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Fungal Communities Associated with Rock Varnishin Black Canyon, New Mexico: Casual Inhabitants orEssential Partners?Kylea J. Parchert a , Michael N. Spilde b , Andrea Porras-Alfaro a c , April M. Nyberg d & DianaE. Northup aa Biology Department , University of New Mexico , Albuquerque , New Mexico , USAb Institute of Meteroritics , University of New Mexico , Albuquerque , New Mexico , USAc Department of Biological Science , Western Illinois University , Macomb , Illinois , USAd National Clonal Germplasm Repository , Corvallis , Oregon , USAAccepted author version posted online: 06 Apr 2012.Published online: 12 Jul 2012.

To cite this article: Kylea J. Parchert , Michael N. Spilde , Andrea Porras-Alfaro , April M. Nyberg & Diana E. Northup (2012)Fungal Communities Associated with Rock Varnish in Black Canyon, New Mexico: Casual Inhabitants or Essential Partners?,Geomicrobiology Journal, 29:8, 752-766, DOI: 10.1080/01490451.2011.619636

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Geomicrobiology Journal, 29:752–766, 2012Copyright © Taylor & Francis Group, LLCISSN: 0149-0451 print / 1521-0529 onlineDOI: 10.1080/01490451.2011.619636

Fungal Communities Associated with Rock Varnishin Black Canyon, New Mexico: Casual Inhabitantsor Essential Partners?

Kylea J. Parchert,1 Michael N. Spilde,2 Andrea Porras-Alfaro,1,3 April M. Nyberg,4

and Diana E. Northup1

1Biology Department, University of New Mexico, Albuquerque, New Mexico, USA2Institute of Meteroritics, University of New Mexico, Albuquerque, New Mexico, USA3Department of Biological Science, Western Illinois University, Macomb, Illinois, USA4National Clonal Germplasm Repository, Corvallis, Oregon, USA

Rock varnish is a darkly pigmented coating rich in manganeseoxides. Though microbes inhabit varnish deposits, it is unclearwhether they are involved in varnish formation. The fungal com-munities of rock varnish and adjacent rock sites with no visible var-nish deposits were examined. Microcolonial fungi were identifiedat all sampling sites, and were associated with manganese oxides inpatches of incipient varnish at non-varnish sites. Fungi were closelyrelated to manganese-oxidizing genera and seventeen isolates ox-idized manganese in culture, producing six distinct manganese-oxide morphologies. Our results indicate that microcolonial fungimay play a crucial role in rock varnish formation. Supplementalmaterials are available for this article. Go to the publisher’s onlineedition of Geomicrobiology Journal to view the free supplementalfile.

Keywords microcolonial fungi, rock varnish, microorganisms, man-ganese oxidation

Received 17 June 2011; accepted 30 August 2011.The authors wish to thank Monica Moya, Matt Garcia,

Jessica Snider, and John Craig for their invaluable assistance in theperformance of laboratory activities as well as Dr. Penny Boston whoassisted with the selection of media recipes for fungal isolation and Jen-nifer Hathaway for help with sequence analysis. In addition, valuablecomments on the manuscript were supplied by anonymous reviewers.This work could not have been done without the generous support of theNational Science Foundation’s Geosciences Directorate (EAR0311932and EAR0311930) and Kenneth Ingham Consulting. We acknowledgetechnical support from the University of New Mexico Department ofBiology’s Molecular Biology Facility, supported by NIH Grant NumberP20RR18754 from the Institute Development Award (IdeA) Programof the National Center for Research Resources.

Address correspondence to Kylea J. Parchert, University of NewMexico, Biology Department, Albuquerque, NM 87131, USA. E-mail:kjodenb@sandia.gov

INTRODUCTIONSurface coatings, called rock varnish, are present on sub-

aerial rock surfaces in many climate regions and are sometimesreferred to as desert varnish in arid and semiarid environments.Rock varnish main components include clay, silica, and oxidesof iron and manganese (Potter and Rossman 1977; Krumbeinand Jens 1981; Liu and Broecker 2000; Gorbushina 2003a).The dark color of varnish may be a function of several materi-als: black from high concentrations of manganese oxides (Dorn2006) and dark brown from various concentrations of iron ox-ides or pigmented opaline silica (Perry et al. 2006). The thinlylaminated coatings rarely exceed 200 µm in thickness and buildup at rates of <1 to 40 µm/k.y. (Krumbein and Jens 1981; Liuand Broecker 2000). The rate of deposition is affected by en-vironmental factors, most notably rainfall (Liu and Broecker2000). For this reason, it has been suggested that varnishes canmap past climatic patterns, especially since it may take up to10,000 years for a “thick” varnish to develop (Nagy et al. 1991;Liu and Broecker 2000; Perry et al. 2006). However, the reliabil-ity of such records has been questioned (Reneau 1993; Kuhlmanet al. 2006).

Desert rock surfaces are inhospitable environments for manymicroorganisms because of the extreme diurnal and seasonaltemperature fluctuations, lack of easily accessible water andnutrients, and constant exposure to intense ultraviolet radi-ation (Gorbushina 2003a; Gorbushina and Broughton 2009).Despite these environmental difficulties, microorganisms havebeen consistently reported as inhabitants of varnish (Krumbeinand Jens 1981; Nagy et al. 1991; Kuhlman et al. 2006; Kuhlmanet al. 2008; Northup et al. 2010). Rock varnishes are colonizedmainly by highly specialized microorganisms that are adaptedto meet the demands of an extreme and variable environment(Gorbushina 2003a; Krumbein et al. 1981).

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The origin of rock varnish is not well understood. Re-searchers have speculated that rock varnish genesis may be aresult of biological processes in which bacteria and fungi facil-itate the accumulation and transformation of minerals (Taylor-George et al. 1983; Golden et al. 1992; Burford et al. 2003; Gadd2007, Wang et al. 2011). Naturally occurring manganese oxidesin the environment are often a result of biological metabolism(Tebo et al. 1997), and manganese-oxidizing bacteria and fungiare known inhabitants of varnishes (Nagy et al. 1991; Gadd2007). Conversely, however, the varnish may be the result ofsolely abiotic processes. In this scenario, rock varnish-inhabitingorganisms could be just chance residents, deposited from nearbysoil by wind dispersal (Schelble et al. 2005; Perry et al. 2006).Varnishes exist in almost all geographic and climatic regions,which could indicate that varnish formation is the result of sedi-mentation processes (Liu and Broecker 2000). Perry et al. (2006)suggested that the formation of a silica glaze might be a crucialstep in an abiotic model for varnish formation.

Compared to bacteria, the study of fungi has been neglected,yet there are many reasons to suspect that the involvement offungi is crucial for rock varnish formation (Gorbushina 2003a;Kuhlman et al. 2006, Gadd and Raven 2010). For example, manygenera of fungi are known to produce manganese-oxide depositsin soil and aquatic environments including Acremonium, Cremo-nium, Coniothyrium, Cladosporium, Penicillium, Phoma, andVerticillium (Tebo et al. 2004; Thompson et al. 2005; Miyataet al. 2006, Gadd and Raven 2010). The mechanics of fungal-mediated manganese oxidation and deposition are not yet fullyunderstood (Miyata et al. 2006; Tebo et al. 2007), although ithas been suggested that manganese in varnish-inhabiting fungicould facilitate resistance to high solar radiation, as is seen inthe cyanobacteria (Krumbein and Jens 1981; Gorbushina 2003a;Daly et al. 2004; Tebo et al. 2007). Additionally, recent stud-ies of bacterial communities associated with varnish coatingsin New Mexico, using scanning electron microscopy (SEM),revealed the presence of putative microcolonial fungi (MCF)in incipient varnish (Northup et al. 2010). MCF are recognizedto be a community of highly specialized organisms that growin compact colonies and are commonly associated with rockvarnish (Gorbushina 2003a).

The purpose of this study was to investigate the role of fungiin rock varnish formation using the production and deposition

of manganese oxides as a model. Using 18S rDNA sequenc-ing we identified common fungi that inhabit rocks with visiblevarnish coatings and compared those with adjacent rocks withno macroscopically visible varnish. To ascertain whether fungipresent in varnish can precipitate manganese oxides, low-carbonfungal media supplemented with reduced manganese was inoc-ulated with chips of varnish-coated rock. To the best of ourknowledge, this is the first study to combine molecular, SEM,and microbiological techniques to reveal the potential role offungal manganese-oxidizers in the formation of rock varnish.

MATERIALS AND METHODS

Sample CollectionSamples from black varnish coatings on andesitic volcanic

rock were aseptically collected in Black Canyon, New Mexicoin November 2003, December 2005, and June 2006. The sam-ple collection site is located within a 33 Ma Socorro volcaniccauldron complex, 9 km southwest of Socorro in central NewMexico, USA (Eggleston et al. 1983). The area is semiarid Chi-huahuan desert, which on average receives less than 250 mm ofprecipitation annually. Samples were collected from five siteson several rock faces along approximately 100 m of road. Sites1, 2, and 5 had abundant, visible deposits of rock varnish (seeTable 1 for description and location of each sampling site, seeSupplementary Materials Figure S1 for sampling site images).

Samples from sites 1 and 2 were taken from a dry, south-facing rock face with scattered patches of black rock varnish.Site 5 is an ephemeral watercourse on a vertical rock face that,during rain events, drains several hundred square meters of rockand scattered soil above it. Sites 3 and 4 had no macroscopicallyvisible varnish coating. Site 3 was an exposed and naturallyweathered bare rock surface, whereas site 4 was a dynamite-blasted road cut, constructed sometime during or slightly afterWorld War II when a mine nearby was opened. At least a dozenrock pieces were sampled from each site, and these varied insize from a few millimeters to several square cm. Several rockpieces were taken from each site because many smaller sampleshave been shown to better capture microbial diversity in soilsthan a single large sample (Grundmann and Gourbieri 1999;Ranjard et al. 2003).

TABLE 1Description and location of varnish and non-varnish sampling site

Site GPS Coordinates Site Details

Site 1 33◦ 58′ 25.94′′ N 106◦ 59′ 36.24′′ W Varnish siteSite 2 33◦ 58′ 26.03′′ N 106◦ 59′ 36.27′′ W Varnish site - dynamite blasted in 1940sSite 3 33◦ 58′ 25.97′′ N 106◦ 59′ 36.12′′ W Non-varnish siteSite 4a 33◦ 58′ 26.91′′ N 106◦ 59′ 36.88′′ W Non-varnish siteSite 5 33◦ 58′ 27.36′′ N 106◦ 59′ 37.06′′ W Varnish site - ephemeral water course

a – Site coordinates were estimated from Site 3 and Site 5 coordinates.

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Varnish from the Black Canyon site is often thin, discon-tinuous and concentrated into micro-pits and pockets in therock surface, unlike the thicker varnish in the Mojave Desert inCalifornia (c.f. Broecker and Liu 2001) and other locations.Black Canyon is part of a mining district where hydrothermalmanganese deposits were mined during and after World WarII. Thus the rock that encompasses the canyon also containselevated manganese levels that may provide a more substantialsource of manganese than most other rock varnish sites, althoughrock varnish composition seems to be independent of the under-lying rock composition (Potter and Rossman 1977, 1979; Dornand Oberlander 1981). Weakly acidic rainwater may leach man-ganese from surrounding rocks, providing a source of Mn(II).Aseptically collected rock pieces from each site were stored insucrose lysis buffer (40 mM EDTA, 400 mM NaCl, 0.75 Msucrose, 50 mM Tris hydrochloride [pH 9.0]; Giovannoni et al.1990), and transported at ambient temperature before storage at−80◦C. A second group of rock samples collected from varnishsites (sites 1, 2, and 5) were embedded into prepared selectivegrowth media on site.

CulturingRock pieces from varnish sites, ranging in size from 1

2to 2 cm2, from the June 2006 sampling date were embed-ded into four different media (all media contained 100 mg/mlampicillin): 1. Mn(II)-one-half PDA (reduced carbon, Mn(II)-enriched potato dextrose agar, Hose et al. 2000), 2. Mn(II)-lowPDA (Mn-enriched media with carbon availability further re-duced, modified from Hose et al. 2000), 3. Mn(II) BHE (Mn(II)-enriched boiled hay extract (BHE) medium, Hose et al. 2000),and 4. Mn (II) trace mineral (a trace mineral media enriched withMn(II), Miyata et al. 2006). The half-PDA medium contained1 L deionized water, 13 g PDA (EMD Biosciences Inc., SanDiego, CA), 10 g BactoAgar (BD Diagnostics, Franklin Lakes,NJ) and 1 g MnCl2. The low-PDA medium is a modified versionof half-PDA medium: we used 6.5 g of PDA instead of 13 g and16 g BactoAgar intead of 10 g.BHE medium was made by boil-ing 100 g of hay in 3 L of water for 3 h. The liquid was then fil-tered using a mesh screen, followed by a 0.22 µm Steriflip filter(Millipore, Billerica, CA). One liter of this hay liquid was com-bined with 2 g nutrient broth (BD Diagnostics, Franklin Lakes,NJ) and 10 g BactoAgar. The trace mineral salt media (Miyataet al. 2004) contained 3 mmol sodium acetate, 150 mg of yeastextract, 50 mg of MgSO4·7H2O, 5 mg of K2HPO4, and 2 mlof a stock solution containing (per liter): 3.7 g of CaCl2·2H2O,2.5 g of H3BO3, 0.87 g of MnCl2·4H2O, 1.0 g of FeCl3·6H2O,0.44 g of ZnSO4·7H2O, 0.29 g of Na2MoO4·2H2O, and 5 mg ofCuSO4·5H2O, per liter of 20 mM HEPES buffer.

Each media type, except for the trace mineral salt media,was enriched with 1 g MnCl2/L before autoclaving. Per site,four plates were embedded with three rock pieces per plate andwere incubated at 25◦C or 37◦C for seven days. During each dayof incubation, the plates were examined and fungal colonieswere sub-cultured until pure cultures were obtained. Cultures

were examined using light microscopy to verify that only oneorganism was present. Though culture media always containedampicillin, cultures were routinely examined for bacterial con-tamination. Fungal isolates were incubated for several weeks atthe respective temperatures before being examined for signs ofvisible manganese oxide production, such as darkly pigmentedhyphae or dark deposits within or beneath the mycelium.

MicroscopyDarkly pigmented cultures were photographed using digital

cameras attached to microscopes (Zeiss Stereoscope DiscoveryV12 model AxioCam MRm [black and white] and AxioCamHRc [color]). Samples were also examined on a JEOL 5800LVscanning electron microscope (SEM) equipped with an OxfordIsis 300 energy dispersive X-ray analyzer (EDX) employingboth secondary electron imaging (SEI) and backscattered elec-tron (BSE) image, which is sensitive to changes in atomic num-ber of the sample. An initial screening of dark pigmented fungalspecimens were dried on glass microscope slides in preparationfor SEM examination. Samples that were found to contain man-ganese using the EDX, were then fixed with 4% glutaraldehydesolution and critical point dried after a series of ethanol and ace-tone dehydration steps (Glauert 1984). All samples were sputtercoated with Au-Pd prior to examination in the SEM.

DNA ExtractionDNA was extracted from rock chip samples at all five sites

using the Powersoil DNA Extraction Kit (MoBio Laboratories,Carlsbad, CA) according to the manufacturer’s protocols, exceptrock pieces (1–3 pieces totaling approximately 1

4 to 12 cm2) were

shaken in a bead beater instead of on a vortexer, and DNA waseluted into 30 µl of EB buffer instead of 50 µl in order toincrease the DNA concentration. For each extraction performed,a negative control without sample was prepared.

DNA was also extracted from pure cultures that precipi-tated manganese oxides (verified by SEM and EDX) usingthe UltraClean Microbial DNA Isolation Kit (Mobio Labora-tories, Carlsbad,CA). Instead of vortexing horizontally as spec-ified in the protocol, tubes containing fungal hyphae (harvestedby aseptically slicing away the culturing media beneath themycelium) were shaken in a Mini-Bead Beater (Biospec Prod-ucts, Bartlesville, OK) for 3 min at a maximum of 3000 rpm.Extracted DNA was stored at −20◦C. A negative control wasincluded for each extraction procedure.

DNA AmplificationExtracted DNA from rock chips was amplified using a

polymerase chain reaction (PCR), and was assayed in trip-licate to minimize the impact of PCR bias. Extracted DNA(1 µl) was combined with 6.5 µl of molecular grade deion-ized water, 0.025 mg bovine serum albumin (BSA) (RocheApplied Science, Indianapolis, IN), 12.5 µl of Takara TaqPremix (Takara Bio Inc., Temecula, CA), and 1 µl eachof 50 µM fungal specific forward and reverse primers

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for the 18S SSU region. The forward primer used wasNS3 (5′GCAAGTCTGGTGCCAGCAGCC3′) and the reverseprimer was NS8 (5′TCCGCAGGTTCACCTACGGA3′) (Whiteet al. 1990). Negative and positive controls were prepared foreach PCR conducted. Touchdown PCR conditions were as fol-lows: 95◦C for 4 min, 95◦C for 30 s, 65◦C for 30 s (increase1◦C each cycle), 72◦C for 45 s, repeat 9 times to step 2, 95◦Cfor 30 s, 58◦C for 30 s, 72◦C for 45 s, repeat 20 times to step6, 72◦C for 10 min, hold at 4◦C (Korbie and Mattick, 2008).Four µl of PCR amplicons were run out on a 1% SeaKem LEAgarose gel (Cambrex Bio Science Rockland Inc., Rockland,ME) and stained with ethidium bromide. The gel was imagedusing the Ingenius System (Syngene, Frederick, MD).

PCR CleanupPCR products from rock extracts were cleaned up with the

MiniElute PCR Purification Kit (Qiagen, Valencia, CA) follow-ing manufacturer’s protocols. If more than one band was visible,three replicate PCRs were combined, run out on a gel preparedwith QiaQuick 1XTAE, and the appropriately sized band wasgel extracted using the QiaQuick Gel Extraction Kit (Qiagen,Valencia, CA). The gel extracted PCR product was quantifiedusing a ND-1000 Spectrophotometer (Nanodrop Technologies,Wilmington, DE), cloned and sequenced (see details below).PCR amplicons from fungal isolates were cleaned using 2 µl ofExosap-IT (USB, Cleveland, OH) and 5 µL of the PCR product.This mixture was heated in a thermocycler to 37◦C for 15 min,then to 80◦C for 15 min and then cooled and held at 4◦C.

Molecular CloningCleaned PCR amplicons were ligated into the pCR4 vector

from the TOPO TA Cloning Kit (Invitrogen, Carlsbad, CA) fol-lowing the manufacturer’s protocols with the exception of theextension of ligation time to one hour. Ligations were trans-formed using Top10 competent cells, according to the protocolsuggested by the kit. The final transformed product was platedonto Luria-Bertani (LB) plates supplemented with 50 ng/mlampicillin and plates were incubated at 37◦C overnight.

Whole Cell PCR MethodColonies from the transformation reaction were sub-cultured

onto LB media and allowed to incubate at 37◦C for severalhours. Colonies were checked for inserts of appropriate sizeusing a whole cell PCR (WCPCR) assay method. Cells werecombined with 12.3 µL of PCR Supermix (Invitrogen, Carls-bad, CA), 0.1 µl of 50 µM T3 vector primer (ATTAACCCT-CACTAAAGGGA), and 0.1 µl of 50 µM T7 vector primer(TAATACGACTCACTATAGGG). The thermocycler protocolused was as follows: 95◦C for 10 min, 55◦C for 1 min, 72◦Cfor 1 min, 94◦C for 1 min, repeat 30 times to step 2, 50◦C for45 s, 72◦C for 10 min, hold at 4◦C. Products were examinedusing the gel electrophoresis method specified above. Coloniesthat did not contain the correctly sized insert of approximately1500 bp were discarded.

Restriction Fragment Length PolymorphismsTo assist in determining which clones to sequence, the 18S

ribosomal DNA was cut with enzymes to produce RFLPs (re-striction fragment length polymorphism) patterns: 1 µl of plas-mid DNA, 2 µl of Reaction Buffer 3, 16 µl of double distilledwater, and one µl of enzyme were used to digest the DNA.Enzymes used were HhaI and RsaI, with both enzymes in eachreaction. Sheared DNA patterns were visualized using a 4%Metaphor (FMC Rockland, Maine) electrophoresis gel in TAE,stained with 1 µl of ethidium bromide and visualized on a UVtransilluminator.

DNA SequencingDNA was prepared from clones using the Qiaprep Spin

Miniprep Kit (Qiagen Inc., Valencia, CA) and an appropriateamount of DNA was added to the sequencing reaction based onreadings using a ND-1000 Spectrophotometer (Nanodrop Tech-nologies, Wilmington, DE). Sequencing reactions contained1 µl Big Dye (Applied Biosystems, Foster City, CA), 1 µl ofthe appropriate 5 µM forward or reverse primer (NS3, NS4[5′CTTCCGTCAATTCCTTTAAG3′], or NS8) (White et al.1990), 3 µl of the 2.5x Big Dye Buffer (Applied Biosystems,Foster City, CA), 100–200 ng of DNA, and enough water tobring the volume up to a total of 10 µl. The following thermo-cycler program was used: 96◦C for 10 s, 45◦C for 15 s, 60◦Cfor 4 min, repeat 24 times, hold at 4◦C. DNA from sequenc-ing reactions was precipitated with anhydrous ethanol and 3 Msodium acetate, washed with 70% ethanol twice, and dried untilall ethanol evaporated. Sequencing reactions were carried out onthe ABI Prism 3100 DNA Sequencer (Perkin Elmer, Waltham,MA).

PCR products (1 to 3 µl) from fungal cultures were alsosequenced following the protocol described above. PCR con-centration was quantified in a gel based on the brightness ofthe band. Sequences of cultures and environmental sampleswere deposited in GenBank under accession numbers JF19074-JF19122.

Phylogenetic AnalysisDNA sequences from clones and cultures were cleaned and

assembled into contigs using Sequencher 4.6 (Gene CodesCorporation, Ann Arbor, MI). Results from a nucleotideBLAST (NCBI; Altschul et al., 1997) search were used tochoose similar sequences and outgroups (James et al. 2006).All sequences were aligned using CLUSTAL W (Thompsonet al. 1994) and the alignment was refined using BioEdit(Tom Hall, www.mbio.ncsu.edu/BioEdit/BioEdit.html). PAUP4.0b10 (Swofford 2000) was used for parsimony analysis thatwas generated using a heuristic search and bootstrap analysis(1000 replicates).

RESULTS AND DISCUSSIONCircumstantial evidence, such as the ubiquitous presence of

rock varnish throughout the world, or the consistent presence

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of microbes in varnish has not offered a satisfactory answer toexplain the origin of rock varnish. Within the last few decades,the advent of culture-independent molecular biology has openedup new opportunities for research that may provide more solidevidence to answer the question “What came first, the microbesor the varnish?”

Though both fungi and bacteria are consistently found inthe complex communities that inhabit rock varnish, few studieshave focused on the fungal inhabitants. Studies that have focusedon the fungal rock varnish community have either exclusivelyemployed molecular techniques (Kuhlman et al. 2006; Schel-ble et al. 2005) or culturing combined with scanning electronmicroscopy (SEM) (Taylor-George et al. 1983; Krumbein andJens 1981). Our study is the first, to our knowledge, to combineall three techniques.

Scanning Electron Microscopy of Black Canyon SitesMicrocolonial fungi (MCF) were present in samples from

all of the sites including sites 3 and 4, which had no macro-scopically visible varnish deposits (Figure 1A–D). MCF wereobserved inhabiting pits in the rock surface, and in some caseswere surrounded by iron and manganese oxides (Figure 1B–D).Hyphae could be observed extending out from some colonies tothe surrounding rock (Figure 1B, E–G). Other studies have alsoobserved MCF and lichens in varnish rock pits (Krumbein andJens 1981; Staley et al. 1982), with hyphae extending out fromthe colonies to the exposed rock surface (Taylor-George et al.1983).

The compact fungal colonies were visible in pits on the rocksurface of the non-varnish rocks, although colonies were morenumerous in the rock varnish samples. The rock face at site 4was dynamite blasted around the time of World War II duringroadway construction to the Black Canyon Mine, providing atime constraint of approximately 60 years during which MCFcould develop. MCF colonies at site 4 were very small, sug-gesting that increase in colony size may occur slowly in theseextreme environments (Figure 1A).

Site 3, the other non-varnish sampling site, possessed largerand more numerous colonies of MCF than those of site 4 (com-pare Figures 1A and 1B). The observation of MCF at the non-varnish sites, the presence of known managanese oxidizers inand the interspersing of rock varnish and non-varnish fungi inthe phylogenetic analysis (see below) suggest that there is in-cipient varnish present at sites that we classified as non-varnishsampling sites. Northup et al. (2010) suggested that varnishis present, but not visible macroscopically at these rock sitesbecause it is in the early stages of formation. Krumbein andJens (1981) suggested that the rock varnish formation beginswith patches of varnish that eventually coalesce. For microor-ganisms that inhabit rock surfaces, dark manganese-oxide coat-ings offer a powerful protection against dangerous UV exposure(Krumbein and Jens 1981; Gorbushina 2003a; Daly et al. 2004;Tebo et al. 2007). MCF at non-varnish sampling sites were ob-served inhabiting incipient varnish patches and were associated

with oxidized metal deposits characteristic of rock varnish, yet,surprisingly, MCF at more developed varnish sites were not as-sociated with metal deposits. This suggests that the MCF actas pioneer organisms in the harsh environment of the exposedrock surfaces (Figure 1C). This alteration of the environmentcould then allow other organisms (e.g. bacteria, including Mn-oxidizers, and [in more humid deserts] algae) that normallywould not be able to survive the high levels of UV exposure, toinhabit the same surface.

Northup et al. (2010) investigated bacterial communities ofthe Black Canyon rock varnish, in which MCF were observedin the varnish near, but not directly associated with, Mn deposits(see Figure 8 in Northup et al. 2010). Northup et al. suggestedthat changes in metal oxidation capacity may happen as a resultof colonial maturation and production of stable biofilms, result-ing in energy being utilized for colonial growth instead of metaloxidation, which is no longer required for survival (Figure 1B,E–G).

It is unclear from SEM images of incipient varnish, how MCFare related to the rock pits that they inhabit or the surroundingdeposits of Mn- and Fe-oxides, particularly since no Mn- or Fe-oxides were detected on the surface of the MCF colonies. Thereare several potential explanations for the presence of MCF inthe midst of the pits and oxidized metal deposits on the rock sur-faces: It is possible that (i) the fungi themselves are eroding therock surface and are enzymatically responsible for the oxidizedmetals surrounding the colonies, (ii) other microorganisms arepresent or were at one time present beneath the MCF and thatthese organisms are responsible for establishing the pits and de-posits that the MCF inhabit, or (iii) that abiotic processes areresponsible for rock erosion and metal oxidation and the MCFhave become established in pre-formed pits or crevices. In or-der to better address this uncertainty, future work is aimed atslicing through the fungal colonies and using SEM to examinethe hidden interface between the MCF and rock surface.

Isolation and Microscopy of Pure Fungal IsolatesRock varnish rock pieces collected in June 2006 were em-

bedded into four Mn-enriched media (Mn(II)-half PDA, Mn(II)-low PDA, Mn(II) BHE, and Mn (II)-trace mineral salt), result-ing in the growth of 119 pure fungal isolates. Multiple mediatypes were utilized for fungal isolation in order to maximizethe number of culturable fungal isolates. Little is understoodabout the growth requirements of rock varnish-inhabiting fungi.Sasaki et al. (2004) found that hot spring fungal isolates were in-creasingly able to oxidize Mn as carbon availability decreased.Therefore, multiple carbon sources and amounts of carbon werechosen (e.g., boiled hay extract [BHE], potato dextrose agars[PDA], and trace amounts in the Miyata et al. 2006 media).

Initial screening of fungal isolates for manganese oxideswas based on the presence of darkly pigmented hyphae and/ordeposits within the mycelial mat or in the media beneath thefungal tissue (Figure 2). Based on these criteria, a total of 26out of 119 fungal isolates were chosen for additional analysis

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FIG. 1. Scanning electron micrographs of microcolonial fungi (MCF) on rock fragments. (A) A few MCF are visible in micro-pits in the site 4 sample, asindicated by the white arrow. Scale bar is 50 µm. (B) More abundant and larger MCF were visible at site 3. A typical colony is indicated with the white arrow;scale bar is 200 µm. (C) A BSE image of the site 3 MCF, showing bright deposits (higher atomic number) surrounding the colonies (scale bar is 500 µm). A blackarrow indicates where EDX spectra was taken, (D) EDX spectrum indicates that the bright area is rich in Mn, Fe, O, Si, and Al. (E) MCF in micro-pits in varnishsite 1; scale bar is 100 µm. White arrows indicate MCF; black arrows indicate hyphae emerging from MCF. (F) Close-up of MCF from site 2; scale bar is 20 µm.(G) Numerous MCF inhabiting micro-pits in the varnish rock surface from site 5; scale bar is 200 µm.

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FIG. 2. Macroscopic and photomicrographs of example pure cultures showing putative manganese oxides. (A) Isolate M36 had small dark spots visible at thepoint of the white arrow and in the region between the black circles. (B) Isolates L16 possessed darkly pigmented (brownish red to black) hyphae. (C) IsolateK11shows large, black deposits of putative manganese oxides. (D) Isolate K5 had discrete dark deposits in the medium beneath the mycelium (4X magnification).(E) Isolate K44 had compact colonies with small clumps of dark deposits visible on hyphae extending from the colonies (2X magnification).

using scanning electron microscopy (SEM). Through SEM andEDX analysis, 17 cultures were found to contain manganeseproducts. The majority of isolates with putative manganese ox-ides were isolated on different specific media (see methods), butall were maintained on PDA media (see Supplementary Materi-als Table S1). The greatest number of fungal morphotypes wasobserved among the isolates on the BHE media, though fewercultures were positive for Mn oxidation (see SupplementaryTable Materials 1). Isolates on the trace-mineral media devel-oped by Miyata et al. (2006) grew more slowly than the isolateson the other media. Isolates on this media were restricted in size,and were generally visible as hyaline mycelia that accumulateddark deposits with age (Figure 2A). M31 and M36 were the onlyisolates on this media in which Mn deposits were detected (seeSupplementary Materials Table 1).

From the EDX analysis, we determined that the Mn by-products associated with fungal growth in cultures were in theform of manganese oxide (MnOx) and manganese phosphate.The deposits were classified into six groups based on theirmorphology. These groups were (I) crumbly masses, (II) traceamounts of manganese oxide, (III) Mn-coated hyphae, (IV) crys-talline Mn-bracelets, (V) Mn-coated nodules and rods, and (VI)radial Mn-structures (Figure 3–5, Table 2). For the first group,which contained cultures K5, K22, K43, K52, and M36, theMn-rich structures were directly associated with hyphae (Fig-ure 3A) and MnOx was detected, as determined by EDX (datanot shown). Isolates K6 and K17 were classified as Group II be-cause only trace amounts of Mn were detected (Figure 3B–C).For K6 (Figure 3B), Mn was present in a structure similar to thecrumbly masses of Group I, while for isolate K17 the structures

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FIG. 3. Electron micrographs Group I, II, and III pure cultures. (A) Group I manganese structures are described as crumbly masses; e.g. BSE image of isolateK5. Scale bar is 1 mm. Black arrow indicates Mn-rich area. (B) Photomicrograph of a Group II culture, isolate K6, showing trace amounts of manganese; scalebar is 10 µm. Arrow indicated Mn-rich structure. (C) Isolate K17 illustrates the coated rods of Group II. Scale bar 20 is µm. (D). The Mn-coated hyphae of isolateK21 of Group III, indicated by the black arrow, were rich in Mn and O; scale bar is 500 µm. EDX spectrum (not shown) of the blob, indicated by the white arrowin, showed that it was rich in Mn and Cl. (E) BSE image of isolate K11, which contained short and rigid Mn-coated hyphae (Group III); scale bar is 200 µm. Arrowindicates Mn-rich structure. (F). BSE-image of Mn-coated hyphae from isolate K31, longer and more flexuous than those found in K11; scale bar is 100 µm.

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FIG. 4. SEI, BSE images and EDX spectra of Group IV and Group VI pure cultures. White arrows in BSE images indicated where EDX spectra were collected.(A, B, C) The Mn-bracelets of Group IV were found exclusively in isolate K11; scale bar is 10 µm. (D, E, F) Group VI radial Mn- structures were found exclusivelyin isolate L27; scale bar is 50 µm).

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FIG. 5. Electron micrographs of example Group V pure cultures. (A) Mn-coated nodules are seen in the SEI of isolate K44; scale bar is 20 µm. Arrow indicatesMn-rich area. (B) Another type of Mn-coated nodule was seen in a BSE of isolate L16; scale bar is 20 µm. Arrow indicates Mn-rich area. (C) Mn-coated rodswere visible in a BSE of isolate M31; scale bar is 50 µm. Arrow indicates Mn-rich area. (D, E) An additional SEI and BSE from isolate L16 show the presence ofcoated nodules where the Mn appears to be internally present in the nodule; scale bar is 100 µm.

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TABLE 2Seventeen fungal isolates out of a total of 119 isolates

produced Mn-oxides observed in electron micrographs and areseparated into six groups based on the type of Mn-structure

Group Mn Structure Isolate

I Crumbly masses K5, K22, K43, K52,M36

II Trace amounts K6, K17III Mn-coated hyphae K11, K18, K21, K31,

K43, L18, L29IV Crystalline bracelets K11V Mn-coated nodules or rods K44, L16, L27, M31VI Radial Mn-structure L27

resembled the coated rods in Group V (Figure 3C). Mn-coatedhyphae (Group III) were found in the largest number of isolates(Figure 3D–F). The Mn-coated hyphae of isolate K21 were at-tached to a Mn blob (Figure 3D). The Mn-coated hyphae fromisolate K11 were rigid (Figure 3E), while among the remainingisolates, the Mn-coated hyphae appeared to be flexuous (Figure3F). In addition to the rigid Mn-coated hyphae, isolate K11 pos-sessed Mn-structures that were not observed in any other isolate.This structure (Group IV) was a crystalline bracelet of MnOxthat completely encircled the hypha (Figure 4A–C). The iso-lates in Group V (K44, L16, and M31) possessed tissue that wascoated by Mn (Figure 5). Isolates K44 and L16 contained coatedstructures that were nodule-like (Figure 5A, 5B), while isolateM31 contained structures that were rod-like (Figure 5C). Addi-tional images of isolate L16 suggest that some of the Mn may beinternal within nodules in the media (Figure 5D–EG–H). L27was the only isolate that was observed with radial Mn-structures(Group VI) (Figure 4D–E). This was a petal-like MnOx struc-ture that appeared to be formed of concentric rings.

Krumbein and Jens (1981) observed a manganese structuresimilar to the Group IV manganese bracelets reported here forisolate K11. The remaining Mn-structure types identified inthis study have not been observed in other studies where SEMwas used to examine rock varnish. Other researchers (Perryand Adams 1978; Nagy et al. 1991) have identified bacterialstructures in desert varnish that also have rarely been seen byother researchers. Rock varnish is a harsh environment withlimited resources and MCF are known to be efficient saprobeswith potential capacity to recycle scarce nutrients within thecompact colonies. The scarcity of resources and consequentrecycling of available nutrients could explain why these fungaland bacterial structures have not been observed or have onlyrarely been observed in situ (Gorbushina 2003a; Gorbushina2003b).

Of the fungi that were isolated from rock varnish in thisstudy, less than a quarter oxidized manganese. It is possible thatsome of the fungal isolates did not oxidize manganese due to

the conditions or media tested. Many of the 18S sequences fromthe cultured isolates and clones were closely related to fungifrom genera that are known to oxidize manganese (e.g., Phoma,Penicillium, and Alternaria) (Tebo et al. 2004; Thompson et al.2005; Miyata et al. 2006). Additional isolation strategies shouldbe explored in future work in order to culture fungi that wereunculturable on media used in this study (see Wollenzien et al.1995 and Rubial et al. 2005). Rock varnish communities arecomposed of a diverse and complex mixture of microorganismsand many of these microbes may contributed to varnish forma-tion, yet our data suggests that fungi may play an important rolein rock varnish formation that has heretofore been overlookedand necessitates further exploration.

Fungal Community AnalysisThe restriction fragment length polymorphism (RFLP) anal-

ysis identified distinct banding patterns in the fungal clonesfrom rock varnish and non-varnish sampling sites. For the rockvarnish sites 1, 2, and 5, there were a total of 10 distinct bandingpatterns and 91 clones, and for the non-varnish sites 3 and 4there were 9 distinct patterns and 57 clones. Not all clones weresequenced because the main objective was to identify the mostcommon fungi present in varnish and an exhaustive analysis ofthe diversity of fungi in varnish was beyond the scope of thisproject. The 18S region was fully sequenced for a total of 15clones from the rock varnish samples (Black Canyon sites 1, 2,and 5). An additional 23 clones were fully sequenced from thenon-varnish sites (Black Canyon sites 3 and 4).

Phylogenic AnalysisThe 18S sequences for the 38 clones from non-varnish and

rock varnish sites were included in a phylogenetic analysis, aswell as 18S sequences from eleven cultured fungal isolates.DNA extraction and/or sequencing was unsuccessful for iso-lates K18, K22, K31, L16, L29, and M31, though multiplesequencing attempts were made, so these isolates were not in-cluded in the analysis. Representative fungal sequences werechosen from near relatives based on BLAST and other knownMn oxidizers such as: Phoma, Cladosporium, Alternaria, andPenicillium (Gadd 2007; Miyata et al. 2006). In addition, fungalrock varnish sequences from a study conducted in California(Kuhlman et al. 2006) were included to assess the relatednessof other fungal varnish communities.

Except for two Basidiomycota clones (NVLL3A1 andRV2bfE3), all of the clones and isolates were Ascomycota(Figure 6). The closest related sequence to the Basidiomycotarock varnish clone (RV2bfE3) was an uncultured Basidiomy-cota clone from a deep-sea hydrothermal vent (EF638629) andthe closest related sequence from a cultured fungus was Filoba-sidium elegans (AB075545).

Isolates K5, K6, K17, K52, L27, and M36 grouped closelywith Penicillium chrysogenum (AF548087), which is known tobe an inhabitant of soil in arid environments. Isolate K11 andclone NVLL3A6 grouped with Cladosporium sp. (EU167592).

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RV12LL2A2RV12LL1A12RV12LL1A11RV12LL1A2

AY635836 Lecophagus sp. NVLL3D10

AY923099 Whipple RV clone

RV12LL1A1DQ066714 Cryomyces minteri

L76614 Cenococcum geophilumRV12LL5A7

NVLL3D7RV12LL2A7RV12LL2A11

AF250818 Phaeosphaeriopsis nolinaeNVLL3B5

AF250819 Phaeosphaeriopsis glauco-punctataK21

K44DQ678009 Cucurbitaria elongata

NVLL3C8NVLL3A7

AY923091 Whipple RV cloneNVLL3A5

EU167560 Ascochyta viciae-villosae AY337712 Phoma herbarum *K43NVLL3A8

AY923098 Whipple RV cloneL18NVLL3C9

NVLL3C2

RV2bfB11NV3bfC5RV2bfF11NV3bfB8NV3bfD7NV3bfC12

NVLL3D2RV12LL2A4

AJ972809 Sarcinomyces sp. EU089664 Guignardia mangiferae

EU167562 Bagnisiella examinansNVLL3D1

NVLL3C7NVLL3C11

NVLL3B9K11

EU167592 Cladosporium sp. *NVLL3A6

DQ066717 Friedmanniomyces simplexRVLL5C5

EU167572 Teratosphaeria microsporaDQ504331 Uncultured ascomycete cloneEU167608 Kabatiella microstictaNVLL3C10EF141324 Aureobasidium pullulans

EU167593 Dothidea muelleri AY220610 Scleroconidioma sphagnicolaRV12LL1A7RV12LL1B4NVLL3A3NVLL3C5

AF548087 Penicillium chrysogenum *K17

L27K5

K6K52M36

NVLL3A1RV2bfE3

EF638629 Uncultured basidiomycete cloneEF638564 Uncultured basidiomycete cloneAB075545 Filobasidium elegans

AY923093 Whipple RV cloneAF113430 Mucor racemosus

AY854021 Synchytrium endobioticumM59758 Chytriomyces hyalinus

5 changes

U05194 Alternaria alternata *

DQ471016 Pezicula carpinea

AY313953 Asteromassaria olivaceohirta

55

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Basidiomycota

ZygomycotaChytridiomycota

FIG. 6. One of the most parsimonious trees. Sequences from rock varnish and non-varnish site clones are presented in black and are prefixed with RV- or NV-respectively. The wide bold font indicates 18S sequences from Mn-oxidizing fungal isolates. Closest relatives are presented in gray with Mn-oxidizing relativesmarked with an asterisk (∗). Sequences within black boxes were selected from Kuhlman et al. (2006), a study of varnish fungi in California. Bootstrap values arebased on 1000 replicates.

Many species of Cladosporium are found in association withliving plants or with senescent plant material. Isolate K43 andclone NVLL3A8 were closely associated with Phoma herbarum(AY337712), a fungus that is known to parasitize humans,plants, and in the case of this AY337712, salmon. ClonesNVLL3C8 and NVLL3A7 grouped with Alternaria alternata(U05194), an opportunistic plant and human pathogen oftenfound as a saprobe on decaying plant material. Studies fromaround the world have consistently identified Phoma, Penicil-lium, Alternaria, Cladosporium, and Aureobasidium (Grishkanet al. 2006; Abdel-Hafez 1982; Oner 1970; Ranzoni 1968) assoil-inhabiting fungi, and these same taxa have been reportedas common inhabitants of semiarid grasslands in New Mexico(Porras-Alfaro et al. 2008, 2010, Khidir et al. 2010). Penicil-lium corylophilum has been demonstrated to be instrumental inlimestone patina formation (Fomina et al. 2010). Representa-

tives of other common soil fungal genera, such as Aspergillusand Fusarium, were not observed. According to Sterflinger andPrillinger (2001) Phoma and Sarcinomyces are common in-habitants on stone monuments in the Mediterranean. Sarcino-myces, among other genera such as Coniosporium, Acrodyctis,Phaeococcomyces, and Phaetheca, are known to produce MCF(Gorbushina 2003a; Gorbushina et al. 2003b).

In the phylogenetic analysis many of the rock varnishand non-varnish clones group close to Sarcinomyces sp.(AJ972809), which was identified as a MCF on marble inTurkey, suggesting that some of these fungi may exhibit mi-crocolonial growth. Cryomyces minteri (DQ066714), whichgrouped closely with RV12LL1A1, was identified in a studythat examined black fungi in the Antarctic desert. Fungal clonesfrom both non-varnish and rock varnish clones have been re-ported forming close associations with plants. For example,

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TABLE 3Nature of the closest relatives of clones from rock varnish (RV) and non-varnish (NV) sampling sites and for cultured fungal

isolates from rock varnish sampling sites

Nature of the closest relativeNumber of clones RV

sites 1 and 2Number of clones RV

site 5Number of clones NV

site 3Number of RV

cultured isolates

Rock-inhabiting 6 7Antarctic desert rock 6 6Other 1

Plant-associated 6 2 13 4Mycorrhizae 1 1Phytopathogens 2 3 3Other 4 1 9 1

Soil inhabitants 2Marine 2 5Other 2 2

Cenococcum geophilum forms mycorrhizal associations. Pezic-ula carpinea, Alternaria alternata, Kabatiella microsticta canbe plant pathogens, while Phaeosphaeriopsis nolinae, P. glauco-punctata, Ascochyta viciae-villosae, Bagnisiella examinans,Tetrasphaeria microspora, Aureobasidium pullulans, Sclero-conidioma sphagnicola, and Filobasidium elegans have beenidentified with living and dead plant material including desertplants such as yucca, although the Aureobasidium pullulanssequence included in this analysis was found in a marine envi-ronment.

Certain Lecophagus spp. (L. longispora, L. muscicola) havebeen found in soils; Friedmanniomyces simplex was isolatedfrom sandstone in Antarctica, and the uncultured ascomyceteclone (DQ504331) was recovered a carbonate, sea-water inter-face. It is possible that many of the fungal clones detected in thisstudy are chance inhabitants that were dispersed by wind to therock surface from the soil or plants. Many of the clones appearedto be novel sequences and did not closely group with any knownsequences (e.g., NVLL3D7, RV12LL2A7, RV12LL2A11). Dif-ferences between rock varnish sequences and non-varnish se-quences were not consistent throughout the tree. Isolates K11,K21, K43, K44 and L18 were more closely related to non-varnish clone sequences than to any of the rock varnish clones.

Although rock varnish and non-varnish clone sequences over-lapped in their placement on the phylogenetic tree, there wasvariability between samples from rock varnish sites 1 and 2, androck varnish site 5, which may be due to the differing amountsof water received by sites 1 and 2 versus site 5, which is anephemeral water course (Table 3). Fungal communities in aridecosystems are known to be highly diverse with high specialand temporal variability (Porras-Alfaro et al. 2010). A more ex-tensive analysis with a barcoding region such as ITS nrDNA isnecessary to describe in detail desert varnish fungal diversity.

Two studies by Kuhlman et al. (2006, 2008) have exam-ined the microbial community present on rock varnish using

molecular techniques. The latter study was conducted usingrock varnish from the Atacama Desert of Chile, which is oneof the driest places on Earth, and though bacteria sequenceswere identified, no fungal DNA appeared to be present, or atleast was not amplified. The earlier study examined rock var-nish from the Whipple Mountains in California and identifiednumerous prokaryote and eukaryotic inhabitants. Several of thefungal sequences from this study were included in our phylo-genetic analysis. The ascomycete sequences seemed to be mostclosely related to the non-varnish clones in our study. Althoughsome of our sequences grouped with genera (e.g., Phoma andAlternaria) represented in the Whipple Mountain study, manyof the clone sequences were not closely related to the sequencesfrom the Whipple Mountain rock varnish or their closest rel-atives, suggesting that the fungal community inhabiting rockvarnishes may be larger than previously expected.

The use of culture-based and culture-independent methodsin this study provides several lines of evidence that support therole of fungi in desert varnish formation. Not only are fungiconsistently found inhabiting rock varnish, but they are alsofound associated with the early stages of rock varnish formation,in many cases are capable of Mn-oxidation, and are closelyrelated to other fungal genera that are known to oxidize Mn.The suite of these findings suggests that at least some of thefungi present in our study sites are active participants in varnishformation. Future studies will explore their precise role.

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