Mycological Society of America

Mycological Society of America
Nuclear Migration in Diploid-Haploid Pairings of Armillaria ostoyae
Author(s): D. M. Rizzo and T. C. Harrington
Source: Mycologia, Vol. 84, No. 6 (Nov. - Dec., 1992), pp. 863-869
Published by: Mycological Society of America
Stable URL: .
Accessed: 02/03/2011 19:16
Your use of the JSTOR archive indicates your acceptance of JSTOR's Terms and Conditions of Use, available at . JSTOR's Terms and Conditions of Use provides, in part, that unless
you have obtained prior permission, you may not download an entire issue of a journal or multiple copies of articles, and you
may use content in the JSTOR archive only for your personal, non-commercial use.
Please contact the publisher regarding any further use of this work. Publisher contact information may be obtained at . .
Each copy of any part of a JSTOR transmission must contain the same copyright notice that appears on the screen or printed
page of such transmission.
JSTOR is a not-for-profit service that helps scholars, researchers, and students discover, use, and build upon a wide range of
content in a trusted digital archive. We use information technology and tools to increase productivity and facilitate new forms
of scholarship. For more information about JSTOR, please contact [email protected]
Mycological Society of America is collaborating with JSTOR to digitize, preserve and extend access to
Mycologia, 84(6), 1992, pp. 863-869.
© 1992, by The New York Botanical Garden, Bronx, NY 10458-5126
D. M. Rizzo
Department of Plant Pathology, University of Minnesota, St. Paul, Minnesota 55108
Department of Plant Pathology, Iowa State University, Ames, Iowa 50011
Diploid-haploid pairings to identify field isolates of Armillaria to species generally rely on morphological differences between diploid and haploid colonies (flat and fluffy colonies, respectively) and
morphological changes in haploid tester colonies. Presumably, the change in morphology of the haploid
tester is due to diploidization. Isozyme markers, aconitase and glucosephosphate isomerase, were used
to follow nuclear migration from putatively diploid isolates into a haploid tester strain of A. ostoyae.
After pairing diploid field isolates with a haploid tester for 3 or 4 weeks, subcultures were taken from
the haploid tester at 2 and 10 mm from the confrontation zone. Subcultures from 2 mm were mostly
flat; subcultures from 10 mm were fluffy, flat, or fluffy with flat sectors. Hyphal tip cultures from the
subcultures were analyzed for the isozyme markers. All fluffy hyphal tip cultures had the isozyme
phenotype of the haploid tester. Flat hyphal tip cultures either had the isozyme phenotype of the diploid
parent or had a hybrid isozyme phenotype with markers of both the diploid parent and the haploid
tester. The data suggest that in conspecific diploid-haploid pairings, the diploid nucleus usually migrates
into the haploid mycelium and eventually displaces the haploid nucleus.
Key Words: Armillaria ostoyae, genetics, identification, isozymes
Identification of species of Armillaria (Agaricales: Tricholomataceae) in the field is difficult
because characteristics of mycelial fans (RollHansen, 1985) and rhizomorphs (Berub6 and
Dessureault, 1988, 1989; Guillaumin et al., 1989)
vary among the species only to a limited extent,
and because basidiomes (mushrooms) are seasonal and rare to uncommon in most regions. In
vitro production of basidiomes is possible for
some species (particularly A. ostoyae (Romagn.)
Herink.), but the process is time-consuming
(Guillaumin, 1986; Guillaumin et al., 1989; Harrington et al., 1992). This limitation and the limited variation among Armillaria species in cultural characteristics (Guillaumin et al., 1985,
1989; Rishbeth, 1986) make identification of field
isolates difficult and have impeded progress on the
elucidation of the biology of the various species.
Electrophoresis of isozymes (Morrison et al.,
1985) and restriction fragments of DNA (Anderson et al., 1987; Jahnke et al., 1987; Smith
and Anderson, 1989), DNA homology (Jahnke
et al., 1987), DNA sequence analysis (Anderson
and Stasovski, 1992) or antibodies (Burdsall et
al., 1990) may prove feasible for species identifications in the future. For the present, critical
identification of cultures relies on pairings with
tester strains.
Although various pairing techniques have been
utilized, most rely on the variation in culture
morphology between single basidiospore strains
(haploid) and isolates from decay, fans or rhizomorphs, which are putatively diploid (Hintikka, 1973). Most diploid isolates produce relatively little aerial mycelium, and the mycelium
often has a reddish-brown crust. Haploid strains
tend to produce more aerial mycelium and the
colonies generally remain white. When sexually
compatible haploid strains of the same species
are placed near each other and allowed to grow
together for 3 or more weeks, the fluffy, white
mycelium may flatten and become darker or
crustose. If the strains are of different species or
are homoallelic at one or both of the two mating
loci (the investigated species of Armillaria are
tetrapolar, with two multiallelic loci determining
sexual compatibility), the mycelia remain white
or light tan, continue to produce abundant aerial
hyphae and do not become crustose (Korhonen,
1978). Evidence suggests that the colony type
arising from conspecific pairings of haploids is
diploid (Anderson and Ullrich, 1982; Korhonen,
1980; Korhonen and Hintikka, 1974; Larsen et
al., 1992; Peabody et al., 1978).
Diploid isolates are not as easy to identify as
haploid strains (Shaw and Loopstra, 1988; Siepmann, 1987; Wargo and Shaw, 1985). Diploidhaploid pairings can be made in a manner analogous to dikaryon-monokaryon pairings used in
dikaryotic hymenomycetes (Korhonen, 1978).
Although this technique has been used by many
investigators (Anderson and Ullrich, 1982; Guillaumin and Berthelay, 1981; Harrington, 1987;
Harrington et al., 1992; Kile, 1983; Mallet, 1990;
Rishbeth, 1982), the genetic evidence for the
conversion of the haploid to a diploid in diploidhaploid pairings has not been established (Ullrich and Anderson, 1988; Guillaumin et al., 1991).
The objective of our study was to determine
the genetic basis for diploidization of diploidhaploid pairings of Armillaria ostoyae. This was
tested by using isozyme markers to follow nuclear migration from diploid isolates into haploid
tester mycelia.
Polymorphic isozymes were searched for in five single basidiospore strains and nine putatively diploid
isolates of A. ostoyaefrom New Hampshire.One haploid strain (B286 from Abies balsamea (L.) Mill., Albany, New Hampshire) and two diploid isolates (B356
and B359 from Tsuga canadensis (L.) Carr. and A.
balsamea, respectively, Bartlett, New Hampshire) were
selected because of differences in electromorphs for
aconitase (ACO; EC number and glucosephosphate isomerase (GPI; EC (FIGS. 1, 2).
Two series of diploid-haploid pairings (B356 x B286
and B359 x B286) were conducted on MEA (1% Difco
malt extract, 1.5% Difco agar) in 50-mm-diam plastic
petri dishes. Inocula were placed 5 mm apart and incubated at room temperature (20 to 25 C) with laboratory lighting. Subcultures were taken from the haploid tester side (B286) after 3 and 4 wk with five replicate
plates for each sampling time. From each plate, one or
two 3-mm-diam plugs were taken from a point 2 mm
and 10 mm (generally the advancing margin) from the
confrontation zone. Where the haploid tester had grown
further than 10 mm, subcultures were also taken from
the advancing margin. Subculture plugs were transferred to MEA and incubated under laboratory conditions for 7 to 10 days before evaluating for culture
type (fluffy, flat, or fluffy with flat sectors).
Hyphal tips were excised using 40 x magnification
from subcultures taken at each sampling time and distance from the confrontation zone. Hyphal tips were
taken without regard to subculture morphology but
were selected on the basis of ease of excision, which
resulted in a bias for fast-growing hyphal tips. A minimum of 40 hyphal tips were taken for each sampling
time and sampling distance. Hyphal tips were transferred to MEA, incubated as above for 14 to 16 days,
and scored for culture type.
Representative samples of hyphal tip cultures (fluffy
and flat from each distance and time) were grown in
liquid media for enzyme extraction. Plugs of mycelia
from MEA plates were inoculated into 30 ml of liquid
medium (20 mg malt extract and 2 mg yeast extract
per ml) in 125 ml erlenmeyer flasks and incubated in
still culture at 20 to 25 C for 3 to 4 wk. Mycelia (and
rhizomorphs, if present) were vacuum-filtered, placed
in prechilled mortars, frozen with liquid nitrogen, and
ground to a fine powder. Enzymes were extracted by
grinding with 0.5 to 0.75 ml of extraction buffer. Extraction buffer was prepared by mixing 0.2 M Na2PO4
(25 ml), glycerol (25 ml), deionized water (50 ml) and
disodium EDTA (0.17 g), and adjusting the pH to 7.2
with 4 N NaOH. Lyophilized bovine serum albumin
(1 g), sodium ascorbate (1 g), glutathione (1 g), NAD
(1 mg), and NADP (1 mg) were added to the buffer
and the pH readjusted to 7.2. Enzyme extracts were
absorbed through miracloth onto 4 x 12 mm wicks of
Whatman No. 3MM chromatography paper and stored
at -80 C.
Twelve percent starch gels were prepared 1 day in
advance by the microwave method of Marty et al.
(1984). Buffer systems D (pH 6.1) and E (pH 8.1) were
continuous morpholine citrate systems run at 250 V
constant voltage for 5 h (Conkle et al., 1982). Gels were
loaded with up to 30 wicks, each representing a different isolate or strain. Wicks were removed after 10
min of electrophoresis at the specified voltage. Staining
procedures were those of Marty et al. (1984).
Although genetic analysis of the segregation of isozyme electromorphs through meiosis was not conducted, the number of coding loci and the monomeric
or dimeric nature of the enzymes could be easily inferred (Micales et al., 1986). Electrophoresis of over 50
putatively diploid field isolates and haploid strains of
A. ostoyae in this study and in other unpublished work
(Rizzo and Harrington) indicated that ACO is monomeric and has two loci. Although only one locus for
ACO has been noted in some organisms (Pasteur et
al., 1988), all haploid strains of A. ostoyae had two
bands (FIG. 1). Putatively diploid field isolates of A.
ostoyae had two, three, or four bands; four bands would
be expected if the isolate was heterozygous at both loci.
No heteromeric isozyme molecules were suggested by
the electrophoretic mobility of the bands (FIG. 1), and
ACO has been found to be monomeric in other organisms (Pasteur et al., 1988).
The enzyme GPI has been shown to be dimeric with
one locus in many organisms (Pasteur et al., 1988),
including fungi (Linde et al., 1990), and appeared to
be dimeric with a single locus in A. ostoyae (FIG. 2).
All haploid strains tested had only one band for GPI,
and a sampling of over 50 putatively diploid field isolates of A. ostoyae had either one band (putatively homozygous) or three bands (putatively heterozygous, with
a strongly staining band migrating halfway between the
~~~~ ~ ~~~~~~
. ..
. ::.
FIG.1. Electromorphsof eight hyphal tip cultures
of Armillaria ostoyae for aconitase (ACO). Cultures
wereeitherfluffy(haploid,H) or flat (putativediploid,
D, or hybrid,HD). The H phenotypewas the same as
B286 and the D phenotypewas the same as B359. The
HD phenotyperesultedfrompairingdiploidB359 with
haploid B286.
FIG.2. Electromorphsof seven hyphaltip cultures
of Armillariaostoyae for glucosephosphateisomerase
(GPI). Cultureswere either fluffy(haploid, H) or flat
(putativediploid, D, or hybrid,HD). The H phenotype
was the same as B286 and the D phenotypewas the
same as B356. The HD phenotyperesultedfrom pairing diploid B356 with haploid B286.
othertwo bands,consistentfor a heteromericisozyme)
tors) grew into fluffy colonies, but many of the
hyphal tip cultures taken from 2 mm subcultures
grew into flat colonies (TABLE I).
In pairing field isolate B359 with B286, 2 mm
subcultures taken from the haploid tester after 3
wk yielded mostly sectored cultures, and those
taken after 4 wk yielded mostly flat cultures (TABLE I). All subcultures from 10 mm were either
flat or sectored (TABLEI). The few subcultures
taken from further than 10 mm were all fluffy
(data not shown). About one-third of the hyphal
tip cultures taken from 2 mm subcultures were
flat (TABLEI). From the 4 wk subcultures taken
at 10 mm, only 3 of 40 hyphal tip cultures were
flat (TABLEI).
Markers for two isozymes allowed for the determination of nuclear phenotype of hyphal tip
(FIG. 2).
After pairing for 3 or 4 wk with a putatively
diploid field isolate, subcultures taken from the
haploid tester mycelia were either fluffy, flat, or
fluffy with flat sectors. In pairing field isolate B356
with haploid tester B286, all subcultures taken
from B286 at 2 mm from the confrontation zone
had a flat culture morphology (TABLEI), suggesting diploidization. Subcultures taken at 10
mm from the confrontation zone were either fluffy
or fluffy with flat sectors (TABLEI). The few subcultures taken from beyond 10 mm were all fluffy
(data not shown). All hyphal tips taken from 10
mm subcultures (even those taken from flat sec-
B286 x B356
B286 x B359
Distance is measuredfrom the confrontationzone betweenthe pairedcultures.
Hyphal tip culturesb
Numbers are the percentage of subcultures (n = 10) or hyphal tip cultures (n = 40) that were fluffy, flat, or
fluffy with flat sectors.
ACO la
~ -i
-~~ -i
- ~-~~ -I
ACO la
GPI la
GPI la
~ -i
-~~ -I
- ~-
FIG.3. Isozyme electromorphs,culture morphology, and putative nuclearstates of Armillariaostoyae
hyphaltip culturesderivedfrompairingB356 (putative
diploid) with B286 (haploid).H = haploid, D = dip-
FIG.4. Isozyme electromorphs,culture morphology, and putative nuclearstates of Armillariaostoyae
hyphaltip culturesderivedfrompairingB359 (putative
diploid) with B286 (haploid).H = haploid, D = diploid, HD = hybrid.
loid, HD = hybrid.
cultures. A diagrammatic representation of the
electrophoretic phenotypes observed in the parent strains and hyphal tip cultures is shown in
FIGS. 3 and 4. Haploid tester strain B286 had
one band for GPI and two bands for ACO (FIGS.
1-4). Because there was only one band resolved
(band lb), diploids B356 and B359 were considered to be homozygous for GPI (FIGS. 3, 4). In
contrast, both B356 and B359 were considered
to be heterozygous for ACO 1 (FIGS.3, 4). Isolate
B356 was considered to be heterozygous for
AC02 but B359 was considered to be homozygous for the AC02 locus.
Isozyme phenotypes of selected flat and fluffy
hyphal tip cultures corresponded in almost all
cases with culture morphology (TABLEII). All
fluffy hyphal tip cultures matched the isozyme
phenotype of the haploid tester (B286). Twentyfive flat, hyphal tip cultures from haploid tester
mycelia had the isozyme phenotype of the respective diploid parent, four cultures had isozyme phenotypes of both the parent and tester
(a hybrid), and three cultures had the isozyme
phenotype of the haploid tester strain (TABLE II,
FIGS. 3, 4).
Consistent with the dimeric nature of GPI, the
putative hybrid culture from pairing B356 x
B286 had three bands: one band from the homozygous diploid, one band from the haploid
tester, and a heteromeric band whose migration
was intermediate between the other two bands
(FIGS. 2, 3). Three putative hybrids of B359 x
B286 also had three bands for GPI, including a
heteromeric band (FIG. 4). In the B359 x B286
hybrid, staining for ACO resulted in four bands:
the three bands found in the diploid and band
2a of the haploid tester (FIGS. 1, 4).
Isozyme markers from diploid parent isolates
moved into the mycelium of a haploid tester
strain of A. ostoyae, supporting the hypothesis
that there is a genetic basis for the changes in
culture morphology used as a criterion for identification of Armillaria species. In those cases
where the flat, hyphal tip cultures showed only
the isozyme phenotype of the diploid parent, it
could be argued that the flat subcultures resulted
from invasive growth of diploid hyphae rather
than from diploidization of the haploid tester
mycelium. This could easily be envisioned where
the subcultures used for hyphal tipping originated from only 2 mm from the confrontation line.
However, in four of the 32 flat, hyphal tip cultures tested, isozyme phenotypes indicative of
hybrids were found, and two of these hybrids
originated from subcultures taken 10 mm from
the confrontation line. In both B356 x B286 and
B359 x B286 hybrids, heteromers of the dimeric
enzyme GPI were found, which suggests that alleles of both the haploid and diploid parents were
in the same hyphal tip and that both were being
expressed in the tested hyphal tip culture.
B286 x B356
B286 x B359
Isozyme phenotypeb
a Distance is measured from the confrontation zone between the paired cultures.
b Numbers
represent the total number of hyphal tip cultures tested. Data from 3 and 4 wk were combined.
It has been suggested that in compatible, diploid-haploid pairings of Armillaria species, the
diploid nucleus replaces the haploid nucleus in
the tester mycelium (Guillaumin et al., 1991;
Ullrich and Anderson, 1988) as has been reported for dikaryon-monokaryon (di-mon) pairings of Trametes versicolor (L.: Fr.) Pilat (Aylmore and Todd, 1984). Our data are consistent
with this hypothesis and indicate that after hyphal fusion, the diploid nucleus slowly migrates
into the haploid mycelium and coresides with
the haploid nucleus in many of the hyphal tips.
Although most subcultures taken from the haploid tester mycelium were flat in morphology,
most hyphal tip cultures from those subcultures
were fluffy and had only the isozyme markers of
the haploid tester strain. Isozyme markers of only
the diploid parent were found in most flat, hyphal tip cultures, which suggests that the diploid
nucleus eventually displaces the haploid nucleus.
The infrequent hybrid isozyme phenotype may
represent a transitional phase. Although our study
was limited in scope, culture morphology and
isozyme patterns suggest that diploidized tester
mycelia are, at least initially, mosaics of haploid,
diploid, and hybrid hyphal tips of both the diploid and haploid parent nuclei.
An alternative hypothesis for diploidization is
that the diploid nucleus undergoes reduction
(haploidization) prior to hyphal fusion or in the
haploid tester mycelium, and one of the resulting
haploid nuclei fuses with the haploid nucleus of
the tester to form a new, recombinant diploid.
Apparent haploidizations of diploid nuclei have
been observed in basidial primordia of Armillaria, which give rise to the dikaryotic hyphae in
the gills, and transient dikaryotic stages in the
life cycle are known (Korhonen and Hintikka,
1974; Korhonen, 1980, 1983; Larsen et al., 1992;
Tommerup and Broadbent, 1975). Guillaumin
(1986) found limited evidence (based on matingtype alleles) of recombinant diploids following
diploid-haploid pairings ofA. ostoyae. However,
vegetative diploids of Armillaria are believed to
be very stable (Anderson and Ullrich, 1982), and
our data further suggest their stability. None of
the 29 flat, hyphal tip cultures had isozyme patterns consistent with a recombination between
the diploid parent and the haploid tester. However, the use of additional nuclear markers (isozyme or molecular) and the testing of more isolates could allow for detection of potential low
frequency recombination events.
Korhonen (1983) observed nuclear migration
through eroded septa in haploid-haploid pairings
of Armillaria. He found that nuclear migration
was very slow (approximately 1.1 mm per day)
compared with migration rates in other hymenomycetes of 3-4 mm per hour (Aylmore and Todd,
1984; Korhonen, 1983). Because of slow nuclear
migration, a long period of time may be required
for the entire mycelium of an Armillaria subculture to become converted to a diploid. It appears, however, that only a portion of a mycelium needs to be diploidized to induce a change
in culture morphology from fluffy to flat.
The reliability of identifications using diploidhaploid pairings is greatly improved if only fresh
tester isolates with exceptionally fluffy mycelia
are used (Korhonen, 1978) and subcultures are
taken from haploid tester mycelium after 4-6 wk
(Harrington et al., 1992). Small plugs of mycelium are excised from the haploid tester at 2 mm
behind the confrontation zone, from the ad-
vancing margin of the tester at the site opposite
the confrontation, and from a point between the
other two samples. These plugs are transferred
to MEA and incubated for 7 to 10 days. Subcultures of the tester with a morphology different
from the unpaired tester indicate that the tester
has been diploidized and that the unidentified
isolate and the tester are conspecific.
Siepmann (1987) found that in many diploidhaploid pairings between different species, the
haploid tester would eventually (after 2 or 3
months) overgrow the diploid mycelium. If the
diploid was of the same species, the haploid tester would change in morphology, but mycelia of
the haploid tester would remain restricted to the
line where the two mycelia met. This technique
may prove useful for identification of field isolates of Armillaria species, but the genetic basis
of the reaction is not known. Similarly, putatively diploid isolates may be identified to species by pairing with other diploid isolates on
MEA. A black line usually develops between diploid isolates if they are different species, and this
can be used to confirm results based on diploidhaploid pairings (Korhonen, 1978; Ullrich and
Anderson, 1978). The black line is more easily
seen if L-DOPA is added to the confrontation
zone (Hopkin et al., 1989). Another alternative
is the induction of haploidization of the diploid
isolates using benomyl (at 25 ppm) and then using the induced haploid strains in haploid-haploid pairings (Anderson and Yacoob, 1984). We
have found, however, that the diploid-haploid
pairing technique with subculturing is the most
reliable method for identification of diploid isolates of North American Armillaria species, and
now a genetic basis for this test has been estab-
ological species of Armillaria mellea. Mycologia
79: 69-76.
, and E. Stasovski. 1992. Molecular phylogeny
of Northern Hemisphere species of Armillaria.
Mycologia 84: 505-516.
, and R. C. Ullrich. 1982. Diploids of Armil-
laria mellea: synthesis, stability, and mating behavior. Canad. J. Bot. 60: 432-439.
, and R. Yacoob. 1984. Benomyl-induced somatic segregation in diploid Armillaria mellea.
Phytopathology 74: 612-615.
Aylmore, R. C., and N. K. Todd. 1984. Hyphal fusion
in Coriolus versicolor. Pp. 103-125. In: The ecology and physiology of the fungal mycelium. Eds.,
D. H. Jenningsand A. D. M. Rayner.Cambridge
University Press, Cambridge,United Kingdom.
Berube,J. A., and M. Dessureault. 1988. Morpho-
logical characterization of Armillaria ostoyae and
Armillaria sinapina sp. nov. Canad. J. Bot. 66:
. 1989. Morphological studies of
, and
the Armillaria mellea complex: two new species,
A. gemina and A. calvescens. Mycologia 81: 216225.
Burdsall, H. H., M. Banik, and M. E. Cook. 1990.
Serological differentiation of three species of Armillaria and Lentinula edodes by enzyme-linked
immunosorbent assay using immunized chickens
as a source of antibodies. Mycologia 82: 415-423.
Conkle, M. T., P. D. Hodgekiss, L. B. Nunnally, and
S. C. Hunter. 1982. Starch gel electrophoresis of
conifer seeds: a laboratory manual. USDA Forest
Serv. General Technical Report PSW-64. 18 pp.
Guillaumin, J. J. 1986. Contribution a l'etude des
Armillaires phytopathogenes, en particulier du
groupe Mellea: cycle caryologique, notion d'espece, role biologique des especes. Th6se, Univ.
Claude Bernard-Lyon I, France. 270 pp.
, J. B. Anderson, and K. Korhonen. 1991. Life
cycle, infertility, and biological species. Pp. 1020. In: Armillaria root disease. Eds., C. G. Shaw
and G. A. Kile. USDA Forest Serv. Agric. Handbook No. 691.
, and S. Berthelay. 1981. Determination specifique des armillaires par la methode des groupes
de compatibilit6 sexuelle. Specialisation ecololished for A. ostoyae.
gique des esp6ces francaises. Agronomie 1: 897908.
, B. Lung, H. Romagnesi, H. Marxmiiller, D.
Lamoure, G. Durrieu, S. Berthelay, and C. MoThe authorsthankRobertEckertof the Department
hammed. 1985. Systematique des Armillaires du
of ForestResources,Universityof New Hampshirefor
groupe Mellea. Consequences phytopatholothe use of laboratoryfacilities for electrophoresisand
giques. Eur. J. Forest Pathol. 15: 268-277.
isozymeanalysis.CharlotteR. Bronson,JamesV. Groth
, C. Mohammed, and S. Berthelay. 1989. Arand GeorgianaMay offereduseful suggestionsfor the
millaria species in the northern temperate hemimanuscript.This material is based upon work supsphere. Pp. 27-43. In: Proceedings 7th IUFRO
ported by the U.S. Departmentof Agriculture,CSRS
conference on root and butt rots. Ed., D. J. MorunderAgreementNo. 86-FSTY-9-0186.JournalPaper
rison. Forestry Canada, Victoria, British Colum-
No. J-14938 of the Iowa Agricultureand Home Economics ExperimentStation, Ames, Iowa ProjectNo.
Anderson, J. B., D. M. Petsche, and M. L. Smith.
1987. Restriction fragment polymorphisms in bi-
bia, Canada.
Harrington, T. C. 1987. Compatibility of New
Hampshire spruce and fir isolates of Armillaria
with A. obscura. Phytopathology 77: 117. (Abstract)
, J. J. Worrall, and F. A. Baker. 1992. Armillaria. Pp. 81-85. In: Methods for research on
soilborne phytopathogenic fungi. Eds., L. L. Sin-
gleton, J. D. Mihail, and C. Rush. American Phytopathological Society Press, St. Paul, Minnesota.
Hintikka, V. 1973. A note on the polarity of Armillariella mellea. Karstenia 13: 32-39.
Hopkin, A. A., K. I. Mallett, and P. V. Blenis. 1989.
The use of L-DOPA to enhance visualization of
the "black line" between species in the Armillaria
mellea complex. Canad. J. Bot. 67: 15-17.
Jahnke, K.-D., G. Bahnweg, and J. J. Worrall. 1987.
Species delimitation in the Armillaria mellea complex by analysis of nuclear and mitochondrial
DNAs. Trans. Brit. Mycol. Soc. 88: 572-575.
Kile, G. A. 1983. Identification of genotypes and the
clonal development of Armillaria luteobubalina
Watling and Kile in eucalypt forests. Austral. J.
Bot. 31: 657-671.
Korhonen, K. 1978. Infertility and clonal size in the
Armillariella complex. Karstenia 18: 31-42.
. 1980. The origin of clamped and clampless
basidia in Armillariella ostoyae. Karstenia 20: 2327.
. 1983. Observations on nuclear migration and
heterokaryotization in Armillaria. Cryptog. Mycol. 4: 79-85.
, and V. Hintikka. 1974. Cytological evidence
for somatic diploidization in dikaryotic cells of
Armillariella mellea. Arch. Microbiol. 95: 187-192.
Larsen, M. J., M. T. Banik, and H. H. Burdsall, Jr.
1992. Clamp connections in North American Armillaria species: occurrence and potential application for delimiting species. Mycologia 84: 214218.
Linde, D. C., J. V. Groth, and A. P. Roelfs. 1990.
Genotypic frequencies of phosphoglucose isomerase allozymes in bean rust (Uromyces appendiculatus) populations estimated by densitometry. Pl.
Pathol. 39: 102-109.
Mallet, K. I. 1990. Host range and geographic distribution of Armillaria root rot pathogens in the
Canadian prairie provinces. Canad. J. Forest Res.
20: 1859-1863.
Marty, T. L., D. M. O'Malley, and R. P. Guries. 1984.
A manual for starch gel electrophoresis: a new
microwave edition. Staff Paper No. 20. Dept. of
Forestry, Univ. of Wisconsin, Madison. 20 pp.
Micales, J. A., M. R. Bonde, and G. L. Peterson. 1986.
The use of isozyme analysis in fungal taxonomy
and genetics. Mycotaxon 27: 405-449.
Morrison, D. J., A. J. Thomson, D. Chu, F. G. Peet,
T. S. Sahota, and U. Rink. 1985. Isozyme patterns of Armillaria intersterility groups occurring
in British Columbia. Canad. J. Microbiol. 31:651653.
Pasteur, N., G. Pasteur, F. Bonhomme, J. Catalan, and
J. Britton. 1988. Practical isozyme genetics. Davidson Halsted Press, New York. 215 pp.
Peabody, D. C., J. J. Motta, and C. D. Therrien. 1978.
Cytophotometric evidence for heteroploidy in the
life cycle ofArmillaria mellea. Mycologia 78: 487498.
Rishbeth, J. 1982. Species of Armillaria in southern
England. PI. Pathol. 31: 9-17.
. 1986. Some characteristics of English Armillaria species in culture. Trans. Brit. Mycol. Soc.
85: 213-218.
Roll-Hansen, F. 1985. The Armillaria species in Europe. Eur. J. Forest Pathol. 15: 22-31.
Shaw, C. G., III, and E. M. Loopstra. 1988. Identification and pathogenicity of some Alaskan isolates of Armillaria. Phytopathology 78: 971-974.
Siepmann, V. R. 1987. Kriterien zur Beurteilung der
Reaktion haploider Tester mit diploiden Armil/aria-Isolierungen. Eur. J. Forest Pathol. 17: 308311.
Smith, M. L., and J. B. Anderson. 1989. Restriction
fragment length polymorphisms in mitochondrial
DNAs of Armillaria: identification of North
American biological species. Mycol. Res. 93: 247256.
Tommerup, I. C., and D. Broadbent. 1975. Nuclear
fusion, meiosis and the origin of dikaryotic hyphae
in Armillariella mellea. Arch. Microbiol. 103: 279282.
Ullrich, R. C., and J. B. Anderson. 1978. Sex and
diploidy in Armillaria mellea. Exp. Mycol. 2:119129.
. 1988. Armillaria mellea, cause
, and
of rots in woody species. Advances PI. Pathol. 6:
Wargo, P. M., and C. G. Shaw, III. 1985. Armillaria
root rot: the puzzle is being solved. PI. Dis. 69:
Accepted for publication July 31, 1992