Research Article

Transcription Factor Amr1 Induces Melanin Biosynthesis and Suppresses Virulence in Alternaria brassicicola

  • Yangrae Cho mail,

    Affiliation: Plant and Environmental Protection Sciences, University of Hawaii at Manoa, Honolulu, Hawaii, United States of America

  • Akhil Srivastava,

    Affiliation: Plant and Environmental Protection Sciences, University of Hawaii at Manoa, Honolulu, Hawaii, United States of America

  • Robin A. Ohm,

    Affiliation: US DOE Joint Genome Institute, Walnut Creek, California, United States of America

  • Christopher B. Lawrence,

    Affiliation: Virginia Bioinformatics Institute and Department of Biological Sciences, Virginia Tech, Blacksburg, Virginia, United States of America

  • Koon-Hui Wang,

    Affiliation: Plant and Environmental Protection Sciences, University of Hawaii at Manoa, Honolulu, Hawaii, United States of America

  • Igor V. Grigoriev,

    Affiliation: US DOE Joint Genome Institute, Walnut Creek, California, United States of America

  • Sharadchandra P. Marahatta

    Affiliation: Plant and Environmental Protection Sciences, University of Hawaii at Manoa, Honolulu, Hawaii, United States of America

    Current address: Science and Math Division, Kaua'i Community College, Lihu'e, Hawaii, United States of America

  • Published: October 25, 2012
  • DOI: 10.1371/journal.ppat.1002974


Alternaria brassicicola is a successful saprophyte and necrotrophic plant pathogen. Several A. brassicicola genes have been characterized as affecting pathogenesis of Brassica species. To study regulatory mechanisms of pathogenesis, we mined 421 genes in silico encoding putative transcription factors in a machine-annotated, draft genome sequence of A. brassicicola. In this study, targeted gene disruption mutants for 117 of the transcription factor genes were produced and screened. Three of these genes were associated with pathogenesis. Disruption mutants of one gene (AbPacC) were nonpathogenic and another gene (AbVf8) caused lesions less than half the diameter of wild-type lesions. Unexpectedly, mutants of the third gene, Amr1, caused lesions with a two-fold larger diameter than the wild type and complementation mutants. Amr1 is a homolog of Cmr1, a transcription factor that regulates melanin biosynthesis in several fungi. We created gene deletion mutants of Δamr1 and characterized their phenotypes. The Δamr1 mutants used pectin as a carbon source more efficiently than the wild type, were melanin-deficient, and more sensitive to UV light and glucanase digestion. The AMR1 protein was localized in the nuclei of hyphae and in highly melanized conidia during the late stage of plant pathogenesis. RNA-seq analysis revealed that three genes in the melanin biosynthesis pathway, along with the deleted Amr1 gene, were expressed at low levels in the mutants. In contrast, many hydrolytic enzyme-coding genes were expressed at higher levels in the mutants than in the wild type during pathogenesis. The results of this study suggested that a gene important for survival in nature negatively affected virulence, probably by a less efficient use of plant cell-wall materials. We speculate that the functions of the Amr1 gene are important to the success of A. brassicicola as a competitive saprophyte and plant parasite.

Author Summary

Induction of cell wall-degrading enzymes, secretion of toxins, enforcement of fungal cell wall architecture, and detoxification of host defense molecules are important or essential for the fungal infection of plants. Genes important for each of these functions have been identified in various fungal species. Understanding how these genes are coordinately regulated and how many regulators are involved in the process is a challenge. We recently discovered a transcription factor gene, AbVf19, which positively regulates hydrolytic enzyme-coding genes. Here, we report on another transcription factor gene, Amr1, which negatively regulates a subset of these genes during late-stage pathogenesis and positively regulates melanin biosynthesis during conidiogenesis. This study adds another dimension to the complex regulation and overall importance of hydrolytic enzyme genes during plant pathogenesis by necrotrophic fungi. In addition, this study provides an example on the evolutionary implication of virulence in the necrotrophic fungus, A. brassicicola. One transcription factor essential for long-term survival because of its role in melanin biosynthesis is used instead to suppress virulence. We speculate that the suppressive functions of Amr1 contribute to the specialized adaptation of A. brassicicola as an efficient and successful facultative parasite.


Alternaria brassicicola is the causal agent of black spot disease of cultivated brassicas (e.g. cabbage, canola, mustard). Pathogenesis in necrotrophic fungi is generally described as a two-step process: killing host cells directly (necrosis) or inducing programmed cell death with toxins, then decomposing host tissues with cell wall-degrading enzymes. The importance of toxins in disease development by other necrotrophs has been clearly demonstrated [1], [2], [3], [4]. Several A. alternata pathotypes produce toxic, host-specific secondary metabolites that are essential for pathogenicity [5], [6], [7], [8]. Our knowledge of toxins produced by A. brassicicola is currently limited. In recent studies, Brassicicolin A emerged as the most selective phytotoxic metabolite produced in liquid cultures of A. brassicicola [9]. The genes responsible for biosynthesis of this toxin, however, have yet to be determined. Diterpenoid toxins called brassicenes have also been described from this fungus and linked to gene clusters in the A. brassicicola genome [10]. In addition, a cluster of five genes responsible for synthesis of the secondary metabolite depudecin, a histone deacetylation inhibitor was recently reported [11]. Mutation of the five genes abolished depudecin synthesis and caused only a small (10%) reduction in virulence compared to wild-type A. brassicicola. A weak protein toxin was also reported [12], [13] but the gene or genes responsible for its production have yet to be identified.

In addition to toxins, several A. brassicicola genes have been linked to pathogenesis [14], [15], [16], [17], [18], [19], [20]. These genes are involved in iron uptake, cell wall integrity, peroxisome-mediated redox homeostasis, hyphal fusion, hydrolytic enzymes, and signal transduction. All mutants of the pathogenesis-related genes showed either a reduction in virulence or loss of pathogenicity. For example, mutants of a mitogen-activated protein (MAP) kinase gene, Amk1, and its downstream transcription factor-coding gene, AbSte12, were nonpathogenic. The mutants of either gene also showed slow vegetative growth and impaired conidium maturation.

Bioinformatics analysis of the genome sequence of A. brassicicola identified many candidate genes important for pathogenesis. They included several genes that are essential for melanin synthesis. Alternaria species produce 1,8-dihydroxynaphthalene (1,8-DHN) melanin, like various other filamentous fungi [21], [22], [23]. It is heavily concentrated in the primary cell walls of conidia and in their septa [24]. Melanin is a ubiquitous pigment that plays an important role in protecting fungi from the damaging effects of environmental stress and so may be considered an advantageous adaptation. It increases the tolerance of fungi to UV irradiation [25], [26], [27], enzymatic lysis [28], and extreme temperatures [26], [29]. Melanin is also required for the mechanical penetration of host plants by other phytopathogenic fungi, such as Magnaporthe grisea and Colletotrichum lagenarium [30].

Synthesis of 1,8-DHN melanin requires polyketide synthase (Pks1 in Bipolaris oryzae and Pks18 in Cochliobolus heterostrophus), T4HN reductase (Brn2), scytalone dehydratase (Scd1), and T3HN reductase (Brn1 or Thr1) genes identified in B. oryzae [31], Col. lagenarium [32], and Coc.heterostrophus [21], [33]. Expression of these genes is mainly regulated by the Cmr1 transcription factor gene in Col. lagenarium. Cmr1 homologs are regulators of the primary melanin biosynthesis pathway in several plant pathogenic fungi but virulence is not affected by the gene mutation in M. grisea, Col. lagenarium, or Coc. heterostrophus [21], [22].

For all pathogenesis-associated genes in A. brassicicola studied to date, their mutants have been either nonpathogenic, or less virulent than the wild type. In this study, we determined that mutants of the Cmr1 homolog in A. brassicicola (Amr1) were melanin-deficient but more virulent than wild-type A. brassicicola. We defined virulence as quantitative host plant damage as measured by lesion diameter. The increase in virulence of the Δamr1 mutants suggested that the loss of the gene was beneficial to pathogenesis. We tested three research questions: whether melanin is important for pathogenesis, if mutants efficiently neutralize host defense chemicals, or if the mutants more efficiently utilize plant cell wall-associated materials such as pectin. Our study provides an example of a transcription factor gene that is important for survival in nature negatively regulates virulence in a necrotrophic plant pathogen. We speculate that the negative regulation of virulence by the Amr1 gene contributed to A. brassicicola's efficiency as a facultative plant pathogen while retaining characteristics of a robust saprophyte.


Identification of putative transcription factor genes

The genome sequence of A. brassicicola has been determined by Washington University School of Medicine, Genome Sequencing Center, and is publicly available from DDBJ/EMBL/GenBank under accession ACIW00000000. In addition, the annotated genome is available in the context of other sequenced Dothideomycetes genomes through the interactive JGI fungal portal MycoCosm [34] at The A. brassicicola genome size is approximately 31.9 Mb, smaller than the average genome size of other dothideomycete genomes [35]. There are 10, 688 predicted genes in the assembled genome. We predicted 421 genes encoding putative transcription factors in the draft genome of A. brassicicola using Pfam searches [36]. These transcription factors excluded the core proteins necessary for the formation of transcription initiation complexes and RNA polymerases. We classified the transcription factors in 13 categories based on their functional domains (Table 1). Members of the largest group (221 of 421) contain at least one zinc-finger domain. The second largest group (72 of 421) contains fungal-specific transcription factor domains.


Table 1. Summary of Alternaria brassicicola transcription factor domains based on Pfam scans.


Discovery of pathogenesis-associated transcription factors

We previously created targeted deletion mutants for 22 transcription factor and signaling related-genes and reported the association of AbSte12, AbPro1, and AbVf19 transcription factors with pathogenesis [15], [20]. In this present study, we generated targeted gene disruption mutants for an additional 117 transcription factor genes (Figure S1). All mutants corresponding to 109 of the 117 genes caused disease symptoms on host plants with lesion sizes similar to those caused by the wild type in multiple pathogenicity assays. Mutants of eight of 117 genes showed unusual changes in virulence. Among the eight mutants, five mutants also grew very slowly on nutrient rich medium. These mutants either failed to cause disease symptoms or their lesions did not expand beyond the initial infection site (Table S1). Gene disruption mutants of the AbPacC transcription factor (abpacc) were nonpathogenic (Table 2), unlike the severely reduced virulence in the loss-of-function mutants of its homologs in Col. acutatum [37] and Sclerotinia sclerotiorum [38] or no changes in Ustilago maydis [39]. Mutants corresponding to two other genes (Amr1and AbVf8) grew normally on nutrient rich media, were novel factors associated with pathogenesis, and are described further in this study.


Table 2. Loss of pathogenicity of the abpacc mutant and decreased virulence of Δabvf8 compared to wild-type Alternaria brassicicola.


The Amr1 gene

In contrast to the loss-of-function mutants of other virulence genes studied thus far in this pathosystem, Amr1 disruption mutants (amr1) caused larger lesions than wild-type A. brassicicola when green cabbage (Brassica oleracea) leaves were inoculated with the same number of conidia (data not shown, see below for detailed study). The predicted Amr1 gene encodes a protein of 997 amino acids with two C2H2-zinc finger motifs and one fungal-specific Zn(II) binuclear motif. Of the 997 predicted amino acids, 869 (87%) were identical (E-value = 0) to the transcription factor Cmr1 in the taxonomically closely related dothideomycete fungus, Coc. heterostrophus [21]. We named this Cmr1 homolog in A. brassicicola, Amr1 (Alternaria melanin regulation, GenBank accession number: JF487829). Only one copy of the gene was present in the draft genome sequence of A. brassicicola (

Replacement mutants of the Amr1 gene

In order to confirm our results with gene disruption mutants and to perform additional studies, deletion mutants of Amr1amr1) were produced by replacing the coding region with either a Hygromycin B (HygB) resistance cassette, or a green fluorescent protein (GFP) coding sequence [40] plus a HygB resistance cassette. Southern hybridization confirmed that the Amr1 gene was absent from nine melanin-deficient transformants but still present in the eight transformants producing melanin (Figure 1). We replaced the Amr1 coding region with a single copy of the HygB resistance cassette in Δamr1-4 and three Δamr1:Amr1p-GFP mutants (d4, D1, D2, and D3 in Figure 1). The replacement construct containing the GFP and HygB resistance cassette was designed so the GFP gene would be regulated by native promoter elements of the Amr1 gene in the replacement mutants (Δamr1:Amr1p-GFP) (Figure 1B). We complemented Δamr1-5 mutant with either the wild-type allele or a chimeric construct with the TrpC promoter to constitutively express the wild-type Amr1 gene.


Figure 1. Verification of Δamr1 deletion mutants.

A. Replacement of the Amr1 coding region with the selectable marker, Hygromycin B transferase (HygB) resistance cassette. On the HygB-probed blot, the expected 4.5 Kb band indicates a single copy insertion of the Hyg B resistance cassette in lane d4. Additional bands in the other lanes suggest multiple insertions of the cassette. B. Replacement of the Amr1 coding region with a green fluorescent protein (GFP) coding region and a HygB cassette. The upper panel of both A and B is a schematic diagram of wild-type (wt) and mutant loci and the lower panel consists of five Southern blots showing loss of the Amr1 gene in selected mutants. Dots (•) indicate lanes of mutants used in this study. Probe regions are marked by Ph and Pa. Abbreviations: E = EcoRI enzyme digestion site. d1 = Δamr1-1, d4 = Δamr1-4, d5 = Δamr1-5, and D3 = Δamr1:Amr1p-GFP. e1-e5, E1, E3, and E4 are ectopic insertion mutants.


Increased virulence of the Δamr1 mutants

Because the increase in virulence in A. brassicicola was unusual and unprecedented, we performed additional pathogenicity assays with four strains of gene deletion mutants (Δamr1-1, Δamr1-4, Δamr1-5, and Δamr1:Amr1p-GFP). All lesions caused by the mutants were 30–80% larger in diameter than the wild type in assays with 5- to-7-week-old plants (Table 3). Generally, wild-type A. brassicicola caused larger lesions on 5-week old plants than on 7-week-old plants and on older leaves compared to younger leaves. Interestingly, all Δamr1-type mutants produced consistent lesion sizes regardless of plant age. Thus, the relative size of lesions caused by the mutants compared to the wild type was greater on 7-week-old plants than on 5-week-old plants and on younger leaves than on older leaves (Figure 2A and Table 3). The difference in lesion size was similar between assays on whole plants and assays on detached leaves, so for convenience we performed a subsequent assay on the detached leaves of 8-week-old plants. In this assay, we compared lesion sizes caused by a gene deletion mutant (Δamr1), one strain from each of the two complemented mutants (Δamr1:Amr1 and Δamr1:TrpCp-Amr1), and the wild type. Lesions caused by the deletion mutant (Δamr1) were 2-fold greater on average than those made by the wild type or both complemented mutants (Δamr1:Amr1 and Δamr1:TrpCp-Amr1) (Figure 2E and Table 4).


Figure 2. Increased virulence of Δamr1 mutants on individual host plants (Brassica oleracea).

A. Lesions on B. oleracea 5 days after inoculation with ~2,000 conidia of the wild type, a Δamr1 mutant, and three complemented mutants with two types of constructs. B. Lesions produced by ~440 and ~400 conidia respectively of the wild type (left) and a Δamr1 mutant (right). C. Lesions produced by ~330 and ~300 conidia respectively of the wild-type (left) and a Δamr1 mutant (right). The three images in panels A–C are replicate experiments with the same pattern of inoculation. Note that the size of lesions caused by the mutant is similar on all leaves, while lesions caused by the wild type are smaller on young leaves than on old leaves. Δamr1-1 and Δamr1-4 produced similar results. D. Pathogenicity assay results for three mutants (Δamr1-1, Δamr1-4, and Δamr1-5) compared to the wild type. E. Pathogenicity assay results with average lesion diameter (mm) and standard deviation. Abbreviations: wt = wild-type A. brassicicola; Δamr1 = Amr1 deletion mutant; compl = mutant complemented with a native allele of the Amr1 gene (Δamr1:Amr1); cons = mutant complemented with chimeric constructs of the TrpC promoter and a native allele of the Amr1 coding region (Δamr1:TrpCp-Amr1).


Table 3. Increased virulence of four strains of Δamr1 deletion mutants compared to wild-type Alternaria brassicicola.


Table 4. Effect of the Amr1 gene on lesion diameters (mm) following inoculation with wild-type Alternaria brassicicola, Δamr1:Amr1p-GFP, Δamr1:Amr1,and Δamr1:TrpCp-Amr1.


We performed additional pathogenicity assays using as few as ~300 spores to test the effect of low inoculum concentration on pathogenesis (Figure 2 B, C). Lesions caused by ~440 wild-type conidia were small and some stopped expanding, while lesion diameters caused by ~400 Δamr1:Amr1p-GFP mutant conidia were on average ~3-fold larger (p<0.0001, df = 29, two tailed t-test) and continued to expand throughout the infection process. Inoculation with approximately ~330 wild-type conidia produced black spots, indicating successful penetration but a failure to colonize host tissue. Lesion diameters created by the same concentration of mutant conidia, however, were on average ~5-fold larger (p<0.0001, df = 29, two tailed t-test) and continued to expand during the five-day experiments. This result demonstrated that the Δamr1 mutants required fewer conidia than the wild type for successful infection.

Amr1 gene expression and its protein localization during plant infection

In order to support the notion that AMR1 is a transcription factor, we monitored the localization of AMR1 protein using a mutant strain expressing an AMR1-GFP fusion protein. The AMR1-GFP fusion protein was primarily located in the nuclei of highly melanized conidia and aerial hyphae (Figure 3A). AMR1 protein expression in conidia was consistent with its proposed regulatory role in melanin biosynthesis during conidiogenesis in B. oryzae, Coc. heterostrophus, Col. lagenarium, and M. grisea [21], [22], [41]. We could not detect the fluorescent fusion protein in hyphae growing on PDA (data not shown) or at 24 hours post-inoculation (hpi) in planta during the early stages of pathogenesis (Figure 3B). In contrast, the fusion proteins were detected in the nuclei of hyphae actively invading host tissues during the late stage of infection (Figure 3C). As a negative control, expression and localization of GFP were also monitored in a promoter-tagged mutant (Δamr1:Amr1p-GFP) in which GFP expression was regulated by the native Amr1 promoter (Figure 3 D–F). GFP in the promoter-tagged mutant was distributed throughout the cytoplasm of all fungal tissues, including conidia. The GFP continued to be expressed at moderate levels in the hyphae of the Δamr1:Amr1p-GFP mutant, even during the early stages of plant infection when AMR1-GFP protein was undetectable (Figure 3 B, E).


Figure 3. Confocal microscopic images showing Amr1 gene expression and its protein localization in fungal nuclei during conidiation and pathogenesis.

A–C. Green fluorescence indirectly shows localization of Amr1 protein in fungal nuclei: GFP is expressed as the Amr1-GFP fusion protein. D–F. Green fluorescence in fungal tissues indicates an abundance of GFP, whose gene is under the regulation of the Amr1 promoter in the Δamr1:Amr1p-GFP mutant. G. Fluorescence shows co-localization of Amr1-GFP proteins, and 4′,6-diamidino-2-phenylindole (DAPI) stains that bind to nucleotides. Pink color represents auto-fluorescence of plant tissues. Arrowheads indicate conidia. Arrows point to fungal hyphae with localized green fluorescence in the nuclei during late plant infection. hpi = hours post-inoculation.


Melanin deficiency and pink pigment secretion by mutants

All strains of verified gene disruption and deletion mutants (amr1, Δamr1, and Δamr1:Amr1p-GFP) produced melanin-deficient conidia. The color of the mutant colonies, however, was orange by 3 days post-inoculation (dpi) when grown on a PDA medium (inset in Figure 4 A, B). The orange color became denser with increased conidia production, or when the culture was exposed to light. The pigment was not accumulated in fungal tissues but was secreted into the medium. Droplets of pink pigment were visible at the tips of many conidial chains on PDA and on plant surfaces by 7 dpi (Figure 4E). The pigment was also secreted during the culture in liquid medium (Figure S3A). The pigment did not have phytotoxic effects on host plants (Figure S3B). Furthermore, addition of pink pigment to inocula of wild-type conidia did not affect lesion size (Figure S3C). The wild-type allele of the Amr1 gene, controlled by its own promoter or by the TrpC promoter, restored the ability of the Δamr1 mutants to accumulate melanin (Figure 4 C, D) and pink pigment was no longer visible. Conidia produced by the mutant complemented with TrpCp-Amr1 constructs were slightly lighter than the taupe-colored wild-type conidia. None of the complemented mutants secreted pink pigment to the medium.


Figure 4. Melanin deficiency and its effects on fungal morphology and responses to stressors.

A–D. Conidial chains produced by each strain. Insets: fungal growth of each strain showing colony color and pink pigment secreted by Δamr1 mutants during saprophytic growth on PDA. E. Pink exudate from the tips of conidial chains of the Δamr1 mutant. F. Germination and hyphal growth after UV irradiation. G. Growth comparisons between mutants and wild type either under high temperature (33°C), or in the presence of the indicated chemical. Abbreviations: Δamr1 = Amr1 deletion mutant; Δamr1:Amr1 = Δamr1 mutant complemented with the Amr1 allele; Δamr1:TrpCp-Amr1 = mutant that constitutively expresses the Amr1 gene under control of the TrpC promoter.


Susceptibility of Δamr1 mutants to abiotic stress

Because melanin plays an important role in protecting fungi against various types of stress, we examined the effect of UV irradiation, glucanase, heat, salt, and osmolites on Δamr1 mutants. UV irradiation at 80 mJ had a marginal effect on conidial germination and hyphal growth of wild-type A. brassicicola (Figure 4F, upper panel). However, 40 mJ of UV irradiation severely affected conidial germination and hyphal growth of the mutants, and 60 mJ almost eliminated germination of the Δamr1mutant (Figure 4F, lower panel). It took more than three hours for β-glucanase to digest the melanized cell walls of wild-type A. brassicicola spores, but less than one hour for it to digest the cell walls of the melanin-deficient Δamr1 mutant (data not shown).

Germination and vegetative growth were not significantly different between the wild type and Δamr1 mutants under high-temperature conditions or in the presence of NaCl, H2O2, or sorbitol. The colony size was comparable for both the Δamr1 and Δamr1:Amr1p-GFP mutants, a complemented mutant, and the wild type on the same media (Figure 4G).

Melanin deficiency and virulence of abpks7 mutants

We produced gene disruption mutants of the melanin-associated polyketide synthase (Pks) homolog in A. brassicicola (AbPks7). The Pks uses acetyl coenzyme A or malonyl coenzyme A as a precursor and produces a melanin-intermediate product, 1,3,6,8-tetrahydroxynaphthalene (T4HN). The abpks7 disruption mutants produced melanin-deficient conidia (Figure S4 A–C). These melanin-deficient mutants caused lesions comparable to the wild type on their host plant, B. oleracea (Figure 4D), further demonstrating that melanin is not a virulence-associated factor in this fungus.

Enhanced growth of the Δamr1mutant in media containing pectin

Carbohydrate-active enzymes, such as glycoside hydrolases, polysaccharide lyases, and carbohydrate esterases are slightly expanded in A. brassicicola as has been found in other plant pathogenic Dothideomycetes [35]. The enzymes encoded by these genes are thought to be involved in the breakdown of complex carbohydrates in the cell walls of host plants. We indirectly evaluated the role of Amr1 in regulating genes encoding putative cell wall-degrading enzymes by comparing the vegetative growth (dry weight of mycelium) of the Δamr1-1 and Δamr1-4 mutants to the wild type on various carbon sources. A minimal broth medium was supplemented with either 1% glucose, or the common cell wall polysaccharides α-cellulose, xylan, or pectin. Vegetative growth of the Δamr1-1 and Δamr1-4 mutants was the same as the wild type in the presence of glucose, xylan, lignin, and α-cellulose (data not shown). However, growth of these mutants was greater than the wild type in the presence of citrus pectin (Figure 5 A, C). To verify this observation, we tested the effects of pectin on the growth of the Δamr1-3, Δamr1-5, and Δamr1:Amr1p-GFP mutants. These mutants also showed a significant increase in vegetative growth over the wild type in the presence of pectin (Figure 5 B, D). Vegetative growth was up to 86% greater than the wild type when mutants were cultured longer than 46 hours (Figure 5 C, D). There was no significant difference in growth between the wild type and the mutants in the presence of glucose. Because all Δamr1 mutants showed similar levels of melanin deficiency, pink pigment secretion, and pectin utilization, we used Δamr1-4 in subsequent gene expression profiling studies.


Figure 5. Effect of pectin on the vegetative growth of four strains of Δamr1 mutants and wild-type Alternaria brassicicola.

A–D. Data show mean dry weight in milligrams. Each graph is the result of an independent experiment. The poor correlation between the absolute amount of dry biomass and the length of the incubation period was partially due to differences in the number of conidia used in each experiment. Values above the bars indicate the percent increase in dry biomass of the mutant compared to the wild type. Hours under each chart show the length of the incubation period. ** indicates p<0.01. Error bars represent standard deviation. wt = wild type.


Regulation of genes by Amr1 during pathogenesis

We used RNA-seq approach to determine the subset of genes regulated by Amr1. Using the RNA-seq approach, gene expression profiles were obtained and then compared between a Δamr1 mutant and the wild type during the late stage of infection on green cabbage. At this time, the AMR1 protein was localized in the nuclei of infecting hyphae. A total of 6.17×107 reads were produced for the wild type and 6.94×107 for the Δamr1 mutant. Of these, 4.14×107 (67.1%) tags for the wild type and 4.78×107 (68.9%) tags for the mutant were mapped to the genome of A. brassicicola. Of A. brassicicola's 10,688 predicted genes, 101 and 163 genes were significantly (p<0.05) up- or down-regulated more than 2-fold, respectively, in the Δamr1 mutant compared to the wild type (Table S2). This respectively represents 0.95% and 1.53% of the genes in the A. brassicicola genome. Functional categories that were overrepresented in the up-regulated genes included putative cell-wall depolymerization enzymes (Table S3). They included 21 glycoside hydrolases, two pectate lyases, and one pectin esterase (Table S2). In addition, two cutinases and one lipase enzyme were also up-regulated in the mutant. One of the two cutinases, CutAb, was up-regulated >32-fold. The necrosis-inducing protein gene (3.1-fold), two P450 genes (2.8- and 2.6-fold), and the homolog of the peptidase aegerolysin (3.3-fold) were also up regulated in the mutant. 23 of 101 up-regulated genes in Δamr1 mutants were expressed at a lower level in the previously characterized Δabvf19 mutants and eleven of the shared genes between the two mutants were glycoside hydrolases (Table 2 and Figure 6). Genes in the melanin synthesis pathway were among the down-regulated genes. As expected, the expression level of Amr1 was close to zero (background noise level) in the Δamr1mutant during the late stage of infection, compared to a high level of induction in the wild type (Table S2). Expression levels of other structural genes in the melanin synthesis pathway—Brn2, Scd1, and Brn1 (Eliahu et al., 2007; Tsuji et al., 2000)were 8- to 30-fold lower in the mutant than in the wild type. Differential expression between the wild type and Δamr1 mutant was verified by quantitative real time PCR (qRT-PCR) for seven differentially and three similarly expressed genes (Table S2). AbPks7 was annotated as three separate genes and their expression in the mutants was reduced about four-fold (Table S2).


Figure 6. Comparison of differentially expressed genes in the Δamr1 and Δabvf19 mutants compared to wild-type Alternaria brassicicola.

A. Comparison between up-regulated genes in the Δamr1 mutant and down-regulated genes in the Δabvf19 mutant during the late stage of infection. B. Number of differentially expressed glycoside hydrolase genes. C. Comparison of expression levels for the 24 genes differentially expressed in the Δamr1 and Δabvf19 mutants compared to the wild type. Y-axis shows Log2 (mutant)expression-Log2 (wild type)expression).


Effect of Amr1 on gene expression during the early infection stage

We examined the expression of the nine genes at two additional time points when Amr1 was expressed at a low level. As expected, the Amr1 gene and its three downstream genes in the melanin synthesis pathway (Brn2, Scd1, and Brn1) were not expressed in the Δamr1:Amr1p-GFP mutant under any of the tested conditions (Figure 7B). They were expressed at a low level in the wild type at 40 and 64 hpi, when Amr1 expression was low. When the Amr1 transcripts were slightly increased at 88 hpi, however, the three genes were highly induced in the wild type (Figure 7A). The expression level of Amr1 was extremely low in the wild type compared to the expression of GFP in the Δamr1:Amr1p-GFP mutant. This may explain why the gene expression level of GFP is high in the Δamr1:Amr1p-GFP mutant, but AMR1-GFP fusion protein was not detected during early infection (Figure 3 B, E). Moreover, this could have been due to the absence of feedback inhibition from Amr1. It may also have been caused by the increased stability of GFP mRNA, its protein, or both as compared to Amr1. The high stability of GFP protein has previously been reported (reviewed in [40]).


Figure 7. Expression of melanin biosynthesis-associated genes and four hydrolytic enzyme-coding genes.

A–C. Relative transcription abundance of each gene was determined in comparison to actin gene transcripts in the same tissue. Y-axes show relative abundance of the transcripts compared to the actin gene. D. Expression ratio between the Δamr1 and wild type during late stage of infection. A total of three biological replicates (N = 3) were used for this study. Bars represent standard error. wt = wild type, Δa = Δamr1:Amr1p-GFP, GY = glucose yeast extract broth. SCD1 = Scytalone dehydratase, Brn1 = T3HN reductase, Brn2 = T4HN reductase, Cbh7 = cellobiohydrolase, Amr1 = Alternaria melanin regulation, chymo = chymotrypsin.


The lipase and cellobiohydrolase Cbh7 (AB06252.1) genes were weakly expressed in the wild type from early pathogenesis until 64 hpi; during this time the Amr1 expression level was low in the wild type. There was no expression of these two genes during saprophytic growth in an axenic medium by either the wild type or the mutant (Figure 7 A, B). At 88 hpi, the time when Amr1 was induced in the wild type, Cbh7 and lipase induction were 4.3- and 4.7-times greater respectively (p<0.001, t-test, df = 2) in the Δamr1:Amr1p-GFP mutant than in the wild type (Figure 7D). The glycoside hydrolases and chymotrypsin genes were not expressed in either the wild type or the mutant during saprophytic growth in glucose yeast extract broth (GYEB) (Figure 7 GY24h). Expression of the chymotrypsin gene was twice as high in the mutant at 64 hpi as in the wild type, but there was little difference in chymotrypsin levels at 88 hpi (Figure 7C). In summary, transcript levels of putative hydrolytic enzyme genes, including lipase and Cbh7, were moderately higher with statistical significance in the mutant than in the wild type during the colonization of host plants (Figure 7). This coincided with the time when the Amr1 gene was induced in the wild type.

The AbVf8 gene is a pathogenesis-associated transcription factor

Another virulence factor identified in these studies was the putative transcription factor AbVf8. The AbVf8 gene showed no sequence similarities to known pathogenesis-associated genes in any fungus. AbVf8 in A. brassicicola and its homologs in other fungi (e.g., in Pyrenophora spp. or Phaeosphaeria nodorum) were annotated as either a hypothetical protein or a predicted transcription factor. The diameters of lesions caused by disruption mutants of AbVf8 (abvf8) were 50–80% smaller than those produced by the wild type. Vegetative growth rates and the formation of conidia were comparable to the wild type. Predicted domains in the protein sequence of the AbVf8 genes were somewhat informative regarding their functions. The predicted protein encoded by AbVf8 contained a putative SET domain. SET domains are thought to be an epigenetic regulator of gene expression during development and modulate chromatin structure.

Because it was a novel transcription factor associated with pathogenesis and had a possible association with epigenetic regulation, we decided to confirm the results of pathogenicity assays using additional mutants. We made gene deletion mutants (Δabvf8) by replacing the coding region with a HygB resistance cassette. Southern hybridization confirmed that the AbVf8 gene was replaced by a single copy of the HygB resistance cassette in nine transformants (Figure S2). One transformant had an ectopic insertion of multiple copies of the HygB resistance cassette. Pathogenicity assays using Δabvf8-2, Δabvf8-3, and an ectopic insertion mutant produced results consistent to those we produced with the gene disruption mutants. Both deletion mutants caused lesions with diameters 55% smaller than lesions caused by the wild type or the ectopic insertion mutant (Table 2). We complemented the Δabvf8-2 mutant with the wild-type allele of the AbVf8 gene. Complemented mutants restored its ability to produce lesions comparable to the wild type (data not shown). We will further characterize the functions of the AbVf8 gene in the future.


Transcription factors associated with pathogenesis

Of 138 A. brassicicola transcription factor genes with zinc finger domains and other regulatory genes analyzed to date in this and previous studies, only six genes were strongly associated with pathogenesis. These six genes were AbSte12, AbPacC, AbVf19, Amr1, AbVf8, and AbPro1 (Table 1). This result was comparable to the comprehensive knockout study of 693 putative transcription factors in Fusarium graminearum, where 62 genes affected virulence [42]. AbSte12 is a homolog of the Ste12 gene with mutants that are either nonpathogenic or reduced in virulence in several fungi (reviewed in [43]). Ste12 is regulated by a MAP kinase, Fus3/Kss1. Mutants of either homologous gene in A. brassicicola showed defective conidial development and were nonpathogenic [14], [15]. The AbPacC gene has a single copy in the genome sequence of A. brassicicola and shows a high sequence similarity (E = 0.0) with the PacC gene in other fungi. PacC is the transcription factor in the pH regulation pathway of Aspergillus nidulans [44]. It also controls the expression of fumonisin toxins in F. verticillioides [45] and endopolygalacturonase and oxalic acid in S. sclerotiorum [38], [46]. Mutation of its homologous genes reduced virulence in S. sclerotiorum and Col. acutatum and increased virulence in F. oxysporum [37], [38], [47]. AbVf19 regulates a suite of genes that are important for decomposing and utilizing plant materials during the late stage of plant infection [20]. We have yet to characterize the functions of AbVf8 and AbPro1. Regardless of their functions, however, all mutants studied to dates except Amr1 were nonpathogenic or showed a reduction in virulence.

Melanin biosynthesis and virulence in A. brassicicola

Melanin is a ubiquitous pigment that plays an important role in protecting fungi from the damaging effects of environmental stress. It accumulates in the cell walls of hyphae and conidia during the late stationary phase of mycelial growth. It also forms under stressful conditions, including ultraviolet irradiation, a hyperosmotic environment, nutrient deprivation, or an accumulation of toxic wastes during in vitro culture. Over-expression of a Cmr1 ortholog in Bipolaris oryzae (Bmr1) caused continuous induction of the three downstream genes in the melanin synthesis pathway and increased melanization of the colonies [41]. The loss of function of Cmr1, its homologs, or their downstream genes in other fungi, resulted in melanin deficiency [21], [22], [25], [48]. We identified a transcription factor in this study, Amr1, which regulates the melanin synthesis pathway in A. brassicicola. The melanin synthesis-associated structural genes Brn1 and Pks are located together in a 30 Kb region and their organization and orientation is the same as in the closely related fungus Coc. heterostrophus [21]. Interestingly, as reported here melanin was not associated with virulence in A. brassicicola as has been reported in other fungi [21]. However, based upon our data there appears to be a linkage between melanin biosynthesis and a reduction in virulence, perhaps due to a lifestyle switch from pathogenesis to reproduction.

Mutation of the Amr1 gene unexpectedly caused increased virulence

Amr1 gene knockout mutants, both gene disruption (amr1) and gene deletion (Δamr1 and Δamr1:Amr1p-GFP), were easily recognized by the lack of melanin in their conidia, orange-colored colonies, and the secretion of a pink pigment (Figure 4). These phenotypes were similar to those found in mutants of its homologs in other fungi [21], [22], [41]. The Δamr1mutants were more susceptible to UV light and enzyme digestion (Figure 4). Unexpectedly, the Amr1 gene knockout mutants were more virulent than the wild type, producing larger lesions (Table 3), and required fewer conidia for successful infection (Figure 2). Mutation of Amr1 homologs in the phytopathogenic fungi M. grisea and Col. lagenarium produced melanin-deficient conidia. The appressoria of these mutants, however, remained melanized suggesting an alternate regulatory mechanism and pathogenicity was not affected [22]. Melanin deficiency caused by a mutation of other genes in the melanin biosynthesis pathway was associated with a loss of pathogenicity in M. grisea and Col. lagenarium [49], [50], [51] and a decrease in virulence in the opportunistic human fungal pathogens Aspergillus fumigatus and Waggiella dermatitidis [52], [53]. In most other plant-pathogenic fungi, melanin deficiency usually had little effect on pathogenesis under controlled laboratory conditions [25], [31], [48], [54]. Melanin deficient mutants of Coc. heterostrophus, however, were nonpathogenic under field conditions possibly due to increased sensitivity to UV light outdoors [54]. In summary, the increased virulence of the Δamr1 mutants was unexpected and unique among melanin-deficient mutants of pathogenic fungi.

The cause of increased virulence in Δamr1 mutants

The pink pigment secreted by Δamr1 mutants was neither toxic to host plants nor beneficial for the wild type A. brasscicola to infect host plants (Figure S3). Furthermore, melanin-deficient mutants of another gene, abpks7, showed no changes in virulence (Figure S4). These data suggested that neither the presence of pink pigment nor the lack of melanin is associated with the increased virulence of the Δamr1 mutants investigated in this study. Additionally, there was no difference in germination or vegetative growth between the Δamr1 mutants and the wild type in the presence of various chemicals (Figure 4G). However, the mutants were more susceptible than wild type to UV light (Figure 4F) and the cell wall-degrading enzyme, glucanase. These data suggested that the mutants' ability to respond to stress was not associated with their increased virulence, since plant glucanases are known to be defense enzymes.

The ability of the mutants to outgrow the wild type when citrus pectin was the major carbon source (Figure 5) suggested that the mutants were more efficient in utilizing the pectin component of plant cell walls and middle lamellae. The expression profile comparisons identified 30 hydrolytic enzyme genes expressed moderately more in the mutant than in wild type A. brassicicola during pathogenesis (Table S2). However, they included only two pectate lyase genes and one pectin esterase gene. Other genes upregulated in the mutant compared to the wild type included genes coding for two cutinases, a lipolytic enzyme and 20 other various glycoside hydrolases (GH). Interestingly, 6 of the 22 genes in the GH61-gene family were expressed at higher levels in the Δamr1 mutant. The GH61 family might have a role in degrading cellulose, lignocellulose, chitin, or other polysaccharides [55]. The GH61 family may be involved in digesting pectins whose side chains are structurally diverse in constituting sugars and glycosidic linkages. They may digest other cell-wall components, such as cellulose and hemicelluloses along with other glycoside hydrolases. It was not possible in this study to evaluate whether the Δamr1 mutants made better use of these cell wall materials because A. brassicicola grew poorly in the presence of α-cellulose, beechwood-xylan, and lignin. Our uninformative growth data was likely due to the non-brassica source of cell wall materials. Nonetheless, the gene expression and growth assay data indirectly support the importance of cell wall-degrading enzymes and possibly those involved in pectin digestion, since pectic polysaccharides constitute about one-third of the cell wall components and most of the middle lamella in dicotyledonous plants.

Induction of the Amr1 gene in A. brassicicola did not occur until 64 hpi (Figure 7). Suppression of the Amr1 gene might be important during pathogenesis when energy is allocated to colonization of host tissues and overcoming host defense mechanisms, rather than producing reproductive dispersal structures like conidia. In the wild type, the transcripts of hydrolytic enzyme genes were slightly increased compared to the actin transcripts, as previously reported [14], [56], [57]. In addition, when conidiation was initiated the induction of Amr1 was accompanied by a sharp increase in transcripts of its downstream genes in the melanin synthesis pathway. There was also a small increase in the transcripts of hydrolytic enzyme-coding genes in the wild type. However, many of these genes, mostly expressing glycoside hydrolases, a few cutinases, and pectate digestion enzymes, were expressed at higher levels in the Δamr1-4 and Δamr1:Amr1p-GFP mutants, which did not express Amr1 (Figure 7D). This suggests that Amr1 suppressed genes predicted to be involved in the digestion of lipids and carbohydrates. This occurred during late pathogenesis (88 hpi) when they were still needed to some extent for either killing [58] or digesting host tissue. Whether the Amr1 gene is involved in the regulation of most hydrolytic enzymes sequentially induced in A. brassicicola during pathogenesis has yet to be investigated [24], [59]. Other genes encoding a necrosis-inducing protein, two P450s, the aegerolysin peptidase homolog, and other hypothetical proteins might have contributed to the increased virulence of the Δamr1 mutant and warrant further investigation. Interestingly, 24 genes expressed at a higher level in Δamr1 were expressed at lower levels in the Δabvf19 deletion mutants that displayed a reduction in virulence (Figure 6C), suggesting opposite roles of the two transcription factor genes in the regulation of pathogenesis-associated genes. This finding may aid in the selection of specific targets for future analyses of their functions in pathogenesis.

Evolution of virulence in A. brassicicola

The melanin-deficient mutant phenotype and expression pattern of the Amr1, Brn1, Brn2, and Scd1 genes in A. brassicicola suggested that the function of the Amr1 gene was to regulate melanin synthesis during conidiogenesis. This function is highly conserved in other fungi [21], [22], [41]. In addition to regulating the melanin biosynthesis pathway, Amr1 negatively regulated virulence. _ENREF_1The evolution of virulence has been a subject of extensive theoretical analysis. The “trade-off hypothesis” [60] produced a number of models showing a contested relationship between virulence and transmissibility. It challenges the hypothesis that strong virulence can seriously damage or kill host plants and threaten pathogen survival, so parasites will ultimately evolve to be less virulent. However, analyses of ecological and epidemiological factors related to virulence has produced inconsistent results [61]. Therefore, a modified trade-off theory predicted that strong parasites became moderately virulent to their hosts over time, but did not completely lose virulence [62]_ENREF_4. Our current work identified a transcription factor suppressing virulence but enhancing the ability of a fungus to preserve itself in space and time via melanin synthesis, primarily in conidia. This study provides an example of a transcription factor gene important for survival in nature that plays a critical role in negatively regulating virulence in the necrotrophic plant pathogen, A. brassicicola. The suppression of virulence we observed is unique and might have been acquired by this fungus, as this phenotype has not been observed in loss-of-function mutants of Amr1 homologs in other species of plant pathogenic fungi. An ancestral fungal strain of A. brassicicola without the ability to suppress virulence by the activity of Amr1 could have been a more virulent pathogen. Negative regulation of virulence is rare but exists in other phytopathogenic fungi. For example, a phosphatase enzyme in the MAP kinase signaling pathway negatively regulates virulence in Ustilago maydis [63]. Currently, A. brassicicola is a prolific saprophyte and necrotroph, efficiently colonizing susceptible, weakened, or dead host plants. Its pathogenicity, however, is inhibited on more resistant host plants. We speculate that the suppressive functions of Amr1 contribute to the specialized adaptation of A. brassicicola as an efficient and successful facultative parasite.

Materials and Methods

Fungal strains, their transformation, maintenance, and primers

We used the facultative plant pathogen Alternaria brassicicola (Schweinitz, Wiltshire) (ATCC96836) in this study. Growth and maintenance of the fungus and its transformation, nucleic acid isolation, mutant purification, and mutant verification by Southern hybridization were performed as described previously [15]. The wild-type fungus and each of the mutant strains created during this study were purified by two rounds of single-spore isolation. The cultures were maintained as glycerol stock in separate tubes, with one tube used for each assay.

Data availability

The genome sequence and gene predictions for A. brassicicola ATCC 96836 were obtained from​ernaria_brassicicola/ and are available from DDBJ/EMBL/GenBank under accession ACIW00000000. In addition, the annotated genome is available in the context of other Dothideomycetes through the interactive JGI fungal portal MycoCosm [34] at

Identification of putative transcription factors

The partially assembled genome sequence was annotated using an EnsEMBL annotation pipeline [64]. The HMM search results of FGENESH predicted peptides against PFAM and TIGRFAM databases were stored in a SQL database. A total of 251 PFAM accession numbers and 13 keywords descriptions were used to manually search for the putative transcription factors using SQL queries. Subsequently, the predicted amino acid, predicted cDNA, and genomic DNA sequences were manually retrieved from the database.

Primer design and generation of targeted gene mutants

Four primers for targeted gene disruption and six primers for targeted gene deletion were synthesized for each gene (Table S4). We followed methods described previously for all steps from producing transformation constructs to verifying purified mutants [15], [65]. Briefly, constructs of up to 12 genes were independently transformed at a time. The transformed protoplasts were plated on a molten regeneration medium (1 M sucrose, 0.5% yeast extract, 0.5% casamino acids, and 1% agar) for 24 hours, followed by an overlay with the same volume of PDA containing 30 ug/ml HygB. On the seventh day after the transformation, four transformants of each gene were transferred to PDA plates containing 30 ug/ml HygB and cultured for five days. The transformants were purified by two rounds of single-spore isolation and verified by PCR using two sets of verification primers for each gene (Table S5).

Generation of the Amr1 deletion mutants, Δamr1 and Δamr1:Amr1p-GFP

All transformation constructs described in this work were produced by a double-jointed PCR method [66] with modifications described previously [15]. We made Amr1 deletion mutants by replacing the gene with a HygB resistance cassette. The replacement construct was produced with the following three sets of primers (Figure 8). The primer set P1 and P2, and the primer set P5 and P6 were used to respectively amplify the 5′ and 3′ flanking regions of the targeted locus. Another set of primers, P3 and P4, was used to amplify the HygB-selectable marker gene cassette (1,436 bp) from pCB1636. In order to create mutants expressing green fluorescence under the control of the Amr1 gene promoter, the Amr1 promoter region and 3′ flanking region were amplified with P7 and P8, and P5 and P6, respectively. Another set of primers, P9 and P4 was used to amplify the 2,423 bp that covered the coding regions of the GFP and the HygB resistance cassette. All primers used Amr1gene are listed in Table S6.


Figure 8. Schematic diagram of the PCR strategy used to make each construct.

A. Construct for replacement of the Amr1 gene with a Hygromycin B resistance cassette. B. Construct for replacement of the Amr1 gene with a GFP coding region and Hygromycin B resistance cassette. C. Amplification of the wild-type allele of the Amr1 gene. D. Construct for the constitutive Amr1 expression cassette. E. Construct for the Amr1-GFP fusion protein expression.


Complementation of Δamr1 mutants

The Δamr1-5 mutant was complemented with either the wild-type Amr1 allele with its native promoter, or a chimeric construct of an Amr1 allele under the control of a constitutive promoter. This promoter was derived from a TrpC gene (TrpCp) [67]. We used two primers, P1 and P6, to make constructs and reintroduce wild-type Amr1 into the Δamr1 mutant. These primers amplified the 6,196 bp wild-type allele of the Amr1 gene using A. brassicicola genomic DNA as a template. The PCR product included a 1,342 bp 5′ flanking region, 3,282 bp complete coding region, and a 1,570 bp 3′ flanking region. Separately, P18 and P19 were used to amplify a 2,226 bp-long nourseothricin-resistant cassette as a selectable marker gene, using a pNR vector as the template [68]. These two products (8 µg of Amr1 and 7 µg of NTC cassette) were mixed to transform the Δamr1 mutant.

To produce constitutive Amr1 expression mutants, three sets of primers were used to replace the 1,840 bp 5′ upstream to the predicted start codon with the TrpC promoter, TrpCp. We used two primers, P10 and P11, to amplify the 943 bp constitutive promoter region that includes a 152 bp partial coding sequence of NAT, 458 bp nonfunctional ToxA promoter, and a 333 bp-long functional TrpC promoter. Another set of promoters, P12 and P6, were used to amplify the 4,852 bp wild-type allele that spans the 3,282 bp coding region from the start codon to the stop codon of the Amr1 gene and the 1,570 bp 3′ flanking region of the gene. The PCR products were mixed and diluted 10 times. The final mixed PCR products were used as template DNA to create TrpCp-Amr1 gene constructs using two primers, P7 and P6. A total of 8 µg of the TrpCp-AMR1 chimeric gene constructs and 7 µg of the 2,226 bp-long nourseothricin-resistant cassette were transformed into the Δamr1 mutant. Among the nourseothricin-resistant transformants, two clones of the former (Δamr1:Amr1) and four clones of the latter (Δamr1:TrpCp-Amr1) were purified by two rounds of single-spore isolation in the presence of nourseothricin. Complementation of the mutant by a single copy of the Amr1 gene was confirmed by Southern blot hybridization. All mutant clones were tested for purity on PDA by checking their growth patterns, growth rates, and uniformity of colony colors (sectoring) in the presence and absence of selectable markers.

Generation of mutants expressing Amr1-GFP fusion proteins

In order to create mutants expressing green fluorescence protein fused to the C-terminal of AMR1, the Amr1 coding region (1,209 bp) and 3′ flanking region (192 bp) were amplified with P13 and P14, and P5 and P15, respectively. Another set of primers, P16 and P17, was used to amplify the 2,384 bp that covered the coding regions of the GFP and the HygB resistance cassette. The final transformation constructs were produced by PCR amplification from the mixture of the PCR products using P1′ and 6′z9219-3R.

Pathogenicity assays

Pathogenicity assays were performed with modifications as described previously using detached leaves harvested from 5- to 8-week-old plants or on the leaves of whole plants [15]. Commercially available seeds (Jonny's, Winslow, ME) of green cabbage (Brassica oleracea) or Arabidopsis thaliana were planted and grown under the same conditions for each pathogenicity assay. We grew green cabbage under 14 hours light 10 hours dark cycle and Arabidopsis under 10 hours light 14 hours dark cycle. Plants of similar heights and with similar-sized leaves were selected for each assay. Leaves were detached and placed in mini-moist chambers and randomly arranged on a laboratory bench for most assays. For the pathogenicity assays on whole plants, potted plants were placed in a semi-transparent plastic trough with adequate water. The troughs and plants were sealed with plastic wrap after inoculation to keep the relative humidity close to 100%. The increased virulence of each mutant was calculated using the formula (∑(Dmi-Dwi)/∑(Dwi))×100, where Dwi was the lesion diameter created by the wild-type for the ith sample and Dmi was the lesion diameter produced by the mutant for the ith sample (Table 2). We also analyzed lesion sizes among the wild type and various mutants (Tables 2). Lesion sizes were subjected to 5×4 (genotype×leaf position (leaf age)) two-way analysis of variance (ANOVA) using the general linear model (GLM) procedure in the Statistical Analysis System (SAS Institute, Cary, NC). Means were separated by the Waller-Duncan k-ratio (k = 100) t-test, or an f-test, whichever was appropriate. The Waller-Duncan test was chosen to minimize the Bayesian risk of the additive loss function.

Confocal microscopy

Infected plant tissues were trimmed with a razor blade, placed on microscope slides, and covered with Gold Seal cover glasses. Confocal images were acquired using a 633 C-Apochromat (numerical aperture 1.2) water-immersion objective lens and an Olympus Fluoview 1000 Laser Scanning Confocal System on a IX-81 inverted microscope. Spectra for fungal tissues expressing standard green fluorescence and for plant cells emitting autofluorescence were collected by simultaneous 488 nm and 543 nm excitation using 30 mW argon and 1 mW helium:neon lasers, respectively. The standard GFP spectrum was collected through 488 nm excitation using a 20 nm window from 505 to 525 nm. Plant tissue, including chloroplasts, was visualized using 543 nm excitation with a 560 nm-long pass filter. Images of fungal tissue grown in nutrient media were captured with 488 nm excitation and DIC-transmitted light. All fluorescent images were composed of multiple layers acquired with the Confocal System.

Measuring the response to stressors

Each fungal strain from glycerol stocks was inoculated on PDA with an appropriate selectable agent and grown in the dark for 5 days at 25°C. In order to test for sensitivity to osmotic stress and oxygen radicals, wild-type and mutant conidia were pipetted onto PDA containing 2% (w/v) or 4% (w/v) NaCl, 0.6 M or 1.2 M sorbitol, or 2.5 mM or 5 mM H2O2. Mutant and wild-type strains were also cultured at 28, 30, and 33°C to measure the effect of temperature on their growth. Colony diameters were measured 4 days post-inoculation (dpi). To study the effect of UV light on germination, 1,000 spores were pipetted onto PDA, irradiated at 20, 40, 60, and 80 millijoules in a UV crosslinker (Agilent Technologies, Inc., CA), and incubated at 25°C for 24 hours. All experiments were conducted three times.

Growth assays in the presence of sole carbon sources

Flasks (250 ml) containing a 50 ml broth of 0.5% (NH4)2SO4, 0.05% yeast extract, 0.15% KH2PO4, 0.06% MgSO4, 0.06% CaCl2, 0.0005% FeSO4 7H2O, 0.00016% MnSO4 H2O, 0.00014% ZnSO4 7H2O, and 0.00037% CoCl2 were supplemented with either 1% glucose or 1% of α-cellulose (cat #, C8002-1KG), or xylan (cat #, X4252-100G), lignin (cat # 471003-100G), or citrus pectin (cat #, P9135-500G) purchased from Sigma (St. Louis, MO). Each flask was inoculated with 4–6×105 conidia of either Δamr1-1, Δamr1-3, Δamr1-4, Δamr1-5, Δamr1:Amr1p-GFP, or the wild type. The flasks were incubated in the dark at 25°C with continuous agitation at 100 rpm. The flasks were shaken vigorously by hand several times during the first eight hours to prevent the conidia from sticking to the flask. Mycelia were harvested at 4 dpi, washed with distilled water, and dried at 70°C overnight. The increased biomass of each mutant and the wild type was calculated using the formula (∑(Wmi-Wwi)/∑(Wwi))×100, where Wwi was the dry weight of the wild type for the ith sample and Wmi was the weight of the mutant for the ith sample.

Gene expression analysis by RNA-seq

To study the regulatory roles of Amr1 in A. brassicicola, we compared gene expression profiles between the wild type and a Δamr1 mutant at 88 hours post-inoculation (hpi), a late stage of infection. We inoculated nine detached leaves harvested from three plants with conidia from the Δamr1-4 mutant and the wild type. Tissue samples containing both host plant tissue and fungal mycelium were harvested. Three biological replicates were produced for both the mutant and the wild type. Total RNA was purified from the tissue using an RNeasy kit (Quiagen, Palo Alto, CA). We used 4 µg of total RNA for each RNA sample to construct strand-specific sequencing libraries with a TruSeq RNA Sample Prep Kit (Illumina, San Diego, CA). Each library was constructed with unique index primers and all were run in a single lane. They were later decoupled using index primer sequences.

Sequence tags were mapped to the genome sequence of A. brassicicola using the programs TopHat 1.3.1 [69] and Bowtie 0.12.7 [70]. Default settings were used, except the segment length was set at 25 nucleotides and the number of allowed segment mismatches was set at 1 nucleotide. Additionally, intron length was designated as a minimum of 10 nucleotides and a maximum of 400 nucleotides. The program Cuffdiff, version 1.0.3, which is part of Cufflinks [71], was used to identify reads overlapping with previously predicted genes. The expression levels of each predicted gene were determined and normalized by the mapped Fragments Per Kilobase of exon model per Million (FPKM). Differentially expressed genes between the wild type and the mutant were determined by comparing FPKMs from three biological replicates for both the wild type and the mutant. We also applied a cutoff of at least a two-fold change in expression value for differential expression. The bias correction method was used while running Cuffdiff [72]. Custom scripts were written in Python to analyze the data.

Representation analysis

Custom scripts were developed in Python and R to analyze over- and under-representation of functional annotation terms in sets of differentially regulated genes using the Fisher Exact test. The Benjamini-Hochberg correction was used to correct for multiple testing using a p-value of <0.05.

Quantitative real-time PCR

Full-length sequences of the three downstream genes in the melanin synthesis pathway regulated by Amr1 were identified in the A. brassicicola genome by Blast search. Primer sequences for each gene are listed in Table S6. The Δamr1:Amr1p-GFP mutant and the wild type were used for qRT-PCR with three biological replicates of infected plant tissues. Each biological sample was collected from three or four leaves and total RNA was extracted using an RNeasy kit with DNAse digestion according to the manufacturer's protocol (Qiagen, Valencia, CA). Two micrograms of total RNA were transcribed to cDNA in a final volume of 20 µl using 50 ng of random pentamers and 200 ng of poly(T)20N with Superscript III (Invitrogen, Carlsbad, CA). Each cDNA was diluted 1:20. Subsequent qRT-PCR reactions were performed in a 20 µl volume containing 120 nM of each primer, 1 µl of diluted cDNA, and 10 µl of FastStart SYBRGreen Master (Roche, Mannheim, Germany). Each reaction was run in a Biorad I-cycler (Bio-Rad, Hercules, CA, USA) as described previously (Cho et al, 2006). Relative amounts of the transcript of each gene were calculated as 2−ΔCt using a threshold cycle (Ct), where ΔCt = (Ct,genei –Ct,actin). Fold changes of each gene between the wild type and Δamr1 strain were calculated as 2−ΔΔCt, where ΔΔCt = (Ct,genei –Ct,actin)Δamr1 (Ct,genei –Ct,actin)wild type.

Accession numbers

RNA-seq data: GEO Series Accession No. GSE36781​king.html

Amr1 (Alternaria melanin regulation) GenBank accession number: JF487829

Supporting Information

Figure S1.

Verification of targeted gene disruption by polymerase chain reaction (PCR). PCR amplification was performed with two pairs of target gene specific and HygB resistance gene primers. Relative primer binding locations are marked below the schematic diagram of the disrupted target gene on the left side of the gel images at the top row of page one. The diagrams were drawn based on a hypothesis that the target gene was disrupted by a single homologous recombination with a circularized linear minimal element construct as described previously (Cho Y, Davis JW, Kim KH, Wang J, Sun QH, et al. (2006) A high throughput targeted gene disruption method for Alternaria brassicicola functional genomics using linear minimal element (LME) constructs. Mol Plant Microbe Interact 19: 7–15.). Hatched box = a selectable marker gene, white box = a target genomic locus, and gray box = partial target gene. Amplification was performed with Taq polymerase for PCR reactions (Invitrogen, Carlsbad, CA, U.S.A.).



Figure S2.

Creation of Δabvf8 deletion mutants and pathogenicity assays. A. Schematic diagram of the wild-type locus of the AbVf8 gene, a replacement construct, and a mutant locus. The mutant locus depicts incorporation of the replacement construct into a wild-type locus by double homologous recombination resulting in gene replacement. B. Replacement of the AbVf8 coding region with the selectable marker, Hygromycin B (HygB) resistance cassette. Absence of a wild-type band in nine lanes on the AbVf8-probe blot (above) indicates loss of the targeted gene. On the HygB-probed blot (below), the expected 4.6 Kb band indicates a single-copy insertion of the Hyg B resistance cassette in 9 lanes. Asterisk (*) indicates lanes of mutants used in this study. Probe regions are marked by p-HygB and p-AbVf8. C. Lesions on Brassica oleracea 5 days after inoculation with ~2,000 conidia of the wild type (wt), two Δabvf8 mutants, and an ectopic insertion mutant. Abbreviations: H = Hind III enzyme digestion site, wt = wild type Alternaria brassicicola, Ect = ectopic insertion mutant.



Figure S3.

Pigment secreted by Δamr1 mutants and its effect on pathogenesis and host plants. A. Mycelia of the Δamr1 mutant at the bottom of the flask (left) is melanin deficient compared to the dark-colored wild-type mycelia (right). Both were cultured in 1% glucose and 0.5% yeast extract broth (GYEB) for three days, in the dark, with continuous agitation at 100 rpm. B. No toxic effects from the pigment. The pigment in 10 ml of GYEB was extracted with 10 ml chloroform, twice. The chloroform was removed by evaporation and the pigment dissolved in 1 ml of sterile water. The concentrated pigment was injected into green cabbage leaves to evaluate its toxic effect. Both the pigment and negative control (water) produced similar small water-soaked areas that disappeared within 24 hours. Injected areas are marked with circles. C. No visible effect of the pigment on plant infection. Wild-type conidia were inoculated either in water (left) or in pigment (right). Lesion size was not significantly different between the two treatments.



Figure S4.

Comparison of phenotypes between abpks7 mutants and wild-type Alternaria brassicicola. A. Stereoscopic view of an abpks7 mutant colony showing melanin-deficient aerial hyphae and conidia. B. Stereoscopic view of a wild-type colony showing melanized aerial hyphae and conidia. C. Hyaline conidia of an abpks7 mutant. Mutant conidia were mixed with a small number of dark, wild-type conidia to illustrate the color difference. D. Pathogenicity assay showing comparable lesions caused by three abpks7 mutants and wild-type A. brassicicola. Average lesion diameters caused by the mutants and wild type were not statistically different.



Table S1.

List of transcription factor genes whose mutants were screened in this study.



Table S2.

List of genes differentially expressed in the Δ amr1 mutants compared to the wild type.



Table S3.

Statistically over-represented functional annotation terms among 101 up-regulated genes in the Δamr1 mutant compared to the wild type. The 101 genes were selected based on the statistical analysis included in Cufflink with at least a two-fold-difference.



Table S4.

Disruption Primers (5′ to 3′ direction).



Table S5.

PCR verification primers (5′ to 3′ direction).



Table S6.

List of primers used for qRT-PCR and transformation constructs.




We thank Washington University School of Medicine, Genome Sequencing Center and Sandra Clifton for determining the Alternaria brassicicola genome sequence and Ann Alvarez, John Hu, Tina M. Carvalho, and Wayne Borth for generously sharing their research equipment. We especially thank Fred Brooks for critical reading of the manuscript. We also thank Steven R. Head and Lana Schaffer for generating RNA-seq data.

Author Contributions

Conceived and designed the experiments: YC AS. Performed the experiments: YC AS. Analyzed the data: YC AS RAO SPM KHW. Contributed reagents/materials/analysis tools: YC AS RAO. Wrote the paper: YC RAO CBL IVG FB.


  1. 1. Churchill A, LU S, Turgeon B, Yoder O, Macko V (1995) Victorin-deficient REMI mutants of Cochliobolus victoriae demonstrate a requirement for victorin in pathogenesis. Fungal Genet Newslett 42A: 41.
  2. 2. Ciuffetti L, Tuori R, Gaventa J (1997) A single gene encodes a selective toxin causal to the development of tan spot of wheat. Plant Cell 9: 135–144. doi: 10.2307/3870536
  3. 3. Del Sorbo G, Scala F, Parrella G, Lorito M, Comparini C, et al. (2000) Functional expression of the gene cu, encoding the phytotoxic hydrophobin cerato-ulmin, enables Ophiostoma quercus, a nonpathogen on elm, to cause symptoms of Dutch elm disease. Mol Plant Microbe Interact 13: 43–53. doi: 10.1094/mpmi.2000.13.1.43
  4. 4. Yun S-H, Turgeon B, Yoder O (1998) REM-induced mutants of Mycosphaerella zeae-maydis lacking the polyketide PM-toxin are deficient in pathogenesis to corn. Physiol Mol Plant Pathol 52: 53–66.
  5. 5. Ito K, Tanaka T, Hatta R, Yamamoto M, Akimitsu K, et al. (2004) Dissection of the host range of the fungal plant pathogen Alternaria alternata by modification of secondary metabolism. Mol Microbiol 52: 399–411. doi: 10.1111/j.1365-2958.2004.04004.x
  6. 6. Johnson RD, Johnson L, Itoh Y, Kodama M, Otani H, et al. (2000) Cloning and characterization of a cyclic peptide synthetase gene from Alternaria alternata apple pathotype whose product is involved in AM-toxin synthesis and pathogenicity. Mol Plant Microbe Interact 13: 742–753. doi: 10.1094/mpmi.2000.13.7.742
  7. 7. Spassieva SD, Markham JE, Hille J (2002) The plant disease resistance gene Asc-1 prevents disruption of sphingolipid metabolism during AAL-toxin-induced programmed cell death. Plant J 32: 561–572. doi: 10.1046/j.1365-313x.2002.01444.x
  8. 8. Tanaka A, Shiotani H, Yamamoto M, Tsuge T (1999) Insertional mutagenesis and cloning of the genes required for biosynthesis of the host-specific AK-toxin in the Japanese pear pathotype of Alternaria alternata. Mol Plant Microbe Interact 12: 691–702. doi: 10.1094/mpmi.1999.12.8.691
  9. 9. Pedras MS, Chumala PB, Jin W, Islam MS, Hauck DW (2009) The phytopathogenic fungus Alternaria brassicicola: phytotoxin production and phytoalexin elicitation. Phytochemistry 70: 394–402. doi: 10.1016/j.phytochem.2009.01.005
  10. 10. Hashimoto M, Higuchi Y, Takahashi S, Osada H, Sakaki T, et al. (2009) Functional analyses of cytochrome P450 genes responsible for the early steps of brassicicene C biosynthesis. Bioorg Med Chem Lett 19: 5640–5643. doi: 10.1016/j.bmcl.2009.08.026
  11. 11. Wight WD, Kim KH, Lawrence CB, Walton JD (2009) Biosynthesis and role in virulence of the histone deacetylase inhibitor depudecin from Alternaria brassicicola. Mol Plant Microbe Interact 22: 1258–1267. doi: 10.1094/mpmi-22-10-1258
  12. 12. Otani H, Kohnobe A, Kodama M, Kohmoto K (1998) Production of a host-specific toxin by germinating spores of Alternaria brassicicola. Physiol Mol Plant Pathol 52: 285–295. doi: 10.1007/978-94-011-5218-1_7
  13. 13. Oka K, Akamatsub H, Kodamab M, Nakajimab H, Kawadab T, et al. (2005) Host-specific AB-toxin production by germinating spores of Alternaria brassicicola is induced by a host-derived oligosaccharide. Physiol Mol Plant Pathol 66: 12–19.
  14. 14. Cho Y, Cramer RA Jr, Kim KH, Davis J, Mitchell TK, et al. (2007) The Fus3/Kss1 MAP kinase homolog Amk1 regulates the expression of genes encoding hydrolytic enzymes in Alternaria brassicicola. Fungal Genet Biol 44: 543–553. doi: 10.1016/j.fgb.2006.11.015
  15. 15. Cho Y, Kim K-H, La Rota M, Scott D, Santopietro G, et al. (2009) Identification of virulence factors by high throughput targeted gene deletion of regulatory genes in Alternaria brassicicola. Mol Microbiol 72: 1316–1333. doi: 10.1111/j.1365-2958.2009.06689.x
  16. 16. Craven KD, Velez H, Cho Y, Lawrence CB, Mitchell TK (2008) Anastomosis is required for virulence of the fungal necrotroph Alternaria brassicicola. Eukaryot Cell 7: 675–683. doi: 10.1128/ec.00423-07
  17. 17. Kim KH, Willger SD, Park SW, Puttikamonkul S, Grahl N, et al. (2009) TmpL, a transmembrane protein required for intracellular redox homeostasis and virulence in a plant and an animal fungal pathogen. PLoS Pathog 5: e1000653. doi: 10.1371/journal.ppat.1000653
  18. 18. Oide S, Moeder W, Krasnoff S, Gibson D, Haas H, et al. (2006) NPS6, encoding a nonribosomal peptide synthetase involved in siderophore-mediated iron metabolism, is a conserved virulence determinant of plant pathogenic ascomycetes. Plant Cell 18: 2836–2853. doi: 10.1105/tpc.106.045633
  19. 19. Kim K-H, Cho Y, La Rota M, Cramer R, Lawrence C (2007) Functional analysis of the Alternaria brassicicola nonribosomal peptide synthetase gene AbNPS2 reveals a role in conidial cell wall construction. Mol Plant Pathol 8: 23–39. doi: 10.1111/j.1364-3703.2006.00366.x
  20. 20. Srivastava A, Ohm RA, Oxiles L, Brooks F, Lawrence CB, et al. (2012) A Zinc-Finger-Family Transcription Factor, AbVf19, Is Required for the Induction of a Gene Subset Important for Virulence in Alternaria brassicicola. Mol Plant Microbe Interact 25: 443–452. doi: 10.1094/mpmi-10-11-0275
  21. 21. Eliahu N, Igbaria A, Rose MS, Horwitz BA, Lev S (2007) Melanin biosynthesis in the maize pathogen Cochliobolus heterostrophus depends on two mitogen-activated protein kinases, Chk1 and Mps1, and the transcription factor Cmr1. Eukaryot Cell 6: 421–429. doi: 10.1128/ec.00264-06
  22. 22. Tsuji G, Kenmochi Y, Takano Y, Sweigard J, Farrall L, et al. (2000) Novel fungal transcriptional activators, Cmr1p of Colletotrichum lagenarium and Pig1p of Magnaporthe grisea, contain Cys2His2 zinc finger and Zn(II)2Cys6 binuclear cluster DNA-binding motifs and regulate transcription of melanin biosynthesis genes in a developmentally specific manner. Mol Microbiol 38: 940–954. doi: 10.1046/j.1365-2958.2000.02181.x
  23. 23. Kimura N, Tsuge T (1993) Gene cluster involved in melanin biosynthesis of the filamentous fungus Alternaria alternata. J Bacteriol 175: 4427–4435.
  24. 24. Thomma BPHJ (2003) Alternaria spp.: from general saprophyte to specific parasite. Mol Plant Pathol 4: 225–236. doi: 10.1046/j.1364-3703.2003.00173.x
  25. 25. Kawamura C, Tsujimoto T, Tsuge T (1999) Targeted disruption of a melanin biosynthesis gene affects conidial development and UV tolerance in the Japanese pear pathotype of Alternaria alternata. Mol Plant Microbe Interact 12: 59–63. doi: 10.1094/mpmi.1999.12.1.59
  26. 26. Rehnstrom AL, Free SJ (1996) The isolation and characterization of melanin-deficient mutants of Monilinia fructicola. Physiol Mol Plant Pathol 49: 321–330.
  27. 27. Wang Y, Casadevall A (1994) Decreased susceptibility of melanized Cryptococcus neoformans to UV light. Appl Environ Microbiol 60: 3864–3866.
  28. 28. Hyakumachi M, Yokoyma K, Ui T (1987) Role of melanin in susceptibility and resistance of Rhizoctonia solani to microbial lysis. Trans Br Mycol Soc 89: 27–33. doi: 10.1016/s0007-1536(87)80053-0
  29. 29. Rosas AL, Casadevall A (1997) Melanization affects susceptibility of Cryptococcus neoformans to heat and cold. FEMS Microbiol Lett 153: 265–272. doi: 10.1016/s0378-1097(97)00239-5
  30. 30. Howard RJ, Ferrari MA (1989) Role of melanin in appressorium function. Exp Mycol 13: 403–418. doi: 10.1016/0147-5975(89)90036-4
  31. 31. Moriwaki A, Kihara J, Kobayashi T, Tokunaga T, Arase S, et al. (2004) Insertional mutagenesis and characterization of a polyketide synthase gene (PKS1) required for melanin biosynthesis in Bipolaris oryzae. FEMS Microbiol Lett 238: 1–8. doi: 10.1111/j.1574-6968.2004.tb09729.x
  32. 32. Takano Y, Kubo Y, Kuroda I, Furusawa I (1997) Temporal transcriptional pattern of three melanin biosynthesis genes, PKS1, SCD1, and THR1, in appressorium-differentiating and nondifferentiating conidia of Colletotrichum lagenarium. Appl Environ Microbiol 63: 351–354.
  33. 33. Kubo Y, Takano Y, Tsuji G, Kenmochi Y, Kuroda I, et al. (1998) Expression and regulation of melanin biosynthetic genes during appressorium formation of Collectotrichum lagenarium. In: Kohmoto K, Yoder OC, editors. Molecular Genetics of Host Specific Toxins in Plant Disease. Dordrecht: Kluwer Academic Publishers. pp. 253–260.
  34. 34. Grigoriev IV, Nordberg H, Shabalov I, Aerts A, Cantor M, et al. (2012) The genome portal of the Department of Energy Joint Genome Institute. Nucleic Acids Res 40: D26–32. doi: 10.1093/nar/gkr947
  35. 35. Ohm RA, Bradshaw RE, Condon BJ, Feau N, Henrissat B, et al. (2012) Diverse lifestyles and strategies of plant pathogenesis encoded in the genomes of eighteen Dothideomycetes fungi. PLoS Pathog In Press. doi: 10.1371/journal.ppat.1003037
  36. 36. Bateman A, Coin L, Durbin R, Finn RD, Hollich V, et al. (2004) The Pfam protein families database. Nucleic Acids Res 32: D138–141. doi: 10.1093/nar/gkh121
  37. 37. You BJ, Choquer M, Chung KR (2007) The Colletotrichum acutatum gene encoding a putative pH-responsive transcription regulator is a key virulence determinant during fungal pathogenesis on citrus. Mol Plant Microbe Interact 20: 1149–1160. doi: 10.1094/mpmi-20-9-1149
  38. 38. Rollins JA (2003) The Sclerotinia sclerotiorum pac1 gene is required for sclerotial development and virulence. Mol Plant Microbe Interact 16: 785–795. doi: 10.1094/mpmi.2003.16.9.785
  39. 39. Arechiga-Carvajal ET, Ruiz-Herrera J (2005) The RIM101/pacC homologue from the basidiomycete Ustilago maydis is functional in multiple pH-sensitive phenomena. Eukaryot Cell 4: 999–1008. doi: 10.1128/ec.4.6.999-1008.2005
  40. 40. Lorang JM, Tuori RP, Martinez JP, Sawyer TL, Redman RS, et al. (2001) Green fluorescent protein is lighting up fungal biology. Appl Environ Microbiol 67: 1987–1994. doi: 10.1128/aem.67.5.1987-1994.2001
  41. 41. Kihara J, Moriwaki A, Tanaka N, Tanaka C, Ueno M, et al. (2008) Characterization of the BMR1 gene encoding a transcription factor for melanin biosynthesis genes in the phytopathogenic fungus Bipolaris oryzae. FEMS Microbiol Lett 281: 221–227. doi: 10.1111/j.1574-6968.2008.01101.x
  42. 42. Son H, Seo YS, Min K, Park AR, Lee J, et al. (2011) A phenome-based functional analysis of transcription factors in the cereal head blight fungus, Fusarium graminearum. PLoS Pathog 7: e1002310. doi: 10.1371/journal.ppat.1002310
  43. 43. Wong Sak Hoi J, Dumas B (2010) Ste12 and Ste12-like proteins, fungal transcription factors regulating development and pathogenicity. Eukaryot Cell 9: 480–485. doi: 10.1128/ec.00333-09
  44. 44. Penalva MA, Arst HN Jr (2004) Recent advances in the characterization of ambient pH regulation of gene expression in filamentous fungi and yeasts. Annu Rev Microbiol 58: 425–451. doi: 10.1146/annurev.micro.58.030603.123715
  45. 45. Flaherty JE, Pirttila AM, Bluhm BH, Woloshuk CP (2003) PAC1, a pH-regulatory gene from Fusarium verticillioides. Appl Environ Microbiol 69: 5222–5227. doi: 10.1128/aem.69.9.5222-5227.2003
  46. 46. Rollins JA, Dickman MB (2001) pH signaling in Sclerotinia sclerotiorum: identification of a pacC/RIM1 homolog. Appl Environ Microbiol 67: 75–81. doi: 10.1128/aem.67.1.75-81.2001
  47. 47. Caracuel Z, Roncero MI, Espeso EA, Gonzalez-Verdejo CI, Garcia-Maceira FI, et al. (2003) The pH signaling transcription factor PacC controls virulence in the plant pathogen Fusarium oxysporum. Mol Microbiol 48: 765–779. doi: 10.1046/j.1365-2958.2003.03465.x
  48. 48. Kihara J, Moriwaki A, Ito M, Arase S, Honda Y (2004) Expression of THR1, a 1,3,8-trihydroxynaphthalene reductase gene involved in melanin biosynthesis in the phytopathogenic fungus Bipolaris oryzae, is enhanced by near-ultraviolet radiation. Pigment Cell Res 17: 15–23. doi: 10.1046/j.1600-0749.2003.00102.x
  49. 49. Chumley F, Valent B (1990) Genetic analysis of melanin-deficient, nonpathogenic mutants of Magnaporthe grisea. Mol Plant Microbe Interact 3: 135–143. doi: 10.1094/mpmi-3-135
  50. 50. Kubo Y, Nakamura H, Kobayashi K, Okuno T, Furusawa I (1991) Cloning of a Melanin Biosynthetic Gene Essential for Appressorial Penetration of Colletotrichum lagenarium. Mol Plant Microbe Interact 4: 440–445. doi: 10.1094/mpmi-4-440
  51. 51. Howard RJ, Ferrari MA, Roach DH, Money NP (1991) Penetration of hard substrates by a fungus employing enormous turgor pressures. Proc Natl Acad Sci U S A 88: 11281–11284. doi: 10.1073/pnas.88.24.11281
  52. 52. Feng B, Wang X, Hauser M, Kaufmann S, Jentsch S, et al. (2001) Molecular cloning and characterization of WdPKS1, a gene involved in dihydroxynaphthalene melanin biosynthesis and virulence in Wangiella (Exophiala) dermatitidis. Infect Immun 69: 1781–1794. doi: 10.1128/iai.69.3.1781-1794.2001
  53. 53. Langfelder K, Jahn B, Gehringer H, Schmidt A, Wanner G, et al. (1998) Identification of a polyketide synthase gene (pksP) of Aspergillus fumigatus involved in conidial pigment biosynthesis and virulence. Med Microbiol Immunol 187: 79–89. doi: 10.1007/s004300050077
  54. 54. Guillen A, Turgeon BG, Thorson PR, Bronson CR, Yoder OC (1994) Linkage among melanin biosynthetic mutations in Cochliobolus heterostrophus. Fungal Genet Newslett 41: 40.
  55. 55. Karkehabadi S, Hansson H, Kim S, Piens K, Mitchinson C, et al. (2008) The first structure of a glycoside hydrolase family 61 member, Cel61B from Hypocrea jecorina, at 1.6 A resolution. J Mol Biol 383: 144–154. doi: 10.1016/j.jmb.2008.08.016
  56. 56. Lev S, Horwitz BA (2003) A mitogen-activated protein kinase pathway modulates the expression of two cellulase genes in Cochliobolus heterostrophus during plant infection. Plant Cell 15: 835–844. doi: 10.1105/tpc.010546
  57. 57. Cramer RA, La Rota CM, Cho Y, Thon M, Craven KD, et al. (2006) Bioinformatic analysis of expressed sequence tags derived from a compatible Alternaria brassicicola-Brassica oleracea interaction. Mol Plant Pathol 7: 113–124. doi: 10.1111/j.1364-3703.2006.00324.x
  58. 58. Noda J, Brito N, Gonzalez C (2010) The Botrytis cinerea xylanase Xyn11A contributes to virulence with its necrotizing activity, not with its catalytic activity. BMC Plant Biol 10: 38. doi: 10.1186/1471-2229-10-38
  59. 59. Fan C-Y, Köller W (1998) Diversity of cutinases from plant pathogenic fungi: differential and sequential expression of cutinolytic esterases by Alternaria brassicicola. FEMS Microbiol Lett 158: 33–38. doi: 10.1111/j.1574-6968.1998.tb12796.x
  60. 60. Anderson R, May R (1982) Coevolution of hosts and parasites. Parasitology 85: 411–426. doi: 10.1017/s0031182000055360
  61. 61. Sacristan S, Garcia-Arenal F (2008) The evolution of virulence and pathogenicity in plant pathogen populations. Mol Plant Pathol 9: 369–384. doi: 10.1111/j.1364-3703.2007.00460.x
  62. 62. Lenski RE, May RM (1994) The evolution of virulence in parasites and pathogens: reconciliation between two competing hypotheses. J Theor Biol 169: 253–265. doi: 10.1006/jtbi.1994.1146
  63. 63. Di Stasio M, Brefort T, Mendoza-Mendoza A, Munch K, Kahmann R (2009) The dual specificity phosphatase Rok1 negatively regulates mating and pathogenicity in Ustilago maydis. Mol Microbiol 73: 73–88. doi: 10.1111/j.1365-2958.2009.06747.x
  64. 64. Potter SC, Clarke L, Curwen V, Keenan S, Mongin E, et al. (2004) The Ensembl analysis pipeline. Genome Res 14: 934–941. doi: 10.1101/gr.1859804
  65. 65. Cho Y, Davis JW, Kim KH, Wang J, Sun QH, et al. (2006) A high throughput targeted gene disruption method for Alternaria brassicicola functional genomics using linear minimal element (LME) constructs. Mol Plant Microbe Interact 19: 7–15. doi: 10.1094/mpmi-19-0007
  66. 66. Yu JH, Hamari Z, Han KH, Seo JA, Reyes-Dominguez Y, et al. (2004) Double-joint PCR: a PCR-based molecular tool for gene manipulations in filamentous fungi. Fungal Genet Biol 41: 973–981. doi: 10.1016/j.fgb.2004.08.001
  67. 67. Hamer JE, Timberlake WE (1987) Functional organization of the Aspergillus nidulans trpC promoter. Mol Cell Biol 7: 2352–2359.
  68. 68. Malonek S, Rojas MC, Hedden P, Gaskin P, Hopkins P, et al. (2004) The NADPH-cytochrome P450 reductase gene from Gibberella fujikuroi is essential for gibberellin biosynthesis. J Biol Chem 279: 25075–25084. doi: 10.1074/jbc.m308517200
  69. 69. Trapnell C, Pachter L, Salzberg SL (2009) TopHat: discovering splice junctions with RNA-Seq. Bioinformatics 25: 1105–1111. doi: 10.1093/bioinformatics/btp120
  70. 70. Langmead B, Trapnell C, Pop M, Salzberg SL (2009) Ultrafast and memory-efficient alignment of short DNA sequences to the human genome. Genome Biol 10: R25. doi: 10.1186/gb-2009-10-3-r25
  71. 71. Trapnell C, Williams BA, Pertea G, Mortazavi A, Kwan G, et al. (2010) Transcript assembly and quantification by RNA-Seq reveals unannotated transcripts and isoform switching during cell differentiation. Nat Biotechnol 28: 511–515. doi: 10.1038/nbt.1621
  72. 72. Roberts A, Trapnell C, Donaghey J, Rinn JL, Pachter L (2011) Improving RNA-Seq expression estimates by correcting for fragment bias. Genome Biol 12: R22. doi: 10.1186/gb-2011-12-3-r22