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Chapter 7. Nonmolecular Methods in Geomicrobiology

Chapter 7. Nonmolecular Methods in Geomicrobiology

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The microcosm will probably contain a mixed population of bacteria, not all of which are likely to

play a role in the geochemical process of interest. Manipulation of the microcosm through qualitative

or quantitative changes in nutrient supply, adjustment of pH or temperature, or a combination of these

factors may cause selective increases of the organisms directly responsible for the geomicrobial process

of interest and intensify the process. Because of enrichment biases, it is possible that a minor member

of the indigenous population may grow to dominate the microcosm, and therefore, care must be taken

before concluding that this member is the causative agent of the geomicrobial process in situ.

In vitro laboratory study of a geomicrobial process may be done by isolating the responsible

microorganism(s) in pure culture, if possible, from a representative sample from the geomicrobially active site. The process originally observed in the field is then recreated with the isolate(s) in

batch or continuous culture. Characterization of the process mechanism will involve qualitative

and quantitative measurements of the biogeochemical transformation(s). It may include genetic and

biochemical studies (see Chapter 8 for more details), where appropriate, as well as an assessment of

environmental effects on the in vitro process. In vitro laboratory study may be important in lending support to field interpretation of geomicrobial processes that are occurring at present or have

occurred in the past.



A geomicrobial process may be the result of a single microbial species or an association of two or

more. An association of microbial species is often called a consortium. The basis for the association may be synergism, in which one type of organism is not capable of carrying out the complex

process but in which each member of the consortium carries out part of the process in a sequential

set of interactions. It is also possible that not all members of an association of microbes contribute

directly to an overall geomicrobial process but instead carry out reactions that create environmental conditions relating, for instance, to pH or Eh that facilitate the geomicrobial process under


Even if a geomicrobial process is the result of the action of a single organism, that organism

rarely occurs as a pure culture in the field. It will usually be accompanied by other organisms, which

may not play a direct role in the geomicrobial process under study although they may compete with

the geomicrobially active agent for living space and nutrients and may even produce metabolites

that stimulate or inhibit the geomicrobial agent to a degree. Three types of microorganisms may be

found in a geomicrobial sample taken in the field: (1) indigenous organisms, whose normal habitat

is being examined and which include the geomicrobially active organism(s); (2) adventitious organisms, which were introduced by chance into the habitat by natural circumstances and which may or

may not grow in the new environment but do survive in it; and (3) contaminants, which were introduced in manipulating the environment during in situ geomicrobial study or sampling. Distinctions

among these groups are frequently difficult to make experimentally.

A criterion for identifying indigenous organisms may be their frequency of occurrence in a given

habitat and in similar habitats at different sites. A criterion for identifying adventitious organisms

may be their inability to grow successfully in the habitat under study and their lower frequency of

occurrence than in their normal habitat. Neither of these criteria is absolute, however. Identification

of a contaminant may simply be based on the knowledge about the organism concerned that would

make its natural existence in the habitat under study unlikely.

7.2.1 IN SITU


To detect geomicrobially active microorganisms in situ, visual approaches including direct observation with the naked eye, light microscopy, or transmission electron microscopy (TEM) or scanning

electron microscopy (SEM), especially environmental SEM (ESEM), are possible. Direct visual

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observation is possible only in rare instances, namely, when the microbes occur so massively as to

be easily seen as, for instance, algal or bacterial mats in hot springs (e.g., in Yellowstone National

Park; see Brock, 1978), or lichen growth on rocks. In most instances, visual observations of microbes

in their natural habitat requires magnification. In soil or sediment, such observation may be made

by the buried slide method or the capillary technique of Perfil’ev (Perfil’ev and Gabe, 1969). In the

buried slide method, a clean microscope slide is inserted in soil or sediment and left undisturbed

for a number of days. It is then withdrawn, washed, and suitably stained. A choice of stains includes

those that discriminate between living and dead cells and those that do not. Examination with a

light microscope, equipped for fluorescence microscopy if fluorescent stains were used (Lawrence

et al., 1997), will then reveal microorganisms, especially bacteria and fungi, which became attached

to the glass surface during burial (Figure 7.1).

In the capillary technique, one or more glass capillaries with optically flat sides are inserted

into soil (pedoscope) or sediment (peloscope). Each capillary takes up soil or sediment solution

and minute soil or sediment particles that become the culture medium for microbes, which entered

the capillary lumen at the moment of emplacement or subsequently. The capillaries can be periodically withdrawn and their contents examined under a light microscope. The capillaries may also be

perfused with special nutrient solutions. The capillaries permit observations of trapped microbes in

a living or nonliving state (Figure 7.2). Using this technique, Perfil’ev and Gabe (1965) discovered

several previously unknown bacteria in soil and sediment, including Metallogenium, Kuznetsovia,

and Caulococcus.

Although the buried slide and capillary methods give an indication of some of the organisms

present in soil or sediment where the slides or capillaries were inserted, they do not indicate whether

the organisms that are seen resided preferentially on the soil or sediment particles or in the pore

fluid. To determine this, direct observation of samples of pore fluid and of soil or sediment particles

is necessary. To observe organisms in pore fluids, axenically collected samples of fluid can be

filtered and any microbial cells in the fluid deposited on suitable filter membranes, which are then

stained and subsequently examined microscopically (e.g., Clesceri et al., 1989).

FIGURE 7.1 Demonstration of microbes in soil by the buried slide method. Slide was buried for 1 week. After

withdrawal from soil and gentle washing, it was stained with crystal violet. Main view is of isolated shorter and

longer rods, and some cocci near soil particles. Inset shows a clump of rods in slime (biofilm?) (×2200).

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FIGURE 7.2 Microbial development in a capillary

tube inserted into lake sediment contained in a beaker and incubated at ambient temperature (×5720).

The oval, refractile structures are bacterial spores.

FIGURE 7.3 Bacteria growing on the surface of

304L stainless steel immersed in tap water (Troy,

New York) for 3 days, stained with fluorescein

isothiocyanate. (Courtesy of Pope DH)

To observe microbes directly on soil or sediment particles or on rock fragments, fluorescence

microscopy may be used in conjunction with staining with fluorescent dyes such as 4′,6-diamidino2-phenylindole (DAPI), acridine orange, or the live/dead stain syto 9/propidium iodide, or with

fluorescently labeled antibodies if specific microbes are being sought (Figure 7.3) (Bohlool and

Brock, 1974; Casida, 1962, 1965, 1971; Edwards et al., 1999; Eren and Pramer, 1966; Huber et al.,

1985; Kepner and Pratt, 1994; Muyzer et al., 1987; Schmidt and Bankole, 1965).

Another approach is to examine sectioned samples by TEM, which is also very useful in detecting fossilized microbes (Barker and Banfield, 1998; Ghiorse and Balkwill, 1983; Jannasch and

Mottl, 1985; Jannasch and Wirsen, 1981; Schopf, 1983). Another approach is the use of SEM (e.g.,

Figure 17.16) (Jannasch and Wirsen, 1981; Sieburth, 1975; LaRock and Ehrlich, 1975; Edwards

et al., 1999).



Recent advances in the techniques of molecular biology have led to the development of powerful

methods for identifying microorganisms and studying their phylogeny. These methods have been

adapted to locate and enumerate microorganisms in environmental samples, even microorganisms

that, though viable, have not yet been cultured in the laboratory (Ward et al., 1990; Stahl, 1997;

Pace, 1997).



To determine the nature of a geomicrobially active organism in terms of its morphology, physiology, and the particular geomicrobial process for which it is responsible, it should be isolated and

cultivated in the laboratory if possible. Some geomicrobial cultures have to be studied in the laboratory in enrichment cultures because of unsuccessful attempts to obtain them in pure culture,

or because the geomicrobial activity of interest is the result of a consortium of microorganisms.

In the case of a consortium, its members should be identified by the application of molecular

techniques, isolated, and characterized, if possible, to determine their particular contribution to

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the overall process. Samples, whatever their nature, brought to the laboratory must be obtained

under conditions as aseptic as possible, that is, with very little or preferably no contamination.

Working surfaces of sampling tools should be thoroughly washed and alcohol-flamed. If a rock

outcrop is to be sampled, use of a rock hammer or chisel may be required. If the interior of a

small rock specimen is to be sampled in the laboratory, the surface of the specimen should be

sterilized if possible. This can be accomplished by using a disinfectant such as carbol-gentian

violet-methanol spray (Bien and Schwartz, 1965) or by flaming briefly with a propane torch, or,

if done in the laboratory, for 1 min with a Bunsen flame (Weirich and Schweisfurth, 1985). If

a sampling device cannot be sterilized, the sample it gathers should be subsampled to obtain a

portion that is least likely to have been contaminated. Rock chips should be collected in sterile

containers. If the collection is done manually, the hands should be covered by sterile surgical

gloves. Weirich and Schweisfurth (1985) devised a special method for obtaining an undisturbed

core of rock with a hollow drill under sterile conditions and with cooling by sterile tap water.

The extracted rock core was aseptically cut into sections with a flamed chisel, and each section was

then aseptically crushed in a flamed mortar mill in sterile dispensing solution and microbiologically tested.



To sample the terrestrial subsurface down to 3000–4000 m or more, drilling methods that depend

on the use of special drilling equipment have been devised, which causes minimal if any contamination during the collection of samples (cores) (Phelps and Russell, 1990; Pedersen, 1993; Griffin

et al., 1997). One method uses modified wireline coring tools, with cores collected in Lexan- or

polyvinyl chloride (PVC)-lined barrels. The drill rig, rods, and tools are steam-cleaned. The drilling

fluid system includes a recirculation tank, the drilling fluid being chlorinated water. Tracers such as

potassium bromide, the dye rhodamine T, fluorescent beads (∼2 µm diameter), and perfluorocarbons

added to the drilling fluid aid in determining to what extent, if any, cores were contaminated during

drilling. The assessment is made by measuring the extent of tracer presence, if any, in the cores.

The extent of bacterial contamination can be determined by quantitative enumeration of bacteria

such as coliforms, which were not expected as part of the normal flora of the core, the enumeration being done on the drilling fluid and core samples (Beeman and Suflita, 1989). If anaerobes as

well as aerobes are sought in subsurface samples, the cores should be kept away from air and be

processed in an oxygen-free atmosphere. Subsamples may then be tested for aerobes and anaerobes

by appropriate culture techniques.

Soil and sediment samples from shallow depths on land surfaces may be collected manually with

an auger or other coring device under aseptic conditions. Cores should be subdivided aseptically for

sampling at different depths. If the cores cannot be obtained with a sterilized sampling device, they

should be subsampled so as to obtain the least contaminated sample.



To obtain aquatic samples, special gear may be required. Water samples at any given depth

below the surface, including deepwater samples, may be obtained with a Van Dorn sampler. It

consists of a piece of large-diameter plastic tube fitted with rubber closures, which can be kept

in an open or closed position at both ends. The plastic tube is mounted on a cable or rope in

such a way that it can be lowered vertically into the water column with the rubber closures held

in an open position by a spring mechanism. While the device is being lowered, water will pass

through it. When a desired depth has been reached, the rubber closures at both ends of the tube

are caused to block the openings of the tube by tripping the spring mechanism by means of a

messenger (brass weight), which slides down the wire or rope to which the sampler is attached.

Below-surface water samples can also be collected with a Niskin sampler, which consists of a

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Ekman dredge. The brass messenger on the rope is 5 cm long.

collapsible, sterile plastic bag with a tubelike opening mounted between two hinged metal plates.

The sampler, with the bag in a collapsed state between the metal plates, is lowered on a cable

or rope to a desired depth. A spring mechanism of the sampler is then activated by a messenger,

as with the Van Dorn sampler, causing the hinged metal plates to open and expanding the bag,

which now draws water into it.

Aquatic sediment samples may be obtained with dredging or coring devices. Lake sediment can be collected with an Ekman (Figure 7.4) or a Peterson dredge (Clesceri et al., 1989,

pp. 10–100) if surface sediment is desired. A corer needs to be used if different depths of a

sediment column are to be examined. Ocean surface sediment may be collected by dragging

a bucket dredge over a desired area of the ocean floor. Such a sample will, however, consist

of combined, mixed surface sediment encompassing the total surface area sampled. To obtain

samples representing different sediment depths at a given location, a gravity (Figure 7.5) or a

box corer (Figure 7.6) has to be used. Such devices are rammed into the sediment. Box corers

of sufficient cross section provide the least disturbed cores. All cores need to be subsampled

to obtain representative, minimally contaminated samples that are representative of the sediment at a given depth. Large rock fragments or concretions on the sediment surface may be

collected with a chain dredge or similar device dragged over the sea bottom in a desired area

(Figure 7.7).


If samples cannot be examined immediately after collection, they should be stored so as to minimize microbial multiplication or loss of viability. Cooling a sample is usually the best way to

preserve it temporarily in its native state, but the extent of cooling may be critical. Freezing

may be destructive to at least some of the microbes. However, icing may not prevent growth of

psychrophiles or psychrotrophs. The duration of storage before examination should not be longer

than absolutely necessary.

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FIGURE 7.5 Gravity corer. This is simply a hollow pipe containing a removable plastic liner and having a

cutting edge at the lower end with a core catcherr to retain the sediment core when the device is pulled out of

the sediment. A heavy lead weight at the top helps to ram the corer into the sediment when allowed to free-fall

just above the sediment surface.

FIGURE 7.6 Box corer. After the frame hits bottom, the coring device is forced into the sediment mechanically. (Courtesy of Sand M)

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FIGURE 7.7 Seafloor samplers. (A) Chain dredge. (B) Dredge for collecting manganese nodules from the

ocean floor. A conical bag of nylon netting is attached to a pyramidal frame.



To study agents active in a geomicrobial process of interest, culture enrichments and pure culture isolations should be attempted as far as possible. Not all cultures obtained by these procedures may be geomicrobially active. Each isolate must be tested for its ability to perform the

particular geomicrobial activity, or a part of it, that is under investigation. Because some geomicrobial processes are the result of the activity of a microbial consortium, no one of the pure

culture isolates may exhibit all of the desired activity but may have to be tested with others of

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the isolates in different combinations. Examples of geomicrobial cooperation among microorganisms in microbial manganese oxidation that have been described include (1) the bacterium

Metallogenium symbioticum in association with the fungus Coniothyrium carpaticum (Zavarzin,

1961; Dubinina, 1970; but see also, Schweisfurth, 1969); (2) the bacteria Corynebacterium sp.

and Chromobacterium sp. (Bromfield and Skerman, 1950; Bromfield, 1956); and (3) two strains

of Pseudomonas (Zavarzin, 1962). More recently, it was discovered that anaerobic methane oxidation depended on the action of a consortium consisting of a methanogen and a sulfate reducer

(e.g., Boetius et al., 2000) (for more detail see Chapter 22).

Enrichment of and isolation from a mixed culture require selective conditions. If a microbial

agent with a specific geomicrobial attribute is sought, the selective culture medium should have

ingredients incorporated that favor the geomicrobial activity of interest. Apart from special nutrients, special pH, Eh, and temperature conditions may also have to be chosen to favor selective

growth of the geomicrobial agents.

Isolation and characterization of pure cultures from enrichments should follow standard bacteriological technique, including determination of the molecular phylogeny of the agent(s) (for details,

see for instance, Gerhardt et al., 1981, 1993; Hurst et al., 1997; and Skerman, 1967).




Past in situ geomicrobial activity in sedimentary deposits that ceased as far back as the early

Precambrian or during the Phanerozoic up to modern times can sometimes be inferred through the

detection of specific organic biomarkers in samples from such deposits. These biomarkers represent

preserved organic derivatives formed from some characteristic cellular constituent, for example,

chlorophylls from photosynthetic prokaryotes (bacteria, cyanobacteria); phototrophic eukaryotic

microbes (Brocks and Pearson, 2005; Brocks et al., 2005; Roselle-Melộ and Koỗ, 1997); carotenoids

of anoxygenic photosynthetic bacteria and the oxygenic cyanobacteria (Hebting et al., 2006); and

lipid cell membrane components from Archaea or Bacteria (Brocks and Pearson, 2005; Hebting

et al., 2006). The process by which these cell constituents were preserved appears to have involved

mostly or exclusively chemical reduction (abiotic) under anaerobic conditions. In situ, the agents of

preservation in the case of preservation of carotenoids such as β- (phytoplankton) and γ-carotene

(cyanobacteria and green nonsulfur bacteria, i.e., Chloroflexaceae), and okenone (purple sulfur

bacteria, i.e., Chromatiaceae appear to have been reducing agents in the form of reduced sulfur

compounds such as H2S. The corresponding products of preservation from these pigments were

β- and γ-carotane and okenane or perhydrookenone (Brocks et al., 2005; Hebting et al., 2006). The

detection of the biomarker okenane, derived naturally from okenane of photosynthetic purple nonsulfur bacteria (Bradyrhizobiaceae) and chlorobactane from green sulfur bacteria (Chlorobiacea) in

sediments of the 1.64-Gyr-old Barney Creek Formation (BCF) of the McArthur Group in northern

Australia, has been used to argue that oxygen levels at this site at that time were well below modern

levels although such levels had been approached or reached elsewhere on Earth (Brocks et al., 2005).

This interpretation was supported by the simultaneous finding of extremely low concentrations of

2α-methylhopanes, biomarkers from cyanobacteria at the BCF site, which are normally considered

to be oxygenic photosynthesizers (Brocks et al., 2005). However, Rashby et al. (2007) have demonstrated that a contemporary strain of the phototroph Rhodopseudomonas palustris (purple nonsulfur bacteria) is capable of synthesizing 2-methylbacteriohopanepolyols from which 2-methylhopane

biomarkers could be formed during fossilization, indicating that this type of biomarker may not be

a unique indicator for cyanobacteria.

Under certain circumstances, the occurrence of past geomicrobial activity can also be identified in terms of isotopic fractionation. Certain prokaryotic and eukaryotic microbes have been

shown to distinguish between stable isotopes of elements such as C, H, O, N, S, Si, Li, and Fe.

These microbes prefer to metabolize molecules containing the lighter isotopes of these elements (12C

in preference to 13C, H in preference to D, 16O in preference to 18O, 14N in preference to 15N, 32S in

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preference to 34S, 28Si in preference to 30Si, 6Li in preference to 7Li, 54Fe in preference to 56Fe), especially under conditions of slow growth (see Jones and Starkey, 1957; Emiliani et al., 1978; Mortimer

and Coleman, 1997; Mandernack et al., 1999; Wellman et al., 1968; Estep and Hoering, 1980; De

La Rocha et al., 2000; Sakaguchi and Tomita, 2000; Croal et al., 2004; Crosby et al., 2007; Beard

et al., 1999). They distinguish kinetically between different stable isotopes of the same element in a

substrate on the basis of a difference in the rate of reaction the substrate undergoes in a specific biochemical step, the reaction rate involving the substrate with the lighter isotope being faster than that

with heavier stable isotope. The isotope fractionation may occur during a specific enzyme-catalyzed

chemical reaction into which the isotope-containing substrate enters, or it may occur during membrane transport of the isotope-containing substrate, or both (see Hoefs, 1997). Thus, the products of

metabolism will be enriched in the lighter isotope compared to the starting compound or to some

reference standard that has not been subjected to isotope fractionation. In practice, isotope fractionation is measured by determining isotope ratios of the heavier isotope of an element to the lighter,

using mass spectrometry, and then calculating the amount of enrichment from the relationship

Isotope ( ‰) ϭ

isotopic ratio of sample Ϫ isotope ratio of standard


isotopic ratio of standard

If the enrichment value (δ) is negative, the sample tested was enriched in the lighter isotope relative

to a reference standard, and if the value is positive, the sample tested was enriched in the heavier

isotope relative to a reference standard. Thus, for instance, to determine if a certain metal sulfide

mineral deposit is of biogenic origin, various parts of the deposit are sampled and δ34S values of

the sulfide are determined. If the values are generally negative (although the magnitude of the δ34S

may vary among the samples and fall in the range of –5 to –50‰), the deposit can be viewed as of

biogenic origin because a chemical explanation for such 32S enrichment under natural conditions is

not likely. If the δ34S values are positive and fall in a narrow range, the deposit is viewed as being

abiogenic in origin.



Ongoing geomicrobial activity may be measurable in situ. Such activity may be followed by the use

of radioisotopes. For instance, bacterial sulfate-reducing activity may be determined by adding a

small quantity of Na35SO4 to water, soil, or sediment sample of known sulfate content in a closed

vessel. After incubation under in situ conditions, the sample is analyzed for loss of 35SO42– and

buildup of 35S2– by separating these two entities and measuring their quantity in terms of their radioactivity. In the case of a water sample, incubation of the reaction mixture in a closed vessel in the

water column may be at the depth from which the sample was taken. An example of a direct application of this method is that of Ivanov (1968). It allows the estimation of the rate of sulfate reduction

in the sample without having any knowledge of the number of physiologically active organisms

present in it. A modified method is that of Sand et al. (1975). Their method allows an estimation of

the sulfate-reducing activity in terms of the number of physiologically active bacteria in the sample

as distinct from an estimation of the sum of physiologically active and inactive bacteria. The assay

for the estimation of active bacteria can be set up either to measure percentage of sulfate reduced in

a fixed amount of time that is proportional to the logarithm of the concentration of active cells, or to

measure the length of time required to reduce a fixed amount of sulfate to sulfide, which is related to

the concentration of physiologically active sulfate-reducing bacteria in the sample. Ivanov’s (1968,

pp. 30–32) method can be adapted to measure the formation of elemental sulfur and sulfates from

sulfide by adding 35S2– to a sample and after incubation in a reaction vessel in situ, separating 35S

and 35SO42– and measuring their quantity in terms of their radioactivity.

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Microbial action on manganese (Mn2+ fixation by biomass; Mn2+ oxidation) in situ is another

example of a geomicrobial process, which can be followed by the use of a radioisotope, 54Mn2+

in this case (LaRock, 1969; Emerson et al., 1982; Burdige and Kepkay, 1983). One approach is to

measure manganese oxidation in terms of the decrease in dissolved 54Mn2+. It assumes that the

oxidized manganese is insoluble. Decreases in dissolved 54Mn2+ are measured on acidified samples

of the reaction mixture from which the oxidized manganese has been removed by centrifugation.

Acidification of samples before centrifugation ensures resolubilization of any adsorbed Mn2+. The

difference in radioactivity counts between a zero-time sample and a sample taken at a subsequent

time is a measure of the amount of Mn2+ oxidized by a combination of biological and chemical processes over this time interval. Manganese oxidation due to biological activity alone can be estimated

by subtracting chemical oxidation from the total Mn2+ oxidation for a corresponding time interval.

An estimate of the amount of chemical oxidation can be obtained from a separate reaction mixture

in which the biological activity is inhibited by use of autoclaved instead of active cells, by adding

one or more chemical inhibitors to the oxidation mixture containing active cells, or by excluding

air (if the enzymatic manganese oxidation is oxygen-dependent). This experimental approach has

been used to assess the manganese-oxidizing activity in lake sediment (LaRock, 1969). It makes no

assumption about binding of unoxidized or oxidized manganese to the bacterial cells.

Another approach is to measure manganese oxidation in terms of product formation. Manganese

binding by metabolically active bacteria in a water sample may be followed in terms of 54Mn accumulation by the cells. A manganese-oxidizing culture is incubated in a suitable reaction mixture

containing added 54Mn2+. After an appropriate length of incubation, the amount of radiomanganese

bound by the cells is determined. For this determination, a measured sample of the bacterial suspension is filtered through a 0.2 µm membrane and washed, and the radioactivity retained on the filter

membrane (assumed to be bound to the cells) is determined on a suitable counter. The results of

this experiment are then compared to those of a parallel experiment in which the cells were inhibited by a poison such as formaldehyde or a mixture of sodium azide, penicillin G, and tetracycline

HCl. The difference in radioactivity between these two experiments represents manganese bound

by actively metabolizing cells. It includes manganese that was bound as Mn2+, presumably a very

minor amount, and that which was bound due to its oxidation by the cells (Emerson et al., 1982).

Manganese binding by bacteria in sediment can be followed by a modified form of this method performed in a special reaction vessel called a peeper (Burdige and Kepkay, 1983). 54Mn2+ adsorbed by

the cells deposited on a filter membrane is displaced by washing with CuSO4 solution. The radioactivity recovered in the wash is then counted. Residual 54Mn associated with the cells (taken as oxidized manganese) is dissolved by washing with hydroxylamine-HCl solution followed by washing

with CuSO4 solution, and the radioactivity in the resultant solution is determined. HydroxylamineHCl is a reducing agent, which converts Mn(IV) to Mn(II).

Biological manganese binding in lake sediments may be studied by a method that requires controls in which biological inhibitors are used to account for abiological manganese binding by sediment constituents in the overall manganese budget (Burdige and Kepkay, 1983).

One advantage of using radioisotopes in the quantitative assessment of a specific geomicrobial

transformation in nature is that their detection is extremely sensitive so that only minute amounts of

radiolabeled substrate, which do not significantly change the naturally occurring concentration of

the substrate, need to be added. Another advantage is that in cases where the rate of transformation

is very slow although the natural substrate concentration is high, spiking the reaction with radiolabeled substrate allows analysis after a relatively brief reaction time because of the sensitivity of

radioisotope detection.

The use of radioisotopes is, however, not essential for quantitative assessment in all instances of

biogeochemical transformation in a natural environment. Other analytical methods with sufficient

sensitivity may be applicable (see, for instance, Jones et al., 1983, 1984; Hornor, 1984; Kieft and

Caldwell, 1984; Tuovila and LaRock, 1987).

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To fully appreciate in situ geomicrobial activity, knowledge of local vertical chemical, pH, and

redox gradients over relatively narrow depth intervals (e.g., millimeters or less) is important. The

application of specific microelectrodes has made such determinations possible, as the following

examples, a few among many, show.

Thus, use of an Au/Hg voltammetric microelectrode made it possible to measure simultaneously and with a spatial resolution in millimeters the vertical three-dimensional distribution of

O2, Mn2+, Fe2+, HS−, and I− in pore water of undisturbed sediment cores from the Canadian continental shelf and slope, and in the sediment surrounding an actively irrigated worm burrow in

a mesocosm (Luther et al., 1998). Solid-state gold amalgam voltammetric microelectrodes have

also made the monitoring of sulfur speciation in situ in sediments, microbial mats, and hydrothermal vent waters possible (Luther et al., 2001). Use of a fiber-optic scalar irradiance microsensor

and oxygen microelectrode spaced 120 µm apart made it possible to measure scalar irradiance

and oxygenic photosynthesis with 100 µm spatial resolution in marine microbial mats dominated

by cyanobacteria with a surface layer of pinnate diatoms on sandy sediment along the coast of

Limfjorden, Denmark (Lassen et al., 1992). Microelectrodes have also been employed in measuring O2, H2S, and pH microgradients at depth increments of 50 µm in Beggiatoa mats from

marine sediments and Thiovulum films above the sediment to determine the microbial response to

O2 and H2S (Jørgensen and Revsbech, 1983). Freshly prepared nitrate-selective microelectrodes

with a liquid ion exchanger have been employed in determining nitrate gradients in sediments from

a mesotrophic lake in the Netherlands (De Beer and Swearts, 1989). Although all the examples

cited here involve sediments in freshwater and marine systems, microelectrodes are equally useful

in studying terrestrial systems (e.g., soil).




It is often important to reconstruct a naturally occurring geomicrobial process in the laboratory

to investigate the mechanism whereby the process operates. Laboratory reconstruction can permit optimization of a process through the application of more favorable conditions than in nature.

Examples are the use of a pure culture or a purifiedd consortium to eliminate interference by competing microorganisms and the optimization of substrate availability, temperature, pH, Eh, and oxygen

and carbon dioxide supply.

The activity of organisms growing on the surface of solid substrates such as soils, sediments,

rocks, and ore may be investigated in batch culture, in air-lift columns, in percolation columns, or

in a chemostat. A batch culture represents a closed system in which an experiment is started with a

finite amount of substrate that is continually depleted during the growth of the culture. Cell population and metabolic products buildup and changes in pH and Eh are likely to occur. Conditions

within the culture are thus continually changing and becoming progressively less favorable. Batch

experiments may be least representative of a natural process, which usually occurs in an open system with continual or intermittent replenishment of substrate and removal of at least some of the

metabolic wastes. A culture in an air-lift column (Figure 7.8) is a partially open system, where the

microbes grow and carry on their biogeochemical activity on a mineral charge in the column. They

are continually fed with recirculated nutrient solution from which nutrients are depleted by the

organisms. The recirculation removes metabolic products from the solid substrate charge in the column. Percolation columns (Figure 7.9) are even more open systems than air-lift columns. In them,

microbes grow on the solid substrate charge in the columns, but they are fed with nutrient solution

that is not recirculated. This fresh nutrient solution is added continually or at intervals, and wastes

are removed at the same time in the effluent without recirculation, while pH, Eh, and temperature

are held constant or nearly so. Steady-state conditions such as in a chemostatt idealize the open

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Chapter 7. Nonmolecular Methods in Geomicrobiology

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