FEMS Microbiology Ecology 38 (1986) 257-265
Pubfished by Elsevier
257
FEC 00078
Fungi of a forest soil nitrifying at low pH values
(Heterotrophic nitrification; nitrifying microorganisms; Verticillium lecanii; sterilized soft; oximes)
Elke Lang
*
and Gerhard Jagnow
Institut far Bodenbiologie, Forschungsanstalt fib" Landwirtschaft, Bundesallee 50, D-3300 Braunschweig F.R.G.
Received 6 May 1986
Revision received and accepted 13 June 1986
1. SUMMARY
No autotrophic nitrifying organisms were found
in a podzolic brown earth forming nitrate. 350
fungi and aerobic heterotrophic bacteria were isolated from this soft and examined for their nitrifying abilities. About one quarter of the isolates
produced 0.05-0.90 nag N . 1-1 nitrite or nitrate in
peptone solution, soft extract mixture or sterilised
soil. The nitrification rate of the most active
fungus, Verticillium lecanii, was highest at pH 3.5
in defined media. The results support the significance of heterotrophic nitrification in acid soils.
2. INTRODUCTION
Nitrification in neutral and slightly acid softs
and in water is carried out by chemoautotrophic
bacteria such as Nitrosomonas and Nitrobacter.
These organisms are sensitive to acidity, and
oxidise nitrogen in nutrient solutions only at pH
values higher than 5.0 [1].
Nitrification occurs in many forest softs under
conditions more acid than pH 5.0 [2,3], and the
* To whom correspondence should be addressed.
question arises which organisms are nitrifying in
these softs. Three hypotheses could answer the
question. (1) The pH of microhabitats differs
widely enough from the average pH measured in
suspension to enable the acid-sensitive nitrifiers to
find microsites of a suitable pH. (2) There are
some acid-tolerant autotrophic nitrifiers which
have not yet been identified. (3) Acid-tolerant
heterotrophic organisms are carrying out nitrification.
Some authors have found autotrophic nitrifiers
in acid forest soils [4-6] and some have not [7-9].
Properties of the site other than soil reaction, such
as humus and clay content or litter quality, may
possibly determine which hypothesis is correct for
acid softs. Knowledge of heterotrophic nitrifying
bacteria and fungi has been reviewed [10,11]. Heterotrophic nitrification has also been identified in
a forest soft [12].
In the present study, a podzolic brown earth
(pH 3.5) was chosen to answer the question about
the nitrifying organisms. We investigated whether
autotrophic nitrifiers were living in the soft,
whether there were heterotrophic bacteria or fungi
present which were capable of nitrification, and
whether the isolated nitrifying organisms could
tolerate acidity and could be active at the low pH
of this soil.
0168-6496/86/$03.50 © 1986 Federation of European Microbiological Societies
258
3. MATERIALS A N D M E T H O D S
3.1. Soil
The podzolic brown earth from loess over triassic sandstone is described in Table 1. The humus
of the beech site in the Soiling mountains (International Biological Program) was a typical moder.
3.2. Sampling
20 cylindrical soil cores (10 cm in diameter and
20 cm in depth) were collected from the beech site.
The organic layers O h plus Of and the upper 5 cm
of the mineral soil of the cores were combined
separately and mixed carefully by hand. The samples were cooled during transport and stored at
3°C.
3. 3. Counting of autotrophic nitrifying organisms
Bacterial counts were carried out by the mostprobable-number method [14]. Within 48 h of
collection, 100 g of the moist soil was weighed
into 400 ml 0.18% sodium hexametaphosphate
(pH 7.2) and shaken for 1 h at 3°C at 35 rev./min
Dilution series were prepared in Ringer's solution
( 1 / 4 concentrated) and 0.1 ml aliquots were inoculated into tubes with 2 ml nutrient solution (pH
7.7) [15]. 5 tubes per dilution were inoculated. For
counting ammonia and nitrite oxidisers, the solution contained 0.472 g (NH4)2HPO4 or 0.7 g
N a N O 2 per 1, respectively. The tubes were incubated at 22°C for 6 weeks. The presence of
Table 1
Some properties of the podzolic brown earth a
Organic layer Mineral soil
Oh+Of
0-5cm
3.4
3.5
20.6:1
20.3:1
24
3.9
1.2
0.2
pH (water
C:N
Total organic (%)
Total nitrogen (%)
Water-holding capacity
(g water-100 g dry weight- l)
230
Net mineralisationof N
(rag N-kg- t. 28 days- l at 22° C) 75
Nitrate production
(% of net mineralised N)
51
a Unpublished data and [13].
72
6.5
78
nitrite was checked by mixing 100 /~1 of the culture with a drop of Griess-Ilosvay reagent ('spot
test', [16]). The nitrate was reduced with a pinch
of zinc powder.
3. 4. Counting oxime-N oxidizing organisms
The method described in section 3.3. was repeated, but using nutrient solution containing
0.625 g pyruvic oxime • 1-1 instead of ammonium
phosphate. Pyruvic oxime was prepared according
to [171.
3.5. Enrichment cultures of autotrophic nitrifiers
2 g of the humus layer was suspended in 20 ml
nutrient solution [15] with ( N H 4 ) 2 H P O 4 or
N a N O 2 at both p H 7.0 and p H 4.5. This technique was repeated substituting the humus by 2 g
of mineral soil or by 1 ml of a 1 : 10 dilution of
soil in sodium hexametaphosphate. The cultures
were incubated for 8 weeks at 22°C and then
tested for nitrite and nitrate content.
3.6. Isolation and identification of heterotrophic soil
fungi and bacteria
Fungi were isolated by a method described in
[18]. Mineral soil was put in a small 20-mesh sieve
and washed with 10 1 water. The residue particles
were washed with streptomycin (150 mg. 1-1) and
put onto agar plates containing carboxymethyl
cellulose [19], 4 particles per Petri dish.
Single strains were isolated from the mycelia
growing out of the crumbs and were transferred
into tubes with agar containing 10 g malt extract
(Maltzin Trocken PT, Diamalt, Munich).
To isolate bacteria, soil was suspended and
diluted as described in section 3.3. and plated on
agar plates with 10 g malt extract or 1 g peptonised
milk (Difco, Detroit, MI, U.S.A.) and 250 mg
cycloheximide (Sigma, St. Louis, MO, U.S.A.) per
1. After 5 days at 22°C, single colonies were
transferred to tubes containing nutrient agar.
Fungal strains were identified by the morphology
of their conidia and conidiophores [20].
3. 7. Determination of the nitrification rates of heterotrophic strains
Heterotrophic isolates were tested for their
nitrification capacity under 3 different growth
conditions:
259
3.7.1. Peptone solution. 50 ml of a peptone
solution [21] with peptone from meat, pepsin-digested (Merck, Darmstadt), pH 6.0 in 250 ml
Erlenmeyer flasks was inoculated with 10 ml of a
5 day old culture of the heterotrophic isolates and
incubated at 22 ° C for 21 days.
3. 7.2. Soil extract mixture. Soil extract was prepared by shaking 500 g moist mineral soil with
700 ml distilled water for 18 h at 2°C and centrifuging at 20000 m. s -2. The supernatant was
mixed 3 : 1 with a peptone solution (as above but
1:100 diluted by distilled water). 10 ml Soil extract mixture in 100-ml flasks was inoculated with
the isolates and incubated for 5 days at 15°C.
50-ml samples of soil extract mixture were put
into 250-ml flasks, and 0.5 g cation exchange resin
was added (Dowex 50W × 4, Dow Chemicals,
Midland, loaded with ammonium [22]). After
autoclaving, the samples were inoculated with the
5-day-old cultures of the organism and incubated
for 21 days at 15°C.
After 4, 9, 14 and 21 days of incubation, 6-ml
samples were taken from the cultures with peptone-solution and with soil extract mixture and
analysed for nitrate plus nitrite.
3.7.3. Sterilized soiL 500 g portions of the
mineral soil (0-5 cm) were enclosed in air-tight
polyethylene bags and exposed to gamma-radiation from a 6°Co-source (5.7 Mrad). 10 g samples
of the sterile soil were weighed into sterile petri
dishes. The isolates were grown in 10 ml peptonesolution for 5 days. 0.25 ml aliquots were dripped
onto the soil and incubated for 28 days at 15°C.
3.8. Determination of the oxidation rate of nitrite by
soil fungi
To examine the oxidation from nitrite to nitrate,
fungi were inoculated in 5 ml Czapek-Dox solution (KH2PO4, 1 g; MgSO4, 0.5 g; KC1, 0.5 g~
CaC12 • 2H20, 0.02 g; FeSO4 • 7H20, in EDTA,
0.01 g; solution of trace elements [23] 10 ml;
glucose 1080 mg C and (NH4)2HPO 4 180 mg N;
distilled water 990 ml), pH 4.5. The cultures were
transferred to 20 ml of the same mineral solution
without ammonium and with only 180 mg C. l-1.
After 4 days of growth, 100 mg N . 1-1 as KNO 2
was added. Nitrite oxidation was measured after 2
and 21 days of incubation at 220C.
3.9. Variation of pH and the carbon and nitrogen
sources during nitrification of V. lecanii
Mineral solution according to Czapek-Dox (pH
4.5) was supplied with 126 mg N and 1080 mg C
per 1. 40-ml portions of the liquid were autoclaved
in 250-ml flasks. A 5-day-old culture of V. lecanii
was homogenised in a sterilised blender (Ultraturrax) for 30 s. All flasks of one experiment were
inoculated with 0.5 ml aliquots of a single homogenised culture and incubated (without shaking)
for 21 days at 22°C.
For examining the influence of the pH on
nitrification, it was necessary to ensure a constant
pH during incubation time. The nutrient solutions
were supplied with pH indicators (methyl orange
pH 3.5; bromocresol green pH 5.0; bromothymol
blue pH 7.0). During growth, pH was corrected
every 3 days with sterile NaOH or HC1 until the
colour of the cultures corresponded to a sterile
control flask. The experiment was run in triplicate. Fungal mycelia were collected by filtration, transferred from the filter papers (No. 1507)
to glass dishes, and dried for 20 h at 70°C. The
total nitrogen in the mycelia was determined after
a Kjehldahl digestion [24].
3.10. Nitrate and nitrite determination
Culture samples were either passed through
filter paper No. 5893 (Schleicher and Schtill,
Einbeck) to remove the fungal mycelia, or centrifuged to remove the bacteria. Usually, the sum of
nitrite and nitrate was measured in the clear filtrate
or supernatant.
10 g soil was extracted by shaking with 50 ml
2N KC1 for 1 h, and the suspension was filtered
through paper No. 512 1/2 (Schleicher and Schtill).
Nitrite was measured colorimetrically (autoanalyser, Technicon Instruments, Terrytown,
U.S.A.) as a diazo-pigment [25]. Nitrate was reduced to nitrite by a cadmium-copper column.
The ingredients of the culture liquids did not
interfere with the analysis of nitrite/nitrate. However, humic substances like 'Humuss~iuren' (Roth,
Karlsruhe), or soil extract obtained by autoclaving
or alkaline extraction of soil, disturbed the determination.
The nitrification rates were calculated by the
difference between nitrite and nitrate concentra-
26O
tions of the samples after incubation and of uninoculated controls.
samples contained chemoautotrophic nitrifiers
within the limits of the method (200 bacteria, g - t).
Two forest soils containing hmestone were tested
by the same method. In these soils, at least 2000
ammonia-oxidising bacteria-g-~ were detected.
The enrichment procedure with the Soiling soil
was not successful, neither nitrite nor nitrate being
produced in the enrichment flasks.
4. RESULTS
4.1. Counting and enrichment of autotrophic nitrifiefs
4.2. Counting oxime-N oxidizing organisms
Ammonia oxidisers were counted in samples
collected monthly between April 1982 and March
1983. Nitrite oxidisers were counted in samples in
March 1983 and August 1984. None of the soil
The most-probable-numbers of oxime-oxidising
bacteria varied during one year between 0.28 × 104
Table 2
Nitrification of complex substrates by heterotrophic isolates from acid mineral soil of a beech site
Substrate
Organisms and number
of strains
tested
Strains forming
nitrite plus nitrate
mg N. 1- ~
Peptone
solution
0.05-0.09
All bacteria
All fungi
62
115
43
26
10
9
28
8
6
2
Trichocladium
2
1
Verticillium
1
A cremonium
1
Phoma
1
Mortierella
Trichoderma
Penicillium
mg N- 1-1
Soil extract
mixture
All bacteria
All fungi
Mortierella
Trichoderma
Penicillium
11
115
34
26
10
0.1-0.19
1
-
-
-
-
1
1
-
-
1
-
-
0.1-0.19
0.2-0.4
1
17
6
4
-
2
11
6
1
1
-
2
1
Ferticillium
1
-
1
A cremonium
1
1
-
Phoma
1
-
0.2-0.5
All bacteria
All fungi
Mortierella
Trichoderma
0.2-0.9
1
7
2
2
1
Trichocladiurn
mg N- 1-1
Sterilised
soil
% of nitrifying
strains
24
110
34
24
1
15
4
4
16
31
27
24
-
0.51-0.8
5
2
Penicillium
9
1
2
Trichocladium
2
1
-
Verticillium
1
-
A cremonium
1
-
1
Phoma
1
-
-
4
18
261
and 31.3 x 104. g-1 in O t plus O h, and between
0.01 x 104 and 12 x 10 4. g-1 in the mineral soft.
4.3. Nitrification by randomly isolated heterotrophic
bacteria and fungi
4.3.1. Oxidation of ammonium or organically bound
nitrogen
290 fungal and 60 bacterial strains were isolated from the brown earth. Fungi were classified
according to their macroscopic structure, and partiaUy according to the morphology of their conidiophores and conidia into 140 strains of Mortierella,
60 of Trichoderma, 32 of Penicillium, 6 of Trichocladium and 52 unidentified strains. 115 representative strains were tested for their nitrifying
abilities.
The lowest detectable production of nitrite plus
nitrate during axenic growth was 0.05 mg N-1-1
in peptone solution, 0.1 mg N . 1-1 in soil extract
mixture, and 0.2 nag N . 1-1 in sterile soft. Table 2
shows the numbers of fungi and bacteria producing nitrite or nitrate. With the peptone medium
supplying the organisms with high amounts of
carbon and nitrogen, 31% of the fungi and 16% of
the bacteria were nitrifying, but only one fungus
produced more than 0.2 nag NO 2 + N O f - N . 1-1.
Generally, nitrite or nitrate formation began only
after 9 days of growth (Fig. 1). In the sterilised
soil, easily available energy and nitrogen sources
were considerably enriched, compared to natural
conditions, by dead biomass resulting from
gamma-radiation. Nevertheless, only 24 of the 62
bacterial strains grew in the soft. In this substrate,
4% of the bacteria and 18% of the fungi produced
more than 0.2 nag N . kg-1.
The most active fungi were identified as Trichoderma harzianum and Penicillium brevicompactum (high rates in sterilised soft), Mortierella
alpina and V. lecanii (in peptone solution and soft
extract mixture). Only Acremonium kiliense showed
significant nitrification in all three substrates examined. Two of the nitrifying bacteria were Bacillus spp.
4.3.2. Oxidation of nitrite
Most of the fungi examined oxidised nitrite to
nitrate at high rates. Within 2 days of incubation,
35-45 mg N . 1-1 was oxidised by 5 out of 10
strains (Table 3).
4.4. The influence of nutrient supply and p H on
nitrification by V. lecanii
Further experiments with the most active fungi
(see section 4.3.1) in defined nutrient solutions
Table 3
Oxidation of nitrite to nitrate by soil fungi
Concentration of nitrite and nitrate (nag N.1-1) in the culture
solution, 2 days and 21 days at 22°C after addition of KNO 2
(100 mg N-I - l ) to growing cultures in Czapek-Dox solution
with glucose (180 mg C.I-1). Mean values of 2 cultures are
given.
2 days
21 days
% of
NO~- a
NO{--N NO~--N NO{--N N0~--N
I
3
.7-
visible growth
,, , "
E2
5
10
15
doys
20
25
30
Fig. 1. Nitrification during growth of V. lecanii. Nutrient
solution according to Czapek-Dox, pH 4.5. It, NH~ as nitrogen source; O, arginine as nitrogen source.
No. 174
102
Phoma sp.
83
Mortierella sp.
57
No. 96
27
No. 192
45
A cremonium
kiliense
35
V. lecanii
27
Trichoderma
harzianum
30
Mortierella
alpina
25
Trichoderma sp. 20
Sterile control 105
0
0
17
35
20
79
77
41
32
27
0
0
21
22
27
0
0
34
41
50
32
37
19
8.7
35
41
65
82
40
7.5
45
86
45
45
0
4.0
3.7
105
47
47
3.7
92
93
3
a ~ nitrate in the sum of nitrate plus nitrite.
262
( C z a p e k - D o x with ammonium plus several carbon
sources) showed that only the nitrification rate of
V. lecanii could be raised. This fungus formed 14
mg N O 3 - N . 1-1 with glucose as energy source,
but less than 0.2 mg NO~--N.1-1 with either
acetamide or ammonium plus tartaric, pyruvic,
succinic or citric acid. Nitrite was not detectable
with glucose. With the other energy sources, up to
0.05 mg NO 2 - N - 1-1 was formed• After arginine,
ammonium was the best promoter of nitrification.
In the experiment on variation of the carbon
source, the fungus was freshly isolated, but the
nitrification rates of the isolate decreased with
increasing storage time of the stock culture on
artificial media (Fig. 2), a phenomenon described
also earlier [26,27]• Most nitrate formation occurred after visible growth of mycelium was
terminated (Fig. 1).
How V. lecanii reacts on variation of the acidity was tested in 4 different nutrient solutions
(Fig. 2). With all the substrates, the differences in
nitrification between p H 3.5 and pH 5.0 w e r e
significant ( P = 0.01). The differences between p H
5.0 and p H 7.0 were only significant ( P = 0.05)
with glucose.
0.7
06
E..
05
pHT.0
pH5.0
0.4
~6 0.2
"
0.1
/
L
•
e
I--']
b
N
d
N
Fig. 2. Effect of pH on mtdficadon of K lecaniL The fungus
was grown for 21 days at 22°C in Czapek-Dox solution with:
a, glucose plus (NH4)2HPO4; b, glucose plus t - a ~ i n ¢ ; c,
glucose plus L-aspa.rtic acid; d, sodium succinate p|us
(NH4)2HPO4.
5. DISCUSSION
5.1. Exclusion of autotrophic nitrification
All attempts to find chemoautotrophic nitrifying bacteria in the podzolic brown earth failed. By
counting every month during one year, we could
exclude the possibility that autotrophic bacteria
were active only during distinct seasons• The
highest rates of autotrophic nitrification in liquid
culture amounted to 0.023 pmol NH~- per cell per
h [28]. To account for the nitrification rates in the
mineral soil of about 6.5 mg N - k g - 1 . 2 8 days -1
(mean per annum at 22°C), at least 3 × 104 nitrifying autotrophs, g - l should have been active in
the soil, a number considerably higher than the
detection limit of 200 autotrophs-g-1. The comparative study of limed soils indicated that the
method was adequate for their detection. Addition
of N-Serve (2-chloro-6-(trichloromethyl)-pyridin),
a specific inhibitor for autotrophic ammonia
oxidisers, to the organic layer or mineral soil, did
not inhibit nitrate formation during incubation
[29]. We conclude from these results that nitrate is
not produced by autotrophic bacteria in the
examined soil.
5.2. Oxime: a suitable substrate for nitrification
research in soils?
Some authors have successfully used oximes as
a substrate to isolate heterotrophic bacteria [30,31]
and fungi [32] which produce large amounts of
nitrite. Many microbes producing nitrite from
pyruvic oxime also lived in the Soiling soil. In
nutrient solutions with different carbon and
nitrogen sources inoculated with soil, nitrification
with pyruvic or acetaldehyde oxime was at least 10
times faster than with any other substrate. We
believe that these results are due to the fact that
with oximes, microorganisms do not have to introduce a first oxygen atom onto a nitrogen of redox
state - 3. During oxidation of ammonia or amines,
this is an endergonic critical reaction [33].
Oximes are found in many organisms [34,35]
but concentrations are as low as 0.05 mg oxime-N
• kg-1 in autolysing plant material. We could find
no suggestion in the literature that oximes or
hydroxylamine might be produced in considerable
amounts during humus degradation [36-38].
263
For these reasons, we conclude that oximes can
only be a minor source of nitrate in this soil, and
are misleading 'model' substrates for the examination of heterotrophic nitrification. Our conclusion
is supported by the fact that oximes were oxidised
to nitrite, never to nitrate.
5.3. Heterotrophic nitrifying organisms
With the exception of oxime-containing media,
enrichment cultures and the strains isolated from
them did not produce more nitrite plus nitrate
than strains isolated from the soil under non-selective conditions. A considerable proportion of these
soil organisms was able to produce nitrite or nitrate
in complex media, although at low rates. This
ability was distributed amongst all the genera
identified. These results support the hypothesis
that nitrate is produced in the brown earth by
many different heterotrophic organisms at low
rates, rather than by a few heterotrophic specialists at high rates. The substrate composition and
nutrient concentration seemed to influence the
nitrification rate of every strain differently, as
some isolates produced nitrate only in peptone,
others only in sterilised soil.
The percentage of nitrifiers among heterotrophic isolates was as high as that found among
actinomycetes (30~) [39] and among fungi isolated from red latosols (28%) [26]. In a previous
study, 7% of 978 strains produced more than 0.2
mg NO 2 - N . 1-t in peptone solution [40], a considerably lower frequency than that reported here.
Aspergillus spp. are the most frequently cited
nitrifying fungi. Penicillium spp. [10] and
Mortierella spp. [41] are also known to nitrify.
Trichoderma, Acremonium, Phoma and Verticillium
species have not been described as nitrifying
organisms until now.
We did not distinguish between nitrate and
nitrite formation in the first experiment. However,
the oxidation of nitrite to nitrate (Table 3) was far
more rapid with most of the fungi tested than the
production of nitrate plus nitrite from ammonium
or organically bound nitrogen. We conclude that
nitrite oxidation to nitrate was not the rate-limiting step in the Solling soil.
5.4. Influence of nutrient supply and pH
The rate of nitrification by V. lecanii increased
when the complex substrates were replaced by
defined media. This fungus nitrified about 0.15
mg N. 1-1 in 21 days in peptone broth, but 5.2 mg
N . 1-1 in mineral solution with glucose and ammonium. V. lecanii did not depend on organically
bound nitrogen for nitrate formation. This corresponds with observations on Aspergillus, Penicillium and Arthrobacter [27,42,43]. Our experiments
did not show whether ammonium was the direct
precursor of nitrate. Nitrification at high rates
began after 10 days of growth in all experiments
(Fig. 1), and nitrate formation probably occurred
during aging or autolysis of the mycelium.
The data showed that nitrate concentrations of
replicate cultures could differ to a great extent. No
explanation has been found for this inconsistency
but it showed as well as the time delay of nitrification that it was not an indispensable (energy-providing) process for growth.
Even though the process of nitrification by
heterotrophs has been well established, as yet little
evidence exists about their ability to nitrify under
acid conditions. Most heterotrophs examined
nitrify optimally under neutral or slightly alkaline
conditions [39,43,44]. Even a fungus isolated from
an acid soil nitrifies optimally in a medium at pH
8.3 [8]. Others show no nitrification at pH values
below 6.0 [45,30]. According to these observations,
also the activity of heterotrophic nitrifiers in acid
soils should depend on neutral microsites. Only a
Micromonospora sp. and a Aspergillus flavus from
acid soils nitrify at the same rate at pH 5.0 as at
pH 7.0 [41,26]. The nitrification rate of V. lecanii,
on the other hand, was greatly enhanced by acid
conditions on different substrates. The influence
of pH could probably explain why V. lecanii produced nitrate only with glucose (pH dropped to
2.7 during growth) and not with salts of organic
acids (pH rose to 6.2-8.0).
That the nitrification rates of all isolates were
higher in the sterile soil (pH 3.5) than in peptone
broth (pH 6.0) implies that V. lecanii is no exception amongst the isolated fungi. These results agree
with those of Lettl [46] who found higher nitrification rates at pH 4.0 than at pH 8.0 among heterotrophic bacteria isolated from the organic layers of
SO2-polluted forest sites.
We have shown that heterotrophic nitrification
264
c a n o c c u r a n d e v e n r e a c h its o p t i m u m at l o w p H
v a l u e s w h i c h s u p p r e s s a u t o t r o p h i c nitrifiers. T h i s
f i n d i n g , b e s i d e s t h e l a c k o f a u t o t r o p h s a n d the
a b i l i t y o f m a n y o f t h e h e t e r o t r o p h s to nitrify,
m a d e it v e r y p r o b a b l e t h a t n i t r i f i c a t i o n in a c i d
soils s u c h as t h e p o d z o l i c b r o w n e a r t h c a n b e
accomplished mainly by heterotrophic organisms.
ACKNOWLEDGEMENTS
W e t h a n k p r o f e s s o r D r . K . H . D o m s c h for h e l p f u l l y s u p p o r t i n g t h e i d e n t i f i c a t i o n o f fungi, a n d
Ilse S c h m i d t a n d K. B r i n k m a n n for t e c h n i c a l
a s s i s t a n c e . T h i s w o r k was s u p p o r t e d b y a g r a n t
from the Deutsche Forschungsgemeinschaft.
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[2] Kriebitzsch, W.-U. (1978) Stickstoffnachlieferung in
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[3] Robertson, G.P. (1982) Nitrification in forested ecosystems. Phil. Trans. R. SOc. Lond. B 296, 445-457.
[4] Hankinson, T.R. and Schmidt, E.L. (1984) Examination
of an acid forest soil for ammonia- and nitrite-oxidizing
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[5] Josserand, A. and Bardin, R. (1981) Nitrification en sol
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(genre Nitrobacter) dans un sol forestier sous rrsineux.
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