APPLIED AND ENVIRONMENTAL MICROBIOLOGY, JUly 1991, p. 1963-19680099-2240/91/071963-06$02.00/0Copyright (C 1991, American Society for Microbiology

Anaerobic Growth and Fermentation Characteristics ofPaecilomyces lilacinus Isolated from Mullet Gut

DOUGLAS 0. MOUNTFORT* AND LESLEY L. RHODES

Cawthron Institute, Private Bag, Nelson, New Zealand

Received 14 January 1991/Accepted 1 May 1991

The anaerobic growth and fermentation of a marine isolate ofPaecilomyces lilacinus is described. The funguswas isolated from mullet gut and grew optimally at 30°C and at a salinity of 210%o. The best growth was

obtained with glucose or laminarin as substrate, and the growth yield was 5.0 g (dry weight of fungus) per molof hexose fermented. Moles of products as a percentage of moles of hexose fermented were acetate, 29.0%;ethanol, 156.6%; C02, 108.0%; and lactate, 4.3%. Together these products accounted for >80% of hexosecarbon. Hydrogen and formate were not detectable as fermentation end products (<0.5%). Other substratesutilized for growth, although less effectively than laminarin or glucose, included the monosaccharidesgalactose, fructose, arabinose, and xylose and the disaccharides maltose and cellobiose. No growth of thefungus occurred on cellulose, and of a variety of other polysaccharides tested only xylan supported growth.

The study of organisms that participate in the degradationof algal material has attracted considerable interest in recentyears because of the potentially useful products that can beobtained from such transformations. To this end marinemicroorganisms have been extensively studied for theirability to produce enzymes capable of degrading algalpolysaccharides (7, 8, 10, 19, 21, 30, 34, 35, 38, 39).However, there has been little or no investigation of micro-organisms that reside in the guts of vegetation-feeding fish,such as algal grazers. Yet, these organisms could be ofconsiderable importance to biotechnology by (i) possessingunique degradative enzymes and (ii) facilitating importantbioconversions.

In this communication we describe the growth of andanaerobic fermentation by Paecilomyces lilacinus isolatedfrom mullet gut and also examine the range of substrateswhich support growth of the fungus. The results are dis-cussed in relation to the possible role this organism couldhave in the mullet gut ecosystem, and the implications of thefindings to the degradation of algal polysaccharides are alsoconsidered.

MATERIALS AND METHODSFungal strain. P. lilacinus described in this communication

was identified by E. MacKenzie, Plant Protection Division,Department of Scientific and Industrial Research, Auckland,New Zealand. The fungus has been deposited in the Inter-national Collection of Micro-organisms at the Plant Protec-tion Division (ICMP no. 10835).

Culture media. Anaerobic techniques for the preparationand use of media were mainly based on those of Hungate (13)as modified by Bryant (4) and Balch and Wolfe (2). The basalmedium contained 30% (vol/vol) seawater, 0.001% (wt/vol)resazurin, 0.01% (wt/vol) NH4NO3, 2% (vol/vol) solution of5% NaHCO3, 2% (vol/vol) solution of 0.2 M sodium phos-phate buffer (pH 6.5), 2% (vol/vol) reducing solution (1.25%Na2S 9H20 and 1.25% cysteine-HCl), and 1% (vol/vol)vitamin solution (20). The gas phase was 80% N2-20% CO2with a final pH of 6.5. Medium containing the above com-

ponents with the exception of NaHCO3, cysteine-sulfide,

* Corresponding author.

and phosphate was boiled, cooled, and dispensed in 8.5-mlamounts into culture tubes (18 by 150 mm) or in 60-mlamounts into 120-ml bottles modified for use with blackrubber septum stoppers and aluminum serum cap closures(2). The remaining components were prepared as separatesolutions, sterilized, and added to individual tubes afterautoclaving, usually several hours before use. Solid mediumfor anaerobic agar plates was prepared by including 2% agarin the basal medium. Medium was autoclaved in a stopperedround-bottom flask, cooled to 45 to 50°C, and poured intoplates in an anaerobic glove box (Coy Laboratory ProductsInc., Ann Arbor, Mich.) after addition of NaHCO3, cysteine-sulfide, and phosphate.Aerobic media were prepared with the same components

as anaerobic media, except that cysteine-sulfide, NaHCO3,and resazurin were omitted. Liquid media were dispensed in10- or 50-ml volumes into conical flasks which were pluggedwith cotton-wool bungs and autoclaved. Solid media for agarplates were prepared with 2% agar.

Soluble sugar substrates were prepared in concentratedsolution and filter sterilized prior to addition to culturemedia. Insoluble substrates were added to culture mediaprior to autoclaving.

Culture techniques and conditions. Cultures were main-tained aerobically by transfer of agar bearing thallus to newplates every 3 weeks or by transfer of thallus from agarslants stored at 2 to 4°C every 2 months. Cultures were alsomaintained anaerobically by transfer of 1/20 the volume ofanaerobic liquid culture at full growth to new culture tubesor by transfer of agar bearing thallus to new anaerobic agarplates. Experimental media were inoculated by the same

procedures as for the maintenance of anaerobic liquid cul-ture, and the inoculum was equivalent to between 1.0 x 106and 1.5 x 106 CFU.

Cultures were routinely incubated at 30°C, and tubes were

kept in a vertical position with no shaking. Anaerobic agarplates after inoculation were placed in an airtight anaerobicsteel jar, the gas phase of which was adjusted to 80%N2-20% CO2.

Quantitative analysis of substrate and products in culturemedia. With the exception of glucose, quantitative determi-nation of sugars was carried out by the anthrone method (1).

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Glucose was assayed with glucose oxidase (Fermcozyme9520M; Finnsugar Biochemicals, Schaumburg, Ill.) (12).Gas analysis for H2 was carried out by gas chromatogra-

phy as previously described (4), and CO2 was determined bycounting 14Co2 produced from [U-14C]glucose (specific ac-

tivity, 2.5 x 103 dpm ,umol`). For the latter, culturemedium in anaerobic growth tubes was acidified to pH 2.0 byinjection of 0.4 ml of 50% (vol/vol) H2SO4 to ensure that allthe CO2 was liberated from the medium. The stopper was

pierced with two syringe needles (19 gauge), and CO2 was

flushed from the tubes by bubbling N2 through the mediumfor 1 h. 14CO2 in the effluent gas stream was trapped inethanolamine-methoxyethanol (2:1, vol/vol), and portionswere counted in toluene-based scintillant by the method ofJeffay and Alvarez (14). The amount of CO2 was calculatedby assuming that the specific activity of 14CO2 released fromthe fermentation was the same as that for [U-14C]glucose on

a gram-atom of C basis.For the determination of volatile fatty acids (except for-

mate) and alcohols, cultures were centrifuged at 6,000 x g

for 15 min at 2°C, and 2-ml portions of supernatant were

acidified with 0.1 ml of 6 N HCl. Analysis of supernatant was

carried out by gas chromatography on an Alltech SuperoxFA column (10 m by 0.53 mm) with a N2 flow rate of 3.1 mlmin- 'in a Hewlett-Packard (model 5890) gas chromatographequipped with a flame ionization detector. After injection ofsample (1 ,ul), the column temperature was held at 35°C for2 min, followed by a ramp of 25°C min-1 up to 150°C andmaintenance of the latter temperature for a further 2 min.Fatty acids and alcohols were identified and quantitated bycomparison of retention times and peak areas with those ofknown standards. Formate was determined by using theformic dehydrogenase assay test kit (Boehringer GmbH,Mannheim, Germany).

Analysis of nonvolatile acids in the supernatant of culturemedia was by gas-liquid chromatography of the correspond-ing methyl esters as previously described (3).Growth determinations. Growth was determined by mea-

surement of chitin by the method of Ride and Drysdale (28)with glucosamine as the standard. The relationship betweenchitin and dry weight of the fungus was found to be linear,with 1.0 mg of glucosamine being equivalent to 39.3 mg (dryweight). Growth rate was determined by measurement of therate of substrate disappearance in cultures.

Electron microscopy. Samples were prepared for electronmicroscopy by fixation in glutaraldehyde-formaldehyde in0.1 M phosphate buffer (pH 7.2) and then in 1% osmiumtetroxide. After repeated buffer washes followed by dehy-dration through a graded ethanol series to 100% ethanol,samples were mounted for scanning electron microscopyand coated with approximately 20 nm of gold. Specimenswere observed with a Cambridge 250 Mark 3 scanningelectron microscope.

Chemicals. All chemicals were obtained from commercialsources and were of reagent grade. The radioisotope[U-_4C]glucose (specific activity, 255 ,uCi/,umol) was ob-tained from the Radiochemical Centre, Amersham, England.

RESULTS

Isolation and description of the fungus. The fungus was

isolated by inoculation of 0.5 ml of 10-1 to 10-7 dilutions ofgut contents of mullet (Aldrichettaforsteri) onto aerobic agar

spread plates in a sterile laminar flow chamber. The platemedium contained as the substrate mixed dried and groundalgae (kelp plus Gracilaria sp.) at 0.5% (wt/vol) and also an

antibiotic mixture of benzylpenicillin (103 IU ml-') andstreptomycin sulfate (102 IU ml-1). Pinkish colonies devel-oped on plates after 1 week of incubation and represented 2x 103 CFU/ml of mullet gut contents; considerably lowernumbers were found in seawater or associated with algaeregularly grazed by mullet (<102 CFU/ml of seawater or perg [wet weight] of algae). The colonies reached a diameter ofbetween 20 and 35 mm after 2 weeks of incubation. Noobvious presence of bacteria was evident. A portion of thethallus of one of the fungal colonies was further transferred,and the culture was successfully maintained on agar platesby being transferred every 3 weeks. Light microscopy wasused to establish the purity of the culture.

Aerobic growth of the culture on agar plates or in liquidmedia resulted in extensive hyphal growth with large num-bers of conidia (Fig. 1A and B), many of which occurred inloosely organized chains. Conidia were ovoid and about 2.5pLm in length and 1.5 ,Lm in width. The size and shape of theconidia together with the shape of the phialides with theswollen basal part tapering into a distinct neck (Fig. 1A) andthe vinaceous color of the conidial heads were consistentwith those of the organism P. lilacinus. In anaerobic growththe number of conidia was smaller, and there was extensivemycelial growth (Fig. 1C). Conidia were similar in size tothose obtained in aerobic growth, although some were moreelongated (Fig. 1D). In actively growing cultures conidiawere shown to germinate and produce new hyphae.Optimal conditions for anaerobic growth. The optimal

temperature for growth of the fungus on glucose was 30°C,and the optimal salinity was .10%7o (Fig. 2). Growth rateswere determined by measurement of the rate of substratedisappearance from the cultures, since the relationship be-tween substrate disappearance and production of fungalbiomass was linear (Fig. 3 and 4). Very little growth of theculture occurred above 37°C, and the lowest salinity givingoptimal growth (10.5%o) was the same as that of mullet gutfluid.

Fermentation products. The fermentation of glucose by thefungus resulted in the formation of three major products,ethanol, acetate, and CO2 (Table 1; Fig. 3). Lactate was alsoproduced in low quantities. Product formation was matchedby a decline in glucose levels and an increase in fungalbiomass, as determined by chitin measurement (Fig. 3).Hydrogen was not detected during the fermentation (detec-tion limit, 5 x 10-4 atm [1 atm = 101.3 kPa]), and otherpossible products which were not detected (<0.1 mol/100mol of substrate fermented) included succinate, isopropanol,acetone, propionate, butyrate, methanol, and formate. Fer-mentation of glucose terminated after about 4 weeks ofincubation, and at the end of fermentation approximately62% of the initial glucose had been fermented. The numberof moles of product per 100 mol of glucose fermented ispresented in Table 1, and together acetate, ethanol, and CO2accounted for >80% of the carbon recovery. The oxidation-reduction index for the products was 0.7, which approachesthe value of 1.00 for the starting substrate.

Substrate specificity during anaerobic growth. Although thebest growth of the fungus was obtained on laminarin andglucose, a variety of other carbohydrates also supportedgrowth (Table 2). These included the monosaccharides ga-lactose, arabinose, mannose, fructose, and xylose togetherwith the disaccharides maltose and cellobiose. Of thepolysaccharides tested, apart from laminarin only xylansupported growth, and no growth occurred with cellulose.Anaerobic versus aerobic growth. When the organism grew

under aerobic conditions the yields of fermentation end

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ANAEROBIC GROWTH AND FERMENTATION OF P. LILACINUS 1965

FIG. 1. Scanning electron microscopy of P. lilacinus grown aerobically (A and B) and anaerobically (C and D). Bars, 10 ,um (A and C) and4 ,um (B and D). Abbreviations: c, conidia; p, phialide.

products declined and only a small proportion of the sub-strate utilized was actually fermented compared with anaer-obic conditions, in which maximal yields of fermentation endproducts were obtained (Tables 1 and 3). Growth yields,however, were substantially higher for aerobic growth thanfor anaerobic growth, reflecting the higher energy yieldsobtained from aerobic metabolism.

DISCUSSIONThe anaerobic growth and fermentation of P. lilacinus has

not previously been documented, although the fungus hasbeen widely reported in nature, including many soils and insediments from estuarine habitats (6). In this communicationwe have described the anaerobic fermentation of a marine

isolate of P. lilacinus. We had earlier observed that thisisolate was the only example of more than 100 fungi isolatedfrom marine environments capable of anaerobic growth (24).The fermentation by the organism appears similar to those ofthe enteric bacteria but is distinct from those carried out bymany yeasts and some other filamentous fungi which resultin only ethanol and CO2 as products (9, 32).

It is of interest that neither hydrogen nor formate wasproduced as a fermentation end product, so in the gutecosystem P. lilacinus and other fungi with similar fermen-tations would be unlikely to participate in any form ofinterspecies hydrogen transfer, which is in contrast to thesituation found with anaerobic fungi in the guts of ruminants(22). It is not yet clear what the likely role of fungi such as P.

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1966 MOUNTFORT AND RHODES

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FIG. 2. Effects of temperature (A) and salinity (B) on the growthrate of P. lilacinus. Culture tubes containing basal medium (8.5 ml)with 0.03 g of glucose were inoculated with fungus equivalent toapproximately 1.5 x 106 CFU. Growth rates were determined bymeasurement of the rate of glucose disappearance in cultures.

lilacinus would be in the gut of mullet, but one possibilitywould be participation in the breakdown of complex marinealgal polysaccharides, rendering these assimilable by thehost. The ability of P. lilacinus to grow on both xylan andlaminarin together with a range of disaccharides wouldindicate that this organism possesses some of the enzymesrequired to fulfill this role. Both xylan and laminarin areimportant components of marine algae (25), the former beingthe main structural component of the cell walls of most greenalgae and the latter being a major component of brown algae.A major component of the mullet's diet consists of greenalgae, namely, Ulva sp. and Enteromorpha sp., and there-fore it is not surprising that organisms isolated from the gutwould possess xylanase activity. Likewise, the presence ofbrown alga fragments found in the gut has led to theestablishment of a microbial population capable of degradinglaminarin (23).The inability of our isolate to grow on cellulose is not

surprising, since this may reflect the small percentage thatcellulose composes in the algae that form the mullet's diet. Itis of interest that with the exception of laminarin, solublepolysaccharides did not support growth of the fungus (Table

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TIME (DAYS)FIG. 3. Time course of glucose fermentation by P. lilacinus.

Media (8.5 ml) were inoculated with 1.5 x 106 CFU. Symbols: 0,

glucose; 0, ethanol; A, acetate; 0, lactate; *, total dry weight offungus.

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cc Iv0 0.2 0-4 0-6 0-8 1-0 1-2GLUCOSE UTILIZED (1021tmoI)

FIG. 4. Dry weight of fungus produced from growth on glucose.Values were determined from the data in Fig. 3 and represent totalamount of fungus produced versus total glucose utilized in culturesat various stages in the time course.

2). This finding may indicate that the organism and othersimilar fungi may be specialized in the gut towards thebreakdown of algal structural polysaccharide in a mannersimilar to anaerobic fungi in the ruminant gut, which have aspecial role in the degradation of plant structural polysac-charide (22). Furthermore, a recent study of more than 80aerobic marine fungi by Rhodes et al. (27) has similarlyrevealed the inability of their isolates to degrade most of thecommon soluble algal polysaccharides, with the exception oflaminarin.Enzymes involved in the breakdown of laminarin have

been examined in some detail in a variety of fungi (11, 17, 33,36), and those involved in the breakdown of xylan have beenwidely studied. Yet, there has been little investigation of thecharacteristics of laminarinases and xylanases in marineisolates. The study of these enzymes in our isolate wouldtherefore offer an opportunity to determine whether theyfacilitate transformations which are different from thosealready known. Also of possible significance to bioconver-sions is the finding that our isolate produces ethanol as themajor product from xylan degradation, which represents oneof the few examples of this type of transformation to becarried out by a fungus. Traditionally, conversions of cellu-lose and hemicellulose to ethanol have involved the coupling

TABLE 1. Fermentation of glucose by P. lilacinus grownunder anaerobic conditionsa

Parameter (unit) Valueb

Product (mol/100 mol of glucose)Acetate ........................................ 29.0 ± 4.9Ethanol ........................................ 156.5 ± 23.0Carbon dioxide ........................................ 108.0 + 18.0Lactate........................................ 4.3 ± 0.3

Carbon recovery (%) ........................................ 82.0Hydrogen recovery (%) ....................................... 89.6Oxidation-reduction index ................................... 0.70

" Determined at the completion of fermentation, as ascertained by nofurther decrease in glucose. The percentage of glucose utilized at the end ofincubation was 62%.

Values are means of at least duplicate determinations and, where the erroris presented, 1 standard deviation.

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ANAEROBIC GROWTH AND FERMENTATION OF P. LILACINUS 1967

TABLE 2. Substrates utilized as energy sources for anaerobicgrowth of P. lilacinusa

Substrate Growthb(,ig [dry wt], 102)

Glucose ...................................... 6.0Arabinose ...................................... 3.9Galactose....................................... 2.9Mannose ...................................... 2.9Fructose ...................................... 2.7Xylose...................................... 4.4Cellobiose ...................................... 3.5Maltose ............ .......................... 3.1Xylan ...................................... 4.5Laminarin .............. ........................ 6.1No addition ................ ...................... 1.0Otherc ...................................... 1.0-1.5

a Culture tubes containing basal medium (8.5 ml) with 0.03 g of substratewere inoculated with fungus equivalent to 1.5 x 106 CFU and incubated for upto 60 days.

b Values were taken when no further growth of the fungus was evident andwere determined from chitin measurements (1 mg of glucosamine is equivalentto 39.3 mg [dry weight]).

c Substrates which did not support anaerobic growth of the fungus includedcellulose, carrageenan, pectin, fucoidin, agarose, and alginic acid.

of the fermentative ability of a yeast with the addition ofcellulases or hemicellulases (5, 29, 37), but more recentlythere have been attempts to carry out this process in a singlestep by using filamentous fungi which possess these enzymes(9, 26), and our study extends the range of fungi capable ofthis type of transformation to include P. lilacinus. Yields ofethanol were higher during anaerobic growth of the fungusthan during aerobic growth (Table 3), a finding not unusualfor fungi which show both respiratory and fermentativecapabilities (32), although some fungal fermentations havebeen reported to be enhanced by the presence of low levelsof oxygen (15, 16, 18, 31).

In this communication we provide the first report ofanaerobic growth of P. lilacinus in a strain isolated from themarine environment. The characteristics of this isolate to-gether with similar organisms should offer new scope towardunderstanding the role that microorganisms play in the gutecosystem of alga-grazing fish. Further investigation of suchorganisms may yield enzymes capable of unique transforma-

TABLE 3. Growth yield and yields of ethanol and acetate fromgrowth of P. lilacinus in various substrates and conditionsa

Yield (moU/100 mol of hexose Growth yieldSubstrate and or pentose) (g [dry wtj/Susrt an mol of

condition hexose orAcetate Ethanol pentose)

GlucoseAnaerobic 29.0 ± 4.9 156.5 ± 23.0 5.0Aerobic 14.8 ± 2.8 41.3 ± 5.0 37.5 + 3.5

Laminarin, anaerobic 33.5 ± 6.3 152.5 ± 14.8 5.1XylanAnaerobic 36.3 ± 5.0 143.8 ± 9.3 2.6Aerobic 13.4 ± 1.2 4.2 ± 0.1 26.0 + 1.4

a Growth of P. lilacinus was carried out in anaerobic medium (8.5 ml) or inaerobic medium (10 ml) containing 0.03 g of substrate. Values are means of atleast duplicate determinations and, where the error is presented, + 1 standarddeviation. Measurements were made at the completion of growth as indicatedby no further utilization of substrate. In anaerobic cultures approximately60%o of substrate was utilized at full growth, and in aerobic cultures xylan andglucose utilization amounted to 70 and 95%, respectively.

tions of algal polysaccharides, giving products with valuablebiotechnological applications.

ACKNOWLEDGMENTS

We gratefully acknowledge the expert technical assistance givenby Rod Asher, Jan Holland, and David White. We also express ourappreciation for helpful comments from Don Grant during thepreparation of the manuscript and to Doug Hopcroft of the Depart-ment of Scientific and Industrial Research, New Zealand, forelectron microscopy. We thank Yvonne Graham for typing themanuscript.The work was supported by the New Zealand government.

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