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Plant and Soil 230: 145–160, 2001.
© 2001 Kluwer Academic Publishers. Printed in the Netherlands.
145
Seasonal fluctuations in the population of denitrifying and N2 -fixing
bacteria in an acid soil of a Norway spruce forest
Alexander Mergel, Karin Kloos & Hermann Bothe1
Botanisches Institut, Universität zu Köln, Gyrhofstr. 15, D-50923 Köln, Germany. 1 Corresponding author∗
Received 3 April 2000. Accepted in revised form 16 November 2000
Key words: denitrification, nitrogen fixation, seasonal fluctuations of soil bacteria, soil bacterial community, soil
DNA isolation
Abstract
The seasonal fluctuations in the concentration of cultured denitrifying and N2 -fixing bacteria were followed in an
ammonium fertilised and a control soil of a Norway spruce forest near Villingen/Black Forest from December
1994 to August 1998. The horizontal distribution of bacteria in three layers was determined by the MPN-method
and by molecular probing (colony hybridisation) using specific 0.4–0.7 kb DNA probes for denitrification steps
(narG, nirS, nirK and nosZ) and for N2 -fixation (nifH). The data showed that highest bacterial counts and higher
numbers of denitrifying and N2 -fixing bacteria were generally detectable in the upper (= 5 cm) soil layer and that
their amount decreased with soil depth. The concentration of these cultured bacteria showed seasonal fluctuations
with highest numbers in autumn/winter/early spring and with low counts in summer. Denitrifying and N2 -fixing
bacteria amounted to less than 10% of the total number of cultured bacteria determined by the MPN-method.
Fertilisation with ammonium did not cause a shift in the population of these bacteria. These findings were corroborated by hybridisation experiments with genomic DNA isolated from the different layers. Strongest DNA–DNA
hybridisation band intensities were obtained in the upper soil layer and their intensities decreased with soil depth.
Soil samples from Villingen assayed in the laboratory produced N2 O (in dependence of nitrate and C2 H2 added
to the vessels) and utilised this gas with higher activities in the assays with the fertilised soil. It is concluded that
molecular techniques can successfully be applied for assessing seasonal fluctuations of bacterial populations in
soil. Relative abundance of denitrifying and N2 -fixing bacteria can be determined from experiments with DNA
isolated from soils. Attempts to transform these results to the total population of soil bacteria on a single cell basis
are faced with many uncertainties.
Introduction
The bacterial populations in soils are said to be complex (Kruske et al., 1997). It was stated that soils
contain 103-104 different genomes/g soil dry weight
(Borneman et al., 1996; Sandaa et al., 1999; Torsvik
et al., 1990) and only a very limited number of these
can be cultivated (Amann et al., 1995; Felske et al.,
1999). Investigations are further complicated by the
fact that the bacterial populations are seemingly variable within short soil distances and may undergo seasonal fluctuations. These aspects, however, have not
∗ Fax No: 221-470-5181. Tel No: 221-470-2760. E-mail:
[email protected]
yet been thoroughly investigated. Information about
seasonal variations of bacterial populations in other
habitats, e.g. in aqueous biofilms or sediments (Joergensen, 1989; MacFarlane and Herbert, 1984), are
also sparse. Whereas determinations were formerly
restricted to activity measurements in soil cores both
in situ and under laboratory conditions, molecular
biological techniques based on DNA gene probing
now offer new avenues for assessing the distribution
of bacteria with special traits like denitrification and
N2 -fixation in soils.
In denitrification, nitrate is reduced via nitrite,
nitric oxide and nitrous oxide finally to molecular nitrogen. The ability to denitrify is distributed
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in phylogenetically diverse groups of unrelated bacteria (Knowles, 1982; Zumft, 1997). Any attempt
to quantify denitrifying bacteria has to take into account that the reduction of nitrite to nitric oxide is
catalysed by two totally different enzymes, either the
cytochrome cd1 or the Cu-containing nitrite reductase.
Denitrifying bacteria contain either one or the other
enzyme (Coyne et al., 1989). In the present study,
gene probes for all steps of denitrification, with the
exception of NO-reductase, have been employed for
monitoring the distribution of denitrifying bacteria in
a selected soil site. Dinitrogen fixation is catalysed by
nitrogenase which is encoded by the structural genes
nifHDK. In spite of phylogenetic differences, nifH
contains highly conserved DNA sequences among the
N2 -fixing micro-organisms and has, therefore, been
employed for developing probes for assessing the occurrence of nitrogenase in bacteria by molecular probing (Linne von Berg and Bothe, 1992; Widmer et al.,
1999).
In the present study, the seasonal fluctuations of the
denitrifying and N2 -fixing bacterial population were
monitored in the period from December 1994 to August 1998 in an acid Norway spruce forest in the
Black Forest/Southern Germany. The location was selected because of low industrial activity in the area,
therefore any N-input to the soil from air pollution
is low. Two different sites in this forest were chosen
for the present investigation. One plot had been fertilised with 150 kg of ammonium sulfate ha−1 in May
1994, and the other served as the non-treated control. The study aimed at determining the seasonal
fluctuations by different microbiological methods. The
initial studies were restricted to DNA hybridisation
studies with cultured bacteria and to activity measurements with soil samples. Recently, we obtained soil
DNA amenable to hybridisation with DNA-probes for
denitrification and N2 -fixation. Data obtained by these
different approaches will be presented.
Materials and methods
area far remote from significant industrial activity.
Two plots were selected for the investigations. One
(at Glasergrenze, see topographic map no. 7916 of
the Landesvermessungsamt Baden-Württemberg from
1994) had been fertilised with the fairly high load of
150 kg ha−1 ammonium sulfate in May 1994, whereas
the other one (at Schlößlebühl, same map) served as
the non-fertilised control. The surface of this fairly
acid soil (pH ∼ 4.0) was covered with bilberry =
whortleberry (Vaccinium myrtillus L.), cowberry (V.
vitis-idea L.), diverse ferns, such as male fern (Dryopteris felix-mas (L.) Schott), mosses and lichens. The
tree canopy contained predominantly Picea excelsior,
some Scots pine (Pinus sylvestris L.) and, remarkably,
considerable amounts silver firs (Abies alba Miller) of
healthy appearance. This latter species is easily affected and indicative of air pollution at other stands.
Other details of the Villingen spruce forest and its soil
are given in Feger and Raspe (1992) and Stoermer et
al. (1997).
Soil sampling from the Norway spruce stand of
Villingen
Samples were taken from the following three layers:
(a) 5 cm depth, thus from the upper part of the Ahorizon, just below the O-layer, consisting of
accumulated decomposed organic matter, colour
black.
(b) 10 cm depths, from the middle of the A-horizon,
brown-blackish decomposed organic matter
(c) 25 cm, from the B-horizon, with an accumulation of brown-yellowish silicate clay, containing
also roots, twigs and smaller stones which were
discarded.
All soil samples from Villingen were transported in
plastic bags, kept at ∼ 10 ◦ C, to the Cologne laboratory and used the following day for activity measurements, DNA extraction or bacterial colony growth.
Sampling dates were December 1994, March, May,
July and September 1995, October 1996, April 1997,
April 1998 and August 1998 (see Figures 1 and 2).
Site investigated
Gene probes used
The soil of a dystric Cambisol (Stoermer et al., 1997)
in a 100 year-old Norway spruce stand near Villingen, Black Forest (altitude about 780 m) was analysed
for its bacterial community. The N-reserves of this
soil have been estimated at 8.000 kg N ha−1 (Feger
and Raspe, 1992) and are thus not low. The atmospheric N–input is regarded as being small in this
The 0.4–0.7 kb DNA probes for denitrification, N2 fixation and for 16S-rRNA were essentially the same
as in preceding publications (Fesefeldt et al., 1998;
Kloos et al., 1995, 1998). They were generated by
PCR using oligonucleotide primers of regions of the
genes and the DNA of the target bacteria, followed
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Figure 1. The seasonal fluctuation in the cell numbers and in water content in the two plots of the Villingen forest. Samples were taken from
the 5, 10 and 25 cm layers of the nonfertilised (A) and the ammonium fertilised soil (B), diluted and determined by the MPN-method after
growth in LB. The sampling dates are given in the Figure. The bars indicate the counts for the layers: black = 5 cm, grey = 10 cm and white,
with small points = 25 cm. The water content of the soil samples (C) was determined as the difference between wet and dry weight of the soil
samples. Straight line: nonfertilised soil. Dashed line: fertilised soil. Samples taken from 5 cm layer, N 10 cm layer, 25 cm layer.
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Figure 2a–b. Hybridisation of the DNA from cultured bacteria with the gene probes for denitrification and for nitrogen fixation (dot blot
analysis). Bacteria were isolated from the nonfertilised and fertilised soil of the Villingen spruce forest at the different sampling dates and
grown on plates with either LB or YEM (both for 1 d) and heterotrophic mineral medium (5 d). Colonies were transferred to the filters which
were used for the hybridisation with the gene probes. The percentage of positive isolates (giving a distinct signal with the probe) was referred
to the absolute number of isolates determined by the MPN- method. Hybridisation with the following gene probes, in each case coding for part
of the apoprotein of: (a) narG: apoprotein of dissimilatory nitrate reductase. (b) nirS: cytochrome cd1 containing nitrite reductase. (c) nirK:
Cu-containing nitrite reductase. (d) nosZ: nitrous oxide reductase. (e) nifH: nitrogenase reductase. Bars represent: Samples from the upper 5 cm
layer (in black), from the 10 cm depth (in darker grey) and for the 25 cm depth (in light grey).
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Figure 2. c–d
by cloning and sequencing of the amplificates. Their
labeling with digoxigenin had been described earlier
(Kloos et al., 1995). Previous studies (Kloos et al.,
1995, 1998) had shown that these probes recognized
the target DNA isolated from a wide range of reference
organisms of all groups of proteobacteria plus some
Gram positive and also several cyanobacteria.
Probes for denitrification
Dissimilatory nitrate reductase (narG)
Template DNA from E. coli, size of the amplificate:
414 bp, homology to the published sequence (McPherson et al., 1984): 100%, both on the DNA and amino
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Figure 2. e
acid level, position of the probe within the DNA
sequence: N 28–N 441.
Cytochrome cd1 containing nitrite reductase (nirS)
DNA from Pseudomonas aeruginosa, size of the
amplificate: 596 bp, homology to the published sequence from Pseudomonas aeruginosa (Silvestrini et
al., 1989): 100%, on the amino acid level, position of
the probe within the DNA sequence: N 199 to N 795.
Cu-containing nitrite reductase (nirK)
DNA from Alcaligenes xylosoxidans NCIMIB 11015,
size of the amplificate: 576 bp, homology to the published sequence from Alcaligenes faecalis (Nishiyama
et al., 1993): 67%, on the amino acid level, position of
the probe within the DNA sequence: N 526–N 1101.
N2 O-reductase (nosZ)
DNA from Pseudomonas stutzeri ZoBell, size of the
amplificate: 598 bp, homology to the published sequence from the same bacterium (Viebrock and Zumft,
1988): 88%, on the DNA and 91% on the amino acid
level, position of the probe within the DNA sequence:
N 630–N 1227.
Probe for N2 -fixation
Nitrogenase reductase (nifH)
DNA from Azospirillum brasilense Sp7, size of the
amplificate: 435 bp, homology to the published sequence from the same bacterium (De Zamaroczy et
al., 1989): 100%, both on the DNA an amino acid
level, position of the probe within the DNA sequence:
N 19–N 453.
A general probe for recognising bacterial DNA
16S-rRNA
DNA from Azospirillum brasilense Sp7, size of the
amplificate: 693 bp, homology to the published sequence from E. coli (Brosius et al., 1981): 85%, on
the DNA level, position of the probe within the DNA
sequence: N 8–N 701.
Isolation of bacteria and colony hybridisations
The method is similar as described previously (Kloos
et al., 1998). Ten g of soil were suspended in 10 ml
H2 O and stirred for 1 h, and larger particles were removed with a sieve of 0.5 µm pore size. The filtrate
was diluted (102–104 fold) and streaked out onto agar
plates containing either LB, YEM or heterotrophic
mineral medium (for the exact composition see Kloos
151
et al., 1998). All media contained 250 µg ml−1 cycloheximide to suppress fungal growth. Plates were
incubated at 30 ◦ C for 1 d (in the case of LB and YEM)
or 5 d (for heterotrophic mineral medium). Colonies grown were transferred to sterile microtiter plates
which contained 250 µl liquids of either LB, YEM
or heterotrophic mineral medium. Samples were analysed from each soil layer and medium. On the average, 250–288 randomly selected colonies for each trial
were grown up to an optical density > 0.1 (620 nm)
and subsequently analysed by colony hybridisation
(Grunstein and Hogness, 1975) using the above mentioned gene probes. Bacteria in 50 µl aliquots were
transferred to nylon+ -membranes (Qiagen, Hilden)
under vacuum blotting. Filters were treated with denaturating solution, neutralising solution, and then
with UV-light for 2 min. The treatment with proteinase
K and PMSF was also exactly the same as described
(Kloos et al., 1998). Hybridisation was done overnight
at 68 ◦ C with 200–300 ng labelled probe/100 cm2
membrane in the mixture 5 × SSC, 0.5% blocking reagent, 0.1% Na-lauroylsarcosine, 0.02% SDS. Filters
were rinsed twice with 2 × SSC, 0.1% SDS (10 min,
68 ◦ C), and signals were detected immunologically
following the standard protocol of Roche-Boehringer,
Mannheim.
Extraction of DNA from the soil samples and
quantifications
Acid soils in particular are contaminated by humic
acids, tannins, carbohydrates, free DNA and other
interfering substances. The following protocol was
developed for the isolation of DNA which was essentially freed from such compounds and which was
amenable to digestion by the restriction enzymes
(BamHI, SmaI, SacI and EcoRI) and to hybridisation with the gene probes for denitrification and N2 fixation.
Soil samples (5 g) were stirred into 10 ml 0.1%
Na4 P2 O7 /10 mm Tris–HCl/ 1 mM EDTA (=TE buffer). After an incubation (10 min, RT) and centrifugation (10 min, 6000 × g), the pellet was washed with 10
ml TE buffer and centrifuged (10 min, 6000 × g). The
pellet was suspended in 5 ml TE/25 mg lysozyme and
incubated (1 h, 37 ◦ C, under shaking), followed by the
addition of 1 ml of 10% SDS (= sodium dodecyl sulphate), incubation (1 h, 65 ◦ C, under shaking) and was
centrifuged (10 min, 6000 × g). Supernatants were
then subjected to phenol extraction (using equal vol
of phenol) and centrifuged (15 min, 12 000 × g). The
aqueous phase was removed and cleaned from residual phenol by treatment with 5 ml chloroform/isoamyl
alcohol (24/1,v/v). The DNA from the upper phase
(∼ 5 ml) was then precipitated with potassium acetate (300 µl) and isopropanol (3 ml) for 16 h at 4 ◦ C.
The sample was centrifuged (10 min, 10 000 × g),
washed with 300 µl 70% ethanol, centrifuged once
more, dried in a vacuum centrifuge and suspended
with 1 ml 1.2 M NaCl. After storage for 2 h at RT, the
sample was supplemented with 1/10 vol 10% CTAB
(= hexadecyl-trimethylammonium bromide) dissolved
in 1.2 M NaCl, stored (10 min, 65 ◦ C), supplemented
with chloroform (1 ml) and spun down in an Eppendorf centrifuge at maximal speed. After repeating this
washing step, the DNA in the aqueous phase was
precipitated with 50 µl 3 m K+ -acetate/0.6 ml isopropanol, stored (16 h, 4 ◦ C), centrifuged (20 min, 18 000
× g), washed once more with 70% ethanol, dried in
the SpeedVac SC 100 (Savant) vacuum centrifuge and
suspended in 20 µl TE buffer. The DNA obtained
was then electrophoresed on 0.6% PeqGold Low
MeltAgaraose (from Peqlab, D-Erlangen) at 40–50 V
for 1–2 h. The DNA bands which were discernible by
UV-light were cut out and transferred to 1.5 ml Eppendorf tubes, supplemented with 3 vol of TE, incubated
at 65 ◦ C to melt the agarose (5–10 min) and extracted
with choroform/ isoamylalcohol (24/1). The DNA was
precipitated with 1/10 vol of 3 m K+ -acetate/0.6 ml
isopropanol (RT, 1 h), centrifuged (15 min, 18 000 ×
g), washed with 70% ethanol, dried in the SpeedVac
and finally suspended in 20 µl TE buffer. This somewhat laborious preparation gave DNA of high purity
(O.D. ratio 260 nm /280 nm ∼ 1.7). Seeding experiments with Azospirillum brasilense Sp7, Alcaligenes
eutrophus H16 or E. coli K12 gave recoveries between
47 and 69% (not documented). Signal intensities of
the genomic DNA isolated from the soil samples or of
the DNA–DNA hybridisation bands were determined
densitometrically using the NIH Image 1.61 picture
analyser program for MacIntosh.
Activity measurements in the soil cores
Samples from the Villingen forest were assayed in
aqueous extract for pH using a PHM 61 laboratory
pHmeter electrode (Radiometer, Copenhavn) and for
nitrate content by the salicylic acid method (Cataldo
et al., 1975). The soil moisture was calculated from
the dry weight of the soil. MPN-determinations after
diluting the soil samples 103 – 107 fold and incubating
in LB for 4 weeks were performed as described by Al-
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exander (1982). Formation or utilization of gases were
assayed with 2 g soil samples in ∼ 7.0 ml Fernbach
flasks covered with gas-tight suba seals. For N2 Oformations, the flasks were supplemented with 2 ml
1 mM NaNO3 , evacuated and refilled with argon, and
0.5 ml C2 H2 was added by a syringe. For determining
N2 O-uptake rates, 1 ml of this gas was injected into
the anaerobic Fernbach flasks. Incubation was generally performed in a shaking water bath (30 ◦ C, 1 d).
Gases (N2 O, CO2 and O2 ) were determined by standard gas chromatography as described earlier (Kloos et
al., 1995).
Results
The seasonal fluctuations in the total number of
cultured bacteria and in the percentages of
denitrifying and N2 -fixing isolates
Any investigation with soil samples is faced with the
problem that soils can show large fluctuations in their
chemical and physical composition within short distances. In the case of the Villingen soil, horizons had
a homogenous appearance judging from looking into
different holes dug within a few m of one another on
the two plots. In the fertilised soil, the upper 10–15 cm
in depth contained black soil of decomposed plant litter material, and then from this depth till 80 cm the
soil was homogeneously brown-yellowish containing
a few small stones. The nonfertilised soil had similar eye-visible appearance, but the black cover spread
from the top to approximately 20 cm, probably due to
the fact that this stand was more extensively covered
with small woody plants like Vaccinium myrtillus, V.
vitis-idaea and Calluna vulgaris. This black upper
cover was more thoroughly interspersed with plant
roots than in the fertilised plot.
In October 1996, samples from five places separated from each other by few meters were collected
from the upper soil layer (5 cm depth) of both the
N-fertilised and the nonfertilised soil, and five independent determinations of nitrate content and cfus
after growth on LB for 1d were performed for each of
the two plots. The nitrate content was 4.4±1.8 µmol
g−1 dry weight of soil for the fertilised plot and
2.4±1.8 for the control. Likewise, the data for the
colony forming units (cfu) after growth on LB for 1
d were variable: 6.7±6.1 × 10 5 cfu g−1 dry weight
of soil in the fertilised plot and 6.2 ± 5.1 × 10 5
cfu g−1 dry weight (n = 5, always) in the case of the
control. Variations in these counts by a factor of maximally 10 had to be encountered when cfus grown on
LB were determined from soil samples taken within
short distances. When the cells had been grown on
plates with yeast extract/mannitol (YEM) for 1 d or
on heterotrophic mineral medium supplemented with
malate, variations in cfu counts were only two- to
threefold. The pH value of the samples taken from the
upper 5 cm of the two plots varied between 3.8 and 4.1
(Table 1).
The seasonal fluctuations of cultured bacteria (determined by the MPN-method after growth in LB for
4 weeks) and of cultured denitrifying and N2 -fixing
microorganisms hybridising with the specific gene
probes was followed during the period from December 1994 till August 1998. Due to the large amount
of data compiled from the different samples (two
plots, of each three horizons, growth of the bacteria
in three different media on the plates in most cases,
then hybridisation with 5 gene probes), only duplicate
determinations for each trial were feasible.
The total cell number showed fluctuations with
peak values in autumn – winter – early spring and with
low values in late spring – summer (Figure 1a, b). In
April 1997, scores were fairly high, whereas they were
low in April 1998. The forest soil was unusually dry
in all three soil layers (5, 10, 25 cm) in April 98 (Figure 1c) which could account for the low MPN at that
date. The soil at other dates was, however, not drier
in summer than in autumn–winter–spring. Thus, there
was no obvious correlation between water content and
bacterial cell counts.
The maximal MPN-values did not exceed 3 × 107
cells g−1 dry weight of soil even in the upper (5 cm)
zone. Highest counts were often, particularly in the
case of the fertilised plot, in the upper layer and lowest
counts were at 25 cm (Figure 1a, b). Hybridisations
with the different gene probes for denitrification (nirS,
nirK, nosZ and narG, experiments with narG could
be performed only from March 1995 on) and for nifH
gave a similar pattern in the seasonal fluctuations of
the counts in the soil layers (Figure 2a–e) as obtained
in the case of the MPN (Figure 1). Scores were generally higher in autumn-winter-early spring and lower
in summer. Highest positive hybridisations were often
obtained with the isolates from the 5 cm or the upper
two (5 cm and 10 cm) layers. Sometimes, however, apparently after prolonged rainfalls (e.g. in March 1995),
high scores were also measured with the isolates from
the 25 cm zone. It should be noted that the nitrate
content decreased with the depth of the soil and was
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Table 1. Parameters of soil samples taken from the two plots of the Villingen forest
Soil
Soil depths [cm]
Non-fertilised
5
10
pH [H2 O]
Water content [%]
Nitrate content
[µmol g−1 dry weight × h−1 ]
N2 O-formation
[nmol g−1 dry weight × h−1 ]
N2 O-utilization
[nmol g−1 dry weight × h−1 ]
CO2 -formation in air
[µmol g−1 dry weight × h−1 ]
O2 -uptake in air
[µmol g−1 dry weight × h−1 ]
CO2 -form., under Ar
[µmol g−1 dry weight × h−1 ]
Cell number [MPN,
105 g−1 dry weight]
cfu × 105 g−1 dry weight]
grown on LB
cfu × 105 g−1 dry weight]
grown on YEM
cfu × 105 g−1 dry weight]
grown on heter. MIN
Total cfu × 105 g−1 dry weight
grown on the three media
3.8
72.7
2.4
3.8
23.4
0.4
4.1
16.6
0.0
4.1
64.9
4.4
4.2
21.7
0.1
4.5
19.3
0.0
3.2
0.5
0.6
15.9
2.2
0.4
167.1
150.3
108.9
447.9
128.7
162.5
1.1
0.25
0.1
0.4
0.15
0.05
1.3
0.4
0.15
1.1
0.25
0.05
0.3
0.1
0.05
0.25
0.1
0.05
48.1
5.1
1.8
41.3
9.1
6.5
6.2
0.2
0.1
6.7
0.7
0.03
8.9
1.2
1.7
5.2
3.7
2.9
6.1
0.7
0.8
4.4
2.0
1.4
21.2
2.1
2.6
16.3
6.4
4.4
25
Fertilised
5
10
25
Non-standard abbreviations: wt = weight, cfu = colony forming units, MPN = most probable number, LB = Luria-Bertani, YEM = yeast extract-mannitol, heterMIN = heterotrophic
mineral medium. The determinations of the parameters are described under ‘Materials and
methods’. The formation of N2 O (in the presence of 1 mM nitrate) and the utilisation of this
gas was performed with the soil samples under laboratory conditions at 20 ◦ C. The total cfu
data represent the sum of the colonies grown on all three media, thus including those isolates
which grow on more than one medium. Data are from October 1996.
low (< 0.1 µmol/g dry weight) at 25 cm (Table 1).
The total number of isolates hybridising with the gene
probes for denitrification and for nifH were low, generally amounting to only 2–5% and rarely exceeding
10 % of the total amounts of cultured bacteria (Figure 2 a–e). Isolates hybridizing with all three probes
for denitrification: narG, nirS or nirK, and nosZ were
under 1% (not documented). DNA of none of the
isolates hybridised with both nirS and nirK (not documented). Scores were considerably higher with nirK
than with nirS (Figure 2b, 2c).
Experiments performed with genomic DNA isolated
from the Villingen soil
Genomic DNA was extracted from the three soil layers
of the two Villingen plots in August 1998. The DNA
preparation obtained had molecular weights between
15 and 25 kb with no smear above and below these
sizes indicating that the portion of sheared DNA was
negligible (Porteous et al., 1994). The yields were
calculated from a standard curve with E. coli DNA
and gave 10.4/10.5 µg DNA g−1 dry weight of soil
for the upper 5 cm zone of the nonfertilised/fertilised
Villingen soil, respectively. The corresponding values for the 10 cm and 25 layers were 5.0/3.7 and
1.4/1.6 µg DNA g−1 dry weight of soil, respectively
(see also Figure 3). The DNA preparation obtained
was suitable for hybridisation with the DNA-probe for
the 16S-rRNA (a general probe recognising only bacterial DNA) and with the other segments representing
parts of the denitrification and nifH genes. DNA–DNA
hybridisation signal intensities can only be compared
154
filterwise because hybridisation conditions cannot be
standardised to such an extent as to get reproducible
signals from one filter to the next. Therefore, the DNA
on the filter was first hybridised with the physiological
gene probe, then the filter was stripped and hybridisation was subsequently performed with the 16S-rRNA
probe (Figure 3). As documented for the hybridisation
with nirK, nirS and nifH (Figure 3), signal intensities
were highest in the upper, 5 cm zone.
For quantitative data, the signals were scanned and
the maximal values, always obtained in the hybridisations with the DNA from the 5 cm layer, were
arbitrarily set to 100% (Table 2). The data indicate that
the signal intensities decreased parallel with the soil
depth in the case of all five physiological gene probes
available and also in parallel with the signal intensity
obtained with the 16S-rRNA probe. Thus, there was
no selective enrichment of denitrifying and N2 -fixing
bacteria in lower soil layer (= 25 cm) of the Villingen
soil.
At the same sampling date (August 1998), the
total number of cfu was determined for comparison.
Total cfu after growth on LB and YEM for 1–2 days
were in the range of 104 – 105 g−1 dry weight of
soil (Table 2). The cfu values were always 1–2 orders
of magnitudes lower than the counts determined by
the MPN-methods (Table 1). Bacteria grown on the
LB and YEM containing plates in August 1998 were
then used for colony hybridisations (Table 2). Positive scores with the physiological gene probes were in
the range of 102 – 103 (Table 2) and thus amounted
to grossly 5% of the bacterial counts. When the percentages of isolates hybridising with one of the gene
probes were referred to the total number of cultured
bacteria obtained by the MPN-method, a clearer picture about the distribution of bacteria in horizontal
layers emerged. (Figure 2). However, it is not clear
whether the percentage of bacteria hybridising with
the probes is the same on growth in suspensions (for
the MPN-method) as on plates (for the determination
of the cfu).
Determination of the potential physiological activities
Soil from Villingen was tested for several physiological activities (Table 1). The rates of CO2 -evolution
and of O2 -uptake were approximately the same when
the soil samples were assayed in air. The CO2 formation activity under anaerobic conditions was
about fourfold lower than in air. The samples utilised
N2 O (Table 1), and this activity was totally inhibited
by approximately 10% C2 H2 in the gas phase (data not
shown). The samples also produced N2 O (Table 1),
however, only in strict dependence on the addition
of nitrate and C2 H2 in the case of the Villingen soil.
In the 5 cm soil layer, samples from the fertilised
soil formed and utilised N2 O with significantly higher
activities than those from the non-fertilised soil in
these experiments performed in October 1996.
The temperature dependence of the N2 O-formation
activity was tested with soil from the 5 cm layer at 9 ◦ ,
20 ◦ and 30 ◦ C. Figure 4 shows a typical experiment
where the assay vessels were supplemented with different concentrations of nitrate. Activities were readily
measurable after 24 h in the 20 ◦ and 30 ◦ C assays, and
unambiguous values were obtained after 48 h in the 9
◦ C assay. The reaction, however, proceeded for at least
60 h. The apparent Vmax –values at 30 ◦ C were 29.5 for
samples from the fertilised plot and 14.5 nmol N2 O
formed h−1 × mg−1 dry weight of soil for those from
the control area. Corresponding data were 19.0/8.2 for
20 ◦ C and 0.5/0.3 nmol N2 O formed h−1 × mg−1
dry weight of soil for 9 ◦ C for samples from the
fertilised/ nonfertilised soil, respectively. Biological
reactions generally show an approximately twofold increase when the assay temperature is raised by 10 ◦ C.
This was, indeed, the case for the data for the fertilised
soil obtained at 20 and at 30 ◦ C, whereas the data at 9
◦ C were 5–6 fold lower than at 20 ◦ C. Remarkably, the
samples from the fertilised plot consistently showed
higher activities than those from the control in this specific test. In these N2 O-evolutions, the S[0.5V ] -values
(= apparent Michaelis constants) for nitrate using the
samples from the fertilised plot were roughly 1/2 of
those obtained with the control soil.
Discussion
Both denitrification and N2 -fixation are widely distributed among diverse bacteria of totally unrelated
systematic affinities. Therefore, general DNA probes
based on 16S-rRNA oligonucleotide sequences cannot be developed for recognising bacteria with these
physiological traits (Braker et al., 1998). The 0.4–0.7
kb probes developed in our laboratory specifically for
the different steps of denitrification and for N2 -fixation
recognize these genes in a wide range of bacteria
by heterologous hybridisations (Kloos et al., 1998)
and have successfully been employed for analysing the populations of Hyphomicrobium isolates from
aqueous systems (Fesefeldt et al., 1998; Kloos et al.,
155
Figure 3. Hybridisation of genomic DNA isolated from the three different soil layers with the gene probes. DNA was isolated as described
under ‘Materials and methods’ and blotted onto the nylon+ -filters. Hybridisation on each filter was first performed with the physiological gene
probe (nirK, nirS or nifH). Then the filters were stripped and hybridisations were performed with the general 16S-rRNA probe recognising
bacterial DNA. DNA isolated from 0.5 g soil (fresh weight) was blotted to the filters for the 5 cm and 10 cm samples and from 1.2 g for the
sample from the 25 cm depth. Band intensities were quantified densitometrically for the data of Table 2.
156
Table 2. Hybridisation of the total genomic DNA isolated from the different layers of the two Villingen plots and of
genomic DNA from cultured bacteria with the gene probes for denitrification and nitrogen fixation
Fertilised
Soil depth
Non-fertilised
5 cm
10 cm
25 cm
5 cm
10 cm
25 cm
Total cfu after growth on
LB (number × 104 × g soil)
YEM (number × 104 × g soil)
8.8±3.9
8.2±1.7
9.5±1.3
3.0±0.4
1.5±0.5
4.9±0.2
13.4±2.8
18.5±9.4
6.0±1.2
17.7±5.5
0.8±0.3
6.7±5.9
100
100
70.2±9.9
79.8±7.8
13.5±0.5
16.9±3.6
100
100
39.9±4.6
48.2±7.0
14.9±1.4
17.8±0.2
6.4
35.8
32.3
14.0
3.6
22.8
34.4
23.0
27.8
30.6
1.2
25.9
100
100
33.0±6.9
56.6±2.8
25.3±1.7
16.3±0.5
100
100
29.1±7.1
20.3±1.0
19.4±7.3
11.3±1.2
6.4
37.3
16.7
4.7
11.7
7.0
159.4
55.6
29.1
46.4
1.1
0.8
100
61.2±6.7
25.0±3.7
100
26.2±2.4
17.6±0.5
100
68.8±18.1
17.5±6.4
100
40.9±3.2
19.1±4.7
24.8
0
49.1
11.4
15.8
29.5
183.7
119.5
17.2
0.0
2.3
0.0
100
100
58.9±3.3
47.5±0.2
18.6±9.1
9.2±0.3
100
100
29.3±1.7
29.2±3.7
15.1±4.1
13.8±3.2
6.3
91.3
41.1
14.4
2.0
63.9
16.5
152.7
37.8
0.0
0.0
0.0
100
100
44.0±3.9
40.3±0.1
21.0±4.1
11.0±3.2
100
100
50.6±2.9
46.1±4.7
20.9±2.7
18.6±3.2
12.7
62.3
67.8
60.4
0.0
0.0
0.0
87.1
0.0
0.0
1.8
1.9
Hybridisation of soil DNA with
NifH
16S-rRNA
Hybridising isolates from medium
LB (number × 102 × g soil)
YEM (number × 102 × g soil)
Hybridisation of soil DNA with
NarG
16S-rRNA
Hybridising isolates from medium
LB (number × 102 × g soil)
YEM (number × 102 × g soil)
Hybridisation of soil DNA with
NirS
16S-rRNA
Hybridising isolates from medium
LB (number × 102 × g soil)
YEM (number × 102 × g soil)
Hybridisation of soil DNA with
NirK
16S-rRNA
Hybridising isolates from medium
LB (number × 102 × g soil)
YEM (number × 102 × g soil)
Hybridisation of soil DNA with
NosZ
16S-rRNA
Hybridising isolates from medium
LB (number × 102 × g soil)
YEM (number × 102 × g soil)
Soil samples from the 5, 10 and 25 cm layer was diluted and plated onto agar containing either LB or YEM. After 1
d, the number of colonies grown (cfu) were counted, and 250–288 colonies were used for hybidization with the gene
probes in the dot- blot analyis. The percentage of positive counts was referred to the total number of cfu. For the other
part of the data of this table, genomic DNA was isolated from the 5, 10 and 25 cm layers of the two Villingen plots and
then hybridised with the gene probes for denitrification or nifH. The signal intensities were quantified densitometrically
and DNA–DNA hybridisation bands were then stripped off the filter, and the same filter was subsequently used for the
hybridisations with the DNA probe coding for the bacterial 16S-rRNA. The signal intensities obtained with the DNA
from the 5 cm soil layers were always set to 100%. All data refer in% to this intensity. The soil was sampled in August
1998.
157
Figure 4. N2 O-formation activity with soil samples from the fertilised and nonfertilised soil. Soil samples from Villingen were brought to
the Cologne laboratory and assayed for N2 O-formation in the presence of varying amounts of nitrate and at different temperatures. The
Fernbach flasks contained 10% C2 H2 to block N2 O-reductase activity. The samples assayed were from the upper (= 5 cm) soil layer. Dashed
lines: samples from the fertilised plot. Straight lines: samples from the nonfertilised control. experiment performed at 30 ◦ C N experiment
performed at 20 ◦ C experiment performed at 9 ◦ C.
1995). Hybridisations with these probes also indicated
seasonal fluctuations in the population of cultured denitrifying and N2 -fixing bacteria in both plots of the
Villingen forest, and these seasonal variations grossly
matched with the fluctuations of the total number of
bacteria determined by the MPN-method.
The high variability of the soil constituents within
short distances is a general problem in soil microbiology. The question arises whether this variability is
larger than any difference in the data obtained or which
definitive conclusions can be drawn from such studies. First of all, it can be stated that the total bacterial
counts in such rather acid soil are – not unexpectedly
– considerably lower than in soils with higher pH values. For example, bacterial numbers in the soil of the
Chorbusch forest (a gleysol, pH around 5) or in chalk
meadows in the vicinity of Cologne are 1–2 orders of
magnitude higher than in the Villingen soil (Kloos et
al., 1998; this study).
Secondly, bacterial life and activity are highest in
the upper zone and decrease with the depth of the
soil. This was shown for all soils investigated so far
(Kloos et al., 1998) and is also seen in the Villingen
plots, although not so distinctly as in the other soils,
possibly because of the low bacterial numbers in the
soil profiles of this spruce forest. Data are clear-cut
for the activity measurements (highest gas formations
or utilisations always in the upper layer) and for the
experiments performed with the isolated DNA (band
intensity also highest in upper 5 cm). In the MPN–
determinations and in the colony hybridisations with
the cultured bacteria, the method allowed to conclude
that seasonal fluctuations occurred with peak activities
in autumn/winter and early spring and with low scores
in summer. The refinement of the data was, however,
not sufficient for more generalisations mainly because
of the low number of cultured bacteria and because
it was not feasible to accumulate more data, e.g. by
monthly determinations.
Such fluctuations with peak activities in winter/early
spring during freezing/thawing conditions have also
been reported for N2 O-formations for grassland and
158
other soils (Bremner et al., 1980; Sommerfeld et al.,
1993). In combination with chemical processes, biological reactions, in particular those performed by
denitrifying bacteria, appear to be responsible for this
large fluxes of the gas (Müller et al., 1997). The population of cultured denitrifying bacteria assessed by
molecular probing are likely too small to account for
these rates in the Villingen soil. Denitrifying bacteria
amounted to less than 10% of the cultured bacteria and
did not exceed 106 cells g−1 soil dry weight (Figure 2).
With soil samples assayed, rates (in the presence of
nitrate and C2 H2 ) amounted to maximally 15–20 nmol
N2 O g−1 soil sample × h−1 at 20 ◦ C in the 5 cm layer
(Figure 4). For comparison, a culture of Alcaligenes
eutrophus in the exponential growth phase produced
about 0.2 nmol N2 O h−1 per 10 6 cells at 20 ◦ C, and
the same cell number of Bacillus cereus formed even
less under the same assay conditions (B. Schmitz and
H. Bothe, unpublished data). Since there is a difference of two orders of magnitude, other bacteria or
chemical processes than the cultured, ‘classical’ denitrifying bacteria must have been major contributors
to the 15–20 nmol h−1 detected in the soil samples.
This consideration is in line with the statement that
only 1% or less of the soil bacteria have been cultured
(Amann et al., 1995). Thus a concentration of at least
108 cells g−1 soil dry weight is to be expected in order
to produce the rates of 15–20 nmol N2 O h−1 × g−1
soil sample.
Which are the organisms (factors) responsible for
this high activity in soil samples? Besides noncultured bacteria, fungi and heterotrophic nitrifiers could
be major contributors to the N2 O-production. Some
fungi have been reported to perform denitrification
(Shoun et al., 1992) but the relative contributions of
fungi to the N2 O-release from soils have not yet been
examined. Heterotrophic nitrifiers may also produce
N2 O, although the major gas formed by them appears
to be nitric oxide (Daum et al., 1998). Such strongly
acid soils have been reported to contain heterotrophic
nitrifiers in the extremely and thus surprisingly high
numbers of 108 to 1011 cells/g soil (Papen and Von
Berg, 1998). A gene probe developed for nitrification
(from the amoA gene of Nitrosomonas europaea) recognised the corresponding gene in the heterotrophic
nitrifier Pseudomonas putida (Daum et al., 1998) but
could not be tested for the occurrence of this gene in
a wide range of heterotrophic nitrifiers because only
few of them can be cultured nowadays. Therefore, the
amoA probe (Kloos et al., 1998) was not used in the
present study.
The data of the present study indicated that the
characterization of cultured bacteria provided some information about the seasonal fluctuations in such soils
with low bacterial counts. The major conclusion is that
cultured denitrifying and N2 -fixing bacteria occur in
the upper soil layer in higher numbers than in depth
which poses the question of their advantage in having
retained these genes for life in such a habitat. The
results also showed that cultured bacteria possessing
the Cu-nitrite reductase (nirK) occur more abundantly
at such a location than those with the cytochrome cd1
nitrite reductase (nirS), whereas it was suggested that
bacteria with nirS predominate in nature (Coyne et
al., 1989). Moreover, the results indicated seasonal
fluctuations of the total number of bacteria and the
percentage of denitrifying and N2 -fixing microorganisms. More details could, however, not be gained,
because the method with its high statistic error was too
coarse. In the present study, 250–300 bacterial colonies were characterized for their hybridisation with the
gene probes at each date and plot. For obtaining more
accurate differences, an at least tenfold higher number
of isolates needs to be investigated which is hardly
feasible. Even then the problem would still remain that
less than 1% of the bacteria detectable by fluorescence
methods in soils (Torsvik et al., 1990) would have
been cultured.
Hybridisation with genomic DNA isolated from
the different layers and with the gene probes indicated
more convincingly that denitrifying and N2 -fixing bacteria mainly occur in the upper soil zone, and that their
concentration decreases in parallel with the total number of bacteria assessed by the MPN-method or by the
cfu-determination. If one assumes that the DNA of the
bacteria from the different layers hybridises with the
same concentration dependence, this approach gives
clear-cut results on the relative abundance. A crucial factor in these assays is, however, the isolation
of DNA in high yield and purity. The preparation
obtained in the present study was amenable to digestion by restriction enzymes and to hybridisations with
the gene probes. The method employed was timeconsuming but allowed the recovery of 50–70% of
the DNA as shown in seeding experiments with Acaligenes eutrophus H16, Azospirillum brasilense Sp7 and
Escherichia coli K12 (A. Mergel, unpublished data).
A recent examination of the different protocols for the
extraction of DNA from soils came to the conclusion
that not more than 6% of the DNA of indigenous bacteria can be recovered from different soils and that the
clay content strongly influences the retrieval of DNA
159
(Frostegard et al., 1999). An even more severe bias is
that such relative signal intensities of the hybridisation
band cannot be transformed to cell numbers or referred to activities based on single cells (Amann et al.,
1997). The quantitative determination of individual
genes present in single copy or low copy numbers in
intact bacterial cells of soils by in situ hybridisation
methods is not possible (Hodson et al., 1995), and
quantitative in situ PCR-techniques for characterising
bacterial populations in habitats are still in their infancy. Such methods are, however, required if one
wants to correlate numbers in the bacterial populations
to global fluxes of NOx or other gases.
Well aware of these facts, we nevertheless venture
to forward the following estimations: The genome size
of E. coli is 4.7 × 106 bp or 3.1 × 109 dalton (standard
table from Boehinger-Roche; see also Torsvik et al.,
1990). A calibration curve with DNA isolated from
E. coli indicated that the nonfertilised/fertilised plot
contained 10.5/10.5 µg DNA g−1 dry weight of soil
in the 5 cm layer and 5.0/3.7 in 10 cm and 1.4/1.6 in
25 cm. This is equivalent to a total number of E. coli
genomes of 2.0/2.0 × 109 g−1 soil in 5 cm, 0.9/0.7
× 109 in 10 cm and 0.25/0.3 × 109 in 25 cm. If the
average genome size of the bacteria from the Villingen soil is similar to that of E. coli, the comparison of
these data with the MPN-determinations shows that 1–
5% of the bacteria of that soil have been cultured. The
same can be deduced from the concentration of DNA
in bacteria cells, varying between 2 and 15 fg DNA
(Fuhrman et al., 1988; Paul and Myers, 1982). If the
average value is 8.5 fg, then the nonfertilised/fertilised
plot contains 1.4/1.4 × 109 cells in 5 cm, 0.65/ 0.5 ×
109 in 10 cm and 0.2/0.2 × 109 in 25 cm. All these
estimations, together with the activity measurements
seem to indicate that such soils contain more than
109 cells g−1 dry weight of soils in the upper layer,
of which grossly 107 to 108 appear to be potential
N2 O-producing microorganisms.
One plot had been fertilised with ammonium
sulfate in May 1994, and the soil samples from this
plot investigated in October 1996 showed higher N2 Oformations and utilizations than those from the control
area. Other parameters (CO2 -evolution and O2 -uptake
activities, MPN-counts, hybridisations with cultured
bacteria, hybridisations with DNA isolated from the
different horizons), however, did not show significant differences between the samples from the two
plots. Fertilisation with nitrate may cause a shift in
the population of denitrifying bacteria in other habitats (Nijburg and Laanbroek, 1997). The ammonium
added to the Villingen soil may have been converted
to nitrate by the action of nitrifying bacteria. As there
was no increase in nitrate content in the horizons of
the fertilised Villingen soil and as the increased denitrification activity in the laboratory experiments was
only detected when nitrate was added to samples, this
higher activity may be caused by the activation of
‘dormant’, noncultured denitrifying or nitrifying bacteria which may have selectively been enriched by the
N-fertilisation.
Acknowledgements
The authors are indebted to Regina Hiltawsky, Karin
Kaiser, Dr Ralf Schlichting and Valer Schulte-Fischedick for participating in some of the experiments,
to Kerstin Nawrath and Barbara Schmitz for skilful technical assistance and to Dr Oliver Schmitz for
helpful comments. This work was kindly supported
by grants from the Deutsche Forschungsgemeinschaft
within the priority program "Microbial Ecology" and
from the BMBF within the program Spurenstoffkreisläufe (through the Fraunhofer Institute in GarmischPartenkirchen).
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Section editor: C. van Kessel