INTRODUCTION

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Although bacteria are the most abundant life forms on earth, knowledge of microbial community structures and population dynamics is still minimal. An estimated 80 to 90% of microorganisms in soil are as yet unidentified (2), and various researchers have detected enormous diversity in such habitats. In particular, Torsvik et al. (37) found evidence for as many as 10⁴ different genomic equivalents in 1 g of forest soil, and in a study of Wisconsin agricultural soil, Borneman et al. (7) found that only 4% of ribosomal DNA (rDNA) clones sequenced were possible duplicates and that several clades of microorganisms had no close relative in the ribosomal database. This limited knowledge of microbial diversity results primarily from our inability to culture and identify the majority of indigenous bacteria. However, an ever-increasing suite of molecular techniques makes it possible to study microbial community structure and compare diversity across habitats (1).

Comparisons of diversity across microbial communities may lead to a knowledge base applicable to a variety of environmental issues. It is necessary to accurately measure changes in populations of microbial community members, especially major components of the community, in response to seasonal, natural, or anthropogenic changes and to identify keystone species (9). Changes in the diversity and structure of a microbial community could become manifested in the ecological processes it mediates. However, difficulties with quantitative investigations of microbial communities lie in the many types of bias which are introduced by culturing or enrichment steps (1, 42), nucleic acid extraction and purification steps (25), and PCR amplification of target genes (1, 17, 28, 36).

It is well recognized that probes targeting 16S rRNAs provide an assessment of microbial community composition. Past studies with such probes have used rRNAs as the hybridization target molecule. It is possible to enumerate cells by using rRNA probes with in situ microscopic techniques or flow cytometry. However, these methods are not currently useful with all types of samples, particularly soils and sediments. Additionally, targeted cells must have high rRNA contents in order to be observed. Studies that employ rRNA probes with nucleic acids extracted from a sample are considered quantitative with respect to the amounts of rRNA measured (31). Hybridizations to rRNA have been used previously to study microbial communities present in anaerobic sewage digesters (31), mixed cultures (29), freshwater sediments (26), biofilms (3), and rumen contents (34). However, since the amount of rRNA per cell may vary according to activity (13, 23), it is difficult to relate the amount of hybridized rRNA to cell numbers. Therefore, a method was sought to estimate cell numbers by hybridization of probes to extracted DNA, thereby providing a different measure of community structure.

Soils high in clay or organic matter, such as marsh sediments, pose particularly tough challenges to obtaining good yields of high-molecular-weight DNA. Compounds present in soils and sediments, particularly humic acids, interfere with molecular reagents. Principally, two approaches are used to recover DNA from environmental samples: (i) concentration of microbial cells from within the environmental sample followed by cell lysis and purification of nucleic acids (19, 20, 21, 33) and (ii) direct lysis of microbial cells within the environmental matrix followed by purification of nucleic acids (6, 8, 30, 38, 40). Separation of cells from soil and sediment samples prior to lysis can be difficult. Differential centrifugation can separate many cells from the surrounding matrix, but many bacteria grow in close association with soil or sediment particles and may be tightly bound to soil colloids (8, 39, 43). Recovery of cells from a sample by this method cannot be expected to be quantitative, representative, or reproducible. In spite of the potential for DNA to adhere to sediment particles, significantly higher yields of DNA are recovered by direct extraction methods than by methods involving cell recovery (6). For these reasons, an approach to estimating DNA from cells lysed within the sample was pursued in the present investigation.

The present study was undertaken to determine the feasibility of using probes directed towards rDNA as a quantitative approach to the estimation of cell numbers when hybridizations with membrane-bound nucleic acids are performed. The approach developed utilizes 16S rDNA probes hybridized to DNA extracted from environmental samples. This method should permit an estimate of "cellular abundance" in a sample, whereas hybridizations to rRNA estimate relative rRNA abundance. A quantitative and reproducible method for isolation and analysis of genomic DNA from marsh sediments high in humic acids was developed and evaluated for efficiency and reproducibility of extraction. This DNA was then used in hybridizations with rDNA-targeted probes to determine the cellular abundance of various groups of sulfate-reducing bacteria (SRB) present in marsh sediments.

	MATERIALS AND METHODS

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Strains used. Desulfococcus multivorans (ATCC 33890), Desulfovibrio vulgaris (ATCC 33405), Desulfovibrio desulfuricans (ATCC 13541), and Desulfobulbus proprionicus (ATCC 33891) were generously provided by Martin Odom of the DuPont Co., Glasgow, Del. Escherichia coli DH5 was obtained from Stratagene. Desulfobacter postgatei (ATCC 33911), Thermus aquaticus (ATCC 31558), Bacillus subtilis (ATCC 27505), and Desulfobacterium autotrophicum (ATCC 43914) were obtained from the American Type Culture Collection.

Study site and sample collection. Marsh sediment samples were taken on 24 June and 30 September 1996 from Canary Creek Marsh in Lewes, Del. This marsh is characterized by a variety of vegetation types and soil characteristics (16). The site is flooded at most high tides. Three 2-g samples, separated horizontally from each other by 2 cm, were taken from each marsh core (14 cm) with a sterile scalpel 3 cm from the top marsh surface and placed in sterile, preweighed 15-ml polypropylene tubes containing 4 ml of phosphate-buffered saline (PBS) (10 mM NaHPO₄ [pH 7.4], 137 mM NaCl, 2 mM KH₂PO₄, 3 mM KCl). These tubes were kept on ice and processed in the laboratory within 3 h. Upon return to the laboratory, samples were adjusted to 2 g of sediment each as necessary by removing portions with a sterile scalpel.

Nucleic acid extraction. Nucleic acid extraction was done by modified versions of the methods of Tsai and Olson (40) and Delgado and Wall (10). Sediment samples were held on ice in 4 ml of PBS in 15-ml tubes. Tubes were shaken at 150 rpm for 15 min at room temperature in a Controlled Environment Incubator Shaker (New Brunswick Scientific) and centrifuged for 10 min at 6,000 × g, and the supernatant was discarded. The process was repeated twice with the addition each time of 4 ml of fresh, cold PBS. Washed sediment was then ground to a fine powder under liquid nitrogen with a porcelain mortar and pestle to assist in the release of cells that were in close association with sediment particles. Each sample was resuspended in 4 ml of lysis solution (0.15 M NaCl, 0.1 M Na₂-EDTA [pH 8.0]) containing 15 mg of lysozyme (Sigma) ml¹ and 15 µg of lysostaphin (Sigma) ml¹ dissolved in TES (20 mM Tris-HCl [pH 7.5], 50 mM NaCl, 10 mM EDTA) and incubated in a 37°C water bath for 2 h with agitation at 20-min intervals. After 1.5 h, 540 µl of 5 M NaCl and 540 µl of a 10% solution of hexadecylmethylammonium bromide (CTAB) in 0.7 M NaCl were added to each tube for the remaining 30 min of incubation. Four milliliters of 0.1 M NaCl-0.5 M Tris-HCl (pH 8.0)-10% sodium dodecyl sulfate (SDS) was then added to each tube, and the samples were incubated at 65°C for 15 min and placed in a 70°C freezer in a dry ice-ethanol bath until further processed. Samples were then cycled four times through freezing at 70°C and thawing at 65°C to lyse the cells and release DNA. After the final thaw, the aqueous phase was extracted twice with 3 ml of 1 M Tris-buffered phenol (pH 8.0) and then twice with 3 ml of chloroform-isoamyl alcohol (24:1 mixture). The phases were separated by centrifugation at 6,000 × g for 10 min. The pellet obtained from the first extraction, which consisted of sediment from the sample, was resuspended in 2 ml of 0.1 M NaCl-0.5 M Tris-HCl (pH 8.0)-10% SDS and extracted once more with 1 ml each of phenol and chloroform-isoamyl alcohol. These extractions were followed by two chloroform-isoamyl alcohol extractions to remove residual phenol and reduce the contaminating iron compounds and humic substances remaining in the sample. Samples were then precipitated overnight at 20°C following the addition of a 10% volume of 3 M NaAc, 5 µl of oyster glycogen (10 mg ml¹), and 2 volumes of 100% ethanol. DNA was pelleted the next morning by centrifugation at 10,000 × g for 15 min, rinsed with 70% ethanol, and resuspended in 130 µl of TE (10 mM Tris, 1 mM EDTA [pH 8.0]). RNA in crude extracts was removed by digestion with 20 µl of 10 mg of RNase A+T₁ ml¹ at 37°C for 0.5 h. RNase was digested by overnight incubation with 40 µl of SDS and 10 µl of proteinase K at 55°C. The DNA preparations at this point were still brownish in color.

Purification of DNA samples. DNA was further purified on MicroSpin Sephacryl S-300 columns (Pharmacia Biotech) according to the manufacturer's instructions with an Eppendorf model 541C variable-speed centrifuge. The 200-µl DNA preparation was loaded onto two MicroSpin columns (100 µl each). Eluents from the two columns were combined, and DNA was stored at 4°C. Samples of purified DNA were analyzed by agarose gel electrophoresis to determine the amount of shearing associated with the purification process.

Oligonucleotide probes. A suite of 16S rDNA oligonucleotide probes was used (Table 1). These probes were previously shown to encompass most members of the gram-negative, mesophilic SRB, and the specificities of these probes were determined previously (14, 15). Probes were labeled at their 5' ends with [-³²P]ATP (3,000 mCi/mmol) (New England Nuclear), as described before (15). Probes were purified on Nick columns (Pharmacia) containing DNA-grade Sephadex G-50, according to the manufacturer. Following purification, a 1-µl aliquot of the preparation was measured in a liquid scintillation counter.

DNA dot blots. Dot blots were prepared with a 96-well dot blot apparatus (Bio-Rad) by use of an Immobilon-N membrane (Millipore). The membrane was prewetted in 95% ethanol for 3 s and rinsed for 2 min in distilled H₂O. The wetted membrane was then placed in 100 ml of 10× SSC (1× SSC is 0.15 M NaCl plus 0.015 M sodium citrate) buffer and allowed to equilibrate for 15 min. The volume of sample loaded onto each well was brought up to 100 µl by addition of 40 µl of 1 M NaOH, 5 µl of 200 mM EDTA (pH 8.2), and double-distilled ddH₂O. For probing with the general eubacterial and general sulfate reducer probes, the signal was expected to be greater and therefore less DNA per sample was loaded onto those membranes. Each DNA sample was boiled in a microcentrifuge tube for 10 min to denature the DNA. Samples were loaded quickly onto the dot blotter and vacuum blotted onto the membrane. Each well was rinsed with 50 µl of 0.4 M NaOH, and the membrane was removed from the blotting apparatus and rinsed for 5 min in 6× SSC, air dried, and then baked for 1 h at 80°C to immobilize the DNA on the membrane.

Hybridization. Membranes were placed in plastic bags and heat sealed approximately 1 cm from the edge of the membrane on all four sides. The prehybridization solution contained 6× SSC, 0.5% SDS, 5× Denhardt's solution (32), and 50 µg of polyadenylic acid ml¹. The hybridization solution was identical to the prehybridization solution and in addition contained 0.01 M EDTA and labeled hybridization probe. Both solutions were vacuum filtered through a 0.2-µm Nalgene filter flask prior to use. Prehybridization, with buffer added to the bag at 100 µl cm of membrane², was carried out for 3 h at 42°C for all probes. The membrane was then gently transferred to a clean Ziploc bag, and hybridization solution was added at 150 µl cm of membrane². Probe was added to the hybridization bag at a concentration of 20 ng of probe ml of hybridization solution¹ and between 5 × 10⁶ and 1 × 10⁷ cpm ml¹ for all experiments. Lower levels produced signal only after prolonged (24 to 120 h) exposure to X-ray film, and higher levels than this produced unacceptable background. Hybridizations proceeded overnight at 42°C. Membranes were washed the next day in 150 ml of a room temperature solution of 6× SSC-1% SDS for 30 min inside a clean plastic container. The wash buffer was changed twice during the 30-min. Membranes were then placed in containers of 1× SSC-0.5% SDS which had been preequilibrated to the optimum final stringency wash temperature for each probe (Table 1). This final wash was performed for 1 h, with two buffer changes during this time. Membranes were blotted gently on a clean paper towel, placed between sheets of plastic wrap, and exposed to Kodak BioMax film with an intensifying screen at 70°C until a clear signal appeared on the X-ray film (between 1 and 16 h).

Image processing. A video image of the autoradiograph was captured with a Gel Doc 1000 Densitometer (Bio-Rad). Signals were quantified by using Molecular Analyst software for Bio-Rad's Image Analysis Systems, version 2.1, on a Macintosh computer, and images were exported to NIH Image and Adobe Photoshop.

Generation of a standard curve to quantify hybridization signals. Amounts of DNA detected in marsh sample extracts were determined by comparison to hybridization signals obtained with DNA standards included on each membrane. The range of standard concentrations used was based on trial runs in which the amount of signal from a sample relative to that from standards of known concentration was compared. E. coli was used as a standard for membranes probed with EUB-338 (bacteria), D. multivorans was used for SRB-814, D. proprionicus was used for SRB-660 (Desulfobulbus spp.), D. postgatei was used for SRB-129 (Desulfobacter spp.), D. autotrophicum was used for SRB-221 (Desulfobacterium spp.), and D. vulgaris was used for SRB-667 (Desulfovibrio spp.).

Conversion of data to cell numbers. Amounts of DNA detected were converted to nanograms of DNA per gram of marsh sediment. An estimate of the average amount of DNA per cell for pure cultures belonging to the groups targeted by the suite of probes used in this study was determined for use as a conversion factor. Four replicate 20-ml cultures of organisms chosen to represent the bacterial types targeted by each probe were grown to mid-log phase. Cells were counted microscopically with a Petroff-Hauser counting chamber, and the results obtained from the four cultures were averaged. DNA preparations were made by the Delgado and Wall protocol (10). The resulting DNA was measured on a Hoeffer Spectrophotometer (260 nm), and the results for the four preparations were averaged. The average DNA yield per milliliter of culture was divided by the average cell count per milliliter to obtain an estimate of DNA per cell. Extraction efficiency was measured by performing the same extraction procedure with three 20-ml control tubes of TE containing 1 µg of E. coli genomic DNA. Estimates of DNA per cell for each representative bacterial type were then adjusted for extraction efficiency. Amounts of DNA obtained per gram of marsh sediment were divided by the adjusted amount of DNA per cell to estimate numbers of cells per gram of marsh sediment.

Reproducibility of DNA preparation and purification. To determine whether equivalent hybridization signals would be produced on X-ray film by replicate environmental samples, three parallel DNA extractions were performed from each of two 2-g sediment samples. Each of the sediment homogenates in PBS was divided into three equal parts by weight. Aliquots (100 µl) of the resulting three DNA preparations were spotted on two different membranes. One membrane was hybridized to the Desulfococcus probe 814 (15) and the other to the general SRB probe 385 (3, 5). The image was imported into Molecular Analyst (Bio-Rad) and quantified by densitometry, as described above. Hybridization signals were compared to signals obtained from a set of DNA standards of known concentration.

	RESULTS

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DNA extraction. The combined supernatants from PBS washes failed to yield DNA that was detectable with a Hoeffer DNA fluorometer or on agarose gels after precipitation with ethanol and concentration (results not shown). Washing sediment samples with PBS therefore caused negligible losses of DNA and/or cells. The average size of the DNA obtained by the extraction procedure was ca. 8.6 kb, as was visualized after agarose gel electrophoresis (Fig. 1).

View larger version (53K): [in this window] [in a new window]   FIG. 1.   DNA preparations examined for shearing. Purified DNA from marsh sediment samples (lanes 1 to 8) is shown; 10 µl of each purified DNA preparation was loaded per lane. Lambda DNA digested with DraI was used as a marker (lane 9). Fragment sizes are given in kilobases.

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View larger version (74K): [in this window] [in a new window]   FIG. 2.   Replicate DNA preparations hybridized with the SRB-385 (general SRB) (A) and SRB-814 (Desulfococcus) (B) probes. From one 2-g sediment sample, three preparations were made and compared to a set of standards of known concentration (data not shown). Images were prepared with NIH Image and Adobe Photoshop 5.0.

Specificity tests and optimization of hybridization conditions. Specificity of the probes had previously been tested only against rRNA (15). It was therefore necessary to evaluate the probes in hybridizations against DNA. DNA purified from pure cultures of target and nontarget species was spotted on a membrane in a range of concentrations and hybridized to a single probe. All probes used in this study, when used at the optimum temperature (Table 1) previously determined with RNAs, yielded strong signals with target DNA and undetectable signals with nontarget DNA. The limit of detection for hybridization to nontarget DNA was <0.1% of the hybridization obtained with the same amount of target DNA. As shown in Fig. 3, the group-specific probes demonstrated the intended specificity when hybridized with DNA. A goal of the optimization study was to maximize the signal detected by X-ray film while preventing nonspecific binding of the probe to nontarget DNA and minimizing the background signal. Parameters experimentally manipulated were temperatures of hybridization and washes, salt concentrations in prehybridization, hybridization and wash solutions, blocking reagents and their concentrations in prehybridization and hybridization solutions, and time durations of washes. The final protocol derived is that described in Materials and Methods.

View larger version (52K): [in this window] [in a new window]   FIG. 3.   Probe specificity tests. Membranes were hybridized with the indicated probe (target group). DNAs (1 µg per spot except in bottom rows of panels B and C, where 500 ng per spot was used) are listed as they appear on membranes from top left to bottom right. (A) EUB-338 (general eubacteria). DNAs: E. coli, D. vulgaris, T. aquaticus, marsh sample, B. subtilis, D. postgatei, D. autotrophicum, D. multivorans, D. desulfuricans. (B) SRB-385 (general SRB). DNAs: E. coli, D. proprionicus, D. desulfuricans, D. postgatei, D. multivorans, B. subtilis, T. aquaticus. (Bottom row shows lower concentrations of the same nucleic acids.) (C) SRB-129 (Desulfobacter spp.). DNAs: D. autotrophicum, D. postgatei, D. vulgaris, D. multivorans, D. proprionicus, E. coli. (Bottom row shows lower concentrations of the same nucleic acids.) (D) SRB-687 (Desulfovibrio spp.). DNAs: D. vulgaris, D. multivorans, D. postgatei, E. coli, D. proprionicus, D. autotrophicum. (E) SRB-221 (Desulfobacterium spp.). DNAs: D. autotrophicum, D. desulfuricans, D. proprionicus, D. multivorans, D. postgatei, E. coli. (F) SRB-660 (Desulfobulbus spp.). DNAs: E. coli, D. multivorans, D. proprionicus, D. vulgaris, D. autotrophicum, D. postgatei, B. subtilis, D. desulfuricans. (G) SRB-814 (Desulfococcus spp.). DNAs: D. vulgaris, D. multivorans, D. postgatei, E. coli, D. proprionicus, D. autotrophicum. Images were prepared with NIH Image and Adobe Photoshop 5.0.

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View larger version (30K): [in this window] [in a new window]   FIG. 4.   Probe sensitivity test. D. autotrophicum DNA was probed with SRB-221. Amounts of genomic DNA spotted (left to right): 100, 80, 40, 20, 10, 5, and 2.5 ng. Image was prepared with NIH Image and Adobe Photoshop 5.0.

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DNA contents. Studies with pure cultures were undertaken to estimate their cellular DNA contents in order to extrapolate amounts of DNA detected in marsh sediments to cell numbers. Results of cellular DNA content determinations are shown in Table 2. The estimated genomic DNA content for E. coli was 6.9 fg. This is 38% higher than the previously reported genome size value of 5.0 fg (17). Estimates of the DNA contents for the SRB examined ranged from 3.1 fg cell¹ (for D. postgatei) to 7.3 fg cell¹ (for D. proprionicus). Devereux et al. (12) estimated the genome sizes of several SRB species by pulsed-field gel electrophoresis. The genome sizes of D. vulgaris and D. proprionicus that they obtained were 3.6 and 3.7 Mb, respectively. Assuming one chromosome per cell, these sizes correspond to 3.9 to 4.1 fg cell¹. The estimates of DNA content per cell determined in this study for D. vulgaris and D. proprionicus were 60 and 94% greater, respectively. The differences in values determined in this study are acceptable considering that actively replicating cells contain greater than one genome equivalent of DNA. Estimation of cell numbers in environmental samples may therefore tend to be conservative.

Estimation of cell numbers. Aliquots of each genomic DNA preparation from marsh sediment samples and DNA from pure cultures, used to generate the standard curve, were spotted on the same membrane when hybridized. The range of DNA standard concentrations (5 to 4,000 ng) and the volume of DNA for all marsh sediment samples on a particular membrane (10 to 30 µl) were selected based upon the expected amount of DNA targeted by the probe used. Reproducibility of the standards was monitored in some cases by including replicate spots of the series. The r² values for regression lines based on signals from the standards were >0.95. The amount of sample DNA loaded onto each membrane was also varied from membrane to membrane to achieve the best signal-to-noise ratio.

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	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

Analysis of microbial community structure is fraught with experimental bias. Although any method is inherently biased, understanding and minimizing the biases will afford the most accurate analysis of sediment microbial communities possible. As with other methods involving hybridization of probes to nucleic acids from environmental samples, probes have been designed based on sequences from cultured organisms, and they will therefore be incapable of detecting an unknown percentage of the natural population. We describe here a quantitative method of microbial community structure analysis which avoids PCR, culturing, and enrichment steps; minimizes some of the sources of bias inherent in soil nucleic acid extractions; and provides reproducible results with sediment samples at a threshold of detection of ca. 8 × 10⁵ cells g of sediment¹.

There are advantages and disadvantages to both rRNA- and rDNA-targeted hybridizations. DNA is more resistant to nuclease attack (enzymatic or divalent metals) and can withstand harsher purification steps. The greatest source of bias in DNA extraction from soil and sediment comes from adhesion and binding of free DNA from cells that have lysed to clay particles. During the DNA preparation protocol, it is difficult to separate this free DNA from the sample and may lead to a lower recovery rate. A portion of the 52% loss of DNA during the purification protocol described above can be attributed to binding of plasmid DNA to clay particles present in the sediment samples. This free DNA can also represent the remnants of dead cells present in the environmental sample. Such DNA is stable and persistent, making it difficult to quantify the contribution to total detected hybridization signal from dead cells. The use of a tritium-labeled plasmid as a standard for DNA recovery from a sediment sample could be improved upon, perhaps with the use of tritiated cells.

RNA is considered a more attractive target than DNA because it is of lesser sequence complexity and is naturally amplified. However, the probability of the random occurrence of a nonspecific target sequence in DNA becomes very small with oligonucleotide probes of the size used in this study. Also, because oligonucleotides are short, mismatches in hybridizations are highly destabilizing so that oligonucleotides have a high degree of specificity (35). However, RNA-based techniques are very useful for evaluating the metabolic status of single cells in environmental samples (27) or of populations, since rRNA content generally increases with growth rate (1). Since the rRNA content of cells is high, rRNA is an excellent target for in situ studies. However, at present in situ observations are generally possible only in relatively clean matrices or with natural samples having highly enriched microbial populations. The high rRNA copy number also enhances the sensitivity of detection with membrane-bound nucleic acids, and because of the smaller size of rRNA, more rigorous nucleic acid extraction techniques can be used, possibly retrieving a larger number of targets (1). However, different bacterial types do not always contain the same amounts of rRNA, and even within one targeted group, more metabolically active cells will have proportionally more RNA, contributing to a stronger signal (11). Additionally, the ribosome content of different species will vary between 10³ and 10⁵ ribosomes per cell (1). For these reasons, it is difficult to estimate cell numbers from hybridizations to rRNA extracted from environmental samples.

Studies with growing cultures have shown E. coli to have an average of 2.1 genome copies cell¹. Similarly, D. vulgaris may contain 4 copies cell¹, and Wall (41) has shown that Desulfovibrio gigas can have between 4 and 17 genome copies cell¹ depending on growth conditions. Bacteria in sediments are not likely to achieve the growth rates attained by those growing in the laboratory. Their genome copy number would be expected to remain low. In a regression analysis of hybridization signals, the amount of DNA detected in an environmental sample should therefore closely correlate with genome equivalents and hence cell numbers used in the array of hybridization standards. Variations in rRNA gene copy number should not introduce significant error, particularly with probes that target a phylogenetically closely related group of organisms. rRNA gene copy numbers should be essentially congruent between detected strains and strains used to generate standards. However, error should be expected when probes that target broader phylogenetic groups are used. Such errors might be greater than severalfold. However, the inferred cell numbers should still be of environmental significance. As previously described, rRNA probes may also underestimate cell numbers by missing an unknown percentage of the target population or by overestimating cell numbers if hybridization to nontarget groups occurs. Evaluation of these errors could be accomplished with the use of nested probes, a suite of probes with varying phylogenetic breadths (1, 13).

The capability of using rDNA probes to estimate actual cell numbers by hybridization to DNA extracted from complex matrices permits initiation of detailed studies on community composition and changes in the community based on cell numbers in formerly intractable environments. In fact, it should not go unnoticed that hybridization of rDNA probes to both DNA and rRNA obtained from a single sample will provide not only quantitative information on community structure but also information about which populations are the most active.