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DOI: 10.3852/mycologia.97.2.444
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Mycologia, 97(2), 2005, pp. 444-453.
© 2005 by The Mycological Society of America

Penicillium dravuni, a new marine-derived species from an alga in Fiji


Jeffrey E. Janso 1
Valerie S. Bernan
Michael Greenstein

     Wyeth Research, Natural Products Microbiology, 401 North Middletown Road, Pearl River, New York 10965

Tim S. Bugni
Chris M. Ireland

     Department of Medicinal Chemistry, University of Utah, Salt Lake City, Utah 84112

   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 LITERATURE CITED
 

Penicillium dravuni is a new monoverticillate, sclerotium-forming species that was isolated from the alga Dictyosphaeria versluyii collected in Dravuni, Fiji. This species morphologically is similar to P. turbatum in the P. turbatum subseries of the P. thomii series of the Monoverticillata. The nuclear ribosomal internal transcribed spacer region exhibited 97% sequence similarity to known Penicillium spp. in the GenBank database. Phylogenetic analyses revealed that P. dravuni is related most closely to Eupenicillium brefeldianum, E. levitum, E. reticulosporum, E. javanicum, E. ehrlichii and P. simplicissimum. However this new species shares only a distant ancestor with this clade because it branches by itself early in the lineage. P. dravuni also is known to produce the secondary metabolites dictyosphaeric acids A and B and carviolin.

Key words: Dictyosphaeria, dictyosphaeric acids, carviolin, ITS, marine-derived, Trichocomaceae


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 LITERATURE CITED
 
The exploration of marine environments for unique microorganisms, especially actinomycetes and fungi, has yielded numerous isolates from algae, sediments, submerged wood and invertebrates. Within the past decade these organisms have been incorporated into culture collections at academic institutions and pharmaceutical companies, where the fermentation products are investigated intensively for bioactive molecules. Often the compounds produced by marine-derived microorganisms are the same as or similar to compounds produced by their terrestrial counterparts (Edrada et al 2002Go, Garo et al 2003Go, Malmstrøm et al 2000Go). However examples of metabolites with unprecedented structures are increasing, such as paecilospirone (Namikoshi et al 2000) and brocaenols A–C (Bugni et al 2003Go), isolated from marine-derived fungi.

Several undescribed, marine-derived Penicillium spp. recently have been isolated from a variety of substrates such as mollusks, sponges, algae and sand. These Penicillia are important producers of new metabolites such as the sculezonones A and B (Komatsu 2000Go), coruscol A (Kagata et al 2000Go), penicillazine (Lin et al 2000Go), and the xestodecalactones A–C (Edrada et al 2002Go). One Penicillium sp., which was isolated from the marine alga Enteromorpha intestinalis, has been shown to produce three different classes of secondary metabolites, the communesins (Numata et al 1993Go), the penostatins (Iwamoto et al 1999Go) and the penochalasins (Iwamoto et al 2001Go).

While screening for compounds that inhibit the growth of antibiotic-resistant bacteria, we found that the fermentation extracts of F01V25, a Penicillium sp. isolated from the alga D. versluyii, had antimicrobial activity against methicillin-resistant Staphylococcus aureus (MRSA). In addition to the known pigment carviolin F01V25 was found to produce two new polyketides, the dictyosphaeric acids A and B (Bugni et al 2004Go). Investigation of the morphology and phylogenetics of the nuclear ribosomal internal transcribed spacer region (ITS) of F01V25 failed to associate it with any described species. The taxonomy of the new species, Penicillium dravuni, is described.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 LITERATURE CITED
 
Isolation.— – A sample of D. versluyii was collected by scuba diving near Dravuni Island, Fiji, in Jan 2001. The inner tissue was separated from the outer tissue with a sterile scalpel and finely ground in sterile seawater with mortar and pestle. An aliquot of 100 µL of the slurry was plated onto several selective agar-based media, some containing antibiotics to inhibit the growth of bacteria. Petri dishes were incubated several weeks at room temperature and examined periodically for new isolates, which were colony purified by plating to fresh agar media.

Morphological analysis.— – Phenotypic identification of F01V25 was performed according to the protocols of Pitt (1988)Go and Ramirez (1982)Go. Culture colors were matched as closely as possible to the Inter-Society Color Council (ISCC) and the National Bureau of Standards (NBS) color name charts (Kelly 1958Go). To determine growth in the presence of salt, F01V25 was plated to malt-extract agar (MEA) prepared with artificial seawater (MEA-ASW), which contains 3% sodium chloride (NaCl) w/v and 1% other salts (potassium chloride [KCl], magnesium chloride [MgCl2], calcium chloride [CaCl2]) w/v. To determine NaCl tolerance, F01V25 was grown on MEA supplemented with 5, 10, 15 or 20% NaCl. To distinguish nonspecific salt from sodium tolerance, F01V25 was cultured on MEA prepared with 5% KCl. All were incubated at 25 C. The maximum and minimum temperatures that would support growth of F01V25 were determined respectively by incubation at 37–40 C and 5, 11 and 15 C.

Macroscopic observations were conducted with a Nikon SMZ1500 stereoscope. A Nikon E600 compound microscope was used with phase-contrast or differential interference contrast (DIC) for microscopic examinations. A Nikon Digital Camera DXM1200 was connected to both instruments to take digital images. The images were viewed and adjusted with Nikon ACT-1 version 2.1 and further modified in Microsoft Photo Editor 3.01.

For scanning electron microscopy (SEM) of the conidia and conidiophores, agar plugs (3 mm2) were cut from an 8 d old culture of F01V25 grown on Difco potato-dextrose agar (PDA). To view the sclerotia by SEM, agar plugs (3 mm2) were cut from a 2 mo old culture of F01V25 grown on malt-extract agar (MEA). Each agar plug was fixed with osmium tetroxide (Sigma) and dehydrated in a successive series of ethanol washes. Critical point drying with carbon dioxide was performed on an Autosamdri-814B sample drier (Tousimis Research Corp.). Samples were stored under desiccation until they were sputter-coated with palladium 50 s (Denton Vacuum Desk II model) and viewed by SEM with a JOEL 5800LV.

DNA extraction.— – Genomic DNA was isolated from cultures grown 7 d on PDA. About 50–100 mg of mycelia were scraped from the agar surface and placed inside a 1.5 mL microcentrifuge tube containing 200 µL of TE buffer at pH 8.0. Mycelia were ground with motorized pellet pestle (Kontes), and an additional 800 µL of TE was added to wash the slurry. Mycelia were pelleted in microcentrifuge 14 000 rpm, and the supernatant was decanted. For cell lysis, 500 µL of lysis buffer (2% Triton X-100, 1% SDS, 100 mM NaCl, 10 mM Tris pH 8.0, 1 mM EDTA) was added to the cell pellet and vortexed 1 min. Five hundred µL of phenol : chloroform : isoamyl alcohol (25:24:1) (Sigma) was added, and the mixture was shaken by hand to precipitate proteins. The aqueous and organic phases were separated by centrifugation at 14 000 rpm 5 min. The top aqueous layer was transferred to a clean 1.5 mL microcentrifuge tube, and the DNA was precipitated by the addition of one-tenth volume 3M sodium acetate followed by 1 volume of isopropyl alcohol (JT Baker). DNA was pelleted by centrifugation at 14 000 rpm 2 min. The isopropyl alcohol was decanted, and the pellet was washed with 1 mL 70% ethanol. The ethanol was decanted, and the pellet was air dried 1 h in a laminar flow hood. DNA was resuspended in 50 µL of 10 mM Tris containing 200 µg/mL RNase (Sigma).

PCR amplification and sequencing.— – The nuclear ribosomal ITS1–5.8S-ITS2 region was amplified with primers ITS1 and ITS4 (White et al 1990Go). The reactions were performed in 100 µL volumes containing 1 µL of genomic DNA, 1 µL 50 µM ITS1, 1 µL 50 µM ITS4, 47 µL sterile deionized water and 50 µL JumpStartTM ReadyMixTM REDTaqTM DNA polymerase (Sigma) as follows: initial denaturation at 95 C for 2 min, followed by 35 cycles of 95 C for 1 min, 53 C for 45 s, 72 C for 1 min 30 s. A final extension at 72 C was done for 5 min. To confirm the amplification of only the ITS, 4 µL of the PCR product was run on an agarose gel. The PCR product was purified with the Eclipse PCR Clean-up DNA purification kit (Tetra Link International), and sequences were determined with the PCR primers ITS1 and ITS4. Direct sequencing was performed with an ABI 3700 sequencer with the ABI Prism DNA sequencing kit and Big Dye terminators version 3.0 (Applied Biosystems). Sequence data was edited with SequencherTM version 4.1.4. ITS was deposited in GenBank with accession number AY494856 [GenBank] .

Phylogenetic analysis.— – To determine the most closely related Penicillium spp., the ITS of F01V25 was compared to other sequences in the GenBank database by BLASTN 2.2.2 analysis. The ITS sequences of morphologically and phylogenetically related Penicillium spp. were obtained from GenBank (TABLE IGo) and aligned in Clustal X 1.81 (Thompson et al 1997Go) with pairwise and multiple alignment parameters set at 15.0 for gap opening and 0.5 for gap extension. Phylogenetic analyses were based on maximum parsimony and distance using PAUP*4.0b10 (Swofford 2002Go). For parsimony, a heuristic search was conducted with simple addition of sequences, tree bisection-reconnection (TBR) branch swapping, collapse option in effect, MulTrees option in effect and MaxTrees set to 100 with automatic increases of 100. Gaps were treated as missing data, branches collapsed (creating polytomies) if the maximum branch length was zero, and groups compatible with 50% majority-rule consensus were included. For distance analysis, matrices were measured using the method of Jukes and Cantor (1969)Go and the tree was inferred by neighbor joining. Bootstrap analyses were performed on both parsimony and neighbor joining trees with 1000 bootstrap replicates. Both trees were rooted with Paecilomyces variotti. Trees were viewed with TreeView 1.6.6 (Page 2001) and edited in Microsoft® PowerPoint® 2000.


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  		TABLE I. Penicillium spp. used in the phylogenetic analysis
 

   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 LITERATURE CITED
 
Penicillium dravuni JE Janso, sp. nov.

Coloniae crescunt 7 dies in CYA ad 25 C, 25–35 mm diam, radialiter sulcatae, textura superficiei velutinosa, flavo-griseae ad pallide griseo-flavae, albescentes ad marginem. Conidia per mediam partem abundanter formantur, sed sparse ad marginem. Sudores aureo-flavi ad fusce crocei vel aurantiaco-rubri per medias colonias rari sunt. Pigmentum solubile et pallide flavum ad croceum sub coloniis ipsis adest. Facies reversa pallide brunnea ad pallide flavo-brunnea in centro, medio-aurantiaca vel medio-flava in marginibus. Conidiophorae fere oriunt a hyphis basalibus vel a hyphis aeriis per medias colonias. Stipites breves, 35 ad 100 µm x 2 ad 3 µm, tenuiter tunicati, exigue vesiculati in CYA, manifeste vesiculati in MEA, rigide monoverticillati, rare ramosi. Phialides ampulliformae, 7 ad 11 µm, frequenter inveniuntur in verticillis 7 ad 13 continentibus. Conidia spherica ad subglobosa, leviter tunicata ad tenuiter asperata, 2.5 ad 3 µm x 2 µm, feruntur in columnis longis laxisque. Sclerotia solum formantur in MEA, 50 ad 100 µm, plerumque globosa, sine colore ad pallide lutea et aliquantum mollia.

Colonies grown 7 d on Czapek’s yeast autolysate agar (CYA) (FIG. 1Go) at 25 C are 25–35 mm diam, radially sulcate, with surface texture strictly velutinous, yellow-gray (ISCC-NBS 93) or gray-yellow (ISCC-NBS 90) to pale gray-yellow (ISCC-NBS 104), becoming white toward the margin. Conidia are produced centrally in abundance but sparsely peripherally. Golden yellow to dark amber or orange-red exudates are sparse in the centers of colonies. A light yellow to amber soluble pigment is present directly under the colonies. No sclerotia or ascomata are produced. The reverse color ranges from a light brown (ISCC-NBS 55) to light yellow-brown (ISCC-NBS 74) centrally to medium orange (ISCC-NBS 71) or medium yellow (ISCC-NBS 87) at the margins.


Figure 1
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  		FIGS. 1–7. 1. F01V25 grown on CYA for 7 d at 25 C. 2. F01V25 grown on MEA for 7 d at 25 C. 3. Conidiophore with vesiculate stipe, phase contrast. 4. Phialides with chains of conidia, SEM. 5. Finely roughened conidia in chains, SEM. 6. Sclerotia of various sizes but all with pseudoparenchymatous cells, phase-contrast. 7. Sclerotia surrounded by nets of mycelia, SEM. Scale bars: 3 = 10 µm; 4 = 5 µm; 5 = 2 µm; 6 = 20 µm; 7 = 10 µm.

 
Colonies grown 7 d on MEA (FIG. 2Go) at 25 C become 25–35 mm diam, are plane, and sporulate abundantly on radial growth. Surface growth is generally thin and floccose with denser growth centrally and submerged mycelia at the margins. Color ranges from grayish green-yellow (ISCC-NBS 105) to yellow-white (ISCC-NBS 92) or pale yellow-gray. The reverse is light yellow-brown (ISCC-NBS 74) to deep orange-yellow (ISCC-NBS 69) at the centers of colonies to light orange-yellow (ISCC-NBS 68) toward the edges. Exudate is absent, but an orange-yellow soluble pigment is present. Sclerotia are produced and can be numerous, white and surrounded by a thin netting of hyphae.

Colonies grown 7 d on G25N at 25 C are somewhat restricted; diam 15–25 mm, surface centrally floccose with sparse conidiation becoming more velutinous or felt-like toward the margins. Surface color is light gray to white, and the reverse is light amber at the centers to yellow peripherally. A yellow soluble pigment is present. Sclerotia and ascomata are absent.

Incubation at 5 C on MEA and CYA did not result in mycelial growth or germination of the conidia. However growth occurred at 37 C as colonies attained diam of 5 mm in 7 d and 8 mm in 14 d. No growth was observed at 38 C.

Conidiophores appear similar on CYA and MEA but show slightly more variability on CYA. In general conidiophores arise from basal hyphae but also from aerial hyphae especially at colony centers. Stipes are short (35–100 µm but commonly 45–65 µm and 2–3 µm wide), smooth-walled, slightly vesiculate (3–4 µm diam) on CYA and obviously vesiculate (4–5 µm diam) on MEA (FIG. 3Go), strictly monoverticillate and rarely branching. Phialides are ampulliform, 7–11 µm long and usually appearing in whorls of 7–13 (FIG. 4Go). Conidia are spherical to subglobose, smooth-walled to finely roughened, 2.5–3 µm long x 2 µm wide and borne in long, loose columns (FIG. 5Go).

Sclerotia are formed only on MEA and are 50–100 µm long, usually globose, colorless to light buff and relatively soft (FIGS. 6, 7Go).

Specimen examined. – HOLOTYPE here designated. FIJI: Dravuni, 18°42.834'S, 178°30.343'E. F01V25, collected by scuba diving at 20–45 ft on the Great Astrolabe Reef, isolated from a sample of Dictyosphaeria versluyii collected by Dr Valerie S. Bernan, Jan 2001. The culture has been deposited in the U.S. National Fungus Collections (BPI 844248).

Etymology. – Dravuni is an island located within the Great Astrolabe Reef.

Some variation in the production of sclerotia for F01V25 was observed. As previously mentioned, sclerotia were formed only on MEA among all of the media used in phenotypic evaluation. However there was a lack of sclerotia on MEA if F01V25 was subcultured several times. Although potato-dextrose agar (PDA) is not a medium regularly used in morphological analyses of Penicillia, it is worth noting that white to buff sclerotia also were produced on PDA.

Because F01V25 was isolated from the marine environment, it seemed appropriate to determine its salt tolerance. The growth rate of F01V25 was equivalent on MEA, MEA-ASW, MEA + 5% NaCl, and MEA + 5% KCl with colonies attaining 30 mm diam after 7 d at 25 C. Increasing the salt concentration above 10% resulted in a decrease in the growth rate (FIG. 8Go). F01V25 grew even in the presence of 20% NaCl; colonies started growing after 14 d at room temperature and attained an 11 mm diam after 21 d.


Figure 2
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  		FIG. 8. Growth of F01V25 for 7 d at 25 C on MEA + 5% NaCl (top), MEA + 10% NaCl (lower left), and MEA + 15% NaCl (lower right). All images are the reverse of colonies. 9. F01V25 grown for 7 d at 37 C on MEA-ASW (left) and MEA (right).

 
The authors found it to be interesting that F01V25 grew optimally on MEA at 37 C if ASW, NaCl or KCl was added to the medium. Colonies on MEA after 14 d at 37 C were only 8 mm diam compared to 38–42 mm diam on MEA-ASW (FIG. 9Go), MEA + 5% NaCl, or MEA + 5% KCl, a fivefold increase. Even in the presence of 15% NaCl, the growth rate was twice that observed on MEA prepared without salt. As might be expected, colony morphologies were altered under these conditions; conidia were produced on MEA-ASW, MEA + NaCl, or MEA + KCl, but they lacked pigmentation and were borne on short chains that were wiry or curly.

If F01V25 is keyed using Pitt (1988)Go, Penicillium dravuni belongs in subgenus Aspergilloides section Aspergilloides based on the presence of vesicles on the stipes (FIG. 3Go) of the monoverticillate conidiophores when grown on MEA. The only two species in section Aspergilloides that produce sclerotia are Penicillium sclerotiorum and Penicillium thomii. However P. dravuni does not resemble P. sclerotiorum, which has a brilliant orange colony color, long conidiophores (100–300 µm), produces sclerotia on CYA, and does not grow at 37 C. Nor does P. dravuni share many phenotypic characteristics with P. thomii, which produces sclerotia on CYA, grows much more rapidly (40–60 mm colony diam on CYA compared to 25–35 mm for P. dravuni), has long, rough-walled stipes, rough conidia, large, hard, apricot sclerotia and grows at 5 C.

P. dravuni should be placed within the Monoverticillata, P. thomii series based on the production of true sclerotia, according to Ramirez (1982)Go. P. dravuni can be categorized further to the Penicillium turbatum subseries, which produce soft, pseudoparenchymatous, thick-walled sclerotia on only certain substrata.

Of the five species that comprise P. turbatum sub-series, P. dravuni most closely resembles P. turbatum. After 14 d at room temperature, both grow somewhat restrictedly on Czapek’s agar (CA) with a dense, felty appearance, spores produced in central areas, short stipes mostly 40–70 µm, smooth or nearly smooth conidia and no sclerotia. However the surface colony color of P. dravuni is yellow-pink, not pale grayish-green as in P. turbatum and P. dravuni is devoid of soluble pigment while P. turbatum produces a pale orange soluble pigment. In addition P. dravuni typically has 7–13 phialides at the end of the stipes; P. turbatum has few (4–8).

These two species phenotypically appear similar on CYA and MEA as well. Both exhibit dense, velvety growth with sulcation, sparse exudate, soluble pigment and no sclerotia on CYA at 14 d at room temperature. However P. dravuni does not grow as robustly, attaining only 38 mm diam compared to 45–50 mm for P. turbatum. Also P. dravuni produces abundant conidial structures, while the conidial structures of P. turbatum are limited. For both species MEA is the only medium used by Ramirez (1982)Go that supports the production of sclerotia, which are soft and crush easily. Once again, however, P. turbatum grows faster (50–60 mm diam) than P. dravuni (30–45 mm) at room temperature after 14 d.

The ITS of F01V25 is 501 base pairs (bp) in length; ITS1 is 174 bp, 5.8S is 113 bp, and ITS2 is 214 bp. Because of high sequence divergence at the end of ITS2, the last 39 bp were trimmed from the multiple alignments, resulting in removal of the last 24 bp of the ITS2 of F01V25. Of the 502 characters that were used in the maximum parsimony analysis, 385 were constant; 49 variable characters were parsimony uninformative, 68 were parsimony informative. Tree statistics were: tree length = 215, consistency index (CI) = 0.6698, retention index (RI) = 0.7321, rescaled consistency index (RC) = 0.4903, homoplasy index (HI) = 0.3302, CI excluding uninformative characters = 0.5671 and HI excluding uninformative characters = 0.4329.

In parsimony analysis P. dravuni forms a mono-phyletic group with Eupenicillium brefeldianum, E. levitum, E. reticulosporum, E. javanicum, E. ehrlichii and P. simplicissimum (FIG. 10Go). However the branching of P. dravuni from this clade is supported only by a bootstrap value of 58%. Distance analysis with neighbor joining produced a tree with a similar topology as that of maximum parsimony, however the same branch is supported by a more confident bootstrap value of 76% (FIG. 11Go). Because bootstrap values are relatively low the exact phylogenetic position of P. dravuni is uncertain, but F01V25 seems to be most closely related to Eupenicillia.


Figure 3
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  		FIG. 10. Phylogenetic tree based on sequence data from the nuclear ribosomal ITS region derived with maximum parsimony analysis. Numbers above the nodes are bootstrap values in percent (based on 1000 bootstrap samples). Values below 50 are not shown. The bar represents number of changes. The tree is rooted with Paecilomyces variottii.

 

Figure 4
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  		FIG. 11. Phylogenetic tree based on the nuclear ribosomal ITS region derived from distance analysis using neighbor joining. Numbers above the nodes are bootstrap values in percent (based on 1000 bootstrap samples). Values below 50 are not shown. The bar represents 10% (0.1) substitutions per site.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
LITERATURE CITED
 
Phylogenetic analyses are reliable methods used to distinguish among species because changes in gene sequences usually are apparent before a change in morphology has become evident (Taylor et al 2000Go). Accordingly, our first approach to identify F01V25 was a BLASTN comparison of the ITS to other ITS sequences from Penicillia in the GenBank database. The most similar ITS sequences were from several Eupenicillium spp. and Penicillium restrictum, but the similarity values were below 97.3%, suggesting that F01V25 is a distinct species.

Phenotypic analysis was the next approach employed to identify F01V25. We used the studies of Pitt (1988)Go and Ramirez (1982)Go to include as many monoverticillate Penicillia as possible. The much abbreviated "Laboratory guide to common Penicillium species" (Pitt 1988Go) lists only two monoverticillate, sclerotigenic species, P. thomii and P. sclerotiorum; Ramirez (1982)Go, however, lists eight. Of the eight species in the "Manual and atlas of the Penicillia" (Ramirez 1982Go), only five are recognized species: P. thomii, P. sclerotiorum, P. syriacum, P. turbatum and P. donkii. P. pusillum is a strain of P. phoeniceum, P. indicum is a strain of P. chermesinum (Pitt et al 2000Go) and P. grancanariae is a strain of P. thomii (Frisvad et al 1990Go). Based on overall morphology and the production of sclerotia exclusively on MEA, the only accepted species is P. turbatum, which F01V25 resembles. However F01V25 and P. turbatum share only about 91.5% similarity in the ITS (data not shown), far below the value that would be expected for intraspecific relatedness.

Secondary metabolite profiles have been used to identify both marine and terrestrial isolates of Penicillium spp. (Malmstrøm et al 2000Go). In fermentation media containing 50% ASW, F01V25 produces the metabolites carviolin and dictyosphaeric acids A and B (Bugni et al 2004Go). The anthraquinone carviolin is a pigment that is produced by P. carminoviolaceum (Hind 1940Go), now P. roseopurpureum (Pitt et al 2000Go), and also by the ascomycete Neobulgaria pura (Eilbert et al 2000Go). F01V25 cannot be a strain of P. roseopurpureum because they share only about 95% sequence similarity in the ITS (data not shown). In addition they do not share many phenotypic characteristics; P. roseopurpureum has cerebriform conidia, reddish pigments diffusing into agar substrates and does not produce sclerotia (Ramirez 1985). The recently described dictyosphaeric acids (Bugni et al 2004Go) are polyketides that thus far are produced only by P. dravuni. In fact the only other known compounds that are similar in structure to the dictyosphaeric acids are the colletofragarones A1 and A2 (Inoue et al 1996Go), which are spore germination self-inhibitors produced by Colletotrichum fragariae. Therefore the uniqueness of the secondary metabolites produced by P. dravuni is further evidence supporting it as a distinct species.

Kohlmeyer and Kohlmeyer (1979)Go stated that one-third of all known higher marine fungi are associated with algae and these relationships may be parasitic, saprobic or symbiotic. Because F01V25 was the only isolate of P. dravuni recovered from the macerated inner tissues of D. versluyii, and the algal sample was not examined microscopically, it is not known what relationship exists between these two organisms. The growth form of this alga is like a flattened cushion that sometimes can be dusted with sediment or rubble. Perhaps conidia of P. dravuni along with some sediment were attached to the bubble-like cells of the thallus or were caught in the crevices between the algal cells of D. versluyii. After all Penicillium spp. are considered to be terrestrial fungi whose spores are washed into the ocean. However Penicillium spp. have been isolated from algae in the tropics from offshore locations (Morrison-Gardiner 2002Go), suggesting that these cultures can adapt to a marine environment and no longer may be considered strictly terrestrial. In any case the true host or niche of P. dravuni within the marine environment cannot be confirmed until additional strains of this fungus are discovered.

P. dravuni is not an obligate halophile. At 25 C this culture grows as well on media prepared without salt as it does with ASW or 5% salt. However after 21 d at 25 C on media containing 20% NaCl, F01V25 remarkably attains a colony diam of 11 mm and produces conidia. This degree of salt tolerance has been reported for fungi isolated from salt-preserved fish (Wheeler et al 1988Go), but one might expect similar behavior for fungi that originate from the marine environment or have adapted to it.

Optimum growth of F01V25 at 37 C occurs on media prepared with ASW or 5% salt. This growth response is similar to the Phoma-pattern (Ritchie 1957Go), which is defined as an increase in salinity tolerance with increasing temperature and is a phenomenon that occurs among marine fungi grown in the laboratory. One theory is that the Phoma-pattern might be an adaptation of fungi enabling them to survive in intertidal zones where water recedes and salinity and temperature rise as the sun evaporates pools (Lorenz and Molitoris 1992Go). Dunn and Baker (1983)Go tested five marine fungi for growth responses to salinity and temperature: Aspergillus tubingensis, Gliomastix murorum, Gymnoascus reesii, A. versicolor and Penicillium corylophilum, which they isolated from the psammon habitat of the Enewetak Atoll. They found that for most isolates the salinity optimum was around the concentration of seawater with temperature optima 25–37 C (Dunn and Baker 1983Go). The ability of F01V25 to grow optimally at higher temperatures only in the presence of some salt may be an adaptation of F01V25 to the fluctuations in salinity and temperature that occur in marine environments.

Penicillium spp. – have been isolated from algae, but to our knowledge they are not described (Numata et al 1993Go) or have been identified as known species (Amagata et al 1998Go). P. dravuni is the first newly described Penicillium sp. isolated from an alga. Phylogenetic data, morphological characteristics and unique secondary metabolite profiles fail to classify F01V25 with any known monoverticillate Penicillia and indicate that P. dravuni is indeed a new species. This unique isolate joins a short list of other Penicillium spp., such as P. limosum (Ueda 1995Go), isolated thus far from the marine environment.


   ACKNOWLEDGMENTS
 
The authors thank Stephen Peterson for valuable answers to frequent questions and for sequencing and depositing many ITS-LSU sequences of Penicillia to GenBank. At Wyeth, we thank Dave Fruhling for providing sequencing and Jennifer Liang for the SEM images. We also thank the National Cooperative Drug Discovery Group (NCDDG) that made the collection possible.


   FOOTNOTES
 
Accepted for publication October 12, 2004.

1 Corresponding author. E-mail: jansoj{at}wyeth.com


   LITERATURE CITED
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 LITERATURE CITED
 
Amagata T, Minoura K, Numata A. 1998. Cytotoxic metabolites produced by a fungal strain from a Sargassum alga. J Antibiot 51:432–434.[Medline]

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