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Canadian Journal of Plant Pathology
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Genetic structure and variation in aggressiveness in
Fusarium virguliforme in the Midwest United States
a
a
b
a
a
G. Y. C. Mbofung , T. C. Harrington , J. T. Steimel , S. S. Navi , X. B. Yang & L. F.
Leandro
a
a
Department of Plant Pathology and Microbiology, 351 Bessey Hall, Iowa State University,
Ames, IA, 50011, USA
b
Pioneer Hi-Bred International Inc., 7300 N.W. 62nd Ave. P.O. Box 1004, Johnston, IA,
50131-1004, USA
Available online: 14 Feb 2012
To cite this article: G. Y. C. Mbofung, T. C. Harrington, J. T. Steimel, S. S. Navi, X. B. Yang & L. F. Leandro (2012): Genetic
structure and variation in aggressiveness in Fusarium virguliforme in the Midwest United States, Canadian Journal of Plant
Pathology, 34:1, 83-97
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Can. J. Plant Pathol. (2012), 34(1): 83–97
Genetics and resistance/Génétique et résistance
Downloaded by [Iowa State University], [Tom Harrington] at 08:44 30 April 2012
Genetic structure and variation in aggressiveness in Fusarium
virguliforme in the Midwest United States
G. Y. C. MBOFUNG1 , T. C. HARRINGTON1 , J. T. STEIMEL2 , S. S. NAVI1 , X. B. YANG1 AND L. F. LEANDRO1
1
2
Department of Plant Pathology and Microbiology, 351 Bessey Hall, Iowa State University, Ames, IA 50011, USA
Pioneer Hi-Bred International Inc., 7300 N.W. 62nd Ave. P.O. Box 1004, Johnston, IA 50131-1004, USA
(Accepted 2 February 2012)
Abstract: Sudden death syndrome of soybean, caused by Fusarium virguliforme, is one of the most damaging diseases affecting soybean
production in the Midwestern region of the USA. To date, there has been very low genetic variation detected in F. virguliforme populations,
although isolates can exhibit differing aggressiveness on soybean. To further investigate the genetics behind this variation in aggressiveness,
multiple fingerprint analyses were conducted on a collection of F. virguliforme isolates from Iowa and neighbouring states, and their
aggressiveness was tested using greenhouse and culture filtrate assays. Twelve RAPD primers identified 13 genotypes, while hybridization of
the (CAT)5 probe to Pst I-restricted genomic DNA yielded eight genotypes. Fingerprint patterns obtained by RFLP analysis of mitochondrial
DNA with the enzyme HaeIII were identical for all F. virguliforme isolates. Combined analysis of the RAPD and (CAT)5 binary data
identified 25 genotypes within F. virguliforme, indicating a greater amount of genetic diversity than was previously known. Disease severity
and plant growth varied significantly among isolates, but isolate aggressiveness was not associated with genetic variation. This study provides
evidence for the existence of genetic variation in F. virguliforme but suggests that minor quantitative traits and environmental interactions are
primarily responsible for the variation in aggressiveness found among isolates within the species.
Keywords: genetic variation, quantitative traits, soybean, sudden death syndrome
Résumé: Le syndrome de la mort subite du soja, causé par Fusarium virguliforme, est une des maladies du soja qui provoque le plus de
dommages dans le Midwest américain. À ce jour, très peu de variations génétiques ont été détectées dans les populations de F. virguliforme,
bien que des isolats puissent afficher des degrés variables de virulence à l’égard du soja. Pour étudier la génétique à l’origine de cette
variation, de multiples analyses de l’empreinte génétique ont été faites sur une collection d’isolats de F. virguliforme provenant de l’Iowa et
des États voisins. Leur virulence a été testée au moyen de filtrats de serre et de culture. Douze sondes RAPD ont identifié 13 génotypes tandis
que l’hybridation entre la sonde (CAT)5 et l’ADN génomique restreint par Pst-I en a produit huit. Les arrangements d’empreintes obtenus par
analyse RFLP de l’ADN mitochondrial avec l’enzyme HaeIII étaient identiques pour tous les isolats de F. virguliforme. L’analyse combinée
des données binaires de la RAPD et de la sonde (CAT)5 a permis d’identifier 25 génotypes chez F. virguliforme, ce qui représente une plus
grande diversité génétique que celle connue à ce jour. La gravité de la maladie et la croissance des plantes variaient significativement selon les
isolats, mais la virulence des isolats n’était pas toujours associée à la variation génétique. Cette étude démontre l’existence de la variation
génétique chez F. virguliforme, mais suggère que des traits quantitatifs mineurs et des interactions environnementales sont principalement
responsables de la variation sur le plan de la virulence chez les isolats de l’espèce.
Mots clés: soja, syndrome de la mort subite, traits quantitatifs, variation génétique
Correspondence to: L. Leandro. E-mail: lleandro@iastate.edu
ISSN: 0706-0661 print/ISSN 1715-2992 online © 2012 The Canadian Phytopathological Society
http://dx.doi.org/10.1080/07060661.2012.664564
G. Y. C. Mbofung et al.
Downloaded by [Iowa State University], [Tom Harrington] at 08:44 30 April 2012
Introduction
Sudden death syndrome (SDS) of soybean (Glycine
max L.) is an economically important disease that causes
major yield losses in North and South America (Roy et al.,
1997). The disease is characterized by root rot, foliar
interveinal chlorosis and necrosis, and premature defoliation (Roy et al., 1997; Aoki et al., 2005). In the USA,
the disease was first detected in Arkansas in 1971 (Hirrel,
1983), later spreading northwards along the Mississippi
river valley and being reported in Indiana and Illinois in
1986 (Rupe et al., 1989), Iowa in 1993 (Yang & Lundeen,
1997), Minnesota in 2002 (Kurle et al., 2003), Wisconsin
in 2007 (Bernstein et al., 2007), and Michigan in 2010
(Chilvers et al., 2010). SDS is currently found in almost
all soybean-producing regions in the USA (Roy et al.,
1997). Yield losses typically range between 5 and 15%,
with up to 80% loss in individual fields (Roy et al.,
1997). Annual losses of 0.3 to 2.0 million tons were estimated for the period between 1997 and 2007 (Wrather &
Koenning, 2009). Worldwide, SDS disease has also been
reported in Canada (Anderson & Tenuta, 1998) and parts
of South America, including Argentina (Ploper, 1993),
Brazil (Yorinori et al., 1993), Bolivia, Paraguay (Yorinori,
1999) and Uruguay (Ploper et al., 2003).
The soilborne fungus Fusarium virguliforme
O’Donnell & T. Aoki (formerly F. solani (Mart.)
Sacc. f. sp. glycines) is the only causal agent of SDS in
North America, whereas four Fusarium species, namely
F. virguliforme, F. tucumaniae T. Aoki, O’Donnell, Yos.
Homma & Lattanzi, F. brasiliense T. Aoki & O’Donnell
and F. crassistipitatum T. Aoki, M. M. Scandiani &
O’Donnell cause the disease in South America (Aoki
et al., 2005; O’Donnell et al., 2010; Aoki et al., 2011).
Various molecular techniques have been used to decipher
the species-limits and infer the evolutionary relationships
among SDS-causing isolates and other members of the
F. solani species complex. These studies have included
sequence analyses of the mitochondrial small subunit
rDNA (Li et al., 2000) and nuclear rDNA (O’Donnell
& Gray, 1995), as well as restriction fragment length
polymorphism (RFLP) analysis of the mitochondrial
genome and internal transcribed spacer of the rDNA
(Rupe et al., 2001; Arruda et al., 2005; Malvick &
Bussey, 2008). Results from these studies showed a
nearly homogeneous population of F. virguliforme exists
in North America. A multiloci genetic assay based on
nucleotide polymorphism within the nuclear ribosomal
intergenic spacer rDNA and two anonymous intergenic
regions differentiated the soybean SDS pathogen from
two closely related pathogens, F. cuneirostrum and F.
phaseoli, which cause root rot on Phaseolus and mung
beans (O’Donnell et al., 2010).
84
Despite the observed genetic homogeneity (Achenbach
& Patrick, 1996; Achenbach et al., 1997; Aoki et al.,
2003), many studies have shown considerable variation
in aggressiveness among F. virguliforme isolates in both
greenhouse and field studies (Roy et al., 1989; Malvick
& Bussey, 2008). The existence of races of F. virguliforme has also been suggested based on differences in
aggressiveness (Lim & Jin, 1991), although there are no
established soybean differentials for race determination.
While SDS development and severity is highly dependent
on environmental conditions (Scherm et al., 1998), this
variation in aggressiveness is surprising given the clonal
reproductive nature of the pathogen (Scherm & Yang,
1996). We hypothesized that differences in aggressiveness
found among F. virguliforme isolates may be caused by
minor genetic variation within the pathogen population,
namely, many quantitative genetic traits.
Some evidence exists for limited genetic variability
in F. virguliforme. In one study, random amplified polymorphic DNA (RAPD) analysis of 55 F. virguliforme
isolates from nine states in the USA showed the existence of two groups, which the authors called amplitypes
(Achenbach & Patrick, 1996). Chromosome length polymorphisms and two distinct karyotypic patterns have also
been reported from preliminary studies with F. virguliforme isolates (Mansouri, 2009). There is a need to further
investigate the genetic variation within the F. virguliforme
genome and determine if it is associated with variations in
isolate aggressiveness. In a recent study, isolates showed
a range of aggressiveness levels, but no genetic variation
was found through sequence analysis of the ITS, IGS and
EF-1 alpha regions (Malvick & Bussey, 2008). DNA fingerprint analyses using RAPDs and microsatellite probes
provide better discrimination among genotypes due to the
large number of characterized fragments from the genome
(Taylor et al., 1999). These methods have been used to
detect previously unknown genetic variations in fungal
pathogens such as Ceratocystis cacaofunesta Engelbrecht
& Harrington, Eutypa lata (Pers : Fr.) Tul. & C. Tul, and
Cadophora gregata Harrington & McNew (Harrington
et al., 2003; Péros & Berger, 2003; Engelbrecht et al.,
2007). The objective of this study was to employ several molecular markers to characterize the intraspecific
genetic variation in 72 F. virguliforme isolates from Iowa
and to test whether genetic variation was associated with
variation in aggressiveness among isolates.
Materials and methods
Fungal isolates
A total of 72 isolates of F. virguliforme collected from
diseased soybean roots (Table 1) were used in this study.
Genetic structure and aggressiveness of Fusarium virguliforme
85
Table 1. Origin, year of collection, genotype classes obtained from analysis of RAPD and (CAT)5 markers, and genetic subgroups of
Fusarium virguliforme isolates used in this study.
a Isolate
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Strain
LL0039
LL0008
LL0041
LL0029
LL0014
LL0038
Mn.Fsg.Br3-ss1
Mn.Fsg.Re1-ss1
Fsg-ISU22
Fsg-ISU22B5
Fsg-ISU5B
LL0130
LL0001
LL0002
LL0003
LL0004
LL0005
LL0006
LL0007
LL0009
LL0010
LL0011
LL0015
LL0020
Fsg-ISU1
Fsg-ISU2A
Fsg-ISU5E
Fsg-ISU9-1C
Fsg-ISU12
Fsg-ISU16
LL0030
LL0031
LL0032
LL0033
LL0037
LL0095
LL0096
LL0106
Fsg-ISU15-2B
Fsg-ISU22B2
Fsg-ISU23
Fsg-ISU24
Fsg-ISU15-2B
Mn.Fsg.Be1-ss1
Mn.Fsg.Fr1-ss1
Mn.Fsg.Ho1-ss1
Mn.Fsg.St1-ss1
Mn.Fsg.Ww1-ss1
Mn.Fsg.St2-ss2
Mn.Fsg.Mo1-ss1
Mn.Fsg.Ls2-ss1
Fsg-ISU11
Fsg-ISU10
Fsg-ISU15-5
LL0131
number
1
2
3
4
8
5
6
7
9
10
11
12
13
14
15
16
17
18
b County/State
Buchanan, Iowa
Story, Iowa
Johnson, Iowa
Boone, Iowa
Story, Iowa
Buchanan, Iowa
Brown, Minnesota
Redwood, Minnesota
Scott, Iowa
Scott, Iowa
Chickasaw, Iowa
Fayette, Iowa
Story, Iowa
Story, Iowa
Story, Iowa
Story, Iowa
Story, Iowa
Story, Iowa
Story, Iowa
Story, Iowa
Story, Iowa
Story, Iowa
Chickasaw, Iowa
Washington, Iowa
Boone, Iowa
Cerro Gordo, Iowa
Chickasaw, Iowa
Floyd, Iowa
Greene, Iowa
Johnson, Iowa
Boone, Iowa
Boone, Iowa
Boone, Iowa
Dallas, Iowa
Buchanan, Iowa
Story, Iowa
Story, Iowa
Boone, Iowa
Jasper, Iowa
Scott, Iowa
Worth, Iowa
Worth, Iowa
Jasper, Iowa
Blue Earth, Minnesota
Freeborn, Minnesota
Houston, Minnesota
Steele, Minnesota
Watonwan, Minnesota
Steele, Minnesota
Mower, Minnesota
LeSueur, Minnesota
Greene, Iowa
Floyd, Iowa
Jasper, Iowa
Fayette, Iowa
Year
(CAT)5
genotype
2006
2006
2006
2006
2006
2006
2006
2006
1995
1995
2000
2007
2006
2006
2007
2006
2006
2006
2006
2006
2006
2006
2006
2006
1993
2000
2000
1995
1996
1994
2006
2006
2006
2006
2006
2006
2006
2006
1994
1995
1995
1995
1994
2006
2006
2006
2006
2006
2006
2006
2006
1994
1994
1994
2007
C5
C4
C4
C3
C1
C1
C1
C1
C6
C6
C8
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C5
C7
C7
C5
C1
C4
C5
RAPD
genotype
R6
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R2
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R1
R3
R1
R5
R11
Combined
genotype
Subgroup
I
II
II
III
IV
IV
IV
IV
V
V
VI
VII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
VIII
IX
IX
X
XI
XII
XIII
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
I
1
1
1
I
1
1
1
1
1
1
1
1
1
1
1
2
(Continued)
G. Y. C. Mbofung et al.
86
Table 1. (Continued.)
a Isolate
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Strain
IlMont-1 (A)
Mont 1
LL0105
LL0019
LL0028
Rupe
LL0036
Fsg-ISU13
Fsg-ISU19
Fsg-ISU17
Fsg-ISU22G2
Fsg-ISU6
Fsg-ISU22B1
Fsg-ISU6A
Fsg-ISU6-1C
number
19
20
21
22
23
24
25
26
27
28
29
30
b County/State
Piatt, Illinois
Monticello, Illinois
Fayette, Iowa
Washington, Iowa
Boone, Iowa
Arkansas
Buchanan, Iowa
Henry, Iowa
Monticello, Illinois
Iowa
Scott, Iowa
Clinton, Iowa
Scott, Iowa
Clinton, Iowa
Clinton, Iowa
Year
(CAT)5
genotype
RAPD
genotype
Combined
genotype
Subgroup
1991
1991
2007
2006
2006
unknown
2006
1994
1991
unknown
1995
1993
1995
1993
1993
C1
C1
C2
C1
C5
C4
C4
C7
C1
C5
C6
C6
C5
C1
C1
R13
R13
R1
R12
R9
R9
R10
R3
R11
R7
R8
R7
R8
R13
R8
XIV
XIV
XV
XVI
XVII
XVIII
XIX
XX
XXI
XXII
XXIII
XXIII
XXIV
XXIV
XXV
2
2
2
3
3
3
3
3
3
4
4
4
4
4
4
a Numbered
isolates were used in aggressiveness assays.
isolates and the ILMont-1(A) isolate were obtained from Dean Malvick, University of Minnesota; isolates from Arkansas, and Illinois were
obtained from X. B. Yang’s collection.
b Minnesota
Fifty-eight of these isolates were collected from 16 different counties in Iowa. Thirty-three of the 58 isolates were
collected from fields surveyed in 2006–2007 and are represented by the LL code, while the remaining 25 isolates
were obtained from a state-wide survey that was conducted in 1994 and 1995 (Yang & Lundeen, 1997) and
are represented by the Fsg-ISU code. During the survey of
2006–2007, F. virguliforme was isolated from roots using
a modified Nash and Snyder medium (MNSM) (Rupe
et al., 1997) and transferred onto potato dextrose agar
(PDA) for identification based on published morphological characteristics (Aoki et al., 2003). All the isolates were
single-spored, and identification was further confirmed by
PCR using primers Fsg 1 and Fsg 2 designed from the
mitochondrial small subunit rDNA and by sequence analysis of the EF-1α gene (Li et al., 2000; Aoki et al., 2003).
Also included in this study were 10 isolates of F. virguliforme from Minnesota, three isolates from Illinois and one
isolate from Arkansas, which were similarly single-spored
and characterized by DNA sequencing (Table 1).
DNA extraction
Isolates were grown in malt yeast extract liquid medium
(3 g malt extract, 3 g yeast extract, 5 g peptone, 20 g
dextrose, 2 g ammonium sulphate in 1 L of water)
(O’Donnell, 1992) at room temperature (20–35 ◦ C) for
4 days on a rotary shaker (115 rpm). Preliminary experiments showed that this medium minimized the production of polysaccharides by the fungus during growth and
yielded the best quality of DNA for all analyses. Mycelial
mats were collected by vacuum filtration on Whatman
#54 filter paper (Whatman International Ltd, Maidstone,
England) and freeze-dried overnight. Total genomic DNA
was extracted using the method described by DeScenzo
and Harrington (1994) and quantified using a TKO100 minifluorometer (Hoefer Scientific Instruments, San
Francisco, CA).
DNA fingerprint analysis
Four molecular techniques were used to generate fingerprints for analysis of genetic variation: mitochondrial
restriction fragment length polymorphisms (RFLP),
minisatellite- and microsatellite-primed PCR, nuclear
DNA analysis using microsatellite probes (CAT)5 and
(CAC)5 , and random amplification of polymorphic DNA
(RAPD). Each of the four fingerprinting methods was
repeated at least once for each isolate, and only bands
that were consistently visible and can be unambiguously
scored in all replicate gels were analyzed.
Mitochondrial DNA RFLP. For generating the
mitochondrial RFLP patterns, total genomic DNA
was digested with the restriction enzymes HaeIII, CfoI
and MspI (Promega, Madison, WI), which recognize the
restriction sites GGCC, GCGC and CCGG, respectively
(Wingfield et al., 1996). For each isolate, 30 µg of
DNA were restricted overnight at 37 ◦ C, with 5 U
of each enzyme per µg of DNA in a 500 µL total
reaction volume. The digested DNA was concentrated
by precipitation with 2 volumes of ethanol and 0.2 M
Genetic structure and aggressiveness of Fusarium virguliforme
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NaCl. The pellet obtained was washed in 70% ethanol
and dried in a speed vacuum for 45 min. The DNA pellet
was then dissolved in TE buffer (10 mM Tris HCL, 1 mM
EDTA) to a concentration of 0.67 µg µL−1 , and 15 µL
of the restricted product was used in electrophoresis. The
restricted samples were separated on a 1.2 % agarose gel
at 100 V for 16 h in 1.25× TBE and visualized under UV
light after staining with ethidium bromide.
Minisatellite and microsatellite analyses. Four DNA fingerprint markers were used to generate binary data for
analysis. The minisatellite primers M13 and T3B, and
the microsatellite or intersimple sequence repeat (ISSR)
primers (GACA)4 and (GTG)5 were used in 50 µL PCR
amplification reactions (McDonald, 1997). PCR amplifications were performed in a 50 µL volume consisting
of 2.5 mM MgCl2 , 0.2 mM deoxyribonucleotide triphosphates, 0.5 µM of each primer, 1× PCR buffer, 0.05 U
of Biolase Taq DNA polymerase (Bioline USA Inc.
Randolph, MA), and 25 ng of template DNA. The reaction conditions were an initial denaturation at 94 ◦ C for
5 min, 40 cycles of 95 ◦ C for 1 min, 55 ◦ C for 90 s and
72 ◦ C for 2 min. The final extension was set at 72 ◦ C
for 5 min. At least two PCR reactions were performed
for each isolate–primer combination to assess the consistency of amplification. Twenty µL of each PCR product
were mixed with 5 µL of bromophenol blue and separated in 1% agarose gels. Electrophoresis was carried out
in 0.5× TBE (Tris-Borate EDTA) buffer run for 6 h at
80 V. The gels were visualized in UV light after staining
with ethidium bromide.
Nuclear DNA analysis using microsatellite probes (CAT)5
and (CAC)5 . The oligonucleotides (CAT)5 and (CAC)5
were end-labelled with 32 P-dCTP using polymerase terminal deoxynucleotidyl transferase (Invitrogen, Carlsbad,
CA) and purified through a Sephadex G-25 column. Total
genomic DNA obtained as described above was digested
with the restriction enzyme PstI following the procedures described by DeScenzo and Harrington (1994). The
digested products were resolved on a 1.2% agarose gel
(Bio-Rad laboratories, Hercules, CA) in 1.25 × TBE. The
gels were run in a system provided with buffer recirculation at 100 V for 16 h. Once the gels were examined
under UV light, they were dried with a gel drier for
45 min at room temperature, sealed in a plastic wrap and
stored at 4 ◦ C for further use. Hybridization and washing conditions were as previously described (DeScenzo
& Harrington, 1994). After hybridization, the gels were
exposed to an imaging screen (Molecular imager Fx,
Biorad, Hercules, CA) for 2 days and scanned with a
Pharos Fx Plus molecular imager (Biorad, Hercules, CA).
87
Each isolate was fingerprinted in at least two gel runs, and
only the bands present in both runs were scored.
RAPD analysis. Nineteen random amplified polymorphic
DNA (RAPD) primers (Achenbach & Patrick, 1996;
Achenbach et al., 1997) were individually screened for the
generation of polymorphic loci. Twelve of these primers,
namely OPA-01, OPA-02, OPA-04, OPA-13, OPA-19,
OPB-05, OPF-01, OPF-13, OPL-12, OPN-13, PM02 and
PM04 were selected for further analysis on the basis of
reproducibility and the number of polymorphic loci generated. Each PCR reaction mix was set up as described
above, except that 100 ng of DNA template and only one
primer was used per reaction. PCR reactions were run
for 40 cycles with the following steps: 94 ◦ C for 3 min,
40 cycles of 94 ◦ C for 1 min, 36 ◦ C for 1 min and 72 ◦ C for
2 min. The amplified products were run on 1.2% agarose
gel in 1.25% TBE buffer for 6 h at 80 V and visualized in
UV after staining with ethidium bromide.
Assessment of isolate aggressiveness
Thirty isolates of F. virguliforme were chosen from
among the genotypic groups identified with the fingerprinting analysis and tested for aggressiveness on the
soybean cultivars ‘Spencer’ and ‘92M91’ using soil inoculation and culture filtrate assays. ‘Spencer’ belongs to
maturity group IV and is susceptible to F. virguliforme
(Wilcox et al., 1989), while ‘92M91’ (Pioneer Hi-Bred,
USA) belongs to maturity group II and is moderately
resistant to F. virguliforme (Schmidt et al., 2008, 2009).
Greenhouse assays. Inoculum was prepared following the
procedure described in Munkvold and O’Mara (2002).
A mixture of sand (1900 mL), cornmeal (380 mL) and
water (110 mL) was autoclaved in 20 cm × 30 cm plastic
autoclave bags (Fisher Scientific, Pittsburgh, PA) for 1 h at
121 ◦ C on two consecutive days. Each bag then received
2 mL of a 1 × 106 spores mL−1 spore suspension. The
bags were incubated in the dark at room temperature
(24 ± 2 ◦ C ) for 10 days, and the inoculum was thoroughly mixed on a daily basis. The inoculum of each
isolate was mixed with sterile sand : soil (1 : 1) at a ratio of
1 part inoculum to 5 parts soil mix. Sterile sand : cornmeal
mix was added at the same ratio for non-inoculated controls. Sterilized 21-cm tall and 4-cm diameter PVC cones
were filled with the inoculum : soil mix and a single seed,
pre-germinated for 3 days, was planted in each cone.
The plants were maintained on a greenhouse bench at
26 ± 4 ◦ C and 24 ± 4 ◦ C with a 14-h photoperiod in
June and September, respectively. The relative humidity of the greenhouse during this period varied between
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G. Y. C. Mbofung et al.
20% in the evening and 70% in morning. The experiment was set up in a completely randomized factorial
design, with 10 replicate plants per isolate by cultivar
combination, and was conducted twice (trial 1 in June
and trial 2 in September, 2009). Foliar disease severity defined as the per cent of total leaf area per plant
showing typical SDS chlorosis or necrosis, was rated
10 days after planting and at 5-day intervals thereafter
by visual assessments. Plants were destructively sampled
30 days after planting and visually assessed for root rot
severity (the per cent root area showing brown to black
discolouration), shoot fresh weight, shoot height and root
dry weight. F. virguliforme density in the soil was quantified before planting (trial 2 only) and at the end of the
experiment (both trials) on three randomly chosen conetainers per isolate using quantitative real-time PCR (Li
et al., 2008).
Culture filtrate assays. Cell-free culture filtrates of F. virguliforme were prepared as described by Li et al. (1999).
Seeds of the soybean cultivars ‘Spencer’ and ‘91M92’
were grown in a growth chamber set to 16 h light at
23 ◦ C and 8 h dark at 16 ◦ C . Fifteen-day-old soybean
seedlings were uprooted, and the roots were washed thoroughly in running tap water to remove soil, disinfested in
10% NaOCl for 1 min, then rinsed in sterile water and
blotted dry. Single seedlings were placed in 50 mL falcon tubes containing 40 mL of the culture filtrate diluted
with sterile water at a ratio of 1 : 5 for the first trial and
1 : 25 for the second trial of the experiment. Control plants
were immersed in sterile culture media and water. The
experiment was established as a completely randomized
design with 10 replicate plants per isolate in trial 1 and
six replicate plants per isolate in trial 2. The plants were
rated visually for severity of SDS foliar symptoms 5 days
after immersion in the culture filtrates and every 4 days
thereafter, until 30 days after immersion. Trials 1 and 2 of
the experiment were conducted in May and June, 2010,
respectively.
88
correlation (r) analysis using the NTSYSpc2.0 program (Exeter Software, Setauket, NY). Bootstrap values (1000 replicates) for each branch were calculated
using the Phylogenetic Analysis Using Parsimony package (PAUP, ver. 4.0b10; Sinauer Associates). Congruency
between the (CAT)5 and RAPD datasets was tested using
partition-homogeneity (P-H) test implemented in PAUP.
Additional tests (Kishino–Hasegawa: KH, Templeton
test: TT, and the Winning-sites test: WS) were performed to assess whether the conflicting dendrogram
topologies were significantly different from each other
before combining the two datasets for further analysis. Population structure and differentiation were estimated using total genetic variation partitioned among
geographic origin. The isolates were partitioned into three
subpopulations in which recent and old isolations from
Iowa formed two separate groups and the rest of the
isolates from other states were combined to form the
third subpopulation. All population analysis coefficients
were computed using the program PopGen 1.31 (Yeh
et al., 1999).
Disease and plant growth data from isolate aggressiveness tests were subjected to analysis of variance using
the generalized linear mixed model (PROC MIXED) of
SAS (SAS Institute, Cary, NC). A nested data structure
was used considering isolates nested within genotype and
genotypes nested within subgroup. Soybean cultivar and
isolate subgroup were treated as fixed effects, while trial,
genotype and isolate were considered random effects.
The area under disease progress curves (AUDPC) for
foliar severity was calculated using the trapezoidal integration method (Campbell & Madden, 1990). For root rot,
Pearson correlation coefficients and statistical significance
were calculated using the PROC CORR procedure of
SAS (SAS Institute, Cary, NC) to determine associations
between foliar severity AUDPC, root rot severity, root dry
weight, shoot height and shoot weight, and inoculum density (log transformed), as well as to test the correlation
between greenhouse and culture filtrate assays for foliar
symptoms.
Data analysis
For all fingerprinting methods, bands were scored as
present or absent to generate a binary matrix. Each
band in a profile was considered a single locus with
two alleles, present or absent. Each unique combination
of polymorphic bands was considered a genotype. Data
were transformed into a matrix to calculate genetic distances among isolates. Cluster analysis was performed by
UPGMA with Jaccard’s similarity coefficient (Sneath &
Sokal, 1973), and the goodness of fit of the phenogram
for the similarity matrix was measured by cophenetic
Results
RFLP, minisatellite and microsatellite analyses
In the RFLP analysis, the restriction enzyme HaeIII generated identical mitochondrial fingerprint patterns for all
F. virguliforme isolates tested in this study. The fingerprinting patterns obtained with the restriction enzymes
Cfo 1 and Msp 1 were not repeatable due to incomplete restriction reactions. The two minisatellite primers
M13 and T3B, the microsatellite primers (GACA)4 and
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Genetic structure and aggressiveness of Fusarium virguliforme
(GTG)5 , and in-gel hybridization of genomic DNA with
the (CAC)5 probe, also generated fingerprint patterns
that were identical among all isolates of F. virguliforme. However, hybridization with the (CAT)5 probe
revealed 30 fingerprint bands, six of which were polymorphic and confined to the high molecular weight DNA
fragments (4, 9, 9.4, 15, 23 and 24 kb). No polymorphic bands were observed within the lower molecular
weight fragments of the (CAT)5 hybridization. Analysis
of the distribution of the six polymorphic bands generated a total of eight unique fingerprint profiles among
the 72 F. virguliforme isolates from Iowa, Minnesota,
Illinois and Arkansas (Table 1). The majority of the
isolates collected in 2006 and 2007 from Iowa (LLcoded isolates) had the same fingerprint profile (C1).
The 10 isolates of F. virguliforme from Minnesota were
grouped in genotype C1, genotype C6 that included
the two isolates from Illinois (Mont 1, and ILMont1A), and genotype C7 that was unique to Minnesota
(Table 1). The isolate from Arkansas (Rupe) belonged to
genotype C4.
RAPD analysis
Two of the 12 RAPD primers generated identical fingerprints among the primers tested for all the F. virguliforme
isolates. The primers OPB-05 and OPA-13 were the most
polymorphic, generating four polymorphic bands each
(data not shown). Seven of the 12 primers identified two
genotypes among the 72 isolates of F. virguliforme, two
primers identified three genotypes, and one primer (PM04) identified four genotypes. Analysis of all 12 RAPD
primers resulted in 13 genotypes among the 72 isolates
of F. virguliforme (Table 1). The most common genotype
(R1) contained more than 70% of the isolates. Among
the most recent collection of isolates from Iowa, 23 isolates belonged to the R1 genotype, while 10 isolates were
distributed among six genotypes other than R1 (Table 1).
Of the 25 isolates collected prior to 2001, nine belonged
to genotypes other than R1. All 10 isolates obtained from
Minnesota had the R1 genotype. Isolate Mont-1 from
Illinois belonged to genotype R8.
Combined fingerprint analysis
The dendrogram generated with the RAPD binary data
conflicted to some degree with that generated with the
(CAT)5 dataset. More genotypes were resolved with the
RAPD dendrogram than within the (CAT)5 dendrogram
(Table 1). The P-value for the P-H test to determine
if there were any differences between the two dendrograms was 0.08 (data not shown). The three additional
89
non-parametric tests for combinability of datasets yielded
different results depending on the comparison being
made. When the (CAT)5 dataset was compared with
the RAPD dendrogram, results of all three tests showed
that both RAPD and (CAT)5 dendrogram topologies
were equally well supported by the (CAT)5 dataset
(P = 0.0901 for KH, P = 0.0679 for TT test, P =
0.1250 for WS test). On the other hand, when the (CAT)5
dendrogram was compared with the RAPD dataset, the
dendrogram was not supported by the RAPD dataset (P ≤
0.0001 for KH test, P = 0.0006 for TT test, P ≤ 0.0001 for
WS test). Based on results of the P-H test alone, the
datasets were combined and analysis resulted in a total
of 25 genotypes among the 72 isolates of F. virguliforme.
Four possible subgroups could be distinguished within
the dendrogram (Fig. 1).
In the combined analysis, genotype VIII within
subgroup 1 was geographically widely distributed and
was found in 12 of the 16 counties from which samples were collected and comprised 43 isolates (Table 1).
However, no isolates belonging to genotype VIII were
identified in the Iowa counties of Henry, Jasper, Clinton
and Fayette. Among the 1993 to 2000 Iowa isolates, 11
(44%) belonged to genotype VIII, while the rest were
distributed among 11 genotypes. Out of the 33 isolates
that made up the 2006 and 2007 Iowa isolates, 21 (63%)
belonged to genotype VIII, while the remaining 12 isolates were distributed among 11 genotypes. Six isolates
from Minnesota belonged to genotype VIII, two isolates
to genotype IX, and two isolates to genotype IV. The
Mont-1 and Fsg-ISU-19 isolates obtained from Illinois
belonged to genotypes XIV and XXI, respectively, while
the single isolate from Arkansas belonged to genotype
XVIII. The cluster pattern showed no association between
genotype and geographic origin.
Gene diversities in the total population and within three
subpopulations (old Iowa isolates, new Iowa isolates and
isolates from other states) were similar (h < 0.200). The
extent of differentiation was low (Gst < 0.06) and the level
of gene flow was relatively high (Nm = 8.431). There
were 16 single-isolate genotypes distributed unequally
among the four subgroups. The genetic distance between
the recent and old isolates from Iowa was larger than
between the old Iowa isolates and isolates from the other
states (0.014 and 0.034, respectively). The genetic identities between the pairs of these three populations were very
similar (0.986, 0.990 and 0.967).
Isolate aggressiveness
Greenhouse assays. Overall analysis of variance showed
a significant effect (P < 0.0001) of trial and trial ×
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G. Y. C. Mbofung et al.
90
Fig. 1. Dendrogram showing relatedness of genotypes of Fusarium virguliforme isolates from Iowa, Minnesota, and Illinois based on combined (CAT)5 and RAPD binary data and cluster analysis (unweighted paired group method with arithmetic means, UPGMA). Roman numerals
at branch ends indicate the different genotypes (I to XXV), and numbers within parentheses represent the number of isolates within each
genotype. The numbers on the branches represent bootstrap support values greater than 50%.
isolate interaction on foliar severity (AUDPC) and root rot
severity; therefore, data were analyzed separately for each
trial. Interveinal chlorotic symptoms on leaves appeared
10 days after planting. In both trials of greenhouse
experiments, foliar severity and root rot severity differed
significantly (P ≤ 0.0004) among isolates, but no differences in these symptoms were observed (P > 0.05) among
genotypes and among subgroups (Table 2). The main
effect of cultivar was significant (P = 0.0483) for root rot
severity in trial 1 and for foliar severity (P = 0.0004) in
Table 2. Analysis of variance on root rot severity and area under disease progress curve (AUDPC) for foliar symptom severity of soybean
sudden death syndrome on plants grown in infested greenhouse soil or exposed to culture filtrates of Fusarium virguliforme in two experimental
trials.
Greenhouse assay
Root rot severity
Culture filtrate assay
Foliar AUDPC
Foliar AUDPC
df
F value
P>F
F value
P>F
df
F value
P>F
Cultivar
Subgroup
Subgroup∗ Cultivar
Genotype
Isolate
Cultivar∗ Isolate
Residual
1
3
3
19
7
25
474
4.31
1.85
1.00
0.71
5.52
1.45
0.0483
0.2267
0.4087
0.7413
0.0004
0.0750
1.49
2.83
1.84
0.45
13.79
0.99
0.2337
0.1667
0.1655
0.9229
<0.0001
0.4822
1
3
3
19
5
24
498
22.70
1.71
0.98
6.74
0.88
1.20
<0.0001
0.2794
0.4198
0.0228
0.5090
0.2370
Cultivar
Subgroup
Subgroup∗ Cultivar
Genotype
Isolate
Cultivar∗ Isolate
Residual
1
1
3
19
7
26
528
0.39
1.98
0.66
1.06
6.01
1.08
0.5350
0.2061
0.5830
0.5050
0.0003
0.0095
16.78
2.14
0.92
0.57
8.17
2.00
0.0004
0.1831
0.4439
0.8470
<0.0001
0.0026
1
3
3
19
7
26
298
14.49
27.44
1.38
0.25
3.17
2.35
0.0008
0.0005
0.2710
0.9921
0.0145
0.0003
Trial
Effect
1
2
trial 2, and there was a significant cultivar × isolate interaction in trial 2 (Table 2). When data were analyzed
separately by cultivar within each trial, the effects of isolate on root rot severity and foliar severity were always
significant (P < 0.001), but there were no differences
among genotype and subgroup, except for a significant
effect (P = 0.019) of subgroup on foliar severity for
cultivar ‘Spencer’ in trial 1.
Both soybean cultivars developed typical SDS symptoms in response to infection by F. virguliforme, but
symptom severity was greater in trial 1 than in trial 2.
Foliar severity at the final assessment (30 days postinoculation) ranged from 50 to 82% and 0 to 77% in trial
1 and 2, respectively, on the susceptible cultivar ‘Spencer’,
and from 47 to 82% and 0 to 48% in trial 1 and 2, respectively, on the moderately resistant cultivar 92M91. Noninoculated control plants did not develop foliar symptoms
in either trial. Foliar severity varied greatly among isolates
a
91
and within each genetic group, with isolates ranging from
high to low aggressiveness within genotypes and subgroups (Fig. 2). However, the relative aggressiveness of
the isolates was not consistent between experimental trials. For example, the isolates LL0001, LL0091, Fsg-ISU
15-5 and Fsg-ISU 22G2 (numbered 11, 6, 17 and 27,
respectively) were among the most aggressive in trial
1 but were among the least aggressive in trial 2 on both
hosts. In contrast, isolates LL0130, LL0009, LL0105 and
Fsg-ISU 6 (numbered 10, 12, 20 and 28, respectively)
were consistently among the most aggressive, and isolates Fsg-ISU 17 and Fsg-ISU 22B1 (numbered 26 and 29,
respectively) were consistently among the least aggressive
isolates, in both trials and on both cultivars. The Mont1 isolate (numbered 19) ranked among the moderately
aggressive isolates in trial 1 and was among the most
aggressive isolates in trial 2. Root rot severity also varied
among isolates and within genetic groups, with a greater
LSD = 202.6
1000
800
600
Area under the disease progress curve
400
200
0
b
LSD = 138.0
600
400
200
0
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
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Genetic structure and aggressiveness of Fusarium virguliforme
Fig. 2. Area under the disease progress curve (AUDPC) for foliar symptom severity on soybean ‘Spencer’ inoculated with 30 Fusarium
virguliforme isolates and incubated for 30 days under greenhouse conditions in (a) trial 1 and (b) trial 2.
G. Y. C. Mbofung et al.
92
a
LSD = 11.5
100
80
60
Root rot severity (%)
20
0
b
LSD = 12.3
80
60
40
20
0
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
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40
Fig. 3. Root rot severity on soybean plants ‘Spencer’ inoculated with 30 Fusarium virguliforme isolates and incubated for 30 days under
greenhouse conditions in (a) trial 1 and (b) trial 2. Per cent root rot was evaluated at the end of the experiment and was calculated as the
percentage of necrotic root area.
range in severity values observed in trial 2 (Fig. 3). Root
rot severity was generally greater than 80% for most isolates in trial 1 and greater than 60% in trial 2 (Fig. 3).
Isolates Fsg-ISU 17 and Fsg-ISU 22B1 (numbered 26 and
29, respectively) were the least aggressive in terms of
causing root rot in both trials (Fig. 3). Root rot severity
rating on non-inoculated controls plants was 0% in trial
1 and 18% in trial 2.
Analysis of variance on root dry weight, shoot dry
weight and shoot height showed significant differences
among isolates (P < 0.05) on both cultivars in both trials
(data not shown). No differences in plant growth variables were found among genotypes or subgroups, except
on cultivar 92M91, where subgroup had an effect on shoot
height (P = 0.0074) in trial 2, and on shoot weight (P =
0.04) in trial 1. The pathogen density in soil at the end
of the experiment varied among isolates (P = 0.002), and
was 1000-fold greater in trial 1 compared with trial 2 (data
not shown).
Significant correlations were found between disease and plant growth parameters in plants from the
greenhouse assay. Foliar disease severity was negatively
correlated with root dry weight, shoot height and shoot
weight for both cultivars in both trials, but was only correlated with root rot severity for cultivar ‘Spencer’ in trial 1
(Table 3). Root rot severity was negatively correlated with
root and shoot dry weight, and shoot height in trial 1, but
was only correlated with shoot weight in trial 2 (Table 3).
In trial 1, final inoculum density was significantly correlated with the AUDPC for foliar symptoms only on
‘Spencer’, and was correlated with other measured disease
and plant growth parameters on both cultivars (Table 3).
In trial 2, the final inoculum density was not correlated
with any disease or growth parameters.
Genetic structure and aggressiveness of Fusarium virguliforme
93
Table 3. Pearson’s correlation coefficient values and significance for Fusarium virguliforme inoculum density in potted mix at the end of the
experiment, foliar severity symptoms (area under disease progress curve, AUDPC), root dry weight, shoot length, shoot weight and root rot
severity ratings of soybean ‘Spencer’ in trials 1 and 2 of the experiment.
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Trial
Final inoculum
density
Foliar severity
(AUDPC)
Root dry
weight
Shoot
length
Shoot
weight
Root rot
severity
1
Final inoculum density
Foliar severity (AUDPC)
Root dry weight
Shoot length
Shoot weight
Root rot severity
1.000
0.383∗
1.000
−0.597∗∗
−0.514∗∗
1.000
−0.216
−0.682∗∗
0.736∗∗
1.000
−0.616∗∗
−0.629∗∗
0.570∗∗
0.581∗∗
1.000
0.630∗∗
0.487∗∗
−0.393∗
−0.595∗∗
−0.416∗
1.000
2
Final inoculum density
Foliar severity (AUDPC)
Root dry weight
Shoot length
Shoot weight
Root rot severity
1.000
0.209
1.000
−0.107
−0.668∗∗
1.000
−0.087
−0.755∗∗
0.460∗
1.000
−0.242
−0.805∗∗
0.802∗∗
0.712∗∗
1.000
0.198
0.112
−0.286
−0.105
−0.366∗
1.000
∗P
≤ 0.05; ∗∗ P ≤ 0.001; n = 30.
Culture filtrate assays. Overall analysis of variance
showed a significant effect of trial (P = 0.0001) and
trial × isolate interaction (P < 0.0001) on foliar symptoms caused by culture filtrates, so the main effects of
cultivars, subgroup, genotype and isolate were tested separately for each trial. The main effect of cultivar was
significant in both trials of the experiment, and there
was a significant cultivar × isolate interaction in trial 2
(Table 2). In trial 1, genotypes had a significant effect
on foliar symptoms, but there was no effect of isolate or
subgroup. In trial 2, isolate and subgroup were significant,
but genotype was not (Table 2). When data were analyzed
separately by cultivar within each trial, there were significant effects of isolate (P = 0.0004 on ‘Spencer’ and P =
0.0001 on ‘91M92’) and subgroup (P = 0.02 on ‘Spencer’
and P = 0.0007 on 92M91) in trial 2, but there were no
effects of isolate, genotype or subgroup in trial 1.
Both soybean cultivars developed typical SDS foliar
symptoms (chlorosis and necrosis) four days after the
roots were dipped in culture filtrates of F. virguliforme isolates. Foliar severity at the final assessment time ranged
from 3 to 31% and 4 to 38% in trial 1 and 2, respectively, on susceptible ‘Spencer’, and from 2 to 57% and
4 to 28% in trial 1 and 2, respectively, on 92M91. Foliar
severity varied greatly among isolates and within genetic
subgroups on both trials (Fig. 4). The isolates LL0039,
LL0038 and LL0019 (numbered 1, 8 and 21, respectively)
were among the most aggressive on both cultivars, while
isolates Fsg-ISU 19 and Fsg-ISU 17 (numbered 25 and
26, respectively) were among the least aggressive (Fig. 4).
Some control plants showed a uniform leaf chlorosis that
was very distinct from the characteristic foliar symptoms
of SDS. This type of chlorosis was 2 and 19% in trial 1 and
2, respectively, on ‘Spencer’ and 0 and 16% in trial 1 and
2, respectively, on 92M91.
Correlations of isolate aggressiveness between assays.
Based on foliar AUDPC values, there was no significant
correlation (P > 0.05) between the two experimental trials
of the greenhouse assay and between the two experimental trials of the culture filtrate assay for either cultivar.
However, trial 1 of the greenhouse assay was correlated
with trial 1 of the culture filtrate assay (P = 0.0007,
r = 0.60) on cultivar ‘Spencer’, and with both trials of
the culture filtrate assay (P = 0.006, r = 0.5 and P =
0.007, r = 0.49 for trials 1 and 2, respectively) on cultivar
92M91. No significant correlations were found between
trials 1 and 2 of the greenhouse assay for root dry weight,
shoot dry weight, shoot height and root rot severity, except
for root rot severity on cultivar ‘Spencer’ (P = 0.0001,
r = 0.64).
Discussion
Despite the genetic homogeneity observed within F.
virguliforme in earlier studies (Achenbach & Patrick,
1996; Achenbach et al., 1997; Aoki et al., 2003, 2005;
Arruda et al., 2005), this study revealed the existence
of intraspecific genetic variation using RAPD and the
(CAT)5 markers. However, no associations were found
between the genetic markers and differences in aggressiveness detected among isolates in greenhouse inoculation and culture filtrate assays. The identification of a
major genotype among the F. virguliforme isolates from
G. Y. C. Mbofung et al.
94
a
LSD = 143.1
b
LSD = 204.7
600
Area under the disease progress curve
200
0
600
400
200
0
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
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400
Fig. 4. Area under the disease progress curve (AUDPC) for foliar symptom severity on soybean ‘Spencer’ with roots immersed in cell-free
culture filtrates of 30 Fusarium virguliforme isolates and incubated for 25 days in a growth chamber. (a) trial 1 and (b) trial 2.
Iowa and Minnesota suggests that a single genotype may
have been introduced into the region, and that minor
genotypes may have arisen from mutations.
RAPD and microsatellite markers have been successfully used to detect intraspecific genetic variations in
other fungi (Wingfield et al., 1996; Doherty et al., 2003;
Harrington et al., 2003; Engelbrecht et al., 2007). An earlier study of genetic variability in F. virguliforme using
RAPDs identified only two genotypes among 55 isolates from nine states, including one isolate from Iowa
(Achenbach et al., 1997). One of the genotypes was represented by only five isolates, including the reference isolate
Mont 1 used in our study. The RAPD markers used in
the present study produced the most polymorphisms, with
a total of 13 genotypes identified among the 72 isolates
of F. virguliforme analyzed, while the (CAT)5 probe generated a total of eight genotypes. Despite being more
conservative than the RAPD markers, the (CAT)5 probe
placed the 10 isolates from Minnesota into three genotypes, although they all belonged to the same genotype
based on the RAPD analysis. It is plausible that a larger
sample from Minnesota might show a more varied population than could be detected with the relatively small
number of isolates analyzed.
Both the separate and combined analysis of the RAPD
and (CAT)5 datasets identified a major genotype within
F. virguliforme. Genotype VIII was found in 12 of the
16 Iowa counties, and was dominant among both older
and more recent isolates from Iowa. This genotype
was also dominant among the isolates from Minnesota.
Genotype VIII may, therefore, have dominated the
original population introduced into Iowa in 1993 (Yang
& Lundeen, 1997), with minor genotypes arising through
mutations. In most studies in which genetic markers such
as (CAT)5 have been successfully used to distinguish
indigenous pathogen populations from introduced ones
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Genetic structure and aggressiveness of Fusarium virguliforme
(Engelbrecht et al., 2007), low levels of genetic variation
have been attributed to a recent introduction of the
pathogen into an area. Often, one or two genotypes are
dominant in the introduced populations and a limited
number of mutations explain the minor variation among
genotypes. For example, low levels of genetic variation
centred around two genotypes of differing aggressiveness
(race A and race M, respectively) were observed among
isolates of the introduced soybean pathogen, Cadophora
gregata f. sp. sojae, using the microsatellite probes
(CAT)5 and (CAC)5 (Harrington et al., 2003). In the case
of F. virguliforme, a larger study including isolates from
other US states would be needed to conclude if genotype
VIII is dominant in other regions and if it might have
been the main genotype introduced into the US from
South America (Aoki et al., 2005).
Although some genetic variation was observed, differences in fingerprint profiles were generally represented by
one to three unique fragments, indicating a very closely
related population of isolates. The similar population
structure and high homogeneity was further suggested by
the low (< 50%) bootstrap support value between two of
the subgroups and the similar gene diversity values for all
subgroups. The low level of genetic variation observed
within the F. virguliforme population further suggests a
recently introduced pathogen (Engelbrecht et al., 2007).
Our study is the first to analyze the relationship
between variation in genetic composition and aggressiveness in F. virguliforme. Variation in isolate aggressiveness
has been previously reported in F. virguliforme (Lim &
Jin, 1991; Achenbach et al., 1997; Li et al., 2000, 2009;
Aoki et al., 2005; Malvick & Bussey, 2008), but the
genetic background of the isolates was not determined
(Lim & Jin, 1991; Roy et al., 1997; Li et al., 2000, 2009),
or no genetic variation was detected among the isolates
studied (Malvick & Bussey, 2008). Isolates from Iowa
varied greatly in their aggressiveness on two soybean
cultivars, as indicated by the ability of some isolates to
cause severe root and foliar symptoms and significant
reductions in plant growth, while other isolates caused
only mild symptoms. However, no association was found
between genetic variation in F. virguliforme and variation
in isolate aggressiveness as assessed in greenhouse
inoculations and by culture filtrate assays. The virulence
of the isolates was very similar on the two soybean
cultivars, suggesting that the inoculum densities used in
both assays were sufficient to cause disease even in the
moderately resistant cultivar ‘92M91’. These results raise
important questions about soybean cultivar resistance and
inoculum thresholds in the field.
Several explanations are possible for the lack of association between isolate aggressiveness and genetic markers.
First, the genetic markers used may not be linked to
95
aggressiveness traits, as has been reported in other fungal
pathogens (Mishra et al., 2003; Trouillas & Gubler, 2010).
According to Pariaud et al. (2009), aggressiveness is
determined by the combination of numerous quantitative traits, and there can be significant differences in
aggressiveness among isolates that belong to the same
genetic group as defined by neutral markers. In a study
on Ceratocystis fimbriata (Harrington et al., 2011), for
example, a large variation in aggressiveness was found
within and among pathogen populations from different
hosts, but aggressiveness was not correlated with genetic
markers. They also reported nearly as much variation in
aggressiveness within lineages and among lineages of
C. fimbriata on some hosts (Harrington et al., 2011).
Similarly, variation in F. virguliforme aggressiveness was
not associated with genetic markers in our study, and variation in aggressiveness was greater among isolates than
among genotypes.
Secondly, the genetic markers used in our study may
have been linked only to a small subset of the minor genes
that define aggressiveness, making it difficult to associate
a definite phenotype to the genotypes. Aggressiveness
components of F. virguliforme include sporulation rate,
infection efficiency (Luo et al., 1999), ability to colonize
the xylem (Navi & Yang, 2008) and ability to produce
toxins that cause foliar symptoms (Li et al., 2009). Since
aggressiveness components are defined by quantitative
traits, variations in several of these components will occur
in natural populations through mutations (Pariaud et al.,
2009). These mutations may have reduced aggressiveness
by affecting any of these components, thereby causing
variations within the genotypes as defined by the markers
used.
Finally, a large portion of variation in aggressiveness
may have resulted from an isolate by environment interaction, which poses a major constraint to the reliability
of aggressiveness tests (Pariaud et al., 2009). The interaction of the environment on aggressiveness may have
been particularly significant in this study since the SDS
pathosystem is well-known for its dependence on environmental factors, such as temperature and soil moisture
(Scherm & Yang, 1996; Roy et al., 1997). The lack of
correlation between trials of the greenhouse and culture
filtrate assays, resulting from the inconsistency in aggressiveness ranking among isolates further illustrates the
challenges in phenotyping soybean responses to infection
by F. virguliforme (Torto et al., 1996; Njiti et al., 2001;
Hashmi et al., 2005). Despite best efforts to incubate
plants under similar conditions, greenhouse temperatures
ranged from 22 to 30 ◦ C in trial 1 and 20 to 28 ◦ C
in trial 2 and natural fluctuations in light intensity also
occurred. These variations may have affected pathogen
growth and sporulation, and consequently the relative
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G. Y. C. Mbofung et al.
aggressiveness of isolates. The marked difference in final
inoculum density in soil between the two trials may also
have contributed to the lack of consistent aggressiveness
ratings between greenhouse experiments.
In the SDS pathosystem, the environment may affect
several components of aggressiveness (Scherm & Yang,
1996; Roy et al., 1997). Measures of disease severity generally correlated between experimental trials of the greenhouse study, except for root rot severity, which was not
well correlated with the other parameters. In related studies (Gongora & Leandro, 2007), ratings of root rot before
maximum severity was reached were better correlated
with foliar severity than when root rot was rated at the end
of the experiment. In the present study, the fact that some
plants showed significant root rot, but escaped the development of foliar symptoms, demonstrates the complexity
of a disease where root rot is caused by pathogen colonization, whereas foliar symptoms result from pathogen toxins
translocated to the leaves (Roy et al., 1997). It has been
proposed that some isolates may be good root colonizers
but poor toxin producers (Li et al., 2009).
The findings of this study do not support the hypothesis that variation in aggressiveness is associated with
intraspecific genetic variation. However, the data suggest
that minor quantitative traits that determine aggressiveness vary greatly within the F. virguliforme population,
and that genetic fingerprinting markers do not strongly
correlate with the measures of aggressiveness. In addition,
variation in aggressiveness may result from significant
interactions between isolate and the environment. Since
aggressiveness in the SDS pathosystem is very difficult
to characterize, it is possible that experimental variation may have masked any associations with genotype
or subgroup, highlighting the importance of controlling
experimental conditions in SDS studies. Continued work
to understand the sources of variation in isolate aggressiveness are needed to understand pathogen biology and
help increase the reliability of SDS resistance screening
assays.
Acknowledgements
This work was supported by funding from the Iowa
Soybean Association. We thank Dean Malvick for providing isolates from Minnesota.
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