INTRODUCTION

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There is heightened interest in using degradative microbes to bioremediate soil and groundwater contaminated with recalcitrant pollutants, a process known as bioaugmentation. Bioaugmentation has been used successfully to remediate groundwater contaminated with chlorinated solvents (10, 28, 40, 42) and is expected to be useful for other compounds as well. Bioaugmentation requires that effective concentrations of microorganisms be predictably transported to and through contaminated areas of the subsurface. Current tracking technologies are of limited use due to the effects of labeling compounds on cell viability or other properties, high detection limits or interferences from indigenous organisms, or regulatory concerns about the release of genetically modified or antibiotic-resistant microbial strains. Therefore, new methods for tracking viable bacterial cells under both laboratory and field conditions are being developed as part of a major research project examining the physical, chemical, and biological controls on bacterial transport under way under the auspices of the Acceleration Element of the Natural and Accelerated Bioremediation Research Program at the U. S. Department of Energy (DOE) South Oyster site (Oyster, Va.). In order to examine the processes in detail, the movement and distribution of introduced bacteria must be monitored.

Several methods have previously been developed to monitor microbial transport through, and microbial interactions with, porous media during laboratory and field experiments. Selective plate counting has been used with some success to detect injected organisms in downgradient monitoring wells (6, 28, 42), but this method was unsuccessful during a field-scale bacterial transport experiment at the DOE North Oyster, Va., site, where an indigenous strain was injected (7). Plating methods can have detection limits around 100 to 1,000 culturable cells per ml of sample, providing that there is no overgrowth of indigenous bacteria on the plates. Selective plating may allow lower detection limits but requires extensive a priori screening of every potential degradative organism for antibiotic sensitivity and carbon source utilization profile. A major disadvantage of this method is that it does not detect "viable but nonculturable" cells (21, 30, 31, 34, 36, 38, 50), which still may be metabolically active but unable to form colonies on solid media.

Stable isotopes are increasingly being used for field experiments. Bacteria grown on ¹³C-enriched glucose prior to injection into the surficial, uncontaminated aquifer at the North Oyster, Va., site were detected in downgradient monitoring wells by converting collected particulate organic matter to carbon dioxide and measuring the ¹³CO₂ by Carlo-Erba isotope ratio mass spectrometry (7). However, additional methods were required to unequivocally establish that the ¹³C represented intact cells of the bacteria injected rather than protozoans or other indigenous microbes which may have incorporated labeled cellular material via predation of live target cells or consumption of dead target cells (W. Holben, University of Montana, personal communication, 1999).

One of the molecular methods available, PCR, has been used to count very low numbers of bacteria in a variety of environments (25, 44, 45, 49, 51, 53). PCR and nucleic acid quantification can usually be performed only in the laboratory, which precludes the use of these methods in the field to obtain near-real-time cell detection and enumeration data. Genetically engineered microorganisms carrying the genes coding for the green fluorescent protein have been developed (11, 12, 29), which has allowed enumeration of these organisms by measuring fluorescence. However, the use of these genetically engineered microorganisms has been restricted to laboratory, greenhouse, and lysimeter experiments because of regulatory concerns.

Labeling cells with fluorescent stains has been employed to examine bacterial attachment to surfaces (13), to count the numbers of total and active cells in a variety of environmental samples (37, 39, 55), and for in situ injection of groundwater bacteria (1, 16-19). One of the stains used, 4',6-diamino-2-phenylindole (DAPI), specifically binds to nucleic acids (RNA and DNA), which enables it to universally label cells in an organism-independent manner. The binding mechanism adversely affects normal cell function (33), resulting in a loss of viability. The potential effects of this compound on the transport properties of an organism have been shown to be minimal for short-term experiments (24). Another stain, 4-phenyl spiro-furan-2-(3H),1 phthalan-3,3'-dione (Fluram, fluorescamine), forms covalent bonds with free amino groups. Fluram was shown not to effect the adhesion of a Rhodococcus strain to titanium-rich particles (13), but its effects on other species have not been examined. For fluorescent stains to be useful for studying bacterial transport and monitoring bioaugmentation, they must have minimal effects on bacterial adhesion, viability, and metabolic activity, while at the same time they must be retained in the cells for at least several weeks.

New fluorescent compounds which may allow cells to be stained without a loss of activity or viability or changes in adhesive properties have been and continue to be developed. Many of these dyes have been developed specifically for eukaryotic cell staining, but the principles underlying their use makes them applicable to prokaryote staining as well. Some of the newer dyes specifically stain cell membranes, while others cross the membrane and covalently bond to intracellular proteins. In either case, some of these dyes have been shown to be retained in cells for at least 3 to 4 weeks without a loss of cell viability or alterations in cell function or adhesion (20). This work was undertaken to critically evaluate the labeling of bacteria with numerous fluorescent stains and to develop an easy and effective high-throughput detection procedure for quantifying fluorescently labeled cells during in situ bacterial transport experiments.

	MATERIALS AND METHODS

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Chemicals and media. The following compounds were purchased from Molecular Probes (Eugene, Oreg.): calcein AM; calcein blue AM; 5-(and 6-)-carboxyfluorescein diacetate, succinimidyl ester (CFDA/SE); 7-amino-4-chloromethylcoumarin (CellTracker Blue CMAC); 5-chloromethylfluorescein diacetate (CellTracker Green) (CMFDA); 4-(4-[dihexadecylamino]styryl)-N-methylpyridinium iodide (DiA); 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI); 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine-5,5'-disulfonic acid [DiIC₁₈(3)-DS]; 1,6-diphenyl-1,3,5-hexatriene (DPH); Oregon Green 488 (carboxy-DFFDA); 1,1'-dioctadecyl-6,6'-diphenyl-3,3,3',3'-tetramethylindocarbocyanine chloride (6,6-Ph2-DiIC₁₈); rhodamine B (hexyl ester, chloride); 5-sulfofluorescein diacetate, sodium salt (SFDA); SlowFade Light mounting fluid; 5-(and-6)-chloromethyl SNARF-1 acetate; (1,1'-dioctadecyl - 6,6' - di(4 - sulfophenyl) - 3,3,3',3' - tetramethyl - indocarbocyanine [SP -DiIC₁₈(3)]; 3,3'-dioctadecyl-5,5'-di(4-sulfophenyl)-oxacarbocyanine, sodium salt [SP-DiOC₁₈(3)]; and 5-(and-6)-carboxytetramethylrhodamine, succinimidyl ester (TAMRA/SE). Fluorescamine and 5-cyano-2,3-ditolyl-tetrazolium chloride (CTC) were purchased from Fluka Chemical (Milwaukee, Wis.) and PolySciences (Warrington, Pa.). DAPI and acridine orange (AO) were obtained from Sigma Chemical (Milwaukee, Wis.). R2A agar was purchased from Fisher Scientific (Fair Lawn, N.J.).

Bacterial strains. An indigenous aquifer organism, DA001, was isolated from groundwater from the South Oyster Narrow Channel Focus Area. DA001 was identified as a member of a Comamonas species by phylogenetic analysis of its 16S rRNA gene sequence (similarity index [SI] = 0.965). This strain has been used extensively for laboratory experiments performed as part of the DOE bacterial transport research project and will be used for field-scale experiments as well. Therefore, the majority of the fluorescent stain screening was focused on finding compounds which would facilitate tracking this organism. Adhesion-deficient variants of DA001 were selected by using a standardized adhesion assay (9) employing South Oyster site sediment. DA001 was used for screening the fluorescent stains when the adhesion values stabilized around 30 to 40%. Routine growth of the organism was performed by using 0.2% (wt/vol) sodium acetate in a basal salts medium (16) at room temperature.

Fluorescent stain screening. Previously published procedures were followed for staining with SFDA (46), fluorescamine (13), and CTC (37, 39). Initial staining with other compounds was performed by using the manufacturers' recommended protocols for carrier solvents, cell suspension buffers, final compound concentrations, and temperature conditions. The compounds were usually added as concentrated stocks in carrier solvents consisting of dimethyl sulfoxide (DMSO) or dimethyl formamide to achieve final concentrations of 10 to 50 µM, with the carrier solvent present at a concentration of 0.2% (vol/vol).

Optimization of CFDA/SE staining protocol and sample handling. The general staining procedure proved to be adequate for the initial screening. However, there was a desire to minimize the amount of CFDA/SE needed to stain a given number of cells and to optimize the overall staining procedure in order to both reduce the costs associated with labeling cells for field-scale experiments and facilitate scaling-up of the staining procedure. The experiments performed included staining in different buffer solutions (NCAGW, PBS, PB), staining of stationary cultures versus staining of log-phase cultures, staining with different temperature regimens (ambient, 37°C, cycling from 25 to 37°C), and staining with different cell densities (10⁹ and 10¹⁰ cells/ml) and with different CFDA/SE concentrations (10, 50, and 100 µM). In general, staining took approximately 3 to 5 h. Specific details are given in the Results and Discussion.

Microcosm fitness experiments. Microcosm experiments were performed to assess the potential effects that CFDA/SE may have on the fitness or survival of the stained organism. A single culture of DA001 was grown and split into two portions. One portion was stained as described above, while the other underwent the same incubation procedure in the absence of CFDA/SE. After the cells were washed and resuspended in mNCAGW, they were incubated at 15°C for 72 h. Both cell solutions were washed and resuspended in fresh mNCAGW prior to addition to aqueous microcosms (50 ml of mNCAGW or Narrow Channel Focus Area [NC] groundwater in 250-ml bottles) and saturated NC sediment microcosms (2 g of wet sediment in 50-ml centrifuge tubes; total liquid volume, 2 ml). Unstained and CFDA/SE stained DA001 cells were added to obtain initial densities of 10⁷ cells per ml or g. The aqueous microcosms were incubated at 15°C with the bottles gently shaken to keep the cells in suspension; the sediment microcosms were incubated statically. Aliquots of the aqueous microcosms were removed over a 3-week period for determination of culturable DA001 concentrations by plating onto R2A agar; CFDA/SE-stained cells were counted by direct microscopic enumeration. Replicates of the sediment microcosms were sacrificed at the same time points and extracted with 18 ml of mNCAGW, and the DA001 concentrations were determined by plating and microscopic examination (CFDA/SE-stained cells only). The indigenous population of DA001 and other microorganisms in the South Oyster sediment is quite small, which results in a low background concentration and allows added DA001 to be enumerated by plating.

Detection methods for CFDA/SE-stained cells. During this work, alternatives to direct microscopic enumeration of CFDA/SE-stained cells were investigated, particularly alternatives which would facilitate high-throughput, low-cost sample analysis. Discrete sample fluorometry was evaluated by using a model 10-AU-005 field fluorometer rented from Turner Designs (Sunnyvale, Calif.). Ninety-six-well black microplate formats were tested by using a model BF10000 FLUOROCOUNT fluorescence microplate reader (Packard Instrument Company, Meriden, Conn.) and a SPECTRAmax GEMINI dual-scanning microplate spectrofluorometer (Molecular Devices Corporation, Sunnyvale, Calif.). Both of these devices measured the total fluorescence emitted by each well of the microplate and reported the fluorescence intensity in relative fluorescence units. Ninety-six-well black OptiPlate HTRF-96 microtiter plates (P/N 6005407) were purchased from Packard. A flowthrough fluorescence quantification system consisting of a Hitachi A11 high-performance liquid chromatography (HPLC) pump and an AL10 108 position autosampler coupled to an FL-750 HPLC spectrofluorometer (McPherson, Inc., Chelmsford, Mass.) equipped with 6-, 12-, and 24-µl flow cells was also evaluated. NCAGW was the mobile phase, and no column was present between the autosampler and the detector.

Intact-core bacterial transport experiments. Intact sediment cores (length, 70 cm; diameter, 7.2 cm) were collected from both the vadose and saturated zones during an excavation at the South Oyster field site in August 1998. The core orientation was parallel to the in situ groundwater flow. Further details regarding the site and core collection can be found elsewhere (7, 14). Intact cores were prepared and operated as described elsewhere (8, 14), with slight modifications to tubing size and configuration as described by DeFlaun et al. (M. DeFlaun, M. Fuller, B. Mailloux, T. Onstott, W. Holben, W. Johnson, P. Zhang, D. Balkwill, and D. White, submitted for publication). The final height of the sediment after core preparation was 50 cm. Initial injections of sodium bromide (50 µg/ml as Br) or [³H]water (approximately 10,000 dpm/ml) were performed as described previously (8, 14). These injections were used to calculate the flow velocity and core porosity, permeability, and dispersivity and served as conservative tracers to which the transport of the bacterial cells was compared.

	RESULTS AND DISCUSSION

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Fluorescent stain screening. A total of 19 fluorescent stains were evaluated; the results of the screening are presented in Table 1. Only seven (37%) of the stains exhibited a high-efficiency staining of Comamonas sp. strain DA001. This indicates that stains developed primarily for use with mammalian cells do not necessarily work with bacterial cells. However, it was encouraging that only one of the stains, fluorescamine, caused significant reductions in cell culturability and significant increases in cell adhesion to sediment. This effect was most likely due to either the acetone carrier solvent or the fact that the fluorescamine reacted with free amine groups at the cell surface.

Microcosm fitness experiments. The survival and culturability of unstained and CFDA/SE-stained DA001 cells were virtually identical in mNCAGW, NC groundwater, and NC sediment microcosms (Fig. 1). Survival was substantial in the NC sediment, possibly due to the protection from predation that the sediment afforded compared to NC groundwater. Cells remained fluorescent in all matrices for up to 35 days. The culturable cell densities in the different microcosms were mirrored by the concentrations of CFDA/SE-stained cells over time (Fig. 1), indicating that fluorescence is likely lost shortly after cells become nonviable. Given this, and the fact that neither the NC groundwater microcosms nor the NC sediment microcosms exhibited a detectable increase in culturable cells, it is reasonable to assume that the introduced DA001 cells remained in a nongrowth state throughout the incubation and did not utilize any labile carbon present. If growth did occur, the number of culturable cells would be expected to increase, while the number of detectable CFDA-stained cells would be expected to decrease due to dilution of the intracellular concentration of the fluor with each cell division. An exciting result was the ability to enumerate CFDA/SE-stained cells in the sediment microcosms by epifluorescence microscopy, even in the presence of brightly autofluorescing mineral grains (Fig. 2).

View larger version (18K): [in this window] [in a new window]   FIG. 1.   Comparison of survival of unstained and CFDA/SE-stained DA001 cells as determined by plate counting of unstained () and CFDA/SE-stained () cells and by epifluorescent direct counting () in NCAGW microcosms (A), NC groundwater microcosms (B), and NC sediment microcosms (C). The error bars represent 1 standard deviation of the mean.

View larger version (57K): [in this window] [in a new window]   FIG. 2.   Micrograph of CFDA/SE-stained DA001 cells in the presence of autofluorescing NC sediment grains.

Optimization of CFDA/SE staining and sample processing. Initial optimization was performed by using only epifluorescence microscopy to qualitatively assess the effects of different protocols on the efficiency and brightness of CFDA/SE staining. Staining was performed by using 1.0-ml (total volume) samples containing 1 × 10⁹ DA001 cells/ml in 2.0-ml amber microcentrifuge tubes shaken vigorously (300 to 600 rpm), with a final CFDA/SE concentration of 100 µM (delivered in 2 µl of DMSO). Staining was most effective when the cells were suspended in PBS instead of 5 or 50 mM PB or mNCAGW. Staining in basal salts medium during growth on acetate (same cell and CFDA concentrations) yielded uniformly faintly fluorescent cells. Incubation at 35 to 41°C yielded better staining than incubation at room temperature. Using a CFDA/SE solution which had been repeatedly frozen and thawed instead of a freshly prepared solution did not affect the quality of staining in any medium. During microscopic examination, cells stained in PBS appeared to contain very bright yellow-green intracellular inclusions; the fluorescence usually dispersed throughout the entire cell after incubation in NCAGW at 15°C for a few days. The use of SlowFade Light mounting fluid proved to be very effective for preventing fading of cell fluorescence during observation and counting.

View larger version (15K): [in this window] [in a new window]   FIG. 3.   Fluorescence intensities of suspensions of CFDA/SE-stained DA001 cells (approximately 5 × 106 cells/ml) buffered to different pH values, as measured with the SPECTRAmax GEMINI scanning microplate spectrofluorometer. The error bars represent 1 standard deviation of the mean based on eight analysis replicates.

Stained-cell detection methods. The Turner Designs fluorometer was able to detect CFDA/SE-stained cells, and the lower detection limit was approximately 1 × 10⁴ cells/ml when the instrument's 25-ml discrete sample chamber was used. However, the instrument was somewhat awkward to set up and operate, and the data capture capabilities were somewhat limited. The use of an HPLC fluorescence detector coupled with small-volume flow cells did not yield consistent results, and several instrument problems precluded further development.

View larger version (29K): [in this window] [in a new window]   FIG. 4.   (A) Expected and observed CFDA/SE-stained DA001 cell concentrations in NC groundwater (pH 6.0) () and NC groundwater preserved with 1% (wt/vol) formaldehyde and buffered to pH 8 (), as determined by epifluorescence microscopy and flow cytometry. The x-axis error bars represent the calculated error based on three replicate direct microscopic counts of the initial cell solution used to prepare each dilution series. The y-axis error bars represent the error associated with the mean from three replicate flow cytometry analyses after the mean from three replicate flow cytometry analyses of the blank had been subtracted to account for background fluorescence. (B and C) Flow cytometric dot plot (B) and histogram (C) of CFDA/SE-stained DA001 cells in formaldehyde-fixed NC groundwater. FL1 is the optical filter in the flow cytometer which allows the sample fluorescence emitted at 530 ± 30 nm to be quantified. TFS, TransFluorSpheres.

Intact-core experiments. Transport of DA001 cells through intact sediment cores was similar regardless of the tracking method used (Fig. 5A and 6). Plate counting of DA001 CFU and liquid scintillation analysis of ¹⁴C-labeled DA001 cells yielded the same results as direct microscopic counting and microplate spectrofluorometry of CFDA/SE-stained cells. During the later time points of experiment 2, the CFDA/SE direct counts matched the CFU very closely, while cell concentrations based on microplate spectrofluorometry and liquid scintillation paralleled each other. In both experiments, cells passed through the cores significantly faster than the conservative tracers passed through, and the peak concentration was observed to elute from the cores sooner for the cells than for the tracers. This is seen more clearly in Fig. 6 than in Fig. 5A because of the multiple peaks in the core NC7-2 cell breakthrough curve.

View larger version (26K): [in this window] [in a new window]   FIG. 5.   (A) Concentrations of CFDA/SE-stained DA001 cells in the effluent of intact sediment core NC7-2, as determined by plate counting (), epifluorescent direct counting (), and microplate spectrofluorometry (). Breakthrough of the conservative bromide tracer is also shown (---). The initial concentrations of DA001 and Br injected into the core were 7 × 107 cells/ml and 50 µg/ml, respectively. The error bars represent 1 standard deviation of the mean. (B) Sediment concentration profile for CFDA/SE-stained DA001 cells along the length of intact core NC7-2, as determined by plate counting () and epifluorescent direct counting ().

View larger version (23K): [in this window] [in a new window]   FIG. 6.   Concentrations of 14C-labeled DA001 cells stained with CFDA/SE in the effluent of intact sediment core NC9-2SAT, as determined by liquid scintillation counting (), plate counting (), epifluorescent direct counting (), and microplate spectrofluorometry (). Breakthrough of the conservative 3H2O tracer is also shown (---). The initial concentration of DA001 injected into the core was 9 × 107 cells/ml. The error bars represent 1 standard deviation of the mean.

Conclusions. This research resulted in the development of a long-term fluorescent staining technique for nondividing cells which has no apparent effects on cell viability, adhesion, and transport characteristics. The stained cells were quantifiable by epifluorescence microscopy, microplate spectrofluorometry, and flow cytometry. Cells stained with CFDA/SE could be distinguished from autofluorescing sediment particles by microscopy, allowing cell densities in sediment samples to be determined. Transport of DA001 cells determined by enumeration of CFDA/SE-stained cells was identical to DA001 transport determined by more established methods. It is believed that the utility of CFDA/SE for tracking microorganisms will become limited if the stained cells begin to grow, since this will result in approximately twofold dilution of the fluorescent moiety with each successive cell division. Additional work with this technique that will be reported elsewhere includes field-scale bacterial transport experiments using CFDA/SE-stained DA001 cells with near-real-time microplate spectrofluorometry enumeration (M. Fuller, B. Mailloux, P. Zhang, S. Vainberg, W. Johnson, T. Onstott, and M. DeFlaun, submitted for publication); experiments examining protozoan predation dynamics using CFDA/SE-stained cells under both laboratory and field conditions; experiments coupling the staining procedure with an immunomagnetic separation-concentration (ferrographic) method to reduce the lower detection limit to 5 to 10 cells/ml (22); and several more intact-core bacterial transport experiments.