MBT_Anaerobic Cultivation Protocols-complete

MOLECULAR BIOLOGY TOOLS (MBT)

WORKSHOP

Anaerobic Cultivation Techniques

Braga, University of Minho

23-24 June 2013

http://www.ad13.org/

Supported by:

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“Everything is everywhere, but, the environment selects.” Baas Becking (1934)

Lourens Baas Becking Martinus Beijerinck

CONTENTS:

1. ENRICHMENT CULTURES 1

2. MINERAL SALTS MEDIUM AND SOLUTIONS 2

3. ISOLATION OF PURE CULTURES 5

4. PURITY CHECK 6

5. ANNEX – VALUABLE LITERATURE EXAMPLES 7

Anaerobic Cultivation Techniques | 1

MBT Workshop, 23-24 June

University of Minho, Braga, Portugal

Some guidelines

1. Enrichment cultures

A definition of enrichment cultures is given by T. Brock in his Textbook Brock,

Biology of Microorganisms

‘‘Use of selective culture media and incubation conditions to isolate

microorganisms directly from nature.’’

Before starting an enrichment culture it is important to think well about (i) the

medium composition, including the selection of the electron donor and electron

acceptor, and (ii) the environmental conditions (such as, salinity, pH, incubation

temperature,...) to supply to the culture. All these factors will constrain the

development of the enrichment culture and eventually the isolates obtained from

this culture. In annex we provide documentation with useful information for

anaerobic microorganisms cultivation using different substrates, electron

acceptors and environmental conditions.

1. Prepare a mineral salts medium that contains the major and trace elements

that are necessary for the growth of a variety of bacteria. Adjust the pH of the

medium. To make the medium anaerobic, boil it and cool it down under a N2

stream. Dispense the medium into serum bottles under a N2 stream and close

them with butyl rubber stoppers and aluminium crimps. Flush the bottles’ headspace with O2-free gas (e.g. N2, N2/CO2, H2/CO2 – depending on the purpose

of the enrichment). Sterilize the medium by autoclaving it at 120ºC for 20 min.

2. Prepare stock solutions to supplement the medium – carbon source, electron

acceptor, salts, trace elements, vitamins, etc. Sterilize stock solutions either by

filtration or autoclaving, depending on the thermal stability and reactivity of the

substance. Note: The optimal concentration of the carbon source depends on its

solubility, availability and toxicity. One advantage of a high carbon concentration is the

high yield in cell mass but the growth of many oligotrophic strains might be suppressed

under such conditions.

3. Get the inoculum from the environment that you want to study, e.g.

(contaminated) soil, anaerobic sludge, mud, decaying vegetation or sediments.

Inoculate the medium with 1–5% (v/v) of the inoculum. Note: The inoculum

volume is merely indicative.

4. Incubate the flasks in the dark at suitable temperature. Incubation can be

performed with or without shaking; when utilizing gaseous carbon sources

shaking enhances mass transfer and normally favours growth.




5. Analyse growth by optical density measurement, if applicable. Alternatively

you can use microscopic techniques, flow cytometry or cell protein quantification.

6. Measure the degradation of the supplemented carbon source and/or formation

of metabolites by suitable methods such as GC, HPLC or spectrophotometry. In

cases where an alternative electron acceptor was added its depletion as well as

the formed reduced compound should be quantified.

7. Once the cultures are grown and the substrate depleted, inoculate a bottle

containing fresh medium with 1–10% (v/v) of the previous culture.

8. Repeat steps 5 to 7 several times until the growth and degradation rates do

not increase anymore and stable lineages are established.

An example of appropriate mineral salts medium and solutions for cultivation of

anaerobic microorganisms at neutral pH is described in the ‘‘Mineral Salts

Medium and Solutions’’ section below.

2. Mineral Salts Medium and Solutions

Basal Medium

Mix the following components/solutions in a 2L Erlenmeyer:

COMPONENT/SOLUTION QUANTITY

Demi Water 900 mL

Solution 1 (KH2PO4) 15 mL

Solution 2 (Na2HPO4.H2O) 15 mL

Solution 6 (trace elements H+) 1 mL

Solution 7 (trace elements OH-) 1 mL

Solution 8 (resazurine) 1 mL

Boil the medium in order to remove the dissolved oxygen. Cool it down in ice

under N2.

Dispense the basal medium into serum bottles (Note: leave 60% of the bottle

volume as gas phase!) and closed them with black butyl rubber stoppers and

aluminium caps. The gas phase (air) is replaced by a gas mixture of N2/CO2

(80/20 % v/v) or H2/CO2 (80/20 % v/v), to an absolute final pressure of 1.75

bar. Sterilise the bottles with the medium by autoclaving at 120C for 20 min.




Before inoculation, the basal medium has to be supplemented with 5%

bicarbonate solution (containing the reducing agent) and 5% salts solution

(containing the vitamins). Carbon and energy sources or alternative electron

acceptors can also be added to the medium from previously prepared anoxic

stock solutions.

Bicarbonate solution:


Solution 4 (NaHCO3) 100 mL

Solution 5 (Na2S.9 H2O) 2 mL

Flush the headspace with N2, pressurize at absolute pressure of 1.75 bar, and

then sterilize at 120 C for 20 min.

Note: Under certain circumstances better growth is obtained if the “reducing solution” (solution 5) is filter sterilized and added to the bicarbonate solution after autoclaving (this happens because H2S can react with the butyl rubber

stoppers at high temperatures).

Salts and vitamins solution:


Demi Water 75 mL

Solution 3 (salts solution) 25 mL

Solution 9 (vitamins solution)

(add filter sterile after

autoclaving)

2 mL

Dilute solution 3 in demi water (as indicated in the table), flush the headspace

with N2, pressurize at absolute pressure of 1.75 bar, and then sterilize at 120 C

for 20 min. After autoclaving add 2 mL of the vitamins solution (filter sterile).

Stock solutions:

Stock solutions to be used to "compose" the basal medium (grams/milligrams are

given for dilution in 1L):

Solution 1: 27.2 g KH2PO4

Solution 2: 35.6 g Na2HPO4.2H2O




Solution 3: 24 g NH4Cl; 24 g NaCl; 8 g MgCl2.6H2O: 8.8 g CaCl2.2H2O

Solution 4: 80 g NaHCO3

Solution 5: 240.2 g Na2S.9 H2O (store under N2 and in the dark)

Solution 6: Trace elements - Acid Stock Solution

50 mM HCl 1.8 g

1 mM H3BO3 61.8 mg

0.5 mM MnCl2 61.25 mg

7.5 mM FeCl2 943.5 mg

0.5 mM CoCl2 64.5 mg

0.1 mM NiCl2 12.86 mg

0.5 mM ZnCl2 67.7 mg

Solution 7: Trace elements - Alkaline Stock Solution

10 mM NaOH 400 mg

0.1 mM Na2SeO3 17.3 mg

0.1 mM Na2WO4 29.4 mg

0.1 mM Na2MoO4 20.5 mg

Solution 8: 0.5 g Resazurin

Solution 9: Vitamins

Biotin 20 mg

Nicotinamid 200 mg

p-Aminobenzoic acid 100 mg

Thiamin (Vit B1) 200 mg

Panthotenic acid 100 mg

Pyridoxamine 500 mg

Cyanocobalamine (Vit B12) 100 mg

Riboflavin 100 mg

(Folate 50 mg)

(Lipoate 50 mg)

P-aminobenzoic acid is dissolved in 120 ml of demi water by addition of NaOH. In

order to dissolve riboflavin add to 200 ml of demi water and heat in a waterbath

(50 C, avoid boiling!!) until completely dissolved. This solution is then mixed

with the solution of p-aminobenzoic acid. The other components should be added

one by one and only upon complete dissolving the previous one. The pH is finally

adjusted to 7 and the volume to 1 litre. Filter sterile the solution to empty sterile

bottles containing N2.




3. Isolation of pure cultures

Working with pure isolates instead of (enriched) consortia provides more defined

conditions for further biochemical and genetic experiments and helps to assign

physiological properties to special microbial taxa.

Each microorganism will ask for a specific treat for isolation. Normally you need

to try several approaches for successfully isolating a new bug.

Some common techniques used for isolation of anaerobes:

- Plating techniques. Plating provides a simple way to isolate pure strains of

anaerobes, though one needs to be aware of their limitations. Anaerobic plating

needs to be performed in the anaerobic chamber, which makes handling of the

cultures more difficult and where aseptic conditions are not so easily kept.

Prepare solid medium using the same recipe as for the medium used for

enrichment but supplement it with 2% agar.

- Roll tubes and soft-agar bottles. In order to work outside of the anaerobic

chamber, one can prepare solid medium in roll tubes or serum bottles. Use a

slightly lower amount of agar than in plates, approximately 1.5%.

- Dilution series using liquid medium. If the microorganism that you want to

isolate is in predominant amount in your enriched culture, you might isolate it by

serially diluting your enrichment in liquid medium.

- Utilization of antibiotics or toxic compounds. If you have an enriched culture

and you know that the main contaminant(s) is susceptible to a certain antibiotic

and/or substance, you can always try to eliminate it by adding this substance to

the medium.

- Heat treatment. In case the microorganisms you want to isolate is able to form

spores, it might be worthy to heat treat your culture in order to eliminate heat-

sensitive contaminants.

Well, these are just some tips. Just be original, and try to follow each small hint

given by your own cultures in order to get you isolate. Good luck!




4. Purity check

Culture purity check can be done by combining different techniques, as for

example (but not only):

- microscopic observation for morphology check; gram staining can also aid in

the detection of possible contaminants;

- molecular diversity profiling, such as DGGE (attention, some microorganisms

have more than one 16S rRNA gene copy, which will result in several bands in a

DGGE – and yet can be a pure culture);

- clone library and sequencing of several 16S rRNA gene clones;

- sub-culturing in different media (including rich media) in order to evaluate if

other microorganisms become predominant;

- colony morphology, when culturing in solid medium;

- design of specific FISH probes or real-time PCR primers.




5. Annex – Valuable literature examples

Anaerobic Cultivation Techniques |8






5.1. DSMZ media

http://www.dsmz.de/catalogues/catalogue-microorganisms/culture-

technology/list-of-media-for-microorganisms.html

http://www.dsmz.de/catalogues/catalogue-microorganisms/culture-technology/list-of-media-for-microorganisms.html

http://www.dsmz.de/catalogues/catalogue-microorganisms/culture-technology/list-of-media-for-microorganisms.html







5.2. DSMZ special instructions for anaerobic cultivation

Cultivation of anaerobes

Cultivation of methanogens

Cultivation of acidophiles

Cultivation of hyperthermophiles




Special Instructions:

Cultivation of Anaerobes

The DSMZ holds a large collection of prokaryotes that thrive only under anaerobic conditions. In our experience beginners in culturing anaerobes or extremophiles encounter often difficulties in handling these cultures appropriately. This technical information should help everybody who is interested to start working with anaerobes. Please read it carefully, it will answer most frequently asked questions about culturing anaerobes!

You will find on this page information to the following topics:

General information about anaerobes

Recommended vials for culturing strict anaerobes

Gassing of media and cultures with oxygen-free gas

Handling of vacuum-dried anaerobic cultures

Handling of actively growing anaerobic cultures

Reducing agents and resazurin

Literature

Notes

General information about anaerobes

In the broadest sense obligate anaerobes can be defined as microorganisms which are unable to utilize molecular oxygen for growth. A further differentiation is possible based on their relationship to the presence of oxygen. Aerotolerant anaerobes are only slightly inhibited by significant levels of oxygen in the atmosphere. For instance Clostridium intestinale DSM 6191

T can grow well on the surface of agar plates incubated in air at

atmospheric pressure. The other extreme is represented by strict anaerobes, which die, or immediately stop growing, upon exposure to low levels of oxygen. It is therefore important to retain anoxic conditions during all steps of handling of these microorganisms. Most strict anaerobes require not only the absence of oxygen to initiate growth, but also a redox potential below -300 mV, which can be only achieved by the supplementation of media with reducing agents (see section on Reducing agents and resazurin). Between both extremes all kinds of adaptation exist.

The majority of anaerobic microorganisms are fastidious and require complex media with many supplements. In the DSMZ catalogue of strains (Internet: http//www.dsmz.de/species) each DSM strain is linked with a specific medium number. It is strongly recommended to use the respective media formulations given for each strain because only those media were tested at the DSMZ for culturing and a transfer to alternative media may cause a delay or complete failure of growth. Before ordering an anaerobe from the DSMZ it is advisable to have a look on the recipe of the medium necessary for growing this strain and to read relevant publications dealing with its cultivation.

It makes only sense to purchase a strain of interest, if you are convinced to be able to handle it properly!

A large number of strict anaerobes are available from the DSMZ only as actively growing cultures. We recommend to use the Hungate technique to culture these strains. Some general remarks on this cultivation technique and required laboratory equipment follow below. Excellent descriptions of the Hungate technique can be found in the reviews of Hungate (1969) and Wolfe (1971), whereas the contribution of Breznak and Costilow (1994) contains more general information on anaerobiosis. However, please keep in mind, that even if described in detail, some steps of the handling of anaerobic cultures are frequently difficult to master without demonstration. For beginners in anaerobic microbiology it is therefore the best to visit a laboratory where anaerobic cultivation techniques are routinely in use.

Anaerobic strains that are available from the DSMZ as lyophilized cultures are normally not sensitive to a short exposure to low oxygen concentrations (nonstringent anaerobes). For instance, a majority of the clostridia and sulfate reducers, but not all of them, belong to this group of strains. If you have received an ampoule from the DSMZ with a vacuum dried sample of a nonstringent anaerobe please read also the instructions given in the section: Handling of vacuum dried anaerobic cultures.

Further technical information on difficult to handle microorganisms, like methanogens or hyperthermophiles are available at the DSMZ web pages and linked with the catalogue entries of the respective strains.

Recommended vials for culturing strict anaerobes

Suitable containers for pre-reduced media are an important prerequisite for the successful culturing of strict anaerobes. For this purpose special glassware has been developed which enables the easy use of completely gas-tight closures. Of crucial importance is the material of the rubber stoppers. Only stoppers or septa made of butyl rubber can efficiently prevent permeation of air into the vial. Nevertheless, a repeated puncturing of stoppers with injection needles could make them become permeable to oxygen. As a rule, the thicker the stopper the more often it is possible to reuse it without loss of impermeability.

Two types of vials are commercially available for anaerobic culturing (Fig. 1):

The Hungate-type tubes are closed with a flange-type butyl rubber septum and a screw cap with 9 mm opening to allow puncturing of the septum with injection needles.

Balch-type tubes are more stable than Hungate-type tubes and recommended if an overpressure of 2 to 3 bar can be expected during culturing. They are closed with a thick butyl rubber stopper which is hold in place by sealing with an aluminum crimp. For sealing and removing of the aluminum crimp special devices (crimper/decapper) are necessary. Serum bottles which are available in various sizes can be used alternatively to Balch-type tubes. However, serum bottles are less stable than Blach tubes and should be handled with special care when strains are cultured that are expected to produce significant amounts of gas during incubation (see below).

Pre-reduced media can be stored in both types of vials at room temperature in the dark for several weeks without becoming oxidized.

Caution: Some microorganisms produce a considerable amount of gas during growth (e. g., Clostridia by fermentation). The formed gas can lead to a substantial overpressure during growth in closed vials. Strains which are known to accumulate gas during growth should be incubated in vessels that are filled only up to 25% with liquid medium. In addition, cultures of fast growing strains should be vented at least on a daily basis to avoid overpressure. Wear protective goggles during handling of glass vessels that might have overpressure!

Suppliers of commercially available glassware and accessories for anaerobic culturing are for instance Bellco Glass Inc. (http://www.bellcoglass.com/) and Ochs GmbH (http://www.glasgeraetebau-ochs.de/).

Fig. 1 Suitable vials for culturing strict anaerobes. (A) Hungate-type tube with screw cap and butyl rubber septum. (B) Balch-type tube with butyl rubber stopper and aluminum crimp seal to hold stopper in place. A crimper is necessary for sealing the vial. Figures are courtesy of Bellco glass Inc.

Gassing of media and cultures with oxygen-free gas

When vials of pre-reduced media or anaerobic cultures are opened a constant flow of oxygen-free gas over the surface of the medium is necessary to avoid exposure to oxygen. The used oxygen-free gas should have the same composition as that used for medium preparation. We recommend to use oxygen-free gasses of high purity (containing less than 5 ppm oxygen), that are delivered as compressed gas cylinders. Normally, oxygen-free gasses of high quality do not require an additional oxygen removal system (e. g., heated copper column) and can be used directly for culturing a broad spectrum of anaerobes. Suppliers of compressed gasses are for instance Messer Griesheim GmbH (http://messergroup.com) or Linde AG (http://www.linde.com).

The Hungate technique is based on the use of Gassing cannulas. Usually, several cannulas are connected by butyl rubber tubing to a manifold supplying oxygen-free gas with an overpressure that should be adjusted to approx. 0.5 bar. At least two cannulas are needed: one for the vessel to be inoculated or filled with medium and one for the vessel containing the inoculum or the medium to be dispensed. When an aseptic gassing of media or cultures is necessary a barrel of a glass syringe is packed with cotton and fitted between the gassing needle and the butyl rubber tubing (Fig. 2A).

Fig. 2 Assembly of cannulas used in the Hungate technique for aseptic gassing. (A) Cannula used for aseptic gassing of opened vials with oxygen-free gas. (B) Overpressurizing of anaerobic cultures with sterile gas mixtures.

After assembly, autoclave the cotton-filled glass syringe and needle, dry in a drying oven at 100 °C, allow to cool, and connect to the manifold. Prior to the first use flush the gassing cannula for approx. 15 min with oxygen-free gas to make it anoxic and then flame the needle to sterilize it.

Caution: Make sure that needles sterilized by flaming are cooled down prior to using gas mixtures containing H2. Hydrogen gas is highly combustible, and even only contact with hot surfaces may cause ignition. Wear protective goggles while overpressurizing vials.

For the overpressurizing of cultures with H2 or H2/CO2 gas mixtures use disposable, sterile injection needles (i. d. 0.4 mm or 27G) connected to cotton-filled glass syringes as described above. To keep the pressure within the glas syringe barrel at a constant level during overpressurizing it is necessary to avoid an imbalance between the inflowing gas stream and the outflow. This can be achieved by puncturing the rubber stopper of the cotton-filled syringe with a steel needle (approx. 20G) which is connected to the rubber tubing by an appropriate valve with Luer-Lock fittings. Adjust the gas pressure to the desired value (in most cases 0.5 to 2 bar overpressure). Turn the vial with the culture up side down and puncture the sterilized septum with the injection needle (Fig 2B). A sputtering of gas bubbles indicates the inflow of gas into the medium and can be observed as soon as the tip of the cannula enters the liquid. When the flow of bubbles slows down the pressure within the vial reaches equilibrium with the external pressure of the gas supply. Withdraw gassing needle immediately when the gas flow stops.

Handling of vacuum-dried anaerobic cultures

The DSMZ delivers lyophilized (freeze-dried) cultures of anaerobic strains exclusively in double-vial preparations, heat-sealed under vacuum. Double-vial preparations have the advantage that a contamination of the atmosphere by aerosols that can be produced by sudden release of the vacuum in single-vial preparations is efficiently prevented. In addition, the cell pellet is protected from contamination, because inflowing air filters through the sterile cotton plug of the inner vial. Before opening of the ampoule please identify the culture by the label on the inner vial which indicates the DSM strain number and date of preservation. Confirm that the ampoule is under vacuum by checking the color of the desiccant at the bottom of the outer vial.

Note: The DSMZ has changed the indicator stain of the desiccant. From January 2002 on the used desiccant is red, instead of blue. The indicator stain will change its color if the outer vial is damaged due to an increase of humidity inside the ampoule. The red color changes to orange and the blue to pink.

It is important to retain anoxic conditions during all steps after the opening of ampoules with freeze-dried anaerobes. This can be achieved in several ways depending on the used anaerobic technique and available equipment in the laboratory. For general information about vacuum dried cultures please visit the following site Opening of ampoules and rehydration of dried cultures.

The freeze-dried pellet of most anaerobic strains available from the DSMZ is protected against short exposure to oxygen by amorphous ferrous sulfide (FeS) which confers a black color to the pellet. However, certain nonstringent or spore forming anaerobes are suspended prior to lyophilization in skim milk without addition of FeS, so that the ampoules display a white pellet.

If the Hungate technique is routinely used in the laboratory, open the ampoules as described in Opening of ampoules and rehydration of dried cultures. After opening keep the inner vial under a flow of oxygen-free gas by inserting a gassing cannula. Add approx. 0.5 ml of the recommended anoxic medium to the vial and resuspend the cell pellet completely (in some cases this may take several minutes). Transfer the cell suspension either by using a 1 ml syringe with hypodermic needle (length at least 38 mm) or a sterile Pasteur pipette, which was made anoxic by flushing with oxygen-free gas. If a Pasteur pipette is used, the Hungate tube containing the appropriate cultivation medium (5 to 10 ml) has to be opened and gassed with a second cannula during transfer of the inoculum.

If an anaerobic gas chamber is available it is recommended to score the ampoule with a sharp file at the middle of its shoulder about one cm from the tip. Transfer the ampoule with the file mark in the anaerobic chamber and strike the ampoule with a file or large forceps to remove the tip. If necessary, wrap the ampoule in tissue paper and enlarge the open end by striking with a file or pencil, then remove the glass wool insulation and the inner vial. Gently raise the cotton plug and sterilize the upper part of the inner vial using an incandescent flaming device (alternatively wipe the upper part of the inner vial with tissue paper soaked in 70% ethanol). Add approx. 0.5 ml of anoxic medium to resuspend the cell pellet and transfer the suspension to a vial with the recommended cultivation medium (5 to 10 ml).

If possible the last few drops of the resuspended cell pellet should be transferred to an agar plate or slant of the recommended medium to obtain single colonies in order to check the purity of the strain. Anaerobic incubation conditions for agar plates can be achieved by placing plates in an anaerobic chamber or an activated anaerobic Gas Pak jar or similar system (e. g., Anaerocult

® bags available from Merck; http://www.merck.de).

We recommend to prepare also 1:10 and 1:100 dilutions of the inoculated medium, because some ingredients of the freeze-dried pellet may inhibit growth in the first tube. Inoculation of only one tube may prevent successful resuscitation of certain lyophilized strains (e.g., Geobacter spp.).

In most cases freeze dried cultures of anaerobic strains exhibit a prolonged lag period upon rehydration and should be given at least twice the normal incubation time before regarding them as non-viable.

Handling of actively growing anaerobic cultures

For the aseptical injection and removal of samples from anaerobic cultures it is recommended to use the Hungate technique which is essentially based on the use of disposable syringes and has the advantage that it allows the use of defined, oxygen-free atmospheres for culturing. The anoxic removal of a sample from an Hungate tube is demonstrated in Fig 3.

First, sterilize the butyl rubber septum by flaming it using a drop of ethanol which has been placed on the septum. If overpressure in the vial can be expected due to microbial growth (e.g., gas production by fermentation) remove excess gas by puncturing the septum with a sterile injection needle Then a sterile, disposable 1 ml syringe with a 25G to 23G hypodermic needle (i. d. 0.50 to 0.65 mm) is made anoxic by displacing the dead space with sterile oxygen-free gas or a reducing agent.

Fig. 3 Anoxic removal of a sample from a Hungate tube:

Penetrate the septum and inject the same volume of oxygen-free gas into the vial as will be subsequently removed as sample from the culture. By doing this the development of an underpressure in the culture tube will be prevented. Then, turn the vial with the culture up side down and fill the syringe with the needed amount of liquid. Finally, withdraw needle and filled syringe carefully.

Reducing agents and resazurin

A redox sensitive dye is usually included in media used for culturing anaerobes to monitor the redox potential. The most commonly used redox indicator is resazurin, because it is generally non-toxic to bacteria and is effective at very low concentrations of 0.5 to 1 g/ml. This indicator dye is dark blue in its inactive form and first has to undergo an irreversible reduction step to resorufin, which is pink at pH values near neutrality (the color may change to blue under alkaline conditions). This first reduction step normally occurs when media containing an excess of organic nutrients are boiled for a few minutes or mineral media are heated under an oxygen-free atmosphere.

In a second reversible reduction step hydroresorufin is formed which is colorless. The resorufin/hydroresorufin redox couple becomes totally colorless below a redox potential of about -110 mV and regains a pink color at a redox potential above -51 mV.

Please note, that some organisms require redox potentials lower than -110 mV and hence may not start to grow even if the medium is colorless. On the other hand a pink color of the medium does not automatically imply that it became oxidized by oxygen (e. g., through an gas permeable rubber septum). For instance, certain nitrate reducers produce nitrite during growth which acts as potent oxidant and so may raise the redox potential above -51 mV.

Reducing agents are added to most anaerobic media to depress and poise the redox potential at optimum levels. The most common reducing agents are sodium thioglycolate, cysteine x HCl, Na2S x 9 H2O, FeS (amorphous, hydrated), dithiothreitol and sodium dithionite.

Sodium thioglycolate is often used in combination with ascorbate and mainly incorporated in some traditional media for culturing anaerobes (e. g., Postgate's media for sulfate reducers, DSMZ medium 63). Thioglycolate as reducing agent has the advantage that it is relatively stable at room temperature and can be therefore included in media prior to flushing with oxygen-free gas. It is only activated by heating above 100 °C and then efficiently removes oxygen. Hence, not so much care has to be taken in avoiding exposure to oxygen of the prepared medium prior to dispensing in anoxic vials. For this reason many commercially available media contain thioglycolate as reducing agent. However, the standard redox potential of thioglycolate alone (around -100 mV) is generally not low enough to allow initiation of growth of a majority of strict anaerobes which need highly reduced media.

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For maximum effectiveness of reducing agents other than thioglycolate, stock solutions under nitrogen gas should be prepared. While preparing stock solutions of reducing agents avoid insertion of the gassing cannula into the liquid, because this can have negative effects on the reducing capacity. Sodium dithionite, which reacts extremely fast with oxygen, has to be dissolved in oxygen-free water and sterilized by filtration. Freshly prepared stock solutions of dithionite can be stored only for up 2-3 weeks at room temperature in the dark.

Add appropriate concentrations of reducing agents to the autoclaved medium just prior to use and allow the medium to sit until it becomes colorless (incubation at 37 °C may accelerate this process). If the medium stays pink despite addition of reducing agent exchange the septum of the vial under a flow of oxygen-free gas, because it might have become permeable to oxygen. Finally, add a small amount of dithionite for final adjustment of the redox potential to a value below -300 mV. If this does not help discard the medium tube.

Literature

Hungate, R. E. 1969. A roll tube method for cultivation of strict anaerobes, pp. 117-132. In J. R. Norris and D. W. Ribbons (eds.), Methods in Microbiology, vol. 3B. Academic Press, New York.

Wolfe, R. S. 1971. Microbial formation of methane. Adv. Microb. Physiol. 6, 107-146.

Breznak, J. A., and Costilow, R. N. 1994. Physicochemical factors in growth, pp. 137-154. In P. Gerhardt (ed.), Methods for general and molecular bacteriology. American Society for Microbiology, Washington.

Notes

1. Abbreviations (excl. chemicals, reagents and measuring units):

approx. = approximately

fig = figure

G = Gauge

i. d. = inner diameter

2. Red colored information indicates an important subject regarding to the content given herein.

3. The information contained herein is offered for informational purposes only and is based on the present state of our knowledge. Recipients of our microorganisms must take responsibility for observing existing laws and regulations. DSMZ does accept no responsibility for the accuracy, sufficiency, reliability or for any loss or injury resulting from the use of the information.

4. Have you any questions or comments to this page? Please send an e-mail to the following address: [email protected]

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Cultivation of Methanogens

Methanogens are a diverse group of strict anaerobes which are widely distributed in nature and can be found in a variety of permanently anoxic habitats like flooded soils, sediments, sewage-sludge digestors or the digestive tract of certain animals. All known methanogens are affiliated to the Archaea and extremely sensitive to oxygen. The hallmark feature of methanogens is the reduction of C-1 compounds (e. g., CO2, methanol, formate, or N-methyl groups) to methane (CH4). Some enzymes and cofactors are unique for this metabolic pathway and therefore only found in methanogens. The coenzyme F420 involved in methanogenesis causes an intense autofluorescence of cells under excitation by shortwave UV light. This autofluorescence is a diagnostic feature and can be used to check cultures of methanogens for contaminants by epifluorescence microscopy.

A detailed description of the cultivation of Methanosarcina barkeri DSM 800T follows below to exemplify the

recommended handling of methanogens. M. barkeri was one of the earliest species of methanogens isolated in axenic culture. It is metabolically very versatile compared to other methanogens and can use also acetate as carbon and energy source (Bryant and Boone, 1987).

M. barkeri DSM 800T is cultured in DSM medium 120 and delivered in 5 ml aliquots in Hungate tubes. After

receipt check a sample of the culture by phase-contrast microscopy. To do this remove aseptically an aliquot of the culture with an 1 ml syringe which was made anoxic as described in the DSMZ Technical Information Cultivation of anaerobes. Under the microscope the cells of this strain appear as nonmotile, large, irregular shaped spheroid bodies which normally occur as packages of several cells. Occasionally, some Methanosarcina strains tend to form large aggregates of up to 1000 m in diameter, which are visible to the naked eye. A phase-contrast micrograph of Methanosarcina barkeri is shown in Fig. 1A, whereas the epifluorescence micrograph in Fig. 1B illustrates the typical blue-green autofluorescence of these cells.

Fig. 1 Phase-contrast (A) and epifluorescence micrograph (B) of DSM 800

T.

Bar, 10 m.

Prepare medium 120 recommended for strain DSM 800T as indicated in the DSMZ catalogue of strains

(Internet: http://www.dsmz.de/media). This methanogen grows well with methanol as substrate and therefore a supply of the culture with H2/CO2 gas mixture is not necessary.

Note: Methanogens other than Methanosarcina spp. that grow only with H2/CO2 as substrate are supplied in media prepared under H2/CO2. Vials of these strains are overpressurized to 0.5 to 1 bar with a gas mixture of 80% H2 and 20% CO2. For the cultivation of these strains it is necessary to supply fresh gas mixture in regular intervals to avoid underpressure by the consumption of H2/CO2 and to remove the produced CH4.

Prior to inoculation reduce medium 120, which contains the redox indicator resazurin, by injection of the reducing agents sulfide and cysteine. Wait until the medium has become completely colorless (redox potential below -110 mV). Then, transfer 0.25 ml of a late logarithmic phase culture of DSM 800

T to 5 ml of freshly

prepared medium 120. Incubate at 30 - 37 °C for up to one week. Growth and methane production (gas bubbles) become visible within 48 hours of incubation.

Caution: Methanosarcina species that grow on methanol can produce a considerable amount of gas according to the formula: 4 CH3OH -> 3 CH4 + CO2 + 2 H2O

Similar to other microorganisms that can produce overpressure during growth in closed vials, Methanosarcina spp. should be incubated in culture vessels (e.g. Balch tubes) that resist substantial pressure and are filled only up to 25% with liquid medium. In addition, fast growing cultures should be vented at least on a daily basis to avoid overpressure. Wear protective goggles during handling of cultures that produce overpressure!

In liquid cultures growth is frequently flocculent and starts with the formation of large cell aggregates that settle down at the bottom of the vial. It is not necessary to agitate tubes during the incubation. Methanosarcina barkeri strains normally grow well in laboratory batch cultures and a turbidity above OD600nm 0.5 is frequently reached at the end of the growth phase.

Cultures of DSM 800T and related strains grown to the early stationary phase are stable at 4 - 10 °C for up to

four weeks.

Literature Bryant, M. P., and Boone, D. R. 1987. Emended description of strain MS

T (DSM 800

T), the type strain of

Methanosarcina barkeri. Int. J. Syst. Bacteriol. 37, 169-170. Notes 1. Abbreviations (excl. chemicals, reagents and measuring units): fig. = figure OD = optical density 2. Red colored information indicates an important subject regarding to the content given herein.




Cultivation of Acidophiles

Acidophiles are adapted to low pH values and show maximum growth rates below pH 5. Many acidophiles are also thermophilic and can be found in thermal vents or hot springs that are acidic. Mesophilic acidophiles, on the other hand, are frequently found in mine drainage water or soils of low pH.

A large number of acidophilic strains do not survive lyophilization and hence are delivered as actively growing cultures from the DSMZ. Cultures grown to stationary phase are usually not very stable and lyse rapidly in spent media. This can be explained by the fact that cells need continuously metabolic energy to maintain a neutral intracellular pH against a steep gradient of excess extracellular protons. Therefore, it is important to transfer the obtained cultures into freshly prepared media immediately upon receipt.

A majority of the fastidious acidophilic strains are aerobic autotrophs and grow either by the oxidation of ferrous iron or sulfur compounds. In order to exemplify the handling of these acidophiles the cultivation of two distinct strains is described in detail: Acidithiobacillus caldus DSM 8584

T grows with elemental sulfur and

Leptospirillum ferrooxidans DSM 2705T with ferrous iron. Additional information on the cultivation of

heterotrophic acidophiles, like e.g. Sulfolobus spp., can be found in the DSMZ Technical Information Hyperthermophiles.

Acidithiobacillus caldus DSM 8584 T

The genus Acidithiobacillus represents a separate lineage within the Proteobacteria. Cells are Gram-negative, small rods (approx. 0.5 x 2.0 m) that are motile by means of polar flagella. While all members of the genus Acidithiobacillus can use sulfur or reduced sulfur compounds for autotrophic growth, some strains can utilize also ferrous iron or hydrogen as energy source.

You will receive from the DSMZ an ampoule with a freeze dried sample of strain DSM 8584T, the type strain of

A. caldus. Prepare medium 150a recommended for this strain according to the instructions given in the DSMZ catalogue of strains (Internet: http://www.dsmz.de/media).

Note: Elemental sulfur melts and then aggregates during autoclaving at 121 °C thereby making it unusable by microorganisms. Hence, it is important to sterilize the sulfur powder separately from the liquid medium. This can be achieved by filling screw cap tubes (18 x 100 mm, borosilicate glass) with sulfur powder (approx. 2/3 volume) to which 1 or 2 drops of distilled water is added. The screw cap should not be tightly closed. The tubes are heated in a water bath to 90-100 °C for 3 hours on each of 3 successive days. The sterilized sulfur can be stored at room temperature in the dark for several months.

Open the ampoule carefully as described in Opening of Ampoules and Rehydration of Dried Cultures. Add approx. 0.5 ml of the freshly prepared medium to the freeze dried pellet and resuspend it. Transfer the cell suspension by using a sterile Pasteur pipette to 5 ml medium in screw cap tubes (16 x 160 mm, borosilicate glass). Prepare 1:10 and 1:100 dilutions in two other tubes and incubate all tubes in a slanted position at 45 °C. Growth should become evident after 2-5 days. As a result of growth most of the sulfur gets coated by bacteria and sinks to the bottom of the tube. In contrast, the sulfur remains at the liquid-air-interface in freshly prepared sterile medium. Fully grown cultures are not very stable and should be weekly transferred in fresh medium. Young cultures can be stored at 4 °C for up to 14 days.

For the cultivation of larger volumes prepare medium in screw cap Erlenmeyer flasks (e. g., 30 ml medium in a 200 ml flask) and incubate with gentle shaking on a rotary shaker. Use 5 - 10% (v/v) of a culture grown to the late logarithmic phase as inoculum.

Leptospirillum ferrooxidans DSM 2705T

Members of the genus Leptospirillum belong to the class Nitrospira within the domain Bacteria and play an important role in the bacterial leaching of sulfidic minerals in acidic environments. Cells are Gram-negative and have a vibroid or spiral-shaped morphology. They are motile by means of a single polar flagellum.

Leptospirilli are strict autotrophs that can use ferrous iron or the reduced sulfur compound of pyrite (FeS2) for growth. Different from Acidithiobacillus strains they are not able to utilize elemental sulfur. In addition, they have generally lower pH optima (pH 1.3 - 2.0) than members of the genus Acidithiobacillus.

L. ferrooxidans DSM 2705T is cultured in DSM medium 882 and delivered in 5 ml aliquots in screw cap tubes.

The density of cells in cultures grown to stationary phase is too low to produce a visible turbidity. Hence, after receipt check a sample of the culture for viable cells by phase-contrast microscopy. Under the microscope you will normally find only several motile vibroid cells per field of view besides numerous large deposits of iron oxide (Fig. 1A).

Prepare medium 882 recommended for strain DSM 2705T as indicated in the DSMZ catalogue of strains

(Internet://www.dsmz.de/media). Please note that the solutions A and B are autoclaved separately at only 112 °C.

Note: In our experience Leptospirillum strains are extremely sensitive to traces of organic substances, which may significantly inhibit growth. To avoid a contamination of medium 882 with organic compounds use only absolutely clean glass vials, chemicals of high purity and double-distilled or MilliQ water for medium preparation.

Screw cap tubes with 5 ml of freshly prepared medium are inoculated with 5 - 10% of the original culture obtained from the DSMZ. Prepare 1:10 and 1:100 dilutions in two other tubes and incubate all tubes in a slanted position at 30 °C without shaking. The purpose of the dilution series is to deplete organic materials, which may have accumulated during culture growth (via cell leakage and lysis) and may inhibit growth. Hence, it is only necessary to prepare dilution series if cultures are used as inoculum that have already reached late stationary phase.

Growth becomes visible after 3 - 10 days by a change of the color of the medium to rusty-brown as shown in Fig. 1B.

Fig 1 Growth of Leptospirillum ferrooxidans DSM 2705

T in liquid

culture. (A) Phase-contrast micrograph of a single cell along with iron oxide deposits. Bar = 5 m. (B) Stationary phase culture in DSMZ medium 882 (left) and uninoculated medium (right).

Young, active cultures of Leptospirillum strains remain active at 15 °C for at least 2 weeks. The viability of cultures can be increased to several months by adding of a sterile suspension of pyrite to stationary-phase ferrous iron-grown cultures (Johnson, 2001).

Literature

Johnson, D. B. 2001. Genus II. Leptospirillum Hippe 2000, 503 VP

(ex Markosyan 1972, 26), pp. 453-457. In Garrity et al. (eds.), Bergey's Manual of Systematic Bacteriology, 2

nd ed., vol. 1. Springer, New York.

Notes

1. Abbreviations (excl. chemicals, reagents and measuring units):


fig = figure





Cultivation of Hyperthermophiles

Hyperthermophiles are defined by a temperature optimum for growth around or above 80 °C. Most representatives of this group belong to the Archaea, whereas only a few are found among the Bacteria (e. g., Aquifex pyrophilus).

Hyperthermophiles are distributed in several physiological groups. Many of them are strictly anaerobic and chemolithoautotrophic, but heterotrophic and microaerophilic or obligately aerobic representatives also exist.

Actively growing strains of hyperthermophiles are delivered from the DSMZ as cultures that have reached late logarithmic or stationary growth phase. Hence, please do not try to increase the cell density of these cultures by further incubation. If, in exceptional cases, a freshly inoculated culture is shipped it is explicitly stated on the culture vial.

In order to exemplify the handling of hyperthermophiles the cultivation of two distinct strains is described in detail: Pyrolobus fumarii DSM 11204

T grows under anaerobic conditions and Sulfolobus solfataricus DSM

1616T under aerobic conditions.

Pyrolobus fumarii DSM 11204T

Pyrolobus fumarii belongs to the Crenarchaeota and represents the most extreme thermophilic organism known to date available in pure culture. It was isolated from material of a black smoker (hydrothermal vent on the sea floor) at the Mid Atlantic Ridge and extends the upper temperature limit for life to 113 °C (Blöchl et al., 1997).

You will receive from the DSMZ an actively growing culture of DSM 11204T in a serum bottle which contains 20

ml of medium 792 under 2 bar overpressure of a 80% H2 and 20% CO2 gas mixture.

Overpressure is applied to culture vials of most anaerobic hyperthermophiles in order to prevent the medium from boiling during incubation and to keep the pH value stable. In addition, numerous anaerobic hyperthermophiles have an autotrophic metabolism and hence H2 and/or CO2 represent important nutrients which have to be supplied in excess to avoid growth limitation.

Caution: Please wear safety goggles during handling of overpressurized cultivation vessels and place vials during incubation in containers that protect against broken fragments in case of explosion. Use culture vessels (e.g. Balch tubes) that resist substantial pressure and are filled only up to 25% with liquid medium. Serum bottles are normally not pressure rated and should be handled with special care.

In general, the maximal cell densities reached by hyperthermophiles in laboratory batch cultures are rather low resulting in a faint or almost non visible turbidity at the end of the growth phase. In addition, some media tend to form precipitates after incubation leading to an increase of turbidity even without growth of the culture. Hence, proliferation of cells should be always checked microscopically.

After receipt of a culture of DSM 11204T first remove overpressure by puncturing the septum (after sterilization

with 70% ethanol) with a sterile injection needle. Remove a sample with a sterile anoxic syringe (see DSMZ Technical Information Cultivation of Anaerobes) and check the culture for cells by phase-contrast microscopy. Then pressurize again the vial to 2 bar overpressure with H2/CO2 gas mixture. Normally, under the microscope only a few nonmotile, irregular coccoid-shaped or lobate cells can be found per field of view (Fig. 1).

Fig. 1 Phase-contrast micrograph of Pyrolobus fumarii DSM 11204

T. Bar, 5 m.

Prepare medium 792 recommended for strain 11204T according to the instructions given in the DSMZ catalogue

of strains (Internet: http://www.dsmz.de/media). Please note that Pyrolobus fumarii is an obligate chemolithoautotrophic microorganism and even traces of organic substances (as impurities of used chemicals or remainings on glass ware) may significantly inhibit growth. Therefore, only use ultrapure chemicals for medium preparation and thoroughly cleaned glass vials rinsed with distilled water as containers for the medium.

Vials with freshly prepared medium are inoculated with 5 - 10% of a culture grown to the late logarithmic phase. It is advisable to prepare a dilution series of all hyperthermophiles sensitive to lysis before incubation. Incubate at 103 °C for 16 to 74 h. After a variable lag phase growth of this strain can be very fast with rapid lysis after stationary phase, thus we do not recommend incubation overnight. Rather, we suggest to check growth at regular intervals during the day. Pyrolobus fumarii couples growth with the reduction of nitrate to ammonium. Hence, proliferation of cells can be checked either by microscopy or monitoring of NH4

+ production

with Nessler's reagent. Often ammonium production starts before a significant number of cells can be found. At the beginning of the stationary phase a white precipitate is formed and the medium can turn slightly pink due to the accumulation of nitrite. That is because nitrite increases the redox potential in the medium which causes the redox indicator resazurin to turn from colorless to pink.

At this phase the culture is not very stable even at room temperature and should be transferred immediately in fresh medium. Only young cultures (early logarithmic growth phase) are relatively stable and can be stored several weeks at 20 °C.

Sulfolobus solfataricus DSM 1616T

The archaeon Sulfolobus solfataricus DSM 1616T belongs phylogenetically to the Crenarchaeota and was

isolated from a volcanic hot spring in Italy (Zillig et al., 1980). All members of the genus Sulfolobus are obligately aerobic, acidophilic, and thermophilic to hyperthermophilic. The preferred substrates for organotrophic growth are complex organic materials, like peptone or yeast extract. Most strains can grow also facultatively lithotrophic by the oxidation of reduced sulfur species (e. g., sulfidic ores or elemental sulfur) to sulfuric acid. The strain Sulfolobus solfataricus DSM 1616

T has lost however the ability to oxidize elemental

sulfur (S0). Members of this genus are adapted to acidic environments and generally do not grow above a pH of

about 6.0. At pH values above 7.5 the cells of most strains lyse. Hence, care has to be taken to adjust the pH of the medium to the correct value. The pH optimum for growth of most strains is between 3.0 and 4.5.

You will receive from the DSMZ an ampoule with a freeze-dried sample of strain DSM 1616T. Prepare medium

182 recommended for this strain according to the instructions given in the DSMZ catalogue of strains (Internet: http://www.dsmz.de/media). Open the ampoule carefully as described in Opening of Ampoules and Rehydration of Dried Cultures. Add approx. 0.5 ml of the freshly prepared medium to the freeze-dried pellet and resuspend it. Transfer the cell suspension by using a sterile Pasteur pipette to 5 ml medium in screw cap tubes (18 x 100 mm, borosilicate glass).

Prepare 1:10 and 1:100 dilutions in two other tubes and incubate all tubes in a slanted position at 70 °C. Growth should become evident after 2 - 5 days. However, sometimes cultures need a longer time for recovery and should be given at least two weeks incubation time before regarding them as non-viable. For the cultivation of larger volumes prepare medium in screw cap Erlenmeyer flasks (e. g., 30 ml medium in a 200 ml flask) and incubate without shaking. Use 10% (v/v) of a culture grown to the late logarithmic phase as inoculum.

Growing cultures are sensitive to lysis and should be checked by phase-contrast microscopy in regularly intervals. Cells of Sulfolobus species have a highly irregular, spherical shape with diameters ranging from 0.7 to 2 m. They occur usually as single cells and stain Gram-negative. A phase-contrast micrograph of cells of DSM 1616

T is shown in Fig. 2.

Fig. 2 Phase-contrast micrograph of Sulfolobus solfataricus DSM 1616

T. Bar, 5 m.

Several methods are possible for the maintainance of grown cultures of Sulfolobus and Acidianus species:

1. Storage at 60 °C:

Cultures generally are viable for about 2 - 4 weeks. Use for transfer to fresh medium at least an inoculum size of 20% (v/v).

2. Storage at room temperature:

Cultures have to be transferred weekly to fresh medium. Use at least 10% inoculum.

3. Storage under N2 gas atmosphere:

According to Grogan (1989) the viability of Sulfolobus species could be significantly increased when cultures were stored anaerobically (under nitrogen) at a pH of about 5.5. Similarily, the facultative aerobic strains of Acidianus survive better anaerobically.

4. Storage after neutralization:

After incubation keep the culture for 1 h at room temperature. Transfer 4.5 ml of the grown culture to another tube containing 20 mg CaCO3, sterilized together with some drops of water. Store neutralized cultures at 4 - 8 °C. Cultures treated in this way have been found viable after 1 month of storage.

5. Preservation in liquid nitrogen:

Incubate 50 ml culture in a 300 ml screw cap Erlenmeyer flask. Keep the grown culture for 2 h at room temperature and add sterile CaCO3 in excess. After a short time allowed for settling of CaCO3 and CaSO4, the supernatant is transferred to sterile tubes and centrifuged. The cell sediment is suspended in a small amount of fresh sterile medium, which has been neutralized with CaCO3 and supplemented with 10% glycerol (sterilized separately). Ampoules with 0.1 ml of the suspension are freezed and stored in the vapour phase of a liquid nitrogen container.

For reviving, ampoules are thawed quickly in a water bath at 37 °C and transferred into about 20 ml medium, which has not been neutralized.

Literature

Blöchl, E., Rachel, R., Burggraf, S., Hafenbradl, D., Jannasch, H. W., and Stetter, K. O. 1997. Pyrolobus fumarii, gen. nov. and sp. nov., represents a novel group of archaea, extending the upper temperature limit for life to 113 °C. Extremophiles 1, 14-21

Grogan, D. W. 1989. Phenotypic characterization of the archaebacterial genus Sulfolobus: comparison of five wild-type strains. J. Bacteriol. 171, 6710-6719

Zillig, W., Stetter, K. O., Wunderl, S., Schulz, W., Priess, H., and Scholz, J. 1980. The Sulfolobus-"Caldariella" group: taxonomy on the basis of the structure of DNA-dependent RNA polymerases. Arch. Microbiol. 125, 259-269

Notes

1. Abbreviations


fig = figure







5.3. “Handbook of Hydrocarbon and Lipid Microbiology”, Springer, 2010.

ISBN 978-3-540-77584-3

Widdel F. Cultivation of Anaerobic Microorganisms with Hydrocarbons as Growth

Substrates.

Kaser F.M. and Coates J.D. Enrichment and Isolation of Metal Respirers and

Hydrocarbon Oxidizers.




Comp. by: PPONRAJ Stage: Revises1 ChapterID: 0000891482 Date:15/6/09

Time:20:09:20

35 Cultivation of AnaerobicMicroorganisms withHydrocarbons as GrowthSubstratesF. WiddelMax Planck Institute for Marine Microbiology, Bremen, [email protected]

1 Principal Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

2 Techniques for Preparing Anoxic Media and Working Under

Anoxic Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

2.1 Some Technical and General Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

2.2 Concentrated Aqueous Stock Solutions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

2.2.1 Trace Element Mixture A (Two Types) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

2.2.2 Trace Element Mixture B . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

2.2.3 Bicarbonate Solution (1.0 M) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

2.2.4 Vitamin Mixture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

2.2.5 Thiamine Solution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

2.2.6 Vitamin B12 Solution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

2.2.7 Sodium Sulfide Solution (0.2 M) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

2.2.8 Sodium Ascorbate Solution (0.5 M) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

2.3 Preparation of Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

2.4 Addition of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7

2.4.1 Gaseous Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7

2.4.2 Liquid Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7

2.4.3 Solid Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

3 Comments on Enrichment and Isolation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

3.1 Enrichment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

3.2 Isolation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

4 Disposal of Hydrocarbon Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

# Springer-Verlag Berlin Heidelberg 2009


Time:20:09:20

Abstract: Microorganisms utilizing hydrocarbons anaerobically as organic growth substrates

are grown in defined anoxic media with sulfate, nitrate or other electron acceptors. Cultivation

of such microorganisms is technically often more elaborated than that of other anaerobes.

Special technical measures include, for instance, dilution of potentially toxic hydrocarbons in

an inert carrier phase, and precautions to aviod diffusion of traces of oxygen through stoppers

during long incubation times. Furthermore, contact of liquid hydrocarbons with stoppers

should be avoided.

1 Principal Considerations

Cultivation of microorganisms with hydrocarbons as growth substrates under anoxic condi-

tions is more demanding than cultivation of conventional anaerobes. Four general points must

be taken into consideration:

1. Anaerobic growth with hydrocarbons is very slow. Whereas doubling times of aerobic

hydrocarbon degraders are in the range of several hours, doubling times of anaerobic

hydrocarbon degraders are in the order of days to several weeks (the latter in the case of

anaerobic methane oxidizers). Most experiments depending on growth therefore include

long ‘‘waiting times.’’ Also cell densities are much lower than those of aerobes.

2. Maintenance of strictly anoxic conditions must be ensured. Especially during the long

incubation times, oxygen can diffuse through stoppers and inhibit growth of anaerobes.

On the other hand, small amounts of O2 may, allow hydrocarbon activation and lead to

partly oxygenated hydrocarbon products that can be degraded further. Such ‘‘pseudoa-

naerobic’’ growth with hydrocarbons may occur especially in enrichment cultures or in

cultures of denitrifiers, which are facultative aerobes.

3. Liquid and solid hydrocarbons are mostly very poorly water soluble. (see Wilkes and

Schwarzbauer in this Volume), and their availability for the bacteria in the mediummay be

strongly limited. So far known, anaerobic hydrocarbon degraders do not produce surfac-

tants to increase hydrocarbon availability in the aqueous phase. Hence, if possible, the

hydrocarbon-water interface should be maximized so as to increase diffusion and/or the

surface area for contact with cells. On the other hand, hydrocarbons strongly adsorb to

stoppers. This may lead to losses of hydrocarbons, which is critical in quantitative growth

experiments, and to deterioration of stoppers. Contact between hydrocarbons and stop-

pers must therefore be kept as minute as possible.

4. Most pure liquid hydrocarbons with noticeable vapor pressure are toxic if added as

pure substances (viz. with chemical activity = 1), in particular due to their interaction

with lipid membranes. Such hydrocarbons must be provided at lower chemical activity.

This can be achieved by their dissolution in hydrophobic inert carriers such as non-volatile

branched-chain alkanes.

2 Techniques for Preparing Anoxic Media and Working UnderAnoxic Conditions

2.1 Some Technical and General Remarks

Techniques involving O2-free N2 or mixtures of N2 and CO2 to prevent entrance of air during

handling of culture flasks and tubes are compulsory. An anoxic chamber is useful but not

2 35 Cultivation of Anaerobic Microorganisms with Hydrocarbons as Growth Substrates


Time:20:09:20

obligatory, unless enzymes are to be purified from anaerobic hydrocarbon degraders. There are

convenient gassing techniques, such as the use of the classical cotton-filled Hungate-syringe

with a bent needle, that allow work with anaerobes at a normal laboratory bench. Details of

these techniques have been published (Balch et al., 1979; Bryant, 1972; Widdel and Bak, 1992)

and are not repeated here.

The use of defined, transparent media that do not contain organic nutrients other than the

hydrocarbon substrate is recommended. An exception is the addition of ascorbate as a mild,

compatible reductant (scavenger of traces of oxygen) to pure cultures of denitrifiers if this does

not serve as a growth substrate (which is usually the case). The medium is prepared in

subsequent steps rather than in one batch. The heat-stable non-volatile major mineral salts

are dissolved and autoclaved. Components that undergo chemical changes (e.g., vitamins) or

volatilization (e.g., hydrogen sulfide) in the heat are added from separately sterilized stock

solutions after autoclaving and cooling of the major mineral salts solution.

Description of the following media is given for sulfate- and nitrate-reducing microorgan-

isms, the most frequently studied anaerobic hydrocarbon degraders, and for methanogenic

cultures which have more recently become of interest. Anaerobic hydrocarbon degradation

coupled to sulfate reduction is presumably quantitatively more important in nature (e.g., in

marine sediments, hydrocarbon seeps, and oil fields) than their degradation coupled to

nitrate. However, denitrifying anaerobic hydrocarbon degraders usually grow much faster,

have higher growth yields, and are therefore more convenient for cultivation in the laboratory,

for instance for enzymatic studies.

2.2 Concentrated Aqueous Stock Solutions

2.2.1 Trace Element Mixture A (Two Types)

A non chelated trace element solution has been frequently used for sulfate-reducing bacteria.

The included acid prevents formation of ferric precipitates during storage. A chelated solution

is common for nitrate-reducing bacteria (> Table 1).

. Table 1

Trace element mixture A

Non-chelated solution Chelated solution

Distilled H2O 987 ml 1,000 ml

HCl (25% = 7.7 M) 13 ml None

EDTA, Na2 salt None 5,200 mg

H3BO3 10 mg 10 mg

MnCl2·4H2O 5 mg 5 mg

FeSO4·7H2O 2,100 mg 2,100 mg

CoCl2·6H2O 190 mg 190 mg

NiCl2·6H2O 24 mg 24 mg

CuCl2·2H2O 2 mg 10 mg

ZnSO4·7H2O 144 mg 144 mg

pH adjustment none to 6.0 (with 1.0 M NaOH)

Cultivation of Anaerobic Microorganisms with Hydrocarbons as Growth Substrates 35 3


Time:20:09:21

Use only fresh, greenish crystals of FeSO4·7H2O; brownish grains indicate weathering and

oxidation. Autoclave, preferentially anoxically under N2.

2.2.2 Trace Element Mixture B

Autoclave. Some colorless flocs that may be formed by reaction of the alkaline solution with

some bottle glasses are harmless.

2.2.3 Bicarbonate Solution (1.0 M)

Dissolve 84 g NaHCO3 in distilled water to a final volume of 1,000 ml. Appropriate portions

may be prepared that can be used as a hole for medium batches (e.g., for 30 ml for 1 l, 60 ml for

2 l; see below). Saturate the solution with CO2 by shaking in a stoppered bottle under a head

space of CO2. Autoclave in closed tubes or bottles with fixed stoppers (butyl rubber or Viton)

under a head space of CO2 (�¼ of total volume).

2.2.4 Vitamin Mixture

Filter-sterilize (pore size, 0.2 mm) and store in the dark (preferentially in brown glass bottles)

at 4�C.

2.2.5 Thiamine Solution


at 4�C.

NaH2PO4 + Na2HPO4 (10 mM total P; pH 7.1) 100 ml

4-Aminobenzoic acid 4 mg

D(+)-Biotin 1 mg

Nicotinic acid 10 mg

D(+)-Pantothenic acid, Ca-salt 5 mg

Pyridoxine dihydrochloride 15 mg

H3PO4 + NaH2PO4 (10 mM total P; pH 3.4) 100 ml

Thiamine Cl·HCl 10 mg

Distilled H2O 1,000 ml

NaOH 400 mg

Na2SeO3·5H2O 6 mg

Na2MoO4·2H2O 36 mg

Na2WO4·2H2O 8 mg



Time:20:09:25

2.2.6 Vitamin B12 Solution


at 4�C.

2.2.7 Sodium Sulfide Solution (0.2 M)

Take a few not too small crystals (ideally a single big crystal) of Ca. 10 g Na2S·9H2O from the

storage jar, determine the exact weight and add to a calibrated cylinder (capacity 250 ml). Use

only colorless, clear crystals of sodium sulfide. In contact with air, stocks of sodium sulfide in

commercial jars are easily oxidized to sodium thiosulfate and other salts. Large crystals with

opaque or milky surface layers (oxidation products) can be briefly washed by rinsing with

distilled water on a plastic sieve. Determine the weight after rinsing and drip-off of water. Add

H2O so as to adjust the solution to 48 g Na2S·9H2O per liter (0.2 M). Dissolve the sodium

sulfide by stirring under an N2 atmosphere (close cylinder with stopper). Aliquot and

autoclave the solution in closed tubes or bottles with fixed stoppers (butyl rubber or Viton)

under a head space of N2 (�¼ of total volume).

2.2.8 Sodium Ascorbate Solution (0.5 M)

Add the NaOH solution slowly while stirring and cooling in an ice water bath, preferentially in

a device that allows gassing with N2 to avoid access of air. Add further NaOH dropwise until

the pH is 8–9. Dilute with distilled H2O to a final volume of 100 ml. Filter-sterilize and store

anoxically under a head space of N2 in the dark at 4�C.

2.3 Preparation of Media

Depending on the physiological type of microorganisms and the salinity of the original source,

one of the following basal mineral media prepared. ‘‘F’’ (freshwater medium) is for micro-

organisms from freshwater habitats. ‘‘S’’ (simple saltwater medium) can be used for marine

microorganisms that do not require magnesium and calcium ion concentrations as high as in

natural seawater; the advantage of the relatively low concentration of these ions (in compari-

son to those in seawater) is that the pH can be increased without significant formation of

precipitates. ‘‘M’’ (full marine medium) is used for marine microorganisms with unknown salt

demands or which require high magnesium and calcium ion concentrations (as in natural

Distilled H2O 40 ml

Ascorbic acid 9 g

NaOH (1.0 M), add slowly 40 ml

Distilled H2O 100 ml

Cyanocobalamin 5 mg



Time:20:09:32

seawater); a certain disadvantage of this medium is the formation of significant precipitates

with increasing pH (> Table 2).

Prepare media in special flasks with tubes for anoxic sterile gassing and a closable outlet

that allows distribution of the complete medium to smaller cultivation tubes or bottles;

devices have been described (Widdel and Bak, 1992). Dissolve the salts subsequently (simul-

taneous addition may lead to chunky precipitates) and autoclave. Cool solution under an

N2–CO2 mixture to prevent redissolution of oxygen. Then, add the other components from

sterile stock solutions as indicated.

Do not use the sodium ascorbate solution for enrichment cultures. Adjust the pH to 7 with

sterile 1 M Na2CO3 or 1 M HCl (autoclaved in closed bottle under gas head space) solution.

Distribute the completed medium in culture tubes and bottles and store anoxically under a

head space of an N2–CO2 mixture. Add the hydrocarbon of interest individually to each tube

or bottle as described in the following section.

. Table 2

Defined media with different salinity (F, freshwater; S, simple saltwater; M, full marine) for

cultivation of anaerobes with hydrocarbons

Sulfate-reducing Nitrate-reducing Methanogenic

F S M F S M F S M

Dissolve subsequently under stirring per liter of dist. H2O (amounts in g)

NaCl – 20.0 26.0 – 20.0 26.0 0.5 20.0 26.0

MgCl2·6H2O 0.5 3.0 10.0 0.5 g 3.0 10.0 0.5 3.0 10.0

CaCl2·2H2O 0.1 0.15 1.4 0.1 g 0.15 1.4 0.1 0.15 1.4

NH4Cl 0.3 0.3 0.3

KH2PO4a 0.2 0.2 0.2

KCl 0.5 0.5 0.5

Na2SO4 4.0 0.05b (0.05)c

After autoclaving and anoxic cooling add per liter (amounts in ml)

Trace elements A 1.0d (non-chelated) 1.0d (chelated) 1.0d (non-chelated)

Trace elements B 1.0d 1.0d 1.0d

NaHCO3-soln.,(1.0 M) 30.0 30.0 30.0

Vitamin mixture 1.0 1.0 1.0

Thiamine soln. 1.0 1.0 1.0

B12-soln. 1.0 1.0 1.0

N2S-soln. (0.2 M) 5.0 – 5.0

Na-ascorbate-soln. (0.5 M) – 3.0 (for pure cultures)e –

NaNO3-soln. (1.0 M) – 4.0–10.0 –

pH, adjusted with 1.0 HCl Usually 6.9–7.1

aFor full marine medium (M) preferentially added from sterile KH2 PO4 stock solution (0.5 M) after autoclaving and

cooling of the other saltsbSulfur source for cell synthesiscMay not be needed as a sulfur source, because sulfide will be addeddA few anaerobic cultures may be stimulated by higher amounts, e.g., 3 ml l�1 (to be tested with each solution)eNot applicable in enrichment cultures, because ascorbate-degrading bacteria will be soon selected



Time:20:09:32

2.4 Addition of Hydrocarbons

2.4.1 Gaseous Hydrocarbons

Gaseous hydrocarbons are obtained from steel bottles via gauges. Aseptic addition is guaran-

teed by passing the hydrocarbon gas through a sterile cotton or a membrane filter. The gaseous

hydrocarbons may be injected through stoppers into the culture head space by means of

syringes with hypodermic needles. The syringes should be pre-flushed anoxically. The added

amount is obvious from the added volume (at 25�C, a volume of 24 ml of the pure gas at

ambient pressure [101 kPa] is approx. 1 mmol).

Waste of hydrocarbon gases and an open gas stream (as in the case of gassing with N2) can

be avoided by using a septum device for filling of syringes (> Fig. 1).

The application of gaseous hydrocarbons with high overpressure (with the exception of

methane) is usually not necessary. In the case of methane, an increased pressure may stimulate

anaerobic methane-oxidizing communities. A safe device has been described that allows

application of high pressure to methane in glass tubes (> Fig. 2; Nauhaus et al., 2002).

2.4.2 Liquid Hydrocarbons

Liquid hydrocarbons can be sterilized by filtration through solvent-resistant cellulose filters

(pore size, 0.2 mm) or be autoclaved in tightly closed bottles with a head space (approx. ½ of

bottle volume); in the case of volatile hydrocarbons, the weight should be controlled to reveal

the tightness of the closure. For storage (as well as for autoclaving), screw caps with Teflon-

coated sealing disks are useful (> Fig. 3a). A special glass flask has been designed for steriliza-

tion and aseptic, anoxic storage of crude oil without loss of volatile components (> Fig. 3b;

Rabus and Widdel, 1996).

Hydrocarbons from stocks may be taken up with anoxic (N2-gassed) syringes and added to

the cultures, for instance by injection through the stoppers (ideally as shown in > Fig. 3c). The

syringes should have plungers with plastic or Teflon sealing. Rubber-sealed plungers are

affected by liquid hydrocarbons such that they stick to the syringe cylinder.

. Figure 1

Sparing filling of a syringe with gaseous hydrocarbons without ‘‘open’’ flushing via a special

glass tube with connection to a bottle, sterile cotton, and a rubber septum. The syringe is initially

flushed with N2; avoid suction of air. For aseptic withdrawal, the rubber stopper may be sterilized

with ethanol (allow to dry).



Time:20:09:36

The problem of low solubility can be minimized by providing a large contact area between

the medium and the overlying hydrocarbon phase. For this purpose, tubes or bottles (prefer-

entially flat bottles) are incubated horizontally. This enlarges the surface area and minimizes

diffusion distances between the hydrocarbon phase and the bacteria in the medium. Flat

bottles (not shown) are particularly useful; however, they may not stand pressure. In a culture

with hexadecane, a-cyclodextrin was used to improve growth (Aeckersberg, 1998). Cyclodex-

trins possess hydrophobic interiors and form inclusion compounds with several hydrocarbons

which in this way are ‘‘transported’’ into the aqueous phase.

The toxicity can in principle be minimized by adding extremely small amounts to keep

the hydrocarbon concentration below saturation. However, such amounts are often below

1 mg l�1 and therefore yield only marginal cell growth. It is therefore much easier to provide

such hydrocarbons from a dilute solution (often 1–20%, v/v) in an inert hydrophobic carrier.

The overlying carrier phase then acts as a reservoir of the hydrocarbon substrate that is

permanently provided at nontoxic concentration. Colorless refined mineral oil (paraffin oil,

pharmaceutical grade; not useful for cultures that degrade long-chain alkanes), 2,2,4,4,6,8,8-

heptamethylnonane, or pristane have been applied as carriers.

Adsorption of hydrocarbons to stopper material can be minimized or prevented in several

ways. Stocks of sterile hydrocarbons can be kept in bottles with screw caps with Teflon-coated

sealing disks. For culture tubes and bottles, Teflon-coated stoppers may be used; however, they

are not easily available and they may be only fabricated by few companies upon special request.

Even if needles penetrate these stoppers, the areas exposed to the hydrocarbon remain

relatively small and adsorption is much slower than at an unprotected stopper surface. In

any case, stoppered culture tubes and bottles containing hydrocarbons should be kept in near-

horizontal position so that the hydrocarbon phase is not in contact with the stopper. This is

achieved by keeping the orifice always lower than the medium level (> Fig. 3c). If the tube or

bottle containing the hydrocarbon phase is initially in an upright position (which is usually the

case), inversion to the horizontal position necessarily brings the hydrocarbon phase in contact

. Figure 2

Anaerobic incubation of a culture with methane at high pressure (high concentration of

dissolved methane). The culture is prepared with a head space of methane. Additional medium is

contained in the syringe. Upon pressurization, this medium is forced into the culture tube and

the methane dissolved under the elevated pressure. The gas volume in the culture tube must

be smaller than the volume in the syringe (Vg < Vs). In this way, a residual gas volume and

implosion of the culture tube is avoided when the syringe is emptied by application of hydraulic

pressure. According to Nauhaus et al. (2002).



Time:20:09:39

. Figure 3

Possibilities to handle liquid hydrocarbons as growth substrates. (a) Use of an autoclaved

hydrocarbon solution. (b) Advanced method for anoxic maintenance of a sterile hydrocarbon.

(c) Special method to add a liquid hydrocarbon to medium without coming into contact with the

stopper.



Time:20:09:39

with the stopper. Shaking the tube or bottle (causing a transient water-hydrocarbon emulsion)

while it is simultaneously being inverted can avoid adherence of large hydrocarbon droplets to

the stopper. An elegant approach is to add the hydrocarbon to the horizontal bottle through

the stopper by means of an anoxic syringe; this can be done in such way that the hydrocarbon

ascends to the medium surface without coming into contact with the stopper (> Fig. 3c).

2.4.3 Solid Hydrocarbons

Solid hydrocarbons as pure substances have been used relatively rarely for cultivation of anae-

robes. The best-known example is the aromatic hydrocarbon, naphthalene (Galushko et al.,

1999) that may be added to a culture tube or bottle from an autoclaved (in closed bottle) stock as

long as this is liquid. Certain inhibitory effects of naphthalene can be avoided and better growth

may be obtained after dissolution in a carrier phase (e.g., 20 mg ml�1). Naphthalene has a

noticeable vapor pressure and is slightly water soluble, so that supply of the slowly growing

bacteria in the medium by diffusion is possible. Alkanes which are solid at room temperature

(e.g., octadecane, C18H38; eicosane, C20H42; and higher) are insoluble and are essentially not

available via diffusion into the medium. Such alkanes may be autoclaved and then added with

pre-warmed pipettes to pre-warmed culture tubes or bottles. Tubes or bottles are rotated so as

to distribute the alkanes and increase their surface area while they are solidifying.

3 Comments on Enrichment and Isolation

3.1 Enrichment

For determination of the natural abundance of anaerobic hydrocarbon degraders, serial

dilution of samples (‘‘dilution to extinction’’) of samples may be attempted, which represents

a special enrichment strategy. However, apart from a few exceptions (Zengler et al., 1999;

Harder et al. 2000) it is unknown how reliable this method is with the slowly growing

anaerobic hydrocarbon degraders. So far, interest was mostly in principles of anaerobic

hydrocarbon degradation (metabolic diversity and capabilities) rather than in natural abun-

dances, and batch enrichments were commonly applied.

Media are provided with 5–10% (V/V) anoxic mud and the hydrocarbon.With the exception

of the gaseous representatives, the consumption of hydrocarbons difficult to quantify. Hydrocar-

bon-dependent microbial activity is detectable by monitoring of the utilization of the electron

acceptor in comparison to a hydrocarbon-free control. Sulfate reduction is easily detectable by

formation of sulfide that can be detected in anaerobically withdrawn samples, for instance with a

copper-containing reagent (Cord-Ruwisch, 1985). Reduction of nitrate and of the intermediate

nitrite can be measured by ion chromatography (with UV-detection). Often, a much simpler

method, the measurement of formed gas (N2) overpressure, is sufficient (> Fig. 4).

3.2 Isolation

If consecutive subcultures have become sediment-free and show clear enrichment of cells

(visible as turbidity and under the microscope), purification via serial agar dilution series

(Widdel and Bak, 1992) or liquid dilutions can be attempted. The agar and the liquid medium



Time:20:09:40

is overlaid with the hydrocarbon, like the preceding enrichment. Both methods are promising

if the hydrocarbon has a noticeable water solubility (e.g., gaseous alkanes, several aromatic

hydrocarbons). With essentially insoluble hydrocarbons (e.g., hexadecane), cells that are not

in direct contact with the hydrocarbon phase may not grow, and this is most likely the case for

the few cells (or a single cell) at high dilution. Such failure of growth may be circumvented by

adding alternative, soluble (polar) substrates. For instance, degraders of long-chain alkanes

are expected to utilize also fatty acids (e.g., 1–2 mM sodium caproate, C5H11COONa).

Pure cultures obtained with the alternative organic substrate are subsequently transferred to

media with hydrocarbons to verify the ability for their utilization. Enrichment cultures with

hydrocarbons often harbor accompanying bacteria that do not utilize the hydrocarbons (they

presumably utilize excreted by-products) but usually grow faster with the non-hydrocarbon

substrate added for isolation. Selection of hydrocarbon degraders with non-hydrocarbon

compounds in liquid enrichment cultures is essentially impossible; they soon select for

different degraders of non-hydrocarbons.

4 Disposal of Hydrocarbon Cultures

Old hydrocarbon cultures which are no longer in use belong to the more ‘‘problematic’’ types

of microbiological waste. Due to the presence of an insoluble and possibly somewhat toxic

hydrocarbon phase, old cultures should not be emptied into a regular sink. Especially cultures

grown with crude oil or petroleum fractions would be problematic in regular waste water. On

the other, collection of the entire culture volumes for special disposal may soon result in big

waste volumes. It is therefore recommended to collected old hydrocarbon cultures in a

separatory funnel. Toxic hydrogen sulfide from sulfate reduction should be oxidized by slow

addition of H2O2 (‘‘titration’’). The aqueous phase can then be emptied into a regular sink (if

there are no ingredients of high environmental concern), while the hydrocarbon phase is

collected in a waste bottle for special disposal.

. Figure 4

Volumetric determination of formed N2 as a simple measure of denitrification and nitrate

consumption.



Time:20:09:40

If cultures contain in addition mud (sediment), which usually harbors hydrocarbon

droplets, the whole culture may be centrifuged. Because centrifuge beakers become ‘‘oily,’’ a

set of these should be kept separately from other beakers and be used only for hydrocarbon

work. The supernatant (aqueous medium and hydrocarbon phase) is then decanted into the

separatory funnel (see above). The mud pellet is removed by means of a spoon, allowed to dry

in a bin under the fume hood, and disposed as special waste.

Stoppers contaminated with hydrocarbons may be used for subsequent cultures with the

same hydrocarbon, or should be disposed.

References

Aeckersberg F, Rainey FA, Widdel F (1998) Growth,

natural relationships, cellular fatty acids and meta-

bolic adaptation of sulfate-reducing bacteria that

utilize long-chain alkanes under anoxic conditions.

Arch Microbiol 170: 361–369.

Balch WE, Fox GE, Magrum LJ, Woese CR, Wolfe RS

(1979) Methanogens: reevaluation of a unique

biological group. Microbiol Rev 43: 260–296.

Bryant MP (1972) Commentary on the Hungate tech-

nique for culture of anaerobic bacteria. Am J Clin

Nutr 25: 1324–1328.

Cord-Ruwisch R (1985) A quick method for the deter-

mination of dissolved and precipitated sulfides in

cultures of sulfate-reducing bacteria. J Microbiol

Meth 4: 33–36.

Galushko A, Minz D, Schink B, Widdel F (1999) Anaer-

obic degradation of naphthalene by a pure culture of

a novel type of marine sulphate-reducing bacterium.

Environ Microbiol 1: 415–420.

Harder J, Heyen U, Probian C, Foss S (2000) Anaerobic

utilization of essential oils by denitrifying bacteria.

Biodegr 11: 55–63.

Nauhaus K, Boetius A, Kruger M, Widdel F (2002)

In vitro demonstration of anaerobic oxidation of

methane coupled to sulphate reduction in sediment

from a marine gashydrate area. Environ Microbiol

4: 296–305.

Rabus R, Widdel F (1996) Utilization of alkylbenzenes

during anaerobic growth of pure cultures of deni-

trifying bacteria on crude oil. Appl Environ Micro-

biol 62: 1238–1241.

Widdel F, Bak F (1992) Gram-negative mesophilic

sulfate-reducing bacteria. In The Prokaryotes,

vol. 4, 2nd edn. A Balows, HG Truper, M Dworkin,

WHarder, K-H Schleifer (eds.). New York: Springer-

Verlag, pp. 3352–3378.

Zengler K, Heider J, Rossello-Mora R, Widdel F (1999)

Phototrophic utilization of toluene under

anoxic conditions by a new strain of Blastochoris

sulfoviridis. Arch Microbiol 172: 204–212.


Comp. by: MGrahalatshmi Stage: Revises1 ChapterID: 0000894224 Date:12/6/

09 Time:19:21:41

39 Enrichment and Isolation ofMetal Respirers andHydrocarbon OxidizersForest M. Kaser . John D. Coates*Department of Plant and Microbial Biology, University of California,Berkeley, CA, USA*[email protected]

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

2 Anaerobic Culturing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

3 Hydrocarbon-Oxidizing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

4 Metal-Reducing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

5 Direct Isolation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

6 Selective Enrichment and Isolation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6

7 Culture Maintenance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8

8 Media Recipes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8

8.1 Freshwater Fe(III)-Oxide Basal Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8

8.1.1 Vitamin Mix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9

8.1.2 Mineral Mix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9

8.2 Freshwater Fe(III)-Citrate Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

8.3 Freshwater Fe(III)-Pyrophosphate Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

8.4 Freshwater Fe(III)-NTA Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

8.5 APW Medium for Marine Isolates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

8.5.1 Salt Solution A . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

8.5.2 Salt Solution B . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

9 Other Techniques for Working with Metal-Reducers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

9.1 Various Fe(III)-Forms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

9.1.1 Amorphous Fe(III)-Oxide Stock . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

9.1.2 Fe(III)-NTA Stock (1 M) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12

9.1.3 Ferric Citrate Stock . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12

9.2 Assaying Fe(III)/Fe(II) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12

# Springer-Verlag Berlin Heidelberg 2009


09 Time:19:21:41

9.2.1 Solutions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12

9.2.2 Reduced Iron Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12

9.2.3 Total Iron Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

10 Most Probable Number Counts (MPN) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

11 Research Needs and Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

2 39 Enrichment and Isolation of Metal Respirers and Hydrocarbon Oxidizers


09 Time:19:21:41

Abstract: The energy conservation strategies of plants and animals are exceedingly homoge-

neous in comparison with those of bacterial cells. Among the wide variety of mechanisms that

exist to provide energy and electrons to support bacterial life, the use of insoluble metallic

elements as electron acceptors is one of the most interesting. The prevalence of metallic

elements in the minerals that comprise soils and sediments raises the possibility that metal-

respiring organisms play a significant role in a variety of biogeochemical transformations. The

environmental abundance of metals also inspires the hope that cells capable of coupling metal

respiration to the oxidation of contaminant hydrocarbons can be captured, studied, and

manipulated to enhance biodegradation processes (Lovley et al., 1994; Wischgoll et al., 2005).

1 Introduction

Only two isolates are known to couple hydrocarbon oxidation to metal reduction,

both classified as species of Geobacter: G. metallireducens (Lovley and Lonergan, 1990) and

G. grbiciae (Coates et al., 2001a). Accordingly, the prescriptions contained herein for the

isolation and characterization of hydrocarbon-oxidizing metal respirers are best regarded as a

starting point rather than a definitive guide. Innovative approaches that account for the

principles underlying these recommendations, but also explore new methodological territory,

are encouraged. To facilitate such exploration, a discussion of general principles precedes a

more detailed explanation of a specific technique. Because of the difficulty of enriching for

both physiological abilities simultaneously, techniques for hydrocarbon oxidizers and metal

reducers are presented separately, with more detail on the latter (for an excellent review of the

former, the reader is referred to Davidova and Suflita [2005]). Descriptions of media and other

useful techniques are included at the end of the chapter.

Two of the most important tasks in characterizing hydrocarbon-oxidizing metal respirers

are linking changes in oxidant concentration to changes in reductant concentration and

establishing that such changes are physiologically rather than chemically mediated. Con-

straints on experiments to determine these facts can include low hydrocarbon concentrations

required to avoid toxicity; technical difficulty of accurately measuring small changes in metal

concentrations; metal species contamination; and long reaction times.

2 Anaerobic Culturing

Nitrogen and carbon dioxide gases used for anaerobic culturing may be contaminated with

small amounts of molecular oxygen. This problem can be minimized by passing the gases

through heated copper filings (350�C) previously reduced by exposure to hydrogen gas

(Hungate, 1969; > Fig. 1). The culturing medium is heated to drive off residual oxygen and

then sparged as it cools with the appropriate gases, whose relative partial pressure can be

adjusted using flow meters. Flowing the same gas through the headspace during sparging and

dispensing prevents the moving liquid from drawing air into the vessel during stirring.

Including a chemical reductant in the medium will help scavenge any residual oxygen and

protect against its reintroduction during dispensing, amendment, or inoculation (Brezak

and Costilow, 1994).

Enrichment and Isolation of Metal Respirers and Hydrocarbon Oxidizers 39 3


09 Time:19:21:41

3 Hydrocarbon-Oxidizing Bacteria

For initial enrichments, liquid from the sampled environment can be deaerated and sterilized

for use as media. Otherwise, the media may be formulated to mimic the environment but also

include an array of vitamins and trace minerals to support any unusual chemistry underlying

anaerobic hydrocarbon transformation reactions.

When used as a sole source of carbon for the enrichment and isolation of anaerobic cells,

hydrocarbons can pose unique challenges due to their toxicity, volatility, and insolubility

(reviewed in Davidova and Suflita, 2005). Over the last 2 decades a great number of method-

ological innovations for the cultivation of anaerobic hydrocarbon-degrading bacteria were

developed by Fritz Widdel’s group at the Max Planck Institute, Bremen, Germany. Hydro-

carbons must be supplied in sufficient quantity to support cell growth, but below that

threshold at which they inhibit growth. Methods to trap hydrocarbon molecules with an

inert organic phase such as 2,2,4,4,6,8,8-heptamethylnonane or mineral oil have proven useful

(Rabus et al., 1993). In such a case, the diffusion rate of hydrocarbons into the aqueous phase

becomes dependent on their removal rate by cells, whereas the concentration is limited by

their aqueous phase solubility. Deaeration and sterilization of the hydrocarbon stock

and carrier phase by autoclaving (less volatile hydrocarbons) or with solvent-resistant filters

(0.2 mm pore size; more volatile hydrocarbons) are helpful in preventing contamination by

molecular oxygen or cells not native to the environment being sampled, respectively. The

volatility of many hydrocarbons has also been exploited by researchers who supply the

substrate in the vapor phase, allowing gas-liquid partitioning to deliver the hydrocarbons to

cells (Fries et al., 1994). For further description of a two-phase delivery system for hydro-

carbons, see the >Chapter 43, Vol. 4, Part 3.

When supplied in an inert overlay, the hydrocarbon of interest generally comprises 2% of

the volume of the carrier phase, but its concentration can easily be monitored by gas

chromatography and replenished if necessary (Davidova and Suflita, 2005). To supply the

. Figure 1

Schematic diagram of equipment used for anaerobic media preparation.



09 Time:19:21:42

hydrocarbon as a vapor, liquid hydrocarbon may be placed in an open container inside a larger

airtight vessel containing cultures in petri dishes (Fries et al., 1994). The volume of liquid

hydrocarbon should be set such that the final concentration in the culture medium does not

exceed inhibitory levels. Different groups of bacteria have different tolerances, but Davidova and

Suflita report that 150 mM benzene or 500 mM toluene generally do not impair the primary

biochemical processes of interest (2005). Especially in the case of the more recalcitrant hydro-

carbons, it may be useful to use water soluble hydrocarbon derivatives such as 4-chlorobenzoate

(0.5 mM) as an electron donor to allow faster growth (Coates et al., 2001b).

4 Metal-Reducing Bacteria

Oxidized metallic elements such as iron that can serve as terminal electron acceptors often

exist in insoluble forms, especially at pH values that support diverse bacterial communities. As

a result, it can be helpful to include a chelating molecule such as nitrilotriacetic acid (NTA) to

distribute the metal throughout the liquid phase (Lovley et al., 1994). Alternatively, the

addition of an electron shuttling agent such as 2,6-anthraquinone disulfonate (AQDS) at

low concentration (100 mM) has been shown to significantly improve hydrocarbon degrada-

tion. However, one must bear in mind that the chelator or electron shuttling agent itself may

be toxic or act as an alternative and more labile electron donor for the microbial enrichment.

This is often the case with ligands such as citrate.

The molecular context of the metallic element should also be considered. The structure and

composition of the mineral species can affect the geometry of electrostatic microenvironment

and the redox potential of the metal, potentially determining the types of cells that can access it.

5 Direct Isolation

Dissimilatory Fe(III)-reducing bacteria can be directly isolated from a broad diversity of envir-

onments using a modified shake tube method. The media of choice uses Fe(III) chelated with

nitrilotriacetic acid (10mM) as the sole electron acceptor and either the hydrocarbon of choice or

any other non-fermentable electron donor such as H2 (101 kPa), acetate (10 mM), or benzoate

(2 mM). If H2 is used a small amount of a suitable carbon source, such as 0.1 g l�1 yeast

extract, should be added to the medium. The media should be further amended with FeCl2(2.5 mM) as a reductant. Freshly collected samples are serially diluted to 10�9 in this medium.

Aliquots (7 ml) of the respective dilutions are transferred anaerobically into anaerobic

pressure tubes containing 3 ml of sterile molten noble agar (Difco) (4% wt/vol) at 55�C

under a gas phase of N2–CO2 (80–20, vol/vol). The sample is mixed by inverting several times

and then solidified by plunging into an ice bath. The solidified dilutions are incubated

inverted. Colonies of Fe(III)-reducing bacteria should be visible in the lower dilutions (10�1–

10�3) after 2 weeks incubation. These can easily be recognized as small (0.5–1 mm diameter)

pink colonies surrounded by a colorless clear zone in the orange-colored agar. In an anaerobic

glove bag colonies can be picked as plugs using a sterile Pasteur pipette, and transferred into

fresh anaerobic medium (5 ml) with Fe(III)-NTA (10 mM) and a suitable electron donor.

These nascent cultures should be further amended with FeCl2 (2.5 mM) as a reductant. Active

cultures can easily be recognized after 2–4 weeks incubation by the color change in the

medium from orange to colorless with a white precipitant in the bottom of the tube. Active

cultures should be transferred through a second dilution shake tube series to ensure isolation.



09 Time:19:21:42

6 Selective Enrichment and Isolation

Dissimilatory Fe(III)-reducing bacteria can be selectively enriched from diverse habitats in

basal media using various forms of Fe(III) as the sole electron acceptor with either H2 or

simple organic acids as the electron donor. Insoluble amorphous Fe(III)-oxide is the prefer-

ential form for enrichment initiation as there are no organic complexing agents which could

potentially be biodegradable and serve as a competing carbon and energy source for fermen-

tative bacteria. Samples collected from the field should completely fill any vessel in which they

are collected to exclude air in the headspace. These should be sealed, transported back to the

laboratory, and used immediately. If not used immediately, samples should not be frozen but

may be stored at 4�C for short periods. Enrichments should be initiated with inoculum sizes of

10% by weight of the culture volume. Incubations should be carried out at environmental

temperatures depending on the source of the sample.

Enrichment for Geobacter species. Geobacter species are mesophilic, complete-oxidizing,

obligate anaerobes. Amorphous Fe(III)-oxide (30 mM) is a suitable electron acceptor for

Geobacter species with acetate (2 mM) as the sole electron donor. Enrichments are generally

incubated at temperatures of 15–30�C. Positive enrichments can be visually identified by a

color change in the amorphous Fe(III) from orange-brown to black as the iron is reduced. If

left for an extended period after complete reduction of the Fe(III) has taken place (1–2 weeks),

the crystalline iron mineral magnetite (Fe2O3) will form which can readily be identified by its

magnetic properties (> Fig. 2). Initial enrichments usually take 1–2 weeks at 30�C. Initial

positive enrichments should be transferred as soon as the iron precipitant in the media has

turned black. Inoculum transfers into fresh media should be 10% of the culture volume.

Sequential transfers should be done three times to remove residual particulates and biode-

gradable organics associated with the original sample. At this stage, the enrichment may be

transferred into basal media amended with a soluble form of Fe(III) such as Fe(III)-NTA

(10 mM). Growth can be recognized by a color change in the media from transparent orange

to colorless and the formation of FeCO3, which is seen as a white precipitant at the bottom of

the culture vessel. Cell yields are generally poor, and an optical increase in cell density will not

be apparent. Once growth has been achieved on Fe(III)-NTA, the enrichment should be

. Figure 2

Iron in the culturing medium that has been converted into magnetite will be attracted by a

magnet through the wall of the culturing tube.



09 Time:19:21:42

transferred to solid media with acetate (2 mM) and Fe(III)-NTA (10 mM) as the sole electron

donor and acceptor respectively amended with 2% by weight noble agar (Difco). Colonies of Fe

(III)-reducing bacteria can readily be recognized on the surface of agar plates as small white

colonies surrounded by a colorless halo in tan-brown colored agar. Colonies should be picked

with sterile glass Pasteur pipettes in an anaerobic glove box and restreaked onto fresh solidmedia.

After incubation several of the restreaked colonies should be picked and used to inoculate a single

tube of fresh anaerobic Fe(III)-NTA media. It is a good idea to amend these nascent cultures

with FeCl2 (2.5 mM) as a reducing agent. A viable Fe(III)-reducing culture should obtained

after 7–10 days.

Enrichment for Desulfuromonas species. Desulfuromonas species can readily be isolated

frommarine environments with Fe(III)-oxide as the sole electron acceptor. The APWmedium

outlined below is a suitable enrichment and growth medium for marine samples. Samples

should be freshly collected and transported back to the laboratory in sealed vials that are filled

to capacity to exclude any O2. Acetate (10 mM) is a suitable electron donor and amorphous Fe

(III)-oxide is the electron acceptor of choice. Enrichments should be initiated as outlined

above for Geobacter species. Active Fe(III)-reducing enrichments should be observed within

7–14 days at 30�C. Active cultures should be passed through at least four transfers of this

medium prior to transfer into medium with a soluble Fe(III) source. Fe(III) chelated with

citrate is the soluble iron form of choice as Fe(III)-NTA precipitates the bound Fe(III) at high

salinities. The highly enriched culture can then be transferred into APWmedium with acetate

and soluble Fe(III)-citrate (50 mM) as the sole electron donor and acceptor respectively.

Growth and Fe(III) reduction can be visually recognized as the medium turns from a deep red

transparent color to dark green and finally to a light green with a white precipitate of FeCO3

visible at the bottom of the tube. The enrichment can then be transferred onto solid medium

with Fe(III)-citrate as the electron acceptor. Fe(III)-reducing colonies will be visible after

7–14 days and can be easily recognized as small pink colonies surrounded by a green halo in

the dark red medium. Colonies can be picked as plugs using a sterile Pasteur pipette and

restreaked onto fresh agar plates. Several colonies can be isolated from these latter plates and

transferred into fresh liquid medium. Active liquid cultures of the isolated Fe(III)-reducers

should be apparent after 5–10 days.

Enrichment for Shewanella species. Shewanella species have been isolated from both

freshwater and estuarine environments. Amorphous Fe(III)-oxide (30 mM) is a suitable iron

form for enrichments of Shewanella species. Lactate (20 mM) or H2 (101 kPa) are the electron

donors of choice. If H2 is used the medium should be amended with yeast extract (0.1 g l�1) as

a suitable carbon source. Positive enrichments can be identified as outlined above for Geo-

bacter and Desulfuromonas species. Once a robust enrichment has been obtained by four or

five passages through an enrichment series the active culture can be streaked on tryptic soy

broth (TSB) (Difco) agar plates and incubated aerobically at 30�C. Colonies of Shewanella

species will appear as pink smooth colonies, 2–4 mm in diameter, after 2–3 days of incubation.

Colonies can be picked and restreaked on TSB plates to further purify the cultures. All isolates

obtained in this fashion should be transferred back into anaerobic medium with Fe(III)-oxide

and lactate or H2 as the electron acceptor and donor, respectively.

Enrichment for Geothrix species. Recent studies (Anderson et al., 1998; Coates et al.,

1999) have shown that Geothrix species may be one of the dominant Fe(III)-reducers found in

mesophilic freshwater environments. These species are relatively slow growing strict anae-

robes. Fe(III)-pyrophosphate (10 mM; Pfaltz and Bauer, Inc.) is the electron acceptor of choice

for their isolation. Enrichment for Geothrix species is similar to that for Geobacter with acetate



09 Time:19:21:42

(10 mM) and amorphous Fe(III)-oxide as the sole electron donor and acceptor respectively.

Incubation periods should be lengthened to 3–4 weeks for each stage of the enrichment to

allow the slow growing Geothrix species to compete. After five passages of the enrichment

through an enrichment series the active culture can be streaked onto anaerobic plates of

Fe(III)-pyrophosphate (10 mM) mediumwith acetate (10 mM) as the sole electron donor and

amended with 2% (wt/vol) noble agar (Difco). The plates should be incubated anaerobically

for 3–4 weeks at 30�C. Fe(III)-reducing colonies can easily be recognized as small (1–2 mm

diameter) white colonies surrounded by a colorless halo in light green agar. The colony is

encrusted with a white mineral precipitate, which is presumably vivianite (Fe3PO4). Colonies

should be picked and restreaked onto fresh agar plates to obtain isolates. After incubation,

several of the restreaked colonies should be picked and used to inoculate a single tube of fresh

anaerobic Fe(III)-pyrophosphate media. The liquid medium should be amended with FeCl2(2.5 mM) as a reducing agent. Active Fe(III)-reducing cultures should be apparent after

3–4 weeks and can be easily recognized by a color change in the medium from light green

to colorless and the appearance of a white precipitate at the bottom of the tube.

7 Culture Maintenance

All mesophilic Fe(III)-reducing cultures can be maintained as frozen stocks at�70�C. The most

reliable technique is to grow the culture in media amended with a soluble electron donor and

acceptor such as acetate (10 mM) and fumarate (50 mM). Once a dense culture has been

obtained, aliquots (1 ml) should be anaerobically transferred into small serum vials (10 ml)

which have previously been gassed out with N2–CO2 (80–20; vol/vol) and heat sterilized. The

vials should be amended with an anaerobic and aqueous glycerol solution (0.1ml) (25% vol/vol)

mixed and frozen at �70�C. Frozen stocks should be checked regularly to ensure viability.

8 Media Recipes

8.1 Freshwater Fe(III)-Oxide Basal Medium

Component Amount

H2O 1.0 l

Amorphous Fe(III)-oxide 30 ml

NH4Cl 0.25 g

NaH2PO4 0.60 g

CH3COONa 3H2O 1.36 g

NaHCO3 2.5 g (primary buffer with CO2 below)

KCl 0.1 g

Vitamin solution 10 ml

Mineral solution 10 ml



09 Time:19:21:43

� Split medium into tubes before sparging with 80% N2 and 20% CO2 (at least 6 min,

the last minute with the stopper in place).

� Final pH should be 6.8–7.0.

� Autoclave for 20 min at 121�C.

� After autoclaving add an appropriate donor.

� The amorphous Fe(III)-oxide can be replaced with alternative Fe(III) forms as needed.

8.1.1 Vitamin Mix

8.1.2 Mineral Mix

Component Amount (mg l�1)

Biotin 2

Folic acid 2

Pyridoxine HCl 10

Riboflavin 5

Thiamin 5

Nicotinic acid 5

Pantothenic acid 5

B-12 0.1

P-aminobenzoic acid 5

Thioctic acid 5

Component Amount (g l�1)

NTA 1.5

MgSO4 3.0

MnSO4 H2O 0.5

NaCl 1.0

FeSO4 7H2O 0.1

CaCl2 2H2O 0.1

CoCl2 6H2O 0.1

ZnCl 0.13

CuSO4 0.01

AlK(SO4)2 12H2O 0.01

H3BO2 0.01

Na2MoO4 0.025

NiCl2 6H2O 0.024

Na2WO4 2H2O 0.025



09 Time:19:21:43

8.2 Freshwater Fe(III)-Citrate Medium

The above outlined basal medium can be prepared with 50 mM soluble Fe(III) chelated with

citrate in replacement of the amorphous Fe(III)-oxide. The Fe(III)-citrate can be added from a

sterile anaerobic stock (1 M) (see Fe(III)-citrate stock solution below) or directly into the basal

medium as follows:

� Check pH (Adjust to 6.0)

� Add medium components outlined for basal medium above

8.3 Freshwater Fe(III)-Pyrophosphate Medium

The amorphous Fe(III)-oxide in the freshwater basal medium can alternatively be replaced

with 3.0 g l�1 soluble Fe(III)-pyrophosphate (Pfaltz and Bauer) added directly to the basal

medium as it is prepared. The resulting Fe(III) concentration will be 10 mM.

8.4 Freshwater Fe(III)-NTA Medium

Fe(III) chelated with nitrilotriacetic acid is only suitable for freshwater media as the Fe(III) will

precipitate out of solution at elevated salinities. In addition Fe(III)-NTA should be filter

sterilized and added from an anaerobic sterile stock (1 M) into heat sterilized media just

prior to inoculation. Fe(III)-NTA stocks are prepared as outlined below.

8.5 APW Medium for Marine Isolates

Component Amount Instructions

H2O 1 l

NaOH 3.4 g l�1 Dissolve

Fe(III)-Citrate 14 g l�1 Boil and cool to room temperature to dissolve

Component Amount

H2O 1.0 l

Ferric citrate 13.7 g (dissolve by heat, cool, and adjust pH to 6.0 with NaOH)

CH3COONa 3H2O 1.36 g

NaHCO3 2.5 g (primary buffer with CO2 below)

NaCl 20 g

KCl 0.67 g

Salt solution A 20 ml

Vitamin solution 10 ml

Mineral solution 10 ml



09 Time:19:21:43

� Split medium into tubes before sparging with 80% N2 and 20% CO2 (at least 6 min, the

last minute with the stopper in place)

� Autoclave for 20 min at 121�C

� After autoclaving add 50 ml salt solution B

� Final pH should be 7.5–7.7

� The vitamin and mineral solution are as outlined above for freshwater basal medium

8.5.1 Salt Solution A

8.5.2 Salt Solution B

9 Other Techniques for Working with Metal-Reducers

9.1 Various Fe(III)-Forms

9.1.1 Amorphous Fe(III)-Oxide Stock

� Dissolve 109 g l�1 FeCl3� Bring pH carefully to pH 7.0 with 10 M NaOH

� Centrifuge for 20 min at 3,000�g

� Pour off supernatant

� Resuspend pellet in distilled water (500 ml) and repeat centrifugation

� Continue this washing procedure until Cl� ion concentration is less than 1.0 mM

� Resuspend final pellet in 100 ml distilled water and store at 4�C

� Use 1 ml of stock solution in 9 ml basal medium

Component Amount (g/100 ml)

NaCl 4.0 g

NH4Cl 5.0 g

KCl 0.5 g

KH2PO4 0.5 g

MgSO4 7H2O 1.0 g

CaCl2 2H2O 0.1 g


MgCl2 6H2O 21.2 g

CaCl2 2H2O 3.04 g



09 Time:19:21:44

9.1.2 Fe(III)-NTA Stock (1 M)

� pH to 6.5

� Degas by sparging with N2/CO2 (80/20) for 10 min

� Filter sterilize

9.1.3 Ferric Citrate Stock

� Heat to dissolve and cool

� Degas with N2 for 20 min and seal with a thick butyl rubber stopper

� Autoclave at 121�C for 15 min

9.2 Assaying Fe(III)/Fe(II)

9.2.1 Solutions

Standards should be prepared from FeSO4(NH4)2SO4 6H2O (0–20 mM)

9.2.2 Reduced Iron Assay

� Add 0.1 ml sample (10–50 mM Fe) to 5 ml 0.5 N HCl

� After dissolution add 0.1 ml of the above to 5 ml ferrozine solution

� Filter through a 0.2 mm filter and read absorbance at 562 nm


NaHCO3 16.4

Nitrilotriacetic acid (sodium salt) 25.6

FeCl3 27.0


NaOH 6.8

Fe(III) Citrate 28.0


HCl 0.5 N

Ferrozine in 50 mM Hepes buffer, pH 7.0 1 g l�1

Hydroxylamine 6.25 N



09 Time:19:21:45

9.2.3 Total Iron Assay

� Add 0.1 ml sample (10–50 mM Fe) to 5 ml 0.5 N HCl and allow it to dissolve

� Add 0.2 ml of the hydroxylamine solution to the dissolved sample

� After 1 h incubation at room temperature in the dark take 0.1 ml of above and add to 5 ml

ferrozine

� Read absorbance at 562 nm

10 Most Probable Number Counts (MPN)

MPNs for Fe(III)-reducing populations in freshwater environments can be performed with a

slight modification to the media outlined above. Amorphous Fe(III)-oxide is the iron form of

choice as no false positives result from the presence of easily biodegraded organics such as

citrate present in the Fe(III)-citrate medium. A non-fermentable electron donor such as H2

(101 kPa), acetate (2 mM), or benzoate (1 mM) should be used. If H2 is being used yeast

extract (0.1 g l�1) should be added as a carbon source.

Basal freshwater medium as outlined previously

Dispense in 9 ml aliquots into pressure tubes and degas individually with N2–CO2 (80–20, vol/

vol) as previously outlined. Autoclave at 121�C for 15 min. Just prior to use add FeCl2 (0.1 ml)

from a sterile anoxic stock (250 mM) as a reductant and 0.1 ml sodium pyrophosphate from a

sterile anaerobic 10% (wt/vol) aqueous stock solution to the initial dilution tubes. This will

serve to release any cells adsorbed onto the soil/sediment particles and significantly improve

the counts obtained. Tubes should be incubated at temperatures suitable to the original sample

environment. Positive MPN tubes should be read after 60 days incubation and can be

identified by measuring the Fe(II) content using the ferrozine assay. An initial visual screening

of the tubes can be done by an observable color change in the iron precipitate from rust-brown

to a dark black color.

11 Research Needs and Outlook

Only two isolates are known to couple hydrocarbon oxidation to metal reduction, both

classified as species of Geobacter: G. metallireducens (Lovley and Lonergan, 1990) and

G. grbiciae (Coates et al., 2001a). Much, therefore, remains to be learned about new

and other bacteria coupling metal reduction to hydrocarbon oxidation. As mentioned

above, the methods described here for the isolation and characterization of hydrocarbon-

oxidizing metal respirers are best regarded as a starting point and there is a definitive need for

Amendment Amount (per 1,000 ml)

Nitrilotriacetic acid (disodium salt) (4 mM) 0.94 g

Amorphous Fe(III)-oxide 30 ml



09 Time:19:21:47

innovative approaches in enrichment and isolation. Apart from having a directmicrobiological

interest, the environmental abundance of metals also inspires the hope that cells capable of

coupling metal respiration to the oxidation of contaminant hydrocarbons can be captured,

studied, and manipulated to enhance biodegradation processes (Lovley et al., 1994; Wischgoll

et al., 2005).

References

Anderson RT, Rooney-Varga JN, Gaw CV, Lovley DR

(1998) Anaerobic benzene oxidation in the Fe(III)

reduction zone of petroleum contaminated aquifers.

Environ Sci Technol 32: 1222–1229.

Brezak J, Costilow R (1994) Physicochemical factors in

growth. In Methods for General and Molecular Bac-

teriology. P Gerhardt (ed. in chief), RGE Murray,

WA Wood, NR Krieg (eds.). Washington, DC:

American Society for Microbiology.

Coates JD, Ellis DJ, Gaw CV, Lovley D (1999) Geothrix

fermentans gen. nov., sp nov., a novel Fe(III)-reducing

bacterium from a hydrocarbon-contaminated aquifer.

Int J Syst Bacteriol 49: 1615–1622.

Coates JD, Bhupathiraju VK, Achenbach LA, McInerney

MJ, Lovley DR (2001a) Geobacter hydrogenophilus,

Geobacter chapellei and Geobacter grbiciae, three new,

strictly anaerobic, dissimilatory Fe(III)-reducers. Int

J Syst Evol Microbiol 51: 581–588.

Coates JD, Chakraborty R, Lack JG, O’Connor SM, Cole

KA, Bender KS, Achenbach LA (2001b) Anaerobic

benzene oxidation coupled to nitrate reduction

in pure culture by two strains of Dechloromonas.

Nature 411: 1039–1043.

Davidova I, Suflita J (2005) Enrichment and isolation of

anaerobic hydrocarbona degrading bacteria. Meth-

ods Enzymol 397: 17–34.

Fries MR, Zhou JH, Cheesanford J, Tiedje JM (1994)

Isolation, characterization, and distribution of deni-

trifying toluene degraders from a variety of habitats.

Appl Environ Microbiol 60: 2802–2810.

Hungate RE (1969) A roll-tube method for cultiva-

tion of strict anaerobes. Methods Microbiol 3B:

117–132.

Lovley D, Lonergan DJ (1990) Anaerobic oxidation of

toluene, phenol, and para-cresol by the dissimilato-

ry iron-reducing organism, GS-15. Appl Environ

Microbiol 56: 1858–1864.

Lovley DR, Woodward JC, Chapelle FH (1994) Stimu-

lated anoxic biodegradation of aromatic-hydrocar-

bons using Fe(III) ligands. Nature 370: 128–131.

Rabus R, Nordhaus R, Ludwig W, Widdel F (1993) Com-

plete oxidation of toluene under strictly anoxic con-

ditions by a new sulfate-reducing bacterium. Appl

Environ Microbiol 59: 1444–1451.

Wischgoll S, Heintz D, Peters F, Erxleben A,

Sarnighausen E, Reski R, Van Dorsselaer A, Boll M

(2005) Gene clusters involved in anaerobic benzoate

degradation of Geobacter metallireducens. Mol

Microbiol 58(5): 1238–1252.


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