Vol. 58, No. 12APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Dec. 1992, p. 4026-40310099-2240/92/124026-06$02.00/0Copyright © 1992, American Society for Microbiology

Soil Bacterial Biomass, Activity, Phospholipid Fatty AcidPattern, and pH Tolerance in an Area Polluted

with Alkaline Dust DepositionE. BAATH,l* A. FROSTEGARD,1 AND H. FRITZE2

Department ofMicrobial Ecology, Lund University, Helgonavagen 5, S-223 62 Lund, Sweden,and Finnish Forest Research Institute, Department ofForest Ecology,

P.O. Box 18, SF-01301 Vantaa, Finland2

Received 23 June 1992/Accepted 29 September 1992

Soil bacterial biomass, phospholipid fatty acid pattern, pH tolerance, and growth rate were studied in aforest area in Finland that is polluted with alkaline dust from an iron and steel works. The pollution raised thepH of the humus layer from 4.1 to 6.6. Total bacterial numbers and the total amounts of bacterial phospholipidfatty acids in the humus layer did not differ between the unpolluted control sites and the polluted ones. Thenumber ofCFU increased by a factor of 6.4 in the polluted sites compared with the controls, while the bacterialgrowth rate, measured by the thymidine incorporation technique, increased about 1.8-fold in the polluted sites.A shift in the pattern of phospholipid fatty acids indicated a shift in the bacterial species composition. Thelargest proportional increase was found for the fatty acid 10Me18:0, which indicated an increase in the numberof actinomycetes in the polluted sites. The levels of the fatty acids i14:0, 16:1w5, cy17:0, 18:1w7, and 19:1 alsoincreased in the polluted sites while those of fatty acids 15:0, i15:0, 1OMel6:0, 16:1w7t, 18:109, and cyl9:0decreased compared with the unpolluted sites. An altered pH tolerance of the bacterial assemblage was detectedeither as a decrease in acid-tolerant CFU in the polluted sites or as altered bacterial growth rates at differentpHs. The latter was estimated by measuring the thymidine incorporation rate of bacteria extracted from soilby homogenization-centriftigation at different pHs.

In a study of the response of soil microbes to emissionsfrom an iron and steel works (9), it was found that themicrobial biomass in the humus horizon of the forest wassimilar in polluted and unpolluted areas if calculated on anorganic-matter basis. Since there was either an increase infungal biomass, measured as total hyphal length (9), or amore or less unchanged fungal biomass, measured as ergos-terol content (10), it appeared that the bacterial biomassmust have decreased or been unaffected by the pollution.This was unexpected, since the main effect of the emissionwas a raised pH, from about pH 4.1 in unpolluted sites to 6.6in polluted ones. It is commonly accepted that bacteria,unlike fungi, are better adapted to a neutral soil pH than amore acidic pH.A drastic change in the microfungal species composition in

the polluted sites was found earlier (10). The extent ofchanges in the bacterial community structure is difficult tostudy, since this usually involves isolation and identificationof bacteria that can grow in pure culture on agar plates.Since these bacteria constitute only a minor part of the totalcommunity (usually about 2 to 4%) (24), we know little aboutthe other, larger part. One way to overcome this problem isto study the phospholipid fatty acid (PLFA) pattern of a soil(29). Since different groups of bacteria are characterized bymore or less specific PFLA profiles, an altered PLFA patternin a soil would indicate an altered species composition.A further indication of a shift in the species composition

can be a changed tolerance pattern of the organisms. Forbacteria, this is usually also studied with organisms that canbe isolated and grown on agar plates. However, a techniqueto measure bacterial growth rates in soil by thymidine

* Corresponding author.

incorporation by bacteria extracted from soil by homogeni-zation-centrifugation was recently described (2). By chal-lenging the extracted bacteria with different concentrationsof heavy metals before measuring their growth rate, ameasure of the heavy metal tolerance of the bacterial com-munity could be made (3). In this way, a Cu-tolerantcommunity was shown to have evolved in a Cu-polluted soil.In the present study, the emissions from the iron and steelworks have resulted in an increased soil pH which hasprobably lasted for about 25 years. One would thereforeexpect a bacterial community with an altered pH tolerancepattern to have evolved here. One objective of the study wastherefore to adapt the thymidine incorporation technique (3)to study the pH tolerance of the bacterial community inthese soils. This article describes the effect of increased pHon the biomass, growth rate, PLFA pattern, and pH toler-ance of the soil bacterial community.

MATERIAIS AND METHODS

Site description and soil sampling. The Koverhar iron andsteel works is located on the Baltic Sea coast in the southernpart of Finland. It was established in 1961. Twenty of 24previously established sites (9) were sampled in September1991. Ten of the sites were influpnced by the deposition ofalkaline dust, while the remaining io sites served as controls.Within each group, five sites were situated south and fivesites were situated north of the factory. The prevailing winddirection is from the south. All the study sites were locatedin a dry Calluna-type Scots pine (Pinus sylvestris L.) forest.For more chemical data on the soil, see reference 9.From each site, 10 individual core samples (AO1/A02 layer)

were combined into a bulk sample. The 20 bulk soil sampleswere then sieved (mesh size, 4 mm), and visible plant

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material was removed before storage at 4°C for 4 to 8 weeks,depending on the analysis. Organic matter was determinedas loss on ignition (LOI) by heating oven-dried (105°C)samples to 550°C for a minimum of 4 h.

Bacterial counts. A 2.5-g (wet weight) amount of soil washomogenized with 200 ml of prefiltered (0.22 ,um) 0.2%Calgon solution (sodium polymetaphosphate) for 1.5 min in aSorvall Omnimixer. CFU were determined on 0.2% tryptonesoy agar (TSA) plates with 50 mg of cycloheximide per literafter appropriate dilution in 0.1% peptone. CFU werecounted after 14 days at 20°C. Five plates were prepared perdilution, and the dilution with 20 to 80 colonies per plate wasused.Samples for total bacterial counts were stored in 2.5%

Formalin until bacteria were counted. The samples wereadded to 9 ml of prefiltered (0.22 p,m) acetate buffer (pH 4.0,150 mM) with 180 pl of acridine orange (0.1%). Filtrationafter 5 min of staining was carried out with 0.2-pum blackNucleopore filters. The filters were washed three times with5 ml of prefiltered buffer. Bacteria were then counted underthe fluorescence microscope. Two replicate filters were usedfor each soil sample, and more than 300 bacteria werecounted per sample by randomly selecting microscopic fieldscovering the whole filter area. Data for bacterial numberswere expressed per LOI in order to relate them to theorganic matter of the soil. Thus, no effect of increased soilweight due solely to pollution of the soil with mineral dustwould be introduced into the calculations.

Bacterial activity. Bacterial growth rates were determinedby the thymidine incorporation technique on bacteria ex-tracted from the soil by homogenization-centrifugation (2).The method can be summarized briefly as follows. One gram(wet weight) of soil was homogenized for 1 min in 200 ml ofdistilled water at 80% full speed in a Sorvall Omnimixer. Thesuspension was then centrifuged for 10 min at 750 x g, andthe supernatant was filtered through glass wool to removesmall organic particles floating on the meniscus of the water.Two milliliters of the resulting bacterial suspension wasincubated with 200 nM methyl-[3H]thymidine (Amersham;25 Ci/mmol, 925 GBq/mmol) at 20°C for 2 h before 1 ml of 5%Formalin was added. Formalin was added to time zerocontrols before addition of the label. The suspension wasthen filtered through Whatman glass fiber filters (GF/F)prewashed with 1% Calgon and washed three times with 5 mlof 80% ice-cold ethanol and three times with 5 ml of ice-cold5% trichloroacetic acid. Radioactivity was counted by usinga Beckman Ready Safe in a Beckman LS 1801 liquidscintillation spectrometer after macromolecules were solu-bilized for 1.5 h in 1 ml of 0.1 M NaOH at 90°C. Quenchingwas corrected with Beckman's H#, an external standardmethod. The number of bacteria in the solutions was deter-mined by acridine orange staining (see above). Only thymi-dine incorporation into macromolecules was measured,since no attempt was made to separate radioactivity incor-poration into RNA, DNA, and protein.The extents of an exogenous pool of thymidine other than

that added and of de novo synthesis were determined by theisotope dilution approach (27). No differences in isotopedilution were found between control and polluted samples,and therefore a mean value of 360 nM was used in thecalculations.pH tolerance. To test the pH tolerance of bacteria, 0.2%

TSA plates were prepared with pH 6.4 and a low pH (4.3).Low-pH plates were prepared by adding HCl.pH tolerance was also determined by using thymidine

incorporation in a similar manner to the way that heavy

metal tolerance was measured by Baath (3). Before radioac-tive thymidine was added the pH of the bacterial suspensionwas varied between 3.5 and 8.5 by using buffers. Distilledwater was added to controls. For pHs of about 3.5 and 4, acitrate-potassium phosphate buffer (3.3 and 6.6 mM citricacid and K2HPO4, respectively, final concentrations) was

used, and for pHs of about 5, 6, 7, 8, and 8.5, a potassiumphosphate buffer (6.6 mM, final concentration) was used.These measurements were made on two occasions withthree different soil samples from each treatment each time.In the first measurement, no citrate-phosphate buffer was

used; instead, the pH of the bacterial solution was changedto 4 with diluted HCI. The actual pH of the soil solutions wasmeasured after the addition of buffers or distilled water, andthese pHs are reported in Results.PLFA. Lipids were extracted by a procedure described

previously (12), which is based on the method of Bligh andDyer (6). Briefly, 0.50 g of fresh soil was extracted for atleast 2 h with a one-phase mixture consisting of chloroform,methanol, and citrate buffer (0.15 M, pH 4.0) in the propor-tions 1:2:0.8 (by volume). After centrifugation, the pelletswere washed once with the one-phase mixture, and thesupernatants were combined. The supernatants were splitinto two phases by adding chloroform and citrate buffer, and1 ml of the lower phase was sampled and dried undernitrogen. The lipid material was fractionated into neutral,glyco-, and polar lipids on silicic acid (100-200 mesh; Unisil)columns by elution with chloroform, acetone, and methanol,respectively. The phospholipid-containing polar fraction wascollected and dried under nitrogen. Methyl nonadecanoatewas added to the phospholipid fractions as an internalstandard. The samples were then subjected to mild alkalinemethanolysis (7). The resulting fatty acid methyl esters wereanalyzed on a Hewlett Packard 5890 gas chromatographequipped with a flame ionization detector and a 50-m HP1capillary column. The preparation of the glassware and thesolvents used were described previously (12).

Fatty acids are designated as total number of carbonatoms: number of double bonds, followed by the position ofthe double bond from the methyl end (X) of the molecule. cisand trans configurations are indicated by c and t, respec-tively. The configuration of the double bond was onlydetermined for 16:1X7. The prefixes a and i indicate anteiso-and isobranching, respectively; lOMe indicates a methylgroup on the 10th carbon atom from the carboxyl end of themolecule; and cy refers to cyclopropane fatty acids.

Statistical analyses. The PLFA pattern was determined forsamples from all 20 sites. Total bacterial counts and bacterialactivity at different pHs were determined for samples fromsix polluted and unpolluted sites each, while recoverablebacteria were only determined for samples from three sitesper treatment.

Differences between measurements were determined byStudent's t test. The PLFA pattern (expressed as molepercent) was analyzed by principal-component analysis withthe program SIRIUS (18). Samples from polluted and unpol-luted sites were also analyzed separately by partial least-squares regression (21) to study whether any differencescould be detected in the PLFA pattern due to samplingdirection (south or north of the factory). Exchangeablecations (Mn, Ca, Al, Mg, Fe, Zn, and K; see reference 9 formethods) were used as independent variables, and PLFAswere used as dependent variables. The soil chemical datahave previously been shown to closely mirror the degree ofpollution (9).To calculate the total amount of bacterial fatty acids, those

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TABLE 1. Characteristics of bacteria and soil from polluted and unpolluted sitesa

LOI/g Total AOC CFU CFU/AO CFUaCr d CFUd/C Bacterial ThymidineSamplesite pHb ~dry wt (1010/g LOI) (108/g LOI) (% 10/ LOT) (% (nmol/g LOI) (10-21 mol/cell/h)

Control 4.13 ± 0.19 62.5 ± 7.4 2.16 ± 0.22 3.39 ± 0.95 1.6 2.21 ± 0.81 62.7 ± 12.1 926 ± 38 26.9 ± 5.1Polluted 6.60 ± 0.41* 28.1 ± 2.9* 2.39 ± 0.19 21.9 ± 0.36* 9.2 2.17 ± 0.75 9.2 ± 2.3* 955 ± 67 47.5 ± 3.5*

Ratio, polluted! NAf NA 1.1 6.4 NA 1.0 NA 1.0 1.8control

a Values are means ± standard error. *, significantly different from control (P < 0.05).b From Fritze (9).AO, acridine orange-stained bacteria.

d CFUacid, CFU on acidified agar plates.I Incorporation into total macromolecules was measured.f NA, not applicable.

suggested to be mainly of bacterial origin (8, 28) weresummed. These fatty acids were ilS:0, alS:0, 15:0, i16:0,16:1w9, 16:17t, 16:1w5, 17:1X8, i17:0, a17:0, cy17:0, 17:0,18:1M7, and cyl9:0. We also considered lOMel6:0, lOMe17:0, and lOMel8:0 to be bacterial fatty acids.

RESULTS

The bacterial characteristics of the polluted and unpol-luted soils are summarized in Table 1. The total number ofbacteria did not differ between the polluted and unpollutedsites, and the total amount of bacterial PLFAs was alsosimilar. This indicated that the bacterial biomass was af-fected to only a minor degree by the increased pH. Thenumber of recoverable bacteria (CFU) increased, however,in the vicinity of the steel works to 6.4 times the numberfound in the control sites. Thus, the percentage of culturablebacteria increased from 1.6% of total bacteria in the acidcontrol soil to more than 9% of total bacteria in the pollutedsoils with more neutral pH.The bacterial growth rate, as measured by the thymidine

incorporation method, was 26.9 x 10-21 mol of thymidineincorporated into total macromolecules per bacterial cell perh in the control samples, but this increased 1.8-fold in thepolluted sites, to 47.5 x 10-21 (Table 1).Twenty-nine different PLFAs, most of them identified,

were used for comparing polluted and unpolluted sites. Ashift in the species composition of the bacterial communitywas evident when the PLFA patterns of samples from thetwo kinds of site were compared by a principal-componentanalysis (Fig. 1). The different sites separated completelyalong the first axis, which accounted for 47.1% of thevariation in the data. Control samples are found to the rightin the plot, and polluted samples are found to the left. Thesecond component accounted for 23.0% of the variation andappeared not to be related to the pollution. The largestproportional increase in the polluted sites was found for thefatty acid lOMel8:0; more than twice as much was present asin the control sites. PLFA i14:0, 16:1w5, cy17:0, 18:107, and19:1 levels also increased in the polluted sites (Table 2). Thelargest decrease in the polluted sites compared with theunpolluted ones was found for the fatty acid ilS:0, but 15:0,lOMel6:0, 16:17t, 18:1X9, and cyl9:0 levels also decreasedwhen the soil pH increased.When data for samples from the polluted and unpolluted

sites were treated separately by partial least-squares regres-sion, data for the sites to the north and south of the factorywere partly separated (data not shown). This was mainly dueto slightly higher levels of Ca, Zn, Mg, and Mn in the sites tothe north of the factory, indicating that the pollution was

more evident downwind than upwind from the factory.However, no differences in the PLFA pattern in areas northand south of the factory could be seen for either the controlor polluted sites.An altered bacterial community was also evident from

changes in the pH tolerance of the bacteria extracted fromthe soil by homogenization-centrifugation (Fig. 2). In thecontrol samples, thymidine incorporation was highest whenthe pH was buffered to ca. 5 to 6, similar to the thymidineincorporation measured when distilled water was used.When the pH was changed from these values, thymidineincorporation decreased. At pH 3.5, it was only 35% of thatin distilled water, and at pH 8.5, it was only about 30%. Inthe polluted soils with a more neutral pH, the pH optimumwas about 6 to 7. An increase to pH 8.5 had only minoreffects (90% of the value found in distilled water), while adecrease in the pH to 3.5 gave a drastic decrease in thymi-dine incorporation, to less than 5% of the optimum value.There was also a shift in the pH tolerance of the culturable

bacteria in the polluted sites, although the number of colo-nies on acidified plates did not differ between samples fromthe two kinds of site (Table 1). However, since morebacterial colonies were found on the normal (neutral) agarplates of samples from the polluted sites, the percentage ofbacteria on acid agar plates compared with that on normalagar plates (CFUaCid/CFU) was less than 10% for the pol-

A

A AA

AA

0*ILI

PC 1

FIG. 1. Principal-component (PC) analyses of the PLFAs inpolluted (A) and unpolluted (0) soils around the Koverhar steelfactory in Finland.

A

A 0 0

A

0

A

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TABLE 2. PLFAs in control and polluted soil samples

Mean mol% of total PLFAs + SE'PLFA Control Polluted Ratio, polluted/

samples samples control

i14:0 0.28 ± 0.02 0.63 ± 0.10* 2.2514:0 1.19 ± 0.11 1.40 ± 0.18 1.17i15:0 6.53 ± 0.40 3.40 + 0.24* 0.52alS:0 2.07 ± 0.13 2.53 ± 0.19 1.23brlS:0 0.88 ± 0.06 1.14 ± 0.10* 1.3015:0 1.55 + 0.26 1.15 + 0.14 0.74i16:0 2.40 ± 0.13 1.97 ± 0.07* 0.8216:1w9 0.79 ± 0.07 0.90 ± 0.07 1.1316:lw7c 7.48 ± 0.18 7.60 ± 0.25 1.0216:lw7t 1.19 ± 0.03 0.75 ± 0.02* 0.6316:lw5 1.72 ± 0.08 3.46 ± 0.11* 2.0116:0 16.78 ± 0.24 18.32 ± 0.27* 1.09lOMel6:0 4.86 ± 0.20 2.93 ± 0.10* 0.60i17:0 0.67 + 0.03 0.89 ± 0.03* 1.33a17:0 0.97 + 0.02 1.52 ± 0.05* 1.5717:1w8 0.97 ± 0.02 1.24 ± 0.03* 1.28cyl7:0 1.71 ± 0.04 3.01 ± 0.08* 1.7617:0 0.79 ± 0.02 0.95 ± 0.02* 1.20lOMel7:0 0.60 ± 0.03 0.70 ± 0.04* 1.1818:2X6 18.67 ± 0.85 15.87 ± 0.49* 0.8518:1X9 10.88 ± 0.35 7.33 ± 0.20* 0.6718:1M 7 6.69 ± 0.22 12.01 ± 0.34* 1.8018:1 0.63 ± 0.04 0.75 ± 0.05 1.2018:0 2.03 ± 0.04 2.63 ± 0.22* 1.3019:1a 0.43 ± 0.01 0.83 ± 0.04* 1.92lOMel8:0 0.59 ± 0.03 1.36 ± 0.07* 2.3119:1b 1.65 ± 0.09 1.47 ± 0.15 0.90cyl9:0 4.05 ± 0.14 2.15 ± 0.10* 0.5320:0 0.98 ± 0.05 1.14 ± 0.08 1.16

a *, significantly different from control (P < 0.05).

luted sites, compared with more than 60%sites.

for the control

DISCUSSION

The effects of alkaline pollution on the bacterial commu-nities were very similar to the effects found after liming of aforest soil. After liming, an increase in the number ofrecoverable bacteria (1, 16, 19, 20, 22) but not in the numberof total bacteria determined by direct counting (4) is alsocommonly found. Increased microbial activity is also usuallydetected as an increase in the soil respiration rate afterliming (1, 15, 16, 22, 25, 31). We found increased bacterialactivity, since the specific thymidine incorporation ratealmost doubled in the polluted soil compared with that in thecontrol soil (Table 1), indicating an increased bacterialgrowth rate.The specific thymidine incorporation rates were 26.9 x

10-21 and 47.5 x 10-21 mol per bacterial cell per h in controland polluted soils, respectively (Table 1). These are some-what higher than the values reported by Baath (2) for a limed(pH 6.1) and unlimed (pH 3.8) forest humus. However,Baath (2) studied thymidine incorporation into DNA aloneand not into total macromolecules, as in the present study.Since thymidine incorporation into DNA was 37.8% of thatinto total macromolecules (2), recalculation of his datawould give 11.5 x 10-21 and 45.0 x 10-21 mol of thymidineincorporated into total macromolecules per bacterial cell perh, which is more similar to the values found in the presentstudy. With a conversion factor of 1018 bacterial cells formedper mol of thymidine incorporated into total macromolecules

(2), a bacterial turnover rate at 20°C of 1.5 days could becalculated for the control soils. In the polluted areas, theturnover time decreased to 0.9 days.The changes in the PLFA pattern were also almost iden-

tical to those found after liming or wood ash treatment ofconiferous forest soils (11). Especially characteristic of anincreased pH was the higher proportions of the fatty acids16:1X5 and 10Mel8:0, while 10Mel6:0, 18:1X9, and cy19:0levels decreased compared with the levels in the control soils(Table 2). Other PLFAs that were less affected, but in similarways in these two studies, were i14:0, i17:0, a17:0, cyl7:0,and 18:1X7, which all increased with increasing pH, whilei15:0 and 16:17t were more abundant in the controls. Theinterpretation of the changes in the PLFA pattern is thussimilar to those reported by Frostegard et al. (11). Anincreased abundance of the fatty acid 10Me18:0 indicated anincreased abundance of actinomycetes in the polluted soils,since this fatty acid is considered unique to this bacterialgroup (17). The increase in 18:1X7 and cyl7:0, fatty acidscharacteristic of gram-negative bacteria (30), and the de-crease in i15:0, 10Me16:0, and 18:1w9, fatty acids character-istic of gram-positive bacteria (23), in the polluted sitescompared with the control sites indicate a shift in theabundance of these two bacterial groups due to the pollution.However, this interpretation is not straightforward, since thelevels of several branched fatty acids (i14:0, i17:0, anda17:0), which are typical in phospholipids from gram-posi-tive bacteria, also increased in the polluted sites comparedwith the unpolluted controls. The similarities between thechanges in the PLFA pattern after lime and wood ashtreatment (11) and the alkaline dust pollution studied in thepresent work support the conclusion by Fritze (9) that thepollution effects were due only to the changed pH, while theincreased metal concentrations had not affected the micro-biota in the area surrounding the steel factory. Furthermore,as suggested previously (11), it is possible that increased pHaffects the soil microorganism community, as estimated byits PLFA pattern, in a similar way in all forest humus.The increase in actinomycetes in the polluted sites could

be one explanation for the differences in fungal biomassestimations which were found when different techniqueswere used. Fritze (9) detected almost twice as much hyphaein polluted sites as in unpolluted ones, but no increase wasfound when ergosterol or the PLFA 18:2w6 was used as anindicator of fungal biomass (10). Actinomycete hyphae aredifficult to distinguish from thin fungal hyphae in soil, andthe increase in hyphal length in the polluted sites could thusbe due, at least partly, to an increase in actinomycetes.An altered PLFA pattern could be due not only to a shift

in the species composition, but also to changing environmen-tal conditions affecting the PLFA composition of the bacte-rial cell membranes. Thus, an increased ratio of trans to cisisomers of the fatty acids 16:1X7 and 18:1X7 has, for severalbacteria, been shown to indicate increased environmentalstress, especially due to nutrient starvation (13, 14). Thetrans/cis ratio of 16:1X7 was lower in the polluted sites (0.10)than in the control sites (0.16), which may indicate that thenutrient stress for the soil bacteria was more severe in theacid control soils than in the polluted soils with a moreneutral pH. This would also be expected, since an increasedpH is known to increase the availability of carbon tomicroorganisms (25), a fact that was also mirrored in theincreased bacterial growth rates (Table 1). A lower trans/cisratio for 16:1X7 was also found in limed soils (0.07) than inunlimed ones (0.12) (calculated from data in reference 11).An altered bacterial assemblage was revealed not only by

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1007-V

50

0~~ ~ ~ ~ ~ ~ 0

0

3 4 5 6 7 8pH

FIG. 2. Effect of different pHs on thymidine incorporation into total macromolecules of bacteria extracted by centrifugation-homogeni-zation from polluted (O, *) and unpolluted (0, 0) soils around the Koverhar steel factory in Finland. The pH was changed by adding variousbuffers to the bacterial suspensions. The data are expressed as a percentage of the values obtained in distilled water only. Solid and opensymbols denote separate experiments; the second (0, 0) was performed 2 months after the first (-, *). Bars indicate standard error (n =3 different soil samples).

the altered PLFA pattern but also by the effect of differentpHs on the growth rate of the bacteria extracted from thesoils (measured by the thymidine incorporation technique,Fig. 2). Bacteria from the unpolluted soil had a lower pHoptimum and less inhibition at low pH than bacteria ex-tracted from polluted soils. For the unpolluted-soil samples,the rate of thymidine incorporation decreased at pHs abovethe optimum (Fig. 2). This was not the case for the polluted-soil samples, for which approximately the same values werefound for pHs between 6 and 8.5. This might indicate thatalthough the bulk pH was 6.6, there existed microhabitatswith higher pH. This is not unlikely, since the pollution wasin the form of alkaline dust, and high pHs might be presentin the immediate surroundings of a dust particle.For the second measurement of bacterial tolerance, the

tolerance curves appeared to be shifted somewhat towardshigher pH, both for the control and for the polluted samples(Fig. 2). The reason for this might be that the secondmeasurement was made 2 months after the first one, andchanges could thus have occurred in the soil pH. It is wellknown that pH increases in humus soils after storage (26).The shift in tolerance curves might also be due to errors inthe method used, but the thymidine incorporation measure-ments were very reproducible, usually with a coefficient ofvariation of less than 5%. Since the measurements are fastand up to 100 samples can easily be processed in a day, thismethod promises to be a good tool with which to study thetolerance of soil bacteria. It might even be possible to studybacterial tolerance with this technique in microenviron-ments. This might give an indication of the pH conditions inhabitats where the actual pH is difficult to measure, such asthe rhizoplane.

An altered bacterial pH tolerance was also evident for theculturable bacteria (CFU, Table 1). The percentages ofbacteria on the acid agar plates and on the normal agar plates(CFUaCid/CFU), 60% for the control and 10% for the pollutedsoils, were very similar to the percentages calculated forbacterial tolerance at pH 4 compared with the optimumconditions by the thymidine incorporation technique (Fig.2). However, it was difficult to monitor the pH on the TSAplates, and the pH of the plates increased over time becauseof bacterial growth. Thus, after 6.5 weeks of incubation,there was little difference in the actual pH of the two types ofagar plates and consequently little difference in the numberof CFU.

In a study of the cell size and viability of soil bacteria (5),it was found that, although only a few percent of the totalbacteria could form colonies on agar plates, they constituteda much larger proportion of the total biomass because theseculturable bacteria usually represented the largest bacterialsize groups. It is also possible that these bacteria represent alarge proportion of the active bacteria, since increases inboth the number of CFU and the specific thymidine incor-poration rate were found in samples from polluted sites.However, the increased number ofCFU in the polluted soilsmight also be due to a changed species composition, as wasindicated by the altered PLFA pattern; bacterial species inthe polluted sites could be better able to form visiblecolonies.

In comparison with measuring pH tolerance by colonycounts on plates with different initial pHs, the use ofthymidine incorporation by bacterial suspensions buffered todifferent pHs has several advantages. Measurements can bemade within a short time interval, 2 h, during which no major

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changes in pH would be expected. Plates, on the other hand,have to be incubated for several weeks, and pH changes inthe vicinity of colonies are likely to occur, as in the presentstudy. The speed and accuracy of the thymidine incorpora-tion technique have been commented on above. Finally, thethymidine incorporation technique measures the pH toler-ance of all bacteria that can incorporate exogenous thymi-dine. These bacteria most likely constitute a larger part ofthe total bacterial community than the number of culturablebacteria.

ACKNOWLEDGMENTS

This work was supported by grants from the Swedish NaturalScience Research Council and the Swedish Environmental Protec-tion Board to E.B. and from the Federation of European Microbi-ological Societies (FEMS) to H.F.

REFERENCES1. Adams, S. N., J. E. Cooper, D. A. Dickson, E. L. Dickson, and

D. A. Seaby. 1978. Some effects of lime and fertiliser on a Sitkaspruce plantation. Forestry 51:57-65.

2. Baath, E. Thymidine incorporation into macromolecules ofbacteria extracted from soil by homogenization-centrifugation.Soil Biol. Biochem., in press.

3. Baath, E. Measurement of heavy metal tolerance of soil bacteriausing thymidine incorporation into bacteria extracted afterhomogenization-centrifugation. Soil Biol. Biochem., in press.

4. Biath, E., B. Berg, U. Lohm, B. Lundgren, H. Lundkvist, T.Rosswall, B. S6derstr6m, and A. Wir6n. 1980. Effects of exper-imental acidification and liming on soil organisms and decom-position in a Scots pine forest. Pedobiologia 20:85-100.

5. Bakken, L. R., and R. A. Olsen. 1987. The relationship betweencell size and viability of soil bacteria. Microb. Ecol. 13:103-114.

6. Bligh, E. G., and W. J. Dyer. 1959. A rapid method of total lipidextraction and purification. Can. J. Biochem. Physiol. 37:911-917.

7. Dowling, N. J. E., F. Widdel, and D. C. White. 1986. Phospho-lipid ester-linked fatty acid biomarkers of acetate-oxidizingsulphate-reducers and other sulphide-forming bacteria. J. Gen.Microbiol. 132:1815-1825.

8. Federle, T. W. 1986. Microbial distribution in soil-new tech-niques, p. 493-498. In F. Megusar and M. Gantar (ed.), Per-spectives in microbial ecology. Slovene Society for Microbiol-ogy, Ljubljana, Yugoslavia.

9. Fritze, H. 1991. Forest soil microbial response to emissionsfrom an iron and steel smelter. Soil Biol. Biochem. 23:151-155.

10. Fritze, H., and E. Baith. Microfungal commumty structure andfungal biomass in a coniferous forest soil polluted by alkalinedeposition. Submitted for publication.

11. Frostegard, A., E. Baith, and A. Tunlid. Shifts in the structureof soil microbial communities in limed forests as revealed byphospholipid fatty acid analysis. Microb. Ecol., in press.

12. Frostegard, A., A. Tunlid, and E. Baath. 1991. Microbial bio-mass measured as total lipid phosphate in soils of differentorganic content. J. Microbiol. Methods 14:151-163.

13. Guckert, J. B., M. A. Hood, and D. C. White. 1986. Phospho-lipid ester-linked fatty acid profile changes during nutrientdeprivation of Vibrio cholerae: increases in the trans/cis ratioand proportions of cyclopropyl fatty acids. Appl. Environ.Microbiol. 52:794-801.

14. Guckert, J. B., D. B. Ringelberg, D. C. White, R. S. Hanson, andB. J. Bratina. 1991. Membrane fatty acids as phenotypic mark-ers in the polyphasic taxonomy of methylotrophs within theProteobacteria. J. Gen. Microbiol. 137:2631-2641.

15. ]Ulmer, P., and F. Schinner. 1991. Effects of lime and nutrientsalts on the microbiological activities of forest soils. Biol. Fertil.Soils 11:261-266.

16. Ivarson, K. C. 1977. Changes in decomposition rate, microbialpopulation and carbohydrate content of an acid peat bog afterliming and reclamation. Can. J. Soil Sci. 57:129-137.

17. Kroppenstedt, R. M. 1985. Fatty acids and menquinone analysisof actinomycetes and related organisms, p. 173-199. In M.Goodfellow and D. E. Minnikin (ed.), Chemical methods inbacterial systematics. Academic Press, London.

18. Kvalheim, 0. M., and T. V. Karstang. 1987. A general-purposeprogram for multivariate data analysis. Chemometrics Intelli-gent Lab. Syst. 2:235-237.

19. Lang, E., and F. Beese. 1985. Die Reaktion der mikrobiellenBodenpopulation auf Kalkungsmassnahmen. Allg. Forstztg. 43:1166-1169.

20. Mai, H., and H. J. Fiedler. 1979. Bodenmikrobiologische Un-tersuchungen an einem Fichtendungungsversuch im Rauch-schadgebiet des Erzgebirges. Zentralbl. Bakteriol. Abt. I 134:651-659.

21. Martens, H., and T. Naes. 1989. Multivariate calibration. JohnWiley & Sons, Inc., New York.

22. Nodar, R., M. J. Acea, and T. Carballas. 1992. Microbiologicalresponse to Ca(OH)2 treatments in a forest soil. FEMS Microb.Ecol. 86:213-219.

23. O'Leary, W. M., and S. G. Wilkinson. 1988. Gram-positivebacteria, p. 117-185. In C. Ratledge and S. G. Wilkinson (ed.),Microbial lipids, vol. 1. Academic Press, London.

24. Olsen, R. A., and L. R. Bakken. 1987. Viability of soil bacteria:optimization of plate-counting technique and comparison be-tween total counts and plate counts within different size groups.Microb. Ecol. 13:59-74.

25. Persson, T., H. Lundkvist, A. Wir6n, R. Hyvonen, and B.Wessen. 1989. Effects of acidification and liming on carbon andnitrogen mineralization and soil organisms in mor humus. WaterAir Soil Pollut. 45:77-96.

26. Persson, T., A. Wiren, and S. Andersson. 1990-91. Effects ofliming on carbon and nitrogen mineralization in coniferousforests. Water Air Soil Pollut. 54:351-364.

27. Pollard, P. C., and D. J. W. Moriarty. 1984. Validity of thetritiated thymidine method for estimating bacterial growth rates:measurement of isotope dilution during DNA synthesis. Appl.Environ. Microbiol. 48:1076-1083.

28. Tualid, A., H. A. J. Hoitink, C. Low, and D. C. White. 1989.Characterization of bacteria that suppress Rhizoctonia damp-ing-off in bark compost media by analysis of fatty acid biomar-kers. Appl. Environ. Microbiol. 55:1368-1374.

29. Tunlid, A., and D. White. 1992. Biochemical analysis of bio-mass, community structure, nutritional status, and metabolicactivity of microbial communities in soil, p. 229-262. In G.Stotzky and J.-M. Bollag (ed.), Soil biochemistry, vol. 7. MarcelDekker, Inc., New York.

30. Wdlkinson, S. G. 1988. Gram-negative bacteria, p. 299-457. InC. Ratledge and S. G. Wilkinson (ed.), Microbial lipids, vol. 1.Academic Press, London.

31. Zelles, L., I. Scheunert, and K. Krentzer. 1987. Bioactivity inlimed soil of a spruce forest. Biol. Fertil. Soils 3:211-216.

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