Journal of Applied Genetics

, Volume 53, Issue 2, pp 237–247 | Cite as

Genetic variation of Fusarium oxysporum isolates forming fumonisin B1 and moniliformin

  • Lidia Irzykowska
  • Jan Bocianowski
  • Agnieszka Waśkiewicz
  • Zbigniew Weber
  • Zbigniew Karolewski
  • Piotr Goliński
  • Marian Kostecki
  • Witold Irzykowski
Open Access
Microbial Genetics · Original Paper

Abstract

Thirty single-spore isolates of a toxigenic fungus, Fusarium oxysporum, were isolated from asparagus spears and identified by species-specific polymerase chain reaction (PCR) and translation elongation factor 1-α (TEF) sequence analysis. In the examined sets of F. oxysporum isolates, the DNA sequences of mating type genes (MAT) were identified. The distribution of MAT idiomorph may suggest that MAT1-2 is a predominant mating type in the F. oxysporum population. F. oxysporum is mainly recognised as a producer of moniliformin—the highly toxic secondary metabolite. Moniliformin content was determined by high-performance liquid chromatography (HPLC) analysis in the range 0.05–1,007.47 μg g−1 (mean 115.93 μg g−1) but, also, fumonisin B1 was detected, in the concentration range 0.01–0.91 μg g−1 (mean 0.19 μg g−1). There was no association between mating types and the mycotoxins biosynthesis level. Additionally, a significant intra-species genetic diversity was revealed and molecular markers associated with toxins biosynthesis were identified.

Keywords

Fusarium oxysporum Fumonisin B1 Moniliformin Mating type idiomorphs Genetic fingerprinting HPLC analysis 

Introduction

Asparagus (Asparagus officinalis L.) is a worldwide well-known vegetable not only due to its high nutritional value, but also because of the biologically active compounds present, including protodioscin and rutin, with a strong cytotoxic activity against many cancer cell lines (Kinghorn et al. 2003; Chin and Garrison 2008). However, the asparagus is often infected by fungi of the Fusarium genus, a causal agent of crown and root rot which decreases the quantity and quality of the yield. A visible chlorosis and wilting of the individual stalks, progressive discolouration of plants and destruction of roots and crown are the main symptoms of this serious disease (Logrieco et al. 2003). F. oxysporum Schlechtend. emend. Snyder & Hansen and F. proliferatum (Matsus.) Nirenberg are the most severe asparagus pathogens (Blok and Bollen 1995; von Bargen et al. 2009). Both species are well-known mycotoxins producers, so their precise identification is of prime concern. New molecular methods have been applied to complement, support and confirm identification based on morphological characters (Geiser et al. 2004).

F. oxysporum is identified worldwide and is known to be phylogenetically diverse and recognised as a moniliformin (MON) producer (Logrieco et al. 2003). Recent studies suggest that some F. oxysporum strains can also biosynthesise fumonisin B1-FB1 (Waśkiewicz et al. 2010a; Stępień et al. 2011a, b). MON exhibits cytotoxic and cardiotoxic activity, causes developmental disorders and may also induce the development of Keshan disease, human myocardial impairment reported in rural areas of China and South Africa (Pineda-Valdes and Bullerman 2000). Fumonisins are polyketide-derived mycotoxins inhibiting a ceramide synthase, the key enzyme in the sphingolipid biosynthetic pathway, resulting in serious mycotoxicoses (Soriano and Dragacci 2004; Voss et al. 2007). Contamination of food with FB1 has been associated with an oesophageal cancer of humans in Africa, China and the United States (Nicholson 2004). Considering an available toxicological evidence, the International Agency for Research on Cancer (IARC) describe FB1 as a probable carcinogenic to humans (IARC 2002). Food processing does not solve the problem because the hydro-thermal stability of fumonisin is high and decomposition products are more toxic than fumonisin itself (Bullerman et al. 2002).

F. oxysporum is a large taxonomic unit, described as a species complex. So far, no known sexual stage of the F. oxysporum is available and the concept of a biological species and the sources of genetic variation are still discussed (Kistler 1997; O’Donnell et al. 2009; Kück and Pöggeler 2009). In most fungi, the mating type locus conferring mating behaviour consists of dissimilar DNA sequences (idiomorphs) in the mating partners (Pöggeler 2001). In heterothallic species, the mating type is controlled by a single locus with two idiomorphic alleles, designated MAT1-1 and MAT1-2 (Kerényi et al. 1999; Turgeon and Yoder 2000). Each MAT idiomorph carries one gene encoding a single MAT-specific DNA binding protein. These proteins probably play a crucial role in the pathways of cell speciation and sexual morphogenesis as regulatory transcription factors. The two MAT alleles contain a conserved alpha (ALPHA) box domain or a high mobility group (HMG) box domain, respectively (Yun et al. 2000). Evolutionary theory indicates that sexual reproduction plays an important role in a pathogen’s evolution, i.e. the development of a new pathogenic race compatible with a disease-resistant cultivar and of a new strain resistant to a fungicide. Recently, it was shown that the presence of the MAT1-2 idiomorph in the F. culmorum genome significantly affects mycelium growth (unpublished data). Assessing the possibility of mating by toxigenic strains is important for the design of successful control strategies, since these strategies are different for clonally and sexually reproducing organisms (McDonald and McDermott 1993). Moreover, there is poor information on the toxicity of pathogenic F. oxysporum isolates with different mating type alleles.

The present study focused on the genetic and toxigenic differentiation of F. oxysporum isolates. The principal aims of the study were to estimate mycotoxins biosynthesis yield and to detect mating type idiomorphs in the F. oxysporum genome. Additionally, the relationship between these features was assessed. Moreover, the level of intra-species genetic diversity were estimated and associations between molecular markers and mycotoxins (FB1 and MON) formation were examined.

Materials and methods

Fungal isolates

Fungi were collected from green (O-02, O-11, O-30) and white (remaining isolates) spears of asparagus ‘Gijnlim’ and ‘Eposs’ cultivated in Western Poland. Fungi were isolated from the basal part of spears (one spear per plant) on the borderline between discoloured and healthy tissues. After disinfection with 1% sodium hypochlorite, five sections (about 1 mm in diameter) of tissue were cut from spears and such a set sections was transferred onto a Petri dish with potato dextrose agar (PDA) medium. One of these fungal cultures from spears was randomly chosen for further work. Thirty single-spore cultures of F. oxysporum were used in the chemical and molecular analyses. All fungal isolates (O-01–O-30) were identified on the basis of their morphology (Booth 1971; Gerlach and Nirenberg 1982; Kwasna et al. 1991). The morphological features of the identified isolates were compared with standard isolates DSM-62287 (Deutsche Sammlung von Mikroorganismen und Zellkulturen, Germany) of F. oxysporum f. sp. asparagi. Isolates of DSM-62287 in this study were designated as O-31.

DNA preparations and molecular identification of F. oxysporum isolates by PCR and sequencing procedures

Mycelia from 9-day-old single-spore cultures of F. oxysporum grown on liquid medium (5 g L−1 of glucose, 1 g L−1 of yeast extract) were separated from solution by vacuum filtration with a Büchner funnel. DNA was extracted and purified using a DNeasy Mini Kit (QIAGEN Inc., Hilden, Germany), according to the manufacturer’s recommendations. Polymerase chain reaction (PCR) with species-specific primers designed based on a partial sequence of calmodulin gene (Mulè et al. 2004a, b) was used to confirm the morphological identification of each fungal isolate. For the PCR analysis, a forward primer CLOX1: 5′-CAGCAAAGCATCAGACCACTATAACTC-3′ and a reverse primer CLOX2: 5′-CTTGTCAGTAACTGGACGTTGGTACT-3′ (Sigma, Pampisford, UK) were used. The amplification reactions were performed with a Taq PCR Core Kit (QIAGEN, Inc., Hilden, Germany). The reaction mixture and amplification conditions were as described previously (Waśkiewicz et al. 2010b). Amplification was carried out in a Biometra Tpersonal 48 thermocycler (Whatman Biometra, Goettingen, Germany).

Additionally, for 12 representative isolates (forming the highest and the lowest amount of FB1 and MON), a part of the translation elongation factor 1-α (TEF) was amplified with the following primer pair: EF1-forward: 3′-ATGGGTAAGGARGACAAGAC-5′ (O’Donnell et al. 1998) and TEF_WI-reverse: 3′- GCTCACGRGTCTGGCCATCCTTG′ (designed in this study). Primer TEF_WI is located about 90 nt downstream of the sequence of the reverse primer designed by O’Donnell et al. (1998). The reaction mixture contained: 20 ng of fungal DNA, 0.2 mM of each dNTP, 1 μM of each primer and 0.5 U of DreamTaq™ polymerase (Fermentas GMBH, St. Leon-Rot, Germany) in 1× reaction buffer with 2 mM MgCl2. Amplification was carried out using the following programme: an initial denaturation for 2 min at 94°C, followed by 45 cycles of denaturation at 94°C for 30 s, primer annealing at 58°C for 30 s and extension at 72°C for 1 min. The amplification was ended with an additional extension at 72°C for 5 min. Amplification products were cleaned and then sequenced on ABI PRISM® 310 Analyzer (Applied Biosystems), according to the procedure described previously (Irzykowska et al. 2005). The resulting sequences were checked manually and were aligned using the computer software package CLUSTAL_X (Thompson et al. 1997). The obtained TEF sequences were compared with all the sequences deposited at the Fusarium ID Database (http://www.fusariumdb.org/index.php).

Diagnostic PCR for mating type detection

To determine the mating type of the F. oxysporum isolates, conserved portions of the ALPHA or HMG boxes of the MAT1-1 and MAT1-2 idiomorphs were amplified with degenerate oligonucleotide primer pairs: forward (ALPHA) 5′-CGCCCTCTKAAYGSCTTCATG-3′ and reverse (ALPHA) 5′-GGARTARACYTTAGCAATYAGGGC-3′ for MAT1-1 and forward (HMG) 5′-CGACCTCCCAAYGCYTACAT-3′ and reverse (HMG) 5′-TGGGCGGTACTGGTARTCRGG-3′ for MAT1-2 (Kerényi et al. 2004). The PCRs were carried out using a Taq PCR Core Kit (QIAGEN, Inc., Hilden, Germany) with a reaction mixture containing: 5 ng of fungal DNA, 0.2 mM of each dNTP, 0.5 μM of each primer and 0.5 U of Taq DNA polymerase in 1× reaction buffer with 2.5 mM magnesium chloride. The PCR profile was as described previously (Irzykowska and Kosiada 2011).

Genetic fingerprinting of F. oxysporum

Random amplified polymorphic DNA PCRs (RAPD-PCRs) were carried out using a Taq PCR Core Kit (QIAGEN, Inc., Hilden, Germany). The reaction mixture was as described previously (Irzykowska and Bocianowski 2008). Eight random 10-mer and one 11-mer primers, OPB-07, OPC-02, OPC-04, OPC-08, OPC-15, OPJ-10 + A, OPL-11, OPL-12 and OPL-19 (Qiagen Operon, Cologne, Germany), were used to screen the isolates for DNA polymorphism. Amplification was carried out using the following programme: an initial denaturation for 2 min at 94°C, followed by 40 cycles of denaturation at 94°C for 30 s, primer annealing at 37°C for 1 min and extension at 72°C for 2 min. The amplification was ended with an additional extension at 72°C for 5 min. PCR was repeated twice in order to check reproducibility. The comparison of each band profile for each primer was performed on the basis of the presence (1) versus the absence (0) of RAPD products of the same length.

Electrophoresis conditions

The PCR products were separated by electrophoresis (4 V cm−1) in 1.5% agarose gels with 1× TBE buffer (89 mM Tris-borate and 2 mM EDTA, pH 8.0) and visualised under UV light following ethidium bromide staining. A Gene Ruler™ 100 bp DNA Ladder Plus (Fermentas GMBH, St. Leon-Rot, Germany) was used as a molecular size standard for PCR products.

Fungal cultures for mycotoxin estimation

For toxin quantification, rice cultures were prepared for individual F. oxysporum isolates. Long-grain white rice (RANI) tested as being mycotoxin-free was used as a medium for the formation of mycotoxins by F. oxysporum isolates. Rice samples of 65 g were soaked for 16 h with 81 ml of distilled water in 500-ml Erlenmeyer flasks and then sterilised for 15 min at 120°C. Such samples were inoculated with 5-mm disks of PDA medium overgrown by F. oxysporum mycelium. On rice substrate, fungal cultures were incubated for 3 weeks at 20°C. All samples for analyses were prepared and cultured in triplicate (three Erlenmeyer flasks with the prepared medium) and the mean values of the results were calculated.

Mycotoxin analyses

Chemicals and reagents

Standards of pure FB1 and MON were purchased from Sigma (St. Louis, MO, USA). Acetonitrile, methanol (high-performance liquid chromatography [HPLC] grade), disodium tetraborate, 2-mercaptoethanol and t-butyl-ammonium hydroxide were purchased from Sigma-Aldrich. Sodium dihydrogen phosphate, acetic acid, n-hexane, o-phosphoric acid and dichloromethane were purchased from POCh (Gliwice, Poland). Water for the HPLC mobile phase was purified using a Milli-Q system (Milipore, Bedford, MA, USA).

Fumonisin B1 analysis

Samples (10 g) of dry 3-week-old cultures were homogenised for 3 min in 20 ml of methanol–water (3:1, v/v) and filtered through Whatman no. 4 filter paper, in accordance with the method described by Sydenham et al. (1990). The detailed procedure of the extraction and purification of FB1 has been reported previously (Waśkiewicz et al. 2010a). Purified FB1 was quantitatively determined by the HPLC method. The o-phosphoric acid (OPA) reagent (20 mg per 0.5 ml methanol) was prepared and diluted with 2.5 ml of 0.1 M disodium tetraborate (Na2B4O7 × 10H2O) and then combined with 25 ml of 2-mercaptoethanol added to the solution. The FB1 standard (5 μl) or extracts (20 μl) were derivatised with 20 or 80 μl of the OPA reagent. After 3 min, the reaction mixture (10 ml) was injected onto an HPLC column. Methanol–sodium dihydrogen phosphate (0.1 M in water) solution (77:23, v/v), adjusted to pH 3.35 with o-phosphoric acid, after filtration through a 0.45-mm Waters HV membrane, was used as the mobile phase with a flow rate of 0.6 ml min−1. A Waters 2695 apparatus (Waters Division of Millipore, Milford, MA, USA), with a C-18 Nova Pak column (3.9 × 150 mm) and a Waters 2475 fluorescence detector (λex = 335 nm, λem = 440 nm), was used in the metabolite quantitative determination. The identification and quantification of FB1 was observed by comparison of the retention times and peak areas in the samples with those observed for FB1 standard and using the relevant calibration curve (correlation coefficients for FB1 was 0.9967). The limit of detection (LOD) was 10 ng g−1 for FB1 and was defined as the concentration that was three times higher than the standard deviation of the blank signal. Recovery for FB1 measured by the mycotoxins extraction from blank samples spiked with 10–100 ng g−1 of the compound was equal to 93%. The relative standard deviations (RSDs) were lower than 8%.

Moniliformin analysis

Culture samples (15 g) of each strain were homogenised with 75 ml of acetonitrile–methanol–water (16:3:1, v/v/v) and filtered (Whatman no. 4 filter paper). Moniliformin was extracted and purified according to the methods described by Waśkiewicz et al. (2010b). MON was quantified using a Waters 501 apparatus (Waters Division of Millipore) with a C-18 Nova Pak column (3.9 × 300 mm) and a Waters 486 UV detector (λmax = 229 nm). Acetonitrile–water (15:85, v/v) buffered with 10 ml of 0.1 M K2HPO4 in 40% t-butyl-ammonium hydroxide in 1 L of solvent (Sharman et al. 1991) was used as the mobile phase (flow rate = 0.6 ml min−1). The MON detection limit was 25 ng g−1. Positive results (on the basis of retention time) were confirmed by the HPLC analysis and by comparison with the relevant calibration curve (the correlation coefficient for MON was 0.9990). The recovery for MON was 90%. The RSD was below 7%.

Statistical analysis

The one-way analysis of variance (ANOVA) of F. oxysporum occurrence was carried out in order to determine the differentiation of isolates regarding the concentrations of FB1 and MON. Tukey’s least significant differences (LSDs) for each trait were calculated. Homogeneous groups (not significantly different from each other) for the analysed traits were determined on the basis of least significant differences. The relationship between FB1 and MON concentrations was estimated using correlation coefficients (Kozak et al. 2010).

The coefficients of genetic similarity (S) of the investigated isolates were calculated using the following formula (Nei and Li 1979):
$$ {S_{{ij}}} = \frac{{2{N_{{ij}}}}}{{\left( {{N_i} + {N_j}} \right)}} $$
where Nij is the number of alleles present at the i-th and j-th isolates, Ni is the number of alleles present at the i-th isolate, Nj is the number of alleles present at the j-th isolate, and i, j = 1, 2,…, 31. On the basis of the calculated coefficients, isolates were grouped hierarchically using the unweighted pair group method of arithmetic means (UPGMA). The relationship among isolates was presented in the form of a dendrogram (Kumar et al. 2004).

The association between RAPD markers and the FB1 and MON concentrations of F. oxysporum isolates was estimated by regression analysis (Bocianowski and Seidler-Łożykowska 2012). The molecular marker observations were treated as independent variables and considered in individual models. The relationship between MAT idiomorphs and the biosynthesis of FB1 and MON were performed by the t-test (Snedecor and Cochran 1989). All statistical analyses were performed using the package GenStat v.7.1. (Payne et al. 2003).

Results

Identification of F. oxysporum isolates

All of the tested isolates displayed morphology typical for F. oxysporum, quite similar to standard F. oxysporum f. sp. asparagi isolates DSM-62287 (here, O-31). Species-specific PCR assay targeting the calmodulin gene confirmed morphological species identification. Specific PCR products obtained for each isolate, including the reference one, was 534 bp in length.

Additionally, for representative isolates producing and non-producing mycotoxins, a 810-bp fragment of the TEF gene was sequenced. The sequences were compared with sequences deposited in the Fusarium ID database. The comparison with the TEF sequence in the Fusarium ID database showed 100% matching of all isolates examined with F. oxysporum, confirming the species identity (Table 1).
Table 1

Isolates displaying the highest and lowest toxin concentrations with 100% sequence matching with Fusarium oxysporum from the Fusarium ID database

Isolate

FB1 (μg g−1)

MON (μg g−1)

Fusarium ID isolates with 100% homology in the TEF sequence

Source of Fusarium ID isolates

O-08

0.5

60.2

FD_00056 (F. oxysporum)

CBS 175.35

O-10

0.9

16.1

FD_00705 (F. oxysporum)

ICMP 5234

O-12

0.5

9.8

FD_00056 (F. oxysporum)

CBS 175.35

O-15

0.3

1,007.5

FD_00805 (F. oxysporum)

ICMP 15895

O-17

0.0

0.6

FD_00446 (F. oxysporum)

CBS 151.27

O-18

0.0

0.5

FD_00446 (F. oxysporum)

CBS 151.27

O-21

0.0

0.0

FD_00070 (F. oxysporum)

CBS171.31

O-25

0.0

2.4

FD_00805 (F. oxysporum)

ICMP 15895

O-27

0.2

0.0

FD_00705 (F. oxysporum)

ICMP 5234

O-28

0.0

0.0

FD_00802 (F. oxysporum)

FRC O-1890

O-29

0.0

171.6

FD_00805 (F. oxysporum)

ICMP 15895

O-30

0.2

182.8

FD_00705 (F. oxysporum)

ICMP 5234

Evaluation of mycotoxins level

HPLC analysis revealed that the majority of F. oxysporum isolates produce both MON and FB1. The results of one-way ANOVA indicate that isolates differ significantly in their toxigenic activity and mycotoxin profile (Table 2). FB1 was determined in the concentration range 0.01–0.91 μg g−1 (mean 0.19 μg g−1), while MON was detected in the range 0.5–1,007.47 μg g−1 (mean 115.93 μg g−1). Fumonisin B1 and moniliformin were formed in the highest concentrations by isolates O-10 and O-15, respectively (Table 3). Reference isolate (O-31 = DSM-62287) produced only FB1 at a concentration of 0.2 μg g−1. The FB1 biosynthesis level allowed to distinguish between four homogeneous groups among 31 isolates (Table 3). Two homogeneous groups (first group: O-15 isolate; second group: other isolates) were distinguished based on differences in the MON level (Table 3). Only four isolates (O-05, O-23, O-24 and O-26) did not form mycotoxins. The correlation between FB1 and MON was not statistically significant (r = 0.113, P = 0.544).
Table 2

Mean squares from the analysis of variance (ANOVA) for FB1 and MON concentrations (μg g−1) for F. oxysporum isolates of asparagus spears origin

Source of variation

Degrees of freedom

Biosynthesis of toxins

FB1

MON

Isolates

30

0.133***

101,576***

Residual

62

0.027

6,174

***Statistically significant (P < 0.001)

Table 3

The average values, standard deviations and homogeneous groups for FB1 and MON concentrations (μg g−1) for 31 F. oxysporum isolates with different mating types

Isolate code

Mating type idiomorph

FB1

MON

Mean*

Standard deviation

Mean*

Standard deviation

O-01

MAT1-2

0.27bcd

0.18

0.00b

0.00

O-02

MAT1-2

0.12bcd

0.04

0.00b

0.00

O-03

MAT1-2

0.06bcd

0.06

35.73b

19.98

O-04

MAT1-1

0.06bcd

0.05

32.60b

20.27

O-05

MAT1-2

0.00d

0.00

0.00b

0.00

O-06

MAT1-2

0.07bcd

0.04

2.86b

2.24

O-07

MAT1-1

0.49abc

0.37

0.00b

0.00

O-08

MAT1-2

0.52ab

0.29

60.23b

34.55

O-09

MAT1-2

0.23bcd

0.12

0.00b

0.00

O-10

MAT1-2

0.91a

0.28

16.07b

11.11

O-11

MAT1-2

0.11bcd

0.07

0.00b

0.00

O-12

MAT1-2

0.52ab

0.29

9.82b

4.58

O-13

MAT1-1

0.10bcd

0.07

105.63b

86.21

O-14

MAT1-2

0.30bcd

0.44

49.00b

27.10

O-15

MAT1-2

0.31bcd

0.21

1,007.47a

400.69

O-16

MAT1-2

0.06bcd

0.04

174.00b

127.06

O-17

MAT1-2

0.00d

0.00

0.62b

0.60

O-18

MAT1-2

0.01d

0.01

0.50b

0.33

O-19

MAT1-2

0.01d

0.01

114.03b

38.99

O-20

MAT1-1

0.03cd

0.03

0.00b

0.00

O-21

MAT1-1

0.04cd

0.04

0.00b

0.00

O-22

MAT1-2

0.28bcd

0.17

5.43b

4.20

O-23

MAT1-2

0.00d

0.00

0.00b

0.00

O-24

MAT1-1

0.00d

0.00

0.00b

0.00

O-25

MAT1-1

0.01d

0.02

2.37b

2.46

O-26

MAT1-1

0.00d

0.00

0.00b

0.00

O-27

MAT1-2

0.2bcd

0.02

0.00b

0.00

O-28

MAT1-1

0.03cd

0.04

0.00b

0.00

O-29

MAT1-2

0.01d

0.01

171.57b

53.31

O-30

MAT1-1

0.23bcd

0.36

182.80b

1.65

O-31

MAT1-2

0.02cd

0.01

0.00b

0.00

LSD0.001

 

0.47

 

217.8

 

*Mean values followed by the same letters are not significantly different. Each letter indicates a single homogenous group

Mating type assessment by PCR

In the examined set of F. oxysporum isolates, DNA sequences encoding MAT1-1 or MAT1-2 genes were detected (Fig. 1). Isolates O-04, O-07, O-13, O-20, O-21, O-24, O-25, O-26, O-28 and O-30 for which a 200-bp fragment was amplified with ALPHA primers were designated the MAT1-1 type. The remaining 21 isolates for which a 260-bp PCR product was amplified with HMG primer pairs were designated MAT1-2.
Fig. 1

Polymerase chain reaction (PCR) amplification of mating type specific sequences from the selected Fusarium oxysporum isolates. Lanes: 1 (O-04), 2 (O-07), 4 (O-13), 6 (O-20), 12 (O-21), 20 (O-24), 22 (O-25), 24 (O-26): amplicon of MAT1-1 specific ALPHA box (200 bp). Lanes: 3 (O-01), 5 (O-02), 7 (O-03), 8 (O-05), 9 (O-06), 10 (O-08), 11 (O-09), 13 (O-10), 14 (O-11), 15 (O-12), 16 (O-14), 17 (O-15), 18 (O-16), 19 (O-17), 21 (O-18), 23 (O-19): amplicon of MAT1-2 specific HMG box (260 bp). Lane: M: a Gene Ruler™ 100 bp DNA Ladder Plus (Fermentas)

The relationships between MAT1-1 and mycotoxins biosynthesis were not statistically significant (t = −1.11, P = 0.275 and t =−0.65, P = 0.524 for FB1 and MON concentrations, respectively). Similarly, the correlation between MAT1-2 and toxins biosynthesis was not statistically significant (t = 1.11, P = 0.275 and t = 0.65, P = 0.524 for FB1 and MON concentrations, respectively).

Genetic variability determination

To assess a genetic variation among F. oxysporum isolates, genetic fingerprinting by RAPD analyses with nine arbitrary primers was carried out. Three of them were excluded from further analysis because of monomorphism or low repeatability. Finally, 33 polymorphic products were obtained, ranging from 4 to 7 per primer (Table 4). The size of the RAPD fragments ranged from 0.15 to 3.2 kbp. The relationship among isolates of F. oxysporum is presented in the form of a dendrogram (Fig. 2). Clustering analysis resolved two groups at the 30% similarity level. Each of these groups was divided into smaller subgroups at 60 and 52% similarity levels, respectively. The highest genetic similarity (equal to 1) was observed between the isolates O-03 and O-10, O-7 and O-30, O-13 and O-16, O-17 and O-27, and O-18 and O-19, whereas the lowest genetic similarity (equal to 0.0667) was found for O-02 and O-12. The average value of genetic similarity was equal to 0.507.
Table 4

Codes and sequences of random amplified polymorphic DNA (RAPD) primers tested, with polymorphic RAPD marker numbers and sizes

Primer code

Sequence 5′-3′

Polymorphic band number

Polymorphic band size (bp)

OPB-07

GGTGACGCAG

7

a-720, b-880, c-1000, d-1470, e-2280, f-2400, g-3200

OPC-02

GTGAGGCGTC

7

a-390, b-800, c-980, d-1300, e-1460, f-1500, g-2600

OPC-04

CCGTATCTAC

Rejected

OPC-08

TGGACCGGTG

0

Monomorphic

OPC-15

GACGGATCAG

4

a-850, b-1150, c-1500, d-1850

OPJ-10 + A

AAGCCCGAGGA

4

a-660, b-1250, c-1370, d-2630

OPL-11

ACGATGAGCC

5

a-340, b-500, c-600, d-810, e-2010

OPL-12

GGGCGGTACT

6

a-700, b-910, c-1000, d-1130, e-1380, f-1840

OPL-19

GAGTGGTGAC

Rejected

Fig. 2

Dendrogram of F. oxysporum isolates based on random amplified polymorphic DNA (RAPD) data analysis; the isolates were grouped hierarchically using the unweighted pair group method of arithmetic means (UPGMA). *Isolates with MAT1-1 idiomorph

Significant associations of eight markers (OPC-02b, OPC-15 d, OPL-11c, OPL-11 d, OPL-11e, OPL-12 d, OPL-12f and OPB-07b) with FB1 biosynthesis (Table 5) as well as one marker (OPC-15b) with MON production were found (Fig. 3). The percentage variation of FB1 concentration accounted by markers ranged from 11.2% (for OPB-07b) to 26.5% (for OPL-11c). In the case of MON concentration, the percentage variation accounted by OPC-15b marker was equal to 56.8%, which makes this marker a good candidate for further studies.
Table 5

Molecular markers significantly (at the α = 0.05 level) associated with FB1 concentration estimated by regression analysis

Marker symbol

Estimates of regression coefficients

P-value

Percentage variation accounted

Standard error of observations

OPC-02b

−0.223

0.003

24.8

0.183

OPC-15 d

0.173

0.023

13.7

0.196

OPL-11c

0.226

0.002

26.5

0.181

OPL-11 d

−0.235

0.005

22.0

0.186

OPL-11e

0.237

0.003

24.4

0.183

OPL-12 d

0.197

0.008

19.2

0.190

OPL-12f

0.184

0.012

16.9

0.192

OPB-07b

−0.167

0.037

11.2

0.199

Fig. 3

Genetic fingerprinting of F. oxysporum isolates applying RAPD-PCR with the OPC-15 primer. Lanes: MM a Gene Ruler™ 100 bp DNA Ladder Plus (Fermentas), N a negative control. *RAPD marker (OPC-15b) associated with high moniliformin level

Discussion

When considering Fusarium fungi as mycotoxin producers, a proper identification of species infecting food and/or feed plants is very important. Traditional diagnostics based on morphology as well as simple PCR in some cases is insufficient. The translation factor 1-α gene has high phylogenetic utility because it is very informative at the species level and non-orthologous copies of them have not been detected in the Fusarium genus (Geiser et al. 2004). Based on species-specific PCR and partial sequencing of the TEF gene, we proved undoubtedly that isolates forming FB1 belong to F. oxysporum species.

The present study showed explicitly that some of the F. oxysporum isolates infecting asparagus spears have the potential to form and accumulate FB1, although, in general, they produced low amounts of fumonisin and significantly higher amounts of MON. The ability of F. oxysporum to form fumonisin has been questioned previously. Wang et al. (2010) reported that F. oxysporum strains isolated from asparagus spears in Zhejiang Province of China did not produce FB1, but only four F. oxysporum isolates were examined. Similarly, it was shown that F. oxysporum isolated from greenhouse-grown jimsonweed produced only MON at a concentration level of 3.5 μg g−1 (Abbas et al. 1992). On the other hand, Abbas et al. (1995) claimed that three of six isolates of F. oxysporum and seven isolates of F. oxysporum var. redolens from root lesions of eastern white pine seedlings produced FB1, ranging from traces (≥100) to 300.60 ng g−1. This statement was also supported by our earlier studies (Waśkiewicz et al. 2009).

Fumonisin formation is positively related to the FUM genes expression (Waalwijk et al. 2004; Stępień et al. 2011a, b). The FUM genes cluster is highly collinear among F. oxysporum, F. proliferatum and F. verticillioides, regarding gene number, orientation and order (Proctor et al. 2008). The amount of FB1 produced in vitro depends on the substrate, genotype and conditions (Desjardins et al. 1992). It was shown that there is a significant impact of interacting environmental factors on the FUM transcript levels (Jurado et al. 2008). The correlation between FB1 and MON levels was not revealed, probably because of the different biosynthetic pathways of these secondary metabolites (Voss et al. 2007). Naturally occurring genetic variance in the fungal population may explain why non-producing isolates were also recorded. The loss of toxigenic ability can occur due to even a single-point mutation that shifts the open reading frame or creates a premature stop codon (Proctor et al. 2006).

The detection of MAT genes in the genomes of mitosporic species is a first step into learning what causes asexuality (Kück and Pöggeler 2009). The obtained results confirm that F. oxysporum has a heterothallic origin and are in agreement with Turgeon’s hypothesis that some Fusarium species may exhibit a cryptic sexual cycle (Turgeon 1998). Similar results were reported by Arie et al. (2000) for F. oxysporum isolates and by Kerényi et al. (2004) for several Fusarium species. The amplification of the MAT idiomorph sequence revealed that 30% of the examined F. oxysporum isolates possess the MAT1-1 idiomorph and as many as 70% possess the MAT1-2 allele. Similar results for mating types distribution in the fungal population were described for F. oxysporum originated from common bean (Karimian et al. 2010). The maximum effective reproductive strategy occurs when the mating type idiomorphs are present in a 1:1 ratio (Britz et al. 1998). In many Fusarium species, the ratio of MAT idiomorphs was significantly different from the theoretical 1:1 ratio expected in an idealised population, i.e. in a population of F. verticillioides, the mating ratio was 2:1 for isolates originated from maize in Italy, Brazil and the Philippines (Cumagun 2007; de Oliveira Rocha et al. 2011; Venturini et al. 2011). The same mating ratio was observed for an F. subglutinans population in South Africa (Britz et al. 1998). Among F. culmorum isolates originated from wheat and rye, the mating ratio was estimated as 2:3 (Irzykowska and Kosiada 2011). The achieved results may suggest that MAT1-2 is a predominant mating type in the F. oxysporum population in Poland, although a larger population of fungus originated from different geographic locations should be analysed before reaching a final conclusion. The predominance of one mating type can cause a limitation or lack of the possibility of sexual reproduction inside the population (Venturini et al. 2011). RT-PCR analysis proved that the F. oxysporum MAT genes are expressed (Yun et al. 2000). Lack of sexual reproduction may also be a result of the functional disorder of the yet-unidentified genes that are involved in successful sexual reproduction (Arie et al. 2000). Alternatively, asexual Fusarium spp. may require environmental conditions for mating that are uncommon when disease epidemics occur (Kerényi et al. 2004). To summarise, the molecular discrimination of MAT1-1 and MAT1-2 in F. oxysporum populations make possible the recognition of potentially compatible isolates that could be used in future experiments revealing whether the species is truly asexual or not.

The genetic variability of F. oxysporum isolates was estimated previously by different molecular methods. Genetic fingerprinting by RAPD-PCR is an effective method for determining an inter- and intra-species genetic variation without prior knowledge of the genome sequence. RAPD has been used successfully to analyse the genetic variation of several Fusarium species, including F. oxysporum (Assigbetse et al. 1994; Clark et al. 1998; Werner and Irzykowska 2007; Karimian et al. 2010), F. proliferatum (von Bargen et al. 2009), F. culmorum (Miedaner et al. 2004; Irzykowska and Baturo 2008), F. graminearum (Ouellet and Seifert 1993; Carter et al. 2002), F. moniliforme, F. solani and F. avenaceum (Khalil et al. 2003). In the present study, a significant genetic variation among F. oxysporum isolates was revealed and shown in the form of a dendrogram. Clustering analysis resolved several fingerprint groups, six of which contained one isolate. This is consistent with the results of another study on the genetic diversity of F. oxysporum, where from 6 to 10 fingerprint groups were found (Cramer et al. 2003; Zanotti et al. 2006; Karimian et al. 2010). The DNA sequences of the most unlike isolates (O-02 and O-12) differ in analysed genome parts by almost 70%. Isolate O-02 originated from green spears of asparagus ‘Gijnlim’, whereas isolate O-12 originated from white spears of asparagus ‘Eposs’, thus, it is possible that the revealed variation between them resulted from race existence. However, we have found five pairs of isolates which were identical in the examined genome parts. All of these isolates originated from white spears of asparagus ‘Eposs’. Such high genetic similarity can also be the result of the relatively low number of markers used in the study.

It is well known that a high level of intra-species genetic diversity is typical for species undergoing sexual cycle with the meiotic recombination (McDonald 1997; Kerényi et al. 2004). The ways available for genetic change in F. oxysporum are still largely unknown, but some possibilities exist beyond simple sexual or clonal reproduction (Kistler 1997). Daboussi and Langin (1994) reported that active fungal transposable elements may comprise up to 5% of the F. oxysporum genome and may have an impact on gene structure and function. Other molecular studies suggest a genetic duplication in the rDNA regions (O’Donnell and Cigelnik 1997).

Moreover, in this study, nine molecular markers associated with MON or FB1 formation levels were found (Table 4). The obtained PCR products will be used in future for the preliminary discrimination of toxigenic isolates. RAPD marker OPC-15b (∼1,150 bp), connected with MON biosynthesis, will be cloned and sequenced to converse them to the specific and more universal SCAR marker. Amplified regions of the F. oxysporum genome may be involved in the study of toxin biosynthesis pathways. A genetic background of toxin biosynthesis is complex and, probably, some genes still remain unknown. Toxicity as the majority of significant physiological traits (i.e. toxin or protein biosynthesis) is inherited quantitatively, so continuous variation arises from the segregation of alleles at many interacting loci, whose effects depend on the environment (Edwards et al. 1987; Irzykowska et al. 2001; Bocianowski and Krajewski 2009). Proctor et al. (2008) reported that the FUM cluster is located in different genome parts in F. oxysporum, F. proliferatum and F. verticillioides. Perhaps the genetic bases of FB1 formation by F. oxysporum strains have not yet been fully elucidated.

In conclusion, we postulate that F. oxysporum present in asparagus spears produces not only MON but also highly toxic FB1, which can be a significant concern for consumers. The mechanism of genetic changes occurring in F. oxysporum populations and the connection between the toxin profile and the genetic variability of fungus demand further attention. More extensive studies are necessary in order to elucidate the distribution of MATs in the fungus population.

Notes

Acknowledgements

This study was partly supported by the project provided by the Ministry of Science and Higher Education of Poland (Project Number NN 3101709 33).

Open Access

This article is distributed under the terms of the Creative Commons Attribution License which permits any use, distribution, and reproduction in any medium, provided the original author(s) and the source are credited.

References

  1. Abbas HK, Vesonder RF, Boyette CD, Hoagland RE, Krick T (1992) Production of fumonisins by Fusarium moniliforme cultures isolated from jimsonweed in Mississippi. J Phytopathol 136:199–203CrossRefGoogle Scholar
  2. Abbas HK, Ocamb CM, Xie W, Mirocha CJ, Shier WT (1995) First report of fumonisin B1, B2, and B3 production by Fusarium oxysporum var. redolens. Plant Dis 79:968CrossRefGoogle Scholar
  3. Arie T, Kaneko I, Yoshida T, Noguchi M, Nomura Y, Yamaguchi I (2000) Mating-type genes from asexual phytopathogenic ascomycetes Fusarium oxysporum and Alternaria alternata. Mol Plant Microbe Interact 13:1330–1339PubMedCrossRefGoogle Scholar
  4. Assigbetse KB, Fernandez D, Dubois MP, Geiger J-P (1994) Differentiation of Fusarium oxysporum f. sp. vasinfectum races on cotton by random amplified polymorphic DNA (RAPD) analysis. Phytopathology 84:622–626CrossRefGoogle Scholar
  5. Blok WJ, Bollen GJ (1995) Fungi on roots and stem bases of asparagus in the Netherlands: species and pathogenicity. Eur J Plant Pathol 101:15–24CrossRefGoogle Scholar
  6. Bocianowski J, Krajewski P (2009) Comparison of the genetic additive effect estimators based on phenotypic observations and on molecular marker data. Euphytica 165:113–122CrossRefGoogle Scholar
  7. Bocianowski J, Seidler-Łożykowska K (2012) The relationship between RAPD markers and quantitative traits of caraway (Carum carvi L.). Ind Crops Prod 36:135–139CrossRefGoogle Scholar
  8. Booth C (1971) The genus Fusarium. CABI Press, Kew, UK, pp 32–185Google Scholar
  9. Britz H, Wingfield MJ, Coutinho TA, Marasas WFO, Leslie JF (1998) Female fertility and mating type distribution in a South African population of Fusarium subglutinans f. sp. pini. Appl Environ Microbiol 64:2094–2095PubMedGoogle Scholar
  10. Bullerman LB, Ryu D, Jackson LS (2002) Stability of fumonisins in food processing. Adv Exp Med Biol 504:195–204PubMedCrossRefGoogle Scholar
  11. Carter JP, Rezanoor HN, Holden D, Desjardins AE, Plattner RD, Nicholson P (2002) Variation in pathogenicity associated with the genetic diversity of Fusarium graminearum. Eur J Plant Pathol 108:573–583CrossRefGoogle Scholar
  12. Chin CK, Garrison SA (2008) Functional elements from asparagus for human health. Acta Hort 776:219–226Google Scholar
  13. Clark CA, Hyun J-W, Hoy MW (1998) Relationships among wilt inducing isolates of Fusarium oxysporum from sweetpotato and tobacco. Plant Dis 82:530–536CrossRefGoogle Scholar
  14. Cramer RA, Byrne PF, Brick MA, Panella L, Wickliffe E, Schwartz HF (2003) Characterization of Fusarium oxysporum isolates from common bean and sugar beet using pathogenicity assays and random-amplified polymorphic DNA markers. J Phytopathol 151:352–360CrossRefGoogle Scholar
  15. Cumagun CJR (2007) Female fertility and mating type distribution in a Philippine population of Fusarium verticillioides. J Appl Genet 48:123–126Google Scholar
  16. Daboussi MJ, Langin T (1994) Transposable elements in the fungal plant pathogen Fusarium oxysporum. Genetica 93:49–59CrossRefGoogle Scholar
  17. de Oliveira Rocha L, Reis GM, da Silva VN, Braghini R, Teixeira MMG, Corrêa B (2011) Molecular characterization and fumonisin production by Fusarium verticillioides isolated from corn grains of different geographic origins in Brazil. Int J Food Microbiol 145:9–21PubMedCrossRefGoogle Scholar
  18. Desjardins AE, Plattner RD, Shackelford DD, Leslie JF, Nelson PE (1992) Heritability of fumonisin B1 production in Gibberella fujikuroi mating population A. Appl Environ Microbiol 58:2799–2805PubMedGoogle Scholar
  19. Edwards MD, Stuber CW, Wendel JF (1987) Molecular-marker-facilitated investigations of quantitative-trait loci in maize. I. Numbers, genomic distribution and types of gene action. Genetics 116:113–125PubMedGoogle Scholar
  20. Geiser DM, del Mar Jiménez-Gasco M, Kang S, Makalowska I, Veeraraghavan N, Ward TJ, Zhang N, Kuldau GA, O’Donnell K (2004) FUSARIUM-ID v. 1.0: a DNA sequence database for identifying Fusarium. Eur J Plant Pathol 110:473–479CrossRefGoogle Scholar
  21. Gerlach W, Nirenberg HI (1982) The genus Fusarium. A pictorial atlas. Mitt Biol Bundesanst Land Forstwirtschaft. Berlin-Dahlem, Germany, pp 9–386Google Scholar
  22. International Agency for Research on Cancer (IARC) (2002) Fumonisin B1. IARC monographs on the evaluation of the carcinogenic risks to humans. Some traditional herbal medicines, some mycotoxins, naphthalene and styrene. IARC, Lyon, France, vol 82, pp 301–366Google Scholar
  23. Irzykowska L, Baturo A (2008) Genetic polymorphism of Fusarium culmorum isolates originating from roots and stem bases of barley. J Plant Protection Res 48:303–312CrossRefGoogle Scholar
  24. Irzykowska L, Bocianowski J (2008) Genetic variation, pathogenicity and mycelial growth rate differentiation between Gaeumannomyces graminis var. tritici isolates derived from winter and spring wheat. Ann Appl Biol 152:369–375CrossRefGoogle Scholar
  25. Irzykowska L, Kosiada T (2011) Molecular identification of mating type genes in asexually reproducing Fusarium oxysporum and F. culmorum. J Plant Protect Res 51:405–409CrossRefGoogle Scholar
  26. Irzykowska L, Wolko B, Swiecicki WK (2001) The genetic linkage map of pea (Pisum sativum L.) based on molecular, biochemical and morphological markers. Pisum Genet 33:13–18Google Scholar
  27. Irzykowska L, Irzykowski W, Jarosz A, Golebniak B (2005) Association of Phytophthora citricola with leather rot disease of strawberry. J Phytopathol 153:680–685CrossRefGoogle Scholar
  28. Jurado M, Marín P, Magan N, González-Jaén MT (2008) Relationship between solute and matric potential stress, temperature, growth, and FUM1 gene expression in two Fusarium verticillioides strains from Spain. Appl Environ Microbiol 74:2032–2036PubMedCrossRefGoogle Scholar
  29. Karimian B, Javan-Nikkhah M, Abbasi M, Ghazanfari K (2010) Genetic diversity of Fusarium oxysporum isolates from common bean and distribution of mating type alleles. Iran J Biotechnol 8:90–97Google Scholar
  30. Kerényi Z, Zeller K, Hornok L, Leslie JF (1999) Molecular standardization of mating type terminology in the Gibberella fujikuroi species complex. Appl Environ Microbiol 65:4071–4076PubMedGoogle Scholar
  31. Kerényi Z, Moretti A, Waalwijk C, Oláh B, Hornok L (2004) Mating type sequences in asexually reproducing Fusarium species. Appl Environ Microbiol 70:4419–4423PubMedCrossRefGoogle Scholar
  32. Khalil MS, Abdel-Sattar MA, Aly IN, Abd-Elsalam KA, Verreet JA (2003) Genetic affinities of Fusarium spp. and their correlation with origin and pathogenicity. Afr J Biotechnol 2:109–113Google Scholar
  33. Kinghorn AD, Su BN, Lee D, Gu JQ, Pezzuto JM (2003) Cancer chemopreventive agents discovered by activity-guided fractionation: an update. Curr Org Chem 7:213–226CrossRefGoogle Scholar
  34. Kistler HC (1997) Genetic diversity in the plant-pathogenic fungus Fusarium oxysporum. Phytopathology 87:474–479PubMedCrossRefGoogle Scholar
  35. Kozak M, Bocianowski J, Sawkojć S, Wnuk A (2010) Call for more graphical elements in statistical teaching and consultancy. Biometrical Lett 47:57–68Google Scholar
  36. Kück U, Pöggeler S (2009) Cryptic sex in fungi. Fungal Biol Rev 23:86–90CrossRefGoogle Scholar
  37. Kumar S, Tamura K, Nei M (2004) MEGA3: Integrated software for Molecular Evolutionary Genetics Analysis and sequence alignment. Brief Bioinform 5:150–163PubMedCrossRefGoogle Scholar
  38. Kwasna H, Chelkowski J, Zajkowski P (1991) Fungi (Mycota) XXII. Polish Academy of Sciences, Warsaw, Poland, pp 30–111Google Scholar
  39. Logrieco A, Bottalico A, Mulé G, Moretti A, Perrone G (2003) Epidemiology of toxigenic fungi and their associated mycotoxins for some Mediterranean crops. Eur J Plant Pathol 109:645–667CrossRefGoogle Scholar
  40. McDonald BA (1997) The population genetics of fungi: tools and techniques. Phytopathology 87:448–453PubMedCrossRefGoogle Scholar
  41. McDonald BA, McDermott JM (1993) Population genetics of plant pathogenic fungi. BioScience 43:311–319CrossRefGoogle Scholar
  42. Miedaner T, Schilling AG, Geiger HH (2004) Competition effects among isolates of Fusarium culmorum differing in aggressiveness and mycotoxin production on heads of winter rye. Eur J Plant Pathol 110:63–70CrossRefGoogle Scholar
  43. Mulè G, Susca A, Stea G, Moretti A (2004a) Specific detection of the toxigenic species Fusarium proliferatum and F. oxysporum from asparagus plants using primers based on calmodulin gene sequences. FEMS Microbiol Lett 230:235–240PubMedCrossRefGoogle Scholar
  44. Mulè G, Susca A, Stea G, Moretti A (2004b) Corrigendum to “Specific detection of the toxigenic species Fusarium proliferatum and F. oxysporum from asparagus plants using primers based on calmodulin gene sequences”. FEMS Microbiol Lett 232:229CrossRefGoogle Scholar
  45. Nei M, Li WH (1979) Mathematical model for studying genetic variation in terms of restriction endonucleases. Proc Natl Acad Sci USA 76:5269–5273PubMedCrossRefGoogle Scholar
  46. Nicholson P (2004) Rapid detection of mycotoxigenic fungi in plants. In: Magan N (ed) Mycotoxins in food: detection and control. Woodhead Publishing, Cambridge, pp 111–152CrossRefGoogle Scholar
  47. O’Donnell K, Cigelnik E (1997) Two divergent intragenomic rDNA ITS2 types within a monophyletic lineage of the fungus Fusarium are nonorthologous. Mol Phylogenet Evol 7:103–116PubMedCrossRefGoogle Scholar
  48. O’Donnell K, Kistler HC, Cigelnik E, Ploetz RC (1998) Multiple evolutionary origins of the fungus causing Panama disease of banana: concordant evidence from nuclear and mitochondrial gene genealogies. Proc Natl Acad Sci USA 95:2044–2049PubMedCrossRefGoogle Scholar
  49. O’Donnell K, Gueidan C, Sink S, Johnston PR, Crous PW, Glenn A, Riley R, Zitomer NC, Colyer P, Waalwijk C, van der Lee T, Moretti A, Kang S, Kim H-S, Geiser DM, Juba JH, Baayen RP, Cromey MG, Bithell S, Sutton DA, Skovgaard K, Ploetz R, Corby Kistler H, Elliott M, Davis M, Sarver BA (2009) A two-locus DNA sequence database for typing plant and human pathogens within the Fusarium oxysporum species complex. Fungal Genet Biol 46:936–948PubMedCrossRefGoogle Scholar
  50. Ouellet T, Seifert KA (1993) Genetic characterization of Fusarium graminearum strains using RAPD and PCR amplification. Phytopathology 83:1003–1007CrossRefGoogle Scholar
  51. Payne RW, Murrey DA, Harding SA, Baird DB, Soutou D, Lane PW (2003) GenStat for Windows (7th edition) Introduction. VSN International Ltd., Oxford, EnglandGoogle Scholar
  52. Pineda-Valdes G, Bullerman LB (2000) Thermal stability of moniliformin at varying temperature, pH, and time in an aqueous environment. J Food Prot 63:1598–1601PubMedGoogle Scholar
  53. Pöggeler S (2001) Mating-type genes for classical strain improvements of ascomycetes. Appl Microbiol Biotechnol 56:589–601PubMedCrossRefGoogle Scholar
  54. Proctor RH, Plattner RD, Desjardins AE, Busman M, Butchko RAE (2006) Fumonisin production in the maize pathogen Fusarium verticillioides: genetic basis of naturally occurring chemical variation. J Agric Food Chem 54:2424–2430PubMedCrossRefGoogle Scholar
  55. Proctor RH, Busman M, Seo J-A, Lee YW, Plattner RD (2008) A fumonisin biosynthetic gene cluster in Fusarium oxysporum strain O-1890 and the genetic basis for B versus C fumonisin production. Fungal Genet Biol 45:1016–1026PubMedCrossRefGoogle Scholar
  56. Sharman M, Gilbert J, Chelkowski J (1991) A survey of the occurrence of the mycotoxin moniliformin in cereal samples from sources worldwide. Food Addit Contam 8:459–466PubMedCrossRefGoogle Scholar
  57. Snedecor GW, Cochran WG (1989) Statistical methods, 8th edn. Iowa State University Press, AmesGoogle Scholar
  58. Soriano JM, Dragacci S (2004) Occurrence of fumonisins in food. Food Res Int 37:985–1000CrossRefGoogle Scholar
  59. Stępień Ł, Koczyk G, Waśkiewicz A (2011a) FUM cluster divergence in fumonisins-producing Fusarium species. Fungal Biol 115:112–123PubMedCrossRefGoogle Scholar
  60. Stępień Ł, Koczyk G, Waśkiewicz A (2011b) Genetic and phenotypic variation of Fusarium proliferatum isolates from different host species. J Appl Genet 52:487–496PubMedCrossRefGoogle Scholar
  61. Sydenham EW, Gelderblom WCA, Thiel PG, Marasas WFO (1990) Evidence for the natural occurrence of fumonisin B1, a mycotoxin produced by Fusarium moniliforme, in corn. J Agric Food Chem 38:285–290CrossRefGoogle Scholar
  62. Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG (1997) The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucl Acids Res 25:4876–4882PubMedCrossRefGoogle Scholar
  63. Turgeon BG (1998) Application of mating type gene technology to problems in fungal biology. Annu Rev Phytopathol 36:115–137PubMedCrossRefGoogle Scholar
  64. Turgeon BG, Yoder OC (2000) Proposed nomenclature for mating type genes of filamentous ascomycetes. Fungal Genet Biol 31:1–5PubMedCrossRefGoogle Scholar
  65. Venturini G, Assante G, Toffolatti SL, Vercesi A (2011) Mating behavior of a Northern Italian population of Fusarium verticillioides associated with maize. J Appl Genet 52:367–370PubMedCrossRefGoogle Scholar
  66. von Bargen S, Martinez O, Schadock I, Eisold A-M, Gossmann M, Büttner C (2009) Genetic variability of phytopathogenic Fusarium proliferatum associated with crown rot in Asparagus officinalis. J Phytopathol 157:446–456CrossRefGoogle Scholar
  67. Voss KA, Smith GW, Haschek WM (2007) Fumonisins: toxicokinetics, mechanism of action and toxicity. Anim Feed Sci Technol 137:299–325CrossRefGoogle Scholar
  68. Waalwijk C, van der Lee T, de Vries I, Hesselink T, Arts J, Kema GHJ (2004) Synteny in toxigenic Fusarium species: the fumonisin gene cluster and the mating type region as examples. Eur J Plant Pathol 110:533–544CrossRefGoogle Scholar
  69. Wang J, Wang X, Zhou Y, Du L, Wang Q (2010) Fumonisin detection and analysis of potential fumonisin-producing Fusarium spp. in asparagus (Asparagus officinalis L.) in Zhejiang Province of China. J Sci Food Agric 90:836–842PubMedGoogle Scholar
  70. Waśkiewicz A, Irzykowska L, Karolewski Z, Bocianowski J, Goliński P, Weber Z (2009) Mycotoxins biosynthesis by Fusarium oxysporum and F. proliferatum isolates of asparagus origin. J Plant Protect Res 49:369–372CrossRefGoogle Scholar
  71. Waśkiewicz A, Golinski P, Karolewski Z, Irzykowska L, Bocianowski J, Kostecki M, Weber Z (2010a) Formation of fumonisins and other secondary metabolites by Fusarium oxysporum and F. proliferatum: a comparative study. Food Addit Contam Part A Chem Anal Control Expo Risk Assess 27:608–615PubMedGoogle Scholar
  72. Waśkiewicz A, Irzykowska L, Bocianowski J, Karolewski Z, Kostecki M, Weber Z, Golinski P (2010b) Occurrence of Fusarium fungi and mycotoxins in marketable asparagus spears. Pol J Environ Stud 19:219–225Google Scholar
  73. Werner M, Irzykowska L (2007) The pathogenicity and DNA polymorphism of Fusarium oxysporum originating from Dianthus caryophyllus, Gypsophila spp. and soil. Phytopathol Pol 46:25–36Google Scholar
  74. Yun S-H, Arie T, Kaneko I, Yoder OC, Turgeon BG (2000) Molecular organization of mating type loci in heterothallic, homothallic, and asexual Gibberella/Fusarium species. Fungal Genet Biol 31:7–20PubMedCrossRefGoogle Scholar
  75. Zanotti MGS, de Queiroz MV, dos Santos JK, Rocha RB, de Barros EG, Araújo EF (2006) Analysis of genetic diversity of Fusarium oxysporum f. sp. phaseoli isolates, pathogenic and non-pathogenic to common bean (Phaseolus vulgaris L.). J Phytopathol 154:545–549CrossRefGoogle Scholar

Copyright information

© The Author(s) 2012

Authors and Affiliations

  • Lidia Irzykowska
    • 1
  • Jan Bocianowski
    • 2
  • Agnieszka Waśkiewicz
    • 3
  • Zbigniew Weber
    • 1
  • Zbigniew Karolewski
    • 1
  • Piotr Goliński
    • 3
  • Marian Kostecki
    • 3
  • Witold Irzykowski
    • 4
  1. 1.Department of PhytopathologyPoznan University of Life SciencesPoznańPoland
  2. 2.Department of Mathematical and Statistical MethodsPoznan University of Life SciencesPoznanPoland
  3. 3.Department of ChemistryPoznan University of Life SciencesPoznańPoland
  4. 4.Institute of Plant Genetics Polish Academy of SciencePoznańPoland

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