springerreference || section 8 update: linking microbial community structure and functioning: stable...

1

SpringerReferenceH. T. S. BoschkerLinking microbial community structure and functioning: stable isotope (13C) labeling in combination with PLFA analysis

12 Dec 2012 11:34http://www.springerreference.com/index/chapterdbid/76362

© Springer-Verlag Berlin Heidelberg 2012

Linking microbial community structure and functioning:stable isotope ( C) labeling in combination with PLFA13

analysis

Introduction

The cutting edge in microbial ecology is to directly link microbial identity to biogeochemical processes (e.g. [ , ]). The4 7use of stable isotope labeled substrates in combination with biomarker analysis offers the unique opportunity to quantifyand identify in an integrated way the degradation rates and pathways of the substrate, and the organisms involved [ , ].2 6The basic idea behind this approach is that a portion of the added stable-isotope tracer is incorporated into the biomass ofthe metabolically active populations. This biomass comprises molecules that can be used as biomarkers. By comparingthe biomarkers that are labeled with known biomarker compositions of microorganisms, active populations can beidentified. A major strength of the method is that the full biomarker fingerprint of an organism can be used in labelingstudies and hence researchers are not restricted to individual specific biomarkers, which are only found in a limitednumber of genera. This greatly extends the use of biomarker identification in natural environments [ ]. The approach4however depends on the availability of biomarker fingerprints from isolates and there is a need for biomarkercompositions of environmentally important but so far uncultured microbes. In addition to identification, estimates of growthrates and yields of functional sub-populations may be obtained, since polar is closely linked to growth inlipid synthesismicroorganisms [ 13].In this manual, I will limit the description of the technique predominantly to labeling of polar lipid derived fatty acids (PLFA)in soils and sediments with C-organic compounds and by gas13 stable isotope analysischromatography-combustion-isotope ratio mass spectrometry (GC-c-IRMS) as this combination has been used mostoften in microbial ecology. The technique is however very versatile and can be used with combinations of other sampletypes, isotopes, biomarkers and . The various applications of this approach have recently beenanalytical techniquesreviewed for a number of fields [ , 17].3

Procedures

Several discrete steps can be identified in the approach namely: 1) incubating the sample with the C-labeled substrate,13

2) extraction of the PLFA biomarkers, 3) analysis of the PLFA extracts by GC-c-IRMS, 4) treatment of GC-c-IRMS resultsand finally 5) data analysis of the isotope distribution in PLFAs.

Labeling and sample handling

The first step in the method is the labeling of a sediment or soil sample with a stable-isotope labeled substrate. A widerange of C-labeled compounds is available for instance from Isotec (www.isotec.com) and Cambridge Isotope13

Laboratories (www.isotope.com). Amounts of C-label needed are mostly small and label costs are in the same range as13

for radio-labeled organic compounds. Final concentrations of label (99% C) between 1 and 3 μg C/ml sample usually13 13

results a maximum labeling in the range of 100 o/oo δ C in PLFA for soils and coastal sediments, which is about 10013

times the . Organic poor sediment and soils (dune soil for instance) contain less microbial biomass anddetection limitlower amounts of label can be used. Incubation times between 8 and 24 hours usually give maximum labeling dependingon the activity in the sample. A simple test series is sufficient to check for actual labeling levels reached and optimalincubation times.Sample handling should be minimal as disturbances like mixing or temperature shifts may affect the active community.The preferred method is to take small sediment or soil cores for instance made from 25 ml disposable syringes and toinject the label dissolved in a small volume of water with an analytical syringe into the core. We found in several casesthat sediment slurries gave different PLFA labeling patterns suggesting that a shift in active populations occurred. In-situ

2




labeling studies are an interesting and feasible option with C labeling [ 12, 14], and may not suffer from possible13

artifacts during sample treatment.Cross feeding may occur in labeling studies for instance by organisms feeding on an excretion product. This is found forinstance in anaerobic communities where fermenting bacteria produce a variety of intermediates used by other groups oforganisms. Cross feeding can be tested by applying specific inhibitors if available or in pulse-chase experiments wherecross-feeding organisms would label later in the experiment than the primary consumers of the label.PLFA are easily lost by degradation. The preferred method of conserving samples is to add them directly to the extractionmixture after incubation and to the first part of the extraction to the total lipid extract within a couple of days. If this isfinishnot possible, sediments can be quickly frozen, freeze-dried and stored dry at −20 °C for several months. Some losses of

found in eukaryotes may however occur in freeze dried sediments.poly-unsaturated fatty acidsThe method is applicable to other types of samples besides sediments and soils. Plant litter usually contains highamounts of PLFA and less than 0.25 g dry weight is needed for analysis. PLFA in suspended material of aquatic samplescan be collected by filtering over precombusted (450 °C for 3 hours) Whatmann GF/F filters. Between 0.5 andglass fiber5 l of water is enough for estuarine and coastal waters and the filters are added directly to the extraction mixture.Amounts of C-label needed for aquatic samples are low in the range of 1 μg C/l. For living tissues like bacterial and13 13

algal cultures about 2 mg of dry weight are mostly sufficient.

Extraction of PLFA

The PLFA extraction procedure consists of three steps: 1) a total lipid extraction followed 2) by a fractionation of the totallipid extract on a silicic-acid column to isolate the polar phospholipids and 3) finally a derivatisation to release the PLFA asfatty acid methyl esters that can be analyzed by various GC methods. The extraction procedure described here is(FAME)used in our lab to accommodate the sometimes-large number of samples that need to be processed in labeling work andmostly uses standard lab ware. Lipid extracts can be stored in the freezer at −20 °C after each step in the procedure.Contamination can be a problem as fatty acids or other lipids that interfere with the analysis are everywhere (hands,plastics, oils, greases etc.). Equipment used should be made of solvent-inert materials like glass, Teflon or metal. Forhandling solvents and lipid extracts, we use glass culture tubes with Teflon lined caps, Pasteur pipettes and glasssyringes of various sizes equipped with whole-metal needles. Plastics other than Teflon should be avoided except forpipeting . All materials should be machine washed and cleaned with methanol and hexane. Organicaqueous solutionssolvents should be of the best analytical quality as used for trace analysis.Other descriptions of the PLFA extraction procedure with variations can be found elsewhere (e.g. [ , 9]). The web site of8the Scottish Crop Research Institute (www.lipid.co.uk/infores/) contains a wealth of general information on fatty acidanalysis.

Total lipid extraction

Total lipids are extracted in this step using an adaptation of the Bligh and Dyer method [ ]. The description is for dry1sediments and soils, but can be used for wet samples if the volume of water in step A is decreased by the amount ofwater added with the sample.

Add 7.5 ml chloroform, 15 ml methanol and 6 ml MilliQ water to a 50 ml tube.Weight approximately 3 g dry sediment into the tube. This amount is sufficient for most sediments and soils. Lessmaterial is needed for organic rich samples like forest soils, whereas for very poor coarse or sandy material up to6 g of sample can be used.Close tube well and shake for 2 hours (100 to 200 rpm).Add 7.5 ml chloroform, shake well by hand, add 7.5 ml Milli-Q water and shake well again. Let the tubes standuntil solvent layers are separated. This separation process can be enhanced by gentle centrifugation (1000 rpm, 5minutes) and/or by leaving the tubes in a fridge (−20 °C) overnight. Collect the lower (chloroform) layer containingthe lipids with a glass syringe and weigh the collected amount of chloroform extract into a pre-weighted 10 mltube. Only about 8 to 10 ml of chloroform can usually be recovered, which needs to be accounted for in thecalculation of PLFA concentrations.Evaporate chloroform extract to complete dryness under a stream of in a manifold thatnitrogen gasaccommodates a number of samples. Pressurized air can also be used preferably from gas bottles and must be

3




free of contaminants. The different evaporation steps in this protocol are time-consuming. Heating the tubes withfor instance a hair accelerates evaporation. After the chloroform has evaporated, re-dissolve the extract in adryerlittle chloroform and dry again. Finally, dissolve the total lipid extract in about 0.5 ml chloroform.

Isolation of phospholipids

Phosphollipids are isolated by separating the total lipid extract on silicic-acid columns into different polarity classes.

Silicic acid (Merck Kieselgel 60, grain size 0.063-0.200 mm) is activated by heating it to 120 °C for 2 hours.Activated silicic acid can be kept in a closed bottle for 2 to 3 days.Columns are made in 10 ml glass, measuring pipettes from which the narrow top is removed. A small glass-woolball is pressed in the tip of the pipette to retain the silicic acid. The pipette is first cleaned with 5 ml chloroform(inside and outside of tip) and the tip is then submerged in tube containing chloroform. Silicic acid (0.5 g) is addedfrom the top and allowed to settle in the chloroform with help of some tapping on the column to remove airbubbles. The column is now ready for use. Ready-made silicic-acid columns ( columns) can also be boughtSPEfor instance from Baker (nr 7493-06, www.jtbaker.com) and used according to the manufacturer instructions.

Prepare a 0.5 g silicic-acid column as above and wash the column with 5 ml chloroform.Add the total lipid extract to top of the column. We often use only half of the total lipid extract and keep the rest asspare for other analysis.Elute with subsequently 7.5 ml chloroform, 7.5 ml acetone and 15 ml methanol. Collect the methanol fraction in a20 ml tube (this is the most polar lipid fraction containing the PLFA). Do not disturb top of column while addingsolvents. The top of the column should also not run dry between solvent additions.Evaporate methanol fraction to dryness under a stream of nitrogen gas. Heating the tubes with for instance a hairdryer accelerates evaporation.

Derivatization

In this step the fatty acids are released from the isolated phopholipids and converted to FAMEs, which can be analyzedby GC methods.

The methanolic-NaOH that is used as the reagent is made by dissolving a piece of metallic sodiumderivatizationwell cleaned with hexane in the appropriate volume of dry methanol (0.58 g Na in 100 g methanol). Cool in waterwhile the metallic sodium is dissolving. The reagent can be stored under nitrogen gas in well-closed bottles forseveral weeks. The presence of water during the derivatization (steps A and B) will result in the formation of freefatty acids instead of FAMEs.Two internal standards are used. The 19:0 internal standard is used for calculating PLFA concentrations and asan internal reference for isotope analysis, and 12:0 to detect possible evaporation losses of the most volatile

in step E. Both are also used as retention time markers for identification of PLFA peaks.FAME

Add 1 ml methanol/toluene (1:1 v/v), 20 μl 0.1 mg 19:0 FAME/ml (internal standard) and 1 ml 0.2 M methanolicNaOH to the dried phospholipid fraction.Incubate for 15 minutes at 37 °C.Add 2 ml hexane, 0.3 ml 1 M acetic acid in MilliQ and 2 ml MilliQ. Shake well by hand and let layers separate.Transfer the upper, hexane layer to a 10 ml tube without taking any of the lower aqueous phase. Repeat thehexane extraction.Add 20 μl 0.1 mg 12:0 FAME/ml (the second internal standard).Evaporate hexane to dryness under a mild flow of . Do not apply heat in this step, as the lowernitrogen gasFAMEs are sensitive to evaporation losses. The hexane surface should not be moved by the nitrogen gas flow.Transfer sample in about 100 μl hexane to a GC-sample vial and keep the sample frozen until analysis.

Analytical methods

PLFA as extracted by the above procedure are analyzed by a number of gas chromatographic techniques (GC). We

4




usually start the analysis with a GC equipped with a flame ionization detector (GC-FID), as it is a robust method that givesconcentrations and a first identification of the PLFA. Further identification is gained by analyzing the samples on twodifferent analytic columns with different separation behavior ( Fig 1) and by organic (see the web site of theGC-MSScottish Crop Research Institute (www.lipid.co.uk/infores/) for details and mass spectra of FAMEs). In this description ofthe method I will however concentrate on the GC-c-IRMS analysis, as it is the current state-of-the-art method for stable

of PLFA. It also yields concentrations and a first identification of the PLFA in the sample based onisotope analysisretention times.

Figure 1 PLFA analysis of an intertidal sediment sample labeled with a C-amino acid mixture to a final concentration of 20 μM as13

analyzed by GC-c-IRMS on an HP 5MS column. For comparison, PLFA analysis of the same sample on the very polar BPXa-polar70 column is shown in Fig 1B.

A GC-c-IRMS comprises a GC equipped with a capillary column that is used to separate the compounds of interest athigh resolution. The outlet of the column is attached to a miniature oxidation reactor where the organic molecules arecombusted to CO and H O gas. Water is removed online and the purified CO is led into an isotope ratio mass2 2 2spectrometer (IRMS). Because of its design, an IRMS measures the isotopic ratios between the heavy and light isotopes (

C/ C for carbon) and results are always calibrated against an international standard or derived .13 12 reference materialDetailed descriptions of the GC-c-IRMS design and operation are found elsewhere [ , 11] and on the ISOGEOCHEM5discussion list (www.uvm.edu∼geology/geowww/isogeochem.html).We use two analytical columns to analyze PLFA as they give different separations (see Fig 1). The a-polar HP5-ms(Hewlett Packard) provides good separation of many PLFA specific for bacteria and the very polar BPX70 column

5




(Scientific Glass Engineering) is better suited for poly-unsaturated PLFA as found in eukaryotes like fungi and algae, butis also used to check the results from the a-polar column. Other companies sell columns with similar characteristics. GCconditions used with the two columns can be found in Box 1.Box 1GC ConditionsOur GC-c-IRMS consists of a Hewlett Packard HP G1530 GC with flow control and equipped with a split/split-less injectorthat is used in the split-less mode. The GC is connected to Thermo Finnigan Delta-plus IRMS via a type III combustioninterface also from Thermo Finnigan (Bremen, Germany).The GC-method used with the a-polar HP 5MS column (60 m length, 0.32 mm diameter, 0.25 μm film) is: Columnpressure: 18 psi. Column flow: 2.2 ml/minute. Injector temperature: 280 °C. Temperature program: initial 60 °C for 2minutes, then to 130 °C with 25 °C/minute, to 280 °C with 3 °C/minute, to 300 °C with 25 °C/minute and hold 3 minutes.Split-less period: 1.5 minutes. Total rum time: about 55 minutes.And for the very polar BPX 70 column (50 m length, 0.32 mm diameter, 0.25 μm film): Column pressure: 14 psi. Injectortemperature: 240 °C. Temperature program: initial 60 °C for 2 minutes, then to 110 °C with 25 °C/minute, to 230 °C with 3°C/minute, to 250 °C with 25 °C/minute and hold 3 minutes. Split-less period: 1.5 minutes. Total rum time: about 45minutes.

>Calculations and data treatment

Identification of PLFA

There are generally about 40 peaks in the PLFA chromatograms and correct identification is an important issue. This isespecially true for many of the smaller peaks, which are often specific for certain groups of microbes. To reach a fullidentification, analysis of the samples on different columns, mass spectrometry and special derivatisation methods areoften necessary (see 2.3). However, identification based on retention from a single column is mostly sufficienttime datafor the more abundant PLFA. Table gives relative retention times for some of the dominant PLFA as found by GC-c-IRMSon the HP5-ms column. These relative retention times can be used as a first guide and may vary somewhat depending onthe GC-configuration used (see box 1 for the system that we use). Mixtures of FAMEs for calibration of retention timesand checking GC performance can be bought from for instance Supelco (www.sigmaaldrich.com/Brands/Supelco_Home.html). Pure cultures of the microbes of interest for which published or PLFA compositions are available alsoFAMEprovide a good source for reference compounds not readily available from commercial sources. A simple mixture ofseveral saturated FAMEs that have been individually analyzed for δ C ratios is also very useful for daily checks on GC13

performance and isotope analysis. Relative retention times for the different PLFA in Table 1 are given in a simplified formof equivalent (ECL), which uses 12:0, 16:0 and 19:0 as retention time markers. The PLFA 12:0 and 19:0 arechain lengthadded as internal standards and 16:0 is always present in samples as one of the main peaks. Relative retention times areused for first identification, as they are less sensitive for small variations in actual retention times between runs. ECL forPLFA with retention times between 12:0 and 16:0 are calculated as:

Table 1. Results of PLFA analysis by GC-c-IRMS of the same sediment sample labeled with C amino acids as in Fig13

1A. Relative retention times given in ECL are for the HP5 ms column. Numbers refer to the peaks in Fig 1. δ C ratios13

are for FAMEs and have not been corrected for the added during derivatizationmethyl group

PLFA ECL Nr δ C ‰13

12:0 12.00 1 −28.5

i14:0 13.54 2 −10.5

14:0 13.91 3 −22.4

i15:0 14.56 4 58.9

a15:0 14.65 5 23.1

15:0 14.94 6 −1.2

i16:0 15.60 7 −16.1

16:1ω9c 15.73 −12.6

6




16:1ω7c 15.79 8 −13.1

16:1ω7t 15.82 −31.6

16:1ω5c + t 15.88 43.2

16:0 16.00 9 −13.5

i17:1ω7c 16.40 89.2

10Me16:0 16.44 −33.2

i17:0 16.64 100.0

a17:0 16.73 4.0

17:1ω8c 16.78 −11.5

17:1ω6c 16.86 56.7

17:0 17.02 −0.7

18:2ω6c 17.71 10 −23.9

18:1ω9c 17.77 11 −19.9

18:1ω7c 17.83 12 −3.2

18:0 18.02 13 −16.6

cy19:0 18.88 14 −23.5

19:0 19.00 15 −32.5

20:4ω6 19.32 16 −22.5

20:5ω3 19.39 17 −22.0

20:0 19.94

22:6ω3 21.08 18 −25.8

22:5ω3 21.21 19 −25.8

22:0 21.73

24:0 23.40

(1)

And for PLFA with longer retention times than 16:0 as:

(2)

Box 2PLFA nomenclatureFatty acids are designated as A:BwC, where A is the number of carbon atoms, B is the number of double bounds and C isthe position first of the double bound from the aliphatic (ω) end. The prefixes "i" and "a" refer to iso- and anteiso-methylbranching, respectively. An iso-methyl branch is situated on the second carbon atom from the aliphatic end, and ananteiso-branch on the third carbon atom. Mid-chain methyl-branches are designated by "Me", preceded by the position ofthe branch from the carboxylic end. A cyclopropyl ring is indicated as "cy". See also [ 15].

PLFA concentrations

Concentrations of PLFA (C) are calculated from the peak area (A) relative the area of the internal 19:0 standard:

(3)

Where C is the amount of carbon in the 19:0 internal standard (1.55 μg C), f is the fraction of the chloroform19:0recovered during lipid extraction (see 2.2.1 step D) and n is the number of carbon atoms in the PLFA.

Isotope data

7




Usually, stable isotope ratios (R) as measured by GC-c-IRMS are given in the δ-notation, which for carbon is defined as:

(4)

The international standard for carbon is Vienna PeeDeeBelemnite (δ C = 0 ‰, R = C/ C-ratio =13standard

13 12

0.0112372). A major advantage of GC-c-IRMS is that very small changes in stable isotopic composition can be detected.The typical precision that is obtained with a GC-c-IRMS system for as little as one nmol carbon is about 0.3 ‰, whichmeans that changes in relative C content of about 0.001 % can be detected. This precision is necessary to study the13

natural variation in C/ C-ratios for which these machines were developed and where the maximum range of isotopic13 12

ratios is usually 20 ‰ or less, but it also means that very low incorporation of label can be detected in tracer studies.Label additions in experiments utilizing GC-c-IRMS methodology can therefore be minimized to concentrations close to orbelow those found in natural environments [ , ].4 6For natural abundance work it is essential that completely separation is achieved between compounds in the GC-c-IRMSchromatogram, because the whole peak must be integrated as isotopic effects on the analytic column cause an isotopicvariation over the peak [ 11]. This problem with not-well-resolved peaks (see Fig 1) is less sever with labeling work, asone is interested in increases in C content compared to a unlabeled, control sample and the range in C-ratios is13 13

larger. It should be regarded as possible label carry over between compounds, which is a function of the overlap and therelative size of the peaks. In my experience, it becomes a problem with minor peaks poorly resolved from major peaks.To obtain the actual PLFA isotope ratio, carbon-isotope ratios of as measured by GC-c-IRMS have to be correctedFAMEfor the one carbon atom in the that has been added during derivatisation:methyl group

(5)

Where n is the number of carbon atoms in the PLFA. The isotopic composition of the methanol (δ ) that was usedMeOHfor derivatisation has to be determined separately for instance by GC-c-IRMS and is usually rather depleted (−40 to −50‰).The δ-notation is based on isotope ratios, which is not very convenient for enriched samples [ ]. Increases in isotope5ratios (Δδ-ratios) that are obtained in tracer work should be regarded as equivalent to increases in specific labeling, anddo therefore not directly indicate the absolute amount of label that was incorporated into a certain biomarker [ ].2Increases in isotope ratios however do provide information on the relative distribution of the label in the different PLFAand therefore indicate if label uptake was either evenly distributed in the microbial community or that specific groups ofmicrobes were active.Absolute amounts of label incorporated ( C), also called excess C, are calculated from the product of biomarker13 13

concentration (C) and the increase in the fraction C after labeling (F ) relative to the control (F ):13 13t

13c

(6)

The fraction C can be calculated from the C/ C-ratios (R) as:13 13 12

(7)

And R is calculated from the δ C ratios as measured with the IRMS equipment using the reverse of equation 4:13

(8)

Another effect of measuring isotope ratios rather than absolute amounts by GC-c-IRMS is that the detectable labelincorporation also depends on pool sizes or concentrations of the components to be analyzed. For instance, labelingsediment with 10 μM C-acetate gave a readily detected increase in δ C ratio of about 10 ‰ in some PLFA [ ], but13 13 4would not be detectable in the much larger sediment organic matter pool due to dilution.

Data analysis

The resulting C-labeling patterns of PLFA as shown for instance in Fig 2 can be analyzed in three ways. First, specific13

8




biomarker PLFA are known for certain groups of microorganisms and labeling of these compounds indicates that thesegroups were active in the degradation of the label. Specific PLFA are for instance known for various genera of sulfatereducing and methane oxidizing bacteria. Major groups of microorganisms like bacteria, fungi and various algae can alsobe separated by PLFA. A further discussion on these specific PLFA can be found in the literature (e.g. [ , 9, 13, 16]).3Second, the whole labeling pattern can be compared with PLFA or fatty acid compositions from bacterial isolates [10].Fatty acid analysis is a common taxonomic method to characterize bacterial isolates and a large data set is available [15].The main advantage of this approach is that also information from more common fatty acids is used to reachidentification. Boschker et al [ ] for instance used cluster analysis compare C-acetate labeling patterns of sediments4 13

with PLFA compositions of sulfate reducers to indicate the most likely active population. Third, the labeling patterns canbe analyzed by various multivariate like principle component analysis to study relationships betweenstatistical techniqueslabeling patterns from different sites or labels. Data are usually log-transformed to put more weight on less abundantcompounds. This technique is also used to compare PLFA concentration profiles from different sites or treatments [ 9].

Figure 2 Incorporation of C -acetate ( Fig 2A) and 3- C -propionate ( Fig 2B) into PLFA in an anoxic sediment. Shown is the132

131

percentage of the C-label recovered in the individual PLFA. After [ ].13 2

Application of the method

An example is taken from Boschker et al [ ], who studied bacterial populations involved in the degradation of acetate and2propionate in anoxic sediment. The mineralisation of organic matter in anoxic sediments is a stepwise process, in whichseveral low-molecular intermediates produced by fermentative bacteria play an important role. Acetate and propionate areamong the most important of these intermediates and are consumed by organisms like sulfate-reducing bacteria.

9




Boschker et al [ ] took undisturbed sediment cores at the Grosser Jasmunder , a brackish bay in the German2 BoddenBaltic Sea. Small sub-cores (diameter 2.4 cm) were taken from larger cores and injected through side holes at 0.5 cmintervals with uniform labeled C-acetate or 3- C-propionate to a final concentration of 100 μM. Control incubations to13 13

which no label was added were run in parallel. Triplicate cores for all treatments were incubated at in-situ temperature (12°C) for 8 hours, after which all label was consumed. The anoxic 1 to 3 cm layer of the sediment was collected andanalyzed for PLFA concentrations and label content as described in this chapter.The C-label from acetate was mainly traced in even-numbered PLFA (16:1ω7c, 16:1ω5, 16:0 and 18:1ω7c, Fig 2A).13

The acetate-labeling pattern resembled PLFA compositions of and recently isolated Desulfotomaculum acetoxidans strains, which are both acetate consuming sulfate reducers, suggesting that one of these organisms wasDesulfofrigus

the dominant, active population. Typical biomarkers for sulfate reducing bacteria like i17:1ω7c, 10Me16:0 and 17:1ωbcontained only minor amounts of label, which suggested that bacteria belonging the genus , orDesulfovibrio Desulfobacter

were not involved in the consumption of acetate.DesulfobulbusThe C-propionate-labeling pattern ( Fig 2B) was almost completely different from the acetate labeling pattern with major13

amounts of label in odd-numbered PLFA (a15:0, 15:0, 17:1ω6 and 17:0). This strongly suggested that different bacterialpopulations were involved in the consumption of acetate and propionate. The complete labeling pattern with propionatedid not resemble any of the known strains, which suggests that it belonged to an unknown type of sulfate . Thisreducerwas one of the first studies where was directly shown in complex microbial communities.niche differentiation

References

7-1. Bligh EG, Dyer WJ (1959) A rapid method of total lipid extraction and purification. Can J Biochem Physiol 31:911-917.7-2. Boschker HTS, De Graaf W, Koster M, Meyer-Reil LA, Cappenberg TE (2001) Bacterial populations andprocesses involved in acetate and propionate consumption in anoxic brackish sediment. FEMS Microbiol Ecol 35:97-103.7-3. Boschker HTS, Middelburg JJ (2002) Stable isotopes and biomarkers in microbial ecology. FEMS MicrobiolEcol 40: 85-95.7-4. Boschker HTS, Nold SC, Wellsbury P, Bos D, De Graaf W, Pel R, Parkes RJ, Cappenberg TE (1998) Directlinking of microbial populations to specific biogeochemical processes by C-labelling of biomarkers. Nature 392:13

801-805.7-5. Brenna JT, Corso TN, Tobias HJ, Caimi RJ (1997) High-precision continuous-flow isotope ratio massspectrometry. Mass Spectrom Rev 16: 227-258.7-6. Bull ID, Parekh NR, Hall GH, Ineson P, Evershed RP (2000) Detection and classification of atmosphericmethane oxidizing bacteria in soil. Nature 405: 175-178.7-7. Cottrell MT, Kirchman DL (2000) Natural assemblages of marine proteobacteria and members of the

cluster consuming low-and high-molecular-weight dissolved organic matter. Appl EnvironCytophaga-FlavobacterMicrobiol 66: 1692-1697.7-8. Dobbs FC, Findlay RH (1993) Analysis of microbial lipids to determine biomass and detect the response ofsedimentary microorganisms to disturbance. In Kemp PF, Sherr BF, Sherr EB, Cole JJ (eds) Handbook ofMethods in Aquatic Microbial Ecology, pp. 347-358. Lewis Publishers, Boca Raton.7-9. Findlay RH & Dobbs FC (1993) Quantitative description of microbial communities using lipid analysis. In:Kemp PF, Sherr BF, Sherr EB & Cole JJ, eds, Handbook of Methods in Aquatic Microbial Ecology, pp 271-284,Lewis Publishers, Boca Raton.7-10. Hanson JR, Macalady JL, Harris D, Scow KM (1999) Linking toluene degradation with specific microbialpopulations in soil. Appl Environ Microbiol 65: 5403-5408.7-11. Hayes JM, Freeman KH, Popp BN, Hoham CH (1990) Compound-specific isotope analysis: a novel tool forreconstruction of ancient biogeochemical processes. Org Geochem 16: 1115-1128.7-12. Middelburg JJ, Barranguet C, Boschker HTS, Herman PMJ, Moens T, Heip CHR (2000) The fate of intertidalmicrophytobenthos carbon: An C-labeling study. Limnol Oceanogr 45: 1224-1234.in situ 13

7-13. Parkes RJ (1987) Analysis of microbial communities within sediments using biomarkers. In Hetcher M, GrayRTG, Jones JG (eds) Ecology of microbial communities, pp. 147-177. Cambridge University Press, Cambridge.

10




7-14. Pombo SA, Pelz O, Schroth MH & Zeyer J (2002) Field-scale C-labeling of phospholipid fatty acids (PLFA)13

and dissolved inorganic carbon: tracing acetate assimilation and mineralization in a petroleumhydrocarbon-contaminated aquifer: FEMS Microbiol. Ecol. 41: 259-267.7-15. Ratledge C, Wilkinson SG (1989) Microbial lipids. Academic Press, San Diego.7-16. Tunlid A, White DC (1992) Biochemical analysis of biomass, community structure, nutritional status, andmetabolic activity of microbial communites in soil. In Bollag JM, Stotzky G (eds) Soil Biochemistry, pp 229-262.Marcel Dekker, New York.7-17. Zhang CLL (2002) Stable carbon isotopes of lipid biomarkers: analysis of metabolites and metabolic fates ofenvironmental microorganisms. Cur Opin Biotechnol 13: 25-30.

Linking microbial community structure and functioning: stable isotope (13C) labeling incombination with PLFA analysis

H. T. S.Boschker

Netherlands institute of Ecology (NIOO-KNAW), Heteren, the Netherlands

DOI: 10.1007/SpringerReference_76362

URL: http://www.springerreference.com/index/chapterdbid/76362

Part of: Molecular Microbial Ecology Manual

Editors:Antoon D. L. Akkermans (deceased), Frans J. de Bruijn, Iian M. Head, Prof. GeorgeA. Kowalchuk and Jan Dirk van Elsas

PDFcreated on:

December, 12, 2012 11:34


springerreference || section 8 update: linking microbial community structure and functioning: stable...

Documents