FEMS Microbiology Ecology 52 (2005) 21–29
www.fems-microbiology.org
Nitrification and occurrence of salt-tolerant nitrifying bacteria
in the Negev desert soils
Ali Nejidat
*
Department of Environmental Hydrology and Microbiology, Institute for Water Sciences and Technologies, The Jacob Blaustein
Institute for Desert Research, Ben-Gurion University of the Negev, Sede Boqer Campus, 84990 Midreshet Ben-Gurion, Israel
Received 17 May 2004; received in revised form 16 August 2004; accepted 12 October 2004
First published online 14 November 2004
Abstract
Ammonia oxidation potential, major ammonia oxidizers and occurrence of salt-tolerant nitrifying bacteria were studied in soil samples collected from diverse ecosystems along the northern Negev desert. Great diversity in ammonia oxidation potential was observed
among the soil samples, and ammonia oxidizers were the rate-limiting step of nitrification. Denaturing gradient gel electrophoresis and
partial 16S rRNA gene sequences indicate that members of the genus Nitrosospira are the major ammonia oxidizers in the natural desert
soil samples. Upon enrichment with different salt concentrations, salt-tolerant nitrifying enrichments were established from several soil
samples. In two enrichments, nitrification was not inhibited by 400 mM NaCl. Electrophoretic analysis and partial 16S rRNA gene
sequences indicate that Nitrosomonas species were dominant in the 400 mM salt enrichment. The results point towards the potential
of the desert ecosystem as a source of stress-tolerant nitrifying bacteria or other microorganisms with important properties.
Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved.
Keywords: Ammonia oxidizers; Denaturing gradient gel electrophoresis; Nitrosospira; Nitrosomonas; Salt tolerance
1. Introduction
Nitrification (sequential oxidation of ammonia to nitrate) is carried out by the lithoautotrophic, Gram-negative nitrifying bacteria classified within the family
Nitrobacteriaceae. This is a physiologically defined
group of ammonia- and nitrite-oxidizing genera that
can be differentiated according to morphological characteristics [1]. Ammonia- and nitrite-oxidizers derive their
energy from the oxidation of ammonia (NH3) and nitrite ðNO
2 Þ, respectively [2]. Nitrification provides the
linkage between ammonia and nitrate in nature and is
thus a key process in nitrogen cycling. Based on 16S
rRNA gene sequences, the ammonia oxidizers that are
*
Corresponding author. Tel.: +972 8 6596832; fax: +972 8 6596831/
6909.
E-mail address: [email protected].
considered to be the rate-limiting step in nitrification
[3–5] are classified [6] into three genera: Nitrosomonas,
Nitrosospira and Nitrosococcus.
With the use of specific PCR primers based on the
16S rRNA gene and amoA gene sequences, together
with denaturing gradient gel electrophoresis (DGGE)
[3,4,7,8], the community structures of ammonia oxidizers have been determined in many ecosystems [8–10].
These include different types of aquatic [11–19] and terrestrial [16,20–22] environments. Although with less
attention, the detection and community structure analysis of nitrite oxidizers in different environments have
also been done in several studies [23–26].
Desert soils are characterized by low water activity,
and poor organic matter- and nitrogen content.
Although nitrogen cycling is a key process in the functioning of desert ecosystems [27–29], the community
structure of nitrifying bacteria in desert soils has not
0168-6496/$22.00 Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved.
doi:10.1016/j.femsec.2004.10.011
22
A. Nejidat / FEMS Microbiology Ecology 52 (2005) 21–29
been studied. The Negev desert occupies almost two
thirds of the land area in Israel. Average annual rainfall
ranges from 200 to 30 mm between Beer Sheva in the
north and Eilat in the south. Winter season in the Negev
is short and rainfall is erratic, causing a cycle of wet and
dry conditions. Rapid increase in moisture kills a significant portion of soil microbial biomass via osmotic
shock of those that have accumulated solutes to resist
desiccation during soil drying [30,31]. Death of microbial biomass supplies nutrients that allow a burst in
microbial activity, which may lead to depletion of soil
oxygen, allowing anaerobic processes such as denitrification to occur [31] and thus causing nitrogen loss. Many
desert ecosystems are characterized by patchiness in distribution of vegetation. In the Negev desert, there are
two types of patches [32]: (1) macrophytic patches consisting of herbs and shrubs that are strongly limited in
nitrogen content, and (2) microphytic patches consisting
of algae, cyanobacteria, lichens and mosses that are rich
in nitrogen due to N2 fixation [29]. The combination of
denitrifcation and NH3 volatilization [29] causes loss of
a significant portion of nitrogen.
In addition to this naturally fragile desert ecosystem,
there has recently been a tendency to relocate major
industrial activities to the Negev desert, away from more
populated areas. Furthermore, intensive agricultural
practices including fertilization and irrigation with saline and wastewater effluents affect the community structure of ammonia-oxidizing bacteria [33,34]. The Negev
desert may thus provide an excellent natural research
field for studies of evolutionary diversity of bacterial
communities as affected by gradients of environmental
conditions combined with anthropogenic activities.
This study was carried out bearing in mind the significance of nitrification in desert soils and the possibility
that the harsh desert environmental (extreme temperature fluctuations, low water activity, high light intensities and nutrient limitations) may select for novel
microbial species that tolerate non-optimal growth conditions. The study aimed at examining nitrification
activity and occurrence of salt-tolerant nitrifying bacteria, and at identifying major ammonia-oxidizing bacteria in natural soil samples collected from the northern
Negev desert. The most important findings of the study
are the presence of salt-tolerant (400 mM NaCl) ammonia- and nitrite-oxidizers and the evidence that Nitrosospira species dominate the ammonia-oxidizer
community in natural desert soil.
Fig. 1. Map of Negev desert, Israel, showing the rainfall gradient
(mm). Asterisks indicate the locations where soil samples were
collected. Modified from Shem-Tov et al. [35].
samples were collected in one week after two rainy days
during the winter of the year 2000.They included samples (0–10 cm deep) from the vicinity of a chemical
industrial park, open fields, agricultural fields, and others. Soil samples were collected into polyethylene bags
and after about 2 h stored at 4 °C until use. Soils were
extracted with distilled water in a 1:1 ratio, and the supernatant was used for the determination of total organic
carbon (TOC) by the combustion-infrared method [36],
pH and conductivity using a TDS/Conductivity Meter.
Total ammonia-nitrogen was determined with the Nessler method [36], nitrite with the sulfanilamide method
[37] and nitrate with the NAS method [37]. Water holding capacity (WHC) test was performed as described
[38]. Relevant physical and chemical characteristics of
the samples are summarized in Table 1.
2.2. Ammonia oxidation potential
Ten grams of soil were added to 20 ml medium (pH
7.8) containing 25 mM K2HPO4, 0.5 mM (NH4)2SO4
and 10 mM KClO3 to inhibit nitrite oxidizers and incubated in a dark room with continuous shaking (200 rpm)
for 6 h. Nitrite accumulation was measured as above.
2. Materials and methods
2.3. Enrichment cultures
2.1. Sample collection and analysis
Eleven natural soil samples were collected from the
area between Beer-Sheva and Sede Boqer (Fig. 1). Soil
Enrichment of nitrifying bacteria was achieved by
incubating 5 g soil samples in 30 ml mineral medium
[39] containing (per liter): 1.3 g (NH4)2SO4, 0.5 g
A. Nejidat / FEMS Microbiology Ecology 52 (2005) 21–29
23
Table 1
Chemical characteristics of the soil samples used in this study
Sample
code
Soil sample origin
pH
Conductivitya
WHC (%)
TOC (ppm C)
NHþ
4 (lM)
NO
2 (lM)
NO
3 (lM)
S1
S2
S3
S4
S5
S6
S7
S8
S9
S11
S12
Wet soil from Nahal Hovav
Soil under Atri plex bush
RH, under Tamarix trees
Soil from agricultural plot
Soil under Acacia trees
Soil under Eucalyptus trees
Olive groove irrigated with wastewater
Pistachio groove irrigated with waste water
Agricultural field, Sede Boqer
Soil covered with snails, Sede Boqer
Open field, Sede Boqer
7.52
7.38
7.95
7.29
7.20
7.54
7.15
7.02
7.37
7.45
7.59
56
330
39
4.9
5.4
7.8
73
34
16.4
27.7
6.1
20.94
28.87
33.68
30.03
27.7
46.05
37.3
32.8
32.09
43.88
29.55
12.46
65.8
110.67
13.7
8.6
183
49
21.16
119
68.4
233
0
795
14.7
0
0
0.02
2.7
0
0
415
0.05
0
0.24
137
0
0
0.03
0.81
0.13
0
23
0.35
89
13
1922
235
18.13
32
1066
236.4
160
6262
28.8
a
Equivalent (mM NaCl).
K2PO4, 0.05 g MgSO4, 4 mg CaCl2, 3.8 mg FeNaEDTA,
0.1 mg NaMoO4 Æ 2H2O, 0.2 mg MnCl2, 0.002 mg CoCl2 Æ 6H2O, 0.1 mg ZnSO4 Æ 7H2 O, 0.02 mg CuSO4 Æ 5H2O and adjusted to pH 7.8–8.0 with 25% K2CO3
solution. Every two weeks, 50% of the supernatant
was replaced with fresh medium and the pH was adjusted to 7.8 using 50% K2CO3 solution. This was done
for two months. Enrichments were performed in three
media: standard mineral medium (SM); low-salt medium containing 200 mM NaCl (LSM), and high-salt
medium containing 400 mM NaCl (HSM). To allow full
nitrification, 5 ml of the supernatant (after mixing and
settling) were added to 45 ml mineral medium (SM,
LSM, HSM) in 100 ml flasks and incubated at 25 °C
in the dark. Ammonia concentration was adjusted (6–
7.5 mM) using (NH4)2SO4. The initial concentrations
of NO
2 and NO3 carried over were taken into consideration in the calculations.
2.4. DNA isolation, PCR amplification and DGGE and
sequence analyses
Total DNA was extracted from soil samples using a
commercial kit (UltraClean Soil DNA Isolation Kit,
MO BIO Lab. Inc., Solana Beach, CA). A fragment of
DNA (465 bp) from the 16S rRNA gene of ammonia
oxidizers belonging to the Betaproteobacteria [40] was
PCR-amplified using the forward primer CTO 189fA
with attached GC clamp: ccgccgcgc ggcgggcgggg cggg
ggcac ggggGGAGRAAAGCAGGGGATCG and a reverse primer CTO654r: CTAGCCTTGTAGTTTCAAACGC. PCR amplification (initial denaturation
step of 94 °C for 4 min and 30 cycles of denaturation
at 94 °C for 1 min, annealing at 55 °C for 1 min, elongation at 72 °C for 2 min and a final elongation step
for 5 min) was carried out in 50 ll volumes [10 mM
Tris–HCl pH 8.3, 50 mM KCl, 1.1 mM MgCl2, 0.01%
gelatin, 200 lM of each of four deoxynucleotide triphosphates, 25 pmol of each primer, 1 ll of template DNA
and 1 unit of REDTaq DNA polymerase (Sigma, St.
Louis, MO)] using a thermocycler (Minicyclerä, MJ Research, Watertown, MA). A single band of the expected
size was obtained when PCR products were visualized in
ethidium bromide-stained 1.5% agarose gels. DGGE
analysis was performed with a Dcodeä Universal Mutation Detection System (Bio-Rad, Herculis, CA) under
the following conditions: 1-mm thick 8% polyacrylamide gels, a denaturant gradient of 35–50% urea–formamide at 60 °C, 240 V for 4 h. Ethidium bromide-stained
gels were photographed on a UV transilluminator table
with a digital camera. Images were captured and inversed by NIH Image version 1.62 (Wayne Rasband,
National Institute of Health, USA) and further processed with Photoshop version 6 (Adobe). Major bands
were carefully excised under UV light and DNA was extracted from gel slices as described elsewhere [34] and
cloned in plasmid pTZ57R using the InsT/Aä PCR
Product Cloning Kit (MBI Fermentas, Hanover, MD).
DNA sequencing was performed at the Ben-Gurion
University of the Negev Equipment Center, The Institute for Applied Biosciences using an ABI Prism 377
DNA sequencer (Perkin–Elmer). To exclude the possibility of mixed sequences in the same band, two to three
clones were randomly picked for sequencing from each
band and identical sequences were obtained. Sequences
were analyzed using the BLAST [41] similarity search
program in order to find the most similar sequences in
the database. The sequences obtained from this study
were deposited in the GenBank and assigned the accession numbers AY605671–AY605680, AY123791 and
AY695808. The accession numbers of other bacteria that
were included in the phylogenetic tree but were not indicated in Table 2 are: L35505, AJ298737, AY123792,
AF272425, X84658, AJ298746, M96405, AY123807,
AB070982,
AJ298727,
AF338213,
AY690336,
AJ243144 and M34131.
Pure cultures of ammonia oxidizers used as controls,
that is Nitrosomonas europaea Nm50, Nitrosospira
sp. L115 and Nitrosospira sp. 40KI, were kindly supplied by Dr. Agot Aakra (Agricultural University of
24
A. Nejidat / FEMS Microbiology Ecology 52 (2005) 21–29
Table 2
The most similar ammonia oxidizers in the database compared to the 16S rRNA gene sequences generated from the major electrophoretic bands
marked in Fig. 3
Sample
Most similar bacterium
Percentage identity
Accession number
S1
S3
S4
Nitrosospira sp. L115
Nitrosospira sp. PJA1
Nitrosospira sp. Nsp41
Nitrosospira sp. Nv6
Nitrosospira sp. R3c5
Nitrosospira sp. Nv6
Nitrosospira sp. Nsp41
Nitrosospira sp. R3c5
Nitrosospira sp. R3c5
Nitrosospira sp. N15
Nitrosospira sp. Nsp57
Nitrosomonas sp. Is343
99
97
99
99
99
99
99
99
96
99
86
98
gb, AY123796a
gb, AF353163
gb, AY123788
gb, AY123805
gb, AF386756
emb, AJ298747b
gb, AY123788
gb, AF386756
gb, AF386756
gb, AY123812
gb, AY123791
emb, AJ621032
Ac
B
S6
S7
S8
S9
S11
S12
a
b
c
gb, Genbank database.
emb, EMBL database.
A and B are the lower and upper band, respectively, marked in Fig. 3 for S6.
Norway). Nitrosomonas eutropha N904 was kindly supplied by Prof. Eberhard Bock (University of Hamburg,
Germany).
3. Results and discussion
3.1. Ammonia oxidation in soil samples
The results presented in Table 1 indicate that ammonia and nitrite did not accumulate in the environmental
samples examined, except in soil sample S2 where most
of the inorganic nitrogen was in the form of ammonia.
This soil sample was collected from beneath an Atriplex
bush. Atriplex plants are halophytes that accumulate
and sequester inorganic salts (Na+, K+, Cl) in their
leaves for osmotic potential adjustment [42]. Upon
Nitrite accumulation (uM)
25
20
15
10
5
0
S1 S2 S3 S4 S5 S6 S7 S8 S9 S11 S12
Soil sample
Fig. 2. Potential for ammonia oxidation in soil samples. Soil samples
correspond to those described in Table 1.
decomposition salts accumulate beneath the bush. These
results may imply that in soil sample S2, ammonia oxidation was totally inhibited, which could be due to high
salt concentration in this soil sample (Table 1) and the
lack of ammonia oxidizers adapted to such salt concentration. It is also possible that ammonia oxidizers, which
were osmotically adapted to the high salt conditions
died due to osmotic shock, since the soil samples were
collected after two rainy days. Some ammonia accumulation was also observed in soil sample S11, which was
collected from soil covered with snails. However, the
high concentration of nitrate indicates that nitrification
was occurring with high rates. Snails have been shown
to excrete ammonia in their feces [43] and this can explain the relatively high concentration of inorganic
nitrogen (ammonia and nitrate). To examine the possibility of the measured quantities of inorganic nitrogen
(ammonia, nitrite, nitrate) reflecting the nitrification
activity in the soil samples, potential for ammonia
oxidation was tested (Fig. 2). The lack of nitrite accumulation (Table 1) indicated that nitrite oxidizers were not
rate-limiting and thus, their activity was not examined.
The results shown in Fig. 2 support the data in
Table 1. Soil sample S2 shows a very low rate of ammonia oxidation, while soil samples S3 and S7 showed high
ammonia oxidation rates. Surprisingly, ammonia oxidation was not exceptionally high in sample S11, as was expected from data in Table 1. High rate of ammonia
oxidation was observed in soil sample S6 although total
inorganic nitrogen (ammonia, nitrate and nitrite) was
not particularly high (Table 1). These results suggest
that ammonia oxidizers are the rate-limiting step of
nitrification in environmental desert soil. They also indicate that no correlation can be made between nitrification potential and soil nitrogenous compounds
(ammonia, nitrite, nitrate content). The nitrification
potential measured in soil samples S3, S6, S7, and S12
A. Nejidat / FEMS Microbiology Ecology 52 (2005) 21–29
25
to that measured in the same soils after fertilization
[21]. However, in another study [33], fertilization had
no effect on nitrification potential. This indicates that,
in addition to substrate availability, other factors such
as soil properties (pH, salinity, moisture), vegetation
and environmental factors (temperature, irradiation)
are crucial to the development of population size and
activity of nitrifying bacteria.
3.2. Identification of ammonia oxidizers
Fig. 3. DGGE analysis of PCR-amplified 16S rRNA gene fragments
from total DNA from soil samples using the CTO189fA and CTO654r
primers. Lanes 1, 2, 3, and 4 are marker strains: Nitrosospira sp. L115,
Nitrosospira sp. 40KI, Nitrosomonas europaea Nm50 and Nitrosomonas eutropha N904, respectively. Soil samples S1–S12 correspond to
those described in Table 1. Asterisks indicate the bands that were
recovered, reamplified, cloned and sequenced. DNA was not successfully amplified from samples S2 and S5 with the CTO primers.
was within the range of that measured in non-fertilized
successive grasslands [22] but was very low compared
Dominant ammonia oxidizers in natural desert soil
samples described in Table 1 were identified based on
partial sequences of the 16S rRNA gene. These sequences (465 bp) were generated from major DNA
bands resolved in DGGE analysis. Denaturing gradient
gel analysis of 16S rRNA gene sequences, amplified
from DNA from several environmental soil samples
and marker strains, is shown in Fig. 3. Major electrophoretic bands (marked on the gel) that showed similar
migration behavior as the marker strains were reamplified, cloned and sequenced. The results of the BLAST
similarity search are summarized in Table 2. All DGGE
bands sequenced were most similar to Nitrosospira-like
sequences except sample S12. It can be noticed that
the major band of S12 migrated significantly less than
the others and was approximately at the same height
Fig. 4. Phylogenetic tree constructed for partial 16S rRNA gene sequences of the environmental clones described in Table 2 and most identical clones
from database. DNA sequences of 380-443 bp were used in the aliment. The tree is based on the results of Neighbor-joining analysis using the RDP
Phylip interface (http://rdp.cme.msu.edu./html/). AOB, ammonia-oxidizing bacteria.
26
A. Nejidat / FEMS Microbiology Ecology 52 (2005) 21–29
as the Nitrosomonas marker. The results in Table 2 were
summarized in a phylogenetic tree (Fig. 4). All Nitrosospira clones grouped to cluster 3 of the Nitrosospira
while clone S12 grouped within cluster 6 of the Nitrosomonas genus [3]. Clone HS61 grouped with the Nitrosomonas communis cluster [8]. These findings conform to
the typical conclusion that Nitrosospira dominate soil
environments [13]. It is difficult to explain the uniqueness of sample S12 in this relation. However, this is
the only sample that was collected from pristine soil
without vegetation and was not touched by any human
activity. Although beyond the range of the marker
strains, in some samples (e.g. S8, S7, S4) other bands
can be identified (Fig. 3), which does not rule out the
presence of other ammonia oxidizers in smaller
numbers.
3.3. Establishment of salt-tolerant nitrifying enrichments
(a)
Ammonia concentration (mM)
7
SM
HSM
LSM
6
5
4
3
2
1
0
0
2
4
6
8
10
12
14
16
5
4
3
2
1
0
0
5
2
8
(d)
4
6
8
10
12
14
6
5
4
3
2
1
0
4
6
8
10
12
14
16
4
6
8
10
12
14
16
4
6
8
10
Time (days)
12
14
16
(e)
1
0.5
0
2
(c)
4
Nitrate concentration (mM)
4
3
2
1
0
2
1.5
0
16
SM
LSM
HSM
7
2
(b)
Nitrite concentration (mM)
Nitrite concentration (mM)
6
Nitrate concentration (mM)
Ammonia concentration (mM)
Enrichment cultures were established from 11 soil
samples in three media (SM, LSM, HSM) as described
in Section 2. The enrichments were screened for nitrification activity at three pH values (pH 5, 8, 10) and three
salt concentrations. All samples tested showed maximal
nitrification activity at pH 8 and significantly lower
activity at pH 5 and 10 (data not shown). However,
nitrification activity showed a great diversity in response
to salt concentration among the soil samples. Two
examples that exhibited either a salt-tolerant (400 mM)
nitrifying enrichment culture (S6) or a gradual inhibition
of nitrification (S4) by increased salt concentration are
presented in Fig. 5. This type of analysis was performed
for all enrichments (three enrichments for each of the 11
soil samples) and the results are summarized in Fig. 6.
Several patterns can be observed concerning the effect
of salt. In the enrichments of samples S1 and S6 both
ammonia- and nitrite-oxidizers are tolerant to 200 and
400 mM salt levels. The natural soil sample S1 did not
show significant ammonia oxidation (Fig. 2). However,
after enrichment it showed high ammonia- and nitriteoxidation. Thus the low activity observed in the natural
sample could be due to nitrification inhibition by con-
0
2
4
6
8 10
Time (days)
12
14
16
(f)
3.5
3
2.5
2
1.5
1
0.5
0
0
2
Fig. 5. Nitrification (ammonia consumption, nitrite and nitrate accumulation) in soil enrichment cultures. SM, LSM and HSM are mineral medium,
mineral medium containing 200 mM NaCl and mineral medium containing 400 mM NaCl, respectively. The initial ammonium sulfate concentration
was adjusted to 5.5–7.5 mM. (a)–(c) are for soil sample S6 and (d)–(f) are for soil sample S4.
A. Nejidat / FEMS Microbiology Ecology 52 (2005) 21–29
Ammonia consumed (mM)
8
(a)
27
SM
LSM
HSM
7
6
5
4
3
2
1
0
S1
6
S2
S3
S4
S5
S6
S7
S8
S9
S11 S12
Soil sample
(b)
Nitrate produced (mM)
5
4
3
2
1
0
S1
S2
S3
S4
S5 S6 S7
Soil sample
S8
S9 S11 S12
Fig. 6. Nitrification in soil enrichments – ammonia concentration and
nitrate concentration after 15 days of incubation. Conditions are as
described for Fig. 4. Soil samples correspond to those described in
Table 1.
taminants, since this sample was collected from a contaminated site, or due to the very low ammonia concentration in the soil (Table 1).
In enriched soil samples S2 and S3, ammonia- and nitrite-oxidizers were highly sensitive to salt concentrations. The enriched sample S2 in SM showed high
ammonia oxidation rates compared to the natural sample (Fig. 6). This supports the conclusion that salt-tolerant nitrifying bacteria did not develop under the
Atriplex bush. Ammonia oxidizers in samples S4, S5,
S7, S8, S9, S11 and S12 showed tolerance to 200 mM
salt and in some samples enhanced activity was observed
(samples S5, S7, S8). However, ammonia oxidation was
inhibited to different degrees at 400 mM salt. Nitrite oxidizers in samples S4 and S5 were inhibited by both salt
concentrations. In samples S8, S11 and S12 nitrite oxidizers were tolerant to 200 mM salt but severely inhibited at 400 mM salt. On the other hand, nitrite
oxidizers in samples S7 and S9 were significantly enhanced at 200 mM salt and inhibited to different degrees
at 400 mM salt. Ammonia- and nitrite-oxidizers thus responded differentially to salt concentrations, which
might result in an unbalanced nitrification in the highly
diverse desert ecosystems. On average, 87.8 ± 9.2% of
Fig. 7. DGGE analysis of cloned bands from the natural soil sample
S6 and HSM enrichment from the same soil sample. Lanes 1 and 2 are
16S rRNA gene fragments amplified from the genome of the marker
strains Nitrosomonas europaea 19718 and Nitrosospira sp. A4, respectively. Lanes 3, 4 and 5 are the DNA bands marked for soil sample S6
in Fig. 3 and a major band that was detected in a DGGE analysis of
16S rRNA gene amplified from total DNA extracted from HSM
enrichment of soil sample S6, respectively. The cloned fragments were
cut from plasmid with restriction enzyme, purified and used in DGGE
analysis. All bands were amplified using the CTO189fA and CTO654r
primers.
the difference between the consumed ammonia and nitrate produced was recovered as nitrite (data not
shown). The rest may have been lost due to ammonia
volatilization, consumption of the nitrogenous compounds by heterotrophs in the enrichments and the production of N2O, NO and N2 gases [10].
Nitrification in the enrichments of S1 and S6 was
highly tolerant to 400 mM salt (2.3% salt), which is close
to seawater salt concentration (3% salt). For comparison,
ammonia oxidation by N. europaea showed 60% inhibition at 200 mM NaCl and about 100% inhibition at 400
mM [44]. On the other hand, nitrite oxidation by Nitrobacter agilis showed only slight inhibition at 200 mM NaCl
as was found for nitrite oxidizers in several soil samples
(S1, S6, S7, S8, S9, S11, S12) and only 20% inhibition at
500 mM salt [45]. It was interesting to investigate whether
the Nitrosospira species identified as major ammonia oxidizers in the natural samples S1 and S6 (Table 2) remained
after salt enrichment. Unfortunately, salt enrichments
from S1 lost their activity after few months. However,
DNA was isolated from S6 enrichment in HSM and analyzed using the CTO primers. The major band (HS61)
cloned from this enrichment and bands A and B (Fig. 3)
were examined in DGGE analysis (Fig. 7). The results
showed that the enrichment band migrated higher than
the natural sample bands. This DNA fragment was
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A. Nejidat / FEMS Microbiology Ecology 52 (2005) 21–29
sequenced and the most similar ammonia oxidizers (98%
identity) were Nitrosomonas nitrosa (accession number
AJ298740) and Nitrosomonas sp. Nm148 (accession number AY123792) and it clustered to Nitrosomonas species
(Fig. 4). This result indicates that Nitrosospira species
dominated the natural soil samples and upon salt enrichment, Nitrosomonas sp. were selected as the major ammonia oxidizers. Shifts in the ammonia-oxidizing
community structure upon enrichments have been observed before in other environmental samples. Nirosomonas species were detected in lake water sediments and
activated sludge [13] and in soil [16] only in enrichments.
The possibility that the high ammonium concentration
used in the enrichment medium compared to its concentration in the soil samples may have affected the species
selection cannot be ruled out. Ammonium concentrations
of 3000 mg of N l1 were shown to cause irreversible
changes in the community structure of nitrifying bacteria
from wastewater compared to lower concentrations [46].
A summary of the salt requirements of cultured ammonia
oxidizers indicated that halotolerant and obligately halophilic species of ammonia oxidizers belonging to the betaproteobacteria are associated with Nitrosomonas sps. [9].
However, in addition to halophilic Nitrosococcus species
(Nitrosococcus oceani andNitrosococcus halophilus), both
Nitrosomonas- and Nitrosospira-like sequences were detected in marine environments [3], suggesting that halotolerance is a cross-genera phenomenon among the
ammonia oxidizers. The results presented in this study
indicate that nitrification shows a great diversity in natural soils of the Negev desert and that activity of ammonia
oxidizers is the rate-limiting step of nitrification. Nitrosospira species dominate the soils of the northern Negev desert and salt-tolerant nitrifying enrichments can be
established, taking into consideration a possible shift in
the community structure. Salt-tolerant nitrifying enrichments can be used in augmentation practices in order to
enhance nitrification in saline environments. Indeed, the
salt-tolerant enrichment S6 was successfully used as a
nitrification starter culture in a laboratory-scale shrimp
aquaria biofilter using saline water pumped from the Negev aquifer [47].
Acknowledgements
[2]
[3]
[4]
[5]
[6]
[7]
[8]
[9]
[10]
[11]
[12]
[13]
[14]
[15]
[16]
[17]
I thank Dr. Zeev Ronen and Lilach Iasur for their
help in preparing the phylogenetic tree and Zoe Grabinar for language improvements in the manuscript.
[18]
References
[1] Bock, E., Koops, H.S. and Harms, H. (1989) Nitrifying bacteria
In: Autotrophic Bacteria (Schlegel, H.G. and Bowien, B., Eds.),
[19]
pp. 81–96. Springer, Berlin, Heidelberg, New York, London,
Paris, Tokyo.
Arp, D.J., Sayavedra-Soto, L.A. and Hommes, N.G. (2002)
Molecular biology and biochemistry of ammonia oxidation by
Nitrosomonas europaea. Arch Microbiol. 178, 250–255.
Kowalchuk, G.A. and Stephen, J.R. (2001) Ammonia-oxidizing
bacteria: a model for molecular microbial ecology. Ann. Rev.
Microbiol. 55, 485–529.
Prosser, J.I. and Embley, T.M. (2002) Cultivation-based and
molecular approaches to characterization of terrestrial and
aquatic nitrifiers. Antonie van Leeuwenhoek 81, 165–179.
Focht, D.D. and Verstraete, W. (1977) Biochemical ecology of
nitrification and denitrification In: (Alexander, Ed.), Advances in
Microbial Ecology, vol. 1, pp. 135–214. Plenum Press, New York,
London.
Head, I.M., Hiorns, W.D., Embley, M., McCarthy, A.J.
and Saunders, J.R. (1993) The phylogeny of autotrophic
ammonia-oxidizing bacteria as determined by analysis of
16S ribosomal RNA gene sequences. J. Gen. Microbiol.
139, 1147–1153.
Taske, A., Alm, E., Regan, J.M., Toze, S., Rittmann, B.E. and
Stahl, D.A. (1994) Evolutionary relationship among ammoniaand nitrite-oxidizing bacteria. J. Bacteriol. 176, 6623–6630.
Purkhold, U. and Wagner, M. (2000) Phylogeny of all recognized
species of ammonia oxidizers based on comparative 16S rRNA
and amoA sequence analysis: implications for molecular diversity
surveys. Appl. Environ. Microbiol. 66, 5368–5382.
Koops, H-P. and Pommerening-Roser, A. (2001) Distribution of
the nitrifying bacteria emphasizing cultured species. FEMS
Micrbiol. Ecol. 37, 1–9.
Bothe, H., Jost, G., Schloter, M. and ward, B.B. (2000) Molecular
analysis of ammonia oxidation and denitrification in natural
environments. FEMS Microbiol. Rev. 24, 673–690.
Nejidat, A. and Abeliovich, A. (1994) Detection of Nitrosomonas
spp. by polymerase chain reaction. FEMS Microbiol. Lett. 120,
191–194.
McCaig, A.M., Embley, T.M. and Prosser, J.I. (1994) Molecular
analysis of enrichment cultures of marine ammonia oxidizers.
FEMS Microbiol. Lett. 120, 363–368.
Hiorns, W.D., Hastings, R.C., Head, I.M., McCarthy, A.J.,
Saunders, J.R., Pickup, R.W. and Hall, G.H. (1995) Amplification of 16S ribosomal RNA genes of autotrophic ammoniaoxidizing bacteria demonstrates the ubiquity of nitrosospiras in
the environment. Microbiology 141, 2793–2800.
Hovanec, T.A. and Delong, E.F. (1996) Comparative analysis of
nitrifying bacteria associated with freshwater and marine aquaria.
App. Environ. Microbiol. 62, 2888–2896.
Burrel, P.C., Phalen, C.M. and Hovanec, T.A. (2001) Identification of bacteria responsible for ammonia oxidation in freshwater
aquaria. Appl. Environ. Microbiol. 67, 5791–5800.
Stephen, J.R., McCaig, A.E., Smith, Z., Prosser, J.I. and Embley,
T.M. (1996) Molecular diversity of soil and marine 16S RNA gene
sequences related to b-subgroup ammonia-oxidizing bacteria.
Appl. Environ. Microbiol. 62, 4147–4154.
McCaig, A.E., Phillips, C.J., Stephen, J.R., Kowalchuk, G.A.,
Harvey, S.M., Herbert, J.R., Embley, T.M. and Prosser, J.I.
(1999) Nitrogen cycling and community structure of proteobacterial b-subgroup ammonia-oxidizing bacteria within polluted
marine fish farm sediments. Appl. Environ. Microbiol. 65, 213–
220.
Phillips, C.J., Smith, Z., Embley, T.M. and Prosser, J.I. (1999)
Phylogenetic differences between particle-associated and planktonic ammonia-oxidizing bacteria of the b subdivision of the class
proteobacteria in the Northwestern Mediterranean sea. Appl.
Environ. Microbiol. 65, 779–786.
Ward, B.B., Martino, D.P., Diaz, M.C. and Joye, S.B. (2000)
Analysis of ammonia-oxidizing bacteria from hypersailine Mono
A. Nejidat / FEMS Microbiology Ecology 52 (2005) 21–29
[20]
[21]
[22]
[23]
[24]
[25]
[26]
[27]
[28]
[29]
[30]
[31]
[32]
[33]
lake, California, on the bases of 16S rRNA sequences. Appl.
Environ. Microbiol. 66, 2873–2881.
Hasting, R.C., Butler, C., Singeleton, I., Saunders, J.R. and
McCarthy, A.J. (2000) Analysis of ammonia-oxidizing bacteria
populations in acid forest soil during conditions of moisture
limitation. Lett. Appl. Microbiol. 30, 14–18.
Kowalchuk, G.A., Stienstra, A.W., Heilig, G.H., Stephen, J.R.
and Woldendrop, J.W. (2000) Changes in the community structure of ammonia-oxidizing bacteria during secondary succession
of calcareous grasslands. Environ. Microbial. 2, 99–110.
Kowalchuk, G.A., Stienstra, A.W., Heilig, G.H., Stephen, J.R.
and Woldendrop, J.W. (2000) Molecular analysis of ammoniaoxidizing bacteria in soil of successional grasslands of Drentsche
A (The Netherlands). FEMS Microbiol. Ecol. 31, 207–215.
Degrange, V. and Bardin, R. (1995) Detection and counting of
Nitrobacter populations in soil by PCR. Appl. Environ. Microbiol. 61, 2093–2098.
Mobarry, B.K, Wagner, M., Urbain, V., Rittman, B.E. and Stahl,
D.A. (1996) Phylogenetic probes for analyzing abundance and
spatial organization of nitrifying bacteria. Appl. Environ. Microbiol. 62, 2156–2162.
Hovanec, T.A., Taylor, L.T., Blakis, A. and Delong, E.F. (1998)
Nitrospira-like bacteria associated with nitrite oxidation in
freshwater aquaria. Appl. Environ. Microbiol. 64, 258–264.
Feray, C., Volat, B., Degrange, V., Clays-Josserand, A. and
Montuelle, B. (1999) Assessment of three methods for detection
and quantification of nitrite-oxidizing bacteria and Nitrobacter in
freshwater sediments (MPN-PCR, MPN-Griess, Immunofluorescence). Microb. Ecol. 37, 208–217.
Skujins, J. and Fulgham, T.Y. (1978) Nitrification in great basin
desert soils In: Nitrogen in Desert Ecosystems (West, N.E. and
Skujins, J.J., Eds.). Dowden, Hutchinson and Ross, Inc.,
Stroudsburg, PA.
Schaeffer, S.M., Billings, S.A. and Evans, R.D. (2003) Responses
of soil nitrogen dynamics in a Mojave desert ecosystem to
manipulations in soil carbon and nitrogen availability. Oecologia
134, 547–553.
Zaady, E., Groffman, P.M. and Shachak, M. (1996) Litter as a
regulator of N and C dynamics in macrophytic patches in Negev
desert soils. Soil Biol. Biochem. 28, 39–46.
Kieft, T.L., Soroker, E. and firestone, M.K. (1987) Microbial
biomass response to rapid increase in water potential when dry
soil is wetted. Soil Biol. Biochem. 19, 119–126.
Peterjohn, W.T. (1991) Denitrification: enzyme content and
activity in desert soils. Soil Biol. Biochem. 23, 845–855.
Shachak, M. and Boeken, B. (1994) Changes in desert plant
communities in human-made patches and their implications for
management of desertified landscapes. Ecol. Appl. 4, 702–716.
Phillips, C.J., Harris, D., Dollhopf, S.L., Gross, K.L., Prosser, J.I.
and Paul, E.A. (2000) Effect of agronomic treatments on structure
and function of ammonia-oxidizing communities. Appl. Environ.
Microbiol. 66, 5410–5418.
29
[34] Oved, T., Shaviv, A., Goldrath, T., Mandelbaum, R.T. and Minz,
D. (2001) Influence of effluent irrigation on community structure
composition and function of ammonia-oxidizing bacteria in soil.
Appl. Environ. Microbiol. 67, 3426–3433.
[35] Shem-Tov, S., Zaady, E., Groffman, P.M. and Gutterman, Y.
(1999) Soil carbon content along a rainfall gradient and inhibition
of germination: a potential mechanism for regulating distribution
of Plantago coronopus. Soil Biol. Biochem., 31,1209–1217.
[36] Clesceri, L.S., Greenberg, A.E. and Trussell, R.R., Eds., (1989).
Standard Methods for the Examination of Water and Wastewater
(APHA). Port City Press, Baltimore, MD, USA.
[37] Gross, A., Boyd, E. and Seo, J. (1999) Evaluation of the
ultraviolet spectrophotometric method for the measurment of
total nitrogen in water. J. World Aquacult. Soc. 30, 388–
393.
[38] Schinner, F., Ohlinger, R., Kandeler, E. and Margesin, R., Eds.,
(1995). Methods in Soil Biology. Springer, Berlin, Heidelberg.
[39] Aakra, A., Utaker, J.B., Nes, I.F. and Bakken, L.R. (1999) An
evaluated improvement of the extinction dilution method for
isolation of ammonia-oxidizing bacteria. J. Microb. Meth. 39, 23–
31.
[40] Kowalchuk, G.E., Stephen, J.R., De Boer, W., Prosser, J.I.,
Embley, T.M. and Woldendorp, J.W. (1997) Analysis of ammonia-oxidizing bacteria of the b subdivision of the class proteobacteria in coastal sand dunes by denaturing gradient gel
electrophoresis and sequencing of PCR-amplified16S ribosomal
DNA fragment. Appl. Environ. Microbiol. 63, 1489–1497.
[41] Altschul, S.F., Madden, T.L., Schaffer, A.A., Zhang, J., Zhang,
Z., Miller, W. and Lipman, D.L. (1997) Gapped BLAST and PSIPLAST: a new generation of protein database search programs.
Nucleic Acid Res. 25, 3389–3402.
[42] Flowers, T.J., Troke, P.F. and Yeo, A.R. (1977) The mechanism
of salt tolerance in halophytes. Ann. Rev. Plant Physiol. 28, 89–
121.
[43] Zaady, E., Groffman, P.M. and Shachak, M. (1996) Release and
consumption of nitrogen by snail feces in Negev Desert soils. Biol.
Fert. Soils 23, 399–404.
[44] Hunik, J.H., Meijer, H.J.G. and Tramper, J. (1992) Kinetics
of Nitrosomonas europaea at extreme substrate, product and
salt concentrations. Appl. Microbiol. Biotechnol. 37, 802–
807.
[45] Hunik, J.H., Meijer, H.J.G. and Tramper, J. (1993) Kinetics of
Nitrobacter agilis at extreme substrate, product and salt concentrations. Appl. Microbiol. Biotechnol. 40, 442–448.
[46] Prinčič, A., Mahne, I., Megušar, F., Paul, E.A. and Tiedje, J.M.
(1998) Effects of pH and oxygen and ammonium concentrations
on the community structure of nitrifying bacteria from wastewater. Appl. Environ. Microbiol. 64, 3584–3590.
[47] Gross, A., Nemirovsky, A., Zilberg, D., Khaimov, A., Brenner,
A., Snir, E., Ronen, Z. and Nejidat, A. (2003) Soil nitrifying
enrichments as biofilter starters in intensive recirculating saline
water aquaculture. Aquaculture 223, 51–62.