Short Title: Vascular plant-Mucoromycotina fungal symbiosis · 65 Glomeromycotina fungi in the same plant host (Desirò et al., 2013; Rimington et al., 2015). 66 Mucoromycotina represents

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Short Title: Vascular plant-Mucoromycotina fungal symbiosis 1

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* Corresponding author: Katie J. Field ([email protected]); Tel: +44 (0)113 343 2849 3

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Mucoromycotina fine root endophyte fungi form nutritional mutualisms with vascular 5

plants 6

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Grace A. Hoysted1, Alison S. Jacob2,3, Jill Kowal4, Philipp Giesemann5, Martin I. Bidartondo2,3, 8

Jeffrey G. Duckett4, Gerhard Gebauer5, William R. Rimington2,3,4, Sebastian Schornack6, Silvia 9

Pressel4, and Katie J. Field1* 10

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1 Centre for Plant Sciences, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 12

9JT, UK 13

2 Comparative Plant & Fungal Biology, Royal Botanic Gardens, Kew, Richmond TW9 3DS, 14

UK 15

3 Department of Life Sciences, Imperial College London, London, SW7 2AZ, UK 16

4 Department of Life Sciences, Natural History Museum, London SW7 5BD, UK 17

5 Laboratory of Isotope Biogeochemistry, Bayreuth Center of Ecology and Environmental 18

Research (BayCEER), University of Bayreuth, Bayreuth, Germany 19

6 Sainsbury Laboratory, University of Cambridge, Cambridge, CB2 1LR 20

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One sentence summary: Mucoromycotina fine root endophyte fungi transfer soil nutrients, 22

notably N, to lycophyte host plants in exchange for photosynthetically-fixed C and colonise 23

diverse neighbouring vascular plants. 24

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Author contributions 26

K.J.F., S.P., S.S., M.I.B., and J.G.D. conceived and designed the investigation. S.P., J.K., 27

J.G.D., M.I.B., A.S.J., G.G., and G.A.H collected plant material. G.A.H. and K.J.F. undertook 28

Plant Physiology Preview. Published on July 29, 2019, as DOI:10.1104/pp.19.00729

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the physiological analyses. A.S.J., W.R.R., and M.I.B. undertook the molecular anlayses. S.P. 29

undertook the cytological analyses with assistance from J.K. P.G. and G.G. analysed and 30

interpreted the stable isotope data. G.A.H. led the writing, all authors discussed results and 31

commented on the manuscript. K.J.F. agrees to serve as the author responsible for contact 32

and ensure communication. 33

34

Abstract 35

Fungi and plants have engaged in intimate symbioses that are globally widespread and have 36

driven terrestrial biogeochemical processes since plant terrestrialisation >500 Mya. Recently, 37

hitherto unknown nutritional mutualisms involving ancient lineages of fungi and non-vascular 38

plants have been discovered, although their extent and functional significance in vascular 39

plants remain uncertain. Here, we provide evidence of carbon-for-nitrogen exchange between 40

an early-diverging vascular plant (Lycopodiella inundata L.) and Mucoromycotina 41

(Endogonales) fine root endophyte fungi. Furthermore, we demonstrate that the same fungal 42

symbionts colonize neighbouring non-vascular and flowering plants. These findings 43

fundamentally change our understanding of the physiology, interrelationships, and ecology of 44

underground plant–fungal symbioses in modern terrestrial ecosystems by revealing the 45

nutritional role of Mucoromycotina fungal symbionts in vascular plants. 46

47

Key Words 48

Arbuscular mycorrhizas, Endogonales, lycophytes, nitrogen, carbon. 49

50

Introduction 51

Plant terrestrialisation >500 Mya (Morris et al., 2018) was facilitated by the formation of 52

mutualistic symbioses with fungi, through which the earliest plants gained access to mineral 53

nutrients in exchange for photosynthetically-fixed carbon (C). It was long hypothesised that 54

this ancient mycorrhiza-like symbiosis was closely related to, and subsequently evolved into, 55

widespread modern-day arbuscular mycorrhizas (AM) formed with plant roots by 56



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Glomeromycotina fungi (Pirozynski and Malloch, 1975; Redecker et al., 2000). However, 57

recent molecular, cytological, physiological, and paleobotanical evidence has strongly 58

indicated that early fungal associates were likely to be more diverse than has previously been 59

assumed (Bidartondo et al., 2011; Field et al., 2015a; Field et al., 2015b). Members of the 60

earliest diverging clade of an ancient land plant lineage, Haplomitriopsida liverworts, are now 61

known to form mycorrhiza-like associations with Mucoromycotina fungi (Bidartondo et al., 62

2011; Field et al., 2012; Field et al., 2015b) which also colonise other early diverging plant 63

lineages, namely hornworts, lycophytes, and ferns, sometimes co-occurring with 64

Glomeromycotina fungi in the same plant host (Desirò et al., 2013; Rimington et al., 2015). 65

Mucoromycotina represents an ancient fungal lineage considered to branch earlier than or 66

sister to the Glomeromycotina AM fungi (James et al., 2006; Lin et al., 2014). The recent 67

identification of Mucoromycotina in a range of modern non-vascular plants (Bidartondo et al., 68

2011) and plant fossils (Krings et al., 2007; Strullu‐Derrien et al., 2014) supports the idea that 69

the colonisation of Earth’s land masses by plants was facilitated not only by Glomeromycotina 70

AM, but also by Mucoromycotina fungal symbionts (Field et al., 2015a). The latest discoveries 71

of putative Mucoromycotina fungi in vascular land plants (Rimington et al., 2015; Rimington et 72

al., 2016; Orchard et al., 2017a) indicate that root symbiotic versatility and diversity (Hoysted 73

et al., 2018) has been grossly underestimated across extant plants. 74

Although Mucoromycotina fungal symbioses in non-vascular plants have received the 75

most attention to date, there are now several reports of their occurrence in vascular plants 76

(Rimington et al., 2015; Rimington et al., 2016; Orchard et al., 2017a; Orchard et al., 2017b; 77

Hoysted et al., 2018). It has been suggested that the globally widespread, arbuscule-forming 78

fine root endophytes (FREs) classified as Glomus tenue (or more recently Planticonsortium 79

tenue (Walker et al., 2018)), which occur across a wide range of vascular plant groups 80

(Rimington et al., 2015; Orchard et al., 2017b), are closely related to the Mucoromycotina 81

fungal symbionts of non-vascular plants (Hoysted et al, 2018; Field and Pressel, 2018). If true, 82

there could be major ramifications for our understanding of the past and present diversity and 83

function of plant-fungal nutritional symbioses (Field and Pressel, 2018), suggesting 84



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Mucoromycotina fungal symbiosis is not limited to ancient plant lineages but is in fact 85

widespread throughout extant land plants. However, it remains unclear whether the putative 86

Mucoromycotina FREs detected in vascular plants to date are comparable in terms of function 87

and identity to the mutualistic Mucoromycotina fungal symbionts detected in non-vascular 88

plants. 89

As lycophytes are considered to be the earliest divergent extant vascular plant lineage 90

(Kenrick and Crane, 1997), the discovery of non-Glomeromycotina fungal associates in 91

lycophyte roots and gametophytes is particularly important. For over 100 years, fungal 92

associations in lycophytes have been thought of as being AM-like but with unique “lycopodioid” 93

features (Duckett and Ligrone, 1992; Schmid and Oberwinkler, 1993). However, global 94

analysis of fungal associates in 20 lycophytes (Rimington et al., 2016) has now shown their 95

colonisation is broadly similar to that of hornworts (Desirò et al., 2013), with many species 96

forming single and/or dual associations with both Glomeromycotina arbuscular mycorrhizal 97

(AM) fungi and Mucoromycotina FRE fungi (Rimington et al., 2016). Remarkably, every 98

sample of Lycopodiella inundata - a species found in wet habitats across the Northern 99

Hemisphere - examined so far appears colonised exclusively by Mucoromycotina FRE fungi 100

(Rimington et al., 2016). The fundamental obstacle to studying function of Mucoromycotina 101

FREs has been finding any plants that are not co-colonized by coarse root endophytes (i.e., 102

Glomeromycotina AM fungi). In fact, so far there is no evidence of nutritional mutualism 103

between any vascular plant and Mucoromycotina FREs (Hoysted et al., 2018). Therefore, L. 104

inundata provides a unique and important opportunity to dissect the symbiotic function of 105

FREs in a vascular plant. 106

Here, we investigate the function, cytology, and occurrence of the fungal associates of 107

L. inundata (Figure 1a, b). We use a combination of radio- and stable isotope tracers 108

(Supplemental Figure S1, S2) to test physiology and functioning of vascular plant-109

Mucoromycotina fungal symbioses, detailed cytological analyses to characterise morphology 110

and colonisation patterns of Mucoromycotina fungi, and molecular techniques to identify the 111

fungal associates of a range of vascular plants (including those used in our experiments) 112



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across several field sites. Furthermore, we used natural abundance 13C and 15N signatures 113

to test under field conditions whether Mucoromycotina fungal association affects the direction 114

of carbon and nitrogen fluxes between plants and associated fungi (Gebauer and Meyer, 115

2003). Specifically, we address the following questions: 116

(1) What is the function of Mucoromycotina fungal associations in lycophytes in terms 117

of carbon-for-nutrient exchange? 118

(2) Are there characteristic cytological signatures or features of Mucoromycotina 119

fungal associations in L. inundata and other vascular plants? 120

(3) Do Mucoromycotina fungal symbionts of L. inundata co-occur in neighbouring 121

angiosperm roots and non-vascular plants? 122

123

Results 124

Mucoromycotina-L. inundata symbioses are nutritional mutualisms 125

Isotope tracing experiments conducted in controlled environment chambers confirmed 126

carbon was transferred from field-collected L. inundata to extraradical hyphae of symbiotic 127

Mucoromycotina FRE fungi (Figure 2). Mucoromycotina fungal symbiont identity was 128

confirmed using sequencing of the fungal 18S ribosomal rRNA gene with the broad specificity 129

fungal primer set NS1/EF3 and a semi-nested approach with Mucoromycotina- and 130

Glomeromycotina-specific primers described in Desirò et al., (2013) (Supplemental Figure 131

S3). Plants transferred an average of 79 g ( 49.3 SE) of recent photosynthate to the external 132

fungal hyphal mycelium within the microcosm during the labelling period (Fig. 2a). This 133

represents 0.28% ( 0.14 SE) of the total amount of carbon that was fixed during the labelling 134

period by L. inundata (Fig. 2b). 135

Mucoromycotina fungi within the experimental microcosms transferred between 3-9% 136

of the supplied 33P tracer and 0.6-1% of the supplied 15N tracer to their plant hosts during the 137

isotope labelling period (Figure 2c, d). Mucoromycotina FREs transferred significantly more 138

15N than 33P to L. inundata both in terms of absolute quantities (Figure 2c, P = 0.05; Student’s 139



6

T-Test) and when normalised to plant biomass (Figure 2d, P = 0.03, Student’s T-Test). 140

To test the potential nutritional role of Mucoromycotina fungi in L. inundata in the field, 141

we analysed the natural abundance 13C and 15N stable isotope signatures of leaves and roots 142

of L. inundata and Juncus bulbosus, both of which were shown to host Mucoromycotina FREs 143

in their roots (Supplemental Figure S4-S9). In addition, five plant species representing three 144

different types of mycorrhizal associations were sampled to serve as reference plants: two 145

ericoid mycorrhizal species (Erica tetralix, collected on six plots; Calluna vulgaris, collected on 146

three plots), two ectomycorrhizal species (Pinus sylvestris and Betula pendula seedlings, both 147

from one plot) and one arbuscular mycorrhizal species (Molinia caerulea from six plots). 148

All leaf δ13C values from plants collected from the same site as those collected for our 149

isotope tracing experiments ranged between -26.2 and -30.1 ‰ and root δ13C values between 150

-24.5 and -28.9 ‰, while leaf δ15N values ranged from 3.3 to -10.0 ‰ and root δ15N values 151

from 3.1 to -5.9 ‰ (Figure 3). Leaves of the three groups, L. inundata (n = 6), Juncus bulbosus 152

(n = 6) and reference plants, five species representing three types of mycorrhizal associations, 153

i.e. arbuscular, ericoid and ectomycorrhizal species (n = 17), also collected from the same site 154

as plants used in our isotope tracing experiments, were significantly different in δ13C (H(2) = 155

8.758; P = 0.013) and δ15N (H(2) = 21.434; P < 0.001, Figure 3a). L. inundata leaves were 156

significantly depleted in 13C compared to J. bulbosus leaves (Q = 2.644, P < 0.05) and a 157

significant depletion of L. inundata leaves compared to reference plant leaves (Q = 2.662, P 158

< 0.05, Figure 3a) was found. The J. bulbosus leaves were not significantly different from 159

reference plants in δ13C. No significant difference was discovered for δ15N in L. inundata and 160

J. bulbosus leaves (Q = 1.017, P > 0.05), while leaves of both species were significantly 161

enriched in 15N compared to the reference plants (Q = 2.968, P < 0.05; Q = 4.205, P < 0.05, 162

Figure 3a). For the roots, only δ15N showed significant differences between the three groups 163

under comparison (F(2) = 34.815; P < 0.001, Figure 3b). The L. inundata and J. bulbosus 164

roots were not significantly distinguished in δ15N, however, roots of both species were 165

significantly enriched in 15N compared to reference plant roots (q = 10.109, P < 0.001; Q = 166

8.515, P < 0.001, Figure 3b). 167



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168

Mucoromycotina fine root endophytes of L. inundata show distinctive cytology 169

Trypan blue staining and SEM of wild-collected plants (a liverwort, two grasses, and a 170

rush) from the same site as the L. inundata plants used in our isotope tracer and stable isotope 171

studies (except for the grass Holcus lanatus, see Supplemental Table S1), revealed two 172

distinct fungal symbiont morphologies. These consisted of either coarse hyphae (>3 µm 173

diameter) and large vesicles (>20 µm diameter) or fine branching hyphae (<2 µm diameter) 174

with small swellings/vesicles (usually 5-10 but up to 15 µm diameter) (Figures 4-5). Both 175

morphologies were regularly observed, often co-occurring in the same sample, in the 176

gametophyte of the liverwort Fossombronia foveolata (Figures 4a, b, 5a; Supplemental Figure 177

S10a), in the roots of the grasses Holcus lanatus (Figure 4f) and Molinia caerulea (Figure 4g, 178

h), and the rush Juncus bulbosus (Figure 5h, i). In the colonised roots of both freshly collected 179

L. inundata and those incubated in growth chambers, only fine hyphae with small 180

swelling/vesicles were invariably detected (Figures 4c-e, 5f, g). As in the other plants 181

analysed, these fine hyphae were aseptate and formed both intercalary and terminal 182

swellings/vesicles but, in contrast to the grasses (Supplemental Figure S10b), never 183

arbuscules. Similar fungal morphology was also observed in protocorm cells of newly 184

developing sporophytes (Figure 5b, c) and in gametophytes of L. inundata (Supplemental 185

Figure S11). However, in these early developmental stages, fungal colonization consistently 186

exhibits a distinct zonation: an outer intracellular zone and a more central, strictly intercellular 187

zone (Figure 5d, e; Supplemental Figure S11b-g). In the intracellular zone, fungal colonization 188

is the same as in the sporophyte roots and consists of fine hyphae with intercalary and terminal 189

swellings/vesicles (Figure 5b, c; see also Supplemental Figure S11i). Unique to the 190

gametophyte generation, in the outermost cortical layers, the fungus also forms tightly wound 191

coils (hyphae up to 2.5 µm in diameter) with larger vesicles (15-20 µm) (Supplemental Figure 192

S11d), as described before in Lycopodium clavatum (Schmid and Oberwinkler, 1993). Both 193

gametophyte and early developmental stages of the sporophyte generation develop a 194

conspicuous central system of large, mucilage-filled intercellular spaces. In this region, the 195



8

fungus becomes strictly intercellular (Figure 5d, e; Supplemental Figures S11e-g). The 196

intercellular hyphae are initially fine and with small swellings/vesicles (Figure 5d, 197

Supplemental Figure S11e), as their intracellular counterparts, but soon enlarge and 198

eventually reach diameters in excess of 3 µm (Supplemental Figure S11f), with no 199

swellings/vesicles present at this stage. 200

201

Mucoromycotina fungal symbionts are shared by neighbouring angiosperms 202

Analysis of L. inundata plants collected from the same site, at the same time as plants 203

used in the above investigations, confirmed that they were colonised by Mucoromycotina 204

fungi. Glomeromycotina sequences were not detected. Mucoromycotina OTUs were detected 205

before and after the experiments (Supplemental Table S2); these same OTUs had previously 206

been identified in wild-collected lycophytes from diverse locations (Rimington et al., 2015). 207

Diverse and shared Mucoromycotina fungi OTUs were detected in wild L. inundata, 208

liverworts, and angiosperms growing adjacently in the same UK locations (Supplemental 209

Table S2, Supplemental Figures S2-8) in the following combinations: L. inundata, F. foveolata, 210

M. caerulea, and J. bulbosus (Thursley Common, Surrey); L. inundata, F. foveolata, and J. 211

bulbosus (Norfolk); F. foveolata and H. lanatus (Lynn Crafnant, Wales). Mucoromycotina 212

OTUs were also detected in L. inundata from Studland Heath, Dorset. 213

214

Discussion 215

Our results show that the symbiosis between L. inundata and Mucoromycotina FREs are 216

nutritionally mutualistic, with the fungus gaining plant-fixed C and the plant gaining fungal-217

acquired P and N (Figure 2a-d). Cytological analyses of the fungus colonising the roots of L. 218

inundata revealed a characteristic morphology consisting invariably of fine, aseptate 219

branching hyphae with terminal and intercalary swellings/vesicles. This morphology matches 220

that described previously in a range of angiosperms colonized by FREs (Orchard et al., 2017a; 221

Orchard et al., 2017b) and here in grasses, a rush, and a liverwort, all harbouring fungi 222

identified molecularly as Mucoromycotina (Supplemental Figure S3). Our results provide 223



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compelling evidence for Mucoromycotina FREs being shared by plants occupying key nodes 224

in the land plant phylogeny - from early liverworts and vascular lycophytes to the later diverging 225

angiosperms - and demonstrate that this association represents a nutritional mutualism as 226

much in vascular plants as it does in non-vascular plants (Field et al., 2015b; Field et al., 227

2016). 228

Our findings raise important questions regarding the ecology and evolution of 229

mycorrhizal associations and the nature of widespread Mucoromycotina FRE fungal 230

symbioses, chief among which is how have these associations persisted and why are they so 231

widespread today? We can now begin to address this with the demonstration that a vascular 232

plant assimilates relatively large amounts of 15N tracer via its Mucoromycotina fungal symbiont 233

when compared to the Mucoromycotina fungal-acquired 33P tracer (Fig. 2c), suggesting a 234

potential role for Mucoromycotina FREs in vascular plant nitrogen uptake, complementary to 235

the role of Glomeromycotina AM fungi (Field et al., 2019). This nutritional role could help to 236

explain the persistence of Mucoromycotina FREs across nearly all modern land plant lineages. 237

238

Costs and benefits of hosting Mucoromycotina fungi 239

Our data demonstrate that L. inundata transfers carbon to symbiotic Mucoromycotina 240

FRE (Figure 2a, b). However, when compared to other vascular plants with Glomeromycotina 241

AM fungal associates in similar experimental systems (Field et al., 2012), it is clear that the 242

relative C “cost” of maintaining Mucoromycotina fungal symbionts in L. inundata is at least on 243

a par with, if not greater than, that of maintaining Glomeromycotina fungi. It is not entirely clear 244

from our experiments why this might be and it is important to note that the C “cost” of hosting 245

Mucoromycotina FREs has only been tested in one vascular plant species to date and thus 246

represents an important area for future research. It is possible that the carbon-for-nutrient 247

exchange dynamics between plant and Mucoromycotina FREs varies according to plant and 248

fungal identity, in addition to abiotic factors, as it does for Glomeromycotina AM (Field and 249

Pressel, 2018). 250



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Lycophytes represent a critical node in land plant phylogeny, widely considered as a 251

diversification point in the mid-Paleozoic (480-360 Ma) characterised by the evolution of roots, 252

leaves, stomata, and associated vasculature (Kenrick and Crane, 1997). Given that all the 253

plants sampled grew in close proximity and all follow the C3 photosynthetic pathway, the trend 254

for lower δ13C values in root tissues versus leaves (Figure 3a, b) is most likely caused by 255

systematic differences in 13C abundance in photosynthetic vs. non-photosynthetic tissues 256

(Gebauer and Schulze, 1991; Cernusak et al., 2009). The depletion of 13C observed in the 257

leaves of L. inundata (Figure 3a) relative to the other, non-Mucoromycotina associated plants 258

sampled is unlikely to be related to C gains from its Mucoromycotina fungal symbiont (Bago 259

et al., 2000) as one would expect (myco)heterotrophic carbon gains to result in 13C enrichment 260

(Press et al., 1987; Schulze et al., 1991; Gebauer and Meyer, 2003); rather, it may indicate 261

that L. inundata leaves regulate their stomata differently from J. bulbosus or the reference 262

plants tested, as δ13C in tissues of terrestrial plants is, among other factors, driven by the 263

water use efficiency of the plant (Farquhar et al., 1982; Farquhar et al., 1989). 264

Alongside increased capacity for regulation of water relations and C capture and 265

fixation, we hypothesise that the increasing size and structural complexity of land plants 266

across the land plant phylogeny and evolutionary time (Field et al., 2012) result in greater 267

plant nutrient demand. Glomeromycotina AM are associated with facilitation of plant P uptake 268

and occur commonly in soils with low P availability (Smith et al., 2015; Albornoz et al., 2016). 269

The amount of 33P transferred to L. inundata plants in our experiments was much less than 270

has previously been recorded for Mucoromycotina-associated liverworts (Field et al., 2016) or 271

for Glomeromycotina-associated ferns and angiosperms (Field et al., 2012) despite the same 272

amount of 33P being made available in comparable experimental systems. This suggests 273

Mucoromycotina fungi may not play a critical role in lycophyte P nutrition. Our results contrast 274

with the view that Mucoromycotina FREs enhance plant P uptake, at least in soils with very 275

low P (Ryan and Kirkegaard, 2012; Orchard et al., 2017b). Previous experiments with 276

Mucoromycotina fungi-associated liverworts suggest that in addition to supplying host plants 277

with P, Mucoromycotina fungal associates also play a role in plant N nutrition (Field et al., 278



11

2015b; Field et al., 2016; Field et al., 2019). 279

Nitrogen is an essential element for plants that is available in soils in plant-inaccessible 280

organic forms and as plant-accessible inorganic nitrate and ammonium (Krapp, 2015). Our 281

results show that Mucoromycotina FRE symbionts transfer significantly more 15N tracer 282

compared to 33P (Figure 2c, d). Up to 145 times more 15N was transferred to L. inundata (0.3-283

1% of the supplied tracer) than to Haplomitriopsida liverworts in comparable experiments 284

which assimilated between 0.05 and 0.2 % of the supplied tracer (Field et al., 2016). Using 285

analysis of natural abundance 15N signatures, we show that Mucoromycotina FRE-associated 286

L. inundata and J. bulbosus were 15N enriched compared to co-occurring reference plants with 287

different mycorrhizal fungal partners (Figure 3). This 15N enrichment could be caused by 288

temporal and spatial variations in N availability and changes in plant N demand over time 289

(Gebauer and Meyer, 2003). However, these factors are unlikely in our case, because all 290

plants sampled from this field collection grew in close spatial proximity and were collected at 291

the same time. Presence of multiple N sources with distinct isotopic values and their utilization 292

by different mycorrhizal associations are known as additional drivers of variations in plant 15N 293

isotope abundance (Bidartondo et al., 2004). While this distinction in N isotope abundance 294

between plants with different mycorrhizas is almost or completely lost in conditions of high N 295

availibility (Gebauer and Meyer, 2003), it may become prominent under severe N limitation 296

(Schulze et al., 1994). Given that our plants were collected at a heathland field site which was 297

likely N-limited (von Oheimb et al., 2010) despite substantial atmospheric N deposition, the 298

separation of L. inundata and J. bulbosus in their natural abundance 15N from neighbouring 299

plants with other fungal associations supports our hypothesis that plants hosting 300

Mucoromycotina symbionts benefit from fungal-acquired N. 301

Some Glomeromycotina AM fungi transfer N to their associated hosts (Farquhar et al., 302

1989); however, the ecological relevance of AM-facilitated N uptake is widely debated, in 303

particular the amounts of N transferred to hosts compared to the overall N requirements of the 304

plant (Smith and Smith, 2011). Exclusive plant-Mucoromycotina FRE symbioses seem to be 305

rare, having been reported before only in the earliest-diverging Haplomitriopsida liverworts 306



12

(Field et al., 2015a; Field et al., 2015b), while all other plants, including other lycophytes 307

(Rimington et al., 2015), that form associations with these fungi appear able to do so also with 308

Glomeromycotina, often simultaneously (Rimington et al., 2015). It is possible that the large 309

input to Lycopodiella N-nutrition and minor contribution to P-nutrition by Mucoromycotina 310

FREs reflect a specialised relationship, particulary pertinent when considering heathland 311

habitats have very low plant-available N. Nevertheless, our present data combined with 312

previous demonstrations of N transfer in liverwort-Mucoromycotina symbioses (Field et al., 313

2015b; Field et al., 2016) and emerging evidence that Mucoromycotina FREs, but not 314

Glomeromycotina AM fungi, are able to transfer N to host liverworts from organic sources 315

(Field et al. 2019), all point to a critical role of Mucoromycotina FREs in host plant N nutrition. 316

Indeed, our cytological analyses show that, differently from Lycopodiella roots where only fine 317

endophytes were observed (Figures 4, 5; Table 1), all other co-occurring plants (F. foveolata, 318

J. bulbosus, and M. caerulea) were also colonised by coarse endophytes with cytology typical 319

of Glomeromycotina (Figures 4, 5; Table 1). The finer functional details, in terms of N and P 320

transfer, of this partnership in other vascular plants from a broader range of habitats remain 321

to be established; the challenge here will be to separate the nutritional contributions of 322

Mucoromycotina FREs and Glomeromycotina to host plants that are co-colonized by both 323

fungi (in addition to the contributions made by any mutualistic Ascomycetes and 324

Basidiomycetes) as that seems to be the prevailing condition in vascular plants, especially 325

angiosperms. 326

327

Mucoromycotina fine root endophytes 328

Mucoromycotina fungi within Endogonales colonising the gametophytes of liverworts 329

(F. foveolata) and lycophytes (L. inundata), the sporophytic protocorms and roots of 330

lycophytes (L. inundata), and the roots of angiosperms (J. bulbosus, M. caerulea, and H. 331

lanatus), all display the same characteristic morphology attributed previously to FREs 332

(Orchard et al., 2017b; Walker et al., 2018). This contrasts with that typical of 333

Glomeromycotina AM fungal associations, consisting of coarse hyphae (>3 µm diameter) and 334



13

larger vesicles, which we observed in Fossombronia, Juncus, Molinia, and Holcus but not in 335

L. inundata (Table 1). These observations, together with the molecular identification of 336

Mucoromycotina clades shared by these phylogenetically distant plant lineages presented 337

here, support previous suggestions that vascular plants’ FREs are closely related to the 338

Mucoromycotina mycorrhizal-like symbionts of non-vascular plants (Rimington et al., 2016). 339

Here, we show that the same Mucoromycotina FREs have the capacity to be nutritionally-340

mutualistic across different land plant phyla. 341

Our demonstration of an extensive intercellular phase of fungal colonisation in the 342

gametophytes and protocorms of L. inundata is in line with other lycophytes (Schmid and 343

Oberwinkler, 1993; Rimington et al., 2015) and strongly recalls the gametophytes of the 344

Haplomitriopsida liverwort Treubia (Duckett et al., 2006) and several hornworts (Desirò et al., 345

2013), all of which have also been shown to associate with Mucoromycotina fungi (Bidartondo 346

et al., 2011; Desirò et al., 2013). Differently from their fine intracellular counterparts, 347

intercellular hyphae become swollen, eventually reaching more than 3 µm in diameter. Tightly 348

wound hyphal coils up to 2.5 µm in diameter with somewhat larger terminal vesicles (up to 20 349

µm in diameter) are also prominent in the outer cortical layers of L. inundata gametophytes 350

but were not observed in either protocorms or roots. Thus, Mucoromycotina FREs display 351

considerable phenotypic plasticity in their interactions with diverse lineages of land plants 352

which appears to relate to the developmental stage of the host and whether it produces an 353

extensive network of mucilage-filled intercellular spaces. The putative occurrence of 354

Mucoromycotina FREs in early land plants and their presence in both extant early and later 355

diverging plant lineages now point to a prominent role of these fungi, not only in plant 356

terrestrialization (Field et al., 2015a), but also in current ecosystem functioning. Indeed, 357

Mucoromycotina FREs have been shown to occur worldwide across many ecosystems, 358

particularly in the roots of crop and pasture species, where colonization levels may be high, 359

even as dense as the biomass of coarse Glomeromycotina arbuscular mycorrhizal fungi 360

(Orchard et al., 2017b). 361

362



14

Conclusions 363

More ammunition for the mycorrhizal revolution 364

Our findings provide conclusive evidence that Mucoromycotina FREs form nutritional 365

mutualisms not only with non-vascular liverworts (Field et al., 2015b; Field et al., 2016), but 366

also with a vascular plant. We have found that the Mucoromycotina FRE associates of L. 367

inundata receive up to 189 times more photosynthesis-derived C from the plant than the 368

Mucoromycotina fungal associates of non-vascular plants (Field et al., 2016). In return, L. 369

inundata hosts receive a relatively large amount of N from their Mucoromycotina FRE partners 370

- ca. 145 times more than non-vascular plants receive from their Mucoromycotina fungal 371

symbionts (Field et al., 2016). Together with our discovery that the same Mucoromycotina 372

fungal symbionts are shared with neighbouring grasses, rushes and liverworts, and recent 373

findings of functional complementarity between Mucoromycotina FREs and Glomeromycotina 374

AM (Field et al., 2019), our findings point towards a unique physiological niche for the 375

persistence of Mucoromycotina fungi, both in single and dual colonisations with 376

Glomeromycotina AM. 377

378

Materials and Methods 379

Plant material and growth conditions 380

Lycopodiella inundata (L.), neighbouring angiosperms (the grasses Holcus lanatus, 381

Molinia caerulea, and the rush Juncus bulbosus), and a liverwort (Fossombronia foveolata) 382

were collected from Thursley National Nature Reserve, Surrey, UK (SU 90081 39754), a 383

heathland site, in June 2017. The L. inundata plants were planted directly into pots (90 mm 384

diameter x 85 mm depth) containing a homogenous mixture of acid-washed silica sand and 5 385

% pot volume compost (Petersfield No.2, Leicester, UK) to aid water-retention properties of 386

the substrate and to provide minimal nutrients. Soil surrounding plant roots was left intact to 387

prevent damage to the roots and to act as a natural inoculum, including symbiotic fungi and 388

associated microorganisms. Pots were weeded regularly to remove other plant species. All 389

plants used throughout this investigation were collected from the wild. Each microcosm was 390



15

a homogenised mixture of acid-washed sand and soil collected from around the plant roots 391

from the collection site. All plants used in this investigation (isotope tracing, cytology, and 392

stable isotope studies) were collected from Thursley Nature Reserve, with additional plants 393

from three other UK field sites (Supplemental Table S1), used for additional cytological and 394

molecular analyses. 395

Based on the methods of Field et al. (2012; 2015a), three windowed cylindrical plastic 396

cores covered in 10 µm nylon mesh (Supplemental Figure S1) were inserted into the substrate 397

within each experimental pot. Two of the cores were filled with the same homogenous mixture 398

of acid-washed silica sand and compost (Petersfield No.2, Leicester, UK), together making up 399

95% of the core volume as made up with microcosm bulk soil together with native soil gathered 400

from around the roots of wild plants (4% of the core volume) to ensure cores contained the 401

same microbial communities as bulk soil in pots, and fine-ground tertiary basalt (1% core 402

volume) to act as fungal bait. The third core was filled with glass wool to allow below-ground 403

gas sampling throughout the 14C-labelling period to monitor soil community respiration. Plants 404

were watered every other day with no additional applications of nutrient solutions. Microcosms 405

shared a common drip-tray within each cabinet throughout the acclimation period which 406

ensured a common pool of rhizospheric microorganisms in each microcosm. 407

A total of 20 L. inundata microcosms were maintained in controlled environment 408

chambers (Micro Clima 1200, Snijders Labs, The Netherlands) with a light cycle of 16 h 409

daytime (20 °C and 70 % humidity) and 8 h night-time (at 15 °C and 70 % humidity). Day-time 410

PAR, supplied by LED lighting was 225 µmol photons m-2 s-1. Atmospheric CO2 (a[CO2]) 411

concentrations were set at 440 ppm. a[CO2] was monitored using a Vaisala sensor system 412

(Vaisala, Birmingham, UK), maintained through addition of gaseous CO2. Cabinet settings and 413

contents were alternated every four weeks, and all pots were rotated within cabinets to control 414

for cabinet and block effects. Plants were acclimated to chamber/growth regimes (see 415

Supplemental Data) for four weeks to allow establishment of mycelial networks within pots 416

and confirmed by hyphal extraction from soil and staining with trypan blue (Brundrett et al., 417

1996). Additionally, roots were stained with acidified ink for the presence of fungi, based on 418



16

the methods of Brundrett et al., (1996). 419

420

Molecular identification of fungal symbionts and phylogenetic analysis 421

All plants (Supplemental Table S1) were processed for molecular analyses within one 422

week of collection. Genomic DNA extraction and purification from all specimens and 423

subsequent amplification, cloning and Sanger sequencing were performed according to 424

methods from Rimington et al., (2015). The fungal 18S ribosomal rRNA gene was targeted 425

using the broad specificity fungal primer set NS1/EF3 and a semi-nested approach with 426

Mucoromycotina- and Glomeromycotina-specific primers described in Desirò et al., (2013) for 427

the experimental L. inundata plants and all other field collected plant material. Resulting partial 428

18S rDNA sequences ca. 400-700bp were edited and preliminarily identified with BLAST in 429

Geneious v. 8.1.7 (Kearse et al., 2012). Chimeric sequences were detected using the 430

UCHIME2 algorithm (Edgar, 2016) in conjunction with the most recent non-redundant SSU 431

SILVA database (SSU Ref NR 132, December 2017, www.arb-silva.de). Sequences identified 432

as Mucoromycotina sp. were aligned with MAFFT prior to removing unreliable columns using 433

the default settings in GUIDANCE2 (http://guidance.tau.ac.il). The best-fit nucleotide model 434

for phylogenetic analysis was calculated using Smart Model Selection (Kumar et al., 2016). 435

Maximum likelihood (ML) with 1,000 replicates was performed using PhyML 3.0 (Guindon and 436

Gascuel, 2003). Bayesian inference analysis was conducted in Mr Bayes version 3.2.6 437

(Ronquist and Huelsenbeck, 2003) with four Markov chain Monte Carlo (MCMC) strands and 438

106 generations. Consensus trees were produced after excluding an initial burn-in of 25% of 439

the samples (Supplemental Figs. S3-9). Representative DNA sequences were deposited in 440

GenBank (see below). 441

442

Cytological analyses 443

Lycopodiella inundata gametophytes (n = 15), young sporophytes (protocorms) (n = 444

15), and roots of mature plants (both wild (n = 20) and experimental (n = 20), roots of 445

angiosperms (Holcus lanatus, Molinia caerulea and Juncus bulbosus), and liverwort 446



17

gametophytes (Fossombronia foveolata) (30 samples for each species) were either stained 447

with trypan blue (Brundrett et al., 1996) and photographed under a Zeiss Axioscope (Zeiss, 448

Oberkochen, Germany) equipped with an MRc digital camera, or processed for scanning 449

electron microscopy (SEM) within 48 hrs of collection. For SEM, we followed the protocol of 450

Duckett et al., (2006) (see Supplemental Data). 451

452

Quantification of C, 33P and 15N fluxes between lycophytes and fungi 453

After the four-week acclimation period, microcosms were moved to individual drip-454

trays immediately prior to isotope labelling to avoid cross-contamination of the isotope tracers. 455

100 µl of an aqueous mixture of 33P-labelled orthophosphate (specific activity 111 TBq mmol-456

1, 0.3 ng 33P added; Hartmann analytics, Braunschweig, Germany) and 15N-ammonium 457

chloride (1 mg ml-1; 0.1 mg 15N added; Sigma, Dorset, UK) was introduced into one of the soil-458

filled mesh cores in each pot through the installed capillary tube (Supplemental Fig. S2a). In 459

half (n = 10) of the pots, cores containing isotope tracers were left static to preserve direct 460

hyphal connections with the lycophytes. Fungal access to isotope tracers was limited in the 461

remaining half (n = 10) of the pots by rotating isotope tracer-containing cores through 90°, 462

thereby severing the hyphal connections between the plants and core soil. These were rotated 463

every second day thereafter, thus providing a control treatment that allows us to distinguish 464

between fungal and microbial contributions to tracer uptake by plants. Assimilation of 33P 465

tracer into above-ground plant material was monitored using a hand-held Geiger counter held 466

over the plant material daily. 467

At detection of peak activity in above-ground plant tissues (21 days after the addition 468

of the 33P and 15N tracers), the tops of 33P and 15N-labelled cores were sealed with plastic caps 469

and anhydrous lanolin and the glass wool cores were sealed with rubber septa (SubaSeal, 470

Sigma, Dorset, UK). Before cabinet lights were turned on at 0800, each pot was sealed into a 471

3.5 l, gas-tight labelling chamber and 2 ml 10% (w/v) lactic acid added to 30 µl NaH14CO3 472

(specific activity 1.621 GBq/mmol-1; Hartmann Analytics, Braunschweig, Germany), releasing 473

a 1.1-MBq pulse of 14CO2 gas into the headspace of the labelling chamber (Supplemental Fig. 474



18

S2b). Pots were maintained under growth cabinet conditions, and 1 ml of headspace gas was 475

sampled after 1 h and every 1.5 h thereafter. Below-ground respiration was monitored via gas 476

sampling from within the glass-wool filled core after 1 h and every 1.5 h thereafter for ~16 h. 477

478

Plant harvest and sample analyses 479

Upon detection of maximum below-ground flux of 14C, approximately 16 h after the 480

release of the 14CO2 pulse, each microcosm compartment (i.e., plant material and soil) were 481

separated, freeze-dried, weighed, and homogenised. The 33P activity in plant and soil samples 482

was quantified by digesting in concentrated H2SO4 (see Supplemental Data) and liquid 483

scintillation (Tricarb 3100TR liquid scintillation analyser, Isotech, Chesterfield, UK). The 484

quantity of 33P tracer that was transferred to the plant by it’s fungal partner was then calculated 485

using previously published equations (Cameron et al., 2007) (see Supplemental Data). To 486

determine total symbiotic fungal-acquired 33P transferred to L. inundata, the mean 33P content 487

of plants which did not have access to the tracer because cores into which the 33P was 488

introduced were rotated, thus severing the hyphal connections between plant and core 489

contents, was subtracted from the total 33P in each plant that did have access to the isotopes 490

within the core via intact fungal hyphal connections where isotope-containing cores remained 491

static throughout the labelling period. This calculation controls for diffusion of isotopes and 492

microbial nutrient cycling in pots, ensuring only 33P gained by the plant via intact fungal hyphal 493

connections is accounted for and therefore serves as a conservative measure of the minimum 494

fungal transfer of tracer to the plant. 495

Between two and four mg of freeze-dried, homogenised plant tissue was weighed into 496

6 x 4 mm2 tin capsules (Sercon Ltd. Irvine, UK) and 15N abundance was determined using a 497

continuous flow IRMS (PDZ 2020 IRMS, Sercon Ltd. Irvine, UK). Air was used as the reference 498

standard, and the IRMS detector was regularly calibrated to commercially available reference 499

gases. The 15N transferred from fungus to plant was then calculated using equations published 500

previously (Field et al., 2016) (see Supplemental Data). In a similar manner as for the 33P 501

tracer, the mean of the total 15N in plants without access to the isotope because of broken 502



19

hyphal connections between plant and core contents was subtracted from the total 15N in each 503

plant with intact hyphal connections to the mesh-covered core to give fungal-acquired 15N. 504

Again, this provides a conservative measure of 15N transfer from fungus to plant as it ensures 505

only 15N gained by the plant via intact fungal hyphal connections is accounted for. 506

The 14C activity of plant and soil samples was quantified through sample oxidation (307 507

Packard Sample Oxidiser, Isotech, Chesterfield, UK) followed by liquid scintillation. Total C 508

(12C + 14C) fixed by the plant and transferred to the fungal network was calculated as a function 509

of the total volume and CO2 content of the labelling chamber and the proportion of the supplied 510

14CO2 label fixed by plants (see Supplemental Data). The difference in total C between the 511

values obtained for static and rotated core contents in each pot is considered equivalent to 512

the total C transferred from plant to symbiotic fungus within the soil core for that microcosm, 513

noting that a small proportion will be lost through soil microbial respiration. The total C budget 514

for each experimental pot was calculated using equations from Cameron et al., (2006) (see 515

Supplemental Data). Total percent allocation of plant-fixed C to extraradical symbiotic fungal 516

hyphae was calculated by subtracting the activity (Bq) of rotated core samples from that 517

detected in static core samples in each pot, dividing this by the sum of activity detected in all 518

components of each microcosm, then multiplying it by 100. 519

520

Stable isotope signatures of neighbouring plants 521

Lycopodiella inundata and J. bulbosus were collected from Thursley National Nature 522

Reserve, Surrey, together with co-occurring reference plants from six 1 m2 plots in May 2018, 523

following the sampling scheme of (Gebauer and Meyer, 2003). Five plant species representing 524

three different types of mycorrhizal associations served as reference plants: two ericoid 525

mycorrhizal species (Erica tetralix, collected on six plots; Calluna vulgaris, collected on three 526

plots), two ectomycorrhizal species (Pinus sylvestris and Betula pendula seedlings, both from 527

one plot), and one arbuscular mycorrhizal species (Molinia caerulea from six plots). Relative 528

C and N isotope natural abundances of dried and ground leaf and root samples were 529

measured in a dual element analysis mode with an elemental analyser (Carlo Erba 530



20

Instruments 1108, Milan, Italy) coupled to a continuous flow isotope ratio mass spectrometer 531

(delta S, Finnigan MAT, Bremen, Germany) via a ConFlo III open-split interface (Thermo 532

Fisher Scientific, Bremen, Germany) as described in Bidartondo et al., (2004). Relative isotope 533

abundances (δ values) were calculated, calibrated and checked for accuracy using methods 534

detailed in Supplemental Data. 535

536

Statistics 537

Isotope tracing data were checked for normality and differences between plant 538

assimilation of 33P and 15N were tested using Student’s t-test in SPSS Version 24 (IBM, New 539

York). Mean values are displayed in figures with standard error. Stable isotope patterns 540

between the groups of Lycopodiella inundata (n = 6), Juncus bulbosus (n = 6) and surrounding 541

angiosperms (n = 17) were tested for normality and equal variance. One-tailed Kruskal Wallis 542

test (Q) was applied for non-parametric data followed by Dunn’s post hoc procedure while 543

One-way ANOVA (F) was applied for parametric data followed by Tukey post hoc procedure 544

(q). Mean values are displayed in figures with standard deviation. 545

546

Accession numbers 547

Representative DNA sequences were deposited in GenBank, accession numbers MK673773-548

MK673803. 549

550

Supplemental Data 551

Supplemental Figure S1. Schematic diagram of mesh-covered core showing dimenions of 552

window (not drawn to scale). 553

Supplemental Figure S2. Schematic diagrams of experimental microcosms showing (a) 15N 554

and 33P and (b) 14C isotope tracing 555

Supplemental Figure S3. Phylogenetic relationships of Mucoromycotina OTUs associated 556

with Lycopodiella inundata grown for isotope tracing and cytology experiments in controlled 557

environment growth chambers. 558



21

Supplemental Figure S4. An overview of phylogenetic relationships of Mucoromycotina 559

OTUs associated with bryophytes, lycophytes and angiosperms from various UK locations 560

based on partial 18S gene sequences 561

Supplemental Figure S5. Phylogenetic relationships of partial 18S DNA sequences classified 562

as OTU 2, corresponding to “group A” in Desirò et al., (2013) 563


as OTU 1, 3 and 4. 565

Supplemental Figure S7. Phylogenetic relationships of partial 18S DNA sequences 566

clustering within OTU 5. 567


as OTU 6, corresponding to “group B” in Desirò et al. ( 2013). 569

Supplemental Figure S9. Phylogenetic relationships of partial 18S DNA sequences 570

clustering with OTU 7, corresponding to “group I” in Desirò et al. (2013). 571

Supplemental Figure S10. Micrographs showing morphology of fungal colonization in 572

Fossombronia foveolata and Holcus lanatus. 573

Supplemental Figure S11. Scanning electron micrographs, except (g) light micrograph of 574

toluidine blue stained semi-thin sections. Gametophyte morphologies in Lycopodiella inundata 575

(from Thursley Common) 576

Supplemental Table S1. Samples of lycophytes, liverworts and angiosperms analysed with 577

their origin. 578

Supplemental Table S2. A summary of Mucoromycotina OTUs associated with liverworts, 579

lycophytes and angiosperms at four UK sites. 580

Supplemental Table S3. A summary of the amounts of carbon, 15N and 33P detected in static 581

and rotated core of microcosms used during carbon-for-nutrient experiments between 582

Lycopodiella inundata and Mucoromycotina FRE fungi. 583

584

Acknowledgements 585

We gratefully acknowledge support from the NERC to KJF, SP, SS (NE/N00941X/1) and MIB 586



22

(NE/N009665/1) and the Leverhume Trust (PLP-2017-079). KJF is funded by BBSRC 587

Translational Fellowship (BB/M026825/1). WRR is supported by a NERC DTP (Science and 588

Solutions for a Changing Planet) studentship. We thank James Giles (Natural England) for 589

field support, Julia Masson and the RSPB for access to the Norfolk site, and The Species 590

Recovery Trust for access to the Dorset site. We are grateful to the anonymous reviewers of 591

this manuscript and to the editor for their constructive comments. 592

593

Table 1. Cytology of colonisation and fungal identity of study species (*) compared to 594

relevant examples from the literature that are referred to in the Discussion. *Results from 595

this study; G = gametophyte generation; S= sporophyte generation; ICSs = intercellular 596

spaces 597

Plant Group G/S Tissue/location Colonization Morphology (diameter) Fungus ID

References

Liverworts

Treubia1 G several ventral cell layers

intracellular coils (0.5-1.5 µm) with ‘lumps’/swellings (up to 15 µm), arbuscule-like short-side branches on coiled hyphae

M2 Duckett et al. 2006 Bidartondo et al. 2011

above intracellular zone

intercellular: large mucilage-filled ICSs

coarse hyphae 2-3 µm, thick-walled fungal structures

Fossombronia* G thallus central strand

intracellular coarse hyphae (2-3 µm); large vesicles (15-30 µm), coils (0.5-1µm), fine hyphae (0.5-1.5 µm) with small swellings/vesicles (5-10 µm), arbuscules

(M&G)*

Lycophytes

Lycopodiella* G outer cortical cell layers several ventral cell layers

intracellular intracellular

coils (up to 2.5µm) with vesicles (15-20µm) fine hyphae (0.5-1.5 µm) with small swellings/vesicles (5-10 µm)

M*

above intracellular zone

intercellular: large mucilage-filled ICSs

coarse hyphae (2->3 µm)

S protocorm: several ventral cell layers central, above intracellular zone

intracellular intercellular: large mucilage-filled ICSs

fine hyphae (0.5-1.5 µm) with small swellings/vesicles (5-10 µm) coarse hyphae (2->3 µm)

M*

S root intracellular and

fine hyphae (0.5-1.5 µm) with small

M1* Rimington et al. 2015



23

intercellular, small ICSs

swellings/vesicles (5-15 µm)

Angiosperms

Holcus* S root intracellular and intercellular, small ICSs

coarse hyphae (>3 µm), large vesicles (20-40 µm), fine hyphae (0.5-1.5 µm) with small vesicles (5-10 µm), arbuscules/arbuscule-like structures

(M&G)*

Molinia* S root intracellular and intercellular, small ICSs

coarse hyphae (>3 µm), large vesicles (20-40 µm), fine hyphae (0.5-1.5 µm) with small vesicles/swellings (5-10 µm), arbuscules/arbuscule-like structures

(M&G)*

Juncus* S root intracellular and intercellular, small ICSs

coarse hyphae (>3 µm), large vesicles (20-40 µm), fine hyphae (0.5-1.5 µm) with small vesicles (5-10 µm), arbuscules/arbuscule-like structures

(M&G)*

Trifolium1 S root intracellular and intercellular, small ICSs

coarse hyphae (>3 m),

large vesicles (>30 m)

fine hyphae (>1.5 m), intercalary and terminal vesicles/swellings (5-10

m) and arbuscules/arbuscule-like structures

(M&G)1 Orchard et al. 2017a

Fossils

Horneophyton1 S aerial axes, cortical cells

intracellular coarse hyphae (>3 m), large vesicles (up to 50

m), arbuscule-like structures

G1 Strullu-Derrien et al. 2014

corm intracellular and intercellular

intracellular coils, intercellular coarse

hyphae (11-13 m), thick-walled fungal structures

M1

Nothia1 S aerial and prostrate axes

Intercellular and intracellular

coarse hyphae (up to 15

m) and intercellular

vesicles (>50 m)

? Krings et al. 2007

598

Figure legends 599

Figure 1. Land plant phylogeny and species used in the present study. (a) Land plant 600

phylogeny showing key nodes alongside commonly associated fungal symbionts (Duckett and 601

Ligrone, 1992; Duckett et al., 2006; James et al., 2006; Bidartondo et al., 2011) (b) 602

Lycopodiella inundata at Thursley Common, Surrey, June 2017. 603

604



24

Figure 2. Figure 2. Carbon-for-nutrient exchange between Lycopodiella inundata and 605

Mucoromycotina fine root endophyte fungi. (a) Total plant-fixed carbon (C) transferred to 606

Mucoromycotina fine root endophyte (FRE) fungi by L. inundata. (b) Percent allocation of 607

plant-fixed C to Mucoromycotina FRE fungi (c) Total plant tissue phosphorus (33P) and 608

nitrogen (15N) content (ng) and (d) tissue concentration (ng g-1) of fungal-acquired 33P and 15N 609

in L. inundata tissue. In all panels, error bars denote SEM. In panels (a) and (b) n = 20. In 610

panels (c) and (d) n = 10. n is equivalent to the number of biological replicates used during 611

carbon-for-nutrient exchange experiements. Experiments were carried out three times. * 612

indicates where P <0.05, Student’s T-Test. 613

614

615

Figure 3. Carbon and nitrogen stable isotope natural abundance of Lycopodiella 616

inundata (L.i., n = 6), Juncus bulbosus (J.b., n = 6) and surrounding angiosperms 617

(arbuscular mycorrhizal: Molinia caerulea (M.c., n = 6); ectomycorrhizal: Pinus sylvestris (P.s., 618

n = 1); Betula pendula (B.p., n = 1); ericoid mycorrhizal: Calluna vulgaris (C.v., n = 3), Erica 619

tetralix (E.t., n = 6)) for leaf (A) and root (B) samples, respectively. Mean values are displayed 620

with standard deviation. One-tailed Kruskal Wallis test, followed by Dunn’s post hoc, found 621

significant differences between L. inundata, J. bulbosus and surrounding angiosperms as 622

references in leaf carbon and nitrogen stable isotope natural abundance and in root nitrogen 623

stable isotope natural abundance. 624

625

Figure 4. Light micrographs of trypan blue stained tissues. (a) Branching fine hyphae with 626

small swellings/vesicles in thallus cells and rhizoid (b) of the liverwort Fossombronia foveolata 627

(from Thursely Common) colonized by both Mucoromycotina fine root endophytes (FREs) and 628

Glomeromycotina, in (b) also note the coarse hyphae (arrowhead). (c-e) Fine hyphae with 629

small swellings/vesicles in the root hairs and root cells of the lycophyte Lycopodiella inundata 630

colonized by Mucoromycotina FREs only (field-collected specimens from Thursley Common). 631

(f) Fine hyphae with small swellings/vesicles and large vesicles in a root of the grass Holcus 632



25

lanatus (from Lynn Crafnant) colonized by both Mucoromycotina FREs and Glomeromycotina. 633

(g-h) Roots of the grass Molinia caerulea (from Thursley Common) colonized by both 634

Mucoromycotina FREs and Glomeromycotina, showing fine hyphae (g) and coarse hyphae 635

with large vesicles (h). Scale bars: (a, b, d-f) 50 µm, (c, g, h) 100 µm. 636

637

Figure 5. Scanning electron micrographs. (a) Fine hyphae (arrows) with a small 638

swelling/vesicle (*) in the thallus cells of Fossombronia foveolata (from Thursley Common), 639

also note the much coarser hyphae (arrowheads). (b-g) Fungal colonization in Lycopodiella 640

inundata. (b, c) Intercalary (b) and terminal (c) small swellings/vesicles on fine hyphae in the 641

ventral cell layers of a protocorm (from Thursley Common). Centrally and above this 642

intracellular colonization zone, the fungus becomes exclusively intercellular, as evidenced by 643

the presence of abundant, tightly-appressed hyphae surrounding the central protocorm cells 644

(arrows) (d) and eventually completely fills the large, mucilage-filled intercellular spaces 645

present in this zone (e, *). (f, g) Cross sections of roots of experimental plants; (f) several cells 646

colonized exclusively (*) by branching fine hyphae with small swellings/vesicles, enlarged in 647

(g). (h, i) Cross sections of roots of Juncus bulbosus (from Thursley Common) showing fine 648

(arrow) and coarse (arrowheads) hyphae (h) and a fine hypha with small swellings/vesicles 649

(i). Scale bars: (a, d, g, i) 20 µm, (b, c, h) 10 µm, (e) 100 µm, (f) 50 µm. 650

651

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795



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Figure 4.

Parsed CitationsAlbornoz FE, Lambers H, Turner BL, Teste FP, Laliberté E (2016) Shifts in symbiotic associations in plants capable of forming multipleroot symbioses across a long‐term soil chronosequence. Ecology and Evolution 6: 2368-2377

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Cernusak LA, Tcherkez G, Keitel C, Cornwell WK, Santiago LS, Knohl A, Barbour MM, Williams DG, Reich PB, Ellsworth DS (2009) Whyare non-photosynthetic tissues generally 13C enriched compared with leaves in C3 plants? Review and synthesis of currenthypotheses. Functional Plant Biology 36: 199-213


Dawson TE, Mambelli S, Plamboeck AH, Templer PH, Tu KP (2002) Stable isotopes in plant ecology. Annual Review of Ecology andSystematics 33: 507-559


Desirò A, Duckett JG, Pressel S, Villarreal JC, Bidartondo MI (2013) Fungal symbioses in hornworts: a chequered history. Proceedingsof the Royal Society of London B: Biological Sciences 280: 20130207


Duckett JG, Carafa A, Ligrone R (2006) A highly differentiated glomeromycotean association with the mucilage-secreting, primitiveantipodean liverwort Treubia (Treubiaceae): clues to the origins of mycorrhizas. American Journal of Botany 93: 797-813


Duckett JG, Ligrone R (1992) A light and electron microscope study of the fungal endophytes in the sporophyte and gametophyte ofLycopodium cernuum with observations on the gametophyte–sporophyte junction. Canadian Journal of Botany 70: 58-72


Edgar R (2016) UCHIME2: improved chimera prediction for amplicon sequencing. bioRxiv: 074252Pubmed: Author and TitleGoogle Scholar: Author Only Title Only Author and Title

Farquhar GD, Ehleringer JR, Hubick KT (1989) Carbon isotope discrimination and photosynthesis. Annual Review of Plant Biology 40:503-537


Farquhar GD, O'Leary MH, Berry JA (1982) On the relationship between carbon isotope discrimination and the intercellular carbondioxide concentration in leaves. Functional Plant Biology 9: 121-137


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https://www.ncbi.nlm.nih.gov/pubmed?term=Cernusak LA%2C Tcherkez G%2C Keitel C%2C Cornwell WK%2C Santiago LS%2C Knohl A%2C Barbour MM%2C Williams DG%2C Reich PB%2C Ellsworth DS %282009%29 Why are non%2Dphotosynthetic tissues generally 13C enriched compared with leaves in C3 plants%3F Review and synthesis of current hypotheses%2E Functional Plant Biology&dopt=abstract

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https://www.ncbi.nlm.nih.gov/pubmed?term=Orchard S%2C Hilton S%2C Bending GD%2C Dickie IA%2C Standish RJ%2C Gleeson DB%2C Jeffery RP%2C Powell JR%2C Walker C%2C Bass D %282017a%29 Fine endophytes %28Glomus tenue%29 are related to Mucoromycotina%2C not Glomeromycota%2E New Phytologist&dopt=abstract

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https://scholar.google.com/scholar?as_q=Fine root endophytes under scrutiny: a review of the literature on arbuscule-producing fungi recently suggested to belong to the Mucoromycotina&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Orchard&as_ylo=2017&as_allsubj=all&hl=en&c2coff=1

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https://scholar.google.com/scholar?as_q=The agronomic relevance of arbuscular mycorrhizas in the fertility of Australian extensive cropping systems&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Ryan&as_ylo=2012&as_allsubj=all&hl=en&c2coff=1

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https://www.ncbi.nlm.nih.gov/pubmed?term=Smith SE%2C Anderson IC%2C Smith FA %282015%29 Mycorrhizal associations and phosphorus acquisition%3A from cells to ecosystems%2E Annual Plant Reviews Online&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Smith&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=Mycorrhizal associations and phosphorus acquisition: from cells to ecosystems&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

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https://www.ncbi.nlm.nih.gov/pubmed?term=Smith SE%2C Smith FA %282011%29 Roles of arbuscular mycorrhizas in plant nutrition and growth%3A new paradigms from cellular to ecosystem scales%2E Annual Review of Plant Biology&dopt=abstract

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https://scholar.google.com/scholar?as_q=Fungal associations in Horneophyton ligneri from the Rhynie Chert (c&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Strulluâ�?�Derrien&as_ylo=2014&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=von Oheimb G%2C Power SA%2C Falk K%2C Friedrich U%2C Mohamed A%2C Krug A%2C Boschatzke N%2C H�rdtle W %282010%29 N%3A P ratio and the nature of nutrient limitation in Calluna%2Ddominated heathlands%2E Ecosystems&dopt=abstract

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https://scholar.google.com/scholar?as_q=N: P ratio and the nature of nutrient limitation in Calluna-dominated heathlands&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=von&as_ylo=2010&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Walker C%2C Gollotte A%2C Redecker D %282018%29 A new genus%2C Planticonsortium %28Mucoromycotina%29%2C and new combination %28P%2E tenue%29%2C for the fine root endophyte%2C Glomus tenue %28basionym Rhizophagus tenuis%29%2E Mycorrhiza&dopt=abstract

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https://scholar.google.com/scholar?as_q=A new genus, Planticonsortium (Mucoromycotina), and new combination (P.&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

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Short Title: Vascular plant-Mucoromycotina fungal symbiosis · 65 Glomeromycotina fungi in the same plant host (Desirò et al., 2013; Rimington et al., 2015). 66 Mucoromycotina represents