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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: k.field@leeds.ac.ukReceived 23 March 2015; revised 8 October 2015; accepted 12October 2015; published online 27 November 2015

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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

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Pleurozia

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lidae

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Basidiomycota

Ascomycota

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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).

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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

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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.

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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

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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).

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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).

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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.

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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

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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).

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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.

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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.

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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

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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.

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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