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Applied and Environmental Microbiology, October 2001, p. 4742-4751, Vol. 67, No. 10
Federal Biological Research Centre for
Agriculture and Forestry, D-38104 Braunschweig,1
and Department of Biological Sciences, Microbiology,
University of Rostock, D-18051 Rostock,2 Germany
Received 20 April 2001/Accepted 6 August 2001
The bacterial rhizosphere communities of three host plants of the
pathogenic fungus Verticillium dahliae, field-grown
strawberry (Fragaria ananassa Duch.), oilseed rape
(Brassica napus L.), and potato (Solanum
tuberosum L.), were analyzed. We aimed to determine the degree to
which the rhizosphere effect is plant dependent and whether this effect
would be increased by growing the same crops in two consecutive years.
Rhizosphere or soil samples were taken five times over the vegetation
periods. To allow a cultivation-independent analysis, total community
DNA was extracted from the microbial pellet recovered from root or soil
samples. 16S rDNA fragments amplified by PCR from soil or rhizosphere
bacterium DNA were analyzed by denaturing gradient gel electrophoresis
(DGGE). The DGGE fingerprints showed plant-dependent shifts in the
relative abundance of bacterial populations in the rhizosphere which
became more pronounced in the second year. DGGE patterns of oilseed
rape and potato rhizosphere communities were more similar to each other
than to the strawberry patterns. In both years seasonal shifts in the
abundance and composition of the bacterial rhizosphere populations were
observed. Independent of the plant species, the patterns of the first
sampling times for both years were characterized by the absence of some
of the bands which became dominant at the following sampling times.
Bacillus megaterium and Arthrobacter sp.
were found as predominant populations in bulk soils. Sequencing of
dominant bands excised from the rhizosphere patterns revealed that 6 out of 10 bands resembled gram-positive bacteria. Nocardia
populations were identified as strawberry-specific bands.
In the future, biological control of
soil-borne fungal or bacterial pathogens will be of increasing
importance for a more sustainable agriculture. Furthermore, fungicides
such as methyl bromides will be phased out, and thus potential
alternatives are needed to control the soil-borne pathogen
Verticillium dahliae Kleb. This has prompted the search for
reliable antagonists which show a high degree of competitiveness and
are active in the rhizospheres of different crops and in different soil
types (3, 4, 39). However, to fully exploit the potentials
of biological control agents, a better understanding of the structural
and functional diversity of microbial populations in the rhizosphere
and their succession during plant development is required
(49). The rhizosphere, defined as the volume of soil
adjacent to and influenced by the plant root (45), is of
great importance to plant health and soil fertility. Root exudates
stimulate the growth of bacterial and fungal populations in the
vicinity of the roots (40). Several studies have indicated
that the structural and functional diversity of rhizosphere populations
is affected by the plant species due to differences in root exudation
and rhizodeposition in different root zones (21, 45).
Furthermore, the soil type, growth stage, cropping practices (such as
tillage and crop rotation), and other environmental factors (8,
14, 20, 22, 27, 52) seem to influence the composition of the
microbial community in the rhizosphere. Rhizosphere microorganisms
exert strong effects on plant growth and health by nutrient
solubilization, N2 fixation, or the production of plant
hormones (19, 36). Increased plant productivity also
results from the suppression of deleterious microorganisms by
antagonistic bacteria, while soil-borne pathogens can greatly reduce
plant growth.
Most studies of the bacterial community structure of rhizospheres
indicating a plant-dependent diversity were performed using cultivation-based techniques (12, 23, 25, 28, 30). The major problem of cultivation-based analysis is that only a small proportion of the bacterial populations can be recovered from the
rhizosphere and soil by traditional cultivation techniques (1,
46). Cultivation-based limitations can be overcome by analyzing
DNA that is directly extracted from rhizosphere and soil samples.
Recently the analysis of 16S rDNA fragments that were PCR amplified
from community DNA was used to unravel bacterial rhizosphere
communities. Molecular fingerprinting techniques, such as denaturing or
temperature gradient gel electrophoresis (16, 31), allow
analysis of large numbers of samples, which is essential for studying
spatial and temporal variations of bacterial rhizosphere populations.
To provide baseline data for biological control of the pathogenic
fungus Verticillium dahliae, the bacterial communities of the rhizospheres of three typical Verticillium host plants
were studied: strawberry (Fragaria ananassa Duch. [family:
Rosaceae]), oilseed rape (Brassica napus L. [family:
Brassicaceae]), and potato (Solanum tuberosum L. [family:
Solanaceae]). We aimed to find out the degree to which the rhizosphere
effect is plant dependent and whether this effect would be increased by
planting the same crops in two consecutive years. Rhizosphere or soil
samples were taken five times over the vegetation periods. To allow a
cultivation-independent analysis, total community DNA was extracted
from the microbial pellets recovered from root or soil samples.
Fragments of the 16S ribosomal DNA (rDNA) were amplified by PCR with
eubacterial primers from soil or rhizosphere DNA, and the PCR products
obtained were analyzed by denaturing gradient gel electrophoresis
(DGGE). Prominent bands were excised and used for sequence
determination in order to obtain further information about the
phylogeny of the dominating or plant-specific bacterial populations.
Field design and sampling.
The field test was performed on
fields belonging to the Federal Biological Research Centre for
Agriculture and Forestry (BBA) in Braunschweig, Germany. The soil type
was classified as loamy sand (pH 7) with an organic matter content of
0.9% and a clay content of 12% (data kindly provided by the Institute
for Weed Research, BBA). Three different plant species
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.10.4742-4751.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Bulk and Rhizosphere Soil Bacterial Communities
Studied by Denaturing Gradient Gel Electrophoresis: Plant-Dependent
Enrichment and Seasonal Shifts Revealed
and
ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
strawberry
(F. ananassa Duch.) cv. Elsanta, oilseed rape (B. napus L.) cv. Licosmos, and potato (S. tuberosum L.)
cv. Elementwere grown with six replicates per plant type and six
unplanted plots (each 3 by 3 m) arranged according to a randomized
Latin square (Fig. 1). Potatoes and strawberries were planted, and oilseed rape (surface sterilization done
only in the first year) was sown on the same field plots for 2 consecutive years. Samples were taken from each of the plots 1, 2, 3, 4, and 5 months after sowing (labeled 1.1 to 1.5 for the first year and
2.1 to 2.5 for the second). One composite soil sample consisting of
five cores (15 cm of topsoil) was taken per fallow plot, and each was
mixed by sieving. One composite rhizosphere sample taken per plot
consisted of roots of 5 randomly selected strawberry or 5 potato plants
or 20 oilseed rape plants, respectively. The roots were shaken
vigorously to separate soil not tightly adhering to the roots. Six
composite samples of each treatment were obtained per sampling time. A
total of 240 composite samples were taken for two consecutive seasons.
View larger version (20K):
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FIG. 1.
Experimental field design. See Materials and Methods for
details.
Extraction of bacterial cells from soil or roots.
Bacterial
cell extraction prior to community DNA extraction was done according to
the recommendations given by Bakken and Lindahl (2).
Briefly, 3 g of soil or plant roots with firmly adhering soil was
resuspended in 9 ml of distilled water and treated in a Stomacher 400 blender (Seward) for 1 min at high speed. After centrifugation at low
speed (2 min, 500 × g), the supernatant was collected
and the resulting pellet was resuspended in 9 ml of distilled water
followed by Stomacher blending and low-speed centrifugation. This step
was repeated once. The supernatants of the three centrifugation steps
were combined before centrifugation at high speed (10,000 × g) for 30 min to collect the microbial pellet. The resulting
pellet was kept at 
70°C.
DNA extraction and PCR amplification of 16S rDNA fragments for DGGE analysis. Total community DNA was extracted from the cell pellet according to the protocol by Smalla and van Elsas (44). The crude DNA was purified by using CsCl and sodium acetate precipitation steps, followed by a purification according to the manufacturer's protocol (GENECLEAN Spin Kit; BIO 101, La Jolla, Calif.).
The 16S rDNA fragments (positions 968 to 1401 [Escherichia coli rDNA sequence]) were amplified by PCR from rhizosphere or soil DNA extracts with the primer pair F984GC and R1378 (17, 18). Amplification was done using the advantage GC-genomic polymerase mixture (25 µl) as described by the manufacturers (Clontech, Palo Alto, Calif.) with 1 M GC-melt and 100 nM of each primer. The template DNA amount was approximately 1 to 5 ng per PCR. Acetamide (50%; 5 µl) was added to the reaction mixture to facilitate the denaturation of double-stranded DNA and to circumvent the formation of secondary structures. After 5 min of denaturation at 94°C and 35 thermal cycles of 1 min at 95°C, 1 min at 53°C, and 2 min at 72°C, PCR was finished by an extension step at 72°C for 10 min. Products were checked by electrophoresis in 1% (wt/vol) agarose gels and ethidium bromide staining (41). A mixture of the DGGE-PCR products from 11 bacterial species was applied two to three times to each DGGE gel as a marker to check the electrophoresis run and to compare fragment migration between gels as described by Heuer et al. (17). These species were (in the order of the migration distance): Clostridium pasteurianum DSM 525, Erwinia carotovora DSM 30168, Agrobacterium tumefaciens DSM 30205, Pseudomonas fluorescens R2f, Pantoea agglomerans, Nocardia asteroides N3, Rhizobium leguminosarum DSM 30132, Actinomadura viridis DSM 43462, Kineosporia aurantiaca JCM 3230, Nocardiopsis atra ATCC 31511, and Actinoplanes philippiensis JCM 3001.DGGE. DGGE analysis was essentially done as described by Heuer et al. (18) with a denaturing gradient of 40 to 58% of the denaturant. Aliquots of PCR samples (4 to 7 µl) were applied on the denaturing gradient gel, and DGGE was performed with 0.5× Tris-acetate-EDTA buffer at 60°C at a constant voltage of 180 V for 4 h. To compare the patterns of all different treatments on one denaturing gradient gel, only PCR products amplified from four replicates per treatment (each representing one composite sample) were loaded on the gel. After silver staining of the gels according to Heuer et al. (18), the gels were air dried and scanned transmissively (pdi 420oe scanner; MWG biotech, Ebersberg, Germany). The GelCompar 4.0 program (Applied Maths, Ghent, Belgium) was used to analyze the bacterial community fingerprints of each denaturing gradient gel as described by Rademaker et al. (37) with a slight modification of some normalization settings. The track resolution was increased to 2,000 pt, curve smoothing was set to 9 pt, and background subtraction was applied using the rolling disk method with an intensity of 8. After normalizing the gel image and background subtraction, the Pearson correlation index (r) for each pair of lanes within a gel was calculated as a measure of similarity between the community fingerprints, and the clustering of patterns was calculated using the unweighted-pair group method using average linkages (UPGMA). According to Rademaker et al. (37), the Pearson product moment correlation coefficient is better suited for identification of fingerprints than band-matching algorithms. The Pearson correlation coefficient is directly applied to the array of densitometric values forming the fingerprint. The coefficient is robust and objective, since whole curves are compared and subjective band scoring is omitted. Moreover, large collections of complex fingerprints can be compared readily using the product moment, since the band-based alternative is more laborious and time consuming. The Pearson correlation coefficient is largely insensitive to relative concentrations of bands between fingerprints, and it is insensitive to differences in the overall intensities of profiles.
For comparison between rhizosphere and soil samples or between rhizosphere samples from different plants, respectively, the similarity of the DGGE profiles within a treatment (natural variability) and between treatments was compared. If the median r values of two treatments (e.g., soil and rhizosphere samples at a certain time) differed more than expected from natural variability (if there is no overlap of interquartile ranges), then a relevant effect was assumed. The approach chosen allowed us to directly compare only lanes (treatments) within one gel and still in one figure to present the comparison for up to 10 different gels.Cloning and sequencing of DGGE bands. Dominant bands were excised from DGGE gels which contained N,N'-bis-acryl(yl)cystamine instead of N,N'-methylenebisacrylamide (32). Staining was performed with SYBR Green 1 (FMC, Vallensbaek Strand, Denmark), and band excision was done as described by Muyzer et al. (32) with an incubation step of 100 µl of 2-mercaptoethanol for 70 min at 37°C to dissolve the gel. Five microliters of the resulting solution was used in a PCR to reamplify the excised 16S rDNA fragment using the PCR conditions described by Heuer et al. (18), using the same primer pair and temperature program described above, with an additional step of 2 h at 72°C after adding fresh dATP (2 mM) and 2 U of AmpliTaq DNA polymerase (Stoffel fragment; Perkin-Elmer) to support the formation of a 3' A overhang for improved cloning efficiency (24). Cloning of the PCR products was necessary because DGGE analysis revealed weak bands in addition to the excised bands after reamplification. After confirming the enrichment of the excised band by DGGE, the PCR product was ligated into the pGEM-T vector (Promega, Madison, Wisc.) and transformed into competent cells (E. coli JM109; Promega) as described by the manufacturers. Plasmids with the correct insert, as determined by DGGE, were selected, and inserts were sequenced with the standard primers SP6 and T7 (IIT GmbH; Bielefeld, Germany). Data analysis was done with ARB software (Department of Microbiology, Technical University of Munich, Munich, Germany [http://www.arb-home.de]).
Nucleotide sequence accession numbers.
Nucleotide sequence
accession numbers of the partial 16S rDNA sequences determined in this
study are given in Table 1.
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RESULTS |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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High-molecular-weight DNA was recovered from all rhizosphere and bulk soil samples. DGGE analysis of 16S rDNA fragments from soil or rhizosphere DNA revealed high similarity of the DGGE patterns obtained from each of the six replicates per treatment and time point, suggesting a low degree of variability caused by the sampling, DNA extraction, PCR amplification, and DGGE analysis.
Rhizosphere effect.
The degree to which bacterial populations
are enriched in the rhizospheres of plants compared to the surrounding
bulk soil, indicating shifts of relative abundance, was analyzed. For
the analysis, 16S rDNA fragments amplified from DNA extracted from the
rhizospheres of strawberries, oilseed rape, potatoes, or bulk soil at
each sampling time were compared, running them in parallel on one
denaturing gradient gel. Since sequencing of prominent bands was done
for samples from sampling time 2.3, the DGGE patterns of this sampling
time are shown as a representative gel picture in Fig.
2. At all sampling times, the bulk soil
patterns consisted of one or two stronger bands and a large number of
less intense bands, indicating that in bulk soil samples the 16S rDNA
fragments of only one or two populations dominated, while many
populations which were less prevalent seemed to be equally abundant. In
contrast, the rhizosphere pattern consisted of several strong bands and a lower number of weak bands. Obviously the relative abundance of
several bacterial populations was enhanced in the surroundings of the
roots, suggesting a rhizosphere effect. To quantify this effect, the
Pearson correlation r value for each pair of lanes within a
gel was calculated as a measure of similarity between the community
fingerprints. For statistical comparison between rhizosphere and soil
samples, the similarity of the DGGE profiles within a treatment (Fig.
3) and between treatments (Fig. 3) was compared. Differences between the DGGE patterns of the rhizosphere and
bulk soil samples could be detected at all sampling times. Only at
sampling time 1.1 did all profiles show a relatively high level of
similarity, indicating that at that time the bacterial community in the
rhizosphere of the different crops and that of the bulk soil were still
rather similar. The patterns from the bulk soil and the rhizosphere
became different for all following sampling times. However, compared
with the first year (1.1 to 1.5), the similarity between the
rhizosphere DGGE patterns and those of the bulk soil clearly decreased
in the second year (2.1 to 2.5).
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Plant-dependent diversity.
In order to compare the DGGE
patterns for strawberry, oilseed rape, and potato at all sampling times
in a more quantitative way, the Pearson indices were determined and the
similarities of the DGGE profiles within a treatment and between the
rhizospheres of two plants were compared (Fig.
4). All DGGE patterns showed a relatively
high level of similarity to each other for samples taken 1 month after
planting in the first year. The levels of similarity between the
profiles of strawberry and oilseed rape were clearly different from
each other for all following sampling times except for samples taken at
1.5 (Fig. 4a). When comparing the similarities of DGGE profiles between
strawberry and potato rhizosphere, a slightly different picture was
found (Fig. 4b). While DGGE patterns for both plants were different at
all sampling times in the second year (2.1 to 2.5), the patterns were
more similar to each other in the first year. Only at sampling times 1.2 and 1.3 did median r values of the two treatments differ
from natural variability more than expected, indicating a relevant effect. Visual inspection of DGGE patterns for all sampling times already indicated that the patterns of oilseed rape and potato rhizosphere communities were more similar to each other than to the
strawberry patterns. This could be confirmed (Fig. 4c), since the
median r-value differences between the two treatments
(potato and oilseed rape) were above variability within each treatment at one sampling time in the first year (1.4) and at two sampling times
in the second year (2.3 and 2.4).
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Sequence analysis.
Prominent bands were excised and sequenced
to get further information about the dominating bacterial populations
in the different treatments from the DGGE gel of sampling time 2.3 (Fig. 2). We failed to reamplify bands from silver-stained gels, and
therefore SYBR Green staining was used. Reamplification products and
clones obtained were screened by DGGE analysis of the respective 16S rDNA fragments. Parallel analysis of 16S rDNA fragments amplified from
clones or from the community DNA on the same gel allowed us to
carefully check whether the cloned 16S rDNA fragments comigrated with
the band of the corresponding community pattern. The results of the
partial sequence analysis of these bands and their tentative phylogenetic affiliation are shown in Table 1. The majority of bands
were excised from the strawberry DGGE patterns, since these patterns
showed the strongest shifts in the relative abundance of dominant
populations. The sequence of the strong bulk soil band 1b showed 100%
similarity to Bacillus megaterium as well as to a dominant
band in the rhizosphere pattern of potato (band 1p), which had the same
migration behaviour as band 1b. This indicates that B. megaterium seems to be a dominant population not only in the bulk
soil but also in the rhizosphere of potato and presumably in the
rhizosphere of strawberry and oilseed rape, where bands with identical
electrophoretic mobility were also detected. The sequence of the other
dominant band in the bulk soil pattern (band 2b) could be assigned to
Arthrobacter sp. with a 97.7% similarity, but with a 100%
similarity to a clone obtained from the potato rhizosphere analyzed in
another project in our laboratory. Although a band with electrophoretic
mobility similar to that of band 2b of the bulk soil pattern was
excised from the potato rhizosphere DGGE patterns and cloned (band 3p),
the sequence obtained showed similarity to that of a
-proteobacterium (clone sequence from paddy soil
[15]). A comparison of the DGGE patterns revealed several bands which were detected in the rhizospheres of all plant species but not in the bulk soil, while a few dominant bands of the
rhizosphere patterns were characteristic for one plant species only. As
plant-specific bands, which dominated only in the rhizospheres of
strawberry plants, sequences related to high-G+C bacteria, such as
Nocardia sp. (band 4s) and Streptomyces galbus
(band 2s), could be identified. Five out of six dominating bands in the
rhizospheres of strawberry plants were assigned to the high-G+C
actinomycetes. Band 2p, which showed a migration behavior similar to
that of band 1s and dominated in the rhizosphere pattern of
potato plants, was related to 
-proteobacteria
(Devosia riboflavina) at 95.2% similarity and
at 98% similarity to clone sequences obtained from the potato
rhizosphere in a different project. Two bands (5s and 1r) observed at
rather high denaturing concentrations in the DGGE rhizosphere patterns
of strawberry and oilseed rape, but not of potatoes, fell in the
high-G+C cluster but showed a rather low similarity to bands of
Promicromonospora citrea. The sequence of 5s had a 97.2%
similarity to the sequence of 1r from oilseed rape. Band 6s, excised
from the rhizosphere pattern of strawberry, also showed a high
similarity to those sequences, but it showed different migration characteristics.
Seasonal shifts.
To study the seasonal shifts in the abundance
and composition of the bacterial rhizosphere populations, we analyzed
the rhizosphere DGGE patterns for all sampling times (1.1 to 1.5 and
2.1 to 2.5) for strawberry, potato, and oilseed rape (Fig. 5). Although
many bands were detected at all sampling times, shifts in the bacterial community could be detected. In all treatments, the strongest shift
occurred at the beginning of the vegetation period. Independently of
the plant species, the patterns of the first sampling time in both
years were characterized by the absence of some of those bands which
became dominant at the following sampling times. In both years and for
all three plants the DGGE patterns of the first sampling time (1.1 and
2.1) formed a separate cluster when UPGMA was used to create a
dendrogram describing the similarities between the DGGE patterns. In
all treatments, some bands with varying intensities could be detected
over time. In the DGGE patterns obtained for strawberry rhizosphere
bacteria in the second year (Fig. 5a),
the relative abundance of several populations seems to be enhanced at
the second sampling time (2.2) compared to the first (2.1). Some very
intense bands in the DGGE patterns of 2.2 were much less intense at
2.1, while other bands, such as 4s, 5s, and 6s, were not detected at
all at 2.1. In the rhizospheres of strawberry plants, bands 5s and 6s
were not observed in the first year. The dominant bands were detected
in most of the replicates of sampling times 2.2 to 2.5. Figure 5b shows
the seasonal shifts in the rhizospheres of potatoes observed in the
second year, and bands showing seasonal shifts in the relative
abundance of the bacterial population are indicated. Several
bands appeared only from sampling time 2.2 and remained dominant
populations in the rhizosphere of potato until the end of the season.
The enrichment of bacterial populations was most pronounced at sampling
times 2.2 and 2.3. Several bands which according to their melting
behavior might belong to high-G+C gram-positive bacteria were detected only at the first three sampling times but not at 2.4 and 2.5. Seasonal
shifts in the relative abundances of respective 16S rDNA targets were
observed for oilseed rape as well (Fig. 5c). As observed also for
strawberry (Fig. 5a) and potato (Fig. 5b), the enrichment of bacterial
populations seemed to be most pronounced for sampling times 2.2 and 2.3 when oilseed rape was flowering. Interestingly, band 1r, which shows a
high similarity to a sequence obtained from strawberry (band 5s) and a
lower similarity to P. citrea, became numerically dominant
only at times 2.3 and 2.4.
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DISCUSSION |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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DGGE fingerprints of PCR-amplified 16S rDNA genes were used to study dominant bacterial populations in the rhizospheres of the three V. dahliae Kleb. host plants, strawberry, potato, and oilseed rape, over two growing seasons. In contrast to other recently published papers, in which DGGE fingerprints have been used to analyze bacterial rhizosphere communities (9, 10, 35, 53), the rhizosphere samples investigated in this study originated from plants grown under field conditions in six replicated plots per treatment. We detected a rhizosphere effect, namely an increased relative abundance of some populations in the vicinity of the roots for all three plants. Several bands observed in rhizosphere DGGE patterns were not detected in patterns of soil from unplanted plots or were detected only as weak bands. The rhizosphere effect became more pronounced in the second year. In contrast, Duineveld et al. (9) observed no differences or only minor differences between bulk soil and the rhizospheres of chrysanthemum plants grown in pots in a growth room. No differences between bulk and rhizosphere patterns were found for barley grown in pots in a growth chamber (35). Due to the cell extraction technique applied in our study, the DNA obtained originated from rhizosphere and rhizoplane bacteria. Although several bands occurred in the DGGE patterns of all plants, a plant-dependent diversity of the bacterial patterns could be shown for most of the sampling times. The differences between the DGGE patterns of the three different plants became more pronounced in the second year, in particular when the similarities of strawberry rhizosphere patterns were compared with potato or oilseed rape. The finding that the patterns of oilseed rape and potato rhizosphere communities were more similar to each other than to the strawberry patterns might be explained by the fact that oilseed rape and potato are annual plants while strawberry is a perennial plant. Our data provide further evidence for the assumption that different plant species select different bacterial communities in the proximity of their roots and that these plant-specific enrichments can be increased by repeated cultivation of the plant species in the same field. Recently, Schwieger and Tebbe (42) reported differences in the single-strand conformation polymorphism fingerprints of 16S rDNA fragments amplified from rhizosphere DNA of Chenopodium album and Medicago sativa grown under field conditions. We observed seasonal shifts of a similar trend in the rhizospheres of strawberry, oilseed rape, and potato plants for 2 years. The most pronounced differences were noticed between the patterns of the first and second samplings of both years. The patterns of rhizosphere samples taken when the plants were flowering (1.2 and 1.3; 2.2 and 2.3) showed the strongest enrichment of some bacterial populations. Lottmann et al. (26) also observed the appearance of additional dominant bands in the DGGE rhizosphere patterns of potatoes at the time of flowering. The variability between replicates was highest for samples taken at the end of both growing seasons. The seasonal shifts observed in our study were less dramatic than those reported for bacterial communities in the rhizosphere of maize grown in tropical soil (13). In contrast, experiments performed in controlled pot experiments showed no changes in relation to the age of the barley plants (35), and only minor shifts in the rhizosphere of chrysanthemum (9) were found. However, both studies followed potential temporal shifts in the composition of bacterial rhizosphere communities for a much shorter period of time (up to 36 days after sowing and up to 10 weeks after planting, respectively) under growth chamber conditions, one factor which might have contributed to the contrasting results. Semenov et al. (43) found a moderate rhizosphere effect in one experiment with soil rich in fresh plant debris and a very pronounced rhizosphere effect in a second experiment with soil low in organic matter content using cultivation techniques (by enumeration of copiotrophic and oligotrophic bacteria).
Sequencing of DGGE bands revealed an astonishingly high proportion of dominant populations in the rhizosphere belonging to high-G+C gram-positive bacteria. Thus, five out of six sequences obtained from dominant bands in the rhizospheres of strawberry plants belonged to different high-G+C gram-positive bacteria, such as Nocardia and Streptomyces. One dominant band which was only detected in the rhizosphere of strawberry showed a 100% similarity to those of four different Nocardia species in the database. Interestingly, the sequences of the two dominant bands from bulk soil and four bands from the rhizosphere shared a similarity of more than 97% with those of cultured isolates. The seasonal shifts that were followed for strawberry, potato, and oilseed rape indicated that the abundance of high-G+C populations was different during the developmental stages of these plants. However, conclusions regarding their activity can hardly be drawn, since our analysis was based on 16S rDNA fragments amplified from DNA.
There is increasing evidence that gram-positive bacteria might be more
dominant in the rhizosphere than previously supposed. Several recently
published papers support this notion. Bacillus species were
found as dominant populations in the rhizospheres of chrysanthemum
(9), of barley (35), and of grass
(11). McCaig et al. (29) reported that in a
clone library obtained from grass rhizospheres,
Actinomycetes spp. were the second-most-abundant group after
the most frequently found -proteobacteria. Arthrobacter spp. were also found as dominant populations in the molecular fingerprints of 16S rDNA fragments amplified from the rhizosphere DNA
of maize grown in tropical soil (13), from the rhizosphere DNA of M. sativa and C. album (42),
and from the rhizosphere DNA of chrysanthemum (9). One
dominant DGGE band obtained at all locations of the barley rhizospheres
grown in controlled pot experiments performed by Yang and Crowley
(53) was identified as Microbacterium. However,
in none of the recently published cultivation-independent studies of
the bacterial diversity of the rhizosphere was such a high proportion
of dominant populations detected belonging to a diverse range of
high-G+C gram-positive bacteria.
In this study DGGE analysis of 16S rDNA fragments amplified from community DNA was used for the molecular analysis of a large number of rhizosphere and bulk soil samples taken over two growing seasons. The DGGE profiles mainly reflect the evenness of populations in an environmental sample, and in this study they indicated that a reduced evenness was found in the rhizosphere compared to soil. Interpretation of DGGE patterns needs to be done cautiously as discussed in several reviews (16, 33). Amplified 16S rDNA fragments of different but phylogenetically related species might have the same electrophoretic mobility because they share the identical or similar sequence in the stretch analyzed, as found for the Nocardia sequence from the strawberry-specific band in this study or for Arthrobacter clones (13) and isolates (5). But phylogenetically nonrelated species also by coincidence might have a similar melting behavior, as observed in this study for bands 1s and 2p as well as 2b and 3p. However, since only one clone per band was sequenced, we cannot exclude the possibility that both sequences 1s and 2p and sequences 2b and 3p are present in the same band. Thus, a rather large diversity of populations sharing the identical 16S rDNA sequence might be hidden behind a DGGE band. Despite several pitfalls of PCR-based rRNA analysis (50), profiling of bacterial rhizosphere and bulk communities by denaturing gradient gels proved to be a powerful method allowing a cultivation-independent analysis of large numbers of rhizosphere and bulk soil samples. Currently we are applying group-specific primers (13, 17) to amplify 16S rDNA fragments of different phylogenetic groups, which should enable a better level of resolution.
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ACKNOWLEDGMENTS |
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This study was funded by a DFG grant to K.S. and G.B. The DGGE approach used had already been established in BMBF research project 0311295.
We are grateful to S. Kropf, Leipzig University, for discussing statistical data treatment with us.
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FOOTNOTES |
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* Corresponding author. Mailing address: Federal Biological Research Centre for Agriculture and Forestry, Messeweg 11-12, D-38104 Braunschweig, Germany. Phone: 49-531-2993814. Fax: 49-531-2993013. E-mail: K.Smalla{at}bba.de.
Present address: Lower Saxony State Agency for Ecology, 31135 Hildesheim, Germany.
Present address: GBF, Abt. Mikrobiologie, D-38124 Braunschweig, Germany.
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REFERENCES |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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| 1. |
Amann, R. I.,
W. Ludwig, and K.-H. Schleifer.
1995.
Phylogenetic identification and in situ detection of individual microbial cells without cultivation.
Microbiol. Rev.
59:143-169 |
| 2. | Bakken, L. R., and V. Lindahl. 1995. Recovery of bacterial cells from soil, p. 9-27. In J. T. Trevors, and J. D. van Elsas (ed.), Nucleic acids in the environment. Springer-Verlag, Berlin, Germany. |
| 3. | Berg, G., S. Kurze, A. Buchner, E. M. H. Wellington, and K. Smalla. 2000. Successful strategy for the selection of new strawberry associated rhizobacteria antagonistic to Verticillium wilt. Can. J. Microbiol. 46:1128-1137[CrossRef][Medline]. |
| 4. | Berg, G., A. Fritze, N. Roskot, and K. Smalla. Evaluation of potential biocontrol rhizobacteria from different host plants of Verticillium dahliae Kleb. J. Appl. Microbiol., in press. |
| 5. | Brim, H., H. Heuer, E. Krögerrecklenfort, M. Mergeay, and K. Smalla. 1999. Characterization of the bacterial community of a zinc-polluted soil. Can. J. Microbiol. 45:326-338[CrossRef][Medline]. |
| 6. |
Chun, J., and M. Goodfellow.
1995.
A phylogenetic analysis of the genus Nocardia with 16S rRNA gene sequences.
Int. J. Syst. Bacteriol.
45:240-245 |
| 7. |
Crocker, F. H.,
J. K. Fredrickson,
D. C. White,
D. B. Ringelberg, and D. L. Balkwill.
2000.
Phylogenetic and physiological diversity of Arthrobacter strains isolated from unconsolidated subsurface sediments.
Microbiology
146:1295-1310 |
| 8. | De Leij, F. A. A. M., J. M. Whipps, and J. M. Lynch. 1994. The use of colony development for the characterization of bacterial communities in soil and on roots. Microb. Ecol. 27:81-97. |
| 9. |
Duineveld, B. M.,
G. A. Kowalchuk,
A. Keijzer,
J. D. van Elsas, and J. A. van Veen.
2001.
Analysis of bacterial communities in the rhizosphere of chrysanthemum via denaturing gradient gel electrophoresis of PCR-amplified 16S rRNA as well as DNA fragments coding for 16S rRNA.
Appl. Environ. Microbiol.
67:172-178 |
| 10. |
Duineveld, B. M.,
A. S. Rosado,
J. D. van Elsas, and J. A. van Veen.
1998.
Analysis of the dynamics of bacterial communities in the rhizosphere of the chrysanthemum via denaturing gradient gel electrophoresis and substrate utilization patterns.
Appl. Environ. Microbiol.
64:4950-4957 |
| 11. |
Felske, A.,
A. Wolterink,
R. van Lis, and A. D. L. Akkermans.
1998.
Phylogeny of the main bacterial 16S rRNA sequences in Drentse A grassland soils (The Netherlands).
Appl. Environ. Microbiol.
64:871-879 |
| 12. | Germida, J. J., S. D. Siciliano, J. Renato de Freitas, and A. M. Seib. 1998. Diversity of root-associated bacteria associated with field-grown canola (Brassica napus L.) and wheat (Triticum aestivum L.). FEMS Microbiol. Ecol. 26:43-50. |
| 13. | Gomes, N. C. M., H. Heuer, J. Schönfeld, R. Costa, L. Hagler-Mendonca, and K. Smalla. 2001. Bacterial diversity of the rhizosphere of maize (Zea mays) grown in tropical soil studied by temperature gradient gel electrophoresis. Plant Soil 232:167-180[CrossRef]. |
| 14. | Grayston, S. J., S. Wang, C. D. Campbell, and A. C. Edwards. 1998. Selective influence of plant species on microbial diversity in the rhizosphere. Soil Biol. Biochem. 30:369-378[CrossRef]. |
| 15. |
Hengstmann, U.,
K. J. Chin,
P. H. Janssen, and W. Liesack.
1999.
Comparative phylogenetic assignment of environmental sequences of genes encoding 16S rRNA and numerically abundant culturable bacteria from an anoxic rice paddy soil.
Appl. Environ. Microbiol.
65:5050-5058 |
| 16. | Heuer, H., and K. Smalla. 1997. Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) for studying soil microbial communities, p. 353-373. In J. D. Van Elsas, J. T. Trevors, and E. M. H. Wellington (ed.), Modern soil microbiology. Marcel Dekker, New York, N.Y. |
| 17. | Heuer, H., M. Krsek, P. Baker, K. Smalla, and E. M. H. Wellington. 1997. Analysis of actinomycete communities by specific amplification of genes encoding 16S rRNA and gel-electrophoretic separation in denaturing gradients. Appl. Environ. Microbiol. 63:3233-3241[Abstract]. |
| 18. |
Heuer, H.,
K. Hartung,
G. Wieland,
I. Kramer, and K. Smalla.
1999.
Polynucleotide probes that target a hypervariable region of 16S rRNA genes to identify bacterial isolates corresponding to bands of community fingerprints.
Appl. Environ. Microbiol.
65:1045-1049 |
| 19. | Höflich, G., W. Wiehe, and G. Kühn. 1994. Plant growth stimulation by inoculation with symbiotic and associative rhizosphere microorganisms. Experientia 50:897-905[CrossRef]. |
| 20. | Horwath, W. R., L. F. Elliott, and J. M. Lynch. 1998. Influence of soil quality on the function of inhibitory rhizobacteria. Lett. Appl. Microbiol. 26:87-92. |
| 21. |
Jaeger, C. H., III,
S. E. Lindow,
W. Miller,
E. Clark, and M. K. Firestone.
1999.
Mapping of sugar and amino acid availability in soil around roots with bacterial sensors of sucrose and tryptophan.
Appl. Environ. Microbiol.
65:2685-2690 |
| 22. | Latour, X., L. Philippot, T. Corberand, and P. Lemanceau. 1996. The establishment of an introduced community of fluorescent pseudomonads in the soil and in the rhizosphere is affected by the soil type. FEMS Microbiol. Ecol. 30:163-170. |
| 23. | Lemanceau, P., T. Corberand, L. Gardan, X. Latour, G. Laguerre, J. M. Boeufgras, and C. Alabouvette. 1995. Effect of two plant species, flax (Linum usitatissinum L.) and tomato (Lycopersicon esculentum Mill.), on the diversity of soilborne populations of fluorescent pseudomonads. Appl. Environ. Microbiol. 61:1004-1012[Abstract]. |
| 24. | Li, Q.-B., and C. L. Guy. 1996. Prolonged final extension time increases cloning efficiency of PCR products. BioTechniques 21:192-196[Medline]. |
| 25. | Liljeroth, E., S. L. G. E. Burgers, and J. A. van Veen. 1991. Changes in bacterial populations along roots of wheat seedings. Biol. Fertil. Soils 10:276-280[CrossRef]. |
| 26. | Lottmann, J., H. Heuer, J. de Vries, A. Mahn, K. Düring, W. Wackernagel, K. Smalla, and G. Berg. 2000. Establishment of introduced antagonistic bacteria in the rhizosphere of transgenic potatoes and their effect on the bacterial community. FEMS Microbiol. Ecol. 33:41-49[CrossRef][Medline]. |
| 27. | Lupwayi, N. Z., W. A. Rice, and G. W. Clayton. 1998. Soil microbial diversity and community structure under wheat as influenced by tillage and crop rotation. Soil Biol. Biochem. 30:1733-1741[CrossRef]. |
| 28. | Mahaffee, W. F., and J. W. Kloepper. 1997. Temporal changes in the bacterial communities of soil, rhizosphere, and endorhiza associated with field-grown cucumber (Cucumis sativus L.). Microb. Ecol. 34:210-223[CrossRef][Medline]. |
| 29. |
McCaig, A. E.,
L. A. Glover, and J. I. Prosser.
1999.
Molecular analysis of bacterial community structure and diversity in unimproved and improved upland grass pastures.
Appl. Environ. Microbiol.
65:1721-1730 |
| 30. | Miller, H. J., G. Henken, and J. A. van Veen. 1989. Variation and composition of bacterial populations in the rhizospheres of maize, wheat, and grass cultivars. Can. J. Microbiol. 35:656-660. |
| 31. |
Muyzer, G.,
E. C. de Waal, and A. G. Uitterlinden.
1993.
Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes encoding for 16S rRNA.
Appl. Environ. Microbiol.
59:695-700 |
| 32. | Muyzer, G., T. Brinkhoff, U. Nübel, C. Santegoeds, H. Schäfer, and C. Wawer. 1998. Denaturing gradient gel electrophoresis (DGGE) in microbial ecology, 3.4.4, p. 1-27. In A. D. L. Akkermans, J. D. van Elsas, and F. J. de Bruijn (ed.), Molecular microbial ecology manual. Kluwer Academic Publishers, Dordrecht, The Netherlands. |
| 33. | Muyzer, G., and K. Smalla. 1998. Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology. Antonie Leeuwenhoek 73:127-141[CrossRef][Medline]. |
| 34. |
Nakagawa, Y.,
T. Sakane, and A. Yokota.
1996.
Transfer of "Pseudomonas riboflavina" (Foster 1944), a gram-negative, motile rod with long-chain 3-hydroxy fatty acids, to Devosia riboflavina gen. nov., sp. nov., nom. rev.
Int. J. Syst. Bacteriol.
46:16-22 |
| 35. |
Normander, B., and J. I. Prosser.
2000.
Bacterial origin and community composition in the barley phytosphere as a function of habitat and presowing conditions.
Appl. Environ. Microbiol.
66:4372-4377 |
| 36. | Patten, C. L., and B. R. Glick. 1996. Bacterial biosynthesis of indole-3-acetic acid. Can. J. Microbiol. 42:207-220[Medline]. |
| 37. | Rademaker, J. L. W., F. J. Louws, U. Rossbach, P. Vinuesa, and F. J. de Bruijn. 1998. Computer-assisted pattern analysis of molecular fingerprints and database construction, 7.1.3. In A. D. L. Akkermans, J. D. van Elsas, and F. J. de Bruijn (ed.), Molecular microbial ecology manual. Kluwer Academic Publishers, Dordrecht, The Netherlands. |
| 38. |
Rainey, F. A.,
N. Weiss, and E. Stackebrandt.
1995.
Phylogenetic analysis of the genera Cellulomonas, Promicromonospora, and Jonesia and proposal to exclude the genus Jonesia from the family Cellulomonadaceae.
Int. J. Syst. Bacteriol.
45:649-652 |
| 39. |
Ross, I. L.,
Y. Alami,
P. R. Harvey,
W. Achouak, and M. H. Ryder.
2000.
Genetic diversity and biological control activity of novel species of closely related pseudomonads isolated from the wheat fields in South Australia.
Appl. Environ. Microbiol.
66:1609-1616 |
| 40. | Rovira, A. D. 1965. Interactions between plant roots and soil micro-organisms. Annu. Rev. Microbiol. 19:241-266[CrossRef][Medline]. |
| 41. | Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. |
| 42. |
Schwieger, F., and C. C. Tebbe.
2000.
Effect of field inoculation with Sinorhizobium meliloti L33 on the composition of bacterial communities in rhizospheres of a target plant (Medicago sativa) and a non-target plant (Chenopodium album)linking of 16S rRNA gene-based single-strand conformation polymorphism community profiles to the diversity of cultivated bacteria.
Appl. Environ. Microbiol.
66:3556-3565 |
| 43. | Semenov, A. M., A. H. C. van Bruggen, and V. V. Zelenev. 1999. Moving waves of bacterial populations and total organic carbon along roots of wheat. Microbiol. Ecol. 37:116-128[CrossRef][Medline]. |
| 44. | Smalla, K., and J. D. van Elsas. 1995. Application of the PCR for detection of antibiotic resistance genes in environmental samples, p. 241-256. In J. T. Trevors, and J. D. van Elsas (ed.), Nucleic acids in the environment. Springer-Verlag, Berlin, Germany. |
| 45. | Sørensen, J. 1997. The rhizosphere as a habitat for soil microorganisms, p. 21-45. In J. D. Van Elsas, J. T. Trevors, and E. M. H. Wellington (ed.), Modern soil microbiology. Marcel Dekker, Inc, New York, N.Y. |
| 46. | Staley, J. T., and A. Konopka. 1985. Measurement of in situ activities of nonphotosynthetic microorganisms in aquatic and terrestrial habitats. Annu. Rev. Microbiol. 39:321-346[CrossRef][Medline]. |
| 47. | Suzuki, T., and K. Yamasato. 1994. Phylogeny of spore-forming lactic acid bacteria based on 16S rRNA gene sequences. FEMS Microbiol. Lett. 115:13-17[CrossRef][Medline]. |
| 48. |
Takeuchi, T.,
H. Sawada,
F. Tanaka, and I. Matsuda.
1996.
Phylogenetic analysis of Streptomyces spp. causing potato scab based on 16S rRNA sequences.
Int. J. Syst. Bacteriol.
46:476-479 |
| 49. | Van Veen, J. A., L. S. van Overbeek, and J. D. van Elsas. 1997. Fate and activity of microorganisms introduced into soil. Microbiol. Mol. Biol. Rev. 61:121-135[Abstract]. |
| 50. | von Wintzingerode, F., U. B. Göbel, and E. Stackebrandt. 1997. Determination of microbial diversity in environmental samples: pitfalls of PCR-based rRNA analysis. FEMS Microbiol. Rev. 21:213-229[CrossRef][Medline]. |
| 51. | Wang, G. C., and Y. Wang. 1995. Rapid differentiation of bacterial species with multiple probes of different lengths in a single slot blot hybridization. Appl. Environ. Microbiol. 61:4269-4273[Abstract]. |
| 52. | Westover, K. M., A. C. Kennedy, and S. E. Kelley. 1997. Patterns of rhizosphere microbial community structure associated with co-occurring plant species. J. Ecol. 85:863-873[CrossRef]. |
| 53. |
Yang, C. H., and D. E. Crowley.
2000.
Rhizosphere microbial community structure in relation to root location and plant iron nutritional status.
Appl. Environ. Microbiol.
66:345-351 |
Applied and Environmental Microbiology, October 2001, p. 4742-4751, Vol. 67, No. 10
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.10.4742-4751.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
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Johansson, P. M., Wright, S. A. I.
(2003). Low-Temperature Isolation of Disease-Suppressive Bacteria and Characterization of a Distinctive Group of Pseudomonads. Appl. Environ. Microbiol.
69: 6464-6474
[Abstract] [Full Text] -
Gomes, N. C. M., Fagbola, O., Costa, R., Rumjanek, N. G., Buchner, A., Mendona-Hagler, L., Smalla, K.
(2003). Dynamics of Fungal Communities in Bulk and Maize Rhizosphere Soil in the Tropics. Appl. Environ. Microbiol.
69: 3758-3766
[Abstract] [Full Text] -
Stamper, D. M., Walch, M., Jacobs, R. N.
(2003). Bacterial Population Changes in a Membrane Bioreactor for Graywater Treatment Monitored by Denaturing Gradient Gel Electrophoretic Analysis of 16S rRNA Gene Fragments. Appl. Environ. Microbiol.
69: 852-860
[Abstract] [Full Text] -
Philippot, L., Piutti, S., Martin-Laurent, F., Hallet, S., Germon, J. C.
(2002). Molecular Analysis of the Nitrate-Reducing Community from Unplanted and Maize-Planted Soils. Appl. Environ. Microbiol.
68: 6121-6128
[Abstract] [Full Text] -
van Dillewijn, P., Villadas, P. J., Toro, N.
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68: 4201-4208
[Abstract] [Full Text] -
Berg, G., Roskot, N., Steidle, A., Eberl, L., Zock, A., Smalla, K.
(2002). Plant-Dependent Genotypic and Phenotypic Diversity of Antagonistic Rhizobacteria Isolated from Different Verticillium Host Plants. Appl. Environ. Microbiol.
68: 3328-3338
[Abstract] [Full Text] -
Krimi, Z., Petit, A., Mougel, C., Dessaux, Y., Nesme, X.
(2002). Seasonal Fluctuations and Long-Term Persistence of Pathogenic Populations of Agrobacterium spp. in Soils. Appl. Environ. Microbiol.
68: 3358-3365
[Abstract] [Full Text] -
Tauch, A., Schneiker, S., Selbitschka, W., Puhler, A., van Overbeek, L. S., Smalla, K., Thomas, C. M., Bailey, M. J., Forney, L. J., Weightman, A., Ceglowski, P., Pembroke, T., Tietze, E., Schroder, G., Lanka, E., van Elsas, J. D.
(2002). The complete nucleotide sequence and environmental distribution of the cryptic, conjugative, broad-host-range plasmid pIPO2 isolated from bacteria of the wheat rhizosphere. Microbiology
148: 1637-1653
[Abstract] [Full Text] -
Reiter, B., Pfeifer, U., Schwab, H., Sessitsch, A.
(2002). Response of Endophytic Bacterial Communities in Potato Plants to Infection with Erwinia carotovora subsp. atroseptica. Appl. Environ. Microbiol.
68: 2261-2268
[Abstract] [Full Text] -
Salles, J. F., De Souza, F. A., van Elsas, J. D.
(2002). Molecular Method To Assess the Diversity of Burkholderia Species in Environmental Samples. Appl. Environ. Microbiol.
68: 1595-1603
[Abstract] [Full Text] -
Heuer, H., Kroppenstedt, R. M., Lottmann, J., Berg, G., Smalla, K.
(2002). Effects of T4 Lysozyme Release from Transgenic Potato Roots on Bacterial Rhizosphere Communities Are Negligible Relative to Natural Factors. Appl. Environ. Microbiol.
68: 1325-1335
[Abstract] [Full Text]
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