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Applied and Environmental Microbiology, December 1998, p. 4950-4957, Vol. 64, No. 12
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Analysis of the Dynamics of Bacterial Communities in the
Rhizosphere of the Chrysanthemum via Denaturing Gradient Gel
Electrophoresis and Substrate Utilization Patterns
Bernadette M.
Duineveld,1,*
Alexandre S.
Rosado,2
Jan Dirk
van Elsas,2 and
Johannes A.
van
Veen1
Institute of Evolutionary and Ecological
Sciences, University of Leiden, 2300 RA Leiden,1
and
DLO-Research Institute for Plant Protection, 6700 GW
Wageningen,2 The Netherlands
Received 24 February 1998/Accepted 23 July 1998
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ABSTRACT |
In order to gain a better understanding of the spatial and temporal
dynamics of bacterial communities of the rhizosphere of the
chrysanthemum, two complementary methods were used: a molecular bacterial community profiling method, i.e., 16S rRNA gene-based PCR
followed by denaturing gradient gel electrophoresis (DGGE), and an agar
plate method in which 11 sole-carbon-source utilization tests were
used. The DGGE patterns showed that the bacterial communities as
determined from direct rhizosphere DNA extracts were largely stable
along developing roots of the chrysanthemum, with very little change
over time or between root parts of different ages. The patterns were
also similar to those produced with DNA extracts obtained from bulk
soil samples. The DGGE patterns obtained by using microbial colonies
from dilution plates as the source of target DNA were different from
those found with the direct DNA extracts. Moreover, these patterns
showed differences among plant replicates but also among replicate
plates. Results obtained with the sole-carbon-source utilization tests
indicated that the metabolic profile of the bacterial communities in
the rhizosphere of the root tip did not change substantially during
plant growth. This suggests selective development of specific bacterial
populations by the presence of a root tip. On the other hand, the
metabolic profile of bacterial communities in the rhizosphere of the
root base changed during plant growth. With eight sole-carbon-source utilization tests, a significant effect of the development stage of the
plant on the number of bacteria which were able to grow on these carbon
sources was observed.
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INTRODUCTION |
The chrysanthemum, an economically
important ornamental plant, is frequently plagued by Pythium
spp., which cause it to decline (32). The oomycete
Pythium is an avid colonizer of young roots (16),
which for the chrysanthemum can result in root rot and a decreased
quality of the plant. The occurrence of an antagonistic microflora
might, however, inhibit the infection of roots by Pythium. As the levels of establishment and survival of introduced antagonistic bacteria are often low (30), the best possibilities for the biological control of root pathogens such as Pythium might
lie in the use or stimulation of indigenous bacteria from the soil in
which the plant is to be grown. Antagonistic organisms that occur in
the same ecological space and time frame, i.e., competing for the
same exudates that play a role in the attraction of an infection
by Pythium, may be the most attractive candidates. To find such antagonists, we need to understand the dynamics of the structure of the bacterial communities in the rhizosphere of the chrysanthemum.
The rhizosphere is the volume of soil adjacent to and influenced by the
plant root (9). Roots are known to excrete several forms of
organic materials. The amounts and composition of these organic
materials are different in different plant species and cultivars,
change during plant development, and are different in old and young
parts of the root system (1, 7, 14). As a result, the
bacterial communities in the rhizosphere, which can use these organic
materials as a substrate, will differ in composition and density
(1, 2). This may result in the buildup of a microflora
specific to a particular plant species and genotype (20,
21), as well as to the plant developmental stage and the root
part (base or tip) (12). The study of the diversity of
bacterial communities in the soil or rhizosphere is inherently difficult, since all methods developed to date have limitations (10, 26, 29). Traditional plating techniques commonly result in assessment of the diversity of less than 10% of the total bacterial community present (23, 29). On the other hand,
microscopic techniques (including those using antisera or
oligonucleotide probes) result in an assessment of total
bacterial numbers, but very little can be said about the
microbial diversity of the sample, due to the lack of a wide range of
probes or antibodies.
Recently, a novel method, PCR followed by denaturing gradient gel
electrophoresis (PCR-DGGE), was proposed for the study of the
phylogenetic diversity of bacterial populations in environmental samples (18). In this method, total microbial DNA is
extracted from soil, after which the bacterial 16S rRNA genes are
amplified by PCR with universal eubacterial primers (8). The
PCR products of the same length but with different internal sequences
can be separated according to their melting properties by DGGE or
temperature gradient gel electrophoresis (TGGE) (8, 18). The
patterns obtained provide information about the underlying bacterial
populations. This molecular method does not have the limitation of
cultivation-based methods, and hence it can assess the diversity
of the bacterial groups, including the nonculturable bacterial groups.
Hence, we have used PCR-DGGE in conjunction with cultivation-based
methods to analyze the diversity of the bacterial community in the
rhizosphere during the growth of chrysanthemum plants. In addition to
information on the total bacterial community structure on a genetic
basis, it is important to gather data on the physiological potential of
bacterial communities. The sole-carbon-source utilization tests of the
culturable populations used here are suitable tools for this purpose,
as they provide a functional measure of metabolic potential.
Information on both the dynamics of the total bacterial community and
the metabolic potential of populations is important for the
identification of suitable biocontrol agents. In this study we aimed to
obtain a better understanding of root effects on the total and the
culturable bacterial community in soil on a general level without
dealing with specific groups of bacteria.
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MATERIALS AND METHODS |
Soil.
Ede loamy sand soil was used throughout. This soil has
been described in detail by van Elsas et al. (27). Briefly,
it is a "beekeerd" soil, slightly acid (pH-KCl 5.5), with about
3.5% organic matter. During the growth of the plants, the soil
moisture content was kept at about 12% (wt/wt). All soil was sieved
prior to use (mesh length, 6 mm; mesh width, 3 mm). To enhance
aeration, the soil was mixed with perlite (20% [vol/vol]). After
mixing, the soil was put into in 1-liter pots (bulk density, about 1.0 g/cm3).
Plant growth conditions.
The chrysanthemum cultivar used in
this study, `Majoor Bosshardt,' was obtained from Fides Inc. (De
Lier, The Netherlands). Cuttings of this cultivar were dipped in the
insecticide Savona (a mixture of natural fatty acids; Koppert,
Berkel-Rodenrijs, The Netherlands), dipped in hormone powder
(3-indolylbutyric acid) to stimulate root growth, and thereafter placed
in 4-cm3 peat blocks. The cuttings were allowed to develop
roots in a growth room (at 20°C day and night, 70% relative
humidity, and a 16/8 h [light/dark] photoperiod).
After 2 weeks, each peat block containing a plantlet was placed in a
pot on top of the Ede loamy sand soil, in such a way that the peat
block was still visible. The plants were watered once a day. Twice a
week, the plants received 75 ml of a nutrient solution (Pokon; 15:20:25
[NPK]; electrical conductivity [EC] = 2 mS/cm); once a week a
trace element mixture (containing iron, manganese, boron, zinc, copper,
and molybdenum ions) was added to the nutrient solution. Three weeks
after the peat blocks with plantlets were placed in the pots, the day
length was changed from 16 to 8 h in order to induce flowering of
the chrysanthemum plants.
Sampling.
Samples of chrysanthemum root parts were collected
2, 4, 6, 8, and 10 weeks after planting. Five plants were harvested at each sampling. The samples collected from each plant were kept separate. Soil and plants were removed from the pots, and subsequently each plant was shaken carefully to remove loose (nonadhering) soil. The
soil adhering to the roots was defined as rhizosphere soil. In a
preliminary experiment, the root development of chrysanthemums was
monitored in Perspex root observation boxes (31) in order to
establish the length of the young root parts used in this study. Young
parts of the root system, collectively called tips, about 1 to 2 days
old, were collected by taking the end 1 cm of the root parts by using a
double-cutting knife with a fixed cut space of 1 cm. Old parts of the
root system, designated base, were collected by taking the first 4 cm
of the roots in the soil just behind the peat block by using a
double-cutting knife with a cut space of 4 cm. All lateral roots that
had developed on parts of the base were cut away as close as possible
to the main root. The root samples with adhering soil were weighed.
Samples of two plants at each sampling date were stored separately at
20°C. These samples were extracted to obtain microbial-community
DNA, which was subsequently analyzed by PCR-DGGE. The samples of the
other three plants were treated as described under "Community carbon
utilization analyses."
In addition, samples of Ede loamy sand bulk soil were collected and
stored at 20°C in order to assess the putative rhizosphere effects
by comparing the patterns obtained from bulk and rhizosphere soils.
Three types of Ede loamy sand bulk soil samples were collected, i.e.,
bulk soil directly from the field mixed with perlite, and bulk soil
with or without perlite from pots which had been kept for 14 days under
the same conditions as the plants used. This period coincided with the
first sampling moment of the plants.
Collection of cultured bacterial communities for DGGE
analysis.
Three 6-week-old plants, grown as described above, were
used to obtain root tip and root base samples. Pooled 1-cm stretches of
each root part (tip versus base) were placed in Eppendorf tubes containing 1 ml of 0.1% sodium pyrophosphate (NaPP) plus gravel. The
samples were shaken on a Vortex mixer (15 min, full speed) and
subsequently diluted 100-fold. Fifty microliters of each sample was
plated on 0.1 strength tryptic soy agar (0.1 TSA) containing cycloheximide (100 µg/ml). After one week of incubation at 20°C, colonies were removed by adding 1.5 ml TE (10 mM Tris-HCl-1 mM EDTA
[pH 8.0]) buffer on each plate and scraping off all growth with a
Drigalski spatula. The cell suspensions thus obtained were subjected to
DNA extraction and PCR-DGGE analysis.
Extraction of microbial DNA.
DNA extraction and purification
from the bulk soil and rhizosphere samples was performed by the method
of Smalla et al. (25) as modified by van Elsas and Smalla
(28) for bulk soil and protocol C of Leung et al.
(11) for rhizosphere soil. Purification of the crude
extracts was by CsCl and potassium acetate precipitation steps followed
by Wizard (Madison, Wis.) spin column treatment. For extraction of DNA
from the samples of the agar plates, the 1.5-ml suspensions were
centrifuged 5 min at maximum speed (Eppendorf centrifuge). The pellet
was resuspended in TE buffer, and glass beads 1 mm in diameter (±100
mg) were added; this procedure was followed by bead beating for 30 s, after which 30 µl of 10% sodium dodecyl sulfate was added. DNA
was extracted by sequential phenol, phenol-chloroform-isoamyl alcohol
(25:24:1), and chloroform-isoamyl alcohol (24:1) extractions. The DNA
was then precipitated by using 0.1 volume 5 M NaCl and 0.6 volume
isopropanol for 5 min at room temperature. After centrifuging at
maximum speed (Eppendorf centrifuge) and washing of the pellet with
70% ethanol, the pellet was resuspended in 50 µl of TE buffer. This
crude extract was finally purified with the Wizard spin column kit.
PCR amplification.
PCR amplifications were performed with
the (clamped) F968 and R1401 universal eubacterial primers described by
Heuer and Smalla (8), with various dilutions of soil-,
rhizosphere-, and plate community-derived mixed templates. The PCR used
40 thermal cycles in a touchdown scheme as described by Rosado et al.
(22). PCR products of approximately 450 bp were analyzed on
conventional agarose gels (24) prior to further analysis on
denaturing gradients. Strong bands of the expected size (450 bp) were
subjected to DGGE analysis.
DGGE analysis.
Denaturing gradients of various degrees of
steepness were prepared in accordance with the methods of Muyzer et al.
(18) and Myers et al. (19). Gradients between 50 and 70% of denaturing agents (urea-formamide) commonly produced
optimal separation of PCR-generated bands and were routinely used.
Samples of 20 µl of PCR product were loaded on gels, which were then
run for 6 or 16 h in an Ingeny (Leiden, The Netherlands) DGGE
setup at 60°C (100 V). After the runs, gels were removed from the
setup and stained for 60 min with SYBR green I nucleic acid gel stain
(Molecular Probes, Leiden, The Netherlands), after which they were
inspected under UV light and photographed. The banding patterns were
analyzed by the 1/0 clustering method of the NT-SYS program (Exeter
Software, New York, N.Y.) by using the unweighted pair group with
mathematical averages (UPGMA). The DGGE banding patterns are considered
to be representative of the dominant bacterial groups (estimated as
0.1 to 1% of the total community) in accordance with the work of
Heuer and Smalla (8) and Muyzer et al. (18).
Community carbon utilization analyses.
Root samples of three
plants, used for analyzing the metabolic potential of the culturable
bacterial populations on agar plates, were placed in Erlenmeyer flasks
containing 95 ml of 0.1% sterile NaPP and 10 g of gravel. The
Erlenmeyer flasks were shaken on a rotary shaker for 20 min at 200 rpm.
Tenfold serial dilutions of the suspensions were made with 0.1% NaPP.
One hundred microliters of the 10
2 and 103
dilutions were plated, in triplicate, on 0.1 TSA for total bacterial counts (CFU). Simmons citrate agar was used as the basic medium (6) for carrying out the carbon utilization tests. The
carbon source citrate (1 g/liter) was replaced with starch, sucrose, maltose, fructose, glucose, fucose, sodium oxalate, sodium succinate, glutamine, serine, or phenylalanine. Medium without a carbon source was
used as a control. On agar plates containing any one of the carbon
sources, 100 µl of the 102 sample dilution was spread.
The plates contained cycloheximide (100 µg/ml) to inhibit the growth
of fungi. After 14 days of incubation at 20°C, colonies with
diameters of 2 mm were enumerated. The total counts on 0.1 TSA were
expressed as CFU per unit of root surface. The root surface was
determined by measuring the diameter and length of the root parts, by
considering the root to be a cylinder.
Statistics.
One-way analysis of variance (ANOVA) was used to
analyze the effect of sampling date on the total CFU counted on 0.1 TSA
plates for samples collected from the root tip and root base. The
numbers of CFU were log transformed before calculations were made. The sign test was used for comparison of the numbers of bacteria on the
root tip and base per plant, since these numbers are not independent per plant. The numbers of colonies with diameters of 2 mm counted on
agar plates with single carbon sources were divided by the numbers of
such colonies found for the same treatment on the 0.1 TSA plates. Per
root tip or base, the fractions of the culturable bacterial populations
that were able to utilize specific single carbon sources were compared
between sampling dates, by using one-way ANOVA.
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RESULTS |
Total microbial community DNA (PCR-DGGE) analyses.
PCR-amplifiable DNA was recovered from all chrysanthemum rhizosphere
samples, as well as from corresponding bulk soil. Repeated DGGE runs of
the same PCR product, as well as repeated PCR amplification of the same
DNA extract followed by DGGE, produced similar banding profiles,
suggesting that the approach was reproducible. In addition, the
variation between profiles obtained from replicates was small.
Samples collected from different root sites during plant development,
i.e., root tip versus root base, showed little variation
in banding
patterns when analyzed by PCR-DGGE (Fig.
1). In all
patterns, 30 to 36 bands of
various intensities were detected
per sample, with about 20 bands
shared among all samples. The
difference in band intensity was presumed
to indicate numerical
differences between the target
molecules (Fig.
1). Clustering
of the profiles revealed that all
profiles were about 82% similar,
with no clear trend with
respect to the clustering above this
level (Fig.
2). This similarity does not take the
intensities
of bands into account. The most obvious differences were
two conspicuous
bands evident in the last sampling, which were mostly
absent from
earlier samplings (Fig.
1, two bands around 62.5%
denaturant).
Moreover, one band present in all profiles generally
showed a
higher intensity in the root base samples than in those from
the
root tip (Fig.
1, band around 63.5% denaturant).
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FIG. 1.
DGGE patterns of 16S ribosomal DNA (rDNA) fragments from
rhizosphere samples of chrysanthemum plants collected at different
development stages and from different sites of the root system. Lanes
1, 3, 5, 7, and 9, root tips of 2-, 4-, 6-, 8-, and 10-week-old plants,
respectively; lanes 2, 4, 6, 8, and 10, root bases of 2-, 4-, 6-, 8-, and 10-week-old plants, respectively. Plants were grown in a loamy
sand. Percent values indicate the percentage of denaturants at each
position.
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FIG. 2.
Dendrogram representing genetic similarity of
microbial-community profiles obtained with PCR-DGGE. Samples were
collected at different moments of plant development and from different
root sites. 1, 3, 5, 7, and 9, root tips of 2-, 4-, 6-, 8-, and
10-week-old plants, respectively; 2, 4, 6, 8, and 10, root bases of 2-, 4-, 6-, 8-, and 10-week-old plants, respectively.
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To assess whether the rhizosphere, per se, induced shifts in the
dominant bacterial groups in soil, comparisons were made
between
patterns generated with community DNA from rhizosphere
versus bulk soil
samples (Fig.
3). In all patterns, 14 to
18 bands
were visible. It became clear that a large number of bands,
about
12, were shared among the different bulk soil and rhizosphere
profiles, and only a few differences were observed. Differences
were
mainly caused by a stronger intensity of bands in the rhizosphere
sample (Fig.
3, the bands around 56, 57, 58.5, and 60% denaturant).
This indicates a low-level impact from the plant root or the
root-excreted
material on the dominant groups in soil. Mixing Ede loamy
sand
with perlite or incubation of Ede loamy sand hardly influenced
the
profiles found (Fig.
3).
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FIG. 3.
DGGE patterns of 16S rDNA fragments from rhizosphere and
bulk soil (loamy sand). Lane 1, bulk soil mixed with perlite, not
incubated; lane 2, bulk soil without perlite incubated for 14 days;
lane 3, bulk soil mixed with perlite and incubated for 14 days under
the same conditions used for plant growth; lane 4, rhizosphere
soil of root tip samples of a 2-week-old chrysanthemum plant. M,
marker, composed of PCR products generated from the following
strains (from top to bottom): Enterobacter cloacae BE1,
Listeria innocua ALM105, Rhizobium
leguminosarum biovar trifolii R62,
Arthrobacter sp., and Burkholderia cepacia P2.
Percent values indicate the percentage of denaturants at each
position.
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In another experiment, all colonies of soil-derived bacteria grown on
agar plates were analyzed together by PCR-DGGE in order
to investigate
whether agar-grown colonies are representative
for the dominant
bacterial groups evidenced by direct PCR-DGGE.
The agar plates used in
this experiment contained about 100 bacterial
colonies of diverse
morphologies. In the DGGE profiles generated
with the mixed growth of
the agar plates, 9 to 21 bands were visible
for the different samples,
with only 3 bands shared among all
samples (Fig.
4). The profiles generated with DNA
obtained from
the colonies of agar plates showed several bands at
different
positions in samples from different plants, but also in
different
agar plate replicates. The profiles also showed about seven
bands
which were present in one sample. The overall variable results
obtained with DGGE-PCR from agar plates suggest that selection
of
bacterial colonies on agar plates can be different at every
occasion. This result was partly supported by UPGMA clustering,
since
replicates of agar plates were similar at levels between
78 and 90%,
with some exceptions that clustered at a lower level
(Fig.
5). The agar plate-derived profiles
clustered together at
a level of 68%.
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FIG. 4.
DGGE patterns of 16S rDNA fragments of microbial growth
on 0.1 TSA plates of rhizosphere samples and directly extracted
bacterial DNA of bulk soil samples. The profiles obtained from agar
plates are from three 6-week-old plants, i.e., plant 1 tip (lanes 1 and
2) and base (lanes 3 and 4), plant 2 tip (lanes 5 and 6) and base
(lanes 7 and 8), and plant 3 tip (lanes 9 and 10) and base (lanes 11 and 12). The bulk soil samples used were treated in three ways: mixed
with perlite and analyzed with no incubation time (lanes 13 and 14),
mixed with perlite and incubated for 14 days under the same conditions
used for plant growth (lanes 15 and 16), and without perlite, incubated
for 14 days (lanes 17 and 18). Lane M, marker composed of PCR products
generated from the following strains (from top to bottom): E. cloacae BE1, L. innocua ALM105, R. leguminosarum biovar trifolii R62,
Arthrobacter sp., and B. cepacia P2. Percent
values indicate the percentage of denaturants at each position.
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FIG. 5.
Dendrogram representing genetic similarity of
PCR-DGGE-obtained profiles of microbial growth on 0.1 TSA plates of
rhizosphere samples and directly extracted bacterial DNA of bulk soil
samples. The profiles obtained from agar plates are from three
6-week-old plants, i.e., plant 1 tip (1 and 2) and base (3 and 4),
plant 2 tip (5 and 6) and base (7 and 8), and plant 3 tip (9 and 10)
and base (11 and 12). The bulk soil samples used were treated in three
ways: mixed with perlite and analyzed with no incubation time (13 and
14), mixed with perlite and incubated for 14 days under the same
conditions used for plant growth (15 and 16), without perlite,
incubated for 14 days (17 and 18).
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The profiles generated with the direct soil DNA extracts revealed
15 to 19 bands, whereas 11 bands were internally consistent
(Fig.
4).
These numbers were in agreement with those obtained
earlier (Fig.
3).
Clustering of these profiles showed similarity
between replicates at a
level of 95 to 100% and a similarity of
80% for all direct soil DNA
extracts (Fig.
5).
Several differences could be observed when the profiles generated with
DNA from the agar plates were compared with those generated
from
directly extracted soil DNA. About 20 bands in the profiles
obtained
with DNA from all agar plates were not visible in the
profiles
generated with the directly extracted soil DNA (Fig.
4). In particular,
about nine strong bands, presumably of high
GC content, observed in the
agar plate profiles (Fig.
4) were
absent or weak in the profiles
generated with soil DNA. On the
other hand, about eight bands present
in the direct DNA extracts
were not present in the profiles based on
plate DNA. In the soil
DNA-derived profiles, bands that occurred at low
denaturant concentrations,
presumably of low GC content, were more
numerous than those in
the plate-generated profiles. This indicates
that these bacterial
groups are not culturable or are poorly culturable
under the conditions
used. In general, this means that bacteria which
are culturable
on 0.1 TSA plates are not representative for the
dominant groups
in the rhizosphere as detected by PCR-DGGE screening.
Clustering
of the profiles from the two methods resulted in a
similarity
of only 48% (Fig.
5).
Cultivation-based analyses.
The total number of bacteria
enumerated on 0.1 TSA plates decreased for both root tip and root base
samples during plant development (Table
1). However, this decrease was
significant only for root base samples (P < 0.01).
Moreover, a per-plant analysis showed that all plants contained
significantly higher numbers of bacteria at the root base than at the
root tip (sign test; P < 0.05).
In the carbon utilization analyses, a large number of bacteria were
able to grow on agar plates to which no carbon sources
had been added.
However, all colonies that developed on these
control plates remained
small (<1 mm in diameter). This limited
growth was probably due to
efficient scavenging of trace nutrient
sources from the agar or the
added suspension by these bacteria.
Hence, only large colonies
(diameter,
2 mm) were taken into account
in enumerating the colonies
on agar plates that contained single
carbon
sources.
The number of colonies from root tip samples which were able to grow on
any one of the carbon sources used, as a fraction
of the total CFU on
0.1 TSA, was similar throughout the entire
plant growth period, except
for those of bacteria that were able
to utilize serine and maltose
(Fig.
6). The relative numbers of
serine
utilizers were significantly enhanced in the last two samplings,
whereas maltose utilizers were significantly reduced in the last
sampling. Glucose and succinate metabolizers also showed large
differences between samplings; however, these differences were
not
significant.
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FIG. 6.
Fraction of bacteria isolated from the tips of
chrysanthemum roots which could use a single carbon source. The total
number of bacteria which could grow on 0.1 TSA was set to 1. Root tip
samples were collected from three chrysanthemum plants at different
developmental stages. Samples were tested on starch (A), sucrose
(B), maltose (C), fructose (D), glucose (E), fucose (F), oxalate (G),
succinate (H), serine (I), glutamine (J), and phenylalanine (K). Bars
marked with the same letters are not significantly different from each
other. (P < 0.05).
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The analysis of the root base populations resulted in numerous
significant differences in relative numbers of bacteria with
specific
carbon utilization capacities between samplings (Fig.
7). The dynamics in time of the specific
metabolic groups on the
root base were different for most carbon
sources used. A significant
increase in bacteria able to utilize starch
was found from the
first to the fourth sampling date. For fucose,
succinate, and
phenylalanine, a significant decrease in the number of
bacteria
able to utilize these carbon sources was observed. A
comparison
of the dynamics of all metabolic groups (defined by the
single
carbon sources used) between root tip and root base resulted in
largely divergent patterns; only glucose utilizers followed similar
patterns.
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FIG. 7.
Fraction of bacteria isolated from the base of
chrysanthemum roots which could use a single carbon source. The total
number of bacteria which could grow on 0.1 TSA was set to 1. Root base
samples were collected from three chrysanthemum plants at different
developmental stages. Samples were tested on starch (A), sucrose (B),
maltose (C), fructose (D), glucose (E), fucose (F), oxalate (G),
succinate (H), serine (I), glutamine (J), and phenylalanine (K). Bars
marked with the same letters are not significantly different
(P < 0.05).
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DISCUSSION |
The results obtained with PCR-DGGE performed on rhizosphere DNA
showed clear profiles that possibly represented the dominant bacterial
fractions in the samples. The picture of relative stability of the
structure of the total (culturable plus nonculturable) bacterial
communities contrasted with the picture of variability obtained in the
cultivation-based analyses. The DNA-based fingerprints indicated that
there are several dominant groups which are relatively stable in soil
and rhizosphere both in time and space. Similar stable patterns of
rhizosphere samples of transgenic and nontransgenic potatoes, as found
by Heuer and Smalla (8), confirm the results of this study.
Obviously, the potential impact of the root on bacterial populations in
soil was not such that major shifts in community structure were
induced, i.e., changes occurred at levels of maximally 0.1 to 1% of
the total bacterial biomass (18). It is likely that the
effect of roots on dominating soil bacterial groups is marginal,
as opposed to the effects of, e.g., soil type (29a). Thus,
each soil type may have its typical set of dominant groups, and
this mainly determines the DNA-based profiles of bacterial communities.
The above conclusions might be influenced by our definition of the
rhizosphere. There are several definitions of the rhizosphere (13). Our definition, the soil adhering to roots after loose soil is gently shaken off, is frequently used and is highly appropriate for biocontrol work. This definition implies that the amount of soil
adhering to roots during the development of the plant and roots may not
be constant. In earlier work we showed that for 10- and 12-week-old
plants, the amount of soil adhering to the collected root parts,
especially the root tip, was small or even not measurable compared to
larger amounts around roots of young plants. So, for root parts of
2-week-old plants, the bacteria present in large numbers in bulk soil
could have a dilution effect on the size of bacterial populations which
are stimulated by the root. In later plant development the dilution
effect is much smaller, and since no large shifts in the composition of
bacterial populations occur during the development of plants, the
conclusion that the influence of the root on dominant bacterial
populations in the rhizosphere may be small still holds.
The dominant bacterial groups analyzed might consist mainly of
the typical, oligotrophic soil bacteria. The stable patterns found can
be explained by the fact that these organisms have a slow
response to changes in the environment, including changes brought
about by root exudates. This does not imply that a rhizosphere effect
is unimportant. In this study we have analyzed the profiles by
considering bands present or not present, which resulted in the rather
stable pattern. However, bands can be present all the time but differ
in intensity. These differences in intensity probably indicate that
bacterial populations are changing in number during plant growth.
Beside this, certain groups, among which are gram-negative copiotrophic
organisms such as Pseudomonas spp., might be strongly stimulated in the rhizosphere. However, presumably these bacteria may
not be stimulated to such a level that they will become a quantitatively dominant population in the rhizosphere. Often, in the
literature, a typical rhizosphere organism is incorrectly considered to
be numerically dominant in the rhizosphere (2). However,
this consideration has generally been based on other methodological approaches. These less-dominant groups might
become apparent by PCR-DGGE only if another level of resolution
is used, for instance, by the use of other, more specific primers.
Our data showed that colonies grown on agar selected with soil
bacterial suspensions are not necessarily a reflection of dominant groups in the soil or rhizosphere. In other studies, it was also concluded that bacteria isolated by cultivation are not representative of the most dominant organisms in the environmental samples analyzed (8). The agar medium used here is considered nonselective
(17) and is widely accepted as a general medium for
isolation of diverse bacterial populations from natural systems such as
root, soil, and water samples. The results obtained have consequences
for the isolation and application of suitable bacteria for use as biological control agents. Antagonists which are to be introduced into
the soil to protect plants against root pathogens should, for practical
reasons, be culturable and occur in high numbers in the rhizosphere.
However, the present results indicate that culturable bacteria cannot
always readily be expected to belong to numerically dominant groups.
This may partly explain the difficulties and inconsistent results with
the use of inoculants (30).
The above-described results obtained by PCR-DGGE provide new and
interesting information. However, we should realize that the
method used here has its limitations (8). In the preparation of the samples, we have aimed for representative rhizosphere community DNA used as a template for PCR amplification. This means efficient dislodging of cells from soil particles or roots and complete lysis of
bacterial cells. As in all PCR-based approaches, selective amplification of genes from mixed communities by PCR may also bias the
analysis. With the interpretation of the profiles obtained by DGGE, it
should further be realized that one band may represent more than
one species. Despite these limitations, the comparative use of
PCR-DGGE as applied in this study justifies the
aforementioned conclusions.
In this study, the dynamics of bacterial communities in the rhizosphere
was also analyzed by plating on agar plates which contained
different single carbon sources. This method gave information about the metabolic properties of the culturable bacterial
communities. There was an obvious problem of growth of efficient
nutrient scavengers even on plates without an added carbon source. The
organic carbon scavenged might have originated from the agar used, from
the air, or from the rhizosphere sample plated. Garland and Mills
(5) already pointed to the interference of organic
material present in rhizosphere samples in subsequent
growth-based analyses.
The root tip samples collected were, in most cases, 1 to 2 days old.
So, if root exudates affect bacterial populations in the rhizosphere,
as is often suggested (2), it is expected that this
will occur in a rather consistent way as a response to the
consistent production of a specific quality of organic compounds.
However, colonization of the root tip might also be a random process;
thus, all or most bacteria that come in contact with the extending root
can settle on the tip, and a more fluctuating pattern between
samples can be expected. The differences in the relative number of tip
isolates capable of utilizing sole carbon sources were not
statistically significant (except for those found for maltose and
serine utilizers). This might indicate that there were no great
differences in the quantity and functional quality of the culturable
bacterial populations that develop on the root tip in a period of 1 to
2 days. Thus, root tip colonization might be a selective process,
stimulating only specific groups of the soil microflora. However, it
should be realized that the results obtained here showed large
variations, so this statement should be considered with caution.
With regard to the bacterial populations at the root base, a
balance between competing populations can be expected, which may result in a low degree of variation between plants of one harvest. Furthermore, the bacterial population of the root base may
consist of species which can utilize more complex carbon sources. These
organisms may grow more slowly than bacteria stimulated at the tip and
might be able to utilize recalcitrant substrates (3). The
number of bacteria collected from the root base that are able to
utilize complex carbon sources, i.e., starch and maltose, increased
during plant development, which is in agreement with our expectations.
Differences in findings for the same treatment were smaller for the
root base than for the root tip, and more significant differences were
found between treatments. So it seems that different root sites exert
different effects on the microbial populations, which agrees with
results found in other studies (12, 15).
The two methods provided complementary results. The carbon utilization
tests showed large differences in the metabolic properties of the
culturable bacteria. PCR-DGGE showed a stable community structure of
the total bacterial population. The stable pattern concerns the
dominant bands in the profile. The weak bands are more difficult to
analyze, since they interfere with the background. It is the weak bands
which are the main factor causing the deviation from 100% similarity.
So it seems that the main effect of the rhizosphere is exerted on the
groups which are represented by the weak bands. From the
sole-carbon-source utilization tests the variable population structure
indicated relations to the dynamics of plant root development. As
these culturable bacteria are generally considered to be a minor
fraction of the total bacterial community in soil, this matches the
changes in weak bands in the DGGE profiles.
We aimed, in this study, to obtain a comprehensive picture of the
dynamics of bacteria in the rhizosphere. Therefore, we did not go into
more detail so as to characterize the DGGE bands by sequencing or to
identify the different isolates. Of course, understanding the
significance of the few distinct differences between the DGGE-obtained patterns of bacterial communities in bulk soil and rhizosphere soil, as
well as on different places on the root during plant development, is
highly relevant. Therefore, we will continue this research by analyzing
the differences in more detail. It is also interesting to know if the
nonculturable dominant groups are actively metabolizing or growing in
the rhizosphere. Felske et al. (4) found a dominant
nonculturable group which may play a active role in a native soil
microbial community. It should be realized that it will be the active
groups which can eventually compete with Pythium for
nutrients and space and so can successfully be applied as biocontrol agents.
| |
ACKNOWLEDGMENTS |
We thank Anneke Keijzer and Ludwina Lankwarten for help in
running the DGGE profiles and Jesse Karthaus for help with the carbon
utilization tests.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Institute of
Evolutionary and Ecological Sciences, University of Leiden, P.O. Box
9516, 2300 RA Leiden, The Netherlands. Phone: 31 (0) 71 527 51 58. Fax: 31 (0) 71 527 49 00. E-mail:
duineveld{at}rulsfb.leidenuniv.nl.
| |
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Applied and Environmental Microbiology, December 1998, p. 4950-4957, Vol. 64, No. 12
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Top
Abstract
Introduction
