 2006, 2007).
Mounting evidence that a large proportion of taxain all extant
early-diverging plant lineages (Desiròet al., 2013; Rimington et
al., 2015; Field et al.,2015a), and likely some Rhynie Chert fossil
plants(Strullu-Derrien et al., 2014), form dual symbiosiswith both
Mucoromycotina and Glomeromycotafungi now corroborates these
simultaneous fungalpartnerships as being an extremely ancient
condition, coincident with the early evolution ofland plants.
Why some Haplomitriopsida liverwortsdo not engage in symbiosis with
the ubiquitousGlomeromycota fungi remains enigmatic given theclear
advantages of dual partnerships demonstratedhere. Even less
comprehensible are the obligateGlomeromycota relationships in
thalloid liverwortssuch as Marchantia and Pressia, given that
recentfunctional studies clearly demonstrated that thesymbiotic
functional efficiency of these partnershipsis severely compromised
by the fall in a[CO2] thatoccurred through land plant
diversification (Fieldet al., 2012). In this context, it is
interesting to notethat liverwort clades harbouring exclusively
Glomer-omycota fungi have much later divergence timesthan those
able to associate with both fungalsymbionts (Cooper et al., 2012;
Feldberg et al.,2013). Marchantia, Conocephalum and
Preissiamostlikely diverged during the Cretaceous (Wikströmet al.,
2009; Villarreal et al., 2015), a period of rapidangiosperm and
polypodiaceous fern radiation(Schneider et al., 2004). We
hypothesize that duringthis period major changes in abiotic and
bioticdynamics, both below ground and above-ground,led to the
predominance of the biotrophic Glomer-omycota fungi in land
plant–fungal interactions. It ispossible therefore that these
Glomeromycota-specificliverworts evolved in
Glomeromycota-dominatedenvironments and never engaged in
associationswith Mucoromycotina fungi.
In this first assessment of the functionality andcytology of the
dual symbiosis of plants withMucoromycotina and Glomeromycota
fungi, wewere not able to distinguish between fungal partnersusing
microscopical techniques nor relative carbonallocation to each
fungal symbiont. Future researchusing axenic cultures of plants and
symbiotic fungimay enable such comparisons to be made and is anarea
for future development. More targeted cytologi-cal techniques, such
as fluorescence in situ hybridi-zation, may provide further novel
insights into theassociations and should be pursued in the
future.With the discovery of dual Mucoromycotina-Glomeromycota
symbioses in early branchinglineages of living vascular plants
(Rimington et al.,2015), it is now critical that we explore how far
thesemight extend into seed plants.
Conflict of Interest
The authors declare no conflict of interest.
AcknowledgementsWe gratefully acknowledge funding from NERC
(NE/1024089/1), a Leverhulme Emeritus Fellowship to JGDand a Royal
Society University Research Fellowship toDDC. We thank Irene
Johnson and Dr Heather Walker fortechnical assistance and stable
isotope analyses. We thankthe New Zealand Department of
Conservation for collect-ing permits. We thank the anonymous
referees and theeditor for their constructive comments on our
manuscript.
Dual fungal symbioses in thalloid liverwortsKJ Field et al
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title_linkIntroductionLiverwort phylogeny and species used in
the present study. (a) Liverwort phylogeny (following Wikström
etal., 2009) showing key nodes alongside commonly associated fungal
symbionts (James etal., 2006; Pressel etal., 2008; Bidartondo and
DuckettMaterials and methodsPlant material and growth
conditionsMolecular identification of fungal
associatesQuantification of fluxes of C, 33P and 15N between
liverworts and fungiPlant harvest and sample
analysesUltrastructural analysesStatistics
ResultsMolecular identification of fungi
Table 1 Summary of differences in mycorrhizal functionality (F
ratio from ANOVA) between Neohodgsonia, Alisonia, Haplomitrium,
Treubia, Preissia and Marchantia at elevated a[CO2] (1500 p.p.m.)
and ambient a[CO2] (440 p.p.m.)Plant biomassLiverwort-to-fungus
carbon transferFungal transfer of 33P and 15N to host
liverwortsNutrient-for-carbon exchange efficiencyCytology of
colonizationNeohodgsonia mirabilis
Mean total plant biomass (dry) at the end of experimental period
in five liverwort species (Field etal., 2012, 2015a) at both
1500 p.p.m. a[CO2] (black bars) and 440 p.p.m. a[CO2] (white bars).
Error bars show s.e.m. (n=4 for all spAllisonia cockaynei
Carbon-for-nutrient exchange between liverworts and their fungal
partners. (a) Percentage allocation of plant-derived carbon to
fungi within soil cores, (b) total measured plant-fixed carbon
transferred to fungi in soil for liverworts with different
fungaDiscussionPhysiological costs and benefits
Nutrient-for-carbon exchange efficiencies between liverworts and
their fungal partners. (a) 33P-for-carbon and (b) 15N-for-carbon
efficiency for different liverwort species with different fungal
associations under both 1500 p.p.m. (black bars) aCytology of
Neohodgsonia mirabilis grown at 440 and 1500 p.p.m. a[CO2].
Scanning (a–c) and transmission (d–f) electron micrographs (TEM).
Both the distribution and cytology of the association remained the
same between a[CO2] treaCytology of Allisonia cockaynei grown at
440 and 1500 p.p.m. a[CO2]. Scanning (a–c) and transmission (e–i)
electron micrographs (TEM). There was no change in the overall
distribution of fungal colonization and in the cytology of
Cytological characteristicsWider perspectives
We gratefully acknowledge funding from NERC (NE/1024089/1), a
Leverhulme Emeritus Fellowship to JGD and a Royal Society
University Research Fellowship to DDC. We thank Irene Johnson and
Dr Heather Walker for technical assistance and stable isotope
analyACKNOWLEDGEMENTS