Vol. 50, No. 3APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Sept. 1985, p. 580-5880099-2240/85/090580-09$02.00/0Copyright C) 1985, American Society for Microbiology

Characterization of Subsurface Bacteria Associated with TwoShallow Aquifers in Oklahoma

DAVID L. BALKWILL1* AND WILLIAM C. GHIORSE2Department of Biological Science, The Florida State University, Tallahassee, Florida 32306%1 and Department of

Microbiology, Cornell University, Ithaca, New York 148532

Received 26 December 1984/Accepted 21 June 1985

The bacterial microflora of two shallow aquifers (saturated subsurface zones) in Oklahoma was character-ized by direct observation with light and electron microscopy, by plating, and by examination of colonymorphology and distribution. Isolated bacterial strains were also examined. Total cell counts varied onlyslightly (2.9 x 106 to 9.8 x 106 g [dry wtf-') from sample to sample, whereas colony counts varied widely (6.3X 102 to 6.5 x 106 CFU g [dry wtf-'). Colony counts on nutritionally rich media were lower than onlow-nutrient media, especially in samples from the saturated zone. The variety of colony types growing onnutritionally rich media decreased with increasing depth and saturation. Colony counts of anaerobic bacteriaalso decreased with depth but were at least 100-fold lower than aerobic counts on most media. Cellmorphologies of bacteria grown aerobically on plates included short rods, cocci, and actinomycete-like forms.Direct light microscopic observation of sediments revealed short, rod-shaped, and coccoid bacterial cells;endospores, actinomycete spores, and eucaryotic forms were not observed by light microscopy. Electronmicroscopic observation of bacteria released from the samples revealed that 85 to 90% of them were coccoid,gram-positive, Arthrobacter-like organisms, some of which were dividing or contained completed division septa;other types of gram-positive and gram-negative bacteria were present in lower numbers. Isolated bacterialstrains were able to grow on both nutritionally rich and low-nutrient media. A higher proportion ofgram-negative organisms was isolated than gram-positive organisms. Most of the isolates were capable ofstoring polyphosphate, poly-Il-hydroxybutyrate, or polysaccharide. The results of this study suggest that themicrobial population of these two shallow aquifers is dominated by aerobic, nutritionally versatile bacteria thatcan subsist on low concentrations of organic compounds without forming specialized resting cells. Other typesof microorganisms, such as facultatively anaerobic bacteria and microeucaryotes, may also be present, but theyrepresent only a small fraction of the microflora.

Our initial microscopic observations on the microflora ofshallow water table aquifers (saturated subsurface zones) inOklahoma, Louisiana, and Texas (7, 8, 15, 16; D. L.Balkwill and W. C. Ghiorse, Proceedings of the Sixth Inter-national Symposium of Environmental Biogeochemistry,1983, in press) showed that bacteria are the predominantforms of life in these subsurface environments. Additionalwork on samples from the same locations suggests that atleast part of the subsurface microflora is metabolically activeand that it has the potential to degrade some of the aromaticand halogenated hydrocarbon pollutants of groundwater (14,16, 17). Based on these initial findings, we have proposed(16) that shallow aquifer sediments are inhabited by stablepopulations of bacteria that can survive in these oligotrophicenvironments (<0.04% total organic matter in the sediment,<10 mg of dissolved organic C liter-' in the groundwater) bymetabolizing low levels of residual organic substances whichhave percolated down to aquifer zones from the soil above.

This proposal immediately raises questions concerning theidentity and characteristics of the aquifer bacteria. Are theymorphologically or metabolically unusual? Or are they un-usual in any other way? Have specific types of bacteria fromthe surface soil been selected by the environmental condi-tions of the subsurface? If so, which types have colonizedthe subsurface and how have they adapted to theoligotrophic conditions therein?

* Corresponding author.

An initial approach to answering some of these questionsis to examine aquifer bacteria by a variety of direct andindirect methods and then to consider the microbial charac-teristics observed by these different techniques. In thepresent study, we report the results of such a study involvingsubsurface samples from two sites in Oklahoma. Thesesampling sites were chosen because they are far-removedfrom potential sources of chemical pollution and becausethey are typical of shallow aquifers used widely in the UnitedStates for drinking water and irrigation.(A preliminary report of this work was presented at the

82nd Annual Meeting of the American Society for Microbi-ology, Atlanta, Ga., 7 to 12 March 1982 [D. L. Balkwill andW. C. Ghiorse, Abstr. Annu. Meet. Am. Soc. Microbiol.1982, N85, p. 192].)

MATERIALS AND METHODS

Subsurface samples. Most of the subsurface sedimentsexamined in this study (Table 1) were obtained from a sitelocated in the margin of the floodplain of a small creek nearLula, Okla. A geological profile of the Lula site is shown inFig. 1 (see also reference 16). A single sample (no. 5) wastaken from a similar site at Pickett, Okla. (8). The depth towater table at both sites was ca. 3 m. The depth to bedrock(i.e., nearly consolidated, very plastic clay) at the Lula sitewas 7 m. The groundwater at both sites was free of organicpollutants and contained <10 mg of dissolved organic Cliter-'. The total organic content of the sediment was

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SUBSURFACE BACTERIA 581

TABLE 1. Total and viable cell counts of bacteria in aquifer sediment samplesViable counts [CFU (SD) g (dry wt)-y] on the following

Samle denifia- Collec- Satu- Moisture Incu- AODC [no. of media":Sample Identifica- Depthmei'tion ration content bation cells (SD) g (dryno. tion no." date (m) statusb (%) atm' wt)-1] PTYG agar Dilute PTYG SSAagar

1 2/81 1.2 U 13.31 AE 6.8 (4.9) x 106e 3.4 (0.9) x 104 1.9 (0.4) x 10 1.3 (0.2) x 1051 2/81 1.2 U 13.31 AN ND 2.1 (0.6) x 103 1.2 (0.5) x 104 5.0 (1.9)X 1032' 6/81 1.2 U 14.35 AE 9.8 (1.3) x 106 3.0 (0.6) x 103 3.1 (0.8) x 104 2.1 (0.5) x 1042E 6/81 1.2 U ND9 AE 6.8 (0.9) x 106e ND ND ND1A 2/81 3.1 I 18.26 AE 3.4 (2.6) x 106e 2.0 (0.5) x 104 2.6 (0.2) x 106 2.9 (0.6) x 1061A 2/81 3.1 I 18.26 AN ND 3.0 (1.0) x 102 4.0 (2.0) X 104 5.0 (1.0) x 1032A 6/81 3.1 I 17.94 AE 3.7 (1.0) x 106 1.2 (0.3) x 103 6.0 (0.2) x 103 7.4 (2.1) x 1032Af 6/81 3.1 I ND AE 4.6 (3.4) x 106 ND ND ND3 9B-1 11/82 2.8 I 17.2 AE 2.9 (3.1) x 106 3.3 (0.3) X 10- 6.1 (0.7) x 105 1.1 (0.1) x 1064 9F-1 9/83 3.1 I 25.0 AE 4.8 (2.3) x 106 5.7 (0.6) X 104 5.8 (0.9) X 104 6.5 (0.9) X 1041B 2/81 4.9 S 19.35 AE 6.8 (4.3) x 106e 2.6 (0.7) X 103 3.5 (0.1) x 106 4.1 (0.2) x 1061B 2/81 4.9 S 19.35 AN ND 2.0 (2.0) x 102 7.0 (5.0) x 102 <1022B 6/81 4.9 S 23.60 AE 3.8 (1.1) x 10O 8.4 (1.1) x 104 7.7 (1.1) X 105 3.8 (1.6) x i052Bf 6/81 4.9 S ND AE 4.8 (1.9) x 10&9 ND ND ND3A 9B-3 11/82 4.5 S 19.3 AE 5.0 (3.8) x 106 2.4 (0.4) x 104 3.0 (1.0) x 10 4.4 (0.3) x 1053B 9B-4 11/82 4.4 S 13.6 AE 3.8 (3.0) x 106 ND ND ND4A 9G-2 9/83 4.7 S 19.3 AE 4.0 (2.0) x 106 2.0 (1.5) x 103 4.7 (2.6) X 104 2.0 (2.0) X 1054B 9G-4 9/83 6.3 S 19.9 AE 9.3 (2.9) x 106 6.4 (1.5) x 104 6.8 (0.8) x 105 1.1 (0.3) x 1065h 4/81 5.5 S 20.46 AE 5.2 (3.1) x 106 6.3 (1.0) x 102 3.1 (1.0) x 106 2.5 (1.0) x 106

c Identification number is the sample designation used by staff scientists (at the U.S. Environmental Protection Agency Robert S. Kerr Environmental ResearchLaboratory in Ada, Okla.) who collected the samples (see text).

b U, Unsaturated sample from above water table; S, saturated sample from within water table; I, sample from interface between saturated and unsaturatedzones.

AE, Aerobic; AN, anaerobic (N,).d Average of triplicate spread plates.eData are from reference 16.f Sample was prefixed with glutaraldehyde in the field.9 ND, Not done.hSample is from Pickett, Okla; all other samples are from Lula, Okla.

approximately 0.1% in the unsaturated zones and 0.04% in theinterface and saturated zones (16).The Lula site was sampled four times over a period of 2.5

years. In February and June 1981, samples were taken fromthe unsaturated zone (located in the B horizon) above thewater table (depth, 1.2 m), from the interface zone (locatedin the C horizon) at the upper surface of the water table (3.1m), and from the saturated zone (unconsolidated material)beneath the water table (4.9 m). In November 1982 andSeptember 1983, samples were taken only from the interface(2.8 to 3.1 m) and saturated (4.4 to 6.3 m) zones. The singlePickett site sample was taken from the saturated zone (5.5m) in February 1981.

All of the samples were obtained and handled aseptically(16) and then shipped to us on ice (0 to 5°C) by staff scientistsat the U.S. Environmental Protection Agency Robert S.Kerr Environmental Research Laboratory in Ada, Okla. Thesamples were stored at 4°C until they were examined as

described below.Total cell counts. An acridine orange direct count (AODC)

method was used to determine the total numbers of micro-bial cells in subsurface sediments and to observe the mor-

phological characteristics of subsurface microorganisms byepifluorescence light microscopy. Details of the AODCprocedure, a modification of the methods of Trolldenier (12)and Conn (5) for examining microorganisms in soil, havebeen described previously (7, 16).

Viable cell counts. Numbers of viable bacteria (CFU) insubsurface sediments were estimated by plate counts within48 h after collection of each sample. Total cell counts did notincrease during this period (8). Plate counts were done byfirst blending 10 g of subsurface sediment with 100 ml of

sterile 0.1% sodium pyrophosphate (Na4P207 11OH20; pH7) for 1 min (two 30-s bursts separated by a 30-s rest interval)in a Waring blender to produce a well-dispersed suspension.Serial dilutions of this suspension were then prepared withsterile 0.1% sodium pyrophosphate, and fractions of theresulting dilutions were spread-plated in triplicate on each ofthree plating media: PTYG agar, dilute PTYG agar, andsurface soil extract agar (SSA). PTYG agar contained thefollowing ingredients per liter of distilled water: glucose, 10g; yeast extract, 10 g; peptone, 5 g; Trypticase (BBLMicrobiology Systems), 5 g; MgSO4 * 7H20, 0.6 g;CaCl2 - 2H20, 0.07 g; agar, 15.0 g. Dilute PTYG was a 1:20dilution of the PTYG formula, except that the concentrationof agar was kept at 15 g/liter. SSA contained the followingingredients per liter: soil extract, 100 ml; distilled water, 900ml; agar, 15 g. The soil extract was made by autoclaving a1:2 suspension of surface soil from the sampling site indistilled water for 2 h, after which the liquid extract wasclarified by centrifugation and filtration. The final pH of SSAwas adjusted to approximate the pH of the subsurfacesample being plated in all cases. Plates were incubatedaerobically for 1 (PTYG agar) or 2 (dilute PTYG and SSA)weeks at 22 to 25°C.

Plating was also used to determine the numbers of viableanaerobic or facultatively anaerobic bacteria in samples 1,1A, and 1B from the Lula site. Plating was performed asdescribed above, except that the aquifer samples were

opened and all of the plating manipulations were carried outin a large, plastic anaerobic (N2 atmosphere) chamber-glovebox. The plates were then incubated anaerobically for 2weeks at 25 to 27°C.

Plates were evaluated by first counting the total number of

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582 BALKWILL AND GHIORSE

colonies and then defining each of the recognizably distincttypes of colonies. The total number of distinct types and thecolony morphologies of the numerically predominant types(all types representing at least 10% of the total number ofcolonies on plates from the countable dilution) were re-corded. The plates from the countable dilution were thenphotographed with Kodak Ektachrome T160 color transpar-ency film to facilitate comparison of colony types obtainedon different collection dates. Cellular morphologies in nu-merically predominant colony types were determined byphase-contrast light microscopy of wet mounts.

Electron microscopy. Subsurface microorganisms wereprepared for electron microscopic examination in two ways:(i) by flotation of surface films and (ii) by release andconcentration for subsequent thin sectioning. For the filmflotation method, surface films were obtained by submergingaquifer sediment particles in sterile water as described byWaid (13). These films were then retrieved on carbon-stabilized, Formvar-coated, 200-mesh, copper specimengrids and prepared for electron microscopy by negativestaining with 1% phosphotungstic acid (pH 7.0) or 0.5%uranyl acetate (pH 5). Release and concentration of micro-bial cells for subsequent thin sectioning was achieved by acentrifugal washing method similar to that originally devel-oped by Balkwill et al. (2). Briefly, glutaraldehyde-prefixedsediment samples were subjected to repeated (5 to 15 times),low-speed (450 to 650 x g) centrifugal washing with 0.1%Na4P207 * 10H20 or 0.1% Na4P207 * 10H20-0.5%polyvinylpyrrolidone-360 (Sigma Chemical Co., St. Louis,Mo.) to release microbial cells from the abiological solids.The supernatant fractions from these washings were pooled(the final pellet was discarded) and centrifuged for 20 min at23,000 x g to concentrate the released microbial cells for

DEPTHm ft

_3 - 10

I ..

-252I

.w3 --10

4 .

9.5 2090.

7 -

-25

.30

95:31.7

--Top soil- loamy sand

*-Fine silty clay with iron stains

/Permanent Water Table

Fawn colored sand with minor clay content

-Mixture of coarse sands and gravels

-Uniform fine sand

-Gravel with sand matrix

*-Very plastic clay (not always present)

VCoarse gravel with sand matrix

Massive, very plastic clay4 olive green with maroon mottling

FIG. 1. Diagram of the vertical geological column at the Lula,Okla., site from which most of the subsurface sediment sampleswere taken (sampling depths indicated by arrows).

TABLE 2. Number of recognizably distinct colony types onenumeration plates

Total no. of distinct colony types onb:Sape SaturationSample statusa PTYG agar Dilute SSA

PTYG agar

1 U >10 7-8 6-72 U >12 8-9 6-7

1A I 3-4 3-4 3-42A I 4-5 3-4 3-4

1B S 1-2 1-2 7-82B S 1-2 2-3 8-93A S 2-3 3-4 >104B S 1-2 2-3 6-75 S 1-2 2-3 7-8a See footnote b in Table 1 for abbreviations.b Number of visibly different colony morphologies present on plates from

the countable dilution.

electron microscopy. The procedures for subsequent fixa-tion, dehydration, embedding, and thin sectioning have beendescribed previously (7, 16).Samples for electron microscopy were viewed with a

JEOL JEM-1OOS or a Philips EM-201 transmission electronmicroscope operating at an accelerating potential of 80 kV.All specimen grids were searched thoroughly for microbialcells, and all of the cells observed were photographed atmagnifications sufficient to reveal detailed ultrastructuralfeatures.

Isolation and cultivation of subsurface bacteria. To isolatesubsurface bacteria, 1 g of aquifer sediment was added to 10ml of a buffered salts solution (pH 7.0) containing thefollowing ingredients per liter of distilled water: NaCl, 6.4 g;KCI, 0.16 g; K2HPO4, 0.92 g; KH2PO4, 0.16 g; CaCl2 * H20,0.13 g; MgCl2 * 6H20, 0.10 g. This mixture was agitated on arotary shaker for 30 min at 160 rpm, after which 0.1 ml wasspread on aquifer sediment extract agar (ASA) prepared asdescribed above for SSA except that a 1:1 suspension ofLula or Pickett aquifer sediment and distilled water wasextracted. The pH of the extracts was ca. 6.3. Colonies thatappeared on ASA were transferred to fresh ASA and to PYGmedium containing per liter of distilled water: peptone, 0.25g; glucose, 0.25 g; yeast extract, 0.25 g; MgSO4 * 7H20, 0.6g; CaC12 * H20, 0.07 g; agar, 15 g. All plates were incubatedaerobically at 22 to 25°C. In some instances the isolates weretested for their ability to grow on Difco nutrient agar (NA).The isolates were characterized by standard microbiologicalmethods (6), and genus designations were made according toBergey's Manual ofDeterminative Bacteriology, 8th ed. (4).

RESULTSTotal cell counts (AODCs). The total cell count (AODC) of

bacteria in all aquifer sediments examined in this studyvaried only slightly, ranging from 2.9 x 106 to 9.8 x 106 g(dry wt)-1 (Table 1). The AODCs of the Lula samples did notchange significantly with increasing depth or saturation, nordid they fluctuate markedly over time. A comparison of thesaturated samples from Lula and Pickett showed that thesesamples contained essentially equal numbers of bacterialcells.

Viable cell counts. Viable cell (plate) counts of subsurfacemicroorganisms varied widely in comparison to the AODCs,ranging from 6.3 x 102 to 6.5 x 106 CFU g (dry wt)-1 (Table

SAMPLENO.

1A,2A,4-3

3A,3B,4A-1B,2B

4B

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SUBSURFACE BACTERIA 583

TABLE 3. Predominant colony and cell morphologies present on enumeration plates of samples collected in February 1981

Frequency (%) of occur-

TypeColony morphology

CellrenceonSb

no. Eelleopa-ogyno.' DiluteColor Size Surface Edge Elevna- PYG PtY SSA

1 Dark green Medium Glossy Smooth Slight Coccoid rod 1 32 44 01A 50 22 0

2 Cream-yellow Small Glossy Smooth Raised Medium-sized rod 1 12 27 01A 0 76 0

3 Light brown Large Glossy' Irregular Raised Pleomorphic rod 1A 49 0 04 Cream Large Glossy Irregular Raised Medium-sized rod 1B 95 98 05 Transparent Small Glossy Smooth Slight Coccoid rod 1 0 0 39

1B 0 0 39

6 Cream Medium Glossy Smooth Slight Pleomorphic rod 1 0 0 397 Brown Small Powdery Irregular Flat Branching filaments 1 0 0 168 Light brown Small Glossy Irregular Partiald Coccoid rod 1A 0 0 969 Blue-gray Small Powdery Irregular Flat Branching filaments 1B 0 0 50

" See Table 1 for further information.b Percentage of total colonies on countable plates of indicated medium; colony types representing less than 10% of the total are not listed.' This type produced excessive amounts of extracellular slime, some of which dripped onto the covers of inverted plates during incubation.d Only the center of the colony was elevated, thereby giving the colony a fried-egg appearance.

1). The fluctuations in viable counts did not correlate withdepth, degree of saturation, time of sampling, or samplingsite (Lula versus Pickett). Even though viable counts were

always lower (usually much lower) than AODCs for thesame sample, they sometimes exceeded 50% of the AODCvalue (e.g., samples 1A and 1B). Plate counts on twonutritionally dilute media (diluted PTYG agar and SSA) were

consistently higher than those on a nutritionally rich medium(undiluted PTYG agar), sometimes by as much as three tofour orders of magnitude. The effect of low-nutrient mediaon plate counts was most pronounced in samples from thesaturated zone (Table 1).Colony counts of anaerobic or facultatively anaerobic

bacteria in samples 1, IA, and 1B from the Lula site(collected February 1981) generally decreased with increas-ing depth (Table 1). Moreover, these anaerobic counts were

always much lower (from one to four orders of magnitude)than the corresponding aerobic counts on the same medium.Because of the very low numbers of anaerobes detected inthese samples, anaerobic colony counts were not performedon any later samples.Colony and cell morphologies on aerobically incubated

plates. An analysis of colony morphological diversity on

aerobically incubated plates (numbers on anaerobic plateswere too low for accurate analysis) in relation to depth andsaturation status is summarized in Table 2. The number ofdifferent colony types appearing on PTYG agar plates de-creased sharply with increasing depth and saturation, even

though AODCs and total viable counts remained relativelyconstant (Table 1). A similar effect was noted on dilutePTYG plates, but it was less pronounced. In contrast, thenumbers of different colony types on SSA plates remainedconstant or increased slightly with increasing depth andsaturation.The numerically predominant colony and cell morpholo-

gies observed on plates of the Lula, Okla., samples collectedin February 1981 are listed and described in Table 3; similarmicrobial forms were predominant on plates of samplestaken at other times. PTYG and SSA media apparentlyselected for different sets of microorganisms. PTYG and

dilute PTYG agar plates contained sizable proportions (12 to95%) of obviously rod-shaped organisms (i.e., organismswhose length was at least twice their diameter). In contrast,very short, coccoid rods predominated on SSA plates,possibly owing to the relatively low nutrient levels in thismedium. The SSA plates also contained actinomycete-like,branching, filamentous organisms (up to 50%) that did notgrow on PTYG or dilute PTYG plates.The predominant colony morphologies on plates (Table 3)

varied both with the plating medium and with the depth andsaturation status of the sample. On PTYG agar, for example,the distinctive green colony type that predominated onplates of the unsaturated sample (1.2 m) did not appear onplates of the saturated sample (4.9 m). This organism pro-duced much less pigment on dilute PTYG agar, an indicationthat nutrients can affect the pigmentation of some subsurfacemicroorganisms. On SSA, an actinomycete-like organismpredominated on plates of the saturated sample, but it did

TABLE 4. Predominant colony morphologies on enumerationplates of samples from the saturated zone at Lula collected on

different dates

Percentage of total coloniesPlating Sample Collection presentb on colony type':medium no.' date

3 4 5 9 10

PTYG 1B 2/81 <1 95 __dPTYG 2B 6/81 <1 96PTYG 3A 11/82 34 60 -PTYG 4B 9/83 10 85SSA 1B 2/81 - 39 50 <1SSA 2B 6/81 - 33 54 <1SSA 3A 11/82 - 24 36 39SSA 4B 9/83 31 65 <1

"See Table 1 for further information.Percentage of total colonies on plates from the countable dilution; colony

types representing less than 10% of the total are not included.C See Table 3 for a description of colony types; colony type 10 is a pale pink,

small, glossy, regular, raised, coccoid rod.d _, Colony type did not grow on this plating medium.

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FIG. 2. Epifluorescence light micrograph of acridine orange-stained subsurface material from sample 1B (see Table 1). Arrow indicatessmall group of cells with similar morphological characteristics.FIG. 3. Electron micrographs of thin sections through the predominant gram-positive, coccoid bacterial form seen in Lula samples. Cells

were released from subsurface sediments by low-speed centrifugal washing. Bars, 0.25 ,um. (a) Representative cell from the unsaturated zone(sample 2). (b) Representative cell from the saturated zone (sample 2B); note the different locations of the nuclear material in Fig. 3a and 3b.(c) Cell from the unsaturated zone (sample 2) with a large, mesosome-like structure (arrow). (d) Cell from the saturated zone (sample 2B) withan Arthrobacter-like division septum (arrow). (e) Cell from the unsaturated zone (sample 2) that lacks most of the usual cytoplasmicconstituents and that may be dying.

not appear on plates of the unsaturated and interface (3.1 m)samples.The predominant colony and cell morphologies growing

on plates of samples from the saturated zone at Lula (Table4) remained reasonably constant over the 2.5-year durationof this study. A medium-sized rod that produced large,cream-colored, glossy colonies predominated on PTYGplates. An actinomycete-like branching filament that pro-duced blue-gray, powdery colonies and coccoid rods thatproduced small, transparent, glossy colonies predominatedon SSA plates. The only significant variation occurred withthe sample taken in November 1982. A previously rare form(a pleomorphic rod that produced exceedingly slimy, light

brown colonies) represented 34% of the colonies on PTYGplates at this tite, and a previously unseen form (a coccoidrod that prodticed small, pink, glossy colonies) represented39% of the colonies on SSA plates. By September 1983,however, the predominant colony types were again similarto those observed on plates of the earlier (February 1981 andJune 1981) samples.

Morphological characteristics of microorganisms in subsur-face samples. Aquifer sediment materials contained primarilyshort rods and coccoid cells that occurred singly or inclusters (Fig. 2). Larger microcolonies were observed rarelyin samples from the unsaturated zone. Bacterial endospores,which exhibit a characteristic epifluorescence image when

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SUBSURFACE BACTERIA 585

5bFIG. 4. Electron micrographs of thin sections through minority bacterial forms seen in Lula samples. Cells were released from subsurface

samples by low-speed centrifugal washing. Bars, 0.25 pLm. Note lack of ribosomes in comparison to cells in Fig. 3. (a) Gram-positivebacterium from the saturated zone (sample 4A) with intracellular inclusion body that resembles a polyphosphate granule (arrow). (b)Gram-negative bacterium from the saturated zone (sample 3B); note the small diameter.FIG. 5. Electron micrographs of negatively stained subsurface microorganisms from the unsaturated zone at Lula (sample 2), as released

from the subsurface sediments with the Waid flotation procedure (13). Bars, 0.25 ,um. (a) Cell with typical gram-negative surface topology.(b) Apparent dividing cell. (c) Cell with polar flagellum.

stained with acridine orange (W. C. Ghiorse, unpublisheddata), were not observed. Structures that resembledactinomycete spores were not observed either; however,these may have been overlooked because their appearancein the epifluorescence microscope would probably be similarto that of the coccoid bacterial cells. Neither fungal sporesnor protozoan cysts were observed microscopically. How-ever, their numbers were probably too low (<105 g [drywt'-1) to detect them microscopically (J. L. Sinclair andW. C. Ghiorse, manuscript in preparation).

Ultrastructural characteristics of microorganisms in subsur-face samples. Transmission electron microscopy of thin-sectioned microorganisms that had been released and con-centrated from subsurface samples by low-speed centrifugalwashing revealed distinct microbial cells surrounded bylarge amounts of abiotic debris (Fig. 3 and 4). The abiotic

materials (probably clay particles) were usually muchsmaller than the cells and sometimes appeared to be clingingto their surfaces.The predominant forms (85 to 90%) in all subsurface

samples, regardless of depth or collection date, were procar-yotic, gram-positive, coccoid rods that ranged from 0.4 to0.9 ,um in diameter (Fig. 3). The cytoplasm in these formswas densely packed with nuclear material and ribosomes,seldom containing inclusion bodies or storage granules. Thenuclear material was usually situated in the center of thecytoplasm in cells from unsaturated or interface samples(Fig. 3a), but was often displaced to a single peripherallocation in the cytoplasm of cells from saturated samples(Fig. 3b). Approximately 50% of the cells from all samplescontained elaborations of the cytoplasmic membrane (Fig.3c) that ranged in complexity from simple loops at the

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TABLE 5. Properties of bacterial strains isolated from Oklahoma subsurface sediment samples

Sugar GrowthStrainSaple Date Satu- Gram Oxi- fer- onrogbenu

Strain Sample Date ration reac- Morphology Flagella dase men- SSA, Storageb Other features Probablestatus' tion tation PYG,

NA

Lula B 1B 2/81 S - Short rod Polar + - + PP Nonfluorescent Pseudomo-nas

Lula D 1B 2/81 S + Rod-coccus None + - + PP, PS Snapping division ArthrobacterLula V 3 11/82 I - Short rod Polar + - + PP, PHB, PS (?) Nonfluorescent Pseudomo-

nasPickett 5 4/81 S - Short rod Polar and + - + PP (?), PHB, PS Purple pigment Chromobac-

1 lateral terium (?)Pickett 5 4/81 S + Long rod Peritri- + - + PP Better growth on Brevibacter-

3 chous PYG and SSA ium (?)Pickett ? ? S - Short rod Polar + - + PHB, PS Nonfluorescent Pseudomo-

3B' tufts nasa See footnote b in Table 1 for abbreviations.b PHB, poly-p-hydroxybutyrate; PP, polyphosphate; PS, polysaccharide.' Isolated by J. T. Wilson.

periphery of the cell to large, centrally located, mesosome-like structures. Dividing cells and cells with completeddivision septa were observed in all samples (Fig. 3d), al-though they were somewhat more prevalent in unsaturatedsamples. The structural aspects of the cell division mecha-nism were similar to those seen in Arthrobacter species in allcases (Fig. 3d). A small proportion of these predominantgram-positive coccoid rods (less than 3%) lacked many ofthe usual cytoplasmic features (Fig. 3e) and appeared to bedead or dying. Consistent with this interpretation was theoccasional presence of what appeared to be empty cell wallsthat no longer contained membranes or cytoplasmic constit-uents.

In addition to the predominant gram-positive coccoid rodsdescribed above, most samples contained a minority (lessthan 15% of ca. 200 cells observed) of other gram-positive(Fig. 4a) and gram-negative (Fig. 4b) bacterial forms. Eu-caryotic forms were not detected by transmission electronmicroscopy in any of the samples. The minority bacterialforms were morphologically more variable than the predom-inant Arthrobacter-like types, ranging in shape from appar-ent spheres to long rods. They were also more variable insize, ranging from 0.25 to 1.5 ,um in diameter. The cytoplasmin most minority forms differed from that of a typicalpredominant cell (see above); its overall electron densitywas much lower because comparatively few ribosomes werepresent. This situation, in turn, caused the nuclear materialto spread throughout the cell and appear as distinctly sepa-rated strands (Fig. 4). In contrast to the predominant forms,the minority cells often contained poly-p-hydroxybutyrategranules or other storage bodies (Fig. 4a) in their cytoplasmand very seldom appeared to be dividing.Transmission electron microscopy of negatively stained

surface films released from subsurface materials by the Waidflotation procedure (13) confirmed that both gram-positiveand gram-negative bacterial cells were present in thesesamples. Again, eucaryotic forms were not detected in anyof the samples examined by this technique. Gram-negativebacteria were easily recognized by their distinctive, wrinkledsurface topology (Fig. 5a). Dividing cells were also presentin the flotation films (Fig. 5b), as were a small number (<5%)of cells with polar flagella (Fig. 5c).

Characteristics of isolated subsurface microorganisms. Itwas generally difficult to obtain stable isolates from thesubsurface specimens examined in this study because many

of the organisms that appeared on plates failed to grow whentransferred to fresh media. Nevertheless, a number of stablebacterial strains were isolated from the samples by plating onASA and then transferring to PYG and fresh ASA. All of theisolates, whose properties are summarized in Table 5, couldalso grow on NA, an indication of their nutritional flexibility.Gram-negative isolates were more common than gram-positive isolates, probably because the media and conditionsemployed for isolation were selective for gram-negativeisolates. Among these isolates were three gram-negative,rod-shaped pseudomonads that were morphologically simi-lar (Fig. 6a) to the polarly flagellated cells seen in flotationfilms (Fig. Sc). One of these isolates (Lula V) stored largeamounts of poly-3-hydroxybutyrate, polyphosphate, and,possibly, polysaccharide. All three pseudomonads werenonfluorescent. At least one of the gram-positive isolateswas an Arthrobacter-like coryneform bacterium that dis-played snapping-type cell division and the bridging wallstructure at the division septum that is typical of coryneformbacteria (Fig. 6b). This isolate (Lula D) underwent thetypical rod-to-coccus morphological change in the late-logphase of growth. Coccoid bacteria with similar morphologi-cal properties were often observed in thin sections of sub-surface samples (Fig. 3d).A purple-pigmented Chromobacterium sp. (Pickett 1), an

unidentified gram-positive rod that may be a Brevibacteriumsp. (Pickett 3), and two uncharacterized gram-negative rods(Pickett 1B and 2B) were isolated from Pickett samples alongwith a pseudomonad (Pickett 3B). These uncharacterizedisolates were nonpigmented, oxidase negative, andnonfermentative.

It should be noted that some of the Lula samples plated onASA consistently yielded powdery white, actinomycete-likecolonies that grew on NA and produced brownish pigments.The frequency with which they were observed and theconsistency of their appearance indicated that actinomycetespores probably were present in these samples in significantnumbers.

DISCUSSION

The data presented here are consistent with our previousfindings (7, 8, 16; Balkwill and Ghiorse, in press) that aquifersediments are inhabited by appreciable numbers (ca. 106 to107 cells [dry wt] g-1) of bacteria but by few eucaryotic

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SUBSURFACE BACTERIA 587

6a _bFIG. 6. Electron micrographs of selected isolates of subsurface microorganisms, as grown in laboratory culture. Bars, 0.25 p.m. (a)

Negatively stained whole-cell preparation of isolate Lula B (see Table 5); note the polar flagella. (b) Thin section through dividing cell ofisolate Lula D (see Table 5); note the typical Arthrobacter-like division septum (arrow).

microorganisms. It can be argued that significant bacterialpopulations probably occur universally in shallow aquifersediments such as these, because comparable bacterial den-sities have now been found in samples from several differentlocations in Oklahoma (this study; 16), Louisiana (7), Texas(14), and Georgia (unpublished data). These findings arecontrary to earlier results that led historically to the errone-ous assumption that microbial life in subsurface soils waseither nonexistent or extremely limited (for a summary of theearlier work, see references 1 and 10). We explain thisapparent discrepancy by attributing it to our use of directmicroscopic methods for enumerating and characterizing thesubsurface microflora, in preference to the usual culturaltechniques employed in the past.Our results (Table 1) show that many subsurface bacteria

grow far more readily on dilute media (especially SSA) thanon typical, nutrient-rich plating media like PTYG agar. Thisfinding is consistent with our previous speculations (7, 16)that most microorganisms in aquifer sediments are adaptedfor growth and survival under low-nutrient (oligotrophic)conditions. The low-nutrient adaptation of these bacteriaappears to be more pronounced in the saturated zone be-cause, at the Lula site, the number and diversity of colonytypes that appeared on PTYG agar decreased sharply withincreasing depth (Table 2), whereas those that grew on SSAplates increased. Such a change may reflect a change in thequality of metabolizable organic carbon compounds in thesaturated zone which could have selected for different typesof bacteria capable of metabolizing the organic matter there.Indeed, these results support our previous suggestion (16)that the deeper saturated zones in aquifers may be inhabitedby relatively few types of oligotrophic bacteria that canmetabolize the recalcitrant degradation products of organicmatter that reaches these zones from above.The present results appear to be in conflict with those of

Hirsch and Rades-Rohkohl (9), who found enormous mor-phological diversity in long-term, low-nutrient enrichmentcultures of the groundwater samples they investigated. Thisdiscrepancy probably is best explained on the basis that theirenrichments were inoculated with groundwater from long-standing wells rather than with sediment material. It remain

to be determined whether the aquifer sediments from whichthe groundwater samples were obtained actually harboredthe same range of morphologically diverse bacteria found inthe wells.

Additional indications of the oligotrophic conditions in theLula aquifer sediment are found in the morphology andultrastructure of the subsurface bacteria observed by directmicroscopic methods. Most of the cells were small ovals,spheres, or coccoid rods, all of which are forms that wouldincrease the surface-area-to-volume ratio and thus improvetransport efficiency (11). In addition, the cytoplasms of manycells appeared to be in various stages of depletion, andinternal membranes were observed frequently in the coccoidArthrobacter-like cells that predominated in Lula samples.The starving Arthrobacter crystallopoietes cells studied byBoylen and Ensign (3) displayed similar cytoplasmic deple-tions and elaborations of internal membranes during long-term starvation.Empty gram-positive cell envelopes were observed fre-

quently in the Lula samples, indicating that measurements ofmuramic acid may not be a good choice as an indicator ofactive microbial biomass in aquifer sediments. However,nearly all of the biochemical indicators of biomass measuredin shallow aquifer sediments from Louisiana by White et al.(15) implied a biomass equivalent to ca. 107 bacteria g (drywt)-1, which was very close to the AODC (106 to 107 g [drywt]-f) of the same samples (7). Whether or not thesebiochemical indicators can reflect active or inactive biomassremains to be determined. In this regard, recent comparisonsof ATP measurements and numbers of 2-(p-iodophenyl)-3-(p-nitrophenyl)-5-phenyl tetrazolium chloride-reducing bac-teria with AODC in samples from the Lula site (14) showedthat at least 1 to 10% of the AODC bacteria were metaboli-cally active, as judged by these methods.

Because measurements of ATP and 2-(p-iodophenyl)-3-(p-nitrophenyl)-5-phenyl tetrazolium chloride-reducing activityreflect only the status of the cells in a sample at the time ofthe test, it can be argued that the proportion of the bacterialpopulation with a potential for metabolic activity in aquifersediments may be higher than the suggested 1 to 10%,especially if the inactive cells in a sample could respond to

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588 BALKWILL AND GHIORSE

added nutrients. The fact that viable counts of the Lulasamples on diluted PTYG agar and SSA occasionally were asmuch as 50% of the AODC (Table 1) suggests that a largeproportion of the subsurface bacteria was viable and, there-fore, capable of metabolic activity. It may be speculatedfrom these results that many of the subsurface bacterianormally are not metabolically active; i.e., they are dormant.However, results of our electron and light microscopicstudies showed that typical dormant bacterial forms such asendospores were not common in the samples examined. It ispossible, however, that actinomycete-like spores and cyst-like forms may have been overlooked during epifluorescencemicroscopy and that these structures did not segregate intothe final pellet (ca. 50% recovery) embedded for thin sec-tioning and transmission electron microscopy. The finding ofpowdery colonies containing branching filaments on SSAplates from saturated zone samples (which often accountedfor 50% or more of the colonies on these plates; Tables 3 and4) shows that actinomycete propagules are common and thatthese bacteria may be important members of subsurfacecommunities. Regardless of the possibility that spores werepresent in some samples, our results demonstrate that mostof the subsurface bacteria in aquifer sediments do not formspecial resistant structures.

It is not possible to draw specific conclusions regardingthe properties of in situ subsurface bacterial populationsfrom the characteristics of the bacteria we isolated fromaquifer samples in this study, because it is not yet knownwhether the isolates we obtained are significant componentsof the total microbial population. Gram-negative bacteriagrew more readily than gram-positive bacteria on the mediawe employed for isolation. Yet, our direct microscopicobservations, like the indirect chemical analyses of White etal. (15), suggest that gram-positive bacteria dominate theaquifer microflora while gram-negative bacteria are in theminority. Nevertheless, the properties of the isolates are atleast consistent with one general conclusion derived fromdirect microscopic examination: subsurface bacteria arecapable of survival under extreme oligotrophic conditions.Indeed, all of the subsurface isolates were capable of growthon very dilute media. That they were also capable of growthon nutrient-rich media such as NA (Table 5) indicates that atleast some subsurface bacteria possess considerable flexi-bility in their ability to utilize low and high levels ofnutrients. It should also be noted that bacteria capable ofgrowth under anaerobic conditions were present in all of thethree samples surveyed, but their numbers were usuallymuch lower than those of aerobes in these samples.We have not studied enough isolates at this point to make

broad generalizations about the microflora of aquifer envi-ronments. However, it is clear from our observations thatshallow aquifer sediments contain a large proportion ofaerobic bacteria that appear to be of the types (such ascoryneform bacteria and pseudomonads) that are known fortheir capacity to degrade aromatic compounds that might bepresent in groundwater as a result of lignin degradation.Such bacteria are also known for their ability to survive longterm under low-nutrient conditions such as those whichapparently exist in most subsurface sediments.

ACKNOWLEDGMENTSThis work was supported by Subcontract No. 6931-5 under United

States Environmental Protection Agency Cooperative AgreementNo. CR806931 and by United States Environmental ProtectionAgency Cooperative Agreement No. CR811148.We thank P. Bowman, B. Eaglesham, K. Echols, R. Garen, D.

Maratea, J. Tugel, and K. Whalen for technical assistance. Thegeological profile of the Lula, Okla., site shown in Fig. 1 wasprepared by Lowell Leach at the U.S. Environmental ProtectionAgency Robert S. Kerr Environmental Research Laboratory atAda, Okla.

LITERATURE CITED1. Alexander, M. 1977. Introduction to soil microbiology, 2nd ed.

John Wiley & Sons, Inc., New York.2. Balkwill, D. L., T. E. Rucinsky, and L. E. Casida, Jr. 1977.

Release of microorganisms from soil with respect to electronmicroscopy viewing and plate counts. Antonie vanLeeuwenhoek J. Microbiol. Serol. 43:73-81.

3. Boylen, C. W., and J. C. Ensign. 1970. Intracellular substratesfor endogenous metabolism during long-term starvation of rodand spherical cells of Arthrobacter crystallopoietes. J. Bacte-riol. 103:578-587.

4. Buchanan, R. E., and N. E. Gibbons (ed.). 1974. Bergey'smanual of determinative bacteriology, 8th ed. The Williams &Wilkins Co., Baltimore.

5. Conn, H. J. 1918. The microscopic study of bacteria and fungi insoil. N. Y. State Agric. Exp. Stn. Tech. Bull. 64:3-20.

6. Gerhardt, P. (ed.). 1981. Manual of methods for general bacte-riology. American Society for Microbiology, Washington, D.C.

7. Ghiorse, W. C., and D. L. Balkwill. 1983. Enumeration andmorphological characterization of bacteria indigenous to sub-surface environments. Dev. Ind. Microbiol. 24:213-224.

8. Ghiorse, W. C., and D. L. Balkwill. 1985. Microbiological char-acterization of subsurface environments, p. 536-556. In C. H.Ward, W. Gieger, and P. L. McCarty (ed.), Ground waterquality. John Wiley & Sons, Inc., New York.

9. Hirsch, P., and E. Rades-Rohkohl. 1983. Microbial diversity in agroundwater aquifer in northern Germany. Dev. Ind. Microbiol.24:183-200.

10. McNabb, J. F., and W. J. Dunlap. 1975. Subsurface biologicalactivity in relation to ground-water pollution. Ground Water13:33-44.

11. Poindexter, J. S. 1981. Oligotrophy, fast and famine existence.Adv. Microb. Ecol. 5:63-89.

12. Trolldenier, G. 1973. The use of fluorescence microscopy forcounting soil microorganisms. Bull. Ecol. Res. Comm.-NFR(Statens Naturvetensk Forskningsrad) 17:53-59.

13. Waid, J. S. 1973. A method to study microorganisms on surfacefilms from soil particles with the aid of the transmission electronmicroscope. Bull. Ecol. Res. Comm.-NFR (StatensNaturvetensk Forskningsrad) 17:103-108.

14. Webster, J., G. J. Hampton, J. T. Wilson, W. C. Ghiorse, andF. R. Leach. 1985. Determination of microbial cell numbers insubsurface samples. Ground Water 23:17-25.

15. White, D. C., H. F. Fredrickson, M. J. Gehron, G. A. Smith,and R. F. Martz. 1981. The ground water aquifer microbiota:biomass, community structure, and nutritional status. Dev. Ind.Microbiol. 24:189-199.

16. Wilson, J. T., J. F. McNabb, D. L. Balkwill, and W. C. Ghiorse.1983. Enumeration and characterization of bacteria indigenousto a shallow water-table aquifer. Ground Water 21:134-142.

17. Wilson, J. T., J. F. McNabb, B. H. Wilson, and M. J. Noonan.1983. Biotransformation of selected organic pollutants in groundwater. Dev. Ind. Microbiol. 24:225-233.

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