Lichen takes more than two to tango


Editor's Introduction

Basidiomycete yeasts in the cortex of ascomycete macrolichens

Lichens are one of the oldest known examples of a symbiotic organism. Lichens are made up of a fungus, which provides structure, and an alga, which provides energy via photosynthesis. However, attempts in the laboratory to re-create a lichen using only these two partners were never successful. In this work, the researchers combine modern techniques to discover a third symbiont that builds the outer cortex of the lichen.

Paper Details

Original title
Basidiomycete yeasts in the cortex of ascomycete macrolichens
Original publication date
Issue name


For over 140 years, lichens have been regarded as a symbiosis between a single fungus, usually an ascomycete, and a photosynthesizing partner. Other fungi have long been known to occur as occasional parasites or endophytes, but the one lichen–one fungus paradigm has seldom been questioned. Here we show that many common lichens are composed of the known ascomycete, the photosynthesizing partner, and, unexpectedly, specific basidiomycete yeasts. These yeasts are embedded in the cortex, and their abundance correlates with previously unexplained variations in phenotype. Basidiomycete lineages maintain close associations with specific lichen species over large geographical distances and have been found on six continents. The structurally important lichen cortex, long treated as a zone of differentiated ascomycete cells, appears to consistently contain two unrelated fungi.

Video. Studies of gene activity have now revealed that many lichens are not a twosome but instead a threesome, with two fungi in the mix (Courtesy Science Magazine).



Most definitions of the lichen symbiosis emphasize its dual nature: the mutualism of a single fungus and single photosynthesizing symbiont, occasionally supplemented by a second photosynthesizing symbiont in modified structures (14). Together, these organisms form stratified, often leafy or shrubby body plans (thalli) that resemble none of the symbionts in isolation, a feature thought to be unique among symbioses (1). Attempts to synthesize lichen thalli from the accepted two components in axenic conditions, however, have seldom produced structures that resemble natural thalli (56). Notably, a critical structural feature of stratified lichens, the cortex, typically remains rudimentary in laboratory-generated symbioses (5). Recently, it has been suggested that microbial players, especially bacteria, may play a role in forming complete, functioning lichen thalli (7). However, although culturing and amplicon sequencing have revealed rich communities of microbes (78), including other fungi (810), no new stably associated symbiotic partners have been found.

The recalcitrance of lichens to form thalli in vitro means that characterizing symbiont gene activity (e.g., through transcriptomics) requires an approach that works with natural thalli. We used metatranscriptomics to better understand the factors involved in forming two macrolichen symbioses, Bryoria fremontii and B. tortuosa. These two species have been distinguished for 90 years by the thallus-wide production of the toxic substance vulpinic acid in B. tortuosa, causing it to appear yellowish, in contrast to B. fremontii, which is dark brown (11). Recent phylogenetic analyses have failed to detect any fixed sequence differences between the two species in either the mycobiont (Ascomycota, Lecanoromycetes, Bryoria) or the photobiont (Viridiplantae, Trebouxia simplex) when considering four and two loci, respectively (1213). We hypothesized that differential gene expression might account for the increased production of vulpinic acid in B. tortuosa.

We first selected 15 thalli (six from B. fremontii and nine from B. tortuosa, all free from visible parasitic infection) from sites across western Montana, USA, for mRNA transcriptome sequencing. An initial transcriptome-wide analysis of single nucleotide polymorphisms (SNPs) for Ascomycota and Viridiplantae transcript subsets showed no correlation between genotype and phenotype in B. fremontii and B. tortuosa, confirming previous results (1213) (Fig. 1, A and B). Next, we estimated transcript abundances by mapping raw reads back to a single, pooled metatranscriptome assembly and binning by taxon. Restricting our analyses to Ascomycota and Viridiplantae revealed little differential transcript abundance associated with phenotype (Fig. 1, C and E). Taken together, these analyses confirm previous conclusions that the two lichen species are nomenclatural synonyms (12) but still provide no explanation for the underlying phenotypes (which we shall continue to refer to by their species names for convenience). However, by expanding the taxonomic range to consider all Fungi, we found 506 contigs with significantly higher abundances in vulpinic acid–rich B. tortuosa thalli. A majority of these contigs were annotated as Basidiomycota (Fig. 1D). These data suggested that a previously unrecognized basidiomycete was present in thalli of both species but was more abundant whenever vulpinic acid was present in large amounts.

Fig. 1 Genome-wide divergence and transcript abundance of fungi and algae, based on symbiont subsets extracted from wild Bryoria metatranscriptomes. (A and B) Unrooted maximum likelihood topologies for (A) the Ascomycota member (lecanoromycete) and (B) the Viridiplantae member (alga) within the lichen pair B. fremontii and B. tortuosa, based on 30,001 and 25,788 SNPs, respectively. Numbers refer to metatranscriptome sample IDs (table S2). Scale bars indicate the average number of substitutions per site. (C to E) Logarithm of the fold change (logFC) between vulpinic acid–deficient (B. fremontii) and vulpinic acid–rich (B. tortuosa) phenotypes in 15 Bryoria metatranscriptomes, plotted against transcript abundance (logCPM, logarithm of counts per million reads). Only transcripts found in all 15 samples were included. Ascomycota transcripts only are shown in (C). All fungal transcripts are shown in (D), with taxonomic assignments superimposed; a plot with statistically significant transcript differential abundance is shown as an inset. Viridiplantae transcripts are shown in (E). Red dots indicate a log fold change with P < 0.05 in (C), (E) (highlighted with arrows), and the inset of (D).

Panels A and B Methods

Phylogenetic trees are used to determine evolutionary relationships between organisms. When a new organism is found, researchers extract gene sequences from the organism to determine “who” it is by comparing the sequence to known organisms. In this phylogeny, the brown circles represent the fungal/algal partner in the Bryoria fremontii (vulpinic acid ) and the yellow circles represent the fungal/algal partner in B. tortuosa (vulpinic acid +). This phylogeny is considered unrooted because all the taxa in the phylogeny are from the same group, and we can’t determine who would be most closely related to a last common ancestor. An unrooted phylogeny is useful for determining how closely related a group of organisms is.

Questions for Panels A and B
  1. Do the trees in panels A and B have the same topology? Explain.
  2. If there were a correlation between a genotype (mycobiont and photobiont genetic identity) and phenotype (production of vulpinic acid), how would you expect trees in Figures 1A and 1B to look? Draw.
  3. Based on the two trees, do you think the association between mycobiont and photobiont happened only once or multiple times?
Panels C, D, and E Methods

A transcriptome is a way to count the number of copies of each gene that is being made. To compare between different samples these counts are "normalized." To normalize, researchers divide the number of copies of the gene by 1 million and then compare this number between the two samples of interest. On the x-axis of these graphs you can see the difference in gene counts between our two samples; on the y-axis you can see how often that gene was found. 

These slides describe the methods behind transcriptome analysis.

Questions for Panels C, D, and E
  1. What does each dot in Figures C, D, and E represent?
  2. What do dots colored in orange (p ≤ 0.05) in Figures 1C and 1E represent?
  3. On panel C, where would you expect to find a hypothetical gene upregulated 100-fold in vulpinic acid-rich lichen?
  4. What additional data is plotted on Figure 1D in comparison to Figure 1C?
  5. What does the cloud of yellow points in Figure 1D signify? Do they correspond to genes that show significant differential expression?

We next sought to determine whether this uncharacterized basidiomycete was specific to the studied Bryoria species or could be found in other lichens. From metatranscriptome contigs containing ribosomal RNA (rRNA) basidiomycete sequences, we designed specific primers for ribosomal DNA [rDNA; 18S, internal transcribed spacer (ITS), and D1D2 domains of 28S) to screen lichens growing physically adjacent to Bryoria in Montana forests. Each assayed lichen species carried a genetically distinct strain of the basidiomycete, indicating a high degree of specificity. Furthermore, we found that Letharia vulpina, a common lichen species growing intermixed with Bryoria, maintained basidiomycete genotypes that were distinct from those in Bryoria, not only in Montana but also in northern Europe (fig. S1). When assaying for the basidiomycete across the seven main radiations of macrolichens in the class Lecanoromycetes, we found related basidiomycete lineages associated with 52 lichen genera from six continents, including in 42 of 56 sampled genera of the family Parmeliaceae (fig. S2). As a whole, these data indicate that basidiomycete fungi are ubiquitous and global associates of the world’s most speciose radiation (14) of macrolichens.

To place the basidiomycete lineages in a phylogenetic context, we generated a 349-locus phylogenomic tree by using gene sequences inferred from our transcriptome data set and other available genomes (table S1). This analysis placed the basidiomycete as sister to Cystobasidium minutum (class Cystobasidiomycetes, subphylum Pucciniomycotina) with high support (Fig. 2A). The only previously known lichen-associated members of Cystobasidiomycetes are two species of Cyphobasidium, which is hypothesized to cause galls on species of Parmeliaceae (15). When incorporated into a broader sample of published cystobasidiomycete rDNA sequence data (1618), the majority of our lichen-derived sequences form a strongly supported monophyletic clade with Cyphobasidium (Fig. 2B). Using current classification criteria (18), the lichen-associated lineages would include numerous new family-level lineages, and we recognize this set of subclades as the new order Cyphobasidiales (19). Applying a relaxed molecular clock to our phylogenomic tree (Fig. 2A) shows the Cystobasidium-Cyphobasidium split occurring around the same time as the origin of three of the main groups of lecanoromycete macrolichens in which Cyphobasidiales species occur, suggesting a long, shared evolutionary history. Two fossil calibrations place this split at around 200 million years before the present (figs. S4 and S5).

Fig. 2 Placement of Cyphobasidiales members and their diversity within lichens. (A) Maximum likelihood phylogenomic tree based on 39 fungal proteomes and 349 single-copy orthologous loci. Dating based on a 58-locus subsample shows relative splits between Cyphobasidiales and Cystobasidium minutum and splits leading to the lecanoromycete genera XanthoriaCladonia, and Bryoria (colored bars indicate 95% confidence intervals; fungi occurring in lichens are shown in green). (B) Maximum likelihood rDNA phylogeny of the class Cystobasidiomycetes, with images of representative lichen species from which sequences were obtained in each clade; thick branches indicate bootstrap support >70%. Shaded triangles are scaled to the earliest branch splits of underlying sequence divergence in each clade. Full versions of the trees are shown in fig. S3.

Rooted Trees

This is a rooted phylogeny of the Basidiomycota (group of the new fungus) and the Ascomycota (group of the old fungus). In a rooted phylogeny, an extra group of distinct or distantly related organisms is added to the phylogeny to act as an out-group. The long branch that connects the out-group to main group is the root of the tree. The root indicates the position of the last common ancestor. A rooted phylogeny is a diagram that looks back in time. The tips of the phylogeny represent modern-day organisms. Nodes (where branches separate) represent a shared ancestor between two or more modern-day organisms. This phylogeny is “calibrated” with dates by assuming a molecular clock.  This means that we assume changes (mutations) happen at a constant given rate. We can use fossil dating of some of the species in the phylogeny to guess when the events depicted in the tree occurred.

Bootstrap Values

Bootstrap support is a way of testing how much evidence there is to support the joining of 2 or more modern-day organisms, represented by a node in the phylogeny. Each node is a hypothesis about the evolutionary pathway of each organism, and all this information is based on similarity or dissimilarity of gene sequences. High bootstrap support values indicate that there is a lot of evidence in the gene sequences that a group of organisms shared an ancestor in the way depicted on the phylogeny. Low bootstrap support values indicate there is not enough evidence to confirm or deny that the group of organisms shared an ancestor.


Two good resources for understanding how to interpret phylogenetic trees are:

CrashCourse on Phylogenies, including some excellent scientific history:

HHMI BioInteractive slides:…

Figure 2 Questions
  1. Is Figure 2A a rooted phylogenetic tree?
  2. Is Figure 2A scaled with respect to time?
  3. How many fossil calibrations did the author used in the phylogenetic analyses of Figure 2A?
  4. Do fungi known to occur in lichen form a monophyletic group?
  5. What is the difference between the data used to reconstruct phylogenies in Figures 2A and 2B?
  6. Why did researchers need to reconstruct the tree shown in Figure 2B after they obtained the tree in Figure 2A? How do the authors use phylogeny in Figure 2B to infer that the Bryoria's unknown fungi belong to genus Cyphobasidium?
  7. What evidence shown in Figure 2B did authors use to make claim that Cyphobasidiales form a "strongly supported monophyletic clade?"

Our initial microscopic imaging failed to reveal any cells that we could assign to Basidiomycetes with certainty. Furthermore, attempts to culture the basidiomycete from fresh thalli were unsuccessful. We therefore developed protocols for fluorescent in situ hybridization (FISH) targeting specific ascomycete and cystobasidiomycete rRNA sequences. Cystobasidiomycete-specific FISH probes unambiguously hybridized round, ~3- to 4-μm-diameter cells embedded in the peripheral cortex of both B. fremontii and B. tortuosa (Fig. 3and movie S1). Consistent with the transcript abundance data, these cells were more abundant in thalli of B. tortuosa (Fig. 3), where they were embedded in secondary metabolite residues (movie S1). Imaging of other lichen species likewise revealed cells of similar morphology in the peripheral cortex (fig. S6). Composite three-dimensional FISH images from B. capillaris show the cells occurring in a zone exterior to the lecanoromycete (Fig. 4 and movie S2) and embedded in polysaccharides (Fig. 4C), explaining why these cells are not observed in scanning electron microscopy (Fig. 4A). In some species, such as L. vulpina, the abundance of hybridized living cells was low, but selective removal of the polysaccharide layer through washing revealed high densities of collapsed, apparently dead cells within the cortex (fig. S7). These dead cells may explain the paucity of the FISH signal in some experiments. The mononucleate single cells (fig. S8C), evidence of budding, and absence of hyphae or clamp connections are consistent with an anamorphic or yeast state in Cystobasidiomycetes. FISH imaging of Cyphobasidium galls on the lichen Hypogymnia physodes, obtained from Norway, confirmed the link to the sexual or teleomorphic state (fig. S8), which appears to develop rarely (15). These data suggest that the gall-inducing form of Cyphobasidium completes its life cycle entirely within lichens.

Fig. 3 Differential abundance of Cyphobasidiales yeasts in B. fremontii and B. tortuosa. (AB. fremontii, with (B) few FISH-hybridized live yeast cells at the level of the cortex. (CB. tortuosa, with (D) abundant FISH-hybridized cortical yeast cells (scale bars, 20 μm).


Fluorescent in situ hybridization (FISH): This is a molecular technique that enables researchers to test if something is present and find out where it is in the sample. Researchers design probes, usually from DNA, that are complementary to a sequence of interest. On the end of the probe is a fluorescent tag. Researchers can incubate their samples with these probes, let the probes bind with the complementary sequence, then wash off loose probes and look for a fluorescent signal to determine if a molecule is present and where it can be found. The green dots in panel three show the probes fluorescing a green light, indicating the presence of the basidiomycete.

FISH is described by Nature Education here:

Video Animation

Animated video for Panel D, showing the overlapping occurrence of vulpinic acid (colored purple) and the basidiomycete yeast cells (green).

Link below will begin download:

Figure 3 Questions
  1. What do green dots in Figures 3B and 3D represent?
  2. Is there a correlation between two strikingly different phenotypes and the presence of yeast cells? 
  3. Do data shown in Figure 3 support or contradict data shown in Figure 1D? Explain.

Fig. 4 Fluorescent cell imaging of dual fungal elements in lichen thalli. (A) Scanning electron microscopy image of a thallus filament of B. capillaris(scale bar, 200 μm). (B) FISH hybridization of B. capillaris thallus, showing Cyphobasidiales yeasts (green) and the lecanoromycete (blue) with algal chlorophyll A autofluorescence (red). The volume within the two vertical lines is visualized on the right; the unclipped frontal view is shown at the top. Movie S2 shows an animation of the three-dimensional ~100-μm z-stack. (C) Detail of yeast cells (scale bar, 5 μm).

Panel A Methods

Scanning electron microscopy (SEM) uses a powerful microscope that uses electrons to get the most detailed possible image of the surface of the sample. Most microscopes use light bouncing off a sample, but this technique limits what is seen due to the power of the lenses and the users’ eyes. A computer can create a highly-detailed image of cell surfaces by bouncing a focused beam of electrons off a sample, and measuring the direction and magnitude of the electron travel. You can see the resolution of this image is phenomenal. The tiny structures on the outside of the thallus are polysaccharides from the cortex.

Panels B and C Methods

Fluorescent in situ hybridization (FISH) can be used with multiple probes. Each probe is designed to stick to a different target and then tagged with different color fluorescent tags. In this figure, the newly discovered Cyphobasidiales is tagged with a green fluorophore, and the lecanoromycete is tagged with a blue fluorophore. The red in the picture is the photosynthetic pigments of the algae. These are activated the same way that the fluorescent tags are so there is no need to use a special probe on them for this experiment. Researchers need to take pictures from many different angles to figure out exactly how close different cell types are in the thallus.

Video Animation

Animated video for Panel D, showing the overlapping occurrence of vulpinic acid (colored purple) and the basidiomycete yeast cells (green).

Link below will begin download:

Figure 4 Questions
  1. What do green dots in Figures 3B and 3D represent?
  2. Is there a correlation between two strikingly different phenotypes and the presence of yeast cells? 
  3. Do data shown in Figure 3 support or contradict data shown in Figure 1D? Explain.

It is remarkable that Cyphobasidium yeasts have evaded detection in lichens until now, despite decades of molecular and microscopic studies of the Parmeliaceae (2022). It seems likely that the failure to detect Cyphobasidium in both Sanger and amplicon sequencing studies (8) is due to multi-template polymerase chain reaction bias. The most widespread clade of Cyphobasidium possesses a 595–base pair group I intron inserted downstream of the primer binding site ITS1F, doubling the template length of ITS, a popular fungal barcode (23). This, combined with low background abundance, can push a template below detection thresholds (24). Also, we cannot rule out that Cyphobasidium yeasts have actually been sequenced and discarded as presumed contaminants.

The lichen cortex layer has long been considered to be key for structural stabilization of macrolichens, as well as for water and nutrient transfer into the thallus interior (625). Most macrolichens possess a basic two-layer cortex scheme consisting of conglutinated internal hyphae and a thin, polysaccharide-rich peripheral layer (25). However, the internal cellular structure is not uniform across lichens (26), and the composition of extracellular polysaccharides is poorly known (27). In Bryoria, the layer in which Cyphobasidium yeasts occur has not been recognized as distinct from the cortex (11), although in other parmelioid lichens, a seemingly homologous layer has sometimes been referred to as the “epicortex” (20). The discovery of ubiquitous yeasts embedded in the cortex raises the prospect that more than one fungus may be involved in its construction, and it could explain why lichens synthesized in vitro from axenically grown ascomycete and algal cultures develop only rudimentary cortex layers (5).

In many lichens, the peripheral cortex layer in which Cyphobasidium yeasts are embedded is enriched with specific secondary metabolites (25), the production of which often does not correlate with the lecanoromycete phylogeny (28). The assumption that these substances are exclusively synthesized by the lecanoromycete must now be considered untested. In B. fremontii, differential transcript and cell abundance data, along with physical adjacency to crystalline residues, implicate Cyphobasidium in the production of vulpinic acid, either directly or by inducing its synthesis by the lecanoromycete. Confirming a link by using transcriptome or genome data is impossible until the enzymatic synthesis pathway of vulpinic acid is described. However, related pulvinic acid derivatives are synthesized by other members of Basidiomycota (29).

The assumption that stratified lichens are constructed by a single fungus with differentiated cell types is so central to the definition of the lichen symbiosis that it has been codified into lichen nomenclature (30). This definition has brought order to the field, but may also have constrained it by forcing untested assumptions about the true nature of the symbiosis. We suggest that the discovery of Cyphobasidium yeasts should change expectations about the potential diversity and ubiquity of organisms involved in one of the oldest known and most recognizable symbioses in science.

Supplementary Materials

Materials and Methods

Figs. S1 to S16

Tables S1 to S12

References (3174)

Movies S1 and S2

References and Notes

  1. A. De Bary, Die Erscheinung der Symbiose (Verlag Karl Trübner, 1879).
  2. A. Gargas, P. T. DePriest, M. Grube, A. Tehler, Multiple origins of lichen symbioses in fungi suggested by SSU rDNA phylogeny. Science 268, 1492–1495 (1995).
  3. F. Lutzoni, M. Pagel, V. Reeb, Major fungal lineages are derived from lichen symbiotic ancestors. Nature 411, 937–940 (2001).
  4. D. L. Hawksworth, The variety of fungal-algal symbioses, their evolutionary significance, and the nature of lichens. Bot. J. Linn. Soc. 96, 3–20 (1988).
  5. V. Ahmadjian, The Lichen Symbiosis (John Wiley & Sons, 1993).
  6. R. Honegger, Developmental biology of lichens. New Phytol. 125, 659–677 (1993).
  7. I. A. Aschenbrenner, T. Cernava, G. Berg, M. Grube, Understanding microbial multi-species symbioses. Front. Microbiol. 7, 180 (2016).
  8. S. T. Bates, D. Berg-Lyons, C. L. Lauber, W. A. Walters, R. Knight, N. Fierer, A preliminary survey of lichen associated eukaryotes using pyrosequencing. Lichenologist 44, 137–146 (2012).
  9. O. Petrini, U. Hake, M. M. Dreyfuss, An analysis of fungal communities isolated from fruticose lichens. Mycologia 82, 444–451 (1990).
  10. J. M. U’Ren, F. Lutzoni, J. Miadlikowska, A. E. Arnold, Community analysis reveals close affinities between endophytic and endolichenic fungi in mosses and lichens. Microb. Ecol. 60, 340–353 (2010).
  11. I. M. Brodo, D. L. Hawksworth, Alectoria and allied genera in North America. Opera Bot. 42, 1–164(1977).
  12. S. Velmala, L. Myllys, P. Halonen, T. Goward, T. Ahti, Molecular data show that Bryoria fremontii and B. tortuosa (Parmeliaceae) are conspecific. Lichenologist 41, 231–242 (2009).
  13. H. Lindgren, S. Velmala, F. Högnabba, T. Goward, H. Holien, L. Myllys, High fungal selectivity for algal symbionts in the genus Bryoria. Lichenologist 46, 681–695 (2014).
  14. G. Amo de Paz, P. Cubas, P. K. Divakar, H. T. Lumbsch, A. Crespo, Origin and diversification of major clades in parmelioid lichens (Parmeliaceae, Ascomycota) during the Paleogene inferred by Bayesian analysis. PLOS ONE 6, e28161 (2011).
  15. A. M. Millanes, P. Diederich, M. Wedin, Cyphobasidium gen. nov., a new lichen-inhabiting lineage in the Cystobasidiomycetes (Pucciniomycotina, Basidiomycota, Fungi). Fungal Biol., 10.1016/j.funbio.2015.12.003 (2015).
  16. M. C. Aime, P. B. Matheny, D. A. Henk, E. M. Frieders, R. H. Nilsson, M. Piepenbring, D. J. McLaughlin,L. J. Szabo, D. Begerow, J. P. Sampaio, R. Bauer, M. Weiss, F. Oberwinkler, D. Hibbett, An overview of the higher level classification of Pucciniomycotina based on combined analyses of nuclear large and small subunit rDNA sequences. Mycologia 98, 896–905 (2006).
  17. Q.-M. Wang, M. Groenewald, M. Takashima, B. Theelen, P.-J. Han, X.-Z. Liu, T. Boekhout, F.-Y. Bai,Phylogeny of yeasts and related filamentous fungi within Pucciniomycotina determined from multigene sequence analyses. Stud. Mycol. 81, 27–53 (2015).
  18. Q.-M. Wang, A. M. Yurkov, M. Göker, H. T. Lumbsch, S. D. Leavitt, M. Groenewald, B. Theelen, X.-Z. Liu,T. Boekhout, F.-Y. Bai, Phylogenetic classification of yeasts and related taxa within Pucciniomycotina.Stud. Mycol. 81, 149–189 (2015).
  19. Cyphobasidiales T. Sprib. & H. Mayrhofer, ord. nov. (MB 816120); diagnosis is the same as type family Cyphobasidiaceae T. Sprib. & H. Mayrhofer, fam. nov. (MB 816119); embedded in lichen thalli; teleomorph filamentous, rarely observed; when fertile, basidia develop thick-walled probasidium and thin-walled, cylindrical meiosporangium; anamorph is the prevalent known form, consisting of budding yeast with round, thin-walled cells, 2.5 to 4.5 μm in diameter, embedded in the upper cortex of lichens, especially Parmeliaceae; exogenous compound utilization not characterized; cell wall constituents unknown. Type genus, Cyphobasidium Millanes, Diederich, and Wedin, Fungal Biology doi:10.1016/j.funbio.2015.12.003, p. 4 (2015).
  20. M. E. Hale Jr., Pseudocyphellae and pored epicortex in the Parmeliaceae: Their delimitation and evolutionary significance. Lichenologist 13, 1–10 (1981).
  21. A. Thell, A. Crespo, P. K. Divakar, I. Kärnefelt, S. D. Leavitt, H. T. Lumbsch, M. R. D. Seaward, A review of the lichen family Parmeliaceae – history, phylogeny and current taxonomy. Nord. J. Bot. 30, 641–664(2012).
  22. A. Crespo, F. Kauff, P. K. Divakar, R. del Prado, S. Pérez-Ortega, G. Amo de Paz, Z. Ferencova, O.Blanco, B. Roca-Valiente, J. Núñez-Zapata, P. Cubas, A. Argüello, J. A. Elix, T. L. Esslinger, D. L.Hawksworth, A. Millanes, C. Molina, M. Wedin, T. Ahti, A. Aptroot, E. Barreno, F. Bungartz, S. Calvelo, M. Candan, M. Cole, D. Ertz, B. Goffinet, L. Lindblom, R. Lücking, F. Lutzoni, J.-E. Mattsson, M. I. Messuti, J.Miadlikowska, M. Piercey-Normore, V. J. Rico, H. J. M. Sipman, I. Schmitt, T. Spribille, A. Thell, G. Thor, D. K. Upreti, H. T. Lumbsch, Phylogenetic generic classification of parmelioid lichens (Parmeliaceae, Ascomycota) based on molecular, morphological and chemical evidence. Taxon 59, 1735–1753 (2010).
  23. C. L. Schoch, K. A. Seifert, S. Huhndorf, V. Robert, J. L. Spouge, C. A. Levesque, W. Chen, E. Bolchacova, K. Voigt, P. W. Crous, A. N. Miller, M. J. Wingfield, M. C. Aime, K.-D. An, F.-Y. Bai, R. W. Barreto, D. Begerow, M.-J. Bergeron, M. Blackwell, T. Boekhout, M. Bogale, N. Boonyuen, A. R. Burgaz, B.Buyck, L. Cai, Q. Cai, G. Cardinali, P. Chaverri, B. J. Coppins, A. Crespo, P. Cubas, C. Cummings, U. Damm, Z. W. de Beer, G. S. de Hoog, R. Del-Prado, B. Dentinger, J. Dieguez-Uribeondo, P. K. Divakar, B.Douglas, M. Duenas, T. A. Duong, U. Eberhardt, J. E. Edwards, M. S. Elshahed, K. Fliegerova, M. Furtado,M. A. Garcia, Z.-W. Ge, G. W. Griffith, K. Griffiths, J. Z. Groenewald, M. Groenewald, M. Grube, M.Gryzenhout, L.-D. Guo, F. Hagen, S. Hambleton, R. C. Hamelin, K. Hansen, P. Harrold, G. Heller, C.Herrera, K. Hirayama, Y. Hirooka, H.-M. Ho, K. Hoffmann, V. Hofstetter, F. Hognabba, P. M. Hollingsworth,S.-B. Hong, K. Hosaka, J. Houbraken, K. Hughes, S. Huhtinen, K. D. Hyde, T. James, E. M. Johnson, J. E.Johnson, P. R. Johnston, E. B. G. Jones, L. J. Kelly, P. M. Kirk, D. G. Knapp, U. Koljalg, G. M. Kovacs, C. P.Kurtzman, S. Landvik, S. D. Leavitt, A. S. Liggenstoffer, K. Liimatainen, L. Lombard, J. J. Luangsa-ard, H. T. Lumbsch, H. Maganti, S. S. N. Maharachchikumbura, M. P. Martin, T. W. May, A. R. McTaggart, A. S.Methven, W. Meyer, J.-M. Moncalvo, S. Mongkolsamrit, L. G. Nagy, R. H. Nilsson, T. Niskanen, I. Nyilasi, G.Okada, I. Okane, I. Olariaga, J. Otte, T. Papp, D. Park, T. Petkovits, R. Pino-Bodas, W. Quaedvlieg, H. A.Raja, D. Redecker, T. L. Rintoul, C. Ruibal, J. M. Sarmiento-Ramirez, I. Schmitt, A. Schussler, C. Shearer,K. Sotome, F. O. P. Stefani, S. Stenroos, B. Stielow, H. Stockinger, S. Suetrong, S.-O. Suh, G.-H. Sung, M.Suzuki, K. Tanaka, L. Tedersoo, M. T. Telleria, E. Tretter, W. A. Untereiner, H. Urbina, C. Vagvolgyi, A.Vialle, T. D. Vu, G. Walther, Q.-M. Wang, Y. Wang, B. S. Weir, M. Weiss, M. M. White, J. Xu, R. Yahr, Z. L.Yang, A. Yurkov, J.-C. Zamora, N. Zhang, W.-Y. Zhuang, D. Schindel, Fungal Barcoding Consortium,Nuclear ribosomal internal transcribed spacer (ITS) region as a universal DNA barcode marker for Fungi. Proc. Natl. Acad. Sci. U.S.A. 109, 6241–6246 (2012).
  24. E. Kalle, M. Kubista, C. Rensing, Multi-template polymerase chain reaction. Biomol. Detect. Quantif. 2, 11–29 (2014).
  25. R. Honegger, “The symbiotic phenotype of lichen-forming ascomycetes and their endo- and epibionts,” in Fungal Associations, B. Hock, Ed., vol. IX of The Mycota, K. Esser, Ed. (Springer, ed. 2, 2012), pp. 287–339.
  26. D. Anglesea, C. Veltkamp, G. H. Greenhalgh, The upper cortex of Parmelia saxatilis and other lichen thalli. Lichenologist 14, 29–38 (1982).
  27. E. S. Olafsdottir, K. Ingólfsdottir, Polysaccharides from lichens: Structural characteristics and biological activity. Planta Med. 67, 199–208 (2001).
  28. C. G. Boluda, V. J. Rico, A. Crespo, P. K. Divakar, D. L. Hawksworth, Molecular sequence data from populations of Bryoria fuscescens s. lat. in the mountains of central Spain indicates a mismatch between haplotypes and chemotypes. Lichenologist 47, 279–286 (2015).
  29. N. Arnold, W. Steglich, H. Besl, Zum vorkommen von pulvinsäure-derivaten in der gattung Scleroderma. Z. Mykol. 62, 69–73 (1996).
  30. W. L. Culberson, Proposed changes in the international code governing the nomenclature of lichens.Taxon 10, 161–165 (1961).
  31. M. G. Grabherr, B. J. Haas, M. Yassour, J. Z. Levin, D. A. Thompson, I. Amit, X. Adiconis, L. Fan, R.Raychowdhury, Q. Zeng, Z. Chen, E. Mauceli, N. Hacohen, A. Gnirke, N. Rhind, F. di Palma, B. W. Birren, C.Nusbaum, K. Lindblad-Toh, N. Friedman, A. Regev, Full-length transcriptome assembly from RNA-Seq data without a reference genome. Nat. Biotechnol. 29, 644–652 (2011).
  32. B. J. Haas, A. Papanicolaou, M. Yassour, M. Grabherr, P. D. Blood, J. Bowden, M. B. Couger, D. Eccles, B.Li, M. Lieber, M. D. Macmanes, M. Ott, J. Orvis, N. Pochet, F. Strozzi, N. Weeks, R. Westerman, T. William,C. N. Dewey, R. Henschel, R. D. Leduc, N. Friedman, A. Regev, De novo transcript sequence reconstruction from RNA-seq using the Trinity platform for reference generation and analysis. Nat. Protoc. 8, 1494–1512 (2013).
  33. C. Stubben, “genomes: Genome sequencing project metadata,” R package version 2.16.1 (2015);
  34. A. Dobin, C. A. Davis, F. Schlesinger, J. Drenkow, C. Zaleski, S. Jha, P. Batut, M. Chaisson, T. R. Gingeras,STAR: Ultrafast universal RNA-seq aligner. Bioinformatics 29, 15–21 (2013).
  35. M. D. Robinson, A. Oshlack, A scaling normalization method for differential expression analysis of RNA-seq data. Genome Biol. 11, R25 (2010).
  36. M. D. Robinson, D. J. McCarthy, G. K. Smyth, edgeR: A Bioconductor package for differential expression analysis of digital gene expression data. Bioinformatics 26, 139–140 (2010).
  37. A. McKenna, M. Hanna, E. Banks, A. Sivachenko, K. Cibulskis, A. Kernytsky, K. Garimella, D. Altshuler, S. Gabriel, M. Daly, M. A. DePristo, The Genome Analysis Toolkit: A MapReduce framework for analyzing next-generation DNA sequencing data. Genome Res. 20, 1297–1303 (2010).
  38. P. De Wit, M. H. Pespeni, J. T. Ladner, D. J. Barshis, F. Seneca, H. Jaris, N. O. Therkildsen, M. Morikawa, S. R. Palumbi, The simple fool’s guide to population genomics via RNA-Seq: An introduction to high-throughput sequencing data analysis. Mol. Ecol. Resour. 12, 1058–1067 (2012).
  39. L. Li, C. J. Stoeckert Jr., D. S. Roos, OrthoMCL: Identification of ortholog groups for eukaryotic genomes. Genome Res. 13, 2178–2189 (2003).
  40. S. van Dongen, “Graph clustering by flow simulation,” thesis, University of Utrecht (2000);
  41. K. Katoh, D. M. Standley, MAFFT multiple sequence alignment software version 7: Improvements in performance and usability. Mol. Biol. Evol. 30, 772–780 (2013).
  42. J. Castresana, Selection of conserved blocks from multiple alignments for their use in phylogenetic analysis. Mol. Biol. Evol. 17, 540–552 (2000).
  43. G. Talavera, J. Castresana, Improvement of phylogenies after removing divergent and ambiguously aligned blocks from protein sequence alignments. Syst. Biol. 56, 564–577 (2007).
  44. A. Stamatakis, RAxML version 8: A tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics 30, 1312–1313 (2014).
  45. D. H. Huson, C. Scornavacca, Dendroscope 3: An interactive tool for rooted phylogenetic trees and networks. Syst. Biol. 61, 1061–1067 (2012).
  46. N. Lartillot, N. Rodrigue, D. Stubbs, J. Richer, PhyloBayes MPI: Phylogenetic reconstruction with infinite mixtures of profiles in a parallel environment. Syst. Biol. 62, 611–615 (2013).
  47. N. Lartillot, H. Philippe, A Bayesian mixture model for across-site heterogeneities in the amino-acid replacement process. Mol. Biol. Evol. 21, 1095–1109 (2004).
  48. S. Tavaré, “Some probabilistic and statistical problems on the analysis of DNA sequences,” in Lectures in Mathematics in the Life Sciences (American Mathematical Society, vol. 17, 1986), pp. 57–86.
  49. J. L. Thorne, H. Kishino, I. S. Painter, Estimating the rate of evolution of the rate of molecular evolution. Mol. Biol. Evol. 15, 1647–1657 (1998).
  50. L. Salichos, A. Stamatakis, A. Rokas, Novel information theory-based measures for quantifying incongruence among phylogenetic trees. Mol. Biol. Evol. 31, 1261–1271 (2014).
  51. D. S. Hibbett, P. B. Matheny, The relative ages of ectomycorrhizal mushrooms and their plant hosts estimated using Bayesian relaxed molecular clock analyses. BMC Biol. 7, 13 (2009).
  52. T. N. Taylor, H. Hass, H. Kerp, M. Krings, R. T. Hanlin, Perithecial ascomycetes from the 400 million year old Rhynie chert: An example of ancestral polymorphism. Mycologia 97, 269–285 (2005).
  53. J. W. Taylor, M. L. Berbee, Dating divergences in the Fungal Tree of Life: Review and new analyses. Mycologia 98, 838–849 (2006).
  54. H. Dörfelt, A. R. Schmidt, A fossil Aspergillus from Baltic amber. Mycol. Res. 109, 956–960 (2005).
  55. D. Hibbett, D. Grimaldi, M. Donoghue, Fossil mushrooms from Miocene and Cretaceous ambers and the evolution of Homobasidiomycetes. Am. J. Bot. 84, 981 (1997).
  56. S. P. Stubblefield, T. N. Taylor, C. B. Beck, Studies of Paleozoic fungi. IV. Wood-decaying fungi in Callixylon newberryi from the Upper Devonian. Am. J. Bot. 72, 1765–1774 (1985).
  57. T. Feuerer, D. L. Hawksworth, Biodiversity of lichens, including a world-wide analysis of checklist data based on Takhtajan's floristic regions. Biodiversity Conserv. 16, 85–98 (2007).
  58. J. Miadlikowska, F. Kauff, F. Högnabba, J. C. Oliver, K. Molnár, E. Fraker, E. Gaya, J. Hafellner, V.Hofstetter, C. Gueidan, M. A. Otálora, B. Hodkinson, M. Kukwa, R. Lücking, C. Björk, H. J. Sipman, A. R.Burgaz, A. Thell, A. Passo, L. Myllys, T. Goward, S. Fernández-Brime, G. Hestmark, J. Lendemer, H. T.Lumbsch, M. Schmull, C. L. Schoch, E. Sérusiaux, D. R. Maddison, A. E. Arnold, F. Lutzoni, S. Stenroos, A multigene phylogenetic synthesis for the class Lecanoromycetes (Ascomycota): 1307 fungi representing 1139 infrageneric taxa, 317 genera and 66 families. Mol. Phylogenet. Evol. 79, 132–168 (2014).
  59. Y. Ohmura, K. Uno, K. Hosaka, T. Hosoya, “DNA fragmentation of herbarium specimens of lichens, and significance of epitypification for threatened species of Japan,” The 10th International Mycological Congress, Bangkok, Thailand, 4 to 8 August 2014 (2014), p. 151.
  60. R. Lanfear, B. Calcott, S. Y. W. Ho, S. Guindon, Partitionfinder: Combined selection of partitioning schemes and substitution models for phylogenetic analyses. Mol. Biol. Evol. 29, 1695–1701 (2012).
  61. S. Altermann, S. D. Leavitt, T. Goward, M. P. Nelsen, H. T. Lumbsch, How do you solve a problem like Letharia? A new look at cryptic species in lichen-forming fungi using Bayesian clustering and SNPs from multilocus sequence data. PLOS ONE 9, e97556 (2014).
  62. E. Paradis, pegas: An R package for population genetics with an integrated-modular approach. Bioinformatics 26, 419–420 (2010).
  63. Y. Yamamoto, R. Mizuguchi, Y. Yamada, Tissue cultures of Usnea rubescens and Ramalina yasudae and production of usnic acid in their cultures. Agric. Biol. Chem. 49, 3347–3348 (1985).
  64. M. del Carmen Molina, A. Crespo, Comparison of development of axenic cultures of five species of lichen-forming fungi. Mycol. Res. 104, 595–602 (2000).
  65. M. Gardes, T. D. Bruns, ITS primers with enhanced specificity for basidiomycetes—application to the identification of mycorrhizae and rusts. Mol. Ecol. 2, 113–118 (1993).
  66. T. J. White, T. Bruns, S. Lee, J. Taylor, “Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics,” in PCR Protocols: A Guide to Methods and Applications, M. A. Innis, D. H. Gelfand, J. J. Sinsky, T. J. White, Eds. (Academic Press, 1990), pp. 315–322.
  67. S. Behrens, C. Rühland, J. Inácio, H. Huber, A. Fonseca, I. Spencer-Martins, B. M. Fuchs, R. Amann, In situ accessibility of small-subunit rRNA of members of the domains Bacteria, Archaea, and Eucarya to Cy3-labeled oligonucleotide probes. Appl. Environ. Microbiol. 69, 1748–1758 (2003).
  68. J. Inácio, S. Behrens, B. M. Fuchs, A. Fonseca, I. Spencer-Martins, R. Amann, In situ accessibility of Saccharomyces cerevisiae 26S rRNA to Cy3-labeled oligonucleotide probes comprising the D1 and D2 domains. Appl. Environ. Microbiol. 69, 2899–2905 (2003).
  69. C. Baschien, W. Manz, T. R. Neu, L. Marvanová, U. Szewzyk, In situ detection of freshwater fungi in an alpine stream by new taxon-specific fluorescence in situ hybridization probes. Appl. Environ. Microbiol.74, 6427–6436 (2008).
  70. B. M. Fuchs, F. O. Glöckner, J. Wulf, R. Amann, Unlabeled helper oligonucleotides increase the in situ accessibility to 16S rRNA of fluorescently labeled oligonucleotide probes. Appl. Environ. Microbiol. 66, 3603–3607 (2000).
  71. D. Anglesea, G. N. Greenhalgh, C. Veltkamp, The cortex of branch tips in Usnea subfloridana. Trans. Br. Mycol. Soc. 81, 438–444 (1983).
  72. G. N. Greenhalgh, A. Whitfield, Thallus tip structure and matrix development in Bryoria fuscescens. Lichenologist 19, 295–305 (1987).
  73. R. Honegger, A. Haisch, Immunocytochemical location of the (1→3) (1→4)-β-glucan lichenin in the lichen-forming ascomycete Cetraria islandica (Icelandic moss). New Phytol. 150, 739–746 (2001).
  74. H. J. Elwood, G. J. Olsen, M. L. Sogin, The small-subunit ribosomal RNA gene sequences from the hypotrichous ciliates Oxytricha nova and Stylonychia pustulata. Mol. Biol. Evol. 2, 399–410 (1985).
  75. Acknowledgments: This project was supported by an incubation grant from the University of Montana to J.P.M. and T.S.; by an Austrian Science Fund grant (P25237) to T.S., H.M., and P.R.; by NSF (IOS-1256680, IOS-1553529, and EPSCoR award NSF-IIA-1443108) and NASA Astrobiology Institute (NNA15BB04A) grants to J.P.M.; by a NSF Graduate Research Fellowship (DGE-1313190) to D.V.; by a grant from the Swedish University of Agricultural Sciences Council for PhD Education (2014.3.2.5-5149) to V.T.; and by a grant (DO2011-0022) from Stiftelsen Oscar och Lili Lamms minne to G.T. Specimens from Glacier Bay National Park, AK, were collected with the support of the U.S. National Park Service as part of CESU (Cooperative Ecosystem Studies Units) project P11AC90513. We thank D. Armaleo and F. Lutzoni of Duke University for allowing us to use unpublished data from the Cladonia grayi proteome, as well as P. Dyer, P. Crittenden, and D. Archer (University of Nottingham, UK) for access to unpublished data from the Xanthoria parietina genome project, which is conducted together with the U.S. Department of Energy Joint Genome Institute (supported by the Office of Science of the U.S. Department of Energy under contract no. DE-AC02-05CH11231). We thank T. Goward, M. Grube, P. Lukasik, J. T. Van Leuven, F. Fernández-Mendoza, A. Millanes, V. Wagner, and M. Wedin for discussions and L. Bergström, C. Gueidan, J. Hermansson, H. Holien, B. Kanz, E. Lagostina, S. Leavitt, B. McCune, J. Nascimbene, C. Printzen, T. Wheeler, and D. Winston for field support and fresh material. C. Björk, S. Gunnarsson, L. Herritt, M. Hiltunen, W. Obermayer, A. de los Ríos, and E. Timdal provided technical help, advice, and photos. We acknowledge the Purdue University Genomic Core Facility for generating transcriptomic data for Cystobasidium minutum and the Institute of Molecular Biosciences–Graz Microscopy Core Facility and S. Kohlwein for providing infrastructural support for imaging. Data are available under accession numbers SRP076577 and SRP073687 in the National Center for Biotechnology Information (NCBI) Sequence Read Archive (transcriptomes), NCBI nucleotide accession numbers KU948728 to KU948928 (single-locus DNA sequences), and the Dryad digital repository at (alignments, scripts, and tree files).