Fungal biomass and productivity

18 Fungal Biomass and Productivity e.

S Y Newell Marine Institute, University of Georgia, Sapelo Island, Georgia, USA

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CONTENTS

Introduction Acetate-to-ergosterol (Ac-~ERG) method Conclusion Future directions

INTRODUCTION The Fungi constitute a Kingdom of evolutionarily closely related microbes that evolved along the same line leading away from the ancestral heterotrophic flagellate as did the animals and choanoflagellates (Kendrick, 1992; H a w k s w o r t h et al., 1995; Alexopoulos et al., 1996; DeLong, 1998). Most species in the Kingdom Fungi are mycelial (but see Chapter 17) eukaryotic organoosmotrophs with non-motile propagules (e.g. ascospores [sexual] and conidia [asexual]) and haploid status for most of the life cycle. The Chytridiomycota, believed to be near the root of the fungal evolutionary tree (Hawksworth et al., 1995), are exceptional in that their propagules are zoospores, and they are not mycelial (though some are rhizomycelial). There is a morphologically strikingly fungal-like group of zoosporic eukaryotic mycelial organoosmotrophs, the Oomycota, which were formerly considered to be primitive fungi, but are now known to be more closely related to diatoms than to fungi (Newell, 1996a; Delong, 1998; Fell and Newell, 1998; Dick et al., 1999), and to be members of the Kingdom Straminipila (Beakes, 1998). Thus, the oomvcotes evolved their mycelial subtrate-pervasion strategy independently of the true fungi. Because fungi digest natural materials such as saltmarsh grass from within the opaque solid substrate (Newell et al., 1996), their masses are best measured via biochemical indices (Newell, 1992). Fungi (including chytrids) and oomycotes are part of the marine microbiota (Fell and Newell, 1998), but biochemical-index methods have been developed for biomass and productivity measurement with natural samples only for METHODS IN MICROBIOLOGY,VOLUME3{) ISBN (1-12 521530-4

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true fungi (Newell, 2000; Gessner and Newell, 2001). Biomasses of zoosporic, non-mycelial true fungi (the chytrids) can be measured using hexosamine techniques, but not with the ergosterol technique that is the focus of this chapter, because chytrids do not synthesize ergosterol (Weete et al., 1989). The oomycotes are also incapable of synthesizing ergosterol, and most do not synthesize chitin, so they are currently beyond the reach of published biochemical-index methods of mycelialmicrobial biomass analysis (Newell, 1994a; Fell and Newell, 1998). Therefore, this chapter, with its sterol-analytical emphasis, is directed at the mycelial true fungi. Mycelial fungi have a negligible presence in the marine plankton (Newell, 1996a) (but see Chapter 17). Fungi have their greatest impact in ecotonal marine ecosystems (marine/terrestrial transitions), where they can target senescing or dead parts of vascular plants for decay (saltmarsh grasses, mangroves) (Newell, 1994a; 1996a; Fell and Newell, 1998). Ascomycetes are the predominant fungi in these types of marine ecosystems (Kohlmeyer and Volkmann-Kohlmeyer, 1991; Fell and Newell, 1998; Kohlmeyer et al., 1999), including those impacted by pollution (Newell and Wall, 1998). Ascomycetous saltmarsh fungi are capable of digesting all parts of saltmarsh grasses, including the lignocellulosic structural framework (Newell at al., 1996). Saltmarsh-fungal production is efficient and can occur at high rates throughout the range of smooth-cordgrass (Spartina alterniflonT) marshes (Newell and Porter, 2000; Newell et al., 2000). There is a clear fungal connection into the saltmarsh foodweb: at least three shredder invertebrates (e.g. saltmarsh periwinkles), which avoid the bitter (ferulic acid) living shoots, are adapted for grazing of the standing, fungal-decayed shoots (Newell and Porter, 2000; Graqa et a[., 2000). It is possible to measure fungal biomass via biochemical proxies in two very different ways: as total mass (living plus dead, evacuated hyphae) or as living mass (membrane-containing mycelium). The hexosamine methods yield total-mass values (since chitin of cell walls is not readily lysed upon hyphal death), and the ergosterol methods yield living-mass values (since ergosterol is a membrane component, and membranes are readily lysed upon hyphal death) (Ekblad et al., 1998; Newell, 2000; Gessner and Newell, 2001). The ergosterol method was first described by agriculturally oriented scientists (Seitz ef al., 1979), but marine ecologists followed closely behind with their own version of the method (Lee el al., 1980), and it was marine microbial ecologists who extended the method beyond biomass measurement to measurement of productivity (Newell and Fallon, 1991). Aquatic microbial ecologists have provided subsequent, invaluable papers on development of the ergosterol-based, fungal-productivity method (Suberkropp and Weyers, 1996; Gessner and Chauvet, 1997). In this chapter the method for finding living-fungal mass and fungal productivity is described; see Newell (2000) and Gessner and Newell (2001) for description and references for hexosamine methods.

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A C E T A T E - T O - E R G O S T E R O L (Ac--~ERG) METHOD

Background and principle "o

Virtually all ascomycetes have ergosterol as the primary membrane sterol (exceptionally brassicasterol may be present in greater quantity than ergosterol), and no plants serving as fungal substrates synthesize ergosterol (Newell, 1992; Gessner and Newell, 2001), so ergosterol is an effective molecular marker for fungal presence in decaying plant material. Ergosterol is also a definitive proxy for fungal mass: only a few microbes other than fungi (green algae and protozoa) have been found to synthesize ergosterol (Newell, 1992; 2000; Gessner and Newell, 2001), and these species are not likely to be firmly bound to decaying leaves or stems of vascular plants in large quantities. The content of ergosterol in ascomycetous mass is rather constant; a recent assay for five species of ascomycetes in two distinct higher taxa (two replicate strains each) and from two different saltmarsh-grass species gave a homogeneous mean ergosterol content of 6.2 l,lg rag' mycelial organic mass (CV = _+8~7,) (Newell, 1996b). This mean value was only 25% different from the suggested general average value of 5 mg ergosterol per g organic fungal mass (Gessner and Newell, 2001; see also Djajakirana et al., 1996). Thus, ergosterol values can be converted into fungal-mass values without the risk of large error due to variation in conversion factors. Ergosterol has basic characteristics that make it a very practicable fungal proxy. It absorbs ultraviolet (UV) light with a peak at 282 nm, a rare feature among sterols (Newell, 1992), reducing the possibility of interference during chromatography. The strong absorbance at 282 nm has permitted development of very sensitive methods for ergosterol assay; e.g. the microwave-extraction method of Young (1995) enables the measurement of living-fungal mass in samples of naturally decaying saltmarsh-grass blades beneath as little as 5 mm" leaf-surface area (1 ng of ergosterol with 50 ~11 injections in HPLC, equivalent to 0.2 1-G of organic fungal mass) (Fell and Newell, 1998). Ergosterol is readily and fully separable from other common sterols (brassicasterol, campesterol, cholesterol, fucosterol, sitosterol, stigmasterol) using liquid chromatography (Newell and Fallon, 199l). Newell and Fallon (1991; Newell, 1993a) expanded the power of the ergosterol method by providing it with the capability for measurement of rates of production of fungal mass. This expansion involved the incubation of decaying-plant samples with radiolabeled acetate (the basic, smallmolecular precursor in ergosterol synthesis: Mercer, 1984; Gessner and Newell, 2001), subsequently separating and measuring the amount of ergosterol in the samples by liquid chromatography, capturing the ergosterol peak during chromatography, measuring the extent to which the ergosterol peak was made radioactive (= the degree of incorporation of acetate) during the incubations, and calculating the rate of production of organic fungal mass (Gessner and Newell, 2001). Acetate was chosen as the tagged precursor because of the possibility that precursors closer to 359

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the final product (e.g. mevalonate) would not be readily taken u p from outside of the cell (see Serrano-Carreon et al., 1993). See Arao (1999) for description of an alternative m e t h o d for monitoring acetate incorporation into fungal lipids.

Detailed methodology Equipment and reagents I include here and in the next subsection the core materials and instructions needed for acetate incubations and ergosterol analyses. See Newell (1993a; 1996a; 2000) for additional details, including methods for obtaining parallel organic densities of natural samples. USE CAUTION: consult local radiation-safety officials for rules for handling radioisotopes and radioactive waste; always work in fume hoods tested for effective function and wear protective clothing w h e n handling radioisotopes and solvents; remember that solvents are highly flammable (especially pentane) - - remove all flame and spark sources. • Carbon-14 acetate (I-['4C]acetate), sodium salt, crystalline solid, specific activity 1-10mCimmol' (37-370 NBq mmol ') (e.g. ICN #12014); store tightly sealed at 0-5°C - - it is hygroscopic and efflorescent and will sublime. The crystalline solid form is recommended here, because the dissolved form is provided in ethanol, which one must remove if one desires to avoid adding ethanol to samples. • Hultiplace filtration manifold for rinsing ['4C]-incubated samples (these can be constructed in-house for much less than scientific-supply prices). • Pure ergosterol for standards; store desiccated under ]'q2 at 0-5°C, or oxidation (can be seen as yellowing) will occur (Newell, 1994b). • HPLC-grade solvents in repipetting containers: methanol, reagent ethanol, pentane; scintillation fluor. • Potassium hydroxide and anhydrous sodium acetate. • Refluxing system (multiple reflux units in an 80°C water bath, plus a source of cooling water with tubing connections to the condensers); grease-free, screw-thread, Teflon connectors (Wheaton Connections®) make this system considerably more user-friendly than condensers with ground-glass connectors. • Dry bath, glass vials to fit bach compartments snugly, and drying-gas distribution manifold. • Sonicating cleaning bath for redissolutions of dried-down neutral lipids. • High-performance liquid chromatographic (HPLC) system (pump, injection valve, column, UV detector [at least 282 and 210 nm detection; 282 for ergosterol, 210 for other sterols], integrator/recorder), with 2-way valve at detector exit (all parts compatible with methanol). • Gas/liquid-tight syringe (5001ul; Teflon plunger) to match injection valve receiver fitting (e.g. luer lock). • Scintillation counter.

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Incubation and assay

1. Place standard samples (e.g. pieces of marshgrass leaf blades of measured surface area, cut from blades, rinsed and stored air-dry [Newell et al., 2000]) in sterile, clean 20 ml, screw-cap vials. Submerge each sample in a measured quantity of bacteria-free seawater (0.2-1amor, ideally, 0.1-1am-filtered; Kirchman and Ducklow, 1993; Blosse et al., 1998; Gasol and Moran, 1999; do not allow filters to run dry: Kiene and Linn, 1999). Cap loosely. Bring two (or more) replicate sample vials to 2% formaldehyde and cap tightly (killed controls). 2. Incubate vials under controlled conditions of temperature and light with slow agitation (e.g. -< 60-75 reciprocations per min through 0.5 cm; violent agitation can have negative effects on fungi: Newell et al., 1996) for 1-3 h to permit adaptation of fungi to submerged conditions (Newell et al., 1985). Slow agitation can be achieved by placing vials loosely in a rack on a rotary shaker run at its slowest maintainable speed, or purchase a slow-speed shaker (see List of suppliers). 3. Add [1-'4C]acetate plus non-radioactive sodium acetate in bacteriafree aqueous solution to bring each incubation vial to 5 mM acetate. Pilot experimentation is needed to find appropriate ratios of radioactive and non-radioactive acetate (Newell, 1993a). Add measured quantity of non-radioactive acetate powder to original vial of radioactive acetate, and add measured volume of bacteria-free water, calculated to be the appropriate amount for micropipetting as aliquots to incubation vials. 5 mM acetate has been repeatedly found to maximize incorporation of acetate into ergosterol (Gessner and Newell, 2001), but ideally, pilot experiments should be run to test whether a particular type of sample conforms to this generalization. Record time of initiation of incubations. Incubate as above for the pre-adaptations for 0.25-4 h (depending on rate of incorporation of acetate, until CPM in ergosterol is clearly measurable, but not long enough to move out of the linearrate period; pilot experiments needed here) (Newell and Fallon, 1991; Gessner and Newell, 2001 ). 4. Terminate acetate incubations by rinsing x2 in bacteria-free seawater (volume = x4 incubation volume) (Suberkropp and Weyers, 1996; fixation and storage at this point can cause loss of label: Newell and Fallon, 1991). Record time of terminations. Immediately place rinsed samples in 5 ml reagent ethanol and store at 4°C in darkness. 5. Extract ergosterol by methanol refluxing and pentane partitioning as described in detail in Newell (2000; Newell, 1993a). This includes: refluxing for 2 h in methanol; addition of KOH/ethanol and 30 min refluxing (lysis of steryl esters); addition of H~O and partioning into pentane (separation from fatty acids and other polar molecules); drying down of pentane in dry bath; redissolution in methanol (cleaning-bath sonication); passage through 0.45 1ira filters into 2 ml, Teflon-capped glass vials. Include procedural standards in the extraction process (add pure ergosterol solutions in methanol to reflux flasks of methanol at a range of final concentrations encompassing those expected from samples, and take them through the whole procedure). 361

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Ideally, one should also add spikes of ergosterol to split natural samples (Newell, 1993a; 2000) to check for sample effects upon ergosterol recovery. See Young (1995) or Eash et al. (1996) for alternative extraction procedures involving microwaving or ultrasonication rather than refluxing. Run samples through HPLC with methanol eluant (Newell, 2000). Inject sufficient sample into injection valve (3 to 4 times injection-loop volume) to ensure that injection loop is sample-rinsed and full of sample (beware of dead volume ahead of injection loop; Newell, 2000; Gessner and Newell, 2001). Inject freshly prepared ergosterol solutions at a concentration in the mid-range of sample concentrations as injection standards (comparison to procedural standards permits calculation of losses of ergosterol during the assay). At 2 ml min ~, 282 nm detection, reverse-phase column (220 x 4.6 mm, 10 1Am), all peaks appear within about 10 min (Newell, 2000). Rinse syringe and injection valve with methanol between samples. During the chromatography of injection standards, record the time of the beginning and ending of the ergosterol peak as shown on the integrator/recorder. During chromatography of samples, at the moment the sample-ergosterol peak appears, open the 2-way valve at the detector exit so that eluant flow is to the open vent (rather than to the vent leading to the waste bottle). Hold a clean scintillation vial under the open vent and collect the sample ergosterol peak. Turn the 2-way valve back to waste just prior to the end of the ergosterol peak. Rinse 2-way valve between samples by allowing clean methanol eluant to flow through the open vent. Collect standard peaks to use to obtain background CPM (scintillation counts per minute) values. Add 10 ml scintillation fluor to vials containing ergosterol peaks. Agitate to mix fluor with methanol, and count vials in a scintillation counter. Subtract counts exhibited by killed controls from live-sample counts to find counts due to fungal incorporation. Find ergosterol concentrations per sample by comparison of sample peak areas with peak areas for procedural standards. Use the CPM for samples, minus tlle background CPM, and the equation of Newell (1993a) to find specific rate (~L, day ~) of ergosterol synthesis, but multiply acetate specific activity (SA) by 0.89 to adjust for losses of carbon atoms when using [l-"C]acetate (Newell, 2000; Gessner and Newell, 2001). Alternatively, use an empirically determined conversion factor to calculate rates of fungal production (Gessner and Newell, 2001; note especially that the conversion factors of Newell (1996b) are high due to a consistent peculiarity of the HPLC injection valve used - - adjust these factors downward by x0.65; the resultant value for the ascomycetes of smooth cordgrass is 12.6 ~g fungal organic mass per nmol acetate incorporated into ergosterol).

Troubleshooting An overriding issue to keep in mind in designing research projects with the goal of studying decay of vascular plants, is that one must avoid

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subjecting the substrate to unnatural conditions that would very likely result in artifactitious experimental outcomes. Severing from the shoot of non-abscised parts, oven-drying, grinding, using green, living shoots or leaves as initial material, litterbagging on sediment surfaces, and permanent submergence of substrates are potentially contraindicated procedures; see Newell (1993b, 1996a), B~rlocher (1997), Gessner et al. (1999). Since ergosterol is a part of the plasma membrane of living hyphal cells, it is susceptible to lysis at cell death. Therefore, sample-preservation methods that slowly kill the fungi within decaying substrates should be avoided. One of these methods is oven-drying (Newell, 2000). For standing-decaying saltmarsh-grass shoots, alternating periods of wetness and dryness are natural, and the fungi of this decay system are not set back by air-drying, so this is an effective method of sample preservation (Newell el al., 2000). This may not be true, however, for samples of persistently submerged leaves such as those of mangroves or riparian leaves in freshwater streams (Gessner and Newell, 2001). gjurman (1994) found that a fungal species that had been grown on wood under damp chamber conditions lost 93% of its ergosterol when humidity was reduced and the wood became desiccated. The safest approach is to move samples directly to methanol or ethanol submergence (Newell, 2000). Losses of ergosterol during extraction and chromatography are characteristically small (< 15~/,; Gessner and Newell, 2001), but there exists the possibility that individual batches could suffer larger losses (accidental temperature shifts, unanticipated interactions among sample chemicals, etc.). Therefore, it is essential that procedural standards be run (see assay step 5), rather than simply standardizing chromatographic runs with a set of injection standards that did not experience the whole extraction~chromatography impact. Although a general conversion factor for calculating fungal organic mass from ergosterol content of decaying substrates has been proposed (200 units organic mass per unit ergosterol; Newell, 2000; Gessner and Newell, 2001), the range of potential conversion factors is large, based on reported ergosterol contents of the fungal species tested (Gessner and Newell, 2001). It is possible that much of this variation is due to genetic change following adaptation to 'luxury' culture conditions during isolate maintenance and transfer. Note that rich culture media can induce low ergosterol contents (Newell, 1992; Bjurman, 1994; Gunnarsson et al., 1996), and consider, for example, the clear cases of constitutive culturinginduced change from the natural state found by Gramss (1991) for wooddecay fungi. When five species of saltmarsh-grass ascomycetes from two distinct major taxa and two decomposition systems were tested immediately upon capture from nature, the five had homogeneous mycelialergosterol content (CV = 8%) only 25~5 different from the proposed general average of 5 mg ergosterol g ' organic fungal mass (Newell, 2000). The empirical conversion factors for calculation of rates of fungal production from rates of incorporation of acetate into ergosterol currently range widely (about 6-19 lJg fungal organic mass per nmol acetate incorporated) (Gessner and Newell, 2001). These values are about 20% to greater than 6-fold the theoretical conversion factor, implying that it is not 363

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possible to force the fungi, in the empirical situations that have been tried, to use offered, radiolabeled precursor exclusively, a situation analogous to that found for bacterial-productivity methods (Jeffrey and Paul, 1988). Only two conversion factors are currently available for marine substrates, 12.6 lJg fungal organic mass per nmol acetate for smooth cordgrass (Spartina alterniflora) and 17.8 lag nmol ~ for black needlerush (]uncus roemerianus) (adjusted from Newell, 1996b; Gessner and Newell, 2001). The needlerush value is very tenuous, since it is based on data for only one species (versus four species for the cordgrass value). Much more work is needed to establish the true range of this conversion factor, and the causes of its variability. See Gessner and Newell (2001) for further discussion of potential problems involved in the acetate-to-ergosterol method.

Application Estimation of fungal-mass content using ergosterol as a proxy has become a standard technique, not only in ecological work but also in agricultural science, plant pathology, industrial applications, and food science (Gessner, 1997; Miller and Young, 1997; Nout et al., 1997; Pereira et al., 1998; Dowell et al., 1999; Joergensen and Scheu, 1999; Kuehn et aI., 1999; Schn6rer et al., 1999; Marstorp et al., 2000; Sridhar and B/irlocher, 2000). In marine science, the ergosterol biomass method has been used extensively in the saltmarsh ecosystem (review: Newell and Porter, 2000; see also Castro and Freitas, 2000), and sparingly in the mangrove ecosystem and on wood decaying in a marine system (Miller et al., 1985; Newell and Fell, 1992). One surprising result of this work is that the saltmarsh and mangrove ecosystems appear to be very different with regard to the major microbial types driving leaf decay: in saltmarshes, ascomycetous fungi accumulate substantial biomass (10-20% or more of decaying-system organic mass) as they decompose standing-decaying leaf blades, but in mangroves, fungi exhibit negligibly small biomasses in submerged decaying mangrove leaves - - oomycotes and bacteria probably do most of this submerged-leaf digestion (Newell, 1996a). Application of the acetate-to-ergosterol method has been much more limited than use of ergosterol as a biomass proxy. Principal ecosystems in which the acetate-to-ergosterol method has been used have been saltmarshes, freshwater marshes, and freshwater streams (Newell et al., 1995; Gessner, 1997; Suberkropp, 1998; Newell and Porter, 2000; Kuehn et al., 2000). For the marine, saltmarsh system, several unexpected results have been obtained. For example: (i) fungal (ascomycetous) productivity per m: of marsh per day in Georgia (USA) marshes has been preliminarily estimated to be about one-half as great as that of total bacteria (bacteria mostly in sediment, to 20 cm depth) per m a marsh in summer, but about xl0 greater than total bacteria in winter (Newell and Porter, 2000; cf. Castro and Freitas, 2000); (ii) rates of fungal production per unit decaying marshgrass blade can be as high (at a standard temperature of 20°C) in northerly marshes (Wells Reserve, Maine, 43°N) as they are in southtemperate marshes (three Florida marshes, 29-30°N) (Newell et al., 2000).

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Many other surprises are likely as the acetate-to-ergosterol method finally enables measurement of fungal productivity in further marine-oriented and other laboratories (Pennanen et al., 1998; Reid et al., 1999). "o ¢.

CONCLUSION The ergosterol biomass-proxy method is a robust one, having been repeatedly challenged and cross-checked with other methods, and having held up well (Newell, 2000; Gessner and Newell, 2001). The acetate-to-ergosterol method of measuring fungal productivity could use more attention from a broader range of scientists to improve circumscription of conversion factors and understanding of other factors affecting the method's accuracy (Newell and Porter, 2000; Gessner and Newell, 2001). There is still a great deal of room for application of the ergosterol-based biomass and productivity methods at the oceans' edges, and in freshwater and terrestrial ecosystems, in order to develop a thorough grasp of patterns of fungal standing crops and rates of fungal throughput.

FUTURE DIRECTIONS Both living and total (empty-hyphal plus cytoplasm-filled hyphal mass) fungal mass could and probably should be measured in parallel (Newell, 1996a) whenever practicable (Ekblad et al., 1998). It may be that these two fungal indices could be measured in one chromatographic run, perhaps by derivatizing either or both glucosamine and ergosterol (Ikemoto et al., 1992), so that they could be measured during one reverse-phase HPLC assay with UV detection (Osswald et al., 1995) (though extractions would still have to be separate and parallel). This combination assay could be made more powerful by the addition of acetate-to-ergosterol application, along with an analogous glucosamine-to-chitin ramification (Roff et al., 1994) or perhaps even an increasingly-fluorescent-chitin method of measuring fungal productivity (Carrano et al., 1997). The cost of [1-1~C]acetate is a hindrance to analyses of large numbers of replicate samples in the acetate-to-ergosterol method (the 1-C-labeled version [ICN/2000 catalog, about $360 mCi '] is 0.4-0.5-fold less costly than the 2-C and 1,2-C versions). It is possible that tritiated acetate could be routinely substituted for [1-'~Clacetate, and the cost ratio, on a mCi basis (1 mCi = 37 MBq), between the two radiochemicals is about 8:1, HC:~H, at the time of writing (ICN catalog for 2000). Newell and Fallon (1991) compared tritiated acetate and [1-"Clacetate in the original acetateto-ergosterol work, and found that pure cultures of two saltmarsh ascomycetes would incorporate measurable quantities of tritium into ergosterol, but the use of tritiated acetate in the acetate-to-ergosterol method has not been further tested. The halophytophthoras are marine-restricted, mycelial eukaryotic organoosmotrophs in the phylum Oomycota, Kingdom Straminipila (see Introduction). They are involved in the decay of leaves that fall into the 365

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marine environment (Newell and Fell, 1997, and references therein), and their zoospores are a part of the coastal-marine plankton (Newell and Fell, 1996). There are no published methods for measurement of biomass or productivity of oomycotes (they do not synthesize ergosterol, and most do not synthesize chitin) (Fell and Newell, 1998). One potential means of measuring h a l o p h y t o p h t h o r a n standing crops might be to use commercially available immunodiagnostic kits designed for testing for the presence of plant-pathogenic species of Phytophthora, which are closely related to the Halophytophthora spp. (Ali-Shtayeh et aI., 1991; Miller et al., 1997; Werres et al., 1997). There m a y well be enough breadth in immunospecificity (Gabler and Richter, 1997; Miller et al., 1997; Wakeham et al., 1997) that the commercial kits could be used in immunoassays for halophytophthoran biomass. Productivity of halophytophthoras might also be measurable analogously to the acetate-to-ergosterol m e t h o d for fungi - the radioprecursors would be to cell-wall glucans not synthesized by fungi (Fell and Newell, 1998) or to the oomycotic phospholipid fatty acid 20:5 (Gessner and Newell, 2001). Gessner and Newell (2001) provide other examples of probable future directions in the evolution of the ergosterol/fungal methods, including the marriage of these non-species-specific methods to DNA-technological or i m m u n o a s s a y methods that are highly species-specific (such as competitive PCR, FISH, and ELISA; Clausen, 1997; Dewey et al., 1997; Nicholson et al., 1998; Kessel et al., 1999; Spear et al., 1999).

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Fell, J. W. and Newell, S. Y. (1998). Biochemical and molecular methods for the study of marine fungi. In: Molecular Approaches to tlre Study of the Oceans (K. E. Cooksey, Ed.), pp. 259-283. Chapman & Hall, London. Gabler, J. and Richter, J. (1997). Cross-reactivity of a polyclonal antiserum against Phytophthora nicotianae. J. Plant Disease Protect. 104, 200-203. Gasol, J. M. and Moran, X. A. G. (1999). Effects of filtration on bacterial activity and picoplankton community structure as assessed by flow cytometry. Aquatic Microbial Ecol. 16, 251-264. Gessner, M. O. (1997). Fungal biomass, production and sporulation associated with particulate organic matter in streams. Limnetica 13, 33-44. Gessner, M. O. and Chauvet, E. (1997). Growth and production of aquatic hyphomycetes in decomposing leaf litter. Limnol. Oceanogr. 42, 496-505. Gessner, M. O. and Newell, S. Y. (2001). Biomass, growth rate, and production of filamentous fungi in plant litter. In: Mamral off Envirormrental Microbiology, 2nd edn (C. J. Hurst, M. McInerne~; L. Stetzenbach, G. Knudsen and M. Walter, Eds), in press. ASM Press, Washington, DC. Gessner, M. O., Chauvet, E. and Dobson, M. (1999). A perspective on leaf litter breakdown in streams. Oikos 85, 377-384. Gra~a, M. A. S., Newell, S. Y. and Kneib, R. T. (2000). Grazing rates of organic matter and living fungal biomass of decaying Spartina alterniflora by three species of saltmarsh invertebrates. Mar. Biol. 136, 281-289. Gramss, G. (1991). 'Definitive senescence' in stock cultures of basidiomycetous wood-decay fungi. J. Basic. Microbiol. 31, 107-112. Gunnarsson, T., Almgren, I., Lyd0n, P., Ekesson, H., Jansson, H.-B., Odham, G. and Gustafsson, M. (1996). The use of ergosterol in the pathogenic fungus Bipolaris sorokiniaHa for resistance rating of barlev cultivars. Eur. J. Plant Pathol. 20, 883-889. Hawksworth, D. L., Kirk, P. M., Sutton, B. C. and Pegler, D. N. (1995). Ainsworth & Bisby's Dictionary of the Frmgi, 8th edn. CAB International, Oxon. lkemoto, N., Lo, L. C. and Nakanishi, K. (1992). Detection of subpicomole levels of

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compounds containing hydroxyl and amino groups with the fluorogenic reagent, 2-naphthoylimidazole. Angew. Chem., Int. Ed. Engl. 31, 890-891. Jeffrey, W. H. and Paul, J. H. (1988). Underestimation of DNA synthesis by l~H]thymidine incorporation in marine bacteria. Appl. Environ. Microbiol. 54, 3165-3168. Joergensen, R. G. and Scheu, S. (1999). Depth gradients of microbial and chemical properties in moder soils under beech and spruce. Pedobiologia 43, 134-144. Kendrick, B. (1992). The Fifth Kingdom, 2nd edn. Focus Information Group, Newburyport, MA. Kessel, G. J. T., de Haas, B. H., Lombaers-van der Plas, C. H., Meijer, E. M. J., Dewey, E M., Goudriaan, J., van der Werf, W. and K6hl, J. (1999). Quantification of mycelium of Botrytis spp. and the antagonist LUocladium atrum in necrotic leaf tissue of cyclamen and lily by fluorescence microscopy and image analysis. Phytopatholo~cy 89, 868-876. Kiene, R. P. and Linn, L. J. (1999). Filter type and sample handling affect determination of organic substrate uptake by bacterioplankton. Aquatic Microbial Ecol. 17, 311-321. Kirchman, D. L. and Ducklow, H. W. (1993). Estimating conversion factors for the thymidine and leucine methods for measuring bacterial production. In: Handbook of Methods in Aquatic Microbial Ecology (P. E Kemp, B. E Sherr, E. B. Sherr and J. J. Cole, Eds), pp. 579-586. Lewis Publishers, Boca Raton, FL. Kohlmeyer, J. and Volkmann-Kohlmeyer, B. (1991). Illustrated key to the filamentous higher marine fungi. Bot. Mar. 34, 1-6l. Kohlmeyer, J., Volkmann-Kohlmeyer, B. and Eriksson, O. E. (1999). Fungi on Juncus tvemerianus 12. Two new species of Mycosphaerella and Paraphaeosphaeria (Ascomycotina). Bot. Mar. 42, 505-511. Kuehn, K. A., Gessner, M. O., Wetzel, R. G. and Suberkropp, K. (1999). Decomposition and CO~ evolution from standing litter of the emergent macrophyte Erh~nthus gi~anteus. Microbial Ecol. 38, 50-57. Kuehn, K. A., Lemke, M. J., Suberkropp, K. and Wetzel, R. G. (2000). Microbial biomass and production associated with decaying leaf litter of the emergent macrophyte ]uncus effusus. Limnol. Oceanogr. 45, 862-870. Lee, C., Howarth, R. W. and Howes, B. L. (1980). Sterols in decomposing Spartina altern(flora and the use of ergosterol in estimating the contribution of fungi to detrital nitrogen. Limnol. Oceanogr. 25, 290-303. Marstorp, H., Guan, X. and Gong, P. (2000). Relationship between dsDNA, chloroform labile C and ergosterol in soils of different organic matter contents and pH. Soil Biol. Biochem. 32, 879-882. Mercer, E. I. (1984). The biosynthesis of ergosterol. Pesticide Sci. 15, 133-155. Miller, J. D. and Young, I. C. (1997). The use of ergosterol to measure exposure to fungal propagules in indoor air. Amer. lndustr. Hyg. J. 58, 39-43. Miller, J. D., Jones, E. B. G., Moharir, Y. E. and Finlay, J. A. (1985). Colonization of wood blocks by marine fungi in Langstone Harbour. Bot. Mar. 28, 251-257. Miller, S. A., Madden, L. V. and Schmitthenner, A. E (1997). Distribution of Phytophthora spp. in field soils determined by immunoassay. Phytopathology 87, 101-107. Newell, S. Y. (1992). Estimating fungal biomass and productivity in decomposing litter. In: The Fungal Community, 2nd edn (G. C. Carroll and D. T. Wicklow, Eds), pp. 521-561. Marcel-Dekker, New York. Newell, S. Y. (1993a). Membrane-containing fungal mass and fungal specific growth rate in natural samples. In: Handbook qf Methods in Aquatic Microbial Ecolo??y(P. E Kemp, B. E Sherr, E. B. Sherr and J. J. Cole, Eds), pp. 579-586. Lewis Publishers, Boca Raton, FL.

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List of suppliers The firms b e l o w are listed b e c a u s e of the a u t h o r ' s familiarity w i t h them, not b e c a u s e they are necessarily the best sources of the m a t e r i a l s that they sell. There is c o n s i d e r a b l e p r o d u c t o v e r l a p a m o n g several of the c o m p a nies b e l o w - - m a n y sell several of the items listed here.

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Alltech Associates 2051 Waukegan Road Deerfield, IL 60015, USA www.alltechweb.com Tel: 847-948-8600 Fax: 847-948-1078

Hamilton Company 4970 Energy Way Reno, N V 89520, USA www.hamiltoncompany.com Tel: 800-648-5950 Fax: 775-856-7259

HPLC gear, including 2-way valves

Gas/liquid-tight syringes ICN Biomedicals 3300 Hyland Avenue Costa Mesa, CA 92626, USA ~ww.icnbionled.co~n Tel: 800-854-0530 Fax: 800-334-6999

Anspec/Ohio 12 W Selby Blvd., Suite 4 Columbus, OH 43085 www.munhall-anspec.com Tel: 800-521-1720 Fax: 614-888-7843

[~4C]acetate

Valves, tubing for HPLC Kimble/Kontes 1022 Spruce Street Vineland, N] 08360, USA www.kimble-kontes.com Tel: 888-546-2351 Fax: 609-692-3242

Beckman Coulter PO Box 3100 Fullerton, CA 92834, USA www.becklIla71co~lter.com Tel: 714-773-6707 Fax: 714-773-8186

HPLC fittings, tubing

Scintillation counters LC.GC Magazine PO Box 6168 Duluth, M N 55806, USA www.lcgcmag.com Teh 888-527-7008

Dionex Corporation 1228 Titan Way Sunnyvale, CA 94086, USA www.dionex.com Tel: 800-723-1161 Fax: 408-739-4398

Contact info. for chromatography suppliers (subscription gratis)

HPLC apparatus N e w Brunswick Scientific PO Box 4005 Edison, NJ 08818, USA

Fisher Scientific 585 Alpha Drive Pittsburgh, PA 15238, USA

WWW.IIbSC.C~}tl

www.fishersci.cotH

Teh 800-631-5417 Fax: 908-287-4222

Teh 800-766-7000 Fax: 800-926-1166

Slow-speed shakers (innova)

Inexpensive HPLC apparatus

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PE Biosystems

Supelco

761 Main Avenue Norwalk, CT 06859, USA www.perkin-elmer.com Teh 203-762-4000 Fax: 203-762-4228

Supelco Park Bellefonte, PA 16823, USA www.sigma-aldrich.com Tel: 800-247-3010 Fax: 800-325-5052

UV detector for HPLC

HPLC columns; other HPLC items

Phenomenex

Wheaton Scientific Products

2320 W. 205th Street Torrance, CA 90501, USA www.phenomenex.com Teh 310-212-0555 Fax: 310-328-7768

1501 N lOth Street Millville, NJ 08332, USA www.wheatonsci.com Teh 800-225-1437 Fax: 609-825-1368

HPLC columns

Grease-free reflux glassware

Sigma-Aldrich PO Box 14508 St. Louis, MO 63178, USA www.sigma-aldrich.com Teh 800-325-3010 Fax: 800-325-5052

Ergosterol, other chemicals

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