evaluation of methods to extract ergosterol for ... - Science Direct

0038.0717/92$5.00+ 0.00 Pergamon Press plc

Soil Bioi. Biochem.Vol. 24, No. 3, pp. 189-198,1992 Printed in Great Britain.

EVALUATION OF METHODS TO EXTRACT ERGOSTEROL FOR QUANTITATION OF SOIL FUNGAL BIOMASS MARK W. DAVIS and RICHARD T. LAMAR Institute for Microbial and Biochemical Technology, USDA Forest Service, Forest Products Laboratory, 1 Gifford Pinchot Drive, Madison, WI 53705, U.S.A.

Summary-Two methods were evaluated to extract ergosterol for quantitation of fungal biomass in Marshan, Zurich, and Batavia soils. Yields of ergosterol from hyphae and from fungal-colonized soil were greater when fungal tissue was extracted with an alkaline solvent mixture than when base was added to neutral extracts following removal of solids. A lyopbilizatjun treatment prior to extraction increased yields from Marshan but not from Zurich and Batavia soils. Losses of ergosterol during lyophil~zation were prevented by a rapid freezing treatment before lyophilization of soil samples. Recoveries from soil fortified with pure ergosterol did not accutately model recoveries from fungal tissue in these substrates. Thus, determinations of extraction efficiencies should be based upon recoveries from fungal tissue added to soils. Ergosterol was quantitatively recovered from Marshan and Zurich soils fortified with fungal tissue; however, only ca 66% was recovered from Batavia soil, a subsoil with a high clay content. The limit of detection of ~~u~era~~~g~~c~~~s~~~~$~~~~ from the three soils ranged from 8 to 15 pg biomass g soil-‘.

Direct strates

quantitation of fungal growth in solid subis not possible because of the difficulty in

isolating fungal hyphae from the substrate. The use of indirect methods to estimate fungal growth have been reviewed by Matcham et al. (1984). Ergosterol content has been shown to be a more sensitive and reliable indicator of fungal growth than other biochemical indices, including glucosamine (Seitz et al., 1979; Matcham et al., 1985; Grant and West, 1986; Johnson and McGill, 1990) and extracellular lactase (Mateham ef ai., 1985). Ergosterol is a particularly useful index of fungal growth because it is endogenous only to fungi (Weete, 1989) and certain green microalgae (Newell er al., 1987). Its maximal absorbance at 282 nm makes ergosterol readily distinguishable from the major sterols of animals, vascular plants, and non-fungal microbes. Seitz et al. (1977, 1979) pioneered the use of ergosterol content as a measure of fungal biomass in solid substrates using fungal-infested grain. Their methodology involved sample homogenization in methanol, filtration, saponification of the extract, partitioning of non-polar extractives to an organic solvent, and quantitation by high pressure liquid chromato~ra~h~~ (HPLC). This procedure has been widely used to measure fungal growth in plant tissue (Miller et nl., 1983; Cordon and Webster, 1984; Newell et al., 1987). A modification of this procedure (Grifiths et al., 198S), wherein the sample is refluxed in basic ethano1, has been variously reported to increase (Nout ef af., 1987) and to reduce (Newell et al., 1958) ergosterol recoveries from those obtained by the method of Seitz et al. (1977). Other researchers

have used simiIar but distinct extraction procedures (Lee et al., 1980; Matcham ef al., 1985). The efficacies of these methods for extracting ergosterol from solid substrates have not been adequately assessed. The cited studies correlated substrate ergosterol contents with quafitative indicators of fungal growth. However, extraction eRicienties were addressed in only a single study (Seitz ef al., 1977), in which 93% of the sterol added to plant tissue was recovered. Subtle modifications of the extraction protocol signi~~ntly affected ergosteroi recoveries from plant tissue (Newell et at., 1988). Application of the method to soils has been reported by Grant and West (1986). In this study, soil ergosterol contents were correlated with fungal biovolume measurements. Although recoveries from soil of 88 to 96% were reported, ergosterol was added with extractants rather than directly to soil prior to the addition of solvents. Addition of the sterol with extractants addresses potential losses during extraction but not extraction efficiencies from the soil. Our primary purpose was to assess the e@icacy of existing methodologies, developed for extracting ergosterol from infected plant tissue, for quantitative extraction of ergosterol from soil. Our approach was to evaluate modi~cations of the procedures developed by Seitz et af. (1977) and GriRiths et al. (1985) using fungal-colonized soil. The modification that exhibited the greatest assay precision and gave the highest yields from funpal-colonized soil was then characterized to establish recoveries and sensitivities from soil fortified with fungal tissue or with pure ergosterol. Secondary goals were to (1) evaluate the procedure for quantifying fungal bioamss of

190

MARK W. DAVIS and RICHARDT. LAMAR

colonized wood chips and (2) develop a procedure rapid analyses of multiple samples.

MATERIALS

for

AND METHODS

Soils

Soils were Marshan sandy loam (fine-silty, mixed, mesic, Mollic-Hapludalf), Zurich sandy loam (finesilty over sandy, mixed, mesic, Typic-Haplaquoll), and Batavia silty clay loam (fine-silty, mixed, mesic, Typic-Hapludalf). Soils were air-dried, sieved (< 2 mm), and stored in polyethylene bags at 4°C. Soil physical and chemical properties were reported by Lamar et al. (1990). Chemicals

Ergosterol was purchased from Sigma Chemical Co. (St Louis, MO) and recrystallized from ethanol. All other chemicals were reagent grade. Fungus

and inoculum

preparation

Phanerochaete chrysosporium Burds. (BKM F1767; ATCC 24725) (Burdsall and Eslyn, 1974) was grown on 2% malt agar slants at 39°C for 1 week and then stored at 4°C. Soil inoculum consisted of aspen (Populus tremuloides Michx.) pulpwood chips (ca 1.5 x 0.5 x 0.25 cm) grown through with P. chrysosporium. Chips were placed in quart canning jars and the moisture content of the chips adjusted to 30% with deionized distilled water. Jars containing chips were covered with aluminum foil and sterilized by autoclaving at 121°C for 1 h on 2 consecutive days. The moisture content of the chips was then adjusted to 60% by the addition of sterile chemically defined medium. Supplemented chips contained the following: glucose (27 g kg-‘), L-glutamic acid (268 mg kg-‘), KH,PO, (2.1 g kg-‘), MgSO, 7H20 (2.1 g kg-‘), CaCl, 2H,O (40 mg kg-‘), MnCl, 4H,O (40 mg kg-‘), ZnCl, (10 mg kg-‘), trace element (1.07 ml kg- I), and vitamin solution solution (1.07 ml kg-‘) (Leatham, 1983). Sterile chips were inoculated by exposing them to pieces of malt agar from P. chrysosporium slants. Inoculated chips were kept at 39C for l-8 weeks. Culture

conditions

Soil cultures were prepared as follows. The water potential of the soils (SWP) was adjusted to - 150 kPa (1 bar = 100 kPa) with deionized distilled water. Moisture contents (dry weight basis) of Marshan, Zurich, and Batavia soils at this SWP were 32.9, 15.6, and 21.3%, respectively. Soils (ca 60 g dry wt) were dispensed into 240 ml canning jars (Ball Corp., Muncie, Ind.) and sterilized by methyl bromide fumigation (Lamar et al., 1990). Sterile soils were inoculated by aseptically placing ca 2% (fresh wt basis) colonized chips on their surfaces. Soil cultures were then covered with modified canning jar lids. Lids were modified by making a 32 mm hole in the center of the lid and gluing a microporous polypropylene film

(Cell-Guard, Hoechst Celarese Corp., Somerville, N.J.) over the hole, on the bottom of the lid. This minimized evaporative losses of moisture while providing adequate aeration (data not shown). Cultures were kept at 32°C until fungal sporulation in the soil was observed (ca 1 week). For liquid cultures, fungi were grown in stationaryphase cultures containing 10 ml of culture medium in 125-ml Erlenmeyer flasks. Culture medium contained the following: glucose (10 g l- ‘), thiamine HCl (1 mg ll’), ammonium tartrate (200 mg 1 ‘), KH,PO, (2 g ll’), MgSO, 7H, 0 (500 mg 1~’ ). Ca(H,PO,), (lOOmgll’), and a trace mineral solution (10 ml 1-l) in 2,2_dimethylsuccinic acid buffer (1.46 g ll’), adjusted to pH 4.5 (Kirk et al., 1978). Cultures were inoculated by adding 1% (v/v) conidial suspensions (2.5 lo6 spores mll’) of P. chrysosporium; conidia were taken from slants stored at 4 ‘C. Flasks were loosely capped with aluminum foil and kept at 39°C. Dry weight determinations

The moisture contents of soils and wood chips were determined gravimetrically after drying at ca 80-C for > 18 h. Hyphal mats from liquid culture were collected by filtration (Whatman No. 1, Fisher Scientific Co., Pittsburgh, Pa), rinsed with water, removed from filters, placed in tared vials, lyophilized for 3 h. and weighed. Analytical

procedures

Dry weights and residual moisture contents of vacuum-dried soils and wood chips were determined by reference to oven-dried replicates. For analyses of ergosterol contents of soil cultures, inoculum chips were removed and the soil in each flask thoroughly mixed. Tests (data not shown) had indicated that milling treatments of Marshan soil did not increase ergosterol yields when subsequent extraction times were of sufficient duration (420 h). However, a milling treatment was necessary to maximize recoveries from chips. Soils were placed in 25 x 150 mm culture tubes, frozen in an acetone-dry ice bath, and lyophilized for 5 h prior to extraction. Wood chips were lyophilized for 16 h and milled (Miracle Mill, Markson Science Inc., Phoenix, Ariz.) to ca 1 mm prior to extraction. Total neutral-extractable (TNE) ergosterol was determined using a procedure modified from that of Seitz et al. (1977). Soil samples (ca 8 g dry wt) in 25 x 150 mm culture tubes with Teflon-lined screw caps (Corning Inc., Corning, N.Y.) were extracted in 25 ml of a mixture of n-hexane-methanol (1:4) for 16 h on a rotating tumbler shaker (ca 33 rev min ‘). Soil was removed by vacuum filtration through G6 glass fiber filters (Fisher Scientific Co., Pittsburgh, Pa) into 25 x 150 mm culture tubes. Soil remaining on the filter was consecutively rinsed with 5 ml methanol and 5 ml n-hexane. Care was taken to avoid channeling in the soil cake, which tended to occur during this

Quantitation of soil ergosterol step. The extract was saponified by adding 2.25 g KOH (pellets), capping the tubes, solubilizing added base by shaking (ca 10 min on a rotating tumbler), and refluxing in a steam cabinet for 30 min. After cooling at room temperature, 5 ml distilled water (ca 20% of methanol volume) was added and the tubes vigorously shaken. Following clarification of the aqueous phase (ca 2 h), the hexane layer was transfered to 12-ml amber screw-cap vials. A second hexane extraction (3 ml) was performed, and the pooled hexane portions were evaporated to dryness under N? Samples were solubilized in 0.25-l .OOml methanol by heating capped vials at 60°C for 30 min. Gravimetric corrections for any losses of solvent volume during solubilization were made prior to filtration through 0.5pm Teflon syringe filters (Corning Inc., Corning, N.Y.). Free ergosterol determinations were performed as except that the saponification described above, step-addition of KOH and refluxing-was omitted. Phase separation of these neutral extracts required the addition of 5 ml 5.5% NaCl. Total alkali-extractable (TAE) ergosterol was determined using modifications of the method of Griffiths et al. (1985). Soil samples were extracted in 25 ml n-hexane-10% KOH in methanol (1:4) on a rotating tumbler shaker for 16 h. Soil was removed from extracts by vacuum filtration as described above, with the exception that the methanol rinse was replaced by a rinse with 10% KOH in methanol. The alkaline extracts were then refluxed for 30min. Subsequent processing was identical to that described for the TNE ergosterol extracts. Aspen chips were processed identically as soils except that chips were not removed from extracts prior to refluxing. Ergosterol in the methanol solutions was quantified by HPLC. The columns were a Vydac 201TP C,, reverse-phase analytical column (0.46 x 25 cm) (Separations Group, Hesparia, Calif.), preceded by a Vydac 2llGCC54T reversephase C,, guard column. Extracts (20~1) were injected and eluted with methanol at 45°C at a flow rate of 0.75 ml min’. Column eluant was monitored for absorbance at 282 nm. An isolated peak eluting at ca 15 min retention time was identified as ergosterol based on its co-chromatography and identical absorption spectrum with pure standard. Pure ergosterol used as standard was weighed and solubilized in 50 ml methylene chloride in a volumetric flask. Measured portions were then dispensed to 2-ml amber screw-top vials, dried under N,, and stored capped at -20°C. Standards were prepared for HPLC by solubilization in methanol and filtration as described above for dried sample extracts. The relationship between ergosterol concentration and peak area was linear in the range 3-100 pg ml-’ (r2 > 0.999). This corresponds to a range of 0.124.16 pg ergosterol g soill’ for the extraction described.

191

Reliability of the detection system (HPLC and integration) was assessed by fortifying Marshan and Zurich soil TNE extracts (ca 16 pg ml-’ background ergosterol) with pure ergosterol. Ergosterol recoveries were calculated as the difference between ergosterol concentrations detected in fortified and non-fortified extracts, divided by the amount added. Quantitative detection of 5 and 25pgmll’ was obtained. Coefficients of variation were 0.14 and 0.05, respectively.

Stability of ergosterol to extraction conditions Potential losses of ergosterol during the extraction procedure were assessed as follows: 25 ml nhexane-10% KOH in methanol (1:4) was added to aliquots of pure ergosterol (10 pg) in methanol. Extraction for TAE ergosterol was performed as described for wood chips. To assess hexane extraction efficiency losses, i.e. losses attributable to the failure of ergosterol to partition to hexane from the alcohol-aqueous phase, pure ergosterol was added to solvent mixtures immediately prior to addition of water or saline and removal of hexane. Duplicate determinations were performed with n-hexane-10% KOH in methanol water, (1:4) with added and with n-hexanee methanol (1:4) with added 5.5% NaCl. Samples obtained by consecutive hexane extractions of the same methanol-aqueous phase were processed and analyzed separately. The percentage increase in ergosterol recovery obtained from a second hexane extraction was calculated.

Recovery of ergosterol from hyphal mats Hyphal mats from liquid cultures were collected, lyophilized, and weighed. Lyophilized mats (ca 10 mg) were ground in 0.5 ml methanol in a scintered glass tissue homogenizer (Bellco Glass Inc., Vineland, N.J.). The resulting suspension was transferred to tared 12-ml amber screw-cap vials. The tissue homogenizer was rinsed twice with 0.5 ml of methanol and these rinses were added to the ground mycelial suspensions. Fungal contents of suspension aliquots (mg fungus ml-‘) were estimated gravimetrically, assuming a suspension density of 1.29 ml g-l. Because the density of fungal suspensions was not a constant, suspension fungal contents were corrected following removals of aliquots for extraction. The percentage of the total suspension represented by each aliquot was determined gravimetrically and multiplied by the total dry weight of the mats homogenized. Determinations of TNE ergosterol contents of hyphal mats (ca 3 mg sample-‘) were identical to that described above. For determination of TAE ergosterol concentrations, aliquots of hyphal mat suspensions were refluxed in 5 ml nhexane-10% KOH in methanol (1:4) for 90120 min.

MARKW. DAVISand RICHARDT. LAMAR

192 Recovery

of ergosterol from

cultured soils

Effects of pre-extraction treatments and extraction procedures on yields of ergosterol from colonized soils were assessed in two experiments. In the first experiment, colonized Marshan soil samples were (1) extracted immediately without prior drying, (2) extracted immediately in the presence of a drying agent, MgSO, (>0.963 gml H,O-‘), (3) lyophilized immediately and then extracted, or (4) frozen in an acetone-dry ice bath, lyophilized, and then extracted. Duration of lyophilization was 5 h. Magnesium sulfate was added directly to soil samples in culture tubes, solvents added immediately thereafter, and tubes vigorously shaken. Subsequent steps were as described above for extraction of TAE ergosterol. In the second experiment, colonized Marshan, Zurich, and Batavia soil samples were (1) extracted immediately without prior drying, (2) extracted in the presence of a drying agent, CuSO, (> 1.8 g ml H,O-‘), or (3) frozen in an acetone-dry ice bath and lyophilized for 5 h prior to extraction. Copper sulfate was added to soil and well mixed, and the mixture was then dispensed to culture tubes for extraction. TAE ergosterol concentrations of all treatment groups and TNE ergosterol concentrations of lyophilized samples were determined as described. Recovery chips

of ergosterol from for@ed

soils and wood

Non-inoculated soils used for fortification studies were kept at 80-90°C for several days to reduce their background ergosterol contents. Tests (data not shown) had indicated that soil ergosterol concentrations were decreased to undetectable amounts by oven drying. Non-inoculated chips had no detectable ergosterol. Wood chips were lyophilized and milled as described above prior to fortification. Recoveries of fungal ergosterol from soils and aspen chips were determined as follows. Fungal suspensions used as fortifications were prepared as described above, except that water was used instead of methanol. The rationale of this substitution was to avoid extracting ergosterol from the fungal membrane into methanol before adding the fungal tissue to the solid substrate. Aliquots of the suspension were added to soil or chip samples, and TAE ergosterol contents of fortified and non-fortified samples were determined. The extent of fortification was determined by simultaneously extracting the fungal suspension in the absence of the substrate. Ergosterol recoveries were calculated as the difference between concentrations from fortified and non-fortified substrates, divided by the amount fortified. For determination of recoveries of pure ergosterol, a solution in methanol was used to fortify aspen chips or soils. Ergosterol recoveries were determined as described above, except that ergosterol contents of fortifying solutions were measured by HPLC without prior extraction.

Stability

of stored ergosterol

Effects of short-term (ca 1 week) storage on soil ergosterol concentrations were examined. TAE ergosterol contents of colonized Batavia, Zurich, and Marshan soils were determined immediately after lyophilization. Lyophilized samples from the same soil pools were stored at -2O”C, either under N? or under 20 ml of methanol. Batavia, Zurich, and Marshan soils were stored for 6, 7, and 8 days, respectively. TAE ergosterol concentrations following storage were then determined as described, except that samples stored under methanol were brought to 10% KOH in methanol by adding 2 g KOH. Statistical

analyses

Data were evaluated by analyses of variance (ANOVA). Differences among more than two qualitative treatments were detected using Fisher’s protected least significance (PLSD). Differences among quantitative treatments were detected using regression analysis. In all cases, a probability level of 0.05 was used. For determination of the limit of detection (Vogelgesang, 1987) of TAE ergosterol, a fungal suspension was added to Marshan, Zurich, and Batavia soils (n = 6) to give a soil ergosterol content (90ngg-‘) near the presumed detection limit. TAE ergosterol recoveries were determined as described above. The pooled standard deviation of values obtained by extraction of non-fortified and fortified replicates was multiplied by two times the Student factor (single-sided, 95% confidence level) and divided by the percentage of recovery. In cases where recoveries exceeded unity, they were assumed to be 100%.

RESULTS Stability

of ergosterol

to extraction

conditions

The procedure for extraction of TAE ergosterol was assessed because this method had the most extreme conditions for oxidative loss of ergosterol. Recovery of pure ergosterol subjected to extraction for TAE ergosterol was 95 k 2%. Losses of ergosterol undergoing extraction were subsequently attributed to incomplete partitioning to hexane from the 10% KOH methanol-aqueous phase by use of two hexane extractions. A second hexane extraction was shown to recover 22% of the ergosterol recovered by the first hexane extraction. Thus, the pattern of recovery from four consecutive hexane extractions can be calculated as 78, 17, 4, and 1% of the total ergosterol contained in the 10% KOH methanol-aqueous phase. Partitioning of ergosterol to the hexane phase of soil extracts was more efficient than that of extracts containing only pure ergosterol. Ergosterol recoveries obtained with a second hexane extraction of Marshan and Zurich soil extracts were 15-19% of those recovered with the first extraction. Thus, recoveries from

193

Quantitation of soil ergosterol soil extracts of 97% were obtained by use of two hexane extractions. The partitioning of ergosterol to the hexane phase of Batavia soil extracts was still more efficient, with two hexane extractions recovering 99% of the ergosterol present in the extracts.

Recovery of ergosterol from hyphal mats

Ergosterol recoveries were increased by saponification of neutral extracts (TNE ergosterol), and increased further by saponification in the presence of fungal tissue (TAE ergosterol), as compared to neutral, non-saponified extracts (free ergosterol). The free ergosterol content of homogenized hyphal mats of P. chrysosporium was 3.82 + 0.07 pg gg’. TNE and TAE ergosterol contents were 27 and 34% greater, respectively, than free ergosterol contents of P. chrysosporium hyphal mats (Table 1). Tests to assess the effect of refluxing in the presence or absence (i.e. filtration of alkaline extracts prior to refluxing) of fungal tissue indicated that identical yields of ergosterol were obtained by these two procedures. TAE ergosterol contents of an aqueous suspension of hyphal mats were 4.59 f 0.11 pgg-’ with refluxing and 4.51 f 0.08 pg gg’ without refluxing. Recoveries from non-inoculated Marshan soil fortified with this suspension were 96 + 5% and 97 f 3% with or without refluxing, respectively. Recovery of ergosterol from cultured soils

Soil-catalyzed thermal degradation of ergosterol was implicated in preliminary experiments. Our original procedure for separating TNE extracts from soil residues was to remove a measured aliquot of the solvent mixture prior to adding base and refluxing. This procedure resulted in the presence of soil fines in the extract. TAE extracts were of necessity filtered in these experiments (prior to refluxing) because they exhibited phase separation and a partitioning of soil components to the hexane phase. In this initial comparison, TAE ergosterol recoveries were much greater and subject to less variation among replicate determinations than were TNE recoveries (data not shown). In subsequent experiments in which TNE extracts were also filtered prior to saponification, the differences in ergosterol yields between the two methods became quite small (Table 1). These observations are consistent with the degradation of ergosterol refluxed under alkaline conditions in the presence of soil fines (non-filtered TNE extracts). Tests (data not shown) also showed that quantitative recovery of TAE ergosterol from filtered soil required the use of a hexane rinse. Two consecutive rinses with alkaline methanol recovered significantly less ergosterol than did consecutive rinses with alkaline methanol and hexane. In contrast, recoveries of TNE ergosterol were identical with either rinsing protocol. Losses of ergosterol following lyophilization have been reported (Newell et al., 1988). Thus, treatments to reduce potential losses of TAE ergosterol during

drying of Marshan soil samples colonized with P. chrysosporium were assessed. Of the methods tested, rapidly freezing samples prior to lyophilization resulted in the geatest recoveries (3.62 f 0.09 pg g-i). Lower concentrations were obtained by lyophilizing without freezing the samples (3.34 f 0.09 pg g-‘, P = 0.004) and by immediately extracting hydrated samples without drying them (3.13 + 0.29 pg gg’, P = 0.019). Recoveries were considerably reduced by including a drying agent, MgSO,, with non-dried soil (1.76 f 0.23 pg g-‘, P = 0.0001). Thus, rapidly freezing and lyophilizing soil samples was adopted as our standard treatment before extraction. There were highly significant differences in ergosterol contents among soil types in recoveries of TNE and TAE ergosterol from Marshan, Zurich, and Batavia soils cultured with P. chrysosporium (Table 1). Thus, soils were analyzed separately to assess the effect of extraction method on ergosterol recovery. Yields of TAE ergosterol from all three soil types were slightly, but not significantly, greater than yields of TNE ergosterol. Statistical evaluation using pooled data from the three soil types did not reveal a significant difference between TAE and TNE ergosterol contents (P = 0.4648). This was because of the presence of high quantities of coextractives absorbing at 282 nm in Zurich soil extracts. The resulting chromatographic noise caused relatively poor reproducibility among measurements of Zurich soils (Table 1). When data from Zurich soil samples were omitted from the analysis, yields of TAE ergosterol from colonized Marshan and Batavia soils were significantly greater than those of TNE ergosterol (P = 0.0367). The effects of treatments to reduce potential losses of TAE ergosterol during drying of colonized Marshan, Zurich, and Batavia soils are shown in Table 2. TAE ergosterol recoveries were increased by lyophilization only for the case of Marshan soil. Although the moisture content of the Marshan soil samples was identical, the amount of material Table I. TNE and TAE ergosterol content of hyphal mats of P. chrysosporium and of fungal colonized soils Substrate* Hyphal mats Marshan soil Zurich soil Batavia soil

TNE$ 4.84 3.79 2.80 I.13

Ergosterol contentt TAW

(0.06) (0.10) (0.39) (0.05)

5. IO (0.04) 3.85 (0.05) 2.84 (0.36) I .20 (0.06)

P valuef 0.040 0.239 0.868 0.055

*Soils were cultured with P. chrysosporium as described in Materials and Methods. tValues are expressed in terms of fig ergosterol g substrate-’ for soils, and in terms of mg ergosterol g fungal tissue-’ for mats (dry weight basis). Soil values are means (SD) of six replicate determinations. Hyphal mat values are means (SD) of duplicate determinations. $P equals the probability of a larger F value for differences between TNE and TAE ergosterol concentrations, as determined by ANOVA. @Total neutral extractable ergosterol pool, operationally defined by the extraction methodology employed (see Materials and Methods). TTotal alkaline extractable.

194

MARK W. DAVIS and RICHARDT. LAMAR Table 2. Effectof strategies to reduce potential losses of TAE ergosterol drying of soils cultured with P. chrysosporium Ergosterol Drying

treatment*

Frozen and lyophilized No drying treatment CuSO, added to wet soil

Marshan 3.85 (0.05) a 3.17 (0.24) b 2.90 (0.14) c

content

during

(pg g-‘)t

Zurich 2.84 (0.36) a 2.82 (0.14) u 2.37 (0.15) b

Batavia

I .20 I .28

(0.06) a (0.14) a 0.80 (0.03) b

*Drying treatments are described in Materials and Methods. tValues are means (SD) of six replicate treatments, expressed on a dry weight basis. Means within columns followed by the same letter are not significantly different (Fisher PLSD, CI= 0.05)

extracted differed among replicate samples. Thus, the amount of water with which the extracting solvents were diluted also differed among replicates. There was an inverse relationship (r2 = 0.818, n = 6) between the amount of water contributed by the Marshan soil samples (1.52-l .82 ml) and recovered TAE ergosterol (75-92% of frozen and lyophilized samples). Non-dried Zurich and Batavia samples contributed 0.85-l .08 and 1.35-l .58 ml H20 extraction tube-‘, respectively. Interestingly, no negative effect on ergosterol recovery was observed from Batavia soil samples, whose contributions to the moisture content of the extracting solvents overlapped those of the Marshan samples (r2 = 0.038, n = 4). Use of CuSO, as a drying agent reduced ergosterol recoveries from all soils. Substantial heat was generated during absorption of soil water by

cuso, . Recovery of ergosterol from fortljied soils and wood chips Recoveries of TAE ergosterol from non-inoculated Marshan soil and aspen chips fortified with pure ergosterol or with an aqueous fungal suspension are shown in Table 3. These results indicate that recoveries from substrates fortified with pure ergosterol may not accurately reflect recoveries obtained from fungal tissue associated with the substrates. Recoveries of TNE ergosterol from soil fortified with ergosterol or fungal tissue were statistically indistinguishable from those of TAE ergosterol.

Limits of detection (LDCs) of TAE ergosterol from the three soil types are shown in Table 4. The limit of determination (LDM), defined as the concentration above which quantitative results are achievable at the 95% confidence level, is commonly estimated as 3.33 times the LDC. Thus, for the soils analyzed, TAE ergosterol concentrations exceeding 0.13-0.25 pg gg’ can be quantitated. Given a TAE ergosterol content of 5.1 mg g fungal tissue-’ (Table l), as little as 26-49 pg gg’ of fungal biomass can be quantitated in these soils. In the experiment to estimate LDCs of TAE ergosterol from Marshan, Zurich, and Batavia soils, recoveries of 111, 127, and 66% (SD range 8-20%), respectively, were obtained from soils fortified with fungal tissue. The uncertainty in these figures was a consequence of the experimental design, which involved fortification with trace amounts of ergosterol (see Vogelgesang, 1987). Fortification concentrations (90 ng gg’) were below the method’s LDM. In addition, non-fortified Zurich samples contained traces of ergosterol that were not detected by integration but were apparent upon visual inspection of chromatograms. Recoveries from Zurich samples in excess of 100% were apparently artifacts resulting from summation of these trace amounts with fortified ergosterol. In light of these factors, we considered the data from Marshan and Zurich soils to be consistent with the quantitative recovery previously shown from Marshan soil. However, lower recoveries from fortified Batavia soil were obtained.

Table 3. Recoveries of TAE ergosterol from soil and wood chips fortified with pure ergosterol or with fungal tissue Ergosterol Substrate’ Marshan soil Aspen chips

Ergosterol$ 76 (2) 89 (2)

Table 4. Limit of detectmn of TAE ergosterol from soils fortified with fungal tissue

recovery (%)f Fungal

tissues

94 (5) 97 (2)

Assay detection

P valuef 0.0023 0.0001

*Non-inoculated substrates were fortified as described in Materials and Methods. fValues are expressed in terms of the percentage recovery of ergosterol from the solid substrate, determined as described in Materials and Methods. Soil values are means (SD) of three replicate determinations. Values for chips fortified with pure ergosterol are means of four replicate determinations, and for chips fortified with fungal tissue are means of four replicate determinations at two fortification concentrations. $P equals the probability of a larger F value for differences between recoveries of pure and fungal TAE ergosterol from each substrate, as determined by ANOVA. $Material used to fortify substrates.

Sorl Marshan Zurich Batavia

Ereosterol 0.040 0.070 0.074

limit (pg g ‘)* Biomasst 8 14 I5

*Calculation of limits of detection, defined as the lowest level statistically different from a blank at the 95% confidence level, were performed as described in Materials and Methods. Values are expressed on a dry weight basis. tConversion of soil ergosterol content to soil fungal biomass content is based on a fungal TAE ergosterol content of 5.1 mg g-’ (see Table I, TOW I).

Quantitation of soil ergosterol Table

5. Effect of

storage on

TAE ergosterol funeal colonized soils Ergosterol

content

concentrations

of

(pg g-‘)t

storage method* NOW Methanol Nitrogen

Zurich

Marshan 3.85 (0.05) a 3.89 (0.08) a 3.63 (0.251 b

2.84 (0.36) a 2.59 (0.23) o 2.75 (0.03) a

Batavia 1.20 (0.06) a 1.20 (0.12) a 1.27 (0.02) a

*Soils were stored as indicated for ca I week at -20°C. None indicates no storage, e.g. samples were extracted immediately after lyophilization. TV&es are means (SD) of four to six replicate determinations, expressed on a dry weight basis. Means within columns followed by the same letter are not significantly different (Fisher PLSD, z = 0.05).

Stability of stored ergosterol Pure ergosterol was stable for at least 6 months when stored dry at - 20°C or at room temperature as a solution in methanol (200pgml-‘). Ergosterol in soil extracts solubilized in methanol was also stable, for at least a week, upon storage at room temperature. However, TAE ergosterol contents of suspensions of hyphal mats in methanol were not stable when stored at room temperature. Concentrations were reduced 14% following 5 weeks of storage under this condition (P = 0.0086, n = 2). The effects of short-term (ca 1 week) storage at -20°C on TAE ergosterol contents of cultured Marshan, Zurich and Batavia soils are shown in Table 5. Recoveries of TAE ergosterol from Zurich and Batavia soils stored under N, were identical to those of freshly extracted samples. However, TAE ergosterol concentrations of Marshan samples stored under N, were 9 1% of those obtained from samples extracted immediately. Storage under methanol prevented losses of TAE ergosterol from all three soils.

DISCUSSION

Yields of ergosterol from hyphal mats and from fungal-colonized soils followed the pattern of TAE > TNE > free. In their initial description of a method to extract fungal ergosterol from cereal grains, Seitz et al. (1977) elucidated the importance of a saponification step to liberate otherwise non-extractable ergosterol from its esterified derivatives. They reported that ergosterol yields from fungal-infested grain samples were increased ca 61% by saponification of neutral (i.e. methanol) extracts. Our results indicate that alkaline extraction conditions cause a further increase in recovered ergosterol. This increase likely occurs as a result of liberation via saponification of ergosterol from esterified derivatives whose linkages to membrane components render them insoluble under neutral extraction conditions. Nout et al. (1987) reported that TNE and TAE ergosterol contents were 3 and 15%, respectively, greater than free ergosterol contents of hyphal mats of Rhizopus oligosporus. These findings are in general agreement with our results, although the magnitudes of increases in ergosterol resulting from

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TNE and TAE extraction procedures differ. In contrast, Newell et al. (1988) reported that the TAE ergosterol contents of dead leaves from a salt marsh were 37% lower than their TNE ergosterol contents. However, their TNE procedure employed more vigorous extraction conditions-refluxing for 2 h or homogenizing in neutral solvent-compared to refluxing for 30min in their TAE procedure. Also, they used a much lower base concentration (0.67%) and a larger particle size than was used in our study, which may have resulted in incomplete extraction of TAE ergosterol. Oxidative degradation of ergosterol is catalyzed by heat (Geras’kina and Mukhina, 1980), and might be catlyzed by soil oxidants as well. Because of these concerns, we wished to modify the procedure of Griffiths et al. (1985) for use with soils by removing soils from their alkaline extracts prior to refluxing. Comparisons of ergosterol yields from hyphal mats indicated that identical recoveries were obtained by alkaline extraction regardless of whether or not the fungal tissue per se was subjected to refluxing. Thus, ergosterol measured by alkaline extraction was designated TAE ergosterol. Processing of soil samples before extraction was shown to affect ergosterol yields. The necessity of rapidly freezing soil samples prior to lyophilizing was shown. However, lyophilization was shown to be unnecessary for extraction of Zurich and Batavia soils, from which identical yields of TAE ergosterol were obtained with or without prior lyophilization. Thus, even the presence of substantial moisture (0.85-1.58 ml H,O extraction tube-‘) had no effect on extraction efficiencies from Zurich or Batavia soils. In contrast, less TAE ergosterol was recovered from moist Marshan soil (1.52-1.82 ml H,O extraction tube-‘) than from lyophilized samples, with the degree of inhibition correlating with the amount of water contributed to the extracting solvents. Taken together, these results indicate that (1) the negative effects of added moisture on extraction efficiency varied with soil type, (2) lyophilization without a prior rapid freezing treatment resulted in losses of ergosterol, and (3) use of a rapid freezing treatment prior to lyophilization prevented drying losses, which occurred when freezing was omitted. These findings are consistent with those of Newell et al. (1988), who reported that recoveries of TNE ergosterol from decayed leaves (< 100 mg) placed directly into methanol exceeded those of samples lyophilized prior to extraction. However, the samples in their study were not rapidly frozen prior to lyophilization, and the authors speculated that lyophilization losses may have been prevented by rapidly freezing samples to prevent fungal autolysis during drying. The homogenization step which has been employed by many groups (Matcham et al., 1985; Newell et al., 1987; Nout et al., 1987) for extracting ergosterol from hyphal mats did not increase yields compared to those of intact mats. TAE ergosterol contents of

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5.4 k 0.4 pg g-’ were obtained from intact hyphal mats (as opposed to homogenized mats, see Materials and Methods). This represents a 6% increase in ergosterol compared to that of homogenized samples (compare to Table 1, row l), and it suggests that losses in material may occur during homogenization. The difference between homogenized and intact samples was not statistically significant because of the larger error in the latter values. A likely source of error in the ergosterol content of intact mats was the dry weight determinations of the ca 8-mg mats. Recovery of extracted ergosterol from an alkaline alcohol-aqueous phase by extraction with non-polar solvent is universally employed. Pentane (Miller et al., 1983; Newell et al., 1987, 1988), diethyl ether (Griffiths et al., 1985) petroleum ether (Seitz et al., 1977; Gordon and Webster, 1984; Nout et al., 1987; Johnson and McGill, 1990), and hexane (Grant and West, 1986) have been used in conjunction with diverse alcohol-aqueous phases. In general, three to four solvent extractions were performed. Newell et al. (1988) reported that losses as a result of poor technique are likely at this step. We found that two extactions with hexane sufficed to recover 97-99% of the ergosterol in soil extracts, but only 95% of the pure ergosteroi added to TAE ergosterol solvent blanks and 92-93% of the pure ergosterol added to free ergosterol solvent blanks. A likely explanation involves the polarity of the methanol-aqueous phase. Any component that increases the polarity of the methanol-aqueous phase, such as alkali or soil ions, can increase the partition coefficient of ergsterol in favor of the hexane phase. The data given here for different extraction methods were not corrected for the efficiency of partitioning of ergosterol to hexane from the different methanol-aqueous phases. However, the variation in partitioning efficiency is not sufficient to explain the differences observed in ergosterol yields. TNE and TAE ergosterol extracts consisted of identical solvents at the time of partitioning to hexane. Although the potential for differences in quantities of soil coextractives exists, ergosterol in TNE and TAE soil extracts partitioned equally well to hexane (data not shown). When data for free, TNE, and TAE ergosterol contents of hyphal mats were corrected for partitioning losses, significantly different values (4.15, were still ob5.09, and 5.37 /lg gg’, respectively) tained. In the fortification studies, partitioning losses of ergosterol from control extracts of fungal suspensions would result in overestimates of ergosterol recoveries from soils (and possibly wood chips) fortified with fungal suspensions. Again, when the data are corrected for potential partitioning losses, significant differences remain significant. A second consequence of solvent partitioning characteristics was the loss of material during filtration of TAE extracts when only KOH in methanol rinses of the soil residue were performed. On the basis of our observation that the volume of

the hexane phase in TAE extracts was reduced by filtration, we demonstrated that quantitative recovery of TAE ergosterol required the use of a hexane rinse. Apparently the association of certain soil components, possibly humic materials, with the hexane phase was sufficiently strong to immobilize the solvent with the soil on the filter. Because ergosterol partitions preferentially to hexane, rinses with KOH in methanol were insufficient. Calculations of extraction efficiency require fortification of the substrate with a known amount of ergosterol. However, recoveries from fortified substrates are valid only to the extent that the interactions to which the added ergosterol is exposed model those experienced by the sterol of the colonizing fungal tissue. Clearly, the molecular environment to which pure ergosterol is exposed in soil is very different from that of ergosterol embedded in the matrix of a fungal membrane. If soil interferes with the liberation of ergosterol from the fungal membrane, then recoveries obtained following fortification with pure ergosterol would overestimate extraction efficiency. If ergosterol is lost as a result of direct interactions (in the absence of extracting solvents) between free ergosterol and soil, then recoveries of pure ergosterol would underestimate extraction efficiency. These concerns led us to compare extraction efficiencies from soil fortified with fungal tissue to those from soil fortified with pure ergosterol. We observed that extraction efficiencies of pure ergosterol from Marshan soil and aspen chips failed to model extraction efficiencies of fungal ergosterol. Recoveries of ergosterol from substrates fortified with fungal tissue were quantitative, whereas significant losses of the pure sterol occurred. Losses of pure ergosterol may have been due to adsorption to soil components or to soil-catalyzed oxidative degradation. Fungal ergosterol is only exposed to soil after solvent extraction. Thus, the forces causing losses of pure ergosterol were obviated, but not reversed, by the solvents. Losses of pure ergosterol from Marshan soil were equally irreversible by TAE and TNE extraction procedures. In addition, soil-catalyzed losses of fungal ergosterol occurred only from Batavia soil, a subsoil with a higher clay content than that of the other soils tested (Lamar et al., 1990). These findings are consistent with a mechanism of soil-catalyzed oxidative degradation. However, adsorptive losses may have also occurred. Extraction efficiencies of pure ergosterol have been reported as 93% from cereal grain (Seitz et al., 1977) and as 88-96% from soil-solvent mixtures (Grant and West, 1986). Johnson and McGill (1990) reported 97% recovery of pure ergosterol, although it is unclear whether the ergosterol was spiked into their substrate (roots) or simply subjected to their extraction procedure. These results demonstrate that ergosterol is stable under the extraction conditions used by these groups, and that losses did not occur as a consequence of interactions of extracted ergosterol

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Quantitation of soil ergosterol with the substrates. However, our results demonstrated that recoveries of pure ergosterol may not accurately predict extraction efficiencies from fungalcolonized solid substrates. LDCs reported here (4&74 ng ergosterol g soil-‘, or 8-l 5 pg g-’ of fungal tissue) are consistent with reported detection limits. Although LDCs from wood were not determined, similar precisions within the working range of the assay were obtained from wood and soils (data not shown). Thus, LDCs from wood are probably similar. In their review of techniques to measure fungal biomass, Matcham et al. (1984) reported that detection limits of 25-200 pg gg’ of fungal biomass were possible by use of an ergosterol assay. Grant and West (1986) reported a detection limit of ca 10 ng ergosterol g soil-‘. However, their procedure involved extraction of ca 5-fold more soil than that extracted in our study. In our study, the ergosterol content for P. chrysosporium grown in liquid culture was 5.1 mg gg ’ (mycelium dry wt). This value is consistent with ergosterol contents reported for other filamentous fungi. Seitz et al. (1979) reported an ergosterol content of 3.84.4mgg-’ for Alternaria alternata, 2.3-3.3 mg g-’ for Aspergillus flaws, and 4.9-5.9 mgg-’ for Aspergillus amstelodami grown in liquid culture. Agaricus bisporus grown in liquid culture had an ergosterol content of 2.2 mg g-’ (Matcham et al., 1985). Two strains of Phaeosphaeria typharum reportedly contained 1.9-2.5 mg g-’ ergosterol (Newell et al., 1987). Grant and West (1986) reported an ergosterol content for endogenous soil fungi of 4-7 mgg-‘, based on soil ergosterol contents and hyphal volume measurements. We observed rapid losses of fungal TNE ergosterol with storage of hydrated sterile soils following fumigation with methyl bromide to kill the fungal biomass (> 95% reduction in 2 weeks, data not shown). West et al. (1987) reported severe losses of ergosterol after storage of air-dried soils and more moderate losses after storage of hydrated soils. These data imply that fungal ergosterol is apparently degraded rapidly in soil after fungal death. Thus, ergosterol contents of soils appear to provide a measure of living fungal biomass. Johnson and McGill (1990) reported that ergosterol contents of roots after mycorrhizal infection may also specifically measure living hyphae. In contrast, ergosterol appears to persist in cereal grains after fungal death (Miller et al., 1983; Cahagnier et al., 1983). The use of ergosterol as a quantitative measure of fungal biomass requires that the ergosterol content of the studied species be constant under the culture conditions employed. Nout et al. (1987) reported that the ergosterol content of Rhizopus oligosporus varied (2-24 mg gg’) as a function of developmental, nutritional, and environmental variables. However, others (Matchem et al., 1985; Newell et al., 1987) demonstrated a constant ergosterol content for mycelial mats harvested from log-phase or stationary liquid

cultures. Also in contrast to the report by Nout et al. (1987), we have observed similar ergosterol contents of fungal hyphae from stationary and log-phase cultures, as well as of hyphae from liquid cultures and agar plates (data not shown). We hope that the evaluation of methodology reported here will stimulate further research on this crucial question. The method for determination of TAE ergosterol described here can be used for routine, multiple analyses. In our laboratory, complete analyses of 36 soil samples, from harvest to HPLC, requires ca 16 h of processing time. Analyses of hyphal and wood samples requires ca 12 h. The ergosterol content of samples and of extracts in methanol is not altered by storage for ca 1 week. Thus, the procedure provides a degree of scheduling flexibility. In summary, we found that the TAE procedure for extraction of ergosterol from soils and wood chips provides a high degree of accuracy and precision. The usefulness of a method to accurately estimate ergosterol recoveries from soils and other solid substrates was demonstrated. Fungal ergosterol was quantitatively recovered from Marshan and Zurich soils and from aspen chips. However, the recovery of fungal ergosterol from Batavia soil, a subsoil with a high clay content, was incomplete. Thus, determinations of extraction efficiency using samples fortified with fungal tissue are required to interpret the apparent ergosterol contents of novel substrates.

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