29 Methods for Psychrophilic Bacteria

JP Bowman School of Agricultural Science, University of Tasmania, Hobart, Tasmania, Australia

C O N T E N T S

Introduction Isolation of psychrophiles Determination of cardinal temperature values Analysis of fatty acids Phenotypic characterization of marine psychrophiles

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Life forms proliferating in perpetual ly cold environments have developed ecophysiological and biochemical adaptat ions to optimize their activity at low temperatures. A broad spectrum of life is cold adapted and includes bacteria, archaea and simple and complex eukaryotes ranging from algae to fish. Organisms which are cold adapted are often referred to as psychrophiles, a word der ived from ancient Greek and Latin meaning literally cold-loving (psychros cold, philus/phile lover, loving). In the eponymous review on bacterial psychrophiles by Morita (1975), a ' true' psychrophile included any organism able to grow optimally at about 15-20°C or less but was unable to grow at room temperature (20-25°C) or higher. Research suggests true psychrophiles are comparat ively uncommon as the majority of bacteria isolated in cold ecosystems are what can be termed psychrotrophic or facultative psychrophiles, or much more appropria te ly psychrotolerant (Nichols et al., 1995). These bacteria have temperature optima at 20°C or more but are able to grow at 0°C. Indeed, the growth rates of psychrotolerant bacteria are usually equiva- lent to or better than that of psychrophiles at low temperatures. Psychrotolerant bacteria abound even in the coldest of environments, s imply because many of them are ecophysiologically resilient and nutri- tionally versatile species.

The aim of this chapter is to provide an overview of various procedures useful for the isolation and s tudy of a specific group of bacteria, the psychrophiles. Most procedures detailed are general methods modified for cold-adapted bacteria but are broadly applicable in the s tudy of

METHODS IN MICROBIOLOGY, VOLUME 30 Copyright © 2001 Academic Press Ltd 1SBN 0-12-521530 4 All rights of reproduction in any form reserved

marine bacteria. The chapter first covers isolation, routine cultivation and maintenance of psychrophiles. Procedures to accurately determine cardinal growth temperatures using temperature gradient incubator (TGI) are then detailed. Techniques for definitively identifying and quan- tifying fatty acids implicated in cold adaptation of cellular membranes including polyunsaturated fatty acids (PUFA) and branched chain fatty acids are explained. Phenotypic characterization for identification and taxonomic purposes is also covered including a list of phenotypic tests applicable to studying marine psychrophiles and marine bacteria in general.

e e e e e e I S O L A T I O N OF P S Y C H R O P H I L E S

Principle and applications Psychrophilic taxa are found only in permanently cold habitats, environ- ments which have constant annual temperatures of less than 4°C (Morita, 1975) and which are not affected by periodic or intermittent solar insola- tion. The marine ecosystem appears to be an excellent place to find psychrophilic bacteria, as most of the oceanic volume is cold (<5°C). Marine psychrophiles almost without exception are able to grow at temperatures down to the limit at which normal seawater stays liquid (-2 to -5°C). Most known psychrophiles originate from the marine environ- ment, isolated from deep waters, sea-ice and sediment (Table 29.1). Some anecdotal evidence also suggests ideal environments for isolating psychrophiles are both permanently cold and relatively eutrophic. For example Burton Lake, a eutrophic marine basin located in Eastern Antarctic, has enriched psychrophilic bacterial populations throughout its waters (Nichols et al., 1995) while adjacent coastal waters have only low psychrophilic bacterial populations (Delille, 1996). Psychrophilic popula- tions, like other marine heterotrophic bacteria, are closely coupled to primary production (Helmke and Weyland, 1995). Psychrophiles may also abound in attached communities on marine snow and organic detritus or on the surfaces of marine fauna and flora. Data from sea-ice communities suggest species of the order Cytophay(ales and gamma proteobacteria make up the majority of marine psychrophiles (Bowman et al., 1997). This has also been borne out in 16S rDNA base cloning library analyses (Brown and Bowman, unpublished). Species of the order Cytophagales appear to be closely associated with phytoplankton while gamma proteobacteria are more likely to be free-living or associated with marine fauna. Relatively few psychrophiles so far have been isolated from the oceanic pelagic zone but isolates from the deep ocean are usually barophilic or barotolerant and specialized methods may be required for their study (described in detail in Chapter 30 of this volume). Overall, there still appears to be huge scope for the isolation of novel psychrophiles from the marine environment.

A small number of psychrophilic species have been isolated from the terrestrial environment.

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There are no specific protocols, such as specific media or enrichment strategies available for the isolation of psychrophiles. Though the methods described below concentrate on the isolation of psychrophilic chemoheterotrophic bacteria, theoretically prokaryotes of any functional group have psychrophilic equivalents, for example psychrophilic methanogens, sulfate reducers and methanotrophs have recently been described (Table 29.1). Thus, existing isolation protocols for bacteria with distinct functionalities can be adapted by applying the general approach described here.

Samples

Seawater, seawater suspensions of marine sediment, suspensions of detritus and marine snow and pieces/swabs of the surfaces of marine fauna and flora samples can be added directly to or spread/placed onto isolation media and incubated at temperatures of 0-4°C. The samples to be investigated should be stored at all times at 0-4°C until used in exper- iments; however, brief exposure to higher temperatures (<25°C for a few hours) will usually have only a limited deleterious effect on psychrophilic populations present. Sample should never be frozen, as many psychrophiles are very easily lysed by freeze-thawing in the absence of cryoprotectants. Incubation at very low temperatures (<0°C) has little if any benefit as psychrotolerant bacteria can grow just as fast as psychrophiles. For sea ice samples, the sea ice must be thawed in seawater at 0-4°C to prevent hypotonic shocking of the bacteria present. Incubation times are dependent on the initial populations present but in most cases, growth should be visible within 7-28 days.

Media

The media useful for isolating chemoheterotrophic marine psychro- philes are shown below. All media can be solidified using agar (at 1.5%) and are sterilized by standard autoclaving (121°C, 15-20 min). The best general media for marine psychrophiles is marine 2216 agar (Difco Laboratories), alternatively it can be prepared from separate constituents in artificial seawater (ASW, see formula below) or natural seawater. If natural seawater is used for making media, ideally it should be filtered (0.21Jm pore-sized filters, Millipore) rather than autoclaved. Reports suggest better recoveries of bacteria are possible on dilute media pre- pared with raw filtered seawater. Dilute media, including the SWC and SWCm media of Irgens et al. (1989) and R2A medium (Oxoid) prepared in seawater, are also effective for isolating a diverse range of marine chemoheterotrophic bacteria including several difficult psychrophilic species (e.g. Colwellia, Polatvmouas and Polaribacter spp.). Media should be kept refrigerated before being used for isolation, purification or sub- culturing. Likewise any significant exposure of psychrophiles to temper- atures greater than 25°C (more than 1-2 h) should be avoided to prevent loss of viability.

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• Marine medium formula: 5 g peptone, 2 g yeast extract and I 0 mg ferric phos- phate dissolved in 1000 ml ASW. The pH is adjusted to approximately 7.5. The formula is based on marine 2216 agar produced by Difco Laboratories.

• SWC medium formula (Irgens et al., 1989): 0.5 g tryptone, 0.5 g yeast extract, 0.2 g beef extract, 0.2 g sodium acetate dissolved in 500 ml of ASW and 500 ml water. Adjust pH to 7.6.

• SWCm medium formula (Irgens et aL, 1989): 0. I g KH2PO 4, 0.001 g ferric citrate, 0.4g NH4CI, 0.4g yeast extract, 0.4g beef extract, 0.4g tryptone, 2ml Humer's mineral salts (see below) and 0.2g carbon source (optional) dissolved in I000 ml of ASW. Adjust pH to 7.6. Vitamin solution (see below) may also be added if desired at I 0 ml I '

• R2A-seawater agar formula: 0.5 g yeast extract, 0.25 g tryptone, 0.25 g peptone, 0.5g casein hydrolysate, 0.5g D-glucose, 0.5 g soluble starch, 0.024g MgSO4.7H20 and 0.3 g sodium pyruvate dissolved in 1000 ml ofASW.The pH is adjusted to about 7.5. The formula based on R2A agar produced by Oxoid.

• Artificial seawater (ASW) formula (ZoBell, 1946): 0.002g NH~NO3, 0.027g H3BO3, I. 14 g CaCI2.2H20, 0.001 g Fe(PO4)3, 5.143 g MgCI~, 0. I g KBr, 0.69 g KCI, 0.2 g NaHCO3, 24.32 g NaCI, 0.003 g NaE 0.002 g Na203Si.9H~O, 4.06 g Na~SO, and 0.026 g SrCI~.6H20 dissolved in I000 ml of distilled water. The chemicals can be added together dry and mixed thoroughly to make a large supply (add 35 g per liter of media). Sea salts can also be purchased directly from the Sigma-Aldrich Chemical Co.

• Hutner's mineral salts solution (Cohen-Bazire et al., 1957): Dissolve 10g of nitriloacetic acid in 950 ml of water and neutralize by adding 7.3 g of KOH. Then add 14.45 g MgSO~, CaCl2.2H20, (NH4)6MoTO~.4H~O, FeSO4.7H20 and 50 ml stock salts solution and adjust the final pH to 6.8.The stock salts solu- tion consists of 2.5 g ethylenediaminetetraacetic acid, 10.95 g ZnSO,.7H~O, 5.0g, FeSO,.7H20, 1.54g MnSQ.H20, 0.392g CuSO~.5H~O, 0.248g Co(NO~)~.6H20 and 0.177 g Na2B,O 7. I 0H~O in 1000 ml of distilled water slightly acidified with a few drops of sulphuric acid to prevent precipitation.

• Vitamin solution (Balch et al., 1979): Dissolve the following vitamins one at a time in 1000 ml of distilled water and adjust pH to 7.0 using NaOH: 5 mg p-aminobenzoic acid, 2 mg folic acid, 2 mg biotin, 5 mg nicotinic acid, 5 mg calcium pantothenate, 5 mg riboflavin, 5 mg thiamine.HCI, I 0 mg pyridoxine, 0.1 mg cyanocobalamin, 5 mg thioctic acid. Store refrigerated in the dark. Filter-sterilize before use or for long-term storage.

Enrichment and enumeration

No direct isolation methods for psychrophilic bacteria are available, however, an initial enrichment of a sample in liquid isolation media at 0-2°C appears to slightly enhance the isolation of psychrophiles (Morita, 1975). The enrichment should be carried out for up to only 24-48 h and the sample immediately plated or transferred to fresh media. The enumera- tion of psychrophiles is only practical where their populations are greater than that of psychrotolerant bacteria, e.g. in sea ice algal assemblages (Bowman et al., 1998d). By determining most probable number (MPN)

596

counts (see Koch, 1994 for a detailed protocol for MPN analysis) at 0-2°C and at 25°C for a given sample, the proportion of psychrophilic versus psychrotolerant bacteria can be revealed.

Routine maintenance and preservation of psychrophilic cultures

General storage on agar

Many psychrophiles can be maintained on agar plates or slants for long periods at 1-2°C in a frost-free cooler or refrigerator. Care must be taken to avoid freezing of the media due to ice nucleation events as it will result in complete loss of viability of the cultures. Media should be supple- mented with suitable antifungal agents such as cycloheximide (at 100 btg ml ', add from a filter-sterilized 10% ethanol stock) a n d / o r nystatin (at 250 U m l ' , add from a 25 000 U m l ' filter-sterilized methanol stock). If agar plates are used, they need to be quite dry to avoid bacterial contami- nation. Most psychrophiles which have been isolated should be sub- cultured every 4-6 months when stored at 2°C. Storage at higher temperatures (4 to 10°C) requires more frequent transfer (once every 1-3 months) as viability is lost at a higher rate. Some psychrophiles, such as the species Colwellia psych~vrythreae and some members of the Cytophagales, are quite delicate and will die on plates in only a few days.

Cryopreservation

For longer term storage, make a dense suspension of cells in about 2-5 ml of growth media which has been supplemented with 20% glycerol or 20~7~ dimethyl sulfoxide. The suspensions should then be frozen initially at -20°C and then stored at -70 to -80°C. For continued recovery of cells from the frozen suspension, repeated thawing should be kept to a minimum. For most psychrophiles, inoculation of frozen culture directly to plates or liquid media is usually sufficient. Large numbers of small aliquots of the cryopreserved culture(s) may also be a convenient safe- guard as they are thawed only once, used and then discarded. Special tubes and boxes for cryopreservation storage are available from a number of laboratory suppliers.

D E T E R M I N A T I O N OF C A R D I N A L TEM PE RATU RE VALU ES

Principle and applications

The square root growth model (Ratkowsky rt al., 1983) has been imple- mented to accurately determine cardinal growth temperatures of a variety of psychrophilic bacteria (Nichols and Russell, 1996; Bowman et al., 1998a,b,c). This model is based on the principal that the square root of the growth rate is linearly related to temperature and can predict growth rates

597

across the entire biokinetic range (Ratkowsky et al. , 1983). The model can be defined as follows:

~, r = b ( T - TMI~,, )(1 - e ~ cr-r.,~.,~/)

where r is the growth rate at temperature T, T.,~,~ is the notional min imum growth temperature (where 5'r = 0), T,>x is the notional maximum growth temperature (where x/r = 0), b is the slope of the regression line and c is the coefficient to be estimated experimentally. Together with the optimal growth temperature (Topr), T~,~ and T~A x are cardinal temperatures for the biokinetic range of a given organism. All cardinal temperatures occur over a cont inuum range including T,,,\,. For marine psychrophilic and psychrotolerant bacteria, TM,,, values are usually in the range of -5°C to -22°C. Temperature gradient incubator (TGI)-based analysis of cardinal temperatures provides a useful set of autoecological data for psychrophilic bacteria and can be used for physiological and environ- mental comparisons and also provides useful data for strain characterization.

Equipment and reagents

• 150 ml flasks, L-tubes with metal or plastic caps (Bellco). L-tubes are custom- altered test tubes made of optical quality glass (original dimensions 25 cm long, 15 mm diameter) which have modified by glass-blowing to form an L- shape ( 18 cm stem, 7 cm side arm) which allows mixing of the cell suspension in the TGI without spillage.

• TGI or several waterbaths/incubators.The TGI should ideally be placed in a room that has a constant air temperature reducing the fluctuation of temper- atures within the tubes.

• Spectrophotometer which can read glass tubes (diameter 0-15mm), e.g. Spectronic 20D.

• Electronic hand-held thermometer with thermocouple.

Assay

1.

2.

Inocula should be grown to the late logarithmic or stationary growth phase in a suitable growth med ium and at a temperature that ensures rapid growth. For many marine chemoheterotrophic psychrophiles, marine 2216 liquid broth and a temperature of about 10°C is quite adequate. L-tubes containing 10 ml of the growth med ium are placed in the TG! for at least 1 h to allow for temperature equilibration. The TGI should be set with a min imum temperature of about 0°C and a max imum temperature of 30°C (for psychrophiles); for psychrotolerant bacteria the maximum temperature end of the TGI should be set to about 45-50°C.

598

3. L-tubes are inoculated with sufficient growth to achieve an absorbance at 540 nm of about 0.1 (if the initial cell concentration is about 10 ~'~ cells ml ' the amount would be 200-300 ~tl).

4. Optical density readings are then taken just following inoculation. 5. L-tubes are agitated at about 40 oscillations per minute to avoid forma-

tion of oxygen gradients. 6. At periodic intervals after inoculation, optical density values at 540 nm

and the time since inoculation are recorded. Between optical densities of 0.01 to 1.5 turbidity increases linearly (Dalgaard et al., 1994).

7. Growth is complete once optical density at 540 nm exceeds about 1.5. Only the exponential growth rate area of the growth curve is needed and recordings into the stationary growth phase are not necessary. A min imum of 15 readings should be recorded for each tube.

8. At approximately one generation intervals and following growth cessation, the temperature is recorded using an electronic ther- mometer fitted with a thermocouple from each L-tube.

9. Optical density versus time should form a sigmoid curve from which maximum specific growth rate (#,,,,,,) and doubling time (t,) can be de termined from the steepest tangent to the fitted curve as follows:

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Temperature (°C)

25 30

Figure 29.1. Square root growth rate-temperature plots of (a) Shewanella gelidima- rina and (b) Glaciecola punicea showing cardinal growth temperatures T ...... T,,,,~ and T,~x. The plotlines are non-linear regressions fitted using the Macintosh program UltraFit (v 3.0). Data was adapted from Nichols and Russell (1996) and Nichols et al. (1999).

599

where B is the slope of the steepest tangent. The doubling time can be thus determined by fitting the linear section of the growth curve (exponential growth region) with a regression line and determining the time interval (in minutes) required for the optical density to double in this region.

10. To determine cardinal temperatures, the reciprocal square roots of either growth rates or generation times are taken and plotted against their respective temperatures. A non-linear regression is then fitted using appropriate software (e.g. SigmaPlot, UltraFit) (Figure 29.1).

Potential problems and limitations

Growth yields at the supra- and sub-optimal temperature extremes decrease markedly. This has the effect of potentially skewing growth rate information. Thus, T ~ and T ~ values are subject to some level of error. To counter this viable count data, performed by serially diluting cultures and plating onto agar (incubated at the T~,pT) can be used to help pinpoint the temperature growth limits.

41,e,e, Hl, l, A N A L Y S I S OF F A T T Y A C I D S

Principle and applications

The ability to maintain cellular membranes in a homeoviscous state is an important adaptation of psychrophilic bacteria (Nichols et al., 1995). In this respect psychrophilic bacteria often express high levels of anteiso- and iso-branched fatty acids and unsaturated fatty acids, depending on the taxonomic group (Nichols et al., 1995). Several psychrophiles have the ability to form polyunsaturated fatty acid (PUFA) a trait unusual among bacteria (Nichols et al., 1995; Russell and Nichols, 1999). PUFAs produced by psychrophiles include eicosapentaenoic acid (EPA, 20:5m3), docosa- hexaenoic acid (DHA, 22:6m3) and arachidonic acid (AA, 20:4c06), fatty acids which are important 'nutriceuticals' (Nichols et al., 1999). In this section methods for the analysis and identification of fatty acids including PUFA are given.

Fatty acid analysis initially uses a modified Bligh and Dyer proce- dure (Bligh and Dyer, 1959; White et al., 1979) to obtain an extract of whole cell fatty acids and of neutral lipids (hydrocarbons, sterols, waxes, etc.). Fatty acids are then transesterified to methyl esters (fatty acid methyl esters, FAME) and analysed by GC-MS techniques. GC- only systems which allow for rapid fatty acid analysis, such as the MID1 system, identify FAME components by retention times alone, and are not able to definitively identify all fatty acids including many mono-unsaturated fatty acids and unusual fatty acids such as PUFAs. Thus, a high proportion of the fatty acid profile can be left unidentified or even misidentified. Using GC-MS accurate identification of fatty acids can be achieved. Identification of the position of double bonds in

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mono-unsaturated fatty acid FAME is possible by dimethyldisulphide (DMDS) derivatization (Dunkleblum et al., 1985). In this method, DMDS in a chemical reaction catalysed by iodine attacks the fatty acid at the double bond resulting in CH~S adducts which can be identified by GC- MS (Figure 29.2). For more complex PUFAs such as EPA, DHA and AA the number of mass fragments derived from DMDS derivitization makes mass spectra too complicated to be interpreted, instead, PUFA FAME can be reacted with 2-amino-2-methylpropano] to create 2- alkenyl-4,4-dimethyloxazoline (DMOX) derivatives (Fay and Richli, 1991) (Figure 29.2). DMOX derivatives have the advantages of having high volatility allowing direct GC analysis and their mass spectra are easily recognizable allowing unambiguous determination of the posi- tions of unsaturation.

(a) , SCH 3 O ' 2 ~ / II

CH3(CH2)mCH=CH(CH2)nCH2COCH 3 ~ CH3(CH2)mCH--CH(CHo)nCH,:,COCH q DMDS I ~ ~

FAME SCH3

MSD SCH3t ~ ~ CH3 CH3(CH2)mCH--(~H(CH2)nCH2COCH 3 ~ CH3(CH2)mCH

I SCH3 (M') (A')

(~ -CH3OH ~H(CH2)nGH2COCH 3 ----t~ ~H(CH2)n CH=G==O SCH 3 (B') SCH3 (C')

(b) RC/N./"O H2NX' ~ 180o(3 z ~ ' O ~

OHCH3 + HO/J ' ~RCNo, ) " FAME 2-amino-2- DMOX derivative

methytpropanol

Figure 29.2. Chemical reaction schemes for fatty acid methyl ester (FAME) deriv- itization for determination of double bond position. (a) DMDS derivitization showing formation of four diagnostic ions for a straight chain FAME (adapted from Dunkleblum et al., 1985); (b) DMOX derivitization (adapted from Fay and Richli, 1991 ).

Equipment and reagents

Lipid extraction and saponification

• Freeze drying unit (optional), fume hood • Sepatory funnels and stand, GC vials with Teflon-lined screwcaps or septa • Waterbath or incubator set to 80°C • Milli-Q water or double distilled water, nanograde methanol, nanograde

chloroform, nanograde hexane, potassium hydroxide, nonadecanoate (C,9 standard)

• Nitrogen gas and manifold for drying and concentrating samples

601

DMDS derivitization

• Dimethyl disulphide, sublimed iodine, diethyl ether, nanograde hexane, sodium thiosulphate

• GC vials with Teflon-lined screw caps • Incubator or waterbath set at 40°C • Nitrogen gas and gas manifold for concentrating samples

D M O X method

2-amino-2-methylpropanol (free base), dichloromethane, nanograde hexane, sodium sulphate

• GC vials with Teflon-lined screw caps • Oven set at 180°C • Nitrogen gas and gas manifold for drying samples

Chemicals, reagents, solvents, GC vials and other minor equipment can be purchased from a variety of companies including Sigma-Aldrich, Mallinckrodt, Alltech, etc. (see List of suppliers for details).

Gas chromatography-Mass spectrometry

The gas chromatograph utilized should be connected to a Mass Selective Detector (MSD) (Hewlett-Packard, among various companies, produce excellent GC-MS equipment). The GC conditions given in the Assay section have been opt imized for a 50 m x 0.32 mm internal diameter cross- linked methyl silicone (0.171am film thickness) fused silica non-polar capillary column. A polar phase capillary column (using the same GC conditions) can also be used to identify co-eluting components. The carrier gas used is hel ium and the injector and the detector are maintained at 290°C and 310°C, respectively. Operat ing conditions for the MSD include: electron multiplier set at 2000-2200 V; transfer line set at 300°C; autotune file DFTPP normalized; electron impact energy set at 70 eV; scan threshold set at 1500; scan rate set at 0.8 s '; and mass range to be analysed set at 40-600 atomic mass units (amu).

Assay

Bligh and Dyer extraction and saponification

1. Lyophil ized cells (about 10-50 mg) are weighed and added to 8 ml of water, 20 ml of anhydrous methanol and 10 ml of chloroform in a sepa- ratory funnel. The mixture is then shaken vigorously and al lowed to extract for at least 6 h. Freeze-dried cells are ideal if quantification is an important issue, otherwise a cell pellet (washed twice with seawater) can also be used.

2. An additional 10 ml of chloroform and 10 ml of water is added to the suspension, mixed and phases allowed to separate.

3. The lower chloroform phase is then decanted into a round bot tom flask.

602

4. Solvents are then removed in vacuo using a rotary evaporator, and redissolved in a small vo lume of chloroform. The concentrated lipid extract can be stored at -20°C under nitrogen.

5. For saponification all of the chloroform is removed by using a stream of N~ and the residue is resuspended in 3 ml of 5c7~ (w/v ) KOH in 80:20 (v /v) methanol:water. The mixture is then incubated for 3 h at 80°C.

6. After cooling, 1 ml of water and 1.5 ml of 1:1 hexane:chloroform are added, and the mixture is shaken vigorously. The suspension is then centrifuged and the organic phase transferred to a clean tube. The extraction is repeated twice. This yields non-saponifiable neutral lipids such as sterols, hydrocarbons and waxes.

7. To the remaining aqueous phase (containing free fatty acids), 0.5 ml of concentrated HC1 and 1 ml of water are added. This is extracted three times with 1.5 ml of hexane:chloroform as shown in step 6. The solvent is evaporated under N~.

8. The residue is dissolved in transesterification reagent (10:1:1 methanol:chloroform:HC1) and heated at 80°C for 1 h.

9. To the cooled mixture add 1 ml of water and extract three times with 1.5 ml of hexane:chloroform to yield FAME.

10. The solvents are removed by using a stream of nitrogen and the residue re-dissolved in hexane. A known amount of C,~, (nonade- canoate) or similar internal s tandard can then be added. The FAME is now ready for GC-MS analysis and can be used for DMDS derivitiza- tion and the DMOX method.

DMDS derivitization

1. Samples (FAME) in 20-50btl hexane are treated with 70-100~1 of DMDS and one drop of iodine reagent (60 mg of iodine in 1 ml diethyl ether) and incubated at 40°C for 24 h.

2. Reaction mixtures are then cooled and diluted with about 200 btl of hexane.

3. The iodine is removed by adding 100 ul of 5% aqueous sodium thio- sulphate with shaking.

4. The organic phase is removed and the aqueous phase re-extracted with 100 ~1 of hexane.

5. The extract is then concentrated to a small vo lume under a stream of N~. The sample is now ready for GC-MS analysis.

D M O X method

1.

2.

.

Samples (FAME) are dissolved in 500 btl of 2-amino-2-methylpropanol and heated at 180°C overnight. After the reaction mix is cooled, 5 ml of dichloromethane is added and mixed thoroughly. The mixture is then washed twice by extracting with 2 ml of distilled water. The organic phase is then dried by adding sodium sulphate and then evaporated under a stream of N2 at room temperature. The residue is dis- solved in a small amount of hexane and is ready for GC-MS analysis.

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Gas chromatograph-mass spectrometry analysis

1. Samples are injected into the GC at 50°C in the splitless mode with a 2 min venting time. The GC oven is p rogrammed to increase in temperature from 50°C to 150°C at 30°C min ', then at 2°C min ~ until 250°C is reached and then at 1°C m i n ' until a final temperature of 300°C is attained which is maintained isothermally for 15 min. MS acquisition should be started after 7 min for FAME and about 10 rain for DMDS/DMOX adducts. Resultant chromatograms and mass spectra are then compared using appropriate software.

2. Compounds are quantified and identified by comparison of relative retention data, peak area (in relation to the internal standard) and mass spectra with other previously reported compounds.

3. For DMDS adducts der ived from mono-unsatura ted fatty acids four diagnostic ions (M', A', B', C') (Figure 29.1, Table 29.2) occur while only three form for mono-unsatura ted fatty alcohols and fatty aldehydes. In the case of DMOX adducts the presence of a double bond is indicated by an interruption of the regular pattern produced by successive chain cleavages of methylene units. In other words, the mass spectra will show a series of fragment clusters that are separated normally by 14 amu. When a double bond occurs the interval becomes only 12 amu between two fragment peaks. The successive fragment peaks contain n and 11-1 carbon atoms of the acid moiety and thus the double bond occurs between carbons n and n+l in the fatty acid (Fay and Richli, 1991) (Figure 29.3).

Table 29.2 Mass spectrometric data of DMDS derivatives of various FAME ~

FAME DMDS derivatized diagnostic ions

M+ A+ B+ C+

12:1o)5 306 117 189 157 12:1o)3 306 89 217 185 13:1o)5 320 117 203 171 14:1o)9 334 173 161 129 14:1c07 334 145 189 157 14:1o)5 334 117 217 185 14:1o)3 334 89 245 213 16:1~09 362 173 189 157 16:1o)7 362 145 217 185 16:1o)5 362 117 245 213 16:1o)4 362 103 259 227 18:1o)9 390 173 217 185 18:1(o7 390 145 245 213

'Adapted from Dunkleblum et a l (1985).

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/ / I / ~ / ~]i ~ ,a, ~ , 4 ,L ~, L.~.~. , . . . .

250 300 350 400

Figure 29.3. Mass spectrum of docosahexaenoic acid (22:6) DMOX derivative. Double bonds (A) positions are indicated by pairs of diagnostic ions (re~z) sepa- rated by 12 amu: A4 (m/z 139), A7 (m/z 166 178), A10 (m/z 206 218), A13 (m/z 246-258), A16 (m/z 286-298) and A19 (m/z 326-338). Mass spectrum and data adapted from Fay and Richli (1991). The abundance for ions of 178 amu and greater has been increased by five times to make the mass peaks more obvious.

P H E N O T Y P I C C H A R A C T E R I Z A T I O N OF MARINE PSYCHROPHILES

Pr inc ip le and app l ica t ions

This section provides descriptions for a variety of phenotypic tests use- ful for the characterization of psychrophilic marine bacteria. The tests are also widely applicable to other aerobic chemoheterotrophic bacteria simply by altering basal media formulat ion and incubation conditions. Characterization data has a variety of uses, first the data is required for taxonomic analysis, particularly if the objective is to place strains into a novel taxonomic group. In this respect other techniques are also required - - the so called polyphasic taxonomic approach - - in which phenotypic, chemotaxonomic and genotypic data are collectively analysed. Methods for chemotaxonomy including fatty acid, phospho- lipid, quinone and cell wall analysis are described in detail in the litera- ture. Genotypic analysis including DNA base composit ion analysis, DNA:DNA hybridizat ion and 16S rDNA sequence analysis (see Chapter 18 of this volume) are also covered in detail in the literature and the methods therein are broadly applicable to a l l prokaryotes. In this sec- tion, tests covered are proven useful for the characterization of marine chemoheterotrophic bacteria. However , phenotypic tests more specific

605

u o m m ° m e- e~ o L ¢, U >,'r"

a - ~ o

"O o ¢- 4-1 I:

for autotrophic and strictly anaerobic bacteria are not shown as very few or no psychrophiles of these physiological types have been isolated so far. Various literature sources can be consulted for tests applied to char- acterize recently described psychrophilic methanogens, sulphate reduc- ers and methanotrophs. Some of the tests are applicable for screening for cold-adapted enzymes. Methods for quantitative analysis of cold- adapted enzymes are much the same as for normal enzymes. The char- acterization, mechanistic characteristics and features of cold-adapted enzymes have been covered recently in numerous reviews (Feller et al., 1996; Feller and Gerday, 1997).

Ecophysiological tests

Temperature range

For large numbers of strains temperature optima and maxima can be roughly estimated using either liquid or solid media. More accurate indi- cations of T;~.~., T~,,., and T,,:, x should be obtained using liquid media as there is an approximate 5°C differential in To,,, values between solid and liquid media (Bowman et al., 1998a) possibly due to the effect of desicca- tion. For example, Psychroflexus torquis and most Polaribacter spp. show growth up to T,~,~ values of 15-20°C in liquid media but will not grow on agar plates at temperatures above 10-12°C. Thus underest imation of optimal temperatures can occur if agar media is used. Experiments should be performed at 5°C intervals starting at 0 to -5°C.

Salinity range

For marine bacteria the range of salinity at which growth occurs can be easily tested on agar plates or in liquid media and should be tested at approximately their growth temperature optima. Media lacking any added NaC1 (a very low level of Na may be der ived from the organic constituents but levels are usually <1 ppt) can be adapted from the media given in the isolation section. Tests should be performed to see whether strains grow well in the absence of divalent cations, Mg e* and Ca >. Usually there is a strong difference, either the strain grows well or it does not grow at all. For strains requiring divalent cations the media can be supplemented with 50 mm MgCI: and 8 mM CaC12.2H20. As known psychrophiles appear to be almost exclusively slightly halophilic the range of salinity tests does not need to exceed a maximum of 1.5-2 M NaCI.

pH range

This data is usually not necessary when marine bacteria are the focus of s tudy as without exception all have optima approaching that of seawater (pH 7-8) and have a plateau of pH tolerance ranging from at least pH 6.0 to pH 8.0.

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Biochemical tests

Rapid tests kits can be used to determine the biochemical properties of selected strains with API 20E, API 32 AN ID and API-ZYM test strips (BioMerieux) being particularly useful. Though the strips are not designed for identification of marine bacteria, they contain a wide range of tests that do not require special media or specific conditions for use. They are also rapid and convenient though on tile other hand they are expensive. The API 20E test strip has the following tests which can be of use in character- izing marine bacteria: nitrate/nitrite reduction, arginine dihydrolase, lysine and ornithine decarboxylase, gelatin hydrolysis, indole production, H~S production from L-cysteine and tryptophan deaminase. The acid pro- duction tests of the API 20E strips are less useful as marine bacteria mav not acidify carbohydrates strongly enough to give definitive results (see below). The AP132A kit contains a large number of potentially useful enzy- matic tests including: arginine dihydrolase, o~-galactosidase, [3-galactosi- dase, [3-galacto(6-phospha te)sidase, cz-glucosidase, [3-glucosidase, o~-arabinosidase, 13-glucuronidase, ~-N-acetylglucosaminidase, R-fucosi- dase, alkaline phosphatase, glutamate decarboxylase, urease, arginine arylamidase, proline arylamidase, leucyl glycine arylamidase, phenylala- nine arylamidase, leucine arylamidase, pyroglutamate arylamidase, tyro- sine arylamidase, alanine arylamidase, glycine arylamidase, histidine arylamidase, glutamyl glutamate arylamidase, serine arylamidase, man- nose and raffinose fermentation, nitrate reduction and indole production tests. Many of the same arylamidases and glycosidases found on the API 32A test strip are also found on the less expensive AP1-ZYM strip which is designed to allow semi-quantitation of enzymatic activity. These tests are quite useful for directly differentiating a wide range of strains from each other, particularly if high levels of discrimination are required. All of the test strips can be easily set up using suspensions of cultures in sterile sea- water and incubating the strips at about 10°C (or lower if necessary) for several days. Subsequent analysis and interpretation of the test strips should be performed according to the lnanufacturer's instructions.

Hydrolysis of complex and simple substrates

Proteins

Gelatin hydrolysis. Gelatin hydrolysis can be tested in two ways.The first way is more subject to error especially if plates are incubated too long, however, the test is very simple and does not require any special preparation. Dissolve I% gelatin in basal growth media and pour as plates. Following incubation, plates are flooded with I M HCI to precipitate unhydrolysed gelatin that appears white, while clear zones around the growth are indicative of hydrolysis. It is critical that the plates are not incubated for too long (for most psychrophiles up to 5-7 d at I 0°C) as the gelatin hydrolysis zones will quickly cover the entire plate. An alternative method is to use commercially available sterile gelatin- charcoal discs (Oxoid) which are added directly to the sterile liquid basal media. As gelatin hydrolysis occurs the charcoal is released into the media.

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• Casein hydrolysis. A 10-20% suspension of casein powder or skimmed milk in distilled water is autoclaved at reduced temperature (I 10°C, 20 min) and is then added to an equal volume of sterile basal agar medium. It is important not to autoclave the casein with the basal media as the casein interacts with the agar and precipitates.

• Elastin and fibrinogen hydrolysis. Both proteins should be tested as thin overlays (0.5%-I% of protein in the basal agar medium) over a base made of unsup- plemented agar. Hydrolysis of the proteins usually occurs within 7-14 days incubation indicated by the appearance of clear zones around growth.

Hydrolysis of polysaccharides

The hydrolysis of polysaccharides such as starch, chitin, alginate and agar are abilities common among marine bacteria and to various saccharolytic psychrophilic bacteria.

• Starch hydrolysis. Starch hydrolysis can be simply tested by supplementing the basal growth agar medium with I% starch and sterilizing the medium at reduced temperature (I 10°C, 20 min). Following sufficient incubation (at least 7 days) plates are flooded with a 1:5 dilution of Lugol's iodine solution (I g KI and I g sublimed iodine in 100 ml distilled water).The areas of the medium containing unhydrolysed starch are stained dark purple while hydrolysed zones around growth are clear.

• Chitin hydrolysis. The test requires prior purification of commercial practical grade crab shell chitin; however, purified chitin can be purchased but it is prohibitively expensive. Add the precipitated purified chitin as a thin overlay in a mineral salts agar medium to achieve about a I% (w/v) concentration. Chitinase activity is indicated by clear zones around the growth.

Chitin agar: 15 g agar, 3 g precipitated chitin, 2 g (NH4)2SO4, 0.7 g KH2PO4, I mg FeSQ and I mg MnSO, added to 1000 ml artificial seawater.

• Alginate hydrolysis. Supplement the basal medium with I% sodium alginate (add with vigorous stirring and heating) and add about I% agar to form a solid medium. Hydrolysis is indicated by clearing zones around the growth.

• Agar hydrolysis. Hydrolysis is indicated by softening, pitting or liquefaction of the agar medium surrounding and beneath growth.

Lu~ol's iodine solution. Grind 1 g of KI and i g of sublimed iodine in a mortar while adding small amounts of water. Once an even solution is formed dilute the iodine solution to 100 ml. Store in the dark.

Chitin purification. Add 40 g of chitin to 400 ml of cold concentrated HC1 and then precipitate the chitin by adding the solution to 2 1 of distilled water at about 5°C. Filter the suspension through Whatman no. 1 filter paper. Re-suspend the chitin in distilled water and dialyse against tapwater overnight. Adjust the pH to 7.0 using KOH.

Lipolytic enzymes

Esterase activity. Tweens 20, 40, 60, 80 (esters of myristic, stearic, palmitic and oleic acids, respectively) or tributyrin can be used as substrates for

608

esterases. For this Tweens are prepared as 10°~ solutions in distilled water and sterilized separately from the basal agar media preventing precipitation. Different Tweens are then added to the sterile molten agar to obtain a 1~/~ concentration and mixed to fully disperse the Tween. Tributyrin can be added directly to the medium (supplemented with 1 ~;{ polyvinyl alcohol to aid dispersal) before autoclaving, to achieve a final concentration of 1~/~. Hydrolysis of the Tweens is indicated by an opaque hazy zone of calcium soap crystals surrounding the growth. Hydrolysis of tributyrin is indicated by clear zones appearing in the initially cloudy medium.

Lecithinase activity. Lecithinase activity can be tested by adding to the basal medium, sterile egg yolk emulsion (about 5~/~ final concentration) (Oxoid) and observing for opaque zones over and surrounding growth.

Lipase activity. True lipase activity using olive oil or cottonseed oil (Sigma), inexpensive triglyceride substrates (other similar plant oils can be used), can be detected most directly using the procedure of Kouker and Jaeger (1987). To sterile molten basal agar medium, held at about 50°C, add a solution of 2.5% olive oil and 0.001(/~ rhodamine B (31.25 ml 1 ' of media) with vigorous shaking. After standing to allow the foaming to subside the medium is poured as plates. After at least 14 days incubation the plates are observed under long wavelength UV light (about 250 am) (e.g. using a hand-held or normal transilluminator or by using a UV light box). Strains producing a lipase develop a bright orange fluorescence. Strains not producing lipase produce no fluorescence.

Other compounds

Tyrosine hydrolysis or uricase activity can be tested by adding 1% tyro- sine or 1'7~ uric acid to the basal agar media and observing for clearing zones around growth. Certain species on tyrosine agar will also produce red-brown diffusible pigments that are catabolites from the tyrosine degradation and can be used as an extra level of characterization. Activity for [3-glucosidase can be assayed by testing for the hydrolysis of esculin. To do this add 0.1% esculin and 0.01% ferric citrate to the basal agar medium. Esculin hydrolysis is indicated by the appearance of dark tan pigment diffusing into the agar. The production of deoxyribonuclease can be tested using DNAse Test agar (Oxoid),which is prepared in natural or artificial seawater and supplemented with 0.01 g toluidine blue. DNAse activity is indicated by the media around growth changing to red while negative strains stay blue.

Oxidation/fermentation of carbohydrates

The medium of choice for testing the ability of marine bacteria, including psychrophiles, to acidify carbohydrates is the Leifson O/F medium which consists of: 1 g casitone, 0.1 g yeast extract, 0.5 g ammonium sulfate, 0.5 g

609

tris buffer, 0.01 g phenol red and 3 g agar in 1000 ml natural or artificial seawater with pH adjusted to about 7.0. The ability to oxidize and ferment carbohydrates can be tested by first dispensing the basal Leifson med ium into 20 ml screw cap tubes (two tubes per strain) and autoclaving. For sugars which are heat labile (e.g. most mono- and disaccharides) these should be added following autoclaving from filter-sterilized 20% stock solutions to achieve a concentration of 0.5-1%. The tubes are inoculated by stabbing them with an inoculating wire right to the base of the tube. One tube is then sealed with sterile liquid paraffin or with molten 3% agar. Acidification is indicated by the med ium color changing from red to yellow. The presence of fermentation is indicated by the bot tom of the tube or the entire tube turning yellow in the sealed tube. A strictly oxida- tive organism produces a distinct color at the top of the tube of the sealed tube. For testing only oxidative acid product ion from carbohydrates, agar plates (made with 1.5% agar) can be used instead of tubes.

Carbon source and nutritional tests

The utilization of sole carbon sources for carbon, energy and in some cases for nitrogen must be tested in a med ium that is defined sufficiently for bacterial growth. Many marine bacteria can grow in either liquid or agar media containing seawater, a simple combined energy source and a carbon substrate while others require extensive supplementat ion including added vitamins, yeast extract, amino acids and possibly other growth factors. This may lead to problems if a large mix of strains is being investigated as some strains may be able to grow quite well on supple- mented media, e.g. oligotrophic growth. Thus suitable controls lacking supplements and lacking carbon sources are very important. Obviously, strains able to hydrolyse agar must be tested in liquid media. A useful seawater mineral salts media broadly applicable to test psychrophiles consists of the following: 1 g ammonium chloride, 0.1 g yeast extract and 2 ml Hutner ' s mineral salts (see above) dissolved in 1000 ml of natural or artificial seawater. Carbon sources should then be added at a concentra- tion of 0.1%, except carbohydrates which should be added at 0.2%. Labile and volatile substrates should be filter sterilized before addit ion to the sterile basal medium. Following addit ion of carbon sources the pH may need adjustment to about 7.0-7.5. After autoclaving, 10ml of vitamin solution stock is then added to media (cooled to about 50°C). If agar is being used for media, a high puri ty grade (Agarose, Agar Noble) should be used to reduce background growth. Incubation should proceed for up to 1 month at 10°C with close comparison made with control plates lacking a carbon source.

R e f e r e n c e s

Balch, W. E., Fox, G. E., Magrum, L. J., Woese, C. R. and Wolfe, R. S. (1979). Methanogens: reevaluation of a unique biological group. Microbiol. Rev. 43, 260-296.

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Bligh, E. G. and Dyer, W. J. (1959). A rapid method of total lipid extraction and purification. Call. J. Biochem. Physiol. 37, 911-917.

Bowman, J. P., McCammon, S. A., Brown, J. L., Nichols, P. D. and McMeekin, T. A. (1997a). Psychroserpens burtonensis gen. nov., sp. nov., and Gelidibacter algeiTs gen. nov., sp. nov., psychrophilic bacteria isolated from Antarctic lacustrine and sea ice habitats, h#. J. Syst. Bacteriol. 47, 670-677.

Bowman, ]. P., McCamnron, S. A., Brown, M. V., Nichols, D. S. and McMeekin, T. A. (1997'o). Diversity and association of psychophilic bacteria in Antarctic Sea ice. Appl. EHviroH. Microbiol. 63, 3068-3078.

Bowman, J. P., McCammon, S. A., Nichols, D. S., Skerratt, J. H., Rea, S. M., Nichols, P. D., and McMeekin, T. A. (1997c). Shewnuella gelidimariua sp. nov. and ShewalTella h'igidimarina sp. nov., novel Antarctic species with the ability to produce eicosapentaenoic acid (20:5o)3) and grow anaerobically by dissimila- tory Fe(Ill) reduction, h#. J. Syst. Bacteriol. 47, 1040-1047.

Bowman, J. P., McCammon, S. A. and Skerratt, J. H. (1997d). Methylosphaera hansonii gen. nov., sp. nov., a psychrophilic, group I methanotroph from Antarctic marine salinity; meromictic lakes. Microbiology 143, 1451-1459.

Bowman, J. P., Gosink, J. J., McCanrmon, S. A., Lewis, T. L., Nichols, D. S., Nichols, P. D., Skerratt, J. H., Staley, J. T. and McMeekin, T. A. (1998a). Novel Colwellia species isolated from Antarctic fast ice: psychrophilic, marine bacteria with the ability to synthesize docosahexaenoic acid (22:60)3). hlt. 1. Syst. Bacteriol. 48, 1171--1180.

Bowman, J. p., McCammon, S. A., Brown, J. L. and McMeekin, T. A. (q998b). Glaciecoh7 putIicen gen. nov., sp. nov. and Glaciecola pallidula gen. nov., sp. nov.: psychrophilic, marine bacteria from Antarctic sympagic habitats, hH. J. Syst. B~Tcteriol. 48, 1213-1222.

Bowman, J. P., McCammon, S. A., Lewis, T. L. and Nichols, D. S. (1998c). Description of Psychroflexus torquis gen. nov. sp. nov., a psychrophilic bacterium from Antarctic Sea ice with the ability to form polyunsaturated fatty acids and the reclassification of Flavobacterium gondwalletlse Dobson, Franzmann 1993 as Psychroflextts ,\~oJidwalletlse gen. nov. comb. nov. Microbiology 144, 1601 1609.

Bowman, J. P., Rea, S. M., Brown, M. V., McCammon, S. A. and McMeekin, T. A. (1998d). Investigation of aspects of community structure and psychrophily in Antarctic microbial ecosystems. Procecdiuss off the 8th IHterluHio~uTl Symposium oH Microbial Ecolo\,y, Halifax, Nova Scotia.

Cohen-Bazire, G., Sistrom, W. R. and Stanier, R. Y. (1957). Kinetic studies of pigment swlthesis by nonsulfur purple bacteria. ]. Cell. Comp. Physiol. 49, 25-68.

Dalgaard, P., Ross, T., Kamperman, L., Neumeyer K. and McMeekin, T. A. (1994). Estimation of bacterial growth rates from turbidimetric and viable count data. h#. J. Food Miclvbiol. 23, 391-404

Delille, D. (1996). Biodiversity and function of bacteria in the Southern Ocean. Biodivers. CoHserv. 11, 1505-1523.

Deming, J. W., Somers, L. K., Straube, W. L., Swartz, D. G. and MacDonald, M. T. (1988). Isolation of an obligately barophilic bacterium and description of a new genus, Colwellia gen. nov. Syst. Appl. Microbiol. 10, 152-160.

Dunkelblum, E., Tan, S. t t. and Silk, P. J. (1985). Double-bond location in monoun- saturated fatty acids by dimethyI disulphide derivitization and mass spectrom- etry: application to analysis of fatty acids in pheromone glands of four lepidoptera. I. Chem. Ecol. 11, 265-277.

Fay, L. and Richli, U. (1991). Location of double bonds in polyunsaturated fatty acids by gas chromatography-mass spectrometry after 4,4-dimethyloxazoline derivitization. ]. Chromatogr. 541, 89-98.

611

Feller, G. and Gerday C. (1997). Psychrophilic enzymes - molecular basis of cold adaptation. Ceil. Mol. Life Sci. 53, 830 841.

Feller, G., Narinx, E., Arpigny, J. L., Aittaleb, M., Baise, E., Genicot, S. and Gerday, C. (1996). Enzymes from psychrophilic organisms. FEMS Microbiol. Rev. 18, 189-202.

Franzmann, P. D. and Dobson, S. J. (1992) Cell wall-less, free-living spirochetes in Antartica. FEMS Microbiol. Lett. 97, 289-292.

Franzmann, P. D, Liu, Y., Balkwill, D. L., Aldrich, H. C., Conway de Macario, E. and Boone, D. R. (1997). Methallogeuiun~ fri\qdum sp. nov., a psychrophilic, H-,- using methanogen from Ace Lake, Antarctica. Int. J. Syst. Bacteriol. 47, 1068-1072.

Gibson, A. M., Bratchell, N. and Roberts, T. A. (1987). The effect of sodium chloride and temperature on the rate and extent of growth of Clostridium botulinum type A in pasteurized pork slurry. J. Appl. Bacteriol. 62, 479-490.

Gosink, J. J., Herwig, R. P. and Staley, J. T. (1997). Octadecabacter arcticus gen. nov., sp. nov., nonpigmented, psychrophilic gas vacuolate bacteria from polar sea ice and water. Syst. Appl. Microbiol. 20, 356-365.

Gosink, J. J., Woese, C. R. and Staley, J. T. (1998). Polaribacter gen. nov., with three new species, P. irgensii sp. nov., P. fraHzmannii sp. nov. and f~ filamentus sp. nov., gas vacuolate polar marine bacteria of the Cytopha~a-Flavobacterium-Bacteroides group and reclassification of 'Fh'ctobacillus glomeratus' as Polaribacter glomeratus comb. nov. Int. J. Syst. Bacteriol. 48, 223 235.

Helmke, E. and Weyland, H. (1995). Bacteria in sea ice and underlying water of the eastern Weddell Sea in midwinter. Mar. Ecol. Prog. Ser. 117, 269-287.

Irgens, R. L., Suzuki, 1. and Staley, J. T. (1989). Gas vacuolate bacteria obtained from marine waters of Antarctica. Curr. Microbiol. 18, 261 265.

Irgens, R. L, Gosink, J. J. and Staley, J. T. (1996). Polaromonas vacuolata gen. nov., sp. nov., a psychrophilic, marine, gas vacuolate bacterium from Antarctica. Int. /. Syst. Baeteriol. 46, 822-826.

Knoblauch, C., Sahm, K. and Jorgensen, B. B. (1999). Psychrophilic sulfate- reducing bacteria isolated from permanently cold Arctic marine sediments: description of Desulfofrigus oceane1~se gen. nov., sp. nov., Desulfofrigus fragile sp. nov., Desulfofaba go/ida gen. nov., sp. nov., Desulfotalea psychrophila gen. nov., sp. nov. and Desulfotalea arctica sp. nov. Int. J. Syst. Bacteriol. 49, 1631-1643.

Koch, A. L. (1994). Growth measurement. In: Methods for General and Molecular Bacteriology (P. Gerhardt, R. G. E. Murray, W. A. Wood and N. R. Krieg, Eds), pp. 257-260. American Society for Microbiology, Washington, DC.

Kouker, G. and Jaeger, K.-E. (1987). Specific and sensitive plate assay for bacterial lipases. Appl. Environ. MicJvbiol. 53, 211-213.

Morita, R. Y. (1975). Psychrophilic bacteria. Bacteriol. Rev. 39, 144-167. Mountford, D. O., Rainey, E A., Burghart, J., Kaspar, H. E and Stackebrandt, E.

(1998). Psychromonas antarcticus gen. nov., sp. nov., a new aerotolerant anaerobic, halophilic psychrophile isolated from pond sediment of the McMurdo Ice Shelf, Antarctica. Arch. Microbiol. 169, 231-238.

Nichols, D., Bowman, J., Sanderson, K., Mancuso Nichols, C., Lewis, T., McMeekin, T. and Nichols, P. D. (1999). Developments with Antarctic microor- ganisms: culture collections, bioactivity screening, taxonomy, PUFA production and cold-adapted enzymes. Curt. Opiu. Bioteclmol. 10, 240-246.

Nichols, D. S. and Russell, N. J. (1996). Fatty acid adaptat ion in an Antarctic bacterium - changes in primer utilization. Microbiology 142, 747-754.

Nichols, D. S., Nichols, P. D. and McMeekin, T. A. (1995). Ecology and physiology of psychrophilic bacteria from Antarctic saline lakes and sea-ice. Sci. Proy,. 78, 3ll-347.

612

Nogi, Y. and Kato, C. (1999). Taxonomic studies of extremely barophilic bacteria isolated from the Mariana Trench and description of Moritella yayaHosii sp. nov., a new barophilic bacterial isolate. Extremophiles 3, 71-77.

Nogi, Y., Kato, C. and Horikoshi, K. (1998a). Moritella japonica sp. nov., a novel barophilic bacteriurn isolated from a Japan Trench sediment. J. Gen. Appl. Microbiol. 44, 289-295.

Nogi, Y., Kato, C. and Horikoshi, K. (1998b). Taxonomic studies of deep-sea barophilic Shewanella strains and description of Shewanella violacea sp. nov. Arch. Microbiol. 170, 331 338.

Ratkowsky, D. A., Lowry, R. K., McMeekin, T. A., Stokes, A. N. and Chandler, R. E. (1983). Model for bacterial culture growth rate throughout the entire biokinetic temperature range. J. Bacteriol. 154, 1222-1226.

Russell, N. J. and Nichols, D. S. (1999). Polyunsaturated fatty acids in marine bacteria--a dogma rewritten. Microbiology 145, 767-779.

Urakawa, H., Kita-Tsukamoto, K., Steven, S. E., Ohwada, K. and Cohvell, R. R. (1998). A proposal to transfer Vibrio marimts (Russell 1891) to a new genus Moritella gen. nov. as Moritella marilu7 comb. nov. FEMS Microbiol. Lett. 165, 373-378.

White, D. C., Davis, W. M., Nickels, J. S., King, J. D. and Bobbie, R. J. (1979). Determination of the sedimentary microbial biomass bv extractible lipid phos- phate. Oecologia 40, 51-62.

ZoBell, C. E. (1946). Matqne Microbiology. Chronica Botanica Co., Waltham, MA, USA.

List of suppliers

Alltech Associates, Inc. 2051 Waukegan Road Deer field IL 60015, USA Tel: +1-847-948-8600 Fax: +1-847-948-1078 http://alltechweb.com

GC vials, GC co lum ns

Becton-Dickinson Biociences (Difco Products) 7 Loveton Circle Sparks MD 21152, USA Tch +1-410-416-3000 (central operator) +1-800-638-8663 (USA only) http://www.bdms.com/difco/index.html

Marine 2216 agar, microbiological media reagents (yeast extract, pep tone , etc.), agar noble

bioM6rieux, Inc. 595 Anglum Road Hazelwood MO 63042-2320, USA Teh +1-314-731-8500 Fax: +1-314-731-8700 http: //www.bionlerieux.con'l

API test strips

Fluke Corporation 6920 Seaway Boulevard Everett WA 98203, USA Tel: +1-425-347-6100 Fax: +1-425-356-5116 http://www.fluke.coni

Electronic hand -he ld t h e r m o m e t e r wi th t h e r m o c o u p l e

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Hewlett-Packard Corporation 3000 Hanover Street Palo Alto CA 94304-1185, USA Tel: +1-650-857-1501 Fax: +1-650-857-5518 h ttp://www.hp.com/

GC-MS, GC columns

Mallinckrodt Inc. 675 McDonnell Boulevard Hazelwood MO 63042, USA Tel: +1-314-654-2000 Fax: +1-314-654-2328 http://www.mallinckrodt.com/

Nanograde organic solvents

Oxoid, USA Inc. 800 Proctor Avenue Ogdensbu~ NY 13669, USA Tel: +1-800-567-8378 Fax: +1-613-226-3728 http://www.oxoid.com/

R2A agar, microbiological media reagents (yeast extract, peptone etc.), egg yolk emulsion, gelatin- charcoal disks, triple-sugar iron agar, DNAse test agar

Sigma-Aldrich Chemical Co. P.O. Box 14508 St. Louis MO 63178, USA Teh +1-314-771-5750 +1-800-521-8956 Fax: +1-314-771-5757 +1-800-325-5052 http://www.sigma-aldrich.cottl

Artificial sea salts; chemicals for reagents, buffers and media; filter paper

Spectronic Instruments Inc. 820 Linden Aveue Rochester NY 24625, USA Tel: +1-716-248-4000 +1-800-654-9955 Fax: +1-716-248-4014 http: //www.spectpvnic.com/

Spectronic spectrophotometers that can read test tubes

Toyo Roshi Kaisha Ltd., Advantec 1510, Bldg., 1-5-10 Kotobuki, Taito-ku Tokyo 111-0042, Japan Teh +81-3-3842-6290 Fax: +81-3-3842-6299 http://www.advm~tec.co.jp/

Temperature gradient incubator

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