1Department of Ecology and Evolutionary Biology, University of Tennessee, Knoxville,
TN 37996. 2Current address - Department of Biology, Box 125, 402 East College
Street, Bridgewater College, Bridgewater, VA 22812. 3Applied Biology and
Biomedical Engineering, Rose-Hulman Institute of Technology, Terre Haute, IN
47803. *Corresponding author - elickey@bridgewater.edu.
The Mushroom TWiG:
A Marvelous Mycological Menagerie in the Mountains
Edgar B. Lickey1,2,*, Shannon M. Tieken3, Karen W. Hughes1,
and Ronald H. Petersen1
Abstract - We present an update on efforts to catalog the basidiomycete taxa, particularly
the mushroom-forming fungi, of Great Smoky Mountains National Park
(GSMNP) for the All Taxa Biodiversity Inventory. The goals of this project are to:
1) collect, identify, and voucher specimens with the help of visiting mycologists and
volunteers; 2) extract DNA, amplify and sequence the nrITS region for barcoding,
and deposit these sequences on GenBank; and 3) create species web pages for general
public use. At present (April 2006), approximately 2000 specimens comprising
about 770 species have been collected. As many as 45% are new Park records, and
several may represent species new to science. DNA has been extracted from about
1000 specimens, and the nuclear ribosomal ITS region has been amplified and sequenced
for about 500 of those. A surprising amount of genetic heterogeneity has
been found, in part due to population migration patterns in response to glacial cycles.
Studies with Artomyces pyxidatus support this hypothesis, showing distinct contributions
from Central America and a second unidentified refugium.
Introduction
The southern Appalachian Mountain region, of which Great Smoky
Mountains National Park (GSMNP) serves as the major protective refuge, is
rich in fl oristic diversity and is an area of high endemism (Estill and Cruzan
2001). This is probably due to the wide variety of habitat types as well as
population-migration patterns caused by successive glacial periods. However,
little is known about how the mycota of this region compares to others
because intensive fungal inventories are uncommon in North America.
While several mycologists visited and collected in the southern Appalachians
in the late 1800s and early 1900s, reports of early forays in the area
of what is now GSMNP are very scarce (Petersen 1979). Among the earliest
was Rev. Moses Ashley Curtis who traversed the mountains in the late
1850s, but reported finding few macrofungi (Petersen 1979). In September
1916, Charles Kauffman spent three weeks in and around Elkmont and reported
283 species of fungi, including two new species (Kauffman 1917). It
was not until 1919, when Dr. Lexemuel Ray Hesler joined the faculty at the
University of Tennessee - Knoxville, that regular forays were made to the
GSMNP area. A preliminary checklist was published in 1937 and included
73
The Great Smoky Mountains National Park All Taxa Biodiversity Inventory:
A Search for Species in Our Own Backyard
2007 Southeastern Naturalist Special Issue 1:73–82
74 Southeastern Naturalist Special Issue 1
752 fungal taxa (Helser 1937). Following a number of years of collecting
by Hesler and a host of visiting mycologists, including forays associated
with the 1939 Mycological Society of America meeting, the checklist grew
to 1975 taxa by 1962, but this list was not published. Taxa were continually
added to the Park list subsequent to formal forays associated with the
Hesler Symposium in 1968, the 2nd International Mycological Congress in
1977, and periodic informal forays by students, faculty, and visiting mycologists.
Updated taxonomic treatments of large genera such as Hygrophorus
Fr. (Hesler and Smith 1963), Entoloma (Fr.) Kummer emend. Donk
(Hesler 1967), and Lactarius Pers. ex S.F. Gray (Hesler and Smith 1979)
added even more taxa to the list, for a total of nearly 2200 when it was
published in its most recent form as a National Park Service management
report (Petersen 1979).
During the establishment of the All Taxa Biodiversity Inventory program
in GSMNP in 1997, a fungal taxonomic working group (TWiG) was formed
marking the start of the current period of intensive collecting. The original
organizational effort was the product of collaboration between Amy Rossman,
Lorelei Norvell, Rod Tulloss, the North American Mycological Association
(NAMA), and the Asheville Mushroom Club. This endeavor was
further bolstered by a National Science Foundation grant awarded in 2004 to
K.W. Hughes and R.H. Petersen to concentrate on the Homobasidiomycete
mycota (mushrooms and their allies). The goals of this grant, in addition to
updating the GSMNP checklist, include gathering sequence data, based on
the nuclear ribosomal inter-transcribed spacer region (nrDNA ITS), which
will be made public on the GenBank database for “barcoding,” and the
creation of species webpages for public use. The timing of this grant conveniently
coincided with the joint Mycological Society of America - North
American Mycological Association meeting in Asheville, NC in July, 2004,
bringing together many mycologists, both amateur and professional. To take
advantage of such a convergence of mycological expertise, two “mycoblitzes”
(intensive collecting of fungi over a four-day period) were executed,
funded in large part by the NSF grant.
Since biodiversity also includes infraspecific genetic diversity, the
preliminary results of an ongoing study with populations of Artomyces
pyxidatus (Pers.: Fr.) Jülich (also known as Clavicorona pyxidata (Pers.: Fr.)
Doty) will also be presented. This work is a continuation of studies by Tieken
(2002) and Lickey et al. (2002), which showed the existence of two major
nrDNA ITS haplotypes in North America, one predominantly northeastern
and one southwestern Central American, with a putative introgression zone
in the southeastern United States.
Methods
Collections
Collecting trips were made at least once a week in the summer and fall
months, and at least once a month in the winter and spring. Locations were
chosen to give a broad spectrum of geography (i.e., east and west ends) and
2007 E.B. Lickey, S.M. Tieken, K.W. Hughes, and R.H. Petersen 75
habitat types (e.g., high and low elevation, cove hardwood and oak-pine forests,
etc.) at approximately the same time of year. Fresh basidiomata were
collected and stored in either fishing tackle boxes or wrapped in wax paper
or aluminum foil for transport back to the lab. In the lab, collections were assigned
a field number, and notes were made concerning location (state, county,
trail or area, and GPS coordinates), date, and collector. Descriptive notes such
as substrate, size, stature, vestiture, color (standards based on Kornerup and
Wanscher 1967), odor, and taste were sometimes included. Photographs of
each collection were taken in natural light on an 18% gray card using Kodak
Elitechrome 100 ASA film. An approximately 100–200-mg tissue sample
of one basidioma was frozen in foil for later DNA extraction. Cultures were
made for some species if their basidiomata were too small or too few to sacrifice voucher material. In this case, either tissue or spores were placed directly
on malt extract agar (15 g malt extract, 20 g agar, 1 L distilled H2O) to obtain
a dikaryotic mycelium which was then grown in potato dextrose broth (24 g
in 1 L distilled H2O) to yield enough tissue for DNA extraction. The rest of
the collection was placed in a food dehydrator for 24–48 hrs and later was
identified, databased, and deposited in the University of Tennessee Herbarium
(TENN). For population studies involving Artomyces pyxidatus haplotype
distribution, multiple collections were made in Cataloochee Cove, NC,
Greenbrier Cove, TN, and near Sugarlands Visitor Center, TN.
Identifications
Collections were cursorily identified using an initial set of keys such as
Largent and Baroni (1988) and Bessette et al. (1997), followed by more specific
keys (e.g., published monographs) to more accurately identify collections to
species. Microscopic characters were examined using typical techniques with
small amounts of tissue and/or thin, hand-sliced sections mounted on slides in
3% aqueous KOH. Iodine-based Melzer’s reagent (see Largent et al. 1988) was
also used where tests for amyloidity were needed. Collections representing taxonomically
difficult genera were sent to experts in those groups for identification.
Molecular analysis
DNA was extracted from frozen, liquid culture, or dried herbarium tissue
using a modified CTAB method (Hughes et al. 1999) or Qiagen DNeasy
Plant Mini kit. The nuclear ribosomal internal transcribed spacer region (ITS1
- 5.8S - ITS2) was PCR-amplified using primers ITS1f (Gardes and Bruns
1993) and ITS4 (White et al. 1990) or ITS4b (Gardes and Bruns 1993), and the
large subunit (LSU) was amplified using primers ITS3 (White et al. 1990) and
LR7 (Moncalvo et al. 2000). Amplification reactions for ITS used the following
parameters: 94 °C for 4 min; 35 cycles of 94 °C for 1 min, 52 °C for 1 min,
and 72 °C for 1 min; 72 °C for 4 min. The same parameters were used for the
LSU region, except that the 72 °C extension step was increased to 2 min.
PCR products were cleaned using ExoSAP-IT (United States Biochemical),
and sequenced with ABI Prism Big Dye Terminator v. 3.1 using primers
ITS5 and ITS4 (White et al. 1990) for the ITS region and LR5 (Moncalvo
et al. 2000) for the LSU region. Sequencing reaction parameters were 25
cycles of 96 °C for 10 sec, 50 °C for 5 sec, and 60 °C for 4 min, and products
76 Southeastern Naturalist Special Issue 1
were electrophoresed on an ABI 3100 automated sequencer (University of
Tennessee Molecular Biology Research Facility). Resulting electropherograms
were viewed and edited using Sequencher 4.1.2 (GeneCodes), and
GenBank BLAST searches were done to determine presence or absence of
similar sequences in that database. When electropherograms were unreadable
due to apparent insertion-deletion events that could not be resolved
with sequencing with different primers, PCR products were cloned and
screened using the pGEM-T vector system and JM109 competent cells following
manufacturer’s directions (Promega).
Restriction enzymes CfoI and BsaJI (Promega) were used in the haplotype
analysis of Artomyces pyxidatus populations following Tieken (2002)
and Lickey et al. (2002). Sequencing as described above was sometimes
used to verify presence or absence of restriction sites.
Results
Collections
A comparison of the numbers of species included in GSMNP checklists
since 1937 of all fungi, Homobasidiomycetes, and agarics and allies is presented
in Figure 1. Since the start of the fungal TWIG of the ATBI in 1998, and
as of April 2006, approximately 1900 collections have been made, representing
about 770 species and bringing the total number of fungi known to occur
in the Park to over 2500 species (Fig. 1). Of those 770 species, 347 (45%) are
new records to GSMNP, and possibly as many as 25 are species new to science,
according to experts in selected fungal groups (pers. comm. with: Bart Buyck,
Museum National D’Histoire Naturelle, Paris, France [Russula Pers. spp.]; Urmas
Koljag, University of Tartu, Estonia [Tomentella Pat. spp.]; Karl-Henrik
Larsson, University of Göteborg, Sweden [Corticium Fr. spp.]; Brandon
Matheny, Clark University, Worcester, MA [Inocybe (Fr.) Fr. spp.]; and Rod
Tulloss, New York Botanical Garden, New York, NY [Amanita Pers. spp.]).
However, a large backlog of collections awaits further study for identification.
Molecular analysis
DNA has been extracted from more than 1000 collections, and the nrDNA
ITS region has been amplified and sequenced for more than 500 collections. An
unexpectedly high proportion of collections were apparently heterozygous for
insertion-deletion mutations (indels). A few examples are included in Table 1.
For most of these, sequences had to be obtained through cloning procedures.
Collections of Artomyces pyxidatus revealed that both the northeastern
and southwestern haplotypes identified by Lickey et al. (2002) and
Tieken (2002) are present in the three low-elevation coves. The northeastern
haplotype is present in much higher frequencies in all three populations, and
apparent heterozygotes have been identified (Tieken 2002).
Discussion
It is difficult at this time to estimate how many basidiomycete species will
be found, much less exist undetected, in GSMNP. The rate of new discoveries
2007 E.B. Lickey, S.M. Tieken, K.W. Hughes, and R.H. Petersen 77
versus accounts of species known from the Park has been outstanding; approximately
45% of the species collected are new Park records. In other words,
for every account of a 1979-checklist species we are finding at least one new
record. In addition, large areas of GSMNP, such as the “Northshore” area in
North Carolina, remain unsurveyed due to their remoteness and inaccessibility.
On the other hand, only about 400 of the species collected since the start
of the ATBI were included on the 1979 checklist which listed almost 1600
basidiomycetes (Petersen 1979). These apparently low re-collection numbers
may be due to several factors. First, the species that have not been re-collected
could be rare or not fruit for long periods of time and by chance have not been
re-encountered, or they might possibly be extirpated. Second, there may be a
better understanding of the taxonomy of some groups where segregate species
that were previously recognized have been synonymized, while others have
only recently been recognized as distinct species. Third, recent collections may
Figure 1. A comparison of the numbers of species of all fungi (Total), Homobasidiomycetes,
and agarics and their allies included in the three previous checklists (Hesler
1937; Hesler, unpubl. data; Petersen 1979) and the current list as of 12 March, 2006.
*The number of agarics for 2006 is an estimate.
Table 1. Numbers of nrDNA ITS sequences for representative mushroom groups and the percentage
exhibiting indel heterozygosities.
Mushroom group # nrITS sequences % indel heterozygous
Hygrophorus s.l. 40 58.0%
Cortinarius 48 31.0%
Lactarius 40 25.0%
Saprobes 309 28.5%
78 Southeastern Naturalist Special Issue 1
have been identified as a different species than what they were identified as on
the 1979 checklist either because of misidentification or differences in character
interpretations. Finally, as seen in Table 2, less than half the number of
species of the largest, but often taxonomically difficult, agaric genera included
on Petersen’s 1979 checklist have been re-collected. While some collections
of Cortinarius Fr., Entoloma and Mycena S.F. Gray have been made and await
identification by experts in these groups, active collecting has been somewhat
limited in favor of other more easily identifiable taxa.
The ephemerality and phenology of most mushroom taxa are not fully
understood, but they can have a large effect on fl oristic studies. In a 21-year
study of a 1500-m2 plot in a Swiss forest, Straatsma et al. (2001) found that of
the more than 400 species of mushrooms, only eight were observed in every
year while an average of 17 were recorded as new occurrences in the last five
years of sampling. The numbers of species they reported per year ranged from
18 in 1989 to 194 in 1992, indicating that productiveness can change drastically
with conditions (Straatsma et al. 2001). As with the present study, rarity
of some taxa and the transient nature of others due to habitat and environmental
changes over time may account for some of the disparity in the numbers
collected from year to year. Even though fruit bodies are the most visible evidence
we have of a mushroom species’ occurrence, fruiting is only a small part
of the life cycle and is often tied to specific environmental conditions. Because
of this, some species may not be seen if they fruited at times when we are not
in the field, if they lie dormant for long periods of time waiting for favorable
fruiting conditions, or even if they forgo sexual reproduction altogether. An
example from the present study involves Collybia (Fr.) Staude subgenus Collybia
section Collybia (Halling 1983), a group of small mushrooms that often
grow on the remains of other mushrooms and referred to here as the “microcollybias.”
While sequencing the nrDNA ITS region for a collection of Lactarius
griseus Peck, a second sequence was isolated that apparently belongs to a
microcollybia. It is likely that the Lactarius fruit body was colonized by the
Collybia with no evidence of fruiting, but based on sequence comparisons
with known microcollybia species, it is apparently undescribed. Had we not
incorporated molecular data in our study, this species might still have gone
Table 2. Number of species of the largest genera listed in Petersen’s 1979 checklist compared
with the numbers that have been re-collected and those that are Park records.
1979 list Re-collectedA New Park records
Cortinarius 103 # 1
Entoloma s.l. 101 # 1
Lactarius 86 34 15
Russula 74 14 15B
Mycena 73 # 2
Hygrophorus s.l. 55 15 4B
Inocybe 43 6 13B
Amanita 34 17 16B
A# indicates that less than 1/3 the number of the 1979 checklist have been re-collected.
BSpecies putatively new to science have been identified, but their number is not included with
the new Park records.
2007 E.B. Lickey, S.M. Tieken, K.W. Hughes, and R.H. Petersen 79
undetected. Likewise, environmental sampling, such as that described by
O’Brien et al. (2005), will undoubtedly uncover many more species that have
been overlooked because their fruiting has yet to be observed.
The numbers of taxa included in the growing checklist may be underestimated
due to the potential existence of cryptic species—those that
are morphologically identical, but in reality represent distinct biological
species. There are a number of mushroom “morphospecies,” such as
Xeromphalina campanella (Batsch: Fr.) Kühner & Maire (Johnson 1997),
Marasmius androsaceus (L.: Fr.) Fr. (Gordon 1994), Gymnopus subnudus
(Ellis: Peck) Halling (Murphy and Miller 1997), which may represent
more than one “biological species” (Petersen 1995). These studies, based
on crosses between collections, have shown that intersterile populations
coexist in the southern Appalachians and further draw attention to possible
intersterility substructuring in morphological species that are assumed to
have intercontinental distributions.
Many genera, for example the genus Cortinarius, may harbor cryptic
species, as over half the species recorded from GSMNP are based on European
names (Table 2). While mating studies are not feasible at present in
mycorrhizal species such as Cortinarius, molecular data, particularly nrDNA
ITS sequences, may shed some light on the degree of genetic differentiation.
Lack of or limited amounts of gene fl ow among geographically separated
populations have been demonstrated using molecular data in several morphospecies
including: Artomyces pyxidatus (Lickey et al. 2002) Lentinellus
castoreus (Secretan apud Fr.) Kühner & R. Maire and L. ursinus (Fr.) Kühner
(Hughes and Petersen 2004) and Flammulina velutipes (Fr.) Karsten (Methven
et al. 2000). With little or no gene fl ow, European and North American taxa
may have diverged genetically. However, the degree of genetic divergence
needed to identify North American taxa as different from their European
counterpart will remain unresolved until many more collections are made and
compared to topotype collections from Europe.
For some fungi, the number of species may be overestimated due to differences
in taxonomic interpretations among researchers, leading some to
recognize multiple taxa where others would simply recognize one. For example,
there are 101 Entoloma s.l. species included in the 1979 checklist.
In his monograph, Hesler (1967) described 59 of those species, 54 of which
are based on type material collected in GSMNP, and of those, 32 are known
only from the type collection. These 32 taxa may not have been encountered
again because they are extremely rare, exhibit infrequent fruiting,
or might simply have been avoided because of the difficulty in identifying
species in this very taxonomically challenging group. On the other hand, it
is possible that some of these may represent aberrations or are simply part
of the range of natural variability, genetic or plastic. Again, more collections
and more complete genetic study are warranted to determine the true
extent of species boundaries.
One genus that has recently received such attention is Amanita, being studied
by Rod Tulloss, one of the original ATBI Fungal TWIG coordinators. Of
80 Southeastern Naturalist Special Issue 1
the 34 species included on the 1979 checklist, only half of those have been recollected,
while at least 16 new Park records have been found along with eight
species putatively new to science (a periodically updated list can be found
at Tulloss’ website - http://pluto.njcc.com/~ret/amanita/key.dir/am_gsmnp.
html). Genetic data in the form of nrDNA ITS sequences are being gathered
in conjunction with morphological studies to aid in species identification and
delimitation. At the rate of collecting new records and putative new species of
Amanita, and with respect to the number already reported from the Park and
the number species known from the southern Appalachian region, Tulloss
(2005 version of website) believes as many as 200 species of Amanita may
inhabit GSMNP.
In the interest of gathering data for barcoding efforts, a large number of
nrDNA ITS sequences are being accumulated and will be deposited in the
publicly accessible GenBank sequence database after identifications have
been verified. Not surprisingly, GenBank BLAST searches on some of the
sequences completed so far have yielded few matches of ≥97%, an arbitrary
but conservative upper estimate of infraspecific ITS sequence divergence
(O’Brien et al. 2005). Only about a third of our sequences have such conspecific counterparts, illustrating how incomplete the database is, but also
emphasizing the need for depositing sequences of correctly identified species.
There are also a number (≈5%) of our sequences that matched (≥97%)
entries listed as unidentified, uncultured basidiomycete from environmental
samples, and as more sequences are generated from identifiable specimens,
names may be attached to these unknown environmental samples.
We have also found that the amount of genetic diversity in the Park is higher
than we had expected. Multiple nrDNA ITS haplotypes have been found
for many different taxa and they often appear as heterozygotes (Table 1).
One explanation for this phenomenon is that present-day populations are
infl uenced by past events, such as glacial cycles. Phylogeographic patterns
associated with glacial cycles have been used to explain population genetic
patterns observed in fungi such as Artomyces pyxidatus (Lickey et al. 2002,
Tieken 2002); Lentinellus ursinus and L. castoreus (Hughes and Petersen
2004); and Rhodocollybia butyracea (Bull.: Fr.) Lennox, Gymnopus biformis
(Peck) Halling, and Flammulina velutipes (K. Hughes, unpubl. data); as
well as in some plants such as Liriodendron tulipifera L. (Parks et al. 1994,
Sewell et al. 1996). It is hypothesized that during the last glacial maximum,
populations of plants (and presumably fungi) tolerated ecological conditions
far south of their present ranges in what have been termed refugia (Delcourt
and Delcourt 1981). We posit, as did Parks et al. (1994), Sewell et al. (1996),
and several others, that populations separated in geographically isolated
refugia accumulated mutations creating new and unique haplotypes. As the
climate ameliorated and glaciers receded, populations expanded and reconnected
on the way to their present-day ranges, and these divergent haplotypes
introgressed. In the case of Artomyces pyxidatus, both the northeastern and
southwestern haplotypes are present in the GSMNP, with the northeastern
haplotype maintained in higher frequencies. The occurrence of these apparent
hybrid haplotypes indicates the presence of interbreeding and introgression.
2007 E.B. Lickey, S.M. Tieken, K.W. Hughes, and R.H. Petersen 81
This same phenomenon may be similar in the many different species of fungi
where multiple haplotypes are being found.
While great strides have been made in cataloging the mycota of the
GSMNP, much work remains to be done. We are continually finding new
Park records and seem to be a long way from reaching the asymptote of the
number of fungal species occurring in the Park. In this regard, herbarium
work is just as important as continued field surveys to identify both new
Park records as well as species new to science. More also needs to be done
to study the amazing amount of genetic diversity that this region harbors.
Species webpages, our public-outreach medium, are still in development,
but will soon provide species descriptions, collection locations, and links to
nrDNA ITS sequences deposited in GenBank.
Acknowledgments
The authors thank two anonymous reviewers for their comments and suggestions.
The following people are thanked for their collections and expert identifications: Cathie
Aime, Joe Ammirati, Rich Baird, Meredith Blackwell, Bart Buyck, Julieta Carranza,
Joaquin Cifuentes, Roy Halling, Rick Kerrigan, Jean Lodge, Juan Mata, Brandon Matheny,
Coleman McCleneghan, Andy Methven, Andy Miller, Lorelei Norvell, Clark
Ovrebo, Amy Rossman, Somsak Sivichai, Rod Tulloss, and Annamieke Verbeken, as
well as Vince Hustad, Sean Jones, Huzefa Raja, and Matt Woods. Others that helped in
lab and/or field include Rob Shepard, Pamela Pokorny, Jason Smith, Beth Helmbrecht,
Stacy Huskins, Kim Kennard, and Jessica Hite. We also thank Pat Cox for the organization
of and invitation to contribute to this special ATBI Symposium. Funding for this
project was provided by NSF DEB grants 9521526 to R.H. Petersen and K.W. Hughes
and 0338699 to K.W. Hughes and R.H. Petersen, as well as the Hesler Endowment Fund.
Literature Cited
Bessette, A.E., A.R. Bessette, and D.W. Fischer. 1997. Mushrooms of Northeastern
North America. Syracuse University Press, Syracuse, NY. 584 pp.
Delcourt, P.A., and H.R. Delcourt. 1981. Vegetation maps for eastern North America:
40,000 yr b.p. to the present. Pp. 123–165, In R.C. Romans (Ed.). Geobotany II.
Plenum Press. New York, NY. 263 pp.
Estill, J.C., and M.B. Cruzan. 2001. Phytogeography of rare plant species endemic
to the Southeastern United States. Castanea 66:3–23.
Gardes, M., and T.D. Bruns. 1993. ITS primers with enhanced specificity for basidiomycetes:
Application to the identification of mycorrhizae and rusts. Molecular
Ecology 2:113–118.
Gordon, S.A. 1994. Intraspecific variation in three species of Marasmius. Ph.D Dissertation.
University of Tennessee, Knoxville, TN. 249 pp.
Halling, R.E. 1983. The genus Collybia (Agaricales) in the northeastern United
States and adjacent Canada. Mycologia Memoir No. 8. J. Cramer Publisher,
Braunschweig, Germany. 148 pp.
Hesler, L.R. 1937. A preliminary checklist of the fungi of the Great Smoky Mountains
National Park. Castanea 2:45–58.
Hesler, L.R. 1967. Entoloma in Southeastern North America. J. Cramer, Stuttgart,
Germany. 244 pp.
Hesler, L.R., and A.H. Smith. 1963. North American Species of Hygrophorus. The
University of Tennessee Press, Knoxville, TN. 416 pp.
82 Southeastern Naturalist Special Issue 1
Hesler, L.R., and A.H. Smith.1979. North American Species of Lactarius. The University
of Michigan Press, Ann Arbor, MI. 841 pp.
Hughes, K.W., and R.H. Petersen. 2004. A phylogenetic reconstruction of the genus
Lentinellus. Pp. 249–268, In R.H. Petersen and K.W. Hughes (Eds.). A Preliminary
Monograph of Lentinellus (Russulales). Bibliotheca Mycologica no. 198, J.
Cramer, Berlin, Germany. 283 pp.
Hughes, K.W., L.L. McGhee, A.S. Methven, J.E. Johnson, and R.H. Petersen. 1999.
Patterns of geographic speciation in the genus Flammulina based on sequences
of the ribosomal ITS1-5.8S-ITS2 area. Mycologia 91:978–986.
Johnson, J.E. 1997. Systematics of the Xeromphalina campanella complex. Ph.D.
Dissertation. University of Tennessee, Knoxville, TN. 337 pp.
Kauffman, C.H. 1917. Tennessee and Kentucky fungi. Mycologia 9:159–166.
Kornerup, A., and J.H. Wanscher. 1967. Methuen Handbook of Color, 2nd Edition.
Methuen and Co., Ltd., London, UK. 243 pp.
Largent, D.L., and T.J. Baroni. 1988. How to Identify Mushrooms to Genus VI:
Modern Genera. Mad River Press, Eureka, CA. 277 pp.
Largent, D.L., D. Johnson, and R. Watling. 1988. How to Identify Mushrooms to
Genus III: Microscopic Features. Mad River Press, Eureka, CA. 148 pp.
Lickey, E.B., K.W. Hughes, and R.H. Petersen. 2002. Biogeographical patterns in
Artomyces pyxidatus. Mycologia 94:461–471.
Methven, A., K.W. Hughes, and R.H. Petersen. 2000. Flammulina RFLP patterns
identify species and show biogeographical patterns within species. Mycologia
92:1064–1070.
Moncalvo, J.-M., F.M. Lutzoni, S.A. Rehner, J. Johnson, and R. Vilgalys. 2000. Phylogenetic
relationships of agaric fungi based on nuclear large subunit ribosomal
DNA sequences. Systematic Biology 49:278–305.
Murphy, J.F., and O.K. Miller, Jr. 1997. Diversity and local distribution of mating
alleles in Marasmiellus praeacutis and Collybia subnuda (Basidiomycetes, Agaricales).
Canadian Journal of Botany 75:8–17.
O’Brien, H.E., J.L. Parrent, J.A. Jackson, J-M. Moncalvo, and R. Vilgalys. 2005.
Fungal community analysis by large-scale sequencing of environmental samples.
Applied and Environmental Microbiology 71:5544–5550.
Parks, C.R., J.F. Wendel, M.M. Sewell, and Y.L. Qiu. 1994. The significance of
allozyme variation and introgression in the Liriodendron tulipifera complex
(Magnoliaceae). American Journal of Botany 81:878–889.
Petersen, R.H. 1979. Checklist of fungi of the Great Smoky Mountains National
Park. Management Report No. 29. Department of the Interior, National Park
Service, Southeast Region, Uplands Field Research Laboratory, Great Smoky
Mountains National Park, Twin Creeks, Gatlinburg, TN. 103 pp.
Petersen, R.H. 1995. Contributions of mating studies to mushroom systematics.
Canadian Journal of Botany 73(Suppl. 1):S831–S842.
Sewell, M.M., C.R. Parks, and M.W. Chase. 1996. Intraspecific chloroplast DNA
variation and biogeography of North American Liriodendron L. (Magnoliaceae).
Evolution 50:1147–1154.
Tieken, S.M. 2002. Artomyces pyxidatus (Auriscalpiaceae, Fungi) Jülich in the
Great Smoky Mountains National Park: Individuals and populations. M.Sc. Thesis.
University of Tennessee, Knoxville, TN. 66 pp.
Straatsma, G., F. Ayer, and S. Egli. 2001. Species richness, abundance, and phenology
of fungal fruit bodies over 21 years in a Swiss forest plot. Mycological
Research 105:515–523.
White, T.J., T. Bruns, S. Lee, and J.W. Taylor. 1990. Amplification and direct sequencing
of fungal ribosomal RNA genes for phylogenetics. Pp. 315–322, In M.A. Innis,
D.H. Gelfand, J.J. Sninsky, and T.J. White (Eds.). PCR Protocols: A Guide to
Methods and Applications. Academic Press, Inc., New York, NY. 482 pp.