![[Feedback]](/icons/banner/feedbackACT.gif)
![[For Subscribers]](/icons/banner/subscriberACT.gif)
![[Archive]](/icons/banner/archiveACT.gif)
![[Search]](/icons/banner/searchACT.gif)
![[Contents]](/icons/banner/tocACT.gif)
Previous Article | Next Article 
Applied and Environmental Microbiology, March 2001, p. 1147-1153, Vol. 67, No. 3
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.3.1147-1153.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Succession of Indigenous Pseudomonas spp. and
Actinomycetes on Barley Roots Affected by the Antagonistic Strain
Pseudomonas fluorescens DR54 and the Fungicide
Imazalil
Laila
Thirup,1
Kaare
Johnsen,2,* and
Anne
Winding1
Department of Microbial Ecology and
Biotechnology, National Environmental Research Institute,
Roskilde,1 and Department of
Geochemistry, Geological Survey of Denmark and Greenland,
Copenhagen,2 Denmark
Received 3 July 2000/Accepted 21 December 2000
 |
ABSTRACT |
In recent years, the interest in the use of bacteria for biological
control of plant-pathogenic fungi has increased. We studied the
possible side effects of coating barley seeds with the antagonistic strain Pseudomonas fluorescens DR54 or a commercial
fungicide, imazalil. This was done by monitoring the number of
indigenous Pseudomonas organisms and actinomycetes on
barley roots during growth in soil, harvest after 50 days,
and subsequent decomposition. Bacteria were enumerated by
traditional plate spreading on Gould's S1 agar
(Pseudomonas) and as filamentous colonies on Winogradsky agar (actinomycetes) and by two quantitative competitive PCR assays. For this we developed an assay targeting Streptomyces and
closely related genera. DR54 constituted more than 75% of the
Pseudomonas population at the root base during the first 21 days but decreased to less than 10% at day 50. DR54 was not successful
in colonizing root tips. Initially, DR54 affected the number
of indigenous Pseudomonas organisms negatively,
whereas imazalil affected Pseudomonas numbers positively,
but the effects were transient. Although plate counts were considerably
lower than the number of DNA copies, the two methods correlated well
for Pseudomonas during plant growth, but after plant
harvest Pseudomonas-specific DNA copy numbers decreased while plate counts were in the same magnitude as before. Hence, Pseudomonas was 10-fold more culturable in a decomposition
environment than in the rhizosphere. The abundance of
actinomycetes was unaffected by DR54 or imazalil amendments, and CFU
and quantitative PCR results correlated throughout the experiment. The
abundance of actinomycetes increased gradually, mostly in numbers of
DNA copies, confirming their role in colonizing old roots.
| |
INTRODUCTION |
Public concern about chemical
pesticides has fostered an interest in application of bacteria for
biological control to protect agricultural crops against
pathogenic fungi (16). There is still a lack of
knowledge concerning environmental risks of such microbial inoculants. Introduced bacteria may outcompete a certain indigenous subpopulation for nutrients and space. For example, the biocontrol strain Pseudomonas fluorescens CHA0 (36)
displaced a part of the indigenous Pseudomonas population
for a short period after application, probably because of competition
for the same ecological niche in the rhizosphere (26).
Further, toxic compounds produced by introduced strains may affect
sensitive nontarget microorganisms.
P. fluorescens DR54 was isolated in Denmark from a sugar
beet rhizosphere and is effective towards preemergence damping-off disease caused by such different fungal pathogens as Pythium
ultimum and Rhizoctonia solani in laboratory pot
experiments (28). The antifungal active compound
viscosinamide has been isolated from DR54 and is believed to be the
main agent of the biocontrol properties of the strain (29,
39).
When studying possible side effects of seed coating with a biocontrol
strain, the effects of the biocontrol organism must be compared to the
effects of the normally applied fungicide. In Denmark, the seed coat
fungicide most used to protect winter and spring barley is the
commercial formulated fungicide Fungazil A. It contains the active
ingredient imazalil, which is a sterol biosynthesis inhibitor
(3), at a concentration of 50 g liter
1.
In Denmark, the recommended dose of Fungazil A is 1 ml kg of seed1.
In this study, we focused on possible side effects of P. fluorescens DR54 and imazalil on two important bacterial groups in soil: Pseudomonas and actinomycetes. The actinomycete group
is a very broad phylogenetic group of gram-positive bacteria with a
high GC content. Traditionally, actinomycetes have been defined as
bacteria with a filamentous, fungus-like growth form, but sequence analysis of 16S rRNA has shown that bacteria with more traditional growth forms, such as the coryneform bacteria and
Micrococcus, also belong to the group (6). In
temperate, well-drained soils with neutral to alkaline pH, the genus
Streptomyces is often the dominating actinomycete genus,
constituting around 95% of the filamentous actinomycetes as determined
by plate spreading (44). The gram-negative genus
Pseudomonas is now a well-defined genus, which formerly was
known as fluorescent pseudomonads or as Pseudomonas rRNA
homology group I (17).
Both groups are important in the degradation of organic matter,
although with different life strategies. Pseudomonas strains are mostly associated with fresh organic matter, rich in easily degradable compounds (14, 20, 34, 38), while actinomycetes traditionally are considered to be most active late in the
decomposition process, where they are strong competitors for complex
organic compounds (4, 19). Pseudomonas
typically has a considerably higher occurrence in the rhizosphere than
in bulk soil (25, 34), while actinomycetes have been found
to have both higher and lower occurrences in rhizosphere soil than in
bulk soil, depending on the plant species (5, 25).
The knowledge of the ecology of Pseudomonas and
actinomycetes, however, is based mainly on traditional cultivation
methods. Since only a small minority of the bacterial cells in soil are culturable (43), it is important to evaluate and compare
results based on culturing with DNA-based methods. Olsen and Bakken
(31) suggested that the ecological significance of
culturable cells is large, as they represent 80 to 90% of the
bacterial biovolume. The unculturable cells are a blend of species that
we cannot culture and species with some cells that are in an
unculturable state because of (e.g.) stresses (33).
We used the quantitative Pseudomonas-selective PCR
method (14), developed an actinomycete (mainly
Streptomyces)-selective PCR, and compared
selective CFU counts with the amounts of specific DNAs from the
two groups.
| |
MATERIALS AND METHODS |
Mesocosm setup.
The soil used was a sandy loam soil from the
Royal Veterinary and Agricultural University experimental field station
in Høje Taastrup, Denmark. Soil characteristics are as follows: coarse sand 69.9%; fine sand, 18.8%; silt, 3.0%; clay, 3.5%, organic matter, 4.8% (dry weight); water holding capacity, 18.9% (dry weight). Soil was sampled in September 1998 and stored at 4°C for 2 weeks before sieving (4 mm). Macronutrients were added in the following
concentrations (milligrams of nutrient kilogram of
soil1): N, 40; P, 8.6; K, 23; Mg, 10. Micronutrients were
added at the following concentrations: (micrograms of nutrient kilogram of soil1): B, 100; Mn, 100; Zn, 100; Cu, 3; Mo, 2. The
soil was thoroughly mixed and distributed in mesocosms for growth of
barley plants.
The mesocosms consisted of nontransparent plastic tubes with a diameter
of 6 cm and a length of either 70 or 35 cm. Each tube was cut into two
halves lengthwise; the halves were held together by strong waterproof
tape, and at the bottom end was closed with polyethylene mesh to hold
back the soil. This enabled destructive sampling of the mesocosms by
opening lengthwise and taking out the soil core. The long tubes
contained 2.40 kg of soil, and the short tubes contained 1.15 kg of
soil. After addition of soil to the mesocosms, the soil water content
was adjusted to 80% of water holding capacity and the soil was
incubated at 10°C for 5 days. The mesocosms were placed in a climate
chamber with light from four halogen-mercury 150-W lamps (HQI-TS NDL;
Osram Sylvania, Danvers, Mass.), with a light intensity of about 300 µE m2 s1 in a 16-h light, 8-h dark cycle.
During the light period the temperature in the growth chamber was
15°C, and during the dark period it was 10°C. After 1 day, seeds
were sown. A batch of spring-barley seeds (Hordeum vulgare
type Lamba) was coated with either imazalil in the form of Fungazil A
(Cillus, Herlev, Denmark) or P. fluorescens DR54. DR54
was chromosomally marked with green fluorescent protein (GFP)
(30), which enabled us to distinguish it from the
indigenous Pseudomonas. The resulting mutant, DR54-BN14, did
not differ from the wild type in various physiological tests and barley
root colonization (30). Imazalil was sprayed on seeds in
the recommended dose (1 ml per kg of seeds) by means of a type HEGE 11 rotator (Hans Ulrich Hege Maschinenbau, Waldenburg, Germany). At the
day of sowing, DR54 was applied to seeds by adding 170 seeds to 120 ml of washed overnight culture with 2.0 × 1010 cells
ml1 (Luria-Bertani medium supplemented with 0.10%
glucose [22], washed twice in 0.010 M phosphate buffer
[pH 7.4]). The cell suspension was gently shaken for 30 min before
sowing, which resulted in 5 × 107 CFU of DR54 per
seed sown. In parallel to this treatment, untreated seeds were shaken
in clean phosphate buffer for 30 min before sowing. Imazalil-coated
seeds were sown dry, but 20 µl of phosphate buffer was pipetted on
top of each seed. This corresponded to the mean amount of phosphate
buffer adhering to DR54-coated seeds and control seeds.
Three seeds were sown per mesocosm at a 3-cm depth. Mesocosms with
different treated seeds were placed randomly in the climate chamber.
Throughout the experimental period mesocosms were regularly weighed and
rewatered to maintain the water content. Where all three seeds
germinated, one was removed, leaving two plants per mesocosm. For the
first sampling occasions more seeds were sown to ensure enough
rhizosphere soil for analysis. For the first four sampling occasions
(days 4, 7, 10, and 14), the short tubes were used, and for all other
sampling occasions the long tubes were used. After 50 days,
above-ground plant material was removed to mimic harvest, initiating
decomposition of the roots.
Sampling.
Sampling was done 4, 7, 10, 14, 21, 35, 50, 63, 91, and 112 days after sowing. On each sampling occasion three
replicate mesocosms from each of the treatments were destructively
sampled. The rhizospheres of the two plants were pooled. At all
sampling days rhizosphere samples from the upper 5 cm of the root
system were collected, and at days 14, 35, 50, 63, and 91 rhizosphere
samples from the root tips were also taken. Bulk soil from mesocosms
with untreated seeds was analyzed on days 0, 10, 21, 50, and 112 by
sampling ca. 0.5 g of root-free soil. Rhizosphere samples of
approximately 30 cm of root with adhering soil and 0.5-g bulk soil
samples were aseptically transferred to glass tubes containing 6 ml of
phosphate buffer and mildly sonicated in an ultrasonic water bath for
30 s (Branson 5210; Merck Eurolab, Albertslund, Denmark). This
soil suspension was used for plate spreading and quantitative PCR. Samples consisting of soil and roots were weighed, as well as roots
alone. Furthermore, the actual water content of the soil in each tube
was measured.
Plate counts.
The number of culturable
Pseudomonas organisms was counted on the
Pseudomonas-selective (15, 18) Gould's S1
medium (9) amended with 50 mg of the fungal inhibitor
delvocid (containing 50% natamycin and 50% lactose) dissolved in 10 ml of methanol liter1. Fifty microliters of appropriate
dilutions was plate spread and incubated at 20°C for 3 days. Three
replicate plates were counted per sample, and the total number of
colonies and the DR54 GFP-positive colonies were determined under blue
light in a microscope.
Filamentous actinomycetes were counted on Winogradsky agar (containing,
per liter, 5.0 g of K2HPO4, 2.5 g of MgSO4 · 7H2O, 2.5 g of NaCl,
0.05 g of MnSO4 · 1H2O, 0.0025 g
of FeSO4 · 7H2O, and 18 g of Bacto
Agar [Difco, Detroit, Mich.]) amended with 25 mg of natamycin
dissolved in 10 ml of methanol liter1 to inhibit fungal
growth. Fifty microliters of appropriate dilutions was spread on agar
plates. Three replicate plates were counted per sample after 4 weeks of
incubation at 20°C. Filamentous actinomycetes were recognized by
their colony morphology.
Quantitative PCR.
DNA extraction and purification from the
soil samples were done with the Fast Soil purification kit (Bio 101, Vista, Calif.) in accordance with the manufacturer's instructions
(1). One DNA extract was used per sample to determine the
rDNA target number. This DNA extraction procedure decreased the number
of intact actinomycete spores to 25 ± 3% of the initial amount.
This was determined by counting suspensions of spores from
Streptomyces lavendulae DSM 40069T and
Streptomyces diastaticus DSM 40446T before and
after bead beating. Following this treatment, the content of the bead
beater tube was resuspended and beads were removed by centrifugation,
leaving spores in the supernatant. Quantitative competitive PCR on
Pseudomonas was done as described elsewhere
(14). Essentially, an internal standard, which is a
shorter fragment with primers identical to the native DNA in the ends,
was used as competitive template DNA in the PCR. Each PCR tube
contained a total volume of 25 µl, with 17.8 µl of DNase-free water, 2.4 µl of AmpliTaq PCR buffer (PE Biosystems, Norwalk, Conn.),
2.4 µl of bovine serum albumin (Amersham Pharmacia, Uppsala, Sweden),
1 µl of deoxynucleoside triphosphate mixture (PE Biosystems), 0.12 µl (0.1 mM) each of the Pseudomonas-specific primer
PSMG (2) and of the
Bacteria-specific 9-27 primer (37) (Life
Technologies, Roskilde, Denmark), 0.12 µl of AmpliTaq Gold polymerase
(PE Biosystems), and 1.0 µl of sample as template. The specificity of
PSMG was verified on 8 August 2000 on the Ribosomal
Database Project (RDP) database (21). Furthermore, the PCR
products from the imazalil treatment after 91 days were cloned using
the TOPA TA Cloning Kit (Invitrogen, Carlsbad, Calif.). Clones
containing the entire insert were identified by PCR using the
Pseudomonas-specific primers as described above. The inserts
in 10 clones were PCR amplified using the primers M13F and M13R as
recommended by the manufacturer and were sequenced at the sequencing
facility at GATC GmbH (Konstanz, Germany). All sequences aligned within
the group of Pseudomonas and relatives when tested in the
RDP database (21). Linear dilutions of the internal
standard were used, and ethidium bromide-stained band intensities on
the gels were quantified by a UV-visible-light recording camera using
the Multi-Analyst software (Bio-Rad, Hercules, Calif.).
A quantitative competitive PCR selective for Streptomyces
and related actinomycetes was developed. The master mix used was as
described above except for the primers. The primer set was the forward
primer F243 (5' GGATGAGCCCGCGGCCTA 3'), which is reported to
be specific for detection of actinomycetes, and the reverse primer R513
(5' CCGCGGCTGCTGGCACGTA 3'), which is less specific, targeting actinomycetes and others (13). The original
primer of Heuer et al. (13) was modified by removing three
bases from the 5' end but retained its specificity. The PCR conditions
used were as follows: 10 min at 95°C; 35 cycles of 30 s at
95°C and 1 min at 72°C; 6 min at 72°C; and final cooling at
5°C. Alignment of primer F243 to 16S ribosomal DNA in the RDP
database (21) showed that strains of
Streptomyces and a few other genera such as
Saccharopolyspora, Mycobacteria, Corynebacterium, and
Rhodococcus had no mismatches to primer F243, while many
other actinomycete genera had at least one mismatch. However, not all
strains within the non-Streptomyces genera mentioned above
were targeted by F243. PCR conditions were specific for the species
without mismatches (Table 1).
Construction of the internal standard was carried out as described by
Hallier-Soulier et al. (10). Two primers within the 284-bp
fragment spanning F243 and R513 in Streptomyces albidoflavus DSM 40455T (InternR [5'
CTGACTCGATGCGTATCCCCACTGCTGCCTCCC 3'] and InternF [5' TACGCATCGAGTCAGGGGATGACGGCCTTCGGGTT 3'])
were constructed. The internal primers had a 15-bp overlapping
region (underlined), where the sequences were complementary. The
constructed internal standard was 255 bp, and the PCR product was
purified with a QIAquick PCR purification kit (Qiagen GmbH, Hilden,
Germany). DNA from S. albidoflavus DSM 40455T
was purified by Fast Soil DNA and mixed with the internal standard in
different mixtures to compare amplification efficiencies for the two
competing templates as described by Suzuki and Giovannoni (37). The internal standard DNA was amplified slightly
preferentially to the unmodified S. albidoflavus DNA. When
the proportions of the internal standard were 0.2, 0.4, 0.6, and 0.8 in
the PCR template they were 0.28, 0.51, 0.71, and 0.82, respectively, in
the PCR product. This was taken into account when calculating the
number of actinomycete-specific DNA copies in the samples by using a standard curve.
Statistical analysis.
Both numbers of CFU and quantitative
PCR data were log transformed and analyzed by a two-way analysis of
variance by the procedure ANOVA or GLM in SAS 6.12 for the effects of
sampling time, antifungal treatment, and interaction between the two
factors. To compare antifungal treatments with the control and the
effect of time on the population development, appropriate
least-significant-difference values at a 95% confidence level were used.
| |
RESULTS AND DISCUSSION |
Fate of P. fluorescens DR54.
In the rhizosphere
near the root base of DR54-treated plants, the strain had a high
relative occurrence (>75% of the total Pseudomonas
population) during the first 21 days, but from day 50 onwards it
constituted less than 10% (Fig. 1). In
the rhizosphere of root tips DR54 constituted less than 0.1% of the
total Pseudomonas population at all sampling days (data not
shown), which shows that DR54 is not a successful root colonizer, in
agreement with the results of Normander et al. (30). Those
authors divided a barley rhizosphere in three parts, upper (near the
seed), middle, and lower, and found that the number of DR54 organisms
was decreasing from the upper to the lower rhizosphere during the first
14 days of rhizosphere development. Natsch et al. (27)
monitored the survival of P. fluorescens CHA0 in wheat
rhizosphere, in which the abundance decreased from ca. 106
root system1 to 104 root
system1 in 42 days. However, the results were not for
root segments but were pooled for the whole root. We found no
GFP-positive colonies in any rhizosphere samples of untreated or
imazalil-treated plants, confirming that there was no contamination by
GFP-positive colonies.
View larger version (14K):
[in this window]
[in a new window]
|
FIG. 1.
P. fluorescens DR54 on root base
rhizosphere during growth, harvest, and decomposition of barley.
The proportion of culturable DR54 to the total number of CFU on
Gould's S1 agar is shown. Error bars show standard errors of the
means.
|
|
Effects of P. fluorescens DR54 and imazalil on the
indigenous Pseudomonas and actinomycete populations.
Both DR54 and imazalil transiently affected the indigenous
Pseudomonas population in the rhizosphere near the root base
(Fig. 2). The DR54 treatment resulted in
significantly higher numbers of CFU and DNA copies of
Pseudomonas at days 4 and 7, compared to the control. This
was not surprising, as both methods include DR54, which was inoculated
at 5 × 107 CFU per seed. If the DR54 CFU counts are
subtracted from the total Pseudomonas counts, a marked
inhibition of the indigenous Pseudomonas is seen the first
21 days. Nevertheless, the appearance of the indigenous
Pseudomonas on Gould's S1 plates has most likely been
suppressed to some degree by the emergence of fast-growing DR54
colonies at samplings when the number of these colonies was high. This
results in a false picture of the suppression. However, at days 10 and
14 both Pseudomonas CFU and DNA show no difference between
treatments (Fig. 2), when DR54 still constituted ca. 80% of the CFU.
If the inhibition of indigenous Pseudomonas on the plates
was pronounced, we would expect a higher number of
Pseudomonas DNA copies in the DR54 treatment group at these
samplings. This was not the case. Therefore, the data suggest that DR54
in a period of ca. 2 weeks displaced a part of the indigenous
Pseudomonas population. Similar results were found by Natsch
et al. (26) for P. fluorescens CHA0, which
probably competed for the same ecological niche as the indigenous
Pseudomonas in the rhizosphere.
View larger version (18K):
[in this window]
[in a new window]
|
FIG. 2.
Numbers of culturable indigenous Pseudomonas
spp. as measured by plate spreading on Gould's S1 agar (A) and numbers
of specific rDNA copies measured by quantitative competitive PCR (B) on
root base rhizosphere during growth, harvest, and decomposition of
barley. Results for untreated seeds ( ), seeds treated with P. fluorescens DR54 ( ), and seeds treated with imazalil ( )
are shown. Error bars show standard errors of the means. dw, dry
weight.
|
|
Four days after sowing, imazalil caused a significantly higher level of
indigenous Pseudomonas compared to the untreated control. This was seen both by CFU, which showed six times more
Pseudomonas colonies than the control, and quantitative PCR,
which showed three times more Pseudomonas-specific DNA
copies (Fig. 2B). As Pseudomonas is known to be able to
degrade many xenobiotic compounds (35),
Pseudomonas bacteria may have been active players in the degradation of the fungicide. Another possibility is that they benefited from the displacement of other taxa, leading to improved conditions for Pseudomonas. In the only study of effects of
imazalil on bacteria, Fisher and Hayes (7) found that the
numbers and function of Rhizobium trifolii were unaffected
by imazalil at field concentrations. However, it is not possible to
draw general conclusions from their observation and hence to determine
what effects the compound had indirectly or directly on
Pseudomonas. There were significantly fewer
Pseudomonas CFU at day 35 in the imazalil treatment group
compared to the untreated control (Fig. 2A). However, this was not
found by quantitative PCR, which, on the other hand, showed
significantly lower numbers in the imazalil treatment group at day 10 and higher numbers at day 21 (Fig. 2B). These changes are probably
repercussions from the effects seen after 4 days and show the risk of
side effects of imazalil and DR54 on soil bacteria.
Imazalil and DR54 did not affect the actinomycete population near the
root base (data not shown). Therefore, the treatments are considered
replicates for presentation of actinomycete CFU and specific DNA copies
(see Fig. 3). Likewise, the treatments did not affect the numbers of
CFU of the two bacterial groups around root tips (see Tables 2 and 3).
The difference in effects between Pseudomonas and
actinomycetes demonstrates that side effects can be specific to certain
taxa, and hence it is recommended not to monitor large groups like
Bacteria exclusively when studying side effects.
Succession of Pseudomonas.
The root base
rhizosphere represented a habitat changing from young to old
rhizosphere to a decomposing hot spot. Across this gradient, a
succession in abundance of Pseudomonas and actinomycetes was observed.
Pseudomonas numbers in untreated samples in rhizosphere
around the root base increased significantly from young to 50-day-old rhizosphere, with respect to both culturable organisms (Fig. 2A) and
the amount of specific DNA copies (Fig. 2B). Furthermore, the level of
Pseudomonas CFU in bulk soil throughout the experiment and
the level of Pseudomonas DNA copies in bulk soil at day 0 (Table 2) were significantly lower than
those in the root base rhizosphere. This supports the view that
Pseudomonas bacteria are rhizosphere competent (25,
34) and hence confirms that the agar plate results in the
literature are not biased by cultivation. We found fewer
Pseudomonas organisms in the root tip rhizosphere (Table 2)
than at the root base, probably because the root tips represented young
rhizosphere in which the Pseudomonas population had not yet
proliferated.
After the plant shoot was removed at day 50 and until day 91, Pseudomonas CFU in the root base rhizosphere were not
significantly different (Fig. 2A). However, the number of
Pseudomonas-specific DNA copies (Fig. 2B) was
significantly reduced right after the plant shoots were removed. In
rhizosphere samples ca. 1,500 DNA copies per CFU were found, while
around decomposing roots ca. 250 DNA copies per CFU were found (Fig.
2). In soil, low culturability (0.1 to 10%), depending on medium,
incubation time, and conditions, has been reported (43),
but to our knowledge this has not been studied for
Pseudomonas alone. Nevertheless, Pseudomonas
introduced into bulk soil or rhizosphere of barley or wheat is
approximately 0.1 to 100% culturable (12, 30, 40, 41). We
consider it unlikely that the smaller amount of DNA present around
decomposing roots is due to changed DNA extraction efficiency compared
to rhizosphere samples. However, it has not been possible to find studies of the efficiency of extraction of Pseudomonas in
different physiological states or for different species within the
genus. The number of 16S ribosomal gene copies in the genomes of the Pseudomonas strains tested varies from four to six
(11, 32). Thus, it is unlikely that changes in
Pseudomonas diversity have influenced the quantitative PCR
results markedly. The decrease in the number of DNA copies following
harvest is thus considered to be due to a decrease in the number of
Pseudomonas organisms. Hence, this indicates that fewer
Pseudomonas cells are present around decomposing roots than
in the rhizosphere but that a higher fraction of them are culturable.
Johnsen et al. (14) reported a ratio of ca. 10 DNA copies
per CFU using the same methods as us. However, this was during the
decomposition of 12-day-old barley roots, which are considerably less
lignified than the roots in this study. Therefore, differences in the
environments presumably control Pseudomonas culturability.
Succession of actinomycetes.
In the rhizosphere near the root
base, the CFU of filamentous actinomycetes showed a small but
significant increase from day 4 to 63 (Fig.
3A). Compared with
Pseudomonas, the actinomycete abundance was stable, though,
as found in wheat rhizosphere by Miller et al. (25). Late
in the decomposition process (days 91 and 112), the number of CFU was
significantly higher. This is in accordance with the classical view
that actinomycetes persist during the microbial succession beyond the
initial phase of the degradation process because of their ability to
penetrate and solubilize many polymers (24). Because of
the fungus-like growth form, CFU from filamentous actinomycetes
originate both from spores and from fragmented hyphae. CFU will
probably often overrepresent the spores, because fragmentation of
vegetative hyphae into single cells is not possible. A harsh
fragmentation treatment will break vegetative hyphae into nonviable
fragments (23). About 95% of the counted colonies of
filamentous actinomycetes can be expected to belong to the genus
Streptomyces (44).
View larger version (13K):
[in this window]
[in a new window]
|
FIG. 3.
Numbers of culturable actinomycetes as measured by plate
spreading on Winogradsky agar (A) and numbers of specific rDNA copies
measured by quantitative competitive PCR (B) on root base rhizosphere
during growth, harvest, and decomposition of barley. Graphs show means
of the three seed treatments, which were not significantly
different. Error bars show standard errors of the means. dw, dry
weight.
|
|
The numbers of actinomycete CFU in bulk soil and root tip rhizosphere
were in the same range as observed in the root base rhizosphere (Table
3). This is in contrast to the results of Miller et al. (25), who found that actinomycete numbers in
wheat rhizosphere were ca. 10-fold higher than those in bulk soil, but many investigations have shown that this varies between plant species
(see, e.g., references 5 and 25).
The results of the quantitative PCR showed an increase in
actinomycete-specific DNA from young to older rhizosphere (Fig. 3B).
After harvest, a significant decrease in specific DNA was observed,
followed by a significant increase in the late decomposition phase,
even though this increase was not as distinct as for the CFU. The level
of specific DNA in bulk soil and root tip rhizosphere was the same as
in the root base rhizosphere (Table 3), supporting the CFU results.
For actinomycetes, there are two differences between the CFU and
quantitative PCR methods. The first concerns the specificity of the two
methods. Because the vast majority of CFU counted on agar plates most
likely are Streptomyces, the quantitative PCR method was
directed towards Streptomyces rather than towards the whole
gram-positive, high-GC group. This group was the target in the work of
Heuer et al. (13), who used the same primers. Our protocol
was more stringent, as we raised the annealing temperature from 65 to
72°C. DNAs from all tested soil isolates of filamentous actinomycetes
and Streptomyces strains, but also DNAs from strains from a
few other actinomycete groups, were amplified by the PCR protocol used.
Second, the specific DNAs conceivably represent spores and hyphae more
equally, even though the DNA contents in and extraction efficiencies
from spores and hyphae may differ (8). We found that the
great majority of spores from two Streptomyces strains were
broken with the bead beating method used in this experiment (data not
shown). Assuming that the specific DNA represents spores and hyphae
more equally than CFU, the larger increase in the rhizosphere seen by
quantitative PCR can be explained by actively growing mycelia,
colonizing the cortex in old roots (42), or a population of actinomycetes not detected on agar plates.
Concluding remarks.
In general, our quantitative PCR results
confirm the plate counting results and hence the conceptions of the
ecology of Pseudomonas and actinomycetes in the rhizosphere.
Thus, quantitative PCR has confirmed that Pseudomonas
spp. are fast colonizers whereas actinomycetes colonize roots at later
stages. However, the increased culturability of Pseudomonas
following barley harvest and the increase in actinomycete-specific DNA
in the rhizosphere stress that plate spreading alone leaves out
important information. These changes may be the result of a rise
of in situ Pseudomonas and actinomycete activity. DR54 and
imazalil transiently affected the number of indigenous
Pseudomonas organisms, but not actinomycetes, demonstrating
that side effects can be specific to certain taxa.
| |
ACKNOWLEDGMENTS |
We thank Svend J. Binnerup for commenting on the manuscript.
This work was funded the Centre for Effects and Risks of Biotechnology
in Agriculture and the Centre for Biological Processes in Contaminated
Soil and Sediments under the Danish Environmental Research Programme.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Geological
Survey of Denmark and Greenland, Department of Geochemistry, Thoravej
8, DK-2400 Copenhagen NV, Denmark. Phone: 45 38 14 20 00. Fax: 45 38 14 20 50. E-mail: kj{at}geus.dk.
| |
REFERENCES |
| 1.
|
Borneman, J.,
P. W. Skroch,
K. M. O'Sullivan,
J. A. Palus,
N. G. Rumjanek,
J. L. Jansen,
J. Nienhuis, and E. W. Triplett.
1996.
Molecular microbial diversity of an agricultural soil in Wisconsin.
Appl. Environ. Microbiol.
62:1935-1943[Abstract].
|
| 2.
|
Braun-Howland, E. B.,
P. A. Vescio, and S. A. Nierzwicki-Bauer.
1993.
Use of a simplified cell blot technique and 16S rRNA-directed probes for identification of common environmental isolates.
Appl. Environ. Microbiol.
59:3219-3224[Abstract/Free Full Text].
|
| 3.
|
Buchenauer, H.
1990.
Physiological reactions in the inhibition of plant pathogenic fungi, p. 217-292.
In
G. Haug, and H. Hoffmann (ed.), Chemistry of plant protection. Springer-Verlag, Berlin, Germany.
|
| 4.
|
Chatterjee, S. K., and B. Nandi.
1981.
Biodegradation of wheat stubbles by soil microorganisms and role of the products on soil fertility.
Plant Soil
59:381-390[CrossRef].
|
| 5.
|
Dickinson, C. H., and M. J. Dooley.
1967.
The microbiology of cut-away peat. 1. Descriptive ecology.
Plant Soil
27:172-186[CrossRef].
|
| 6.
|
Embley, T. M., and E. Stackebrandt.
1994.
The molecular phylogeny and systematics of the actinomycetes.
Annu. Rev. Microbiol.
48:257-289[Medline].
|
| 7.
|
Fisher, D. J., and A. L. Hayes.
1982.
Effects of some systemic imidazole and triazole fungicides on white clover and symbiotic nitrogen fixation by Rhizobium trifolii.
Ann. Appl. Biol.
101:19-24.
|
| 8.
|
Frostegård, Å.,
S. Courtois,
V. Ramisse,
S. Clerc,
D. Bernillon,
F. le Gall,
P. Jeannin,
X. Nesme, and P. Simonet.
1999.
Quantification of bias related to the extraction of DNA directly from soils.
Appl. Environ. Microbiol.
65:5409-5420[Abstract/Free Full Text].
|
| 9.
|
Gould, W. D.,
C. Hagedorn,
T. R. Bardinelli, and R. M. Zablotowicz.
1985.
New selective media for enumeration and recovery of fluorescent pseudomonads from various habitats.
Appl. Environ. Microbiol.
49:28-32[Abstract/Free Full Text].
|
| 10.
|
Hallier-Soulier, S.,
V. Ducrocq,
N. Mazure, and N. Truffaut.
1996.
Detection and quantification of degradative genes in soils contaminated by toluene.
FEMS Microbiol. Ecol.
20:121-133[CrossRef].
|
| 11.
|
Hartmann, R. K.,
H. Y. Toschka,
N. Ulbrich, and V. A. Erdmann.
1986.
Genomic organization of rDNA in Pseudomonas aeruginosa.
FEBS Lett.
195:187-193[CrossRef][Medline].
|
| 12.
|
Hase, C.,
F. Mascher,
Y. Moënne-Loccoz, and G. Défago.
1999.
Nutrient deprivation and the subsequent survival of biocontrol Pseudomonas fluorescens CHA0 in soil.
Soil Biol. Biochem.
31:1181-1188[CrossRef].
|
| 13.
|
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].
|
| 14.
|
Johnsen, K.,
Ø. Enger,
C. S. Jacobsen,
L. Thirup, and V. Torsvik.
1999.
Quantitative selective PCR of 16S ribosomal DNA correlates well with selective agar plating in describing population dynamics of indigenous Pseudomonas spp. in soil hot spots.
Appl. Environ. Microbiol.
65:1786-1788[Abstract/Free Full Text].
|
| 15.
|
Johnsen, K., and P. Nielsen.
1999.
Diversity of Pseudomonas strains isolated with King's B and Gould's S1 agar determined by repetitive extragenic palindromic-polymerase chain reaction, 16S rDNA sequencing and Fourier transform infrared spectroscopy characterisation.
FEMS Microbiol. Lett.
173:155-162[CrossRef][Medline].
|
| 16.
|
Johnsson, L.,
M. Hökeberg, and B. Gerhardson.
1998.
Performance of the Pseudomonas chlororaphis biocontrol agent MA 342 against cereal seed-borne diseases in field experiments.
Eur. J. Plant Pathol.
104:701-711[CrossRef].
|
| 17.
|
Kersters, K.,
W. Ludwig,
M. Vancanneyt,
P. de Vos,
M. Gillis, and K.-H. Schleifer.
1996.
Recent changes in the classification of the pseudomonads: an overview.
Syst. Appl. Microbiol.
19:465-477.
|
| 18.
|
Kragelund, L.,
K. Leopold, and O. Nybroe.
1996.
Outer membrane protein heterogeneity within Pseudomonas fluorescens and P. putida and use of an OprF antibody as a probe for rRNA homology group I pseudomonads.
Appl. Environ. Microbiol.
62:480-485[Abstract].
|
| 19.
|
Lacey, J.
1973.
Actinomycetes in soils, composts and fodders, p. 231-251.
In
G. Sykes, and F. A. Skinner (ed.), Actinomycetales: characteristics and practical importance Academic Press, London, United Kingdom.
|
| 20.
|
Lambert, B.,
P. Meire,
H. Joos,
P. Lens, and J. Swings.
1990.
Fast-growing, aerobic, heterotrophic bacteria from the rhizosphere of young sugar beet plants.
Appl. Environ. Microbiol.
56:3375-3381[Abstract/Free Full Text].
|
| 21.
|
Maidak, B. L.,
J. R. Cole,
T. G. Lilburn,
C. T. J. Parker,
P. R. Saxman,
J. M. Stredwick,
G. M. Garrity,
B. Li,
G. J. Olsen,
S. Pramanik,
T. M. Schmidt, and J. M. Tiedje.
2000.
The RDP (Ribosomal Database Project) continues.
Nucleic Acids Res.
28:173-174[Abstract/Free Full Text].
|
| 22.
|
Maniatis, T.,
E. F. Fritsch, and J. Sambrook.
1982.
Molecular cloning: a laboratory manual.
Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
|
| 23.
|
Mayfield, C. I.,
S. T. Williams,
S. M. Ruddick, and H. L. Hatfield.
1972.
Studies on the ecology of actinomycetes in soil. IV. Observations on the form and growth of streptomycetes in soil.
Soil Biol. Biochem.
4:79-91[CrossRef].
|
| 24.
|
McCarthy, A. J., and S. T. Williams.
1990.
Methods for studying the ecology of actinomycetes.
Methods Microbiol.
22:533-563.
|
| 25.
|
Miller, H. J.,
E. Liljeroth,
G. Henken, and J. A. van Veen.
1990.
Fluctuations in the fluorescent pseudomonad and actinomycete populations of rhizosphere and rhizoplane during the growth of spring wheat.
Can. J. Microbiol.
36:254-258.
|
| 26.
|
Natsch, A.,
C. Keel,
N. Hebecker,
E. Laasik, and G. Défago.
1997.
Influence of biocontrol strain Pseudomonas fluorescens CHA0 and its antibiotic overproducing derivative on the diversity of resident root colonizing pseudomonads.
FEMS Microbiol. Ecol.
23:341-352[CrossRef].
|
| 27.
|
Natsch, A.,
C. Keel,
H. A. Pfirter,
D. Haas, and G. Défago.
1994.
Contribution of the global regulator gene gacA to persistence and dissemination of Pseudomonas fluorescens biocontrol strain CHA0 introduced into soil microcosms.
Appl. Environ. Microbiol.
60:2553-2560[Abstract/Free Full Text].
|
| 28.
|
Nielsen, M. N.,
J. Sørensen,
J. Fels, and H. C. Pedersen.
1998.
Secondary metabolite- and endochitinase-dependent antagonism toward plant-pathogenic microfungi of Pseudomonas fluorescens isolates from sugar beet rhizosphere.
Appl. Environ. Microbiol.
64:3563-3569[Abstract/Free Full Text].
|
| 29.
|
Nielsen, T. H.,
C. Christophersen,
U. Anthoni, and J. Sørensen.
1999.
Viscosinamide, a new cyclic depsipeptide with surfactant and antifungal properties produced by Pseudomonas fluorescens DR54.
J. Appl. Microbiol.
87:80-90[CrossRef][Medline].
|
| 30.
|
Normander, B.,
N. B. Hendriksen, and O. Nybroe.
1999.
Green fluorescent protein-marked Pseudomonas fluorescens: localization, viability, and activity in the natural barley rhizosphere.
Appl. Environ. Microbiol.
65:4646-4651[Abstract/Free Full Text].
|
| 31.
|
Olsen, R. A., and L. R. Bakken.
1987.
Viability of soil bacteria: optimization of plate-counting technique and comparison between total counts and plate counts within different size groups.
Microb. Ecol.
13:59-74.
|
| 32.
|
Ramos-Díaz, M. A., and J. L. Ramos.
1998.
Combined physical and genetic map of the Pseudomonas putida KT2440 chromosome.
J. Bacteriol.
180:6352-6363[Abstract/Free Full Text].
|
| 33.
|
Roszak, D. B., and R. R. Colwell.
1987.
Survival strategies of bacteria in the natural environment.
Microbiol. Rev.
51:365-379[Free Full Text].
|
| 34.
|
Rovira, A. D., and D. C. Sands.
1971.
Fluorescent pseudomonads a residual component in the soil microflora?
J. Appl. Bacteriol.
34:253-259[Medline].
|
| 35.
|
Sayler, G. S.,
S. W. Hooper,
A. C. Layton, and J. M. H. King.
1990.
Catabolic plasmids of environmental and ecological significance.
Microb. Ecol.
19:1-20.
|
| 36.
|
Stutz, E. W.,
G. Défago, and H. Kern.
1986.
Naturally occurring fluorescent pseudomonads involved in suppression of black root rot of tobacco.
Phytopathology
76:181-185.
|
| 37.
|
Suzuki, M. T., and S. J. Giovannoni.
1996.
Bias caused by template annealing in the amplification of mixtures of 16S rRNA genes by PCR.
Appl. Environ. Microbiol.
62:625-630[Abstract].
|
| 38.
|
Thirup, L.,
F. Ekelund,
K. Johnsen, and C. S. Jacobsen.
2000.
Population dynamics of the fast-growing sub-populations of Pseudomonas and total bacteria, and their protozoan grazers, revealed by fenpropimorph treatment.
Soil Biol. Biochem.
32:1615-1623[CrossRef].
|
| 39.
|
Thrane, C.,
S. Olsson,
T. H. Nielsen, and J. Sørensen.
2000.
Vital fluorescent stains for detection of stress in Pythium ultimum and Rhizoctonia solani challenged with viscosinamide from Pseudomonas fluorescens DR54.
FEMS Microbiol. Ecol.
30:11-23.
|
| 40.
|
Troxler, J.,
M. Zala,
Y. Moënne-Loccoz,
C. Keel, and G. Défago.
1997.
Predominance of nonculturable cells of the biocontrol strain Pseudomonas fluorescens CHA0 in the surface horizon of large outdoor lysimeters.
Appl. Environ. Microbiol.
63:3776-3782[Abstract].
|
| 41.
|
van Overbeek, L. S.,
J. A. van Veen, and J. D. van Elsas.
1995.
Induced reporter gene activity, enhanced stress resistance, and competitive ability of a genetically modified Pseudomonas fluorescens strain released into a field plot planted with wheat.
Appl. Environ. Microbiol.
63:1965-1973[Abstract].
|
| 42.
|
Watson, E. T., and S. T. Williams.
1974.
Studies on the ecology of actinomycetes in soil. VII. Actinomycetes in a coastal sand belt.
Soil Biol. Biochem.
6:43-52.
|
| 43.
|
Winding, A.,
S. J. Binnerup, and J. Sørensen.
1994.
Viability of indigenous soil bacteria assayed by respiratory activity and growth.
Appl. Environ. Microbiol.
60:2869-2875[Abstract/Free Full Text].
|
| 44.
|
Xu, L.-H.,
Q.-R. Li, and C.-L. Jiang.
1996.
Diversity of soil actinomycetes in Yunnan, China.
Appl. Environ. Microbiol.
62:244-248[Abstract].
|
Applied and Environmental Microbiology, March 2001, p. 1147-1153, Vol. 67, No. 3
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.3.1147-1153.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
This article has been cited by other articles:
-
de Lipthay, J. R., Tuxen, N., Johnsen, K., Hansen, L. H., Albrechtsen, H.-J., Bjerg, P. L., Aamand, J.
(2002). In Situ Exposure to Low Herbicide Concentrations Affects Microbial Population Composition and Catabolic Gene Frequency in an Aerobic Shallow Aquifer. Appl. Environ. Microbiol.
69: 461-467
[Abstract]
[Full Text]
-
Aagot, N., Nybroe, O., Nielsen, P., Johnsen, K.
(2001). An Altered Pseudomonas Diversity Is Recovered from Soil by Using Nutrient-Poor Pseudomonas-Selective Soil Extract Media. Appl. Environ. Microbiol.
67: 5233-5239
[Abstract]
[Full Text]
-
Moenne-Loccoz, Y., Tichy, H.-V., O'Donnell, A., Simon, R., O'Gara, F.
(2001). Impact of 2,4-Diacetylphloroglucinol-Producing Biocontrol Strain Pseudomonas fluorescens F113 on Intraspecific Diversity of Resident Culturable Fluorescent Pseudomonads Associated with the Roots of Field-Grown Sugar Beet Seedlings. Appl. Environ. Microbiol.
67: 3418-3425
[Abstract]
[Full Text]
-
Fredslund, L., Ekelund, F., Jacobsen, C. S., Johnsen, K.
(2001). Development and Application of a Most-Probable-Number-PCR Assay To Quantify Flagellate Populations in Soil Samples. Appl. Environ. Microbiol.
67: 1613-1618
[Abstract]
[Full Text]
Top
Abstract
Introduction
