New 1,3-benzodioxin-4-ones from Synnemapestaloides ericacearum sp. nov., a biosynthetic link to remarkable compounds within the Xylariales

Joey B. Tanney; Justin B. Renaud; J. David Miller; David R. McMullin

doi:10.1371/journal.pone.0198321

Abstract

Surveys of foliar endophytes from the Acadian forest region over the past three decades have identified numerous phylogenetically diverse fungi producing natural products toxic to forest pests and diseases. The life histories of some conifer endophytes can be restricted to plant foliage or may include saprotrophic phases on other plants tissues or even alternate hosts. Considering the potentially broad host preferences of conifer endophytes we explored fungi isolated from understory species and their metabolites as part of an ongoing investigation of fungal biodiversity from the Acadian forest. We report a hitherto unidentified Xylariomycetidae species isolated from symptomatic Labrador tea (Rhododendron groenlandicum) leaves and mountain laurel (Kalmia latifolia) collected in coastal southern New Brunswick, Canada. Morphological and phylogenetic evidence demonstrated the unknown species was a novel Synnemapestaloides (Sporocadaceae) species, described here as Syn. ericacearum. A preliminary screening assay indicated that the culture filtrate extract of the new species was potently antifungal towards the biotrophic pathogen Microbotryum violaceum, warranting an investigation of its natural products. Two natural products possessing a rare 1,3-benzodioxin-4-one scaffold, synnemadoxins A-B (1–2), and their postulated precursor, synnemadiacid A (3), were characterized as new structures and assessed for antimicrobial activity. All isolated compounds elicited in vitro inhibitory antifungal activity towards M. violaceum at 2.3 μg mL^-1 and moderate antibiotic activity. Further, the characterization of synnemadoxins A-B provided a perspective on the biosynthesis of some related 1,3-benzodioxin-4-ones produced by other fungi within the Xylariales.

Citation: Tanney JB, Renaud JB, Miller JD, McMullin DR (2018) New 1,3-benzodioxin-4-ones from Synnemapestaloides ericacearum sp. nov., a biosynthetic link to remarkable compounds within the Xylariales. PLoS ONE 13(6): e0198321. https://doi.org/10.1371/journal.pone.0198321

Editor: Petr Karlovsky, Georg-August-Universitat Gottingen, GERMANY

Received: February 7, 2018; Accepted: May 12, 2018; Published: June 27, 2018

Copyright: © 2018 Tanney et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: All relevant data are within the paper and its Supporting Information files. Synnemapestaloides ericacearum strains were deposited with the Canadian National Mycological Herbarium (DAOM). ITS and LSU sequences for studied strains were also accessioned with GenBank: Synnemapestaloides ericacearum holotype (DAOM 867445), and ex-type culture (DAOMC 251621), ITS barcode= MG687266 and LSU = MG687267. Synnemapestaloides ericacearum DAOMC 250336, ITS barcode= MG687268 and LSU = MG687269.

Funding: Funding was provided by the Natural Science and Engineering Research Council (NSERC) of Canada, MITACS, and JD Irving, Limited. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Foliar endophytes of conifers in the Acadian forest of Eastern Canada have proven an exceptionally rich source of structurally diverse biologically active natural products [1,2]. Our studies have revealed that the life histories of conifer endophytes can be restricted to plant foliage or may include saprotrophic phases on other host tissues or even alternate hosts. For example, Phialocephala scopiformis is a common Picea endophyte that produces its sexual reproductive structures (apothecia) on decaying Picea wood and fallen branches [3]. In contrast, apothecia of Phialocephala piceae, a foliar endophyte of Picea, are reported only from decaying hardwood [3–5]. A griseofulvin-producing Xylaria endophyte of Pinus strobus was also isolated as an endophyte of Vaccinium angustifolium, indicating a plurivorous life cycle involving both overstory and understory species [6].

While considering the potentially broad host preferences of conifer endophytes, an unidentified species in Xylariomycetidae was isolated from symptomatic Rhododendron groenlandicum leaves. The resulting strain, tentatively identified as a Seimatosporium species, generated a potently antifungal culture filtrate extract, prompting further investigation. Morphological and phylogenetic analyses indicated this strain, as well as a conspecific strain isolated from symptomatic Kalmia latifolia leaves, represented a novel Synnemapestaloides (Sporocadaceae) species. Herein, we describe the new species as Synnemapestaloides ericacearum, providing morphological and phylogenetic evidence supporting its distinction from Synnemapestaloides and other morphologically similar species within the Sporocadaceae. From the culture filtrate extract of Syn. ericacearum DAOMC 250336, three new natural products, including two metabolites possessing a rare 1,3-benzodioxin-4-one scaffold, were structurally characterized and assessed for antimicrobial activity.

Results

Strains from symptomatic leaves of Rhododendron groenlandicum (collected on Taylors Island, Saint John, NB, 45.212018, -66.136745) and Kalmia latifolia (see below) were conspecific based on morphology and identical ITS and LSU sequences. Both isolates were sterile on all media tested; however, sporulation by one strain (DAOMC 250336) was induced when malt extract agar (MEA) blocks containing mycelia were floated in sterile water. This species is morphologically similar to Seimatosporium (Sporocadaceae), a plurivorous genus including several species described from hosts in the Ericaceae, for example Seim. arbuti, Seim. ledi, and Seim. rhododendri on Rhododendron [7,8]. The ITS-LSU phylogeny places both strains in a strongly supported monophyletic clade sister to Synnemapestaloides rhododendri and Syn. foliicola (Fig 1). Based on morphological and phylogenetic evidence, we consider these two strains to represent a new species, described herewith as Synnemapestaloides ericacearum.

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Fig 1. Most likely tree from a RAxML analysis of ITS-LSU dataset containing representative Sporocadaceae species.

Culture collection accession numbers or specimen identifiers follow the species name, with sequences from type specimens or ex-type strains in bold. RAxML bootstrap support percentages ≥50 from a summary of 1000 replicates are presented at the branch nodes. Thickened branches indicate Bayesian posterior probability values ≥0.95. The tree was rooted with Phlogicylindrium uniforme CBS 131312 and the scale bar represents the number of substitutions per site.

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Based on a NCBI GenBank BLAST query using Syn. ericacearum ITS sequences, three unidentified endophyte sequences (Xylariomycetidae sp.) were included in the ITS-LSU phylogeny. These endophyte sequences are of strains isolated from surface-sterilized photosynthetic tissues of Equisetum arvense (JQ759638, JQ759542) and the Ericaceae host Cassiope tetragona (JQ759462) in Alaska, U.S.A. [9]. The unidentified endophyte ITS sequences were very similar to, and probably conspecific with, Syn. ericacearum, e.g.: JQ759638: identities = 734/738 (99%), gaps = 1/738 (0%).

Taxonomy

Synnemapestaloides ericacearum J.B. Tanney, sp. nov. (Figs 1 and 2)

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Fig 2. Synnemapestaloides ericacearum.

A, C, E. Leaf spots and acervuli on Rhododendron groenlandicum (DAOM 745782). B, D, F. Coalescing leaf spot and acervuli on Kalmia latifolia (DAOM 867445, holotype), triangular acervuli in F. G. Acervulus with intact host cuticle. H. Acervulus with ruptured host cuticle. I–K. Conidiophores and conidia, arrows denoting conidiogenous cells exhibiting obvious successive percurrent extensions. L–M. Conidia. Scale bars: G–M = 10 μm.

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MycoBank MB 824126

Typification. CANADA. NEW BRUNSWICK: Charlotte County, Roosevelt Campobello International Park, near Upper Duck Pond, 44.849370, -66.965173, from symptomatic Kalmia latifolia leaves, 26 Sep 2016, J.B. Tanney NB-846 (holotype DAOM 867445). Ex-type culture DAOMC 251621.

Etymology: Named for its association with Ericaceae hosts.

ITS barcode: MG687266 (LSU = MG687267)

Leaf spots mostly epiphyllous, sometimes hypophyllous, punctiform to effused, circular to lobate, discrete, eventually becoming irregularly confluent and covering most of the leaf lamina, distinct on upper surface but less conspicuous on lower surface, brown, delimited from healthy leaf tissue by thin purple-brown margin.

Colonies 40–45 mm diam after 7 d in 12:12 h light/dark at 20 °C on MEA; flat, sparse to moderately abundant hyaline aerial mycelia, margin entire, flat, wide, hyaline; surface and reverse white to reddish blond (5C4) in center. Exudates and soluble pigments absent. Mycelium consisting of hyaline, smooth, septate, branched, hyphae 2–8 μm diam.

Conidiomata acervular, pulvinate, circular or triangular in outline, up to 275 μm diam, blackish-brown to black, glabrous, epiphyllous, subcuticular, scattered, initially covered by the host cuticle, which becomes raised then irregularly ruptures laterally or radially to release black mass of conidia; basal stroma well-developed, up to 30 μm thick, pale brown textura angularis composed of thick-walled, pale brown cells (3–)4–6(–7.5) μm diam, inner wall becoming hyaline with thinner-walled cells.

Conidiophores aggregated into a compact layer, up to 30–55 μm long, 2–3(–5.5) μm wide, septate, branched, hyaline, smooth, thin-walled. Conidiogenous cells holoblastic, annellidic, cylindrical to slightly ampuliform, hyaline, smooth, thin-walled, bearing up to six successive percurrent extensions, (7–)8.5–11.5(–12.5) × 2–2.5(–3) μm (length: n = 30, μm, SD = 1.5 μm, SE = 0.29 μm, 95% CI = 0.57; width: n = 30, μm, SD = 0.3 μm, SE = 0.05 μm, 95% CI = 0.1).

Conidia fusiform-ellipsoid to fusiform-cylindrical or clavate, smooth, 3-septate, pale brown to brown, concolorous when young, becoming versicolorous with age, (10.5–)11.5–13.5(–16.5) × (3.5–)4–5(–5.5) μm (length: n = 73, μm, SD = 1 μm, SE = 0.12 μm, 95% CI = 0.24; width: n = 73, μm, SD = 0.36 μm, SE = 0.04 μm, 95% CI = 0.08); apical cell elongate-conic, sometimes apiculate, usually same wall-thickness but sometimes paler than median cells, (2–)3–4(–4.5) μm long (n = 55, μm, SD = 0.5 μm, SE = 0.07 μm, 95% CI = 0.14); two median cells doliiform to subcylindrical, moderately thick-walled, pale brown to brown; penultimate cell 2.5–3.5(–4) μm long (n = 55, μm, SD = 0.5 μm, SE = 0.06 μm, 95% CI = 0.13); antepenultimate cell (2.5–)3–3.5(–4) μm long (n = 55, μm, SE = 0.4 μm, SE = 0.05 μm, 95% CI = 0.10); basal cell obconic, thin-walled, subhyaline or occasionally hyaline, subtruncate to truncate with unthickened, 1–1.5 μm diam basal scar that is sometimes inconspicuous, (2–)2.5–3.5(–4) μm long (n = 55, μm, SD = 0.4 μm, SE = 0.06 μm, 95% CI = 0.12); appendages absent.

Host range: Causing leaf spots on Kalmia latifolia and Rhododendron groenlandicum.

Distribution: Canada (New Brunswick).

Additional specimens and cultures examined: CANADA. NEW BRUNSWICK: Saint John County, Saint John, Taylors Island, 45.212018, -66.136745, from symptomatic Rhododendron groenlandicum leaves, 26 Sep 2016, A.K. Walker DAOM 745782/DAOMC 250336, ITS barcode: MG687268 (LSU = MG687269).

Based on ITS-LSU phylogeny, Synnemapestaloides ericacearum is sister to a clade containing Syn. foliicola and Syn. rhododendri. Synnemapestaloides rhododendri produces synnemata that give rise to five-septate conidia with appendages [10], while Syn. ericacearum produces three-septate conidia lacking appendages from acervuli. Synnemapestaloides foliicola differs from Syn. ericacearum by its five-septate conidia bearing terminal appendages produced from sporodochia with similar ontogeny to that of Syn. rhododendri [10,11]. The absence of conidial appendages distinguishes Syn. ericacearum from some other Seimatosporium species reported from Ericaceae hosts, e.g.: Seim. arbuti and Seim. azalea conidia possess terminal appendages [12,13] and Seim. ledi conidia possess basal appendages [7]. Seimatosporium rhododendri conidia are three-septate and unappendaged, but significantly larger than those of Syn. ericacearum (15.5–20 × 6.5–8.5 μm) [7]. Similarly, the conidia of Seim. vaccinii are three-septate and unappendaged, but are somewhat longer (13–18 × 4.5–5.5 μm) than those of Syn. ericaearum, and also born from shorter conidiophores (up to 28 μm long vs. 30–55 μm long) [14]. The mostly three-septate, unappendaged conidia of Seim. lichenicola are similar to those of Syn. Ericacearum; however, they are considerably larger: 18–20 × 5–7 um versus (10.5–)11.5–13.5(–16.5) × (3.5–)4–5(–5.5) μm [15], although Sutton [14] reported Seim. lichenicola conidial dimensions more similar to those of Syn. ericacearum: 13–15 × 4.4–6.5 μm. Norphanphoun et al. [16] designated a reference specimen of Seim. lichenicola from Cotinus coggygria with conidial dimensions similar to Syn. ericacearum and Seim. lichenicola sensu Sutton [14], (10–)12–14 × 4–5(–6) μm; however, it is phylogenetically distinct.

Natural products

The incubation of Syn. ericacearum DAOMC 250336 in 2% malt extract broth yielded ethyl acetate culture filtrate extracts that exhibited strong in vitro antifungal activity using a modified Oxford diffusion assay. Three new natural products including two rare 1,3-benzodioxin-4-ones were structurally characterized and assessed for antimicrobial activity (Fig 3). The first metabolite (1) purified was isolated as a dull brown oil with the molecular formula C₁₅H₁₈O₆ determined by an [M-H]—ion at m/z 293.1033 indicative of seven units of unsaturation. Its UV spectrum showed absorption maxima at 224, 268, and 306 nm suggesting the presence of a conjugated structure. Its ¹H NMR spectrum displayed a single aromatic methine at δ 6.51 (s), deshielded methine at δ 3.18 (q, 7.0), and five methyl groups at δ 2.58 (s), 2.11 (s), 1.73 (s), and 1.28 (d, 7.0) as well as a methoxy group at δ 3.87 (s). Interpretation of the HSQC and ¹³C NMR spectra of 1 revealed eight of the fifteen carbon signals were quaternary. These signals were resultant of two carbonyls at δ 175.4 and 161.9, five sp² carbons at δ 165.0, 157.7, 143.2, 122.7, and 106.2 and one sp³ carbon at δ 107.2 (Table 1). The presence of eight highly deshielded quaternary carbons and four singlet methyl groups supported a highly substituted, conjugated structure. As expected from the ¹H spectrum, only one COSY cross-peak was observed for compound 1, between δ 1.28 (9-Me) and δ 3.18 (H-9; Fig 4). HMBC correlations from the singlet methyl at δ 1.73 (2-Me) to δ 107.2 (C-2) and 47.8 (C-9), the cross peak from 9-Me to the carbonyl at δ 175.4 (C-10) and the chemical shift of C-2 suggested the presence of a 3,3-dioxy-2-methylbutanoic acid moiety in 1 (Fig 4). The presence of a single carboxylic acid (C-10) was supported by methylation of 1 with excess diazomethane that yielded a hexamethyl derivative (1a), identifying one exchangeable acidic proton within the structure (Figure J and Table E in S1 File). The high number of deshielded quaternary carbons and extensive HMBC coupling for 1 suggested a pentasubstituted aromatic ring. A lactone or other ring system fused to the aromatic moiety possessing a carbonyl at δ 161.9 (C-4) was theorized to constitute the remaining degrees of unsaturation. HMBC correlations from the singlet methyl at δ 2.11 (6-Me) to δ 143.2 (C-5), 122.7 (C-6) and (C-7), and from δ 3.87 (7-OMe) to δ 165.0 (C-7) and 98.1 (C-8) revealed a methoxy functionality vicinal to the only aromatic methine proton. HMBC cross-peaks were also observed from δ 2.58 (5-Me) to δ 161.9 (C-4), 106.2 (C-4a) and C-5, and from δ 6.51 (H-8) to C-4a, C-6, C-7 and C-8a (δ 157.7; Fig 4). These data and the chemical shift of C-8a revealed that compound 1 possess a modified 6-hydroxy-4-methoxy-2,3-dimethylbenzoic acid core structure. Together, the chemical shift of C-2, lack of other exchangeable acidic protons and an HMBC correlation from 2-Me to C-4 fuses the previously characterized 3,3-dioxy-2-methylbutanoic acid moiety to the aromatic functionality yielding a rare 1,3-benzodioxin-4-one structure (Figs 3 and 4). The 1,3-dioxin-4-one ring satisfies the seventh unit of unsaturation dictated by the molecular formula and NMR data.

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Table 1. ¹H (400 MHz) and ¹³C (100 MHz) NMR data for synnemadoxins A-B (1–2) in CD₃OD.

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Fig 3. Structures of new 1,3-benzodioxin-4-ones synnemadoxins A-B (1–2) and synnemadiacid A (3) characterized from Syn. ericacearum DAOMC 250336.

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Fig 4. Observed COSY (bold) and key HMBC (blue) correlations for synnemadoxin A (1) and synnemadiacid A (3).

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The planar 1,3-benzodioxin-4-one structure of 1 was elucidated as a new natural product; however, the configurations of the adjacent chiral centers at C-2 and C-9 remained elusive. Due to the lability of the 3,3-dioxy-2-methylbutanoic acid moiety, conventional NOESY experiments used to resolve stereochemical assignments were not discriminatory. Attempts to generate crystals of compound 1 or its potassium salt were also not successful. Chiral HPLC of 1 revealed an approximate 55:45 mixture of enantiomers, dominated by the positive enantiomer according to the optical rotation values, [α]²³_D 9.4 (c 0.3, MeOH), [α]²³_D 9.8 (c 0.3, CHCl₃). To determine the relative configurations of C-2 and C-9, the in silico approach described by Willoughby et al. 2017 [17] was used. Briefly, molecular dynamics were calculated with AMBER (University of California) to generate a library of all possible conformers for each stereochemical configuration. Density function theory was used to calculate frequency and free energy values for each conformation to generate in silico computed chemical shifts. Comparisons of mean absolute error (MAE) between experimental and computed chemical shifts of all reported ¹H and ¹³C resonances for 1 and its methyl ester 1a revealed the relative configurations of the enantiomeric mixture of 1 as 2R,9R and 2S,9S (Tables A-D in S1 File). This in silico approach is not able to differentiate between enantiomers of 1; however, is discriminatory of diastereomers. The HRMS and spectroscopic data support 1 as a new rare 1,3-benzodioxin-4-one structure reported here as synnemadoxin A (Fig 3).

The second metabolite characterized from Syn. ericacearum DAOMC 250336 was also isolated as dull brown oil with the same UV absorption maxima as synnemadoxin A (1). Metabolite 2 was more polar and assigned the molecular formula C₁₅H₁₈O₇ based on an HRMS peak at m/z 309.0981 [M-H]^- corresponding to the addition of an oxygen atom compared to synnemadoxin A (1). The NMR spectroscopic data for 2 were very similar to synnemadoxin A (1), indicating 2 was a structurally similar 1,3-benzodioxin-4-one metabolite (Table 1). The primary difference between the two 1,3-benzodioxin-4-ones NMR data was the substitution of a methyl group in synnemadoxin A (1) with an oxygenated methylene in 2 revealed by ¹H and ¹³C signals at δ 4.70 (s) and δ 55.0, respectively (Table 1). Together, the differences in NMR chemical shifts, increase in molecular weight, and HMBC correlations from δ 4.70 (6-CH₂OH) to δ 145.7 (C-5), 125.4 (C-6) and 165.6 (C-7), are indicative of C-6 bearing a hydroxymethyl moiety in 2 instead of a methyl group (Fig 3). The two 1,3-benzodioxin-4-ones (1–2) similar ¹H NMR chemical shift data, and positive optical rotation values, [α]²³_D 10.4 (c 0.3, MeOH) for 2, are evidence for the adjacent chiral centers having the same configurations as synnemadoxin A (1). Here we report 2 as a new structure, synnemadoxin B.

Compound 3 was purified as the major constituent of the Syn. ericacearum DAOMC 250336 culture filtrate extract as a yellow oil with the same molecular formula as synnemadoxin A (1) determined by an HRMS peak at m/z 293.1032 [M-H]^-. Based on resonances in the ¹H and ¹³C NMR spectra, compound 3 possesses a pentasubstituted aromatic ring similar to synnemadoxin A (1) where primary differences in the NMR spectra of synnemadoxin A (1) and 3 appeared in the acyclic moiety (Tables 1 and 2). Chemical shifts for 3 at δ 162.8 (C-1’) and 113.0 (C-2’), the absence of an NOESY correlation between vicinal methyl protons at δ 2.20 (1’-Me) and 1.90 (2’-Me), and magnitude of their coupling constant (J = 1.4 Hz) reveals vicinal methyl protons trans- configuration on a tetrasubstituted double bond (Fig 3). An additional carbonyl carboxylic acid chemical shift at δ 171.8 (C-1”) and trans-double bond together with the HRMS data, and similar NMR spectra suggested a reduction of the 1,3-dioxin-4-one ring in 3 (Fig 4). Here were report compound 3 as a new structure, (E)-6-((3-carboxybut-2-en-2-yl)oxy)-4-methoxy-2,3-dimethylbenzoic acid, or synnemadiacid A (Fig 3).

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Table 2. ¹H (400 MHz) and ¹³C (100 MHz) NMR data for synnemadiacid A (3) CD₃OD.

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Natural products 1–3 were individually tested for in vitro antifungal activity against Microbotryum violaceum and Saccharomyces cerevisiae, and antibiotic activity towards Bacillus subtilis and Escherichia coli (Table 3). Synnemadoxins A-B (1–2) and synnemadiacid A (3) were potently inhibitory to M. violaceum (formerly Ustilago violacea), a proxy for rust needle diseases [18], each with a MIC of 2.3 μg mL^-1. However, no activity was observed for each metabolite at the highest concentration tested (150 μg mL^-1) against S. cerevisiae despite the crude culture filtrate extract showing activity at 50 mg mL^-1 using a modified Oxford diffusion assay. Moderate antibiotic activity for compounds 1–3 against B. subtilis were observed at 9.3 μg mL^-1; and 1 and 3 had MIC values of 18.8 μg mL^-1 against E. coli.

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Table 3. Minimum inhibitory concentrations for 1–3 isolated from Syn. ericacearum DAOMC 250336.

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Discussion

Synnemapestaloides was established to describe Syn. rhododendri, the causal agent of a leaf and twig blight disease of Rhododendron brachycarpum in Japan [10]. Handa et al. [10] provided morphological evidence distinguishing the monotypic Synnemapestaloides from other pestaloid genera, namely the presence of columnar synnemata. A later phylogenetic study confirmed the distinction of Synnemapestaloides from Seimatosporium and other Sporocadaceae genera, while also establishing the new combination Syn. foliicola for a presumably saprotrophic species known from Chamaecyparis and Juniperus, previously placed in Seimatosporium [19,20]. The unappendaged three-septate conidia of Synnemapestaloides ericacearum distinguish it from the five-septate conidia of both Syn. foliicola and Syn. rhododendri. Synnemapestaloides rhododendri conidia are fusiform and more-or-less pestaloid, with unbranched or dichotomously branched apical appendages and unbranched or irregularly branched basal appendages, while Syn. foliicola conidia are broadly ovate with unbranched apical and basal appendages [10,20]. Watanabe et al. [19] described similar conidiomata ontogeny for both Syn. rhododendri, which produces up to ca. 500 μm tall synnemata, and Syn. foliicola, which produces short synnemata (sporodochia). The authors hypothesized that this conidiomata ontogeny might be a synapomorphy for Synnemapestaloides. We interpret the conidiomata of Syn. ericacearum as acervular, with the conidiogenous cells and conidia observed under the ruptured leaf cuticle (Fig 2G and 2H). However, detailed time-lapse observations would be necessary to elucidate the conidiomata ontogeny of Syn. ericacearum. There are no reports of sexual states in Synnemapestaloides.

Currently, Syn. ericacearum is only known as an associate of leaf spots on the Ericaceae hosts Kalmia latifolia and Rhododendron groenlandicum. However, it is conceivable that the life cycle of Syn. ericacearum contains an endophytic phase, which is common for species from the Sporocadaceae (e.g., [13, 21, 22]). The association of Syn. ericacearum with leaf spots characteristic of those caused by Syn. rhododendri, Seimatosporium, and other Sporocadaceae species indicate it is likely a pathogen, although its pathogenicity was not tested.

Many strains of fungi within Sporocadaceae representing the genera Pestalotiopsis, Bartalinia, Monochaetia, Robillarda, Seimatosporium, and Strickeria have been critically evaluated for natural products [23]. This interest resulted from the putative identification of the plant anticancer compound paclitaxel (taxol) from Taxomyces andreanae, collected as an endophyte of Taxus brevifolia, and subsequently from several related species including endophytic strains of Bartalinia, Monochaetia, and Pestalotiopsis [24]. Horizontal transfer of biosynthetic genes from plants to fungi was postulated to explain paclitaxel production by these fungi. However, more recent evidence shows the genomes of the putative endophyte producers’ do not possess any significant homology to plant paclitaxel biosynthetic genes, questioning its biosynthesis by these fungi [25].

Strains of species within Sporocadaceae collected as endophytic or plant-associates have proved to be an abundant source of structurally diverse natural products. The polyhydroylated macrolides seimatopolides A and B with peroxisome proliferative activity were characterized from a Seimatosporium discosioides (GenBank: EF600969) strain isolated from Rosa multiflora [26,27]. A Seimatosporium endophyte of Salsola oppositifolia synthesized several aromatic compounds acid together with metabolites structurally similar to terreic acid, an antiinsectan metabolite isolated from a Picea endophyte [1,28]. From the same plant host, the acaranoic acids, seimatoporic acid A-B were reported together with (R)-mellein and five of its derivatives [29]. The simple isocoumarin (R)-mellein and its derivatives have been reported from phylogenetically diverse plant-associated fungi including Pezicula, Diaporthe and Rhytismatales species [30–34]. Pestalotiopsis, Robillarda and other related but unidentified species likely associated with woody plants (collected from three-toed sloth hair) generated cytotoxic and antibiotic culture filtrate extracts; however, metabolite structures were not elucidated [35].

Fungal natural products possessing a 1,3-benzodioxin-4-one scaffold present within synnemadoxins A-B (1–2) are uncommon. An unidentified Xylaria isolate collected from the Malaysian rain forest produced the chlorinated spirocyclohexadienone maladoxin which contains a 1,3-benzodioxin-4-one moiety together with the related chlorinated depsidone maldoxone and diphenyl ether biosynthetic intermediates dihydromaldoxin (= pestheic acid), isodihydromaldoxin, and dechlorodihydromaldoxin (Fig 5A) [36]. A Pestalotiopsis fici (Sporocadaceae) strain isolated as an endophyte of Camellia sinensis (Ericales) related to Syn. ericacearum biosynthesizes the most compounds reported to date containing this uncommon structural element (Fig 5) [37]. Genomic data for this strain of P. fici (GenBank: ARNU00000000) strain revealed a high number of extracellular pectinases. Pectin is a major carbohydrate between plants cells suggesting adaptation to its endophytic state [37]. P. fici metabolites possessing the 1,3-benzodioxin-4-one core structure include chloropestolides A-G [38,39], chloropupukeanolides A-E [40,41], and dechloromaldoxin [39]. Much of the natural product diversity from this strain (> 80 metabolites) is thought to arise from the stereo-unspecific intermolecular Diels-Alder cycloaddition of intermediates followed by carbonyl-ene reactions [37,39,41]. Proposed intermediates include the same biphenyl ether, dihydromaldoxin (= pestheic acid), and the sesquiterpene (+)-iso-A82775C, isolated together with larger 1,3-benzodioxin-4-one containing metabolites (Fig 5B) [39,41]. The disruption of epigenetic regulators in this P. fici strain resulted in the characterization of additional new polyketides including eleven new macrolides and metabolites sharing structural features of the proposed precursors noted above [42]. Due to the unusual, highly functionalized structures and reported biological activities of P. fici metabolites possessing a 1,3-benzodioxin-4-one scaffold, their biogenesis has been critically evaluated [39,41,43,44]. Similar Diels-Alder biosynthetic schemes have also been reported for sorbicillin-derived metabolites from the Trichoderma section Longibrachiatum [45–47].

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Fig 5.

A) Proposed biosynthesis of maladoxin from the oxidation of dihydromaldoxin by a Xylariales fungus (unidentified Xylaria sp.; adapted from Adeboya et al. 1996). B) Proposed biosynthetic scheme of chloropestolides from P. fici from the oxidation of the diphenyl ether pestheic acid to maladoxin followed by the Diels-Alder addition of iso-A82775C (adapted from Liu et al. 2013). C) Proposed biosynthesis of synnemadoxin A (1) by Syn. ericacearum from the oxidation of synnemadiacid A (3), analogous to the schemes proposed for maldoxin from and unidentified Xylaria sp. (Xylariales) and P. fici.

https://doi.org/10.1371/journal.pone.0198321.g005

The biogenesis of these highly functionalized P. fici metabolites are proposed to originate with the oxidation of dihydromaldoxin (= pestheic acid) to maladoxin, which possesses a reactive diene functionality. Diels-Alder cascades from the reaction of the maladoxin diene and terminal alkene of (+)-iso-A82775C generate the observed structural diversity for this strain [37]. Due to the reactivity of the maladoxin diene moiety, the Diels-Alder reaction in the presence of the terminal diene of (+)-iso-A82775C (Fig 5B) is speculated to occur almost spontaneously, without major enzymatic involvement. Maladoxin was not isolated from rice culture extracts of P. fici, whereas the less reactive dechloro congener was [41]. The recovery of maladoxin from the mycelium of the previously noted Xylariales (unidentified Xylaria sp.) species together with dihydromaldoxin (= pestheic acid) and other related diphenyl ether intermediates suggests a similar biosynthetic pathway, absent the presence of a reactive diene moiety or additional enzymatic input [36]. Since 1,3-benzodioxin-4-one containing natural products are rarely reported, we speculate synnemadoxins A-B (1–2) arise from the oxidation of synnemadiacid A (3) similar to the oxidation of diphenyl ether intermediates to maladoxin (Fig 5A–5C) by related fungi within the Xylariales [36,39,41]. Oxidative cyclization of the diphenyl ether dihydromaldoxin is reported to yield enantiomeric mixtures of the 1,3-benzodioxin-4-ones maldoxin and dechloromaldoxin [39,48]. A similar oxidative scheme using synnemadiacid A (3) as a precursor would help explain the observation of enantiomeric synnemadoxins A-B (1–2).

In summary, we have described a new species, Syn. ericacearum (Sporocadaceae) from the ericaceous hosts R. groenlandicum and Kalmia latifolia. The culture filtrate extract from Syn. ericacearum DAOMC 250336 was potently antifungal. Two natural products possessing a rare 1,3-benzodioxin-4-one scaffold, synnemadoxin A-B (1–2), and their postulated precursor, synnemadiacid A (3), were characterized as new structures and assessed for biological activity. All isolated compounds elicited in vitro inhibitory antifungal activity towards M. violaceum at 2.3 μg mL^-1 and moderate antibiotic activity. This study helps to link rare 1,3-benzodioxin-4-ones produced by Syn. ericacearum and other fungi within the Xylariales.

Materials and methods

General experimental section

NMR spectra of purified metabolites were recorded with a Bruker Avance 400 Spectrometer (Milton, Ontario) at 400.1 (¹H) and 100 MHz (¹³C) using a 5 mm auto-tuning broadband probe with a Z-gradient. Secondary metabolites were dissolved in CD₃OD (CDN Isotopes, Point Claire, Quebec) and referenced to the solvent peak (δ_H 3.31 and δ_C 49.1). LC-HRMS spectra were acquired with a Thermo Q-Exactive Orbitrap mass spectrometer (Thermo Scientific, Waltham, MA), coupled to an Agilent 1290 HPLC system. Extracts and metabolites were separated by an Agilent Zorbax Eclipse Plus, RRHD C₁₈ (2.1 x 50 mm, 1.8 μm) column and a mobile phase consisting of acetonitrile (ACN) -ddH₂O with 0.1% formic acid at a flow rate of 0.3 mL min^-1. Normal phase and semi-preparative fractions were screened by LC-UV-MS with a Waters 2795 separation module, Waters 996 diode array detector, and Micromass Quatro LC mass spectrometer. Fractions were separated by a Phenomenex Kinetix C₁₈ (100 × 4.60 mm, 2.6 μm) column (Torrance, CA, USA) using a mobile phase also consisting of ACN-ddH₂O with 0.1% formic acid at a flow rate of 1.0 mL min^-1. Semi-preparative HPLC was achieved with an Agilent 1100 HPLC system equipped with a diode array detector, Phenomnex Luna C₁₈ (250 x 10.00 mm, 5 μm) column, and a mobile phase consisting of ACN-ddH₂O. Gradients were programmed accordingly for each purified compounds with a flow rate of 4 mL min^-1. Silica gel (Silicycle; 40–63 μm) was utilized for normal phase flash chromatography. Optical rotations were acquired with an Autopol IV polarimeter (Rudolph Analytical, Hackettown) and UV spectra were recorded with a Varian Cary 3 UV-VIS spectrophotometer scanning from 190–800 nm.

Isolation and identification of fungal strains

Two collections of symptomatic leaves of Kalmia latifolia and Rhododendron groenlandicum were made during an ongoing survey of endophytes and other plant-associated fungi in Eastern Canada. Conidia from acervuli associated with leaf spots were transferred to Petri dishes containing 2% malt extract agar (MEA; 20 g Bacto malt extract, Difco Laboratories, Sparks, Maryland; 15 g agar, EMD Chemicals Inc., New Jersey; 1 L distilled water) and incubated at 20 °C. To induce sporulation, DAOMC 250336 was also grown on cornmeal agar (CMA; Acumedia Manufacturers Inc., Lansing, MI), oatmeal agar [49], and water agar (1.5% water agar (WA, with 1 mL trace metal solution; [50]) under 12:12 h fluorescent light and ambient natural light conditions. Additionally, MEA blocks (ca. 1 cm³) containing mycelia were placed in 9 cm Petri dishes containing ca. 25 μL sterile autoclaved tap water and exposed to ambient light.

Total genomic DNA was extracted from MEA cultures using the Ultraclean Microbial DNA isolation kit (Mo Bio Laboratories, Carlsbad, CA, USA) following the manufacturer’s protocol. The ITS region was amplified and sequenced using the primer pair V9G and LS266 [51,52]. Partial LSU was amplified using the primer pair LROR and LR5 and sequenced using LROR, LR3, LR3R, and LR5 [53]. PCR and sequencing protocol were carried out according to McMullin et al. [33]. Sequence contigs were assembled and trimmed using Geneious R8 v. 8.1.5 (Biomatters, Auckland, New Zealand).

ITS and LSU sequences of related Sporocadaceae species were obtained from GenBank by conducting BLAST searches using DAOMC 250336 and DAOMC 251621 as query sequences. A concatenated ITS and LSU dataset containing 59 sequences was aligned using MAFFT v. 7 [54] and visually examined in Geneious. MrModeltest v. 2.2.6 was used to determine the most suitable sequence evolution model (GTR+I+G) using the Akaike information criterion [55]. The ex-type culture of Phlogicylindrium uniforme (CBS 131312; Phlogicylindriaceae) was selected as outgroup because of its phylogenetic position outside of Sporocadaceae [19]. MrBayes v. 3.2 was used to perform Bayesian inference (BI) phylogenetic reconstruction [56]. Three independent Metropolis-coupled Markov chain Monte Carlo (MCMCMC) samplings were performed with 11 heated chains and one cold chain. Sampling occurred every 500 generations until the standard deviation of split frequencies reached a value < 0.01 (commands: stoprule = yes stopval = 0.01). The first 25% of trees were discarded as burn-in and the remaining trees were retained and combined into one 50% majority rule consensus tree. A maximum likelihood (ML) analysis was performed using RAxML v. 8.2.4 [57] in PAUP* v. 4.0b10 [58] starting from a random starting tree with 1000 bootstrap replicates. Trees were visualized in FigureTree 1.4.2 (available at http://tree.bio.ed.ac.uk/software/Figuretree/) and exported as SVG vector graphics for assembly in Adobe Illustrator v10 (Adobe Systems, San Jose, CA). Novel sequences used in this study were accessioned in GenBank and the novel species and associated metadata were deposited in MycoBank (www.mycobank.org). Collecting on Campobello Island was part of the 2016 Campobello Island Mycological Foray hosted by the New Brunswick Museum and J.D. Irving, Limited gave permission to conduct fungal collections on their sites.

Nomenclature

The electronic version of this article in Portable Document Format (PDF) in a work with an ISSN or ISBN will represent a published work according to the International Code of Nomenclature for algae, fungi, and plants, and hence the new names contained in the electronic publication of a PLOS ONE article are effectively published under that Code from the electronic edition alone, so there is no longer any need to provide printed copies. In addition, new names contained in this work have been submitted to MycoBank from where they will be made available to the Global Names Index. The unique MycoBank number can be resolved and the associated information viewed through any standard web browser by appending the MycoBank number contained in this publication to the prefix http://www.mycobank.org/MB/. The online version of this work is archived and available from the following digital repositories: PubMed Central, LOCKSS.

Fermentation and extraction

For inoculation into culture medium, Syn. ericacearum DAOMC 250336 growing on a 2% malt extract agar (MEA; Difco Laboratories) was excised and macerated in sterile ddH₂O under aseptic conditions. Aliquots were used to inoculate (5% v/v) fifteen 250 mL Erlenmeyer flasks each containing 50 mL of 2% malt extract (Difco Laboratories) broth. First stage cultures were incubated for one week on a rotary shaker (100 RPM) in the dark at 25 °C. Resulting mycelia were macerated under aseptic conditions, and individually transferred to Glaxo bottles containing 1 L of the same culture medium. Second stage cultures were incubated stationary as described above for eight weeks.

After the incubation period, Syn. ericacearum mycelia were separated from the culture filtrate by suction through Whatman #4 (Whatman GE Healthcare, UK) filter papers. Mycelia were lyophilized and stored at -20 °C. The culture filtrate was saturated with NaCl and extracted with ACS grade EtOAc. The aqueous culture filtrate was discarded and the organic layer was filtered through anhydrous Na₂SO₄ and Whatman #1 filter papers prior to drying by rotary evaporation. The resulting culture filtrate extracts were dissolved in HPLC grade MeOH, passed through 0.2μm PTFE (25 mm) syringe filters (Tisch Scientific, USA), dried under a gentle stream of nitrogen gas and weighed. The resulting culture filtrate extract (1.20 g) was stored dry in an amber vial in a freezer at -20 °C.

Metabolite isolation

The culture filtrate extract of Syn. ericacearum DAOMC 250336 was assessed for in vitro antifungal activity with a modified Oxford disc assay and subsequently chromatographed using a bioassay-guided approach. Flash column chromatography implementing a short Silica column and step gradient elution system of hexanes-EtOAc (0–100% v/v) in 10 percent increments followed by 5%, 10%, 25% and 50% EtOAc-MeOH (v/v) yielded 14 fractions screened for antifungal activity. Fractions 5–8, and 12–14 all exhibited antifungal activity with the former group showing a greater inhibitory effect in vitro. Based on LC-UV-MS, fractions with similar LC-UV-MS spectra were combined and metabolites were purified by reverse phase semi-preparative HPLC. Fractions 6–7 which eluted with 60–70% hexanes-EtOAc (v/v) were combined (48 mg) and further separated by semi-preparative HPLC with a linear ACN-ddH₂O gradient programmed from 5–100% ACN over 25 minutes to afford compound 1 (14.9 mg). Compound 3 (46.0 mg) and additional compound 1 used for chemical derivatization were purified using the same semi-preparative HPLC method from fraction 8 (149.0 mg) that eluted with 80% hexanes-EtOAc. Fraction 12 (39.0 mg) eluted with 10% MeOH-EtOAc and provided compound 2 (6.9 mg) prior to further purification with a linear semi-preparative HPLC method programmed from 5–100% ACN over 20 minutes. ¹H, ¹³C and HMBC spectra for natural products (1–3) can be found in the supporting information (Figures A-I in S1 File).

Synnemadoxin A (1): 14.9 mg; dull brown oil; [α]²³_D 9.4 (c 0.3, MeOH), [α]²³_D 9.8 (c 0.3, CHCl₃); UV (MeOH) λ_max (log ε) 224 (4.23), 268 (3.84), 306 (3.23); for ¹H and ¹³C NMR spectroscopic data, see Table 1; HRMS m/z 293.1033 [M-H]^- (calc. for [C₁₅H₁₇O₆]^- 293.1031).
Synnemadoxin B (2): 6.9 mg; dull brown oil; [α]²³_D 10.4 (c 0.3, MeOH), [α]²³_D 10.2 (c 0.3, CHCl₃); UV (MeOH) λ_max (log ε) 224 (4.34), 268 (3.58), 306 (3.21); for ¹H and ¹³C NMR spectroscopic data, see Table 1; HRMS m/z 309.0981 [M-H]^- (calc. for [C₁₅H₁₇O₇]^- 309.0980).
Synnemadiacid A (3): 46.0 mg; yellow oil; UV (MeOH) λ_max (log ε) 221 (4.12), 270 (3.53), 310 (3.14); for ¹H and ¹³C NMR spectroscopic data, see Table 2; HRMS m/z 293.1032 [M-H]^- (calc. for [C₁₅H₁₇O₆]^- 293.1031).

In silico computed chemical shifts

Theoretical chemical shifts were calculated for the four possible stereoisomers of compounds 1, and 1a based on a computational method described by Willoughby et al. [17], with some minor modifications. Briefly, the AMBER12 program suite [59] using the Generalized AMBER Force Field (GAFF) was used to explore the conformational space of each stereoisomer [60]. Minimized structures were heated to 500K over 5 ps (1 fs time step) and equilibrated at 500K for 1000, 100 ps cycles (0.5 fs time step). Following each cycle, structures were cooled to 0K over 5 ps (1 fs time step). Unique conformations were identified manually based on the C-2-Me, C-2, C-9, C-9-Me dihedral angles, and potential energy recorded following each cycle. Up to 20 conformations for each stereoisomer were removed for geometric optimization and frequency calculations using the B3LYP functional with a 6–31+G(d,p) basis set [61]. NMR shielding tensors were subsequently calculated from the geometry optimized structures with the B3LYP functional with the 6-311G (2d,p) basis set. Duplicate conformations with identical shielding tensors and zero point energies were removed and the chemical shift contribution for each conformation were averaged based on Boltzmann distributions. The scaling factor and intercept were determined using a linear fit of the theoretical tensors of all protons not predicted to be affected by the stereocenters of interest (positions 8, 5-Me, 6-Me and 7-OMe). The scaling factor and intercept were -0.99972 and 31.8085, respectively.

Antimicrobial assays

Initial antifungal activity of the Syn. ericacearum DAOMC 250336 culture filtrate extract (50 mg mL^-1) and flash chromatography fractions were screened using a modified Oxford diffusion assay against Microbotryum violaceum and Saccharomyces cerevisiae homogeneously spread on 2% MEA as previously described by McMullin et al. [33]. Purified natural products 1–3 were tested for in vitro antimicrobial activity against M. violaceum, S. cerevisiae, Bacillus subtilis (ATCC 23857) and Escherichia coli (ATCC 67878). M. violaceum was grown in 20 g L^-1 malt extract (Bacto), 2.5 g L^-1 peptone (Bacto) and 2.5 g L^-1 yeast extract (Sigma, St. Louis, MO, U.S.A.) whereas S. cerevisiae was inoculated and grown in 1 g L^-1 yeast extract supplemented with 10 g L^-1 glucose. Bacteria were inoculated and grown in 5 g L^-1 yeast extract, 10 g L^-1 peptone, and 10 g L^-1 NaCl. Nystatin was the positive control for antifungal assays and chloramphenicol was the antibacterial positive control. DMSO was the negative control for all in vitro assays. Isolated metabolites and positive controls were individually tested at 150, 75, 37.5, 18.8, 9.3, 4.7, 2.3 and 1.2 μg mL^-1 in sterile 96-well microplates, respectively (Falcon 353072 Microtest-9, Franklin Lakes, NJ, U.S.A.). A 10 μL aliquot of each individual metabolite solution dissolved in DMSO was added to 200 μL fungal or bacterial suspension. Assays were performed in triplicate and incubated at 25 °C with a rotary table shaker providing gentle agitation (500 rpm). Optical density (OD) measurements were made at 600 nm with a Molecular Devices Spectra Max 340PC reader (Sunnyvale, CA, U.S.A.). Antimicrobial OD data were subsequently analyzed by ANOVA followed by Tukey’s test (p < 0.05) for significant differences (Systat V13.1; Systat Software Inc., Chicago, IL, U.S.A.) compared to the negative control DMSO. Positive controls inhibited the growth of all test organisms at the second lowest concentrations tested. Previous studies had shown the MIC of nystatin in the S. cerevisiae culture used was 4 μM and for M. violaceum, 2 μM. In the antibiotic assays, for B. subtilis chloramphenicol had an MIC of 2.5 μM. All test organisms grew in the presence of the negative control DMSO.

Supporting information

S1 File.

Supporting Information: Figure A. ¹H (400 MHz, CD₃OD) NMR spectrum for 1. Figure B. ¹³C (100 MHz, CD₃OD) NMR spectrum for 1. Figure C. HMBC NMR spectrum for 1 in CD₃OD. Figure D. ¹H (400 MHz, CD₃OD) NMR spectrum for 2. Figure E. ¹³C (100 MHz, CD₃OD) NMR spectrum for 2. Figure F. HMBC NMR spectrum for 2 in CD₃OD. Figure G. ¹H (400 MHz, CD₃OD) NMR spectrum for 3. Figure H. ¹³C (100 MHz, CD₃OD) NMR spectrum for 3. Figure I. HMBC NMR spectrum for 3 in CD₃OD. Table A. Comparison of experimental and computed ¹H chemical shift data (ppm) for 1. Table B. Comparison of experimental and computed ¹³C chemical shift data (ppm) for 1. Table C. Comparison of experimental and computed ¹H chemical shift data (ppm) for 1a. Table D. Comparison of experimental and computed ¹³C chemical shift data (ppm) for 1a. Figure J. Methylation of synnemadoxin A (1) and synnemadiacid A (3) to their respective hexamethyl (1a) and heptamethyl (3a) derivatives using excess diazomethane. Table E. 1H (400 MHz) and 13C (100 MHz) NMR data for hexamethyl derivative of 1 (1a) in CD3OD. Table F. 1H (400 MHz) and 13C (100 MHz) NMR data for heptamethyl derivative of 3 (3a) in CD3OD. Text A. Methylation of synnemadoxin A (1) and synnemadiacid A (3) to their respective hexamethyl (1a) and heptamethyl (3a) derivatives using excess diazomethane.

https://doi.org/10.1371/journal.pone.0198321.s001

(DOCX)

Acknowledgments

We thank Dr. Allison Walker (Acadia University) for isolating DAOMC 250336 and Natasha C. Prince (Carleton University) for preliminary work on this strain. Dr. Stephen Clayden, Dr. David Malloch, Amanda Bremner (New Brunswick Museum) and Greg Adams (JD Irving, Ltd.) facilitated collection trips. Dr. Mark Sumarah is thanked for providing helpful comments on the manuscript.

References

1. Sumarah MW, Miller JD (2009) Anti-insect secondary metabolites from fungal endophytes of conifer trees. Natural product communications 4: 1497–1504. pmid:19967982
- View Article
- PubMed/NCBI
- Google Scholar
2. Tanney JB, McMullin DR, Miller JD (in press) Toxigenic foliar endophytes from the Acadian Forest. In: Frank A.C. AMP, editor. Endophytes of Forest Trees: Biology and Applications Volume II: Springer Forestry Series.
3. Tanney JB, Douglas B, Seifert KA (2016) Sexual and asexual states of some endophytic Phialocephala species of Picea. Mycologia: 15–136.
- View Article
- Google Scholar
4. Grünig CR, Queloz V, Duò A, Sieber TN (2009) Phylogeny of Phaeomollisia piceae gen. sp. nov.: a dark, septate, conifer-needle endophyte and its relationships to Phialocephala and Acephala. Mycological Research 113: 207–221. pmid:19015028
- View Article
- PubMed/NCBI
- Google Scholar
5. Menkis A, Allmer J, Vasiliauskas R, Lygis V, Stenlid J, Finlay R (2004) Ecology and molecular characterization of dark septate fungi from roots, living stems, coarse and fine woody debris. Mycological Research 108: 965–973. pmid:15449602
- View Article
- PubMed/NCBI
- Google Scholar
6. Richardson SN, Walker AK, Nsiama TK, McFarlane J, Sumarah MW, Miller JD (2014) Griseofulvin-producing Xylaria endophytes of Pinus strobus and Vaccinium angustifolium: evidence for a conifer-understory species endophyte ecology. Fungal Ecology 11: 107–113.
- View Article
- Google Scholar
7. Pirozynski K, Shoemaker R (1970) Seimatosporium leaf spot of Ledum and Rhododendron. Canadian Journal of Botany 48: 2199–2203.
- View Article
- Google Scholar
8. Shoemaker R (1964) Seimatosporium (= Cryptostictis) parasites of Rosa, Vitis, and Cornus. Canadian Journal of Botany 42: 411–421.
- View Article
- Google Scholar
9. U’Ren JM, Lutzoni F, Miadlikowska J, Laetsch AD, Arnold AE (2012) Host and geographic structure of endophytic and endolichenic fungi at a continental scale. American Journal of Botany 99: 898–914. pmid:22539507
- View Article
- PubMed/NCBI
- Google Scholar
10. Handa T, Fujita T, Harada Y (2004) Synnemapestaloides rhododendri, a new genus and new species of synnematous hyphomycete, causing synnemapestaloides twig blight disease of Rhododendron brachycarpum in Japan. Mycoscience 45: 137–142.
- View Article
- Google Scholar
11. Nag Raj TR (1993) Coelomycetous anamorphs with appendage-bearing conidia. Waterloo, Canada: Mycologue Publications. 1101 p.
12. Bonar L (1928) Studies on some California fungi. Mycologia 20: 292–300.
- View Article
- Google Scholar
13. Okane I, Nakagiri A, Ito T (1996) Discostroma tricellulare, a new endophytic ascomycete with a Seimatosporium anamorph isolated from Rhododendron. Canadian journal of botany 74: 1338–1344.
- View Article
- Google Scholar
14. Sutton BC (1980) The Coelomycetes: fungi imperfecti with pycnidia, acervuli and stromata. Kew, England: Commonwealth Mycological Institute. 696 p.
15. Shoemaker R, Müller E (1964) Generic correlations and concepts: Clathridium (= Griphosphaeria) and Seimatosporium (= Sporocadus). Canadian Journal of Botany 42: 403–410.
- View Article
- Google Scholar
16. Norphanphoun C, Maharachchikumbura SS, Daranagama A, Bulgakov TS, Bhat DJ, Bahkali AH, et al. (2015) Towards a backbone tree for Seimatosporium, with S. physocarpi sp. nov. Mycosphere 6: 385–400.
- View Article
- Google Scholar
17. Willoughby PH, Jansma MJ, Hoye TR (2014) A guide to small-molecule structure assignment through computation of (1H and 13C) NMR chemical shifts. Nature protocols 9: 643–660. pmid:24556787
- View Article
- PubMed/NCBI
- Google Scholar
18. Sumarah MW, Walker AK, Seifert KA, Todorov A, Miller JD (2015) Screening of fungal endophytes isolated from eastern white pine needles. The Formation, Structure and Activity of Phytochemicals: Springer. pp. 195–206.
19. Watanabe K, Sekiguchi M, Sato T, Hsiang T, Kaneko S, Tanaka K, et al. (2016) Phylogenetic analysis of the synnema-producing genus Synnemapestaloides. Journal of Fungi 2: 28.
- View Article
- Google Scholar
20. Sutton BC (1963) Coelomycetes. II. Neobarclaya, Mycohypallage, Bleptosporium and Cryptostictis. Mycological Papers 88: 1–50.
- View Article
- Google Scholar
21. Purmale L, Apine I, Nikolajewa V, Grantina L, Verkley G, Tomsone S (2012) Endophytic fungi in evergreen Rhododendron cultivated in vitro and in vivo. Environmental and Experimental Biology 10: 1–7.
- View Article
- Google Scholar
22. Maharachchikumbura SS, Hyde KD, Groenewald JZ, Xu J, Crous PW (2014) Pestalotiopsis revisited. Studies in Mycology 79: 121–186. pmid:25492988
- View Article
- PubMed/NCBI
- Google Scholar
23. Maharachchikumbura SS, Guo L-D, Chukeatirote E, Bahkali AH, Hyde KD (2011) Pestalotiopsis—morphology, phylogeny, biochemistry and diversity. Fungal Diversity 50: 167.
- View Article
- Google Scholar
24. Stierle AA, Stierle DB (2015) Bioactive secondary metabolites produced by the fungal endophytes of conifers. Natural product communications 10: 1671. pmid:26669101
- View Article
- PubMed/NCBI
- Google Scholar
25. Heinig U, Scholz S, Jennewein S (2013) Getting to the bottom of Taxol biosynthesis by fungi. Fungal Diversity 60: 161–170.
- View Article
- Google Scholar
26. Hiep NT, Choi Y-h, Kim N, Hong SS, Hong S-B, Hwang BY, et al. (2012) Polyhydroxylated macrolides from Seimatosporium discosioides and their effects on the activation of peroxisome proliferator-activated receptor gamma. Journal of Natural Products 75: 784–788. pmid:22424300
- View Article
- PubMed/NCBI
- Google Scholar
27. Reddy CR, Dilipkumar U, Reddy MD, Rao NN (2013) Total synthesis and revision of the absolute configuration of seimatopolide B. Organic & biomolecular chemistry 11: 3355–3364.
- View Article
- Google Scholar
28. Hussain H, Root N, Jabeen F, Al-Harrasi A, Al-Rawahi A, Ahmad M, et al. (2014) Seimatoric acid and colletonoic acid: Two new compounds from the endophytic fungi, Seimatosporium sp. and Colletotrichum sp. Chinese Chemical Letters 25: 1577–1579.
- View Article
- Google Scholar
29. Hussain H, Krohn K, Schulz B, Draeger S, Nazir M, Saleem M (2012) Two new antimicrobial metabolites from the endophytic fungus, Seimatosporium sp. Natural product communications 7: 293–294. pmid:22545398
- View Article
- PubMed/NCBI
- Google Scholar
30. Schulz B, Sucker J, Aust H, Krohn K, Ludewig K, Jones PG et al. (1995) Biologically active secondary metabolites of endophytic Pezicula species. Mycological Research 99: 1007–1015.
- View Article
- Google Scholar
31. Sumarah MW, Puniani E, Blackwell BA, Miller JD (2008) Characterization of polyketide metabolites from foliar endophytes of Picea glauca. Journal of Natural Products 71: 1393–1398. pmid:18636777
- View Article
- PubMed/NCBI
- Google Scholar
32. Claydon N, Grove JF, Pople M (1985) Elm bark beetle boring and feeding deterrents from Phomopsis oblonga. Phytochemistry 24: 937–943.
- View Article
- Google Scholar
33. McMullin DR, Green BD, Prince NC, Tanney JB, Miller JD (2017) Natural Products of Picea Endophytes from the Acadian Forest. Journal of Natural Products 80: 1475–1483. pmid:28398744
- View Article
- PubMed/NCBI
- Google Scholar
34. Tanney JB, McMullin DR, Green BD, Miller JD, Seifert KA (2016) Production of antifungal and antiinsectan metabolites by the Picea endophyte Diaporthe maritima sp. nov. Fungal Biology.
- View Article
- Google Scholar
35. Higginbotham S, Wong WR, Linington RG, Spadafora C, Iturrado L, Arnold AE (2014) Sloth hair as a novel source of fungi with potent anti-parasitic, anti-cancer and anti-bacterial bioactivity. PloS one 9: e84549. pmid:24454729
- View Article
- PubMed/NCBI
- Google Scholar
36. Adeboya MO, Edwards RL, Lassøe T, Maitland DJ, Shields L, Whalley AJS (1996) Metabolites of the higher fungi. Part 29. Maldoxin, maldoxone, dihydromaldoxin, isodihydromaldoxin and dechlorodihydromaldoxin. A spirocyclohexadienone, a depsidone and three diphenyl ethers: keys in the depsidone biosynthetic pathway from a member of the fungus genus Xylaria. Journal of the Chemical Society, Perkin Transactions 1: 1419–1425.
- View Article
- Google Scholar
37. Wang X, Zhang X, Liu L, Xiang M, Wang W, Sun X, et al. (2015) Genomic and transcriptomic analysis of the endophytic fungus Pestalotiopsis fici reveals its lifestyle and high potential for synthesis of natural products. BMC genomics 16: 28. pmid:25623211
- View Article
- PubMed/NCBI
- Google Scholar
38. Liu L, Li Y, Liu S, Zheng Z, Chen X, Zhang H, et al. (2009) Chloropestolide A, an antitumor metabolite with an unprecedented spiroketal skeleton from Pestalotiopsis fici. Organic letters 11: 2836–2839. pmid:19496604
- View Article
- PubMed/NCBI
- Google Scholar
39. Liu L, Li Y, Li L, Cao Y, Guo L, Liu G et al. (2013) Spiroketals of Pestalotiopsis fici provide evidence for a biosynthetic hypothesis involving diversified Diels–Alder reaction cascades. The Journal of organic chemistry 78: 2992–3000. pmid:23489375
- View Article
- PubMed/NCBI
- Google Scholar
40. Liu L, Niu S, Lu X, Chen X, Zhang H, Guo L, et al. (2010) Unique metabolites of Pestalotiopsis fici suggest a biosynthetic hypothesis involving a Diels–Alder reaction and then mechanistic diversification. Chemical Communications 46: 460–462. pmid:20066325
- View Article
- PubMed/NCBI
- Google Scholar
41. Liu L, Bruhn T, Guo L, Götz DC, Brun R, Stich A, et al. (2011) Chloropupukeanolides C–E: cytotoxic pupukeanane chlorides with a spiroketal skeleton from Pestalotiopsis fici. Chemistry-A European Journal 17: 2604–2613.
- View Article
- Google Scholar
42. Wu G, Zhou H, Zhang P, Wang X, Li W, Zhang W, et al. (2016) Polyketide production of pestaloficiols and macrodiolide ficiolides revealed by manipulations of epigenetic regulators in an endophytic fungus. Organic letters 18: 1832–1835. pmid:27015125
- View Article
- PubMed/NCBI
- Google Scholar
43. Suzuki T, Watanabe S, Kobayashi S, Tanino K (2017) Enantioselective Total Synthesis of (+)-Iso-A82775C, a Proposed Biosynthetic Precursor of Chloropupukeananin. Organic letters 19: 922–925.
- View Article
- Google Scholar
44. Suzuki T, Miyajima Y, Suzuki K, Iwakiri K, Koshimizu M, Hirai G, et al. (2013) Unexpected Diels–Alder/carbonyl-ene cascade toward the biomimetic synthesis of chloropupukeananin. Organic letters 15: 1748–1751. pmid:23530662
- View Article
- PubMed/NCBI
- Google Scholar
45. McMullin DR, Renaud JB, Barasubiye T, Sumarah MW, Miller JD (2017) Metabolites of Trichoderma species isolated from damp building materials. Canadian Journal of Microbiology.
- View Article
- Google Scholar
46. Washida K, Abe N, Sugiyama Y, Hirota A (2009) Novel secondary metabolites, spirosorbicillinols A, B, and C, from a fungus. Bioscience, biotechnology, and biochemistry 73: 1355–1361. pmid:19502726
- View Article
- PubMed/NCBI
- Google Scholar
47. Andrade R, Ayer WA, Trifonov LS (1996) The metabolites of Trichoderma longibrachiatum. Part II The structures of trichodermolide and sorbiquinol. Canadian journal of chemistry 74: 371–379.
- View Article
- Google Scholar
48. Yu M, Snider B (2011) Syntheses of chloroisosulochrin and isosulochrin and biomimetic elaboration to maldoxin, maldoxone, dihydromaldoxin, and dechlorodihydromaldoxin. Organic letters 13:4224–4227. pmid:21793536
- View Article
- PubMed/NCBI
- Google Scholar
49. Crous PW, Verkley GJ, Groenewald JZ, Samson R (2009) Fungal Biodiversity: Centraalbureau voor Schimmelcultures, Utrecht.
50. Pitt JI (1979) The genus Penicillium and its teleomorphic states Eupenicillium and Talaromyces. London and New York: Academic Press. 634 p.
51. Masclaux F, Guého E, De Hoog G, Christen R (1995) Phylogenetic relationships of human-pathogenic Cladosporium (Xylohypha) species inferred from partial LS rRNA sequences. Journal of Medical and Veterinary Mycology 33: 327–338. pmid:8544086
- View Article
- PubMed/NCBI
- Google Scholar
52. Gd Hoog, Ende A (1998) Molecular diagnostics of clinical strains of filamentous Basidiomycetes. Mycoses 41: 183–189. pmid:9715630
- View Article
- PubMed/NCBI
- Google Scholar
53. Vilgalys R, Hester M (1990) Rapid genetic identification and mapping of enzymatically amplified ribosomal DNA from several Cryptococcus species. Journal of Bacteriology 172: 4238–4246. pmid:2376561
- View Article
- PubMed/NCBI
- Google Scholar
54. Katoh K, Standley DM (2013) MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Molecular biology and evolution 30: 772–780. pmid:23329690
- View Article
- PubMed/NCBI
- Google Scholar
55. Nylander J (2004) MrModeltest version 2.3. http://www.abcse/~nylander/.
56. Ronquist F, Teslenko M, Van Der Mark P, Ayres DL, Darling A, et al. (2012) MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Systematic biology 61: 539–542. pmid:22357727
- View Article
- PubMed/NCBI
- Google Scholar
57. Stamatakis A (2014) RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics 30: 1312–1313. pmid:24451623
- View Article
- PubMed/NCBI
- Google Scholar
58. Swofford DL (2003) PAUP*: phylogenetic analysis using parsimony, version 4.0 b10.
59. Case D, Babin V, Berryman J, Betz R, Cai Q, Cerutti DS, et al. (2006) AMBER 14, University of California, San Francisco, 2014 Search PubMed;(b) V. Hornak, R. Abel, A. Okur, B. Strockbine, A. Roitberg and C. Simmerling. Proteins 65: 712–725.
- View Article
- Google Scholar
60. Wang J, Wolf RM, Caldwell JW, Kollman PA, Case DA (2004) Development and testing of a general amber force field. Journal of computational chemistry 25: 1157–1174. pmid:15116359
- View Article
- PubMed/NCBI
- Google Scholar
61. Becke AD (1992) Density-functional thermochemistry. I. The effect of the exchange-only gradient correction. The Journal of chemical physics 96: 2155–2160.
- View Article
- Google Scholar

[ref1] 1. Sumarah MW, Miller JD (2009) Anti-insect secondary metabolites from fungal endophytes of conifer trees. Natural product communications 4: 1497–1504. pmid:19967982
View Article
PubMed/NCBI
Google Scholar

[2] View Article

[3] PubMed/NCBI

[4] Google Scholar

[ref2] 2. Tanney JB, McMullin DR, Miller JD (in press) Toxigenic foliar endophytes from the Acadian Forest. In: Frank A.C. AMP, editor. Endophytes of Forest Trees: Biology and Applications Volume II: Springer Forestry Series.

[ref3] 3. Tanney JB, Douglas B, Seifert KA (2016) Sexual and asexual states of some endophytic Phialocephala species of Picea. Mycologia: 15–136.
View Article
Google Scholar

[7] View Article

[8] Google Scholar

[ref4] 4. Grünig CR, Queloz V, Duò A, Sieber TN (2009) Phylogeny of Phaeomollisia piceae gen. sp. nov.: a dark, septate, conifer-needle endophyte and its relationships to Phialocephala and Acephala. Mycological Research 113: 207–221. pmid:19015028
View Article
PubMed/NCBI
Google Scholar

[10] View Article

[11] PubMed/NCBI

[12] Google Scholar

[ref5] 5. Menkis A, Allmer J, Vasiliauskas R, Lygis V, Stenlid J, Finlay R (2004) Ecology and molecular characterization of dark septate fungi from roots, living stems, coarse and fine woody debris. Mycological Research 108: 965–973. pmid:15449602
View Article
PubMed/NCBI
Google Scholar

[14] View Article

[15] PubMed/NCBI

[16] Google Scholar

[ref6] 6. Richardson SN, Walker AK, Nsiama TK, McFarlane J, Sumarah MW, Miller JD (2014) Griseofulvin-producing Xylaria endophytes of Pinus strobus and Vaccinium angustifolium: evidence for a conifer-understory species endophyte ecology. Fungal Ecology 11: 107–113.
View Article
Google Scholar

[18] View Article

[19] Google Scholar

[ref7] 7. Pirozynski K, Shoemaker R (1970) Seimatosporium leaf spot of Ledum and Rhododendron. Canadian Journal of Botany 48: 2199–2203.
View Article
Google Scholar

[21] View Article

[22] Google Scholar

[ref8] 8. Shoemaker R (1964) Seimatosporium (= Cryptostictis) parasites of Rosa, Vitis, and Cornus. Canadian Journal of Botany 42: 411–421.
View Article
Google Scholar

[24] View Article

[25] Google Scholar

[ref9] 9. U’Ren JM, Lutzoni F, Miadlikowska J, Laetsch AD, Arnold AE (2012) Host and geographic structure of endophytic and endolichenic fungi at a continental scale. American Journal of Botany 99: 898–914. pmid:22539507
View Article
PubMed/NCBI
Google Scholar

[27] View Article

[28] PubMed/NCBI

[29] Google Scholar

[ref10] 10. Handa T, Fujita T, Harada Y (2004) Synnemapestaloides rhododendri, a new genus and new species of synnematous hyphomycete, causing synnemapestaloides twig blight disease of Rhododendron brachycarpum in Japan. Mycoscience 45: 137–142.
View Article
Google Scholar

[31] View Article

[32] Google Scholar

[ref11] 11. Nag Raj TR (1993) Coelomycetous anamorphs with appendage-bearing conidia. Waterloo, Canada: Mycologue Publications. 1101 p.

[ref12] 12. Bonar L (1928) Studies on some California fungi. Mycologia 20: 292–300.
View Article
Google Scholar

[35] View Article

[36] Google Scholar

[ref13] 13. Okane I, Nakagiri A, Ito T (1996) Discostroma tricellulare, a new endophytic ascomycete with a Seimatosporium anamorph isolated from Rhododendron. Canadian journal of botany 74: 1338–1344.
View Article
Google Scholar

[38] View Article

[39] Google Scholar

[ref14] 14. Sutton BC (1980) The Coelomycetes: fungi imperfecti with pycnidia, acervuli and stromata. Kew, England: Commonwealth Mycological Institute. 696 p.

[ref15] 15. Shoemaker R, Müller E (1964) Generic correlations and concepts: Clathridium (= Griphosphaeria) and Seimatosporium (= Sporocadus). Canadian Journal of Botany 42: 403–410.
View Article
Google Scholar

[42] View Article

[43] Google Scholar

[ref16] 16. Norphanphoun C, Maharachchikumbura SS, Daranagama A, Bulgakov TS, Bhat DJ, Bahkali AH, et al. (2015) Towards a backbone tree for Seimatosporium, with S. physocarpi sp. nov. Mycosphere 6: 385–400.
View Article
Google Scholar

[45] View Article

[46] Google Scholar

[ref17] 17. Willoughby PH, Jansma MJ, Hoye TR (2014) A guide to small-molecule structure assignment through computation of (1H and 13C) NMR chemical shifts. Nature protocols 9: 643–660. pmid:24556787
View Article
PubMed/NCBI
Google Scholar

[48] View Article

[49] PubMed/NCBI

[50] Google Scholar

[ref18] 18. Sumarah MW, Walker AK, Seifert KA, Todorov A, Miller JD (2015) Screening of fungal endophytes isolated from eastern white pine needles. The Formation, Structure and Activity of Phytochemicals: Springer. pp. 195–206.

[ref19] 19. Watanabe K, Sekiguchi M, Sato T, Hsiang T, Kaneko S, Tanaka K, et al. (2016) Phylogenetic analysis of the synnema-producing genus Synnemapestaloides. Journal of Fungi 2: 28.
View Article
Google Scholar

[53] View Article

[54] Google Scholar

[ref20] 20. Sutton BC (1963) Coelomycetes. II. Neobarclaya, Mycohypallage, Bleptosporium and Cryptostictis. Mycological Papers 88: 1–50.
View Article
Google Scholar

[56] View Article

[57] Google Scholar

[ref21] 21. Purmale L, Apine I, Nikolajewa V, Grantina L, Verkley G, Tomsone S (2012) Endophytic fungi in evergreen Rhododendron cultivated in vitro and in vivo. Environmental and Experimental Biology 10: 1–7.
View Article
Google Scholar

[59] View Article

[60] Google Scholar

[ref22] 22. Maharachchikumbura SS, Hyde KD, Groenewald JZ, Xu J, Crous PW (2014) Pestalotiopsis revisited. Studies in Mycology 79: 121–186. pmid:25492988
View Article
PubMed/NCBI
Google Scholar

[62] View Article

[63] PubMed/NCBI

[64] Google Scholar

[ref23] 23. Maharachchikumbura SS, Guo L-D, Chukeatirote E, Bahkali AH, Hyde KD (2011) Pestalotiopsis—morphology, phylogeny, biochemistry and diversity. Fungal Diversity 50: 167.
View Article
Google Scholar

[66] View Article

[67] Google Scholar

[ref24] 24. Stierle AA, Stierle DB (2015) Bioactive secondary metabolites produced by the fungal endophytes of conifers. Natural product communications 10: 1671. pmid:26669101
View Article
PubMed/NCBI
Google Scholar

[69] View Article

[70] PubMed/NCBI

[71] Google Scholar

[ref25] 25. Heinig U, Scholz S, Jennewein S (2013) Getting to the bottom of Taxol biosynthesis by fungi. Fungal Diversity 60: 161–170.
View Article
Google Scholar

[73] View Article

[74] Google Scholar

[ref26] 26. Hiep NT, Choi Y-h, Kim N, Hong SS, Hong S-B, Hwang BY, et al. (2012) Polyhydroxylated macrolides from Seimatosporium discosioides and their effects on the activation of peroxisome proliferator-activated receptor gamma. Journal of Natural Products 75: 784–788. pmid:22424300
View Article
PubMed/NCBI
Google Scholar

[76] View Article

[77] PubMed/NCBI

[78] Google Scholar

[ref27] 27. Reddy CR, Dilipkumar U, Reddy MD, Rao NN (2013) Total synthesis and revision of the absolute configuration of seimatopolide B. Organic & biomolecular chemistry 11: 3355–3364.
View Article
Google Scholar

[80] View Article

[81] Google Scholar

[ref28] 28. Hussain H, Root N, Jabeen F, Al-Harrasi A, Al-Rawahi A, Ahmad M, et al. (2014) Seimatoric acid and colletonoic acid: Two new compounds from the endophytic fungi, Seimatosporium sp. and Colletotrichum sp. Chinese Chemical Letters 25: 1577–1579.
View Article
Google Scholar

[83] View Article

[84] Google Scholar

[ref29] 29. Hussain H, Krohn K, Schulz B, Draeger S, Nazir M, Saleem M (2012) Two new antimicrobial metabolites from the endophytic fungus, Seimatosporium sp. Natural product communications 7: 293–294. pmid:22545398
View Article
PubMed/NCBI
Google Scholar

[86] View Article

[87] PubMed/NCBI

[88] Google Scholar

[ref30] 30. Schulz B, Sucker J, Aust H, Krohn K, Ludewig K, Jones PG et al. (1995) Biologically active secondary metabolites of endophytic Pezicula species. Mycological Research 99: 1007–1015.
View Article
Google Scholar

[90] View Article

[91] Google Scholar

[ref31] 31. Sumarah MW, Puniani E, Blackwell BA, Miller JD (2008) Characterization of polyketide metabolites from foliar endophytes of Picea glauca. Journal of Natural Products 71: 1393–1398. pmid:18636777
View Article
PubMed/NCBI
Google Scholar

[93] View Article

[94] PubMed/NCBI

[95] Google Scholar

[ref32] 32. Claydon N, Grove JF, Pople M (1985) Elm bark beetle boring and feeding deterrents from Phomopsis oblonga. Phytochemistry 24: 937–943.
View Article
Google Scholar

[97] View Article

[98] Google Scholar

[ref33] 33. McMullin DR, Green BD, Prince NC, Tanney JB, Miller JD (2017) Natural Products of Picea Endophytes from the Acadian Forest. Journal of Natural Products 80: 1475–1483. pmid:28398744
View Article
PubMed/NCBI
Google Scholar

[100] View Article

[101] PubMed/NCBI

[102] Google Scholar

[ref34] 34. Tanney JB, McMullin DR, Green BD, Miller JD, Seifert KA (2016) Production of antifungal and antiinsectan metabolites by the Picea endophyte Diaporthe maritima sp. nov. Fungal Biology.
View Article
Google Scholar

[104] View Article

[105] Google Scholar

[ref35] 35. Higginbotham S, Wong WR, Linington RG, Spadafora C, Iturrado L, Arnold AE (2014) Sloth hair as a novel source of fungi with potent anti-parasitic, anti-cancer and anti-bacterial bioactivity. PloS one 9: e84549. pmid:24454729
View Article
PubMed/NCBI
Google Scholar

[107] View Article

[108] PubMed/NCBI

[109] Google Scholar

[ref36] 36. Adeboya MO, Edwards RL, Lassøe T, Maitland DJ, Shields L, Whalley AJS (1996) Metabolites of the higher fungi. Part 29. Maldoxin, maldoxone, dihydromaldoxin, isodihydromaldoxin and dechlorodihydromaldoxin. A spirocyclohexadienone, a depsidone and three diphenyl ethers: keys in the depsidone biosynthetic pathway from a member of the fungus genus Xylaria. Journal of the Chemical Society, Perkin Transactions 1: 1419–1425.
View Article
Google Scholar

[111] View Article

[112] Google Scholar

[ref37] 37. Wang X, Zhang X, Liu L, Xiang M, Wang W, Sun X, et al. (2015) Genomic and transcriptomic analysis of the endophytic fungus Pestalotiopsis fici reveals its lifestyle and high potential for synthesis of natural products. BMC genomics 16: 28. pmid:25623211
View Article
PubMed/NCBI
Google Scholar

[114] View Article

[115] PubMed/NCBI

[116] Google Scholar

[ref38] 38. Liu L, Li Y, Liu S, Zheng Z, Chen X, Zhang H, et al. (2009) Chloropestolide A, an antitumor metabolite with an unprecedented spiroketal skeleton from Pestalotiopsis fici. Organic letters 11: 2836–2839. pmid:19496604
View Article
PubMed/NCBI
Google Scholar

[118] View Article

[119] PubMed/NCBI

[120] Google Scholar

[ref39] 39. Liu L, Li Y, Li L, Cao Y, Guo L, Liu G et al. (2013) Spiroketals of Pestalotiopsis fici provide evidence for a biosynthetic hypothesis involving diversified Diels–Alder reaction cascades. The Journal of organic chemistry 78: 2992–3000. pmid:23489375
View Article
PubMed/NCBI
Google Scholar

[122] View Article

[123] PubMed/NCBI

[124] Google Scholar

[ref40] 40. Liu L, Niu S, Lu X, Chen X, Zhang H, Guo L, et al. (2010) Unique metabolites of Pestalotiopsis fici suggest a biosynthetic hypothesis involving a Diels–Alder reaction and then mechanistic diversification. Chemical Communications 46: 460–462. pmid:20066325
View Article
PubMed/NCBI
Google Scholar

[126] View Article

[127] PubMed/NCBI

[128] Google Scholar

[ref41] 41. Liu L, Bruhn T, Guo L, Götz DC, Brun R, Stich A, et al. (2011) Chloropupukeanolides C–E: cytotoxic pupukeanane chlorides with a spiroketal skeleton from Pestalotiopsis fici. Chemistry-A European Journal 17: 2604–2613.
View Article
Google Scholar

[130] View Article

[131] Google Scholar

[ref42] 42. Wu G, Zhou H, Zhang P, Wang X, Li W, Zhang W, et al. (2016) Polyketide production of pestaloficiols and macrodiolide ficiolides revealed by manipulations of epigenetic regulators in an endophytic fungus. Organic letters 18: 1832–1835. pmid:27015125
View Article
PubMed/NCBI
Google Scholar

[133] View Article

[134] PubMed/NCBI

[135] Google Scholar

[ref43] 43. Suzuki T, Watanabe S, Kobayashi S, Tanino K (2017) Enantioselective Total Synthesis of (+)-Iso-A82775C, a Proposed Biosynthetic Precursor of Chloropupukeananin. Organic letters 19: 922–925.
View Article
Google Scholar

[137] View Article

[138] Google Scholar

[ref44] 44. Suzuki T, Miyajima Y, Suzuki K, Iwakiri K, Koshimizu M, Hirai G, et al. (2013) Unexpected Diels–Alder/carbonyl-ene cascade toward the biomimetic synthesis of chloropupukeananin. Organic letters 15: 1748–1751. pmid:23530662
View Article
PubMed/NCBI
Google Scholar

[140] View Article

[141] PubMed/NCBI

[142] Google Scholar

[ref45] 45. McMullin DR, Renaud JB, Barasubiye T, Sumarah MW, Miller JD (2017) Metabolites of Trichoderma species isolated from damp building materials. Canadian Journal of Microbiology.
View Article
Google Scholar

[144] View Article

[145] Google Scholar

[ref46] 46. Washida K, Abe N, Sugiyama Y, Hirota A (2009) Novel secondary metabolites, spirosorbicillinols A, B, and C, from a fungus. Bioscience, biotechnology, and biochemistry 73: 1355–1361. pmid:19502726
View Article
PubMed/NCBI
Google Scholar

[147] View Article

[148] PubMed/NCBI

[149] Google Scholar

[ref47] 47. Andrade R, Ayer WA, Trifonov LS (1996) The metabolites of Trichoderma longibrachiatum. Part II The structures of trichodermolide and sorbiquinol. Canadian journal of chemistry 74: 371–379.
View Article
Google Scholar

[151] View Article

[152] Google Scholar

[ref48] 48. Yu M, Snider B (2011) Syntheses of chloroisosulochrin and isosulochrin and biomimetic elaboration to maldoxin, maldoxone, dihydromaldoxin, and dechlorodihydromaldoxin. Organic letters 13:4224–4227. pmid:21793536
View Article
PubMed/NCBI
Google Scholar

[154] View Article

[155] PubMed/NCBI

[156] Google Scholar

[ref49] 49. Crous PW, Verkley GJ, Groenewald JZ, Samson R (2009) Fungal Biodiversity: Centraalbureau voor Schimmelcultures, Utrecht.

[ref50] 50. Pitt JI (1979) The genus Penicillium and its teleomorphic states Eupenicillium and Talaromyces. London and New York: Academic Press. 634 p.

[ref51] 51. Masclaux F, Guého E, De Hoog G, Christen R (1995) Phylogenetic relationships of human-pathogenic Cladosporium (Xylohypha) species inferred from partial LS rRNA sequences. Journal of Medical and Veterinary Mycology 33: 327–338. pmid:8544086
View Article
PubMed/NCBI
Google Scholar

[160] View Article

[161] PubMed/NCBI

[162] Google Scholar

[ref52] 52. Gd Hoog, Ende A (1998) Molecular diagnostics of clinical strains of filamentous Basidiomycetes. Mycoses 41: 183–189. pmid:9715630
View Article
PubMed/NCBI
Google Scholar

[164] View Article

[165] PubMed/NCBI

[166] Google Scholar

[ref53] 53. Vilgalys R, Hester M (1990) Rapid genetic identification and mapping of enzymatically amplified ribosomal DNA from several Cryptococcus species. Journal of Bacteriology 172: 4238–4246. pmid:2376561
View Article
PubMed/NCBI
Google Scholar

[168] View Article

[169] PubMed/NCBI

[170] Google Scholar

[ref54] 54. Katoh K, Standley DM (2013) MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Molecular biology and evolution 30: 772–780. pmid:23329690
View Article
PubMed/NCBI
Google Scholar

[172] View Article

[173] PubMed/NCBI

[174] Google Scholar

[ref55] 55. Nylander J (2004) MrModeltest version 2.3. http://www.abcse/~nylander/.

[ref56] 56. Ronquist F, Teslenko M, Van Der Mark P, Ayres DL, Darling A, et al. (2012) MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Systematic biology 61: 539–542. pmid:22357727
View Article
PubMed/NCBI
Google Scholar

[177] View Article

[178] PubMed/NCBI

[179] Google Scholar

[ref57] 57. Stamatakis A (2014) RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics 30: 1312–1313. pmid:24451623
View Article
PubMed/NCBI
Google Scholar

[181] View Article

[182] PubMed/NCBI

[183] Google Scholar

[ref58] 58. Swofford DL (2003) PAUP*: phylogenetic analysis using parsimony, version 4.0 b10.

[ref59] 59. Case D, Babin V, Berryman J, Betz R, Cai Q, Cerutti DS, et al. (2006) AMBER 14, University of California, San Francisco, 2014 Search PubMed;(b) V. Hornak, R. Abel, A. Okur, B. Strockbine, A. Roitberg and C. Simmerling. Proteins 65: 712–725.
View Article
Google Scholar

[186] View Article

[187] Google Scholar

[ref60] 60. Wang J, Wolf RM, Caldwell JW, Kollman PA, Case DA (2004) Development and testing of a general amber force field. Journal of computational chemistry 25: 1157–1174. pmid:15116359
View Article
PubMed/NCBI
Google Scholar

[189] View Article

[190] PubMed/NCBI

[191] Google Scholar

[ref61] 61. Becke AD (1992) Density-functional thermochemistry. I. The effect of the exchange-only gradient correction. The Journal of chemical physics 96: 2155–2160.
View Article
Google Scholar

[193] View Article

[194] Google Scholar