Reproductive structures



Most protist species in the soil are inactive at the time of sampling. If the sample has been air dried for storage, species need to be activated first. Therefore, to enumerate and identify species, it is necessary to excyst the individuals. Even before the soil sample is stimulated to excyst, changes in storage conditions (a pre-treatment) will affect species excystment. Different protozoa species have different preferences in pre-treatment requirements for excystment. For instance, some Gymnamoebae cysts require exposure to slightly elevated temperature as a stimulus. Other cysts require a period of desiccation before they can be stimulated to excyst. Thus, pre-treatment storage conditions (especially moisture and temperature) and duration of storage affect the excystment response. Usually soil is moistened, with or without powdered litter amendment, to provide an excystment stimulus. Cysts and spores of various species will respond differently. Some may not be stimulated to initiate excystment under particular conditions. Some species can abort excystment if the conditions are not adequate. Therefore, one protocol is inadequate to excyst all species. Common stimuli to initiate excystment are shifts in temperature (up or down), or changes in moisture, aeration, or nutrient quality or quantity. Variations in temperature, amount of moisture and aeration, or litter additions can help bring out other species, gradually over several weeks. Once excysted, cells remain active only for a certain time. The duration of activity and sequence of species excystment result in a sequence of species activity in the soil samples, over several weeks. Species which are stimulated to excyst do not represent the active community in the soil, because some species can remain inactive or encysted for several years. Therefore, conclusions about an experiment at the time of sampling cannot be obtained from excysted cultures. However, some conclusions pertaining to the soil in general can be derived from analysis of total species composition.

In general, manipulation of excysted cells is carried out by hand with a micropipette or with a bench-top centrifuge. Micropipettes are made from Pasteur pipettes pulled into a thin tube over a small but hot flame. Known volumes of the suspension can be obtained with a pipet-tor. The cone of the pipette can be cut to obtain a wider bore, to prevent clogging by the soil suspension. For collection of live ciliates and testate amoebae on to a membrane, gentle suction can be applied. Soil suspensions can be filtered with nylon mesh or cheese cloth, without suction, to prevent cell lysis. Nucleopore membranes or nitrocellulose membranes are useful because they are transparent under the microscope. Nitrocellulose membranes are resistant to the fixatives used and do not stain. They are made transparent at the end of the staining procedures by dehydration in alcohols and clearing in xylene. Protocols for general histology and cell handling are provided in standard texts in protozoology (Kirby, 1950; Sonneborn, 1970; Lee and Soldo, 1992). Use of ocular grids and micrometre scales in the microscope eye-piece objective is needed for estimating abundances or measuring cell dimensions.

Once soil samples have been prepared to stimulate excystment and organisms extracted, the next step is species identification. Most taxa are large enough to identify under a compound microscope. However, staining of cell structures for ciliate identification or scanning electron microscopy (SEM) for smaller species of protozoa are necessary. Many species of nanoflagellates are unknown or indistinguishable from each other unless thin sections are obtained with transmission electron microscopy (TEM). New species require TEM sections to reveal the ultrastructure of organelles, especially of the basal body and cytoskeleton. As a rule, new species require SEM and TEM for formal descriptions and identification. These are accompanied by descriptions (or video footage) of locomotion, sampling site description, food preferences, growth rates, nuclear and somatic division, and other potentially useful observations. Storing a DNA extract in ethanol is a valuable additional reference.

Ciliates are best obtained from the 'non-flooded Petri plate' technique (see Lee and Soldo, 1992). The soil sample is moistened and, at regular intervals over several days to weeks, more water is added and drained from the soil into a Petri plate. The free gravitational water will carry organisms with it. Many larger flagellated species can also be obtained with this procedure. The extract can be scanned with an appropriate microscope, or fixed and collected on a membrane for staining. The quantitative protargol staining protocol can be modified for other stains or soil suspensions, to obtain quantitative estimates (Montagnes and Lynn, 1987a,b). Species identification of ciliates requires staining of the ciliature and oral structures with protargol (Foissner, 1991). The general antibody against a-tubulin (Amersham, bovine brain a-tubulin) is also useful for immunofluorescence of the basal bodies and cilia. Many of the smaller species can only be identified accurately by SEM.

The amoeboid species can be observed from soil droplets on agar, supplemented with 0.1% yeast and malt extracts or other appropriate organic nutrient solutions. The medium provides a source of nutrients for osmotrophy and for bacterial growth. Different species will excyst over several weeks of observations. A variation based on serial dilutions can be used to obtain estimates of abundances (Anderson and Rogerson, 1995). Individuals are identified based on locomotion, overall shape and dimensions, and details of the pseudopodia. These observations can provide identification of the family or genera. However, confirmation requires TEM, and final species identification will require DNA sequencing of selected genes. Several genera require hyphae, yeast or other protozoa as food sources. Testate species are not easily cultured or activated. On agar plate preparations, excystment and activation occur after >10 days, and some species will only excyst after 3 weeks or more. However, the tests, which are used for species identification, can be found in the litter and soil. It is best to sift through the organic horizon and surface litter under a dissecting microscope. Two useful protocols are recommended for species in the organic horizon. One provides a preparation on a filtration membrane (Couteaux, 1967), and the other provides a fixed preparation on a microscope slide (Korganova and Geltzer, 1977). The preparations can be stained to highlight test features and to differentiate between living individuals and empty tests. The addition of leaf powder or fragmented macrodetritus can stimulate growth of some species on agar plates.

Many species have more specialized food requirements, and several sources must be tried. This approach is called 'baiting' with an appropriate selection of organic nutrients or other food sources supplied to agar plates. Although the technique can be applied to many taxa, it is used primarily in obtaining preparations of Labyrinthulea (including Thraustochytrids), Oomycetes and Hyphochytrea (chromista), Chytridiomycetes (fungi) and specialized species. More details about collection and isolation of these species are provided in Fuller and Jaworski (1987). These protists with filamentous and reticulate growth forms are obtained in liquid or on agar media supplemented with a source of organic nutrient. Common organic matter 'bait' includes pollen, cellulose, chitin, keratin or powdered leaf litter. The soil subsample spread on the plate is incubated for several weeks on various agar-bait combinations and observed regularly. Reticulate and phyllose forms, such as Vampyrellidae and the Eumyxa (protozoa), are also obtained on agar plates. They can be fed with oats, hyphae or macrodetritus.

Most other species, primarily heterotrophic nanoflagellates, can be obtained from soil suspension dilutions. The suspension can be pre-fil-tered through 50-25 ^m mesh and centrifuged to sediment cells. This may be repeated several time to separate cells from soil particles and microdetritus, although with significant loss of cell numbers. The sus pension can be observed under a compound microscope or fixed for SEM for species identifications. These small flagellates are mainly osmotrophic and bacterivorous species. The preparations can be supplemented with bacterized soil extract, 0.1% yeast extract or a similar dilute organic medium such as 1% wheat grass powder infusion.

Many species of protists across phyla are tolerant to aerobic and anaerobic conditions. Other species are adapted for growth in anaerobic conditions and cannot tolerate long exposures to oxygen. These species include yeasts, several species from various phyla of protozoa, such as free-living Archamoebae, and uncharacterized nanoflagellates. They occur in anaerobic soils, submerged soil or deeper in the profile of aerated soil. In order to isolate or observe these species, the soil must be anaerobic for excystment and enumeration. For isolates kept a few days only, an open tube of soil kept saturated with water may be sufficient. For isolates that need to be kept longer, alternative methods exist. The anaerobic environment can be obtained by keeping the soil or the organisms in a sealed container deprived of oxygen. The oxygen can be removed by placing a small lit candle inside a holding jar sealed tight (Fig. 3.3), or with commercially available apparatus.

Lastly, one final group of anaerobic protists needs to be considered in samples. These are endosymbionts of several soil invertebrate taxa. The symbionts live in the intestinal tract of host species, and contribute to the host's digestion and assimilation of nutrient derived from microdetritus. The most common are species of the phyla Trichozoa and Metamonada (both protozoa) that reside in the gut of several woodfeeding termite families, along with specialized bacteria. The protozoan

Fig. 3.3. Anaerobic conditions can be recreated easily in the laboratory. The diagram illustrates soil in an open container or tissue culture flask, inside a sealed canning jar. The candle burns until the oxygen is exhausted.

symbionts ingest the wood microdetritus ingested by the termite host. The cellulose is partly digested and some nutrients assimilated for growth of the symbionts. The remainder is excreted back into the gut, where the termite can continue the digestion and further assimilate the wood. The Termitidae family do not contain protozoa, but have bacterial and fungal symbionts. One subclass of ciliates, the Astomatia with 10 families, is specific to the intestinal tract of annelids, especially oligochaete worms. It is likely that many other species of wood- or detritus-feeding invertebrates, such as some Scarabaeidae, depend on bacteria or protozoa endosymbionts. Endosymbionts are obtained from dissection or squashed preparations of gut contents. However, most species are not tolerant of oxygen and will die soon after exposure, so they must be observed immediately under a coverslip.

Estimates of bacterivorous protists

One popular technique is to assay for total protist species from one functional group only, the bacterivorous species. Generally referred to as the 'most probable number' (MPN) method, it was devised initially to estimate the number of bacteria species that could be cultured under one set of conditions. This approach was abandoned by bacteriologists. Several modifications were proposed to apply the technique to protists (see Coleman et al., 1999). The method relies on serial dilutions of a soil suspension, incubated in a bacterized suspension, and observed at fixed intervals for enumeration. The method does not assay species that do not grow in aerated conditions, in liquid, on the inoculum bacteria or that have been damaged in pre-culture treatments. Furthermore, as explained earlier, excystment conditions vary between species. Soil handling and storage conditions after sample collection affect excystment. Several methodological parameters also affect excystment and growth. These include solution pH (which should be close to that of the collected soil), heat or cold treatments, duration and extent of soil desiccation, and soil dissolved nutrients in the solution. As these conditions vary between laboratories and are not always reported, it is difficult to compare results between laboratories. Moreover, the assumptions of the method are not met when applied to protists. These assumptions are that cells can be evenly distributed in the soil suspension, that all species will grow under the set laboratory conditions on the food provided, and that encysted cells can be differentiated from excysted cells. None are met, and growth of small flagellates can be hidden by growth of predators on them. Among the bacterivorous species, testate amoebae are not cultured, naked amoebae are grossly underestimated, and ciliates are overestimated. Early studies showed that direct counts yielded far more amoebae and nanoflagellate species than culture methods (Sherman, 1914; Martin and Lewin, 1915). The MPN method has poor resolution between collected soils from varied environments, and between trials with the same soil samples, and fails to correlate between variants of the method (Berthold and Palzenberger, 1995). Because the variations in protocols between laboratories are not reported, and because they affect the results, it is impossible to compare results between research groups. This method is not recommended by protozoologists or bacteriologists.

Besides problems in its resolution and repeatability with collected soil, the method is also applied to microcosm and laboratory experiments. This is an erroneous application of the method. The soil for these laboratory treatments is derived from one collected site, and it is then subjected to the same culture conditions for MPN. It is not surprising, therefore, that little statistical difference is obtained between treatments in MPN results from air-dried samples. All the treatments contain the same species diversity, from the same collected soil and, under MPN conditions, the same species are excysted and enumerated each time. The variability is then due to systematic errors, variability inherent to the technique and variations in pre-treatment conditions that affect excyst-ment rates. Attempts at discriminating between encysted and excysted species are also not successful with the MPN approach. For example, HCl-treated subsamples, to kill active species and preserve cysts, were compared with non-treated subsamples. The study showed that almost all cysts were also inactivated by the HCl treatment (Bodenheimer and Reich, 1933). An adequate discussion of problems with this method is provided by Foissner (1987) and L├╝ftenagger et al. (1988).

Fungal hyphae

The mycelium of fungal hyphae must be separated from the soil matrix and collected on a surface prior to examination. It is impossible to release the hyphae intact from the soil, as they are in large part responsible for holding peds and soil-litter aggregates together. Soils rich in hyphae will hold >100 m hyphae/g soil. The soil will also hold yeasts, >1 X 104/g in organic horizons and 600-1800/g in subsurface horizons. The yeast forms are more easily separated with bacteriological protocols. Their species identification is by morphology of growth on agar media (if culturable), with consideration for the range of metabolized substrates and ability for anaerobic fermentation (see Slavikova and Vadkertiova, 2000). However, for stringency and accuracy, molecular markers are essential, such as rDNA sequence, mitochondrial DNA restriction fragment length polymorphism (RFLP) patterns or nuclear magnetic resonance (NMR) spectra of cell wall mannans (Spencer and Spencer, 1997).

Separation of the mycelium from the soil can be achieved rapidly by homogenizing a soil-water suspension briefly (~1 min) in a Waring blender. This will shred the hyphae into short fragments, but it is suitable for many molecular and biochemical protocols. More gentle homogenization of the soil by hand in tissue homogenizers provides less damaged preparations. Alternatively, the soil can be disaggregated with a suspension of suitable density, and hyphae separated from the heavy mineral particles by density and collected on a fine mesh or membrane. This procedure is similar to the invertebrate extraction by density sedimentation/decantation protocols (see below). The latter is possible in soil with dense mycelia.

For estimates of abundance, a known volume of the suspension or weighed cellular material is smeared on a microscope slide for visualization of hyphae and slide preparation. Useful stains include calco-fluor white M2R (a fluorescence brightener which stains fungal cell walls), and fluorescent DNA stains such as 4',6-diamidino-2-phenylin-dole (DAPI) for nuclei. Basidiomycete hyphae can be differentiated from the Ascomycetes with diazonium blue stains after a KOH pre-treatment, which stains cell walls of Basidiomycetes red-purple. The Ascomycete's simple septate pores between cells in the filaments can be detected by trypan blue and congo red in NH4 or sodium dodecylsul-phate (SDS). The Ascomycetes cell wall is bi-layered, with a thin outer and thick inner component. The fluorescent stain 3,3''-dihexyloxacar-bocyanin (DiOC6(3)) is specific for Ascomycetes. Other morphological characters are useful to distinguish between these two classes (Fig. 1.21). Basidiomycetes can be recognized from dolipore septa and clamp connections, with monokaryotic and dikaryotic hyphae, while Ascomycetes have simple septa. However, from most hyphal lengths, it will be impossible to determine any identity. Hyphae inside macrodetri-tus, leaf litter or in mycorrhizal associations require staining of the host material. Common techniques and detailed procedures are described in Johnson et al. (1999) and Norris et al. (1994). Trypan blue is the recommended stain, but acid fuchsin, chlorazol black E, cotton blue in lactic acid and rose Bengal can be useful with some root preparations.

Hyphal preparations can be assayed for biomass from estimates of length/g soil. Common microscope procedures use the line transect or variable area transect method to scan the slide preparation (Paul et al., 1999; Krebs, 2000). The ratio of nuclei/m is a useful index of living hyphae material. However, the presence of nuclei does not indicate that the isolate was active when sampled. Many species are only seasonally active and become transiently inactive when conditions are unsuitable for growth.

A culture-based approach to identification of fungal species involves baiting the soil with appropriate litter or specific plant roots (Johnson et al., 1999). The idea is to stimulate the growth of hyphae from spores, sclerotia or inactive hyphae. Species that are not abundant can be detected under suitable conditions. Relatively intact soil cores are necessary so that the hyphal network is not shredded. The soil is then analysed at regular intervals for the presence of identifiable reproductive structures, spores or mycorrhizae.

In order to identify hyphae, traditionally one relied on descriptions of conidiospores, basidiospores and sporocarps. This procedure requires the visual identification of morphology to a corresponding species (see Johnson et al., 1999). The method does not provide quantitative estimates of species abundance, because some species produce abundant spores, whereas others sporulate infrequently or in small amounts. One study recommends that at least 5 years of repeated observations are required to identify most species in an area (Arnolds, 1992). Therefore, on short sampling periods, even abundant species could be missed if they are not sporulating during that time.

Modern methods based on DNA hybridization with oligonucleotide probes provide an alternative approach. The taxonomic resolution to family or species level is determined by specificity of the oligonucleotide primers chosen. Extracted DNA from soil samples is purified and the DNA can be amplified by PCR techniques. Extraction of DNA requires bead-beating in a lysis buffer. This has been shown to be adequate to break even dehydrated cysts and spores. Lysis buffers such as Trizol allow for sequential extraction of DNA, RNA and proteins from the lysate. Separation of amplified products from nested PCR techniques by DGGE was able to resolve banding patterns between fungal species and can follow species dynamics and succession trends (van Elsas et al., 2000). Detection of known sequences is by Southern hybridization with labelled probes. To quantify known local species of mycorrhizal fungi, RFLP of amplified rDNA has been useful for identification against standards (Kernaghan, 1997; Viaud et al., 2000). The usual chromosomal region for detection of species is the internally transcribed spacer (ITS) region of the nuclear rDNA. This ITS region is flanked by the conserved 18S and 28S genes, which are present in hundreds of copies in each nucleus. It is therefore possible to obtain amplification products from very few nuclei. These PCR-based methods theoretically are more efficient at detecting rare species if the protocol is optimized. However, it is more difficult to quantify species abundances with PCR protocols. Unknown species can be identified from PCR amplification using more general primers (such as one targeting the fungi or the Ascomycetes), separated by DGGE and unknown bands cut out for cloning or direct sequencing. The detection and identification of endomycorrhizal fungi pose a difficulty, in that the hyphae must be identified from within the plant host tissue. A combination of staining, in situ enzyme reaction assay and molecular protocols is recommended, as discussed by Dodd et al. (2000).


Interstitial and litter invertebrates are best separated from the soil by distinguishing between microarthropods and other invertebrates. The microarthropods are obtained from soil cores placed on to modified

Tullgren or Burlese funnel extractors (Farrar and Crossley, 1983; Coleman et al., 1999). The soil core or litter is heated gently with a low-wattage light bulb, and the organisms migrate down, away from the heat and desiccating surface, and fall into a container of 70% alcohol (Fig. 3.4). The efficiency of these extractions is variable, depending on the soil, and some Collembola families (such as the Onychiuridae) are poorly extracted. Specimens may remain adhered to the collecting funnel and not be counted. The extraction is improved using floatation methods, which are more labour intensive (Walter et al., 1987). These require dis-aggregation of the soil in an organic solvent or mineral oil. The cuticle of microarthropods has an affinity for the organic molecules, and individuals float to the surface. Alternatively, density gradient sedimentation with salt or sugar solutions can separate the denser mineral particles from the less dense biota. These methods are discussed in detail elsewhere (Coleman et al., 1999). The most abundant individuals are usually the Oribatida, with up to 40 species in a 20 cm diameter core to 5 cm depth, and 150-200 species represented in temperate forest soil and litter.

In a comparative study of extraction procedures, Snider and Snider (1997) examined samples collected over several years. The micro-arthropods were first Tullgren extracted, followed by three extractions in saturated sugar solution floatation-decanting. Very little organic matter was recovered by the final extraction, on 200 mesh sieve, so that it can be assumed that the invertebrates were also fully removed. The efficiency of the Tullgren heat gradient extraction was compared with total recovery after a further three floatation extractions. The results showed that even though Acarina were in general well extracted (>90%) by the

Fig. 3.4. Extraction of invertebrate species from soil and litter samples. (A) Mist chamber above the sample for gravimetric extraction with water flow. (B) Baermann funnel with a light bulb as heat source. (C) Density sedimentation and separation of organisms from soil in a suitable liquid, followed by decanting. (D) Modified Tullgren and Burlese dry extraction of microarthropods with a light bulb as heat source.

Fig. 3.4. Extraction of invertebrate species from soil and litter samples. (A) Mist chamber above the sample for gravimetric extraction with water flow. (B) Baermann funnel with a light bulb as heat source. (C) Density sedimentation and separation of organisms from soil in a suitable liquid, followed by decanting. (D) Modified Tullgren and Burlese dry extraction of microarthropods with a light bulb as heat source.

heat gradient, it was more variable for some taxa. For example, a Mesostigmata species was only recovered at 14.4 to 42.1% between samples. The results were more variable for Collembola, with the Onychiuridae being poorly extracted by the Tullgren method. In particular, the authors note that greatest variations between heat extraction efficiencies were obtained between samples on the same sampling date. There was also date-specific variability at the same site on the same soil. These observations argue against using one correction factor for all sampling dates, or all soil samples.

Other small invertebrates, namely the tardigrades, nematodes, rotifers, gastrotrichs and enchytraeids, are wet extracted. The soil or litter is placed in water in a funnel, and the organisms are collected into a recipient. The set-up is called a Baermann funnel extraction, and relies on organisms migrating down through the sample and sinking into a recipient. For interstitial nematodes, the Baermann funnel is the easiest method, but it is also the least quantitative (see Grasse, 1965; Nagy, 1996). Alternative and efficient methods are also recommended at the Society of Nematology homepage manual. A variety of centrifugation and elutriation devices have been described that are more quantitative, but require an initial investment in set-up (Freckman and Baldwin, 1990; Coleman et al., 1999). The cen-trifugation method relies on separation of the nematodes from the soil sediment by floatation in a sucrose solution (454 g/l in water), followed by sedimentation of the organisms. It is noteworthy that different families of nematodes are extracted with different efficiency between techniques, so that choice of procedure should reflect the taxa studied (Nagy, 1996). One can expect about 50-100 species represented in a temperate forest, with 30-80 individuals/g of soil.

The wet extraction for the other taxa requires a low-wattage light bulb above the water, to drive individuals away from the heat source. More accurate enumeration requires floatation, differential sedimentation, elutriation or hand sorting, as for the nematodes.

Collection and hand sorting of larger litter invertebrates are described in detail in Sumner (2000). For the macroarthropods, pit-fall traps of varying diameter are used. The traps consist of a collecting funnel or conical-shaped container placed into the soil, with the opening below the litter surface. A trap is placed into the soil below the collector to capture organisms that fall in. The trap consists of a receptacle that can be capped tight, partially filled with a fixative. Both 70% ethanol and polyethyleneglycol (PEG) are commonly used. The latter has the advantage of not evaporating or becoming diluted with dew and rain, but it is very toxic. Salt water is also an effective and non-odorous alternative that is non-toxic. The collected samples are hand sorted and enumerated in the laboratory at a dissection microscope or with an inverted microscope.

Lastly, earthworms are best sampled by hand sorting soil. Samples from pedons (25 X 25 X 25 cm and deeper) are hand sifted with some washing and sieving to remove the mineral particles. One can expect to find 10-100 individuals/m2 in temperate regions, and about six species in a given area of forest or agricultural land. Active individuals can be differentiated from inactive species, cocoons and juveniles. Depth of sampling varies with soil type, and the anecic and deeper burrowing species are always difficult to sample. Absence of a species in samples during certain seasons is probably due to deeper burrowing. A variety of other approaches exist, such as pouring formalin to irritate individuals to the surface, and electro-shock pulses. The efficiency of these methods is usually poor, especially with deeper species, and they cause a sampling bias for certain functional groups and species. Mark recapture has been possible using dyes, fluorescent markers and radioactivity. Species identification can be done visually, but may require dissection, especially in smaller species. Useful keys are provided in Sumner (2000) and references therein.

Active species at time of sampling

The aim is to estimate living and feeding organisms at the time of sampling. This provides a snap-shot of soil activities at a particular moment. Soil species populations are dynamic, and both the active species and the number of individuals of each species are continually changing. The number of individuals of each species will vary over time as cells grow and divide in favourable conditions, or will encyst or disperse in unfavourable conditions. The soil environment itself is a continuously changing environment. It undergoes diurnal warming and cooling, wetting and drying, variable litter input and localized disturbance of microhabitats. Superimposed on these changes, there are seasonal climatic variations. There is a physiological lag between cells detecting a significant change in their immediate environment and responding. There is a further lag in the numerical adjustment of the active population. Thus, species composition and the number of active individuals reflect past conditions, and individuals are adjusting to current soil conditions. Therefore, soil organisms are permanently in a state of adjusting. To obtain an idea of the rate of change and the direction of change, one must sample at regular intervals. The correct interval depends on the resolution required and the nature of the investigation.

For rapid estimation of general cytoplasmic activity by soil organisms, extraction and quantification of ATP can provide a useful index. This assay does not give any indication of the species type or functional groups which are active. However, because ATP is unstable outside cells, with a short half-life, it is a suitable marker for cellular activity. The levels of ATP

in inactive cells, cysts and spores are very low, and both cysts and spores are unlikely to be extracted without bead-beating and lysis buffer. The method needs to be standardized between laboratories, and efficiency of ATP extraction optimized for the soil condition (Martens, 2001).

For ecological studies, one needs to focus on species abundances and estimates of species activity at the time of sampling. Therefore, it is necessary to extract organisms and enumerate the abundance of different species that are interacting. Several precautions are necessary to obtain living specimens and to prevent the inadvertent inactivation or activation of species after sampling soil. Rapid changes in osmotic pressure with desiccation, or in temperature, will lyse cells before they encyst. Gradual desiccation (over several hours) after sampling will affect species composition, as some will no longer be active when the sample is analysed. As protists respond within hours to changes in conditions, to observe species active at the time of sampling, it is necessary to analyse the samples within hours of collection. Some taxa, such as the filamentous hyphae and reticulate species, cannot be isolated in active form. They are either destroyed in collection or are too entangled with the soil matrix and damaged in extraction. These precautions are unnecessary for bacteria, which are too small to be damaged mechanically by sampling, and cannot be lysed by desiccation as they are protected by a cell wall.


When sampling for bacteria, it is often assumed that a composite sample of about 1 g accumulated from multiple milligram quantities of soil adequately represents a sampling area or pedon. Bacteria can be dislodged from particles of a soil suspension in water using a Waring blender, soni-cation, mechanical shaker and mild surfactant solutions, with minimal mechanical damage. However, if applied to eukaryotic cells, these techniques produce lysed homogenates, but cysts and spores will survive. Once the bacteria are dislodged from the soil mineral particles and organic matter, differential sedimentation by centrifugation with or without filtration steps can yield clean separation of the bacterial assemblage. These protocols vary with the soil types and purpose of isolation. The protocol must seek to balance how many bacteria are extracted and how much damage to cells can be tolerated (see Mayr et al., 1999).

Physiologically, bacteria will responds in minutes to new conditions, so that they can be inactivated or activated rapidly. Therefore, standard extraction procedures will tend to inactivate species through dilution of the soil solution, temporary desiccation, disturbance of the soil pore microhabitats and changes in temperature. It is a challenge to determine which species were active before sampling. Probably the least questionable procedure is to use tracers and assay uptake into cells, before the sample is disturbed. Suitable markers with fluorescence, or stable or radioactive isotopes should be added in solution to the soil core and quantified after extraction. The tracer molecule should be able to enter bacteria by active transport reflecting cytoplasmic activity. These could be amino acids, nucleotides or fluorescent molecules. Common fluorescent markers include fluorescein diacetate (FDA) which is activated by a membrane esterase through cleavage of the acetate, releasing the fluorescent molecule in the cytoplasm (Soderstrom, 1979). Another molecule is 5-cyano-2,3 ditolyl tetrazolium chloride which is reduced by electron transfer in metabolism (Rodriguez et al., 1992). Another redox activated dye is tetrazolium chloride (INT) which releases formazan in the cytoplasm. The latter is not fluorescent, but is visible under transmitted light microscopy. One must verify that the species under investigation do take up the markers under controlled active conditions.

Enumeration of bacteria from soil samples by microscopy usually requires staining of nucleic acids and cytoplasmic proteins to distinguish between living cells (with nucleic acids and cytoplasm) and dead cells (cell wall without nucleic acids inside). This provides an estimate of the fraction of cells that are potentially active and inactive, or dead. Suitable stains for nucleic acids include acridine orange, DAPI and ethidium bromide. Common protein stains for the cytoplasm include 5-(4,6-dichloro-triazin-2-yl)aminofluorescein (DTAF) and fluorescein isothiocyanate (FITC). A soil suspension extracted for bacteria is smeared and stained on a microscope slide, or observed in a haemocytometer counting chamber. Appropriate microscope settings include a magnification of 60-100 with phase contrast or fluorescence setting.

An exciting approach to identification of bacteria, without prior culture of the organisms, exploits a combination of these methods. This protocol combines the metabolic uptake of radioactively labelled substrates, together with microautoradiography of smears on slides, and direct microscopy for visualization of fluorescence from rRNA probes against specific genera (Lee et al., 1999). The procedure is amenable to modifications to provide microenvironments suitable for the habitat and organisms studied. Substrate utilization profiles, conditions for activity and the identity and abundance of the bacteria can be quantified. These more focused, experimental and functional approaches are the more informative approach, rather than enumeration by taxonomy or biomass estimations.

Motile protists

The protist taxa are usually separated from the soil matrix based on their locomotion. Although the mode of locomotion is not related to ecologically relevant functional groups, it provides a convenient method of separating the cells from the matrix for identification. The following locomotion categories are proposed: (i) amoeboid with lobose or conose pseudopodia; (ii) amoeboid with filose pseudopodia; (iii) reticulate or phyllose amoeboid species; (iv) ciliated and swimming; (v) with cilia but at least partially amoeboid and surface associated; and (vi) filamentous or hyphal growth. To avoid technical problems associated with sampling fragile cells, soil desiccation and excessive perturbation, the soil portion to be analysed is briefly and gently mixed with a spatula in a watch glass or suitable container. Once evenly distributed, a subsample is removed and weighed for microscopy, and another to obtain the oven-dried weight (Fig. 3.5). Deionized water is added to the subsample for microscopy, to prevent further desiccation, which occurs within minutes if the soil only contains capillary water. Table 3.4 summarizes common methods that have been shown to be suitable to obtain these species.



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