In Aspergilli, mycotoxin production and sporulation are governed, in part, by endogenous oxylipins (oxygenated, polyunsaturated fatty acids and metabolites derived therefrom). In Aspergillus nidulans, oxylipins are synthesized by the dioxygenase enzymes PpoA, PpoB and PpoC. Structurally similar oxylipins are synthesized in seeds via the action of lipoxygenase (LOX) enzymes. Previous reports have shown that exogenous application of seed oxylipins to Aspergillus cultures alters sporulation and mycotoxin production. Herein, we explored whether a plant oxylipin biosynthetic gene (ZmLOX3) could substitute functionally for A. nidulans ppo genes. We engineered ZmLOX3 into wild-type A. nidulans, and into a ΔppoAC strain that was reduced in production of oxylipins, conidia and the mycotoxin sterigmatocystin. ZmLOX3 expression increased production of conidia and sterigmatocystin in both backgrounds. We additionally explored whether A. nidulans oxylipins affect seed LOX gene expression during Aspergillus colonization. We observed that peanut seed pnlox2–3 expression was decreased when infected by A. nidulansΔppo mutants compared with infection by wild type. This result provides genetic evidence that fungal oxylipins are involved in plant LOX gene expression changes, leading to possible alterations in the fungal/host interaction. This report provides the first genetic evidence for reciprocal oxylipin cross-talk in the Aspergillus–seed pathosystem.
Hydroxylated C18 unsaturated fatty acids (which fall into the class of oxygenated, polyunsaturated fatty acids called oxylipins) are endogenously produced by Aspergilli and well studied in the genetic model, Aspergillus nidulans (Champe and El-Zayat, 1989; Mazur et al., 1990; 1991). These substances, called psi factor in A. nidulans (for precocious sexual inducer), alter the ratio of sexual : asexual spore production in this species (Champe et al., 1987; Champe and El-Zayat, 1989). Psi factor oxylipins have hormone-like characteristics: two of the components (psiA1 and psiB1) had opposing effects (Champe and El-Zayat, 1989), and psiA1 demonstrated bioactivity at concentrations as low as 0.2 μm (Champe and El-Zayat, 1989) – comparable to bioactive concentrations of trisporic acid, a fungal sex hormone of Mucor spp. (Gooday, 1983). Simultaneous deletion of the three dioxygenase genes (ppoA, ppoB and ppoC) of A nidulans severely reduced production of at least two oxylipin species comprising psi factor (Tsitsigiannis et al., 2005a), implicating these dioxygenases in oxylipin biosynthesis. Genetic studies showed that ppoA, ppoB and ppoC help regulate the asexual : sexual spore ratio (Tsitsigiannis et al., 2004a,b, 2005a). Oxylipins also govern secondary metabolite production: early work demonstrated that psi factor regulated production of an unidentified yellow pigment (Champe and El-Zayat, 1989); later genetic studies showed that ppoA, ppoB and ppoC govern production of the aflatoxin precursor, sterigmatocystin, the antibiotic penicillin (Tsitsigiannis and Keller, 2006), and an octaketide, shamixanthone (J. Frisvald and N.P. Keller, unpubl. data).
Sporulation and mycotoxin production by Aspergillus is affected not only by endogenous oxylipins, but also by plant-derived oxylipins. Linoleic and α-linolenic acid (18:3) can be converted to several oxylipin species by plant LOX enzymes (for review, see Liavonchanka and Feussner, 2006). These plant oxylipins are 13S-hydroperoxy linoleic (α-linolenic) acid (13S-HPODE/TE), and 9S-hydroperoxy linoleic (α-linolenic) acid (9S-HPODE/TE). 9S-HPODE/TE and 13S-HPODE/TE exhibited growth-independent effects on sporulation and mycotoxin production in Aspergillus spp. – filter disks soaked with 0.1 mg of 13S-HPODE in confluent plate cultures induced conidial development whereas 9S-HPODE induced ascospore production (Calvo et al., 1999). Exogenous application of pure 13S-HPODE to Aspergillus cultures repressed aflatoxin and sterigmatocystin gene expression by A. nidulans and Aspergillus parasiticus cultures, while pure 9S-HPODE had a positive effect (Burow et al., 1997). Reciprocally, Aspergillus colonization altered seed LOX gene expression, leading to changes in levels of bioactive oxylipins (Burow et al., 2000; Wilson et al., 2001; Tsitsigiannis et al., 2005b). The maize 9-LOX gene was upregulated and peanut 13-LOX genes were downregulated by Aspergillus infection of plant tissues (Wilson et al., 2001; Tsitsigiannis et al., 2005b). Taken together, these studies show that plant oxylipins directly or indirectly affect Aspergillus developmental processes, and that colonization by Aspergillus, in turn, alters plant oxylipin production. We postulated that these effects stem from the potential for plant oxylipins to mimic or interfere with biological activities of endogenous fungal oxylipins.
Based on the hypothesis that plant oxylipins are sensed by, and bioactive in Aspergillus, we performed genetic experiments to test whether plant oxylipin-generating dioxygenases might substitute functionally for fungal dioxygenases. A. nidulans deleted of the ppoA and ppoC genes (ΔppoAC mutants) produce reduced levels of at least two fungal oxylipins relative to the wild type (WT): the oleic acid-derived 8-hydroxy oleic acid or (9Z)-8-hydroxy-octadecenoic acid (8-HOE or psiBβ), and the linoleic acid-derived 8-hydroxy linoleic acid or (9Z,12Z)-8-hydroxy-octadecadienoic acid (8-HODE or psiBα) (Tsitsigiannis et al., 2004a). The ΔppoAC mutants are also unable to produce sterigmatocystin and generate fewer conidia than the WT. We reasoned that seed-Aspergillus cross-talk would be supported if a plant dioxygenase could restore a WT phenotype(s) to the ΔppoAC mutant. Simply growing the ΔppoAC mutant on seed did not restore the phenotype (Tsitsigiannis and Keller, 2006), and neither did applying crude WT A. nidulans psi extract (oxylipins) to A. nidulansΔppoAC cultures (data not shown). The latter observation implies that the ΔppoAC is a pleiotrophic mutant, compromised not only in its ability for oxylipin biosynthesis, but also for oxylipin perception/uptake. To circumvent this problem, we expressed a gene for a plant oxylipin-forming enzyme inside the fungal cells. We engineered into the ΔppoAC mutant a maize gene (ZmLOX3; formerly named cssap92). ZmLOX3 encodes an enzyme with 9-LOX activity, is expressed in maize embryo tissues and is upregulated during Aspergillus infection of maize seed (Wilson et al., 2001). We herein report the phenotypic consequences of expressing ZmLOX3 in WT and ΔppoAC genetic backgrounds of A. nidulans. We also report genetic evidence for the reciprocal interaction: colonization of peanut seeds with ppo mutants of A. nidulans resulted in decreased levels of plant LOX gene expression. To our knowledge, this is the first report providing genetic support for plant-Aspergillus oxylipin cross-talk.
Expression of maize ZmLOX3 in A. nidulans
To verify that the maize ZmLOX3 gene was expressed in A. nidulans, RNA was extracted and analysed by Northern hybridization. A single transcript of approximately 2.6 kb was expressed in all WT, ΔppoAC, and ΔppoB strains carrying the ZmLOX3 construct (Fig. 1 and M. Brodhagen and N. Keller, unpubl. data). Oxylipin effects on A. nidulans sporulation are known to be light-dependent (Tsitsigiannis et al., 2004a,b, 2005a). However, the gpdA promoter that we used to drive ZmLOX3 expression is governed by a circadian rhythm (Greene et al., 2003), and in the WT/ZmLOX3 and ΔppoAC/ZmLOX3 strains, ZmLOX3 expression was noticeably higher in dark-grown cultures (data not shown). Because this gene expression difference could confound our analysis of any potential light-dependent phenotypes, we limit our discussion of phenotypic differences among strains to those observed under a single light condition appropriate for each experiment.
Next, we verified that ZmLOX3 was expressed as active enzyme in the WT and ΔppoAC background. Lysates were obtained from fungal cultures that had been grown either in darkness or under continuous light. The lysates were incubated with linoleic acid, and subsequently extracted in order to analyse linoleic acid derivatives via high-performance liquid chromatography (HPLC) (Fig. 2). Linoleic acid was oxidized by an unknown enzyme in the WT and ΔppoAC strains into an almost equal ratio of 9- and 13-HODE, both occurring as racemates (equal R- to S-ratios). In contrast, the strains expressing the 9-LOX enzyme, ZmLOX3, showed a strong bias towards formation of 9S-HODE, indicating that the transgenic strains harbour 9-LOX activity. However, this was not reflected by the endogenous accumulation of 9-LOX-derived oxylipins (Fig. 3). We observed endogenous production of 9- and 13-HODE and 13-HOTE in all strains; however, their equal R- to S-ratios in all strains imply either that these products were not derived from LOX activity, or that ZmLOX3 derivatives of linoleic or α-linolenic acids were metabolized as earlier studies hinted (Tsitsigiannis et al., 2004a, 2005a). The presence of ZmLOX3 did not lead to significant differences in the amount of the native A. nidulans oxylipins, psiBβ (8-HOE) or psiBα (8-HODE) (data not shown).
Effect of maize ZmLOX3 on growth and spore viability of A. nidulans
We observed no differences between colony diameters of uniformly point-inoculated WT (RDIT9.32) and WT/ZmLOX3 (RDIT99.2) strains (P ≥ 0.05 for all time points, one-way anova; Fig. S1). However, at 5 days, ΔppoAC/ZmLOX3 (RDIT106.5) colonies were 8.6% smaller than ΔppoAC (RDIT54.7) colonies (P = 0.002; t-test; Fig. S1). Recently, examination of an Aspergillus fumigatus ppoC mutant revealed that endogenous oxylipins can affect spore structure and integrity (T. Dagenias, D. Chung and N.P. Keller, unpubl. data). However, we did not observe differences between ZmLOX-harbouring and near-isogenic strains in viability (measured by per cent germination) of ascospores (all P > 0.12; t-test; data not shown) or conidiospores (all P > 0.10; t-test; Fig. S2).
Expression of ZmLOX3 gene affected spore numbers
Earlier studies (Calvo et al., 1999) demonstrated that exogenous application of 9S-HPODE (the primary product of ZmLOX3; Wilson et al., 2001) altered the asexual to sexual spore ratio in a concentration-dependent manner: 1 mg on a filter disk applied to confluent agar plate cultures increased conidiation, while 0.5 mg increased ascospore production. Therefore, we examined both sexual and asexual spore production in strains expressing ZmLOX3.
Conidial production. For comparison of conidial production, strains were cultured as described in the Experimental procedures, under continuous light. ZmLOX3+ strains of A. nidulans produced more asexual spores than near-isogenic, LOX– strains in the WT and ΔppoAC background (Fig. 4A and C). This difference was not apparent at 3 days, but by 7 days, ZmLOX3 strains always accumulated more conidia than control strains (Fig. 4A and C).
Ascospore production. For comparison of ascospore production, strains were cultured as described in the Experimental procedures, under continuous darkness. WT/ZmLOX3 strains showed no significant difference in ascospore production at 5 or 7 days (Fig. 4B). In contrast, cultures harvested at 5 days revealed that ascospore production was lower in the ΔppoAC/ZmLOX3 strain (P < 0.05; t-test) than in the ΔppoAC isogenic control, but this reduction was remediated by the 7 day harvest (Fig. 4D).
Heterologous expression of plant ZmLOX3 gene slowed ascospore production but increased cleistothecial size
Delayed production of ascospores by ZmLOX3 strains in the ΔppoAC background (Fig. 4D) suggested that cleistothecial development might be altered in these strains. We therefore examined cultures for cleistothecial abundance, diameter and ascospore load. In the WT and ΔppoAC backgrounds, 7-day-old cleistothecia of ZmLOX3+ strains were larger than those of near-isogenic lines (Fig. 5A and B). These larger cleistothecia contained proportionally more ascospores (data not shown). However, there were fewer cleistothecia along a given transect (Fig. 5C). These opposing effects appear to negate one another, explaining why, despite larger cleistothecia (Fig. 5A and B), ZmLOX3+ strains do not accumulate more ascospores (Fig. 4B and D).
Heterologous expression of plant LOX genes increased sterigmatocystin production
In previous experiments, exogenous application of 9S-HPODE enhanced accumulation of a sterigmatocystin biosynthetic gene transcript (Burow et al., 1997). The 9-LOX encoded by ZmLOX3 produces mainly 9S-HPODE (Wilson et al., 2001). Therefore, we hypothesized that addition of the ZmLOX3 gene to sterigmatocystin-deficient ΔppoAC mutants would restore sterigmatocystin production. Expression of the ZmLOX3 gene increased sterigmatocystin production in the WT background (Fig. 6A) and, as predicted, ΔppoAC mutants expressing the ZmLOX3 gene were restored in sterigmatocystin production (Fig. 6B).
Effects of fungal ppo gene expression on plant LOX gene expression
Previous studies have indicated that oxylipins (of both plant and fungal origin) regulate Aspergillus developmental processes at the level of gene transcription (Burow et al., 1997; Tsitsigiannis et al., 2004b, 2005a, 2006). We wondered whether fungal oxylipins, conversely, could alter plant gene expression and participate in Aspergillus–seed cross-talk. The most direct method for measuring seeds' response to fungal oxylipins would be application of pure oxylipins to seeds followed by measurement of seed LOX gene expression. However, pure fungal oxylipins are not available, so we addressed the question in a more indirect manner. Because Aspergillus infection alters LOX gene expression in both corn (Wilson et al., 2001) and peanut (Burow et al., 2000; Tsitsigiannis et al., 2005b), we compared peanut LOX gene expression in peanut seeds infected with Aspergillus WT and oxylipin-reduced mutant strains. Here, we looked at the expression of two peanut LOX, PnLOX2 and PnLOX3 (whose nearly identical transcripts are indistinguishable in our study, and referred to jointly as pnlox2–3). We infected peanut seeds with A. nidulans WT and three near-isogenic Δppo mutant strains with deficiencies in production of one or more oxylipins (Tsitsigiannis et al., 2004a,b, 2005a). Quantitative RT-PCR confirmed that ppoA, ppoB and ppoC were expressed in A. nidulans during infection of peanut seeds (Fig. 7A). Notable are the higher relative abundances of ppoB and ppoC transcripts recovered from A. nidulans grown in vivo (Fig. 7A) as opposed to in vitro (Fig. 7B). Infection of seeds with mutant A. nidulans strains, defective in one or more ppo gene and resultant oxylipin production, resulted in less accumulation of the peanut 13-LOX transcript(s) pnlox2–3 compared with seeds inoculated with WT fungus (Fig. 7C). Expression of pnlox2–3 was lowest during infection by the ΔppoAC and ΔppoABC strains of A. nidulans.
Effects of ZmLOX3 in a ΔppoB background
Besides expressing ZmLOX3 in WT and ΔppoAC genetic backgrounds, we also successfully expressed ZmLOX3 in a ΔppoB background. With regard to the phenotypes discussed herein, the A. nidulansΔppoB mutant is opposite to the ΔppoAC mutant in that it overproduces conidia and sterigmatocystin (Tsitsigiannis et al., 2005a; Tsitsigiannis and Keller, 2006), and thus we expected little if any phenotype difference from the WT in a ΔppoB/ZmLOX3 strain. Indeed, in contrast to the results seen in the WT and ΔppoAC backgrounds, expression of ZmLOX3 in the ΔppoB background did not alter the phenotypic parameters that we measured in this study. ΔppoB/ZmLOX3 strains (RMLB2.01 and RMLB2.50) did not differ significantly from the near-isogenic strain, RDIT59.1, in colony diameter, spore viability and production, cleistothecial size or sterigmatocystin production (data not shown).
Although ZmLOX3 expression clearly conferred 9-LOX activity to fungal cell extracts after incubation with linoleic acid (Fig. 2A), analysis of lipid extracts from fungal mycelium did not indicate that endogenous, enzymatically produced 9S-HPODE accumulated to a higher degree in ZmLOX3-harbouring strains than in WT strains. Levels of linoleic acid were equivalent between ZmLOX3-expressing and near-isogenic strains (Fig. S5), and thus the ZmLOX3 substrate was not likely limiting. It is possible that the failure to observe high endogenous 9S-HPODE production in A. nidulans ZmLOX3-expressing mycelia is due to low in vivo activity or intracellular turnover of this heterologously expressed protein or its products. It also is possible that our analysis – performed on lipid extracts from bulk fungal mycelia grown in liquid culture for 72 h – may have missed early, transient or tissue-localized accumulation of this plant oxylipin. Lack of observation of enhanced 9S-HPODE in ZmLOX3 strains suggests tight control of endogenous oxylipins, both native and foreign. We did not observe changes in the fungal oxylipins 8-HODE or 8-HOE associated with ZmLOX3 expression. However, it is possible that ZmLOX3 affected sporulation and sterigmatocystin phenotypes indirectly by affecting levels of other oxylipins not monitored in this study. Certainly deletion or overexpression of specific ppo genes can lead to altered expression of other ppo genes and, ultimately, oxylipin production – presumably via regulatory feedback loops (Tsitsigiannis et al., 2004a, 2005a).
Plant and fungal oxylipins alter Aspergillus sporulation and secondary metabolism and ppo expression itself at the transcriptional level (Burow et al., 1997; Tsitsigiannis et al., 2004b, 2005a; Tsitsigiannis and Keller, 2006). To examine the converse situation, we examined whether Aspergillus ppo mutants altered in oxylipin production could alter plant LOX expression. When we compared pnlox2–3 expression in response to infection by Aspergillus WT and oxylipin-reduced (Δppo) mutant strains, we observed that colonization with A. nidulansΔppoB, ΔppoAC and ΔppoABC mutants all resulted in decreased accumulation of pnlox2–3 compared with the WT strain. One interpretation of this result is that fungal oxylipins are recognized by plant cells and induce changes in plant oxylipin gene expression. This interaction could result from direct sensing of A. nidulans oxylipins by peanut tissues, resulting in changes in pnlox2–3 expression. Another explanation is that peanut tissues altered pnlox2–3 expression in response to a non-oxylipin fungal product, the biosynthesis of which requires intact A. nidulans Ppo enzymes. Regardless of mechanism, this observation provides the first genetic evidence that fungal oxylipins mediate the response of peanut LOX expression to Aspergillus colonization, and suggests that oxylipin cross-talk in the seed-Aspergillus pathosystem may be reciprocal.
The reduced expression of pnlox2–3 in peanuts following colonization by Δppo mutants of A. nidulans is notable because pnlox2–3 expression is profoundly downregulated during colonization by the WT strain of a related fungus, A. flavus (Tsitsigiannis et al., 2005b). The current results suggest that Aspergillus ppo-derived oxylipins may ameliorate the repression of the peanut 13-LOX genes, pnlox2–3, to some degree. The ΔppoAC and ΔppoABC mutants are crippled in the colonization process (Tsitsigiannis and Keller, 2006); further studies should clarify whether this disability reflects a loss in fungal virulence as a result of the pleiotrophic ppo mutant phenotypes, or an effect on host defense responses including LOX gene expression changes, or both.
The molecular mechanism for oxylipin cross-talk is, as yet, unknown. One hypothesis, based on known mechanisms in mammalian cells, is that oxylipins bind cell-surface receptors (Brodhagen and Keller, 2006), but whether 9S-HPODE is taken up by tissues or sensed by fungal cell-surface receptors remains to be demonstrated. Presumably, in our studies, the ZmLOX3 oxylipin product(s) were produced in the cytoplasm, and thus their effects on fungal physiology are more likely to mimic a scenario in which 9S-HPODE is taken up by the fungus rather than perceived by cell-surface proteins. Also still unknown is the bioactive concentration of 9S-HPODE required to elicit physiological responses by fungal tissues. Previous studies showed that effects of exogenous 9S-HPODE varied, depending on the 9S-HPODE concentration (Calvo et al., 1999).
In summary, we have shown that expression of the maize ZmLOX3 transgene elicits changes in the developmental programme of A. nidulans that mimic developmental changes governed by endogenous fungal oxylipins. Further, we present genetic evidence that Aspergillus oxylipins contribute to alterations in peanut LOX gene expression that occur upon fungal colonization (Tsitsigiannis et al., 2005b), bolstering the idea that Aspergillus–seed oxylipin cross-talk is a two-way conversation. Ultimately, understanding the mechanisms of Aspergillus–seed oxylipin cross-talk should lead to novel methods for control of mycotoxin production by Aspergillus spp. Taking a broader view, the widespread occurrence of oxylipins (Brodowsky and Oliw, 1993; Su et al., 1995; Nakayama et al., 1996; Bareetseng et al., 2004) and LOX genes (Tsitsigiannis et al., 2005a) among filamentous fungi genera suggests a conserved role among life cycles. It is possible that cross-kingdom oxylipin signalling may be a phenomenon common to numerous host–microbe interactions, pathogenic and otherwise.
Fungal strains and growth conditions
Aspergillus nidulans strains used in this study are listed in Table 1. Strains were cultured at 37°C on glucose minimal medium (GMM) (Calvo et al., 2001) containing appropriate supplements for auxotrophies. Cultures were grown inside cardboard boxes to achieve continuous dark, or 50 cm below General Electric 15-watt-broad spectrum fluorescent light bulbs (F15T12CW) to achieve continuous white light. For RNA quantification and oxylipin analysis, A. nidulans cultures were grown by inoculating GMM broth with 106 conidia per ml, aliquotting 30 ml of this suspension into individual Petri plates, and incubating suspensions at 37°C without shaking. Cultures were harvested at 72 h and lyophilized. Mycelial growth, spore production and sterigmatocystin production were measured from cultures grown on GMM agar. To assess growth, we point-inoculated 103 conidia onto the centre of 30 ml GMM agar plates and measured colony diameters over time. For analyses of spore and sterigmatocystin production, 25 ml of solid (1.5%) GMM agar was overlaid with 5 ml of a soft (0.7%) agar suspension containing 106 conidia of each appropriate strain. Plates were incubated at 37°C.
Table 1. A. nidulans strains used in this study.
veA indicates the WT veA allele (as opposed to veA1).
Plasmid construction, fungal transformation and fungal strain generation
Construction of a vector for targeting genes to the pyroA locus of A. nidulans. A truncated 1790 kb fragment of the 3′ portion of the A. nidulans pyroA gene (PstI-pyroA3/4-EagI), which spans the site of the pyroA4 mutation, was amplified via PCR from plasmid p14 (Osmani et al., 1999), and ligated between the PstI and EagI sites of pBluescript II SK (–) to create plasmid pTMH76.2. Then, a 1.8 kb HindIII-pyroA3/4-PstI fragment was released from pTMH76.2, blunt-ended with the Klenow DNA polymerase fragment (New England Biolabs) and ligated into the SmaI site of pBluescript II SK (–) to create plasmid pTMH77.9. Finally, a 1.8 kb EcoRI-pyroA3/4- EcoRI fragment was released from pTMH77.9, and ligated into the EcoRI site of pTMH44.2 (McDonald et al., 2005) which contains the constitutive A. nidulans glyceraldehyde-3-phosphate dehydrogenase (gpdA) promoter and a transcriptional terminator from the trpC gene of the tryptophan biosynthetic pathway (Punt et al., 1991). This final transformation vector was named pTMH78.1 (Fig. S3).
Construction of a vector for targeting ZmLOX3 to A. nidulans pyroA. Plasmid pMLB1.7 (T. Hammond and N.P. Keller, unpubl. data) harbours cDNA of the maize 9-LOX gene ZmLOX3 (previously named cssap92; GenBank accession number AF329371). A fragment extending from the ZmLOX3 start codon to 361 bp downstream of the stop codon was amplified via PCR from pMLB1.7 using the modified primers Corn-lox-F1-NcoI (5′-ATTCCTGCAGCCCCATGGTGAGCGG-3′) and Corn-lox-R1-BamHI (5′-TGTTGTGATCGAGGATCCCTGCGAA-3′) and digested with NcoI and BamHI. The resulting 2958 bp NcoI–BamHI fragment was subcloned into the corresponding sites of pTMH78.1 (described above) between the gpdA promoter and the trpC transcriptional terminator.
Fungal transformations and sexual crosses. The resultant plasmid, pDWC2.2, was used to transform RDIT55.24 to pyridoxine prototrophy. Fungal transformations were performed as previously described (Miller et al., 1985). Transformants were screened for presence of the gpdA(p)::ZmLOX3 construct by PCR. Southern analysis confirmed single-genome insertions of pDWC2.2 at the A. nidulans pyroA locus (Fig. S4 and data not shown). A single transformant, TDWC 2.15, was sexually crossed as previously described (Pontecorvo et al., 1953) with RDIT55.37 to generate progeny bearing the gpdA(p)::ZmLOX3 allele in a prototrophic background with a WT oxylipin profile (strains RDIT98.5 and RDIT99.2, referred to as WT/ZmLOX3. TDWC2.15 was also crossed with the oxylipin-deficient strain RDIT62.15 (ΔppoA; ΔppoB; ΔppoC), generating as progeny strains RDIT106.5 and RDIT106.6, referred to as ΔppoAC/ZmLOX3). Finally, TDWC2.15 was crossed with the oxylipin-reduced strains RDIT64.23 and RDIT64.29 (ΔppoB), generating among their progeny strains RMLB2.01 and RMLB2.50, referred to as ΔppoB/ZmLOX3. Recombinant strains were verified by PCR and Southern analysis to confirm ppo allele genotype and the presence of gpdA(p)::ZmLOX3 (data not shown).
The WT and WT/ZmLOX3 strains are herein referred to as ‘near-isogenic’: although they differ in that the WT strain does not bear the ZmLOX3 gene at the pyroA locus, they were derived by crossing genetically related strains, and all auxotrophies in the parental strains have been repaired by sexual recombination. The same is true for the ΔppoAC and ΔppoAC/ZmLOX3 strains, and the ΔppoB and ΔppoB/ZmLOX3 strains.
Gene expression analysis
Expression of ZmLOX3. Total RNA was extracted from lyophilized mycelia using TRIzol reagent (Invitrogen) according to manufacturer's recommendations. Approximately 25 μg per sample of total RNA was separated on a 1.2% agarose/1.5% formaldehyde gel, and transferred to a Hybond-XL membrane (GE Healthcare Bio-Sciences Corp.). Probes for Northern analysis of ZmLOX3 were amplified via PCR from pDWC2.2 using the primers Corn lox F3 (5′-TCTTCTTCTCCAACGATACGTACCT-3′) and Corn lox R3 (5′-AGCAGCATGTAGGGGAACTCGGCGG-3′). PCR products were gel-purified, radiolabeled, hybridized to membranes and visualized using a Phosphorimager SI equipped with ImageQuaNT software (Molecular Dynamics, Inc.). RNA was extracted from duplicate or triplicate cultures, and gene expression for all strains was confirmed by two independent experiments.
Expression of peanut PnLOX2-3 and endogenous Aspergillus ppo genes. For peanut infection studies, seed infections and RNA extractions were carried out as previously described (Tsitsigiannis et al., 2005b). The peanut LOX-encoding genes, PnLOX2 and PnLOX3, are nearly identical to one another in nucleic acid sequence, and we were not able to design primers capable of distinguishing these genes in our assay. Therefore, transcripts from both genes were measured jointly by amplification of cDNA using the primers RT-PnLOX2-3-F1 and RT-PnLOX2-3-R1 (Tsitsigiannis et al., 2005b), and the transcripts are referred to as PnLOX2-3. Expression of PnLOX2-3 was normalized to the peanut actin-DF gene. Design and analysis of experiments to quantify transcripts of PnLOX2-3 and actin-DF via quantitative RT-PCR are described in detail elsewhere (Tsitsigiannis et al., 2005b). Transcripts of A. nidulans genes were measured by amplification of cDNA using the primers An-ppoA-F1-539 (CTGGTGTTGTTGTGGAAGAAG) and An-ppoA-R1-688 (GGTGCTTCGGAGTGTAGTC) for ppoA, An-ppoB-F1-176 (CTGAAGCATTGGAAGCACTC) and An-ppoB-R1-319 (CGTCTGAGAGAATGGCAAAC) for ppoB, An-ppoC-F1-421 (CAGATCGTTAGCAGCCTTC) and An-ppoC-R1-569 (TTGTTAGAACCATCAGCAGAG) for ppoC, and An-tubulin-F1-1000 (CAGAGCAAGAACCAGTCCTAC) and An-tubulin-R1-1147 (CACCGACACGCTTGAAGAG) for tubulin. Mycelium used for RNA extraction from A. nidulans was obtained by inoculating GMM broth with 106 conidia per ml and shaking at 300 r.p.m. in darkness and at 37°C for 72 h.
Analysis of ZmLOX enzymatic activity A. nidulans
A total of 1.5 g of fungal tissue (harvested from different growth stages) was homogenized under liquid nitrogen, vortexed with 3 ml of lysis buffer [50 mM Tris/HCl, pH 8.0, 300 mM NaCl, 0.1% (v/v) Tween20, 200 μm PMFS], and incubated at 4°C for 40 min. The lysate was centrifuged for 10 min at 5000 g. A total of 900 μl of the supernatant was incubated with 250 μg of linoleic acid for 30 min at room temperature (RT). The reaction was stopped by adding 100 μl of glacial acid, and fatty acids were extracted according to the method of Bligh and Dyer (1959). The fatty acid-harbouring organic phase was further analysed as described below for the determination of hydroxy fatty acids.
Hydroxy fatty acid analysis
Hydroxy fatty acids were analysed as described previously (Göbel et al., 2002; Weichert et al., 2002), but with some modifications. About 0.5 g of lyophilized, frozen fungal tissue was homogenized in 20 ml extraction medium [hexane : 2-propanol, 3:2 (v/v), with 0.0025% (w/v) butylated hydroxytoluene, including (6Z,9Z,11E,13S)-13-hydroxy-6,9,11-octadecatrienoic acid (13γ-HOTE) as internal standard] with an Ultra Turrax (13 000 r.p.m) under streaming argon for 45 s. The extract was shaken at 4°C for 10 min and then centrifuged at 3200 g at 4°C for 10 min. To the clear upper phase, a 6.7% (w/v) solution of potassium sulphate was added up to a volume of 32.5 ml. After vigorous shaking at 4°C for 10 min, the extract was centrifuged at 3200 g at 4°C for 10 min. Subsequently, the upper hexane-rich layer was dried under streaming nitrogen. The remaining lipids were re-dissolved in 0.2 ml methanol and stored under argon atmosphere at −20°C until further use.
For analysis of the putative LOX-derived hydroxides, 0.16 ml of the sample was dried under streaming nitrogen, re-dissolved in 0.08 ml of methanol : water : acetic acid (80:20:0.1, v/v/v) and subjected to HPLC analysis using an 1100 HPLC system (Agilent) coupled to a diode array detector. For the detection of hydroxy fatty acids, absorbance of the conjugated diene system at 234 nm was recorded. First, hydroxy fatty acids were purified by reverse-phase (RP) HPLC. This was performed on an ET250/2 Nucleosil 120-5 C18 column (250 × 2.1 mm, 5 μm particle size; Macherey and Nagel) with a methanol : water : acetic acid (80:20:0.1, v/v/v) solvent system at a flow rate of 0.18 ml min−1. Hydroxy fatty acid containing fractions were collected. To analyse the regio-isomers of hydroxy fatty acids, purified by straight-phase (SP) HPLC was performed on a Zorbax Rx-SIL column (150 × 2.1 mm, 5 μm particle size; Agilent) with n-hexane : 2-propanol : acetic acid (100:1:0.1, v/v/v) as a solvent system at a flow rate of 0.2 ml min−1. Fractions corresponding to the regio-isomers were collected and subjected to chiral phase HPLC analysis to measure enantiomeric composition. This analysis was carried out on a Chiral OD-H column (150 × 2.1 mm, 5 μm particle size; Baker) with n-hexane : 2-propanol : acetic acid (100:5:0.1, v/v/v) as a solvent system at a flow rate of 0.1 ml min−1. For quantification, calibration curves (5-point measurements) for 13-HODE and 13-HOTE were established. 13γ-HOTE was used as internal standard to determine the recovery of the hydroxy fatty acids.
For analysis of the dioxygenase-derived hydroxides, 0.36 ml of methanol and 6.5 μl of 2 M trimethylsilyldiazomethane (in hexane; Sigma) were added to the remaining 0.04 ml of the sample. The sample was shaken for 30 min, and 0.2 μl of acetic acid was added to degrade the remaining trimethylsilyldiazomethane. The sample was dried under streaming nitrogen, the remaining hydroxy fatty acid methyl esters were re-dissolved in 0.08 ml of methanol : water : acetic acid (80:20:0.1, v/v/v), and were purified by reverse-phase (RP)-HPLC as described above. Hydroxy fatty acid methyl ester containing fractions were collected, dried under streaming nitrogen and re-dissolved in 3 μl of acetonitrile. After adding 1 μl of N,O-bis(trimethylsilyl)trifluoroacetamide for derivatization, sample was subjected to gas chromatography/mass spectrometry (GC/MS) using a 5973 Network mass selective detector connected to a 6890 gas chromatograph (Agilent) equipped with a capillary DB-23 column (30 m × 0.25 mm; 0.25 μm coating thickness; J and W Scientific). Helium was used as a carrier gas at a flow rate of 1 ml min−1. The temperature gradient was as follows: 150°C for 1 min, 150–200°C at 4 K min−1, 200–250°C at 5 K min−1 and 250°C for 10 min. Electron energy of 70 eV, an ion source temperature of 230°C, and a temperature of 260°C for the transfer line was used. For quantification, the ion m/z 73 was used and calibration curves (5-point measurements) for methyl ricinoleate were established. 13γ-HOTE was used as internal standard to determine the recovery of the hydroxy fatty acids.
Growth and spore viability analyses
To assess growth, we compared colony diameters of ZmLOX and near-isogenic, ZmLOX- strains that had been point-inoculated with 103 conidia onto 30 ml of GMM agar daily over 5 days. To assess spore viability, conidia were harvested by submerging 5-day-old cultures in 0.01% Tween 80, and scrubbing with a sterile bent glass rod. Ascospores were harvested by rolling cleistothecia on 3% water agar to remove mycelia and conidiospores, and crushing in 10 μl of 0.01% Tween 80. Spores were enumerated using a haemacytometer at 400×. Using the mean of four separate spore counts, spore suspensions were diluted in 0.01% Tween 80 to approximate 50 spores in 50 μl, spread evenly onto 30 ml GMM agar plates, and incubated for 1–3 days at 37°C in darkness. Individual colonies (representing single germinated spores) were enumerated.
Spore enumeration and sterigmatocystin analysis
A total of 106 conidia of each appropriate strain were overlaid evenly on GMM agar as described above and incubated at 37°C. On days 3, 5 or 7, three 12.5 mm cores from each plate were removed and homogenized in 4 ml of 0.01% Tween 80. Ten μL aliquots were diluted and spores enumerated using a haemacytometer at 400×. Four mL of CHCl3 were added to the remaining homogenate of 7 day cultures, and samples were vortexed for 30 s, extracted for 1 h at RT, and vortexed again prior to centrifugation at 2000 r.p.m. for 10 min. The organic phase was transferred to eight dram glass vials and allowed to evaporate to dryness at RT. Sample residues were then resuspended in 100 μl CHCl3, and 20 μl of each sample was separated via thin-layer chromatography (TLC) using polyester-backed, 250 μm silica gel plates (Whatman) using 4:1 (v/v) hexane : ethyl acetate as a mobile phase. TLC plates were sprayed with AlCl3 (15% in 95% EtOH) to enhance sterigmatocystin fluorescence, baked at 65°C for 10 min, and visualized under long-wave (365 nm) UV light (Stack and Rodricks 1971). Spot intensity of sterigmatocystin on TLC plates was quantified using a TLC Scanner II densitometer, equipped with CATS (version 3.20) TLC software (CAMAG).
For spore production data, we present the means of five replicate cultures. Sterigmatocystin production was compared across three to five replicate cultures. Each experiment was performed at least twice; and the results from a representative experiment are presented. After initial experiments that demonstrated similar sporulation and secondary metabolite phenotypes for WT/ZmLOX3 strains (RDIT98.5 and RDIT99.2), and for ΔppoAC/ZmLOX3 strains (RDIT106.5 and 106.6), further experiments were conducted with only RDIT99.2 and RDIT106.5 as representative of ZmLOX3+ genotypes.
Treatment means were compared using an analysis of variance (Fisher's LSD) or a Student's t-test. Two-tailed P-values are reported. Statistical analyses were performed using JMP, version 3 (SAS Institute).
The authors gratefully acknowledge financial support for M.B. from an NIEHS NRSA training grant through the Molecular and Environmental Toxicology Center at the University of Wisconsin and funding from NSF IOB-0544428, subagreement S060039 to N.P.K. We are grateful for technical assistance by Theres Riemekasten (Goettingen), DaWoon Chung (Madison) and Meghan Walters (Madison).