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Root colonization by
Pseudomonas putida : love
at first sight
Fluorescent pseudomonads are an important
part of the soil microbiota, frequently found
in association with plant roots. This interaction between bacteria and plants is often
beneficial for both partners. The surface of
roots and the surrounding soil areas (the
rhizosphere) constitute an environment where
nutrients released by the plant in the form of
root exudates are available to the microorganisms. In turn, some Pseudomonas
strains can improve plant growth and play a
protective role against pathogens. These activities and their potential use in agriculture
have prompted an increasing interest in the
study of mutualistic plant–Pseudomonas interactions. One of the most relevant aspects is
the process of bacterial establishment in the
rhizosphere, since an effective biocontrol depends on the efficiency of root colonization
(2). Most studies of factors potentially involved in root colonization efficiency draw
their conclusions from looking at wellestablished populations, i.e. after one to four
weeks of seedling inoculation and sowing.
Comparatively less attention has been paid to
the early stages of colonization. How do these
root–Pseudomonas interactions begin ?
Like in any love story, attraction is probably key for the initiation of mutualistic
plant–bacterial interactions. Bacterial chemotaxis on plates towards different nutrients
known to be present in root exudates has been
demonstrated (9). In Azospirillum brasilense,
non-flagellated and non-chemotactic mutants
showed reduced ability to colonize wheat
roots (10). Although the question of bacterial
motility in soil is still under debate, a series of
experiments performed by Bashan (1) with
Pseudomonas fluorescens and A. brasilense,
pointed out the existence of a directed motion
of bacterial cells towards wheat roots in soil.
However, this motion was heavily influenced
by soil composition and humidity (1). The
role of motility in attachment and colonization has been recently examined in detail by
Turnbull et al. (7, 8). Motility seems to be
important for competitive root colonization
by P. fluorescens (8), as well as for attachment
of Pseudomonas putida to wheat roots under
conditions of nutrient limitation (7).
As part of our on-going interest in mutualistic plant–Pseudomonas interactions, we
have now examined the initiation of root
colonization by P. putida in an in vitro system
that allows direct observation by time-lapse
microscopy.
The bacterium Pseudomonas putida
KT2440, a derivative of the soil isolate mt-2
(5), is able to colonize the root system of a
number of different plants, establishing and
persisting in the rhizosphere at a relatively
high population density (6). The initial establishment of P. putida KT2440 around corn
roots was followed by phase-contrast microscopy using a Nikon Diaphot 200 inverted
microscope. Corn seedlings (germinated for 4
days) were placed on Petri plates with 0n4 %
agar (w\v in sterile deionized water). A small
amount of bacterial culture was then inoculated " 1 mm away from the root with a
sterile toothpick. Images were captured at
20 s intervals with a CCD72 camera integrated with a Power Macintosh 8600\300
computer, processed using SCION IMAGE
software (Scion), a modification of NIH
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IMAGE (NIH), and mounted as three different movies, 25 s long, each corresponding to
1n5 h periods. These movies are available at
http :\\mic.sgmjournals.org
The bacteria immediately responded to
the presence of the plant, migrating from the
point of inoculation towards the root. During
the first hour after inoculation, motion of the
bacterial cells towards the root could be
clearly observed (Fig. 1a, b), suggesting a
rapid chemotactic response of KT2440 to the
presence of the plant (which of course would
depend on the distance from the root). After
this first stage of bacterial motion, microcolonies start to form on the surface and in
the vicinity of the root, their size steadily
increasing during the following hours (Fig. 1c,
d). This increase in size may result both from
the recruitment of new members into the
colony as well as from cell growth and
division. Dividing cells were in fact observed
during this period, suggesting that there is a
relatively abundant release of nutrients in the
form of root exudates. An intriguing question
raised by these microscopy data is if bacterial
aggregation and microcolony formation responds to the relative accumulation of nutrients around the root, or if there is a specific
signal triggering it.
After this fast establishment of the bacteria
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Editor-in-Chief via the Editorial Office.
341
Microbiology Comment
ROOT
(a)
(b)
(c)
(d)
ROOT
log c.f.u. g–1
8
(e)
7
6
5
4
3
0
4
8
12 16 20 24
Time (h)
................................................................................................................................................................................................................................................................................................................................
Fig. 1. Establishment of P. putida KT2440 around corn roots. Microscopy images were captured every 20 s for 4n5 h from the time of inoculation.
Selected images are shown. (a, b) Motility of KT2440 towards the root (dark area) (a) 5 min and (b) 1 h after inoculation. (c, d) Establishment and
growth of microcolonies of KT2440 around the corn root tip (c) 2n5 h and (d) 4n5 h after inoculation. Examples are indicated by arrows. (e) Corn root
colonization by P. putida KT2440 from inoculated seedlings. , No. cells in the bacterial suspension used to inoculate seedlings [log c.f.u. (g seed)−"] ;
$, bacteria associated with the seedling or root (log c.f.u. g−"). Inoculation and determination of the number of cells associated with the seeds\roots
was done as described (3, 4). Results are the average of two assays where each data point corresponds to duplicate samples from three seedlings. Error
bars are indicated.
in the vicinity of the root, the population
reached an equilibrium. Two days after inoculation, microscopy revealed a stable and
somewhat static population of bacteria
around the root, with only a small proportion
of free-swimming cells visible (not shown).
We compared these microscopy observations with a short-term study of bacterial
growth on the root from inoculated seedlings,
determined by viable counts. In this system,
the initial association takes place during
incubation in liquid medium, and is reflected
in the number of bacteria attached to the
seedlings.
Corn seedlings (germinated for 3 d) were
inoculated with a suspension of KT2440
[5i10' c.f.u. (g seedling)−", Fig. 1e], and
sown in pots containing vermiculite. At different times, plants were removed and the
number of bacteria associated with the root
was determined by plating on selective medium as described (3, 4). Around 1 % of the
inoculated bacteria attached to the seedlings
after 1 h incubation in minimal medium.
Interestingly, after sowing the seedlings, there
was a drop in the number of viable cells
recovered from them with respect to the initial
attached population. This phenomenon was
observed in all the experiments and could be
explained either by the bacteria spreading in
the solid substrate or by death of part of the
population. To determine which of these
possibilities was true, samples of vermiculite
were taken and the number of viable cells was
also estimated. Dispersal of KT2440 in the
vermiculite accounted almost completely for
the reduction in the number of cells recovered
from the seedlings (data not shown). The
reasons why this ‘ detachment ’ from the
seedlings takes place in the solid matrix
remain to be clarified.
Following this initial drop, the population
associated with the seedlings rapidly grew (τ
" 85 min during the first 3 h and " 60 min
the following 3 h), reaching " 2i10' c.f.u.
(g root)−" after 24 h and a maximum of
" 5i10( c.f.u. (g root)−" after 48 h. From
then on the number of c.f.u. (g root)−" stayed
essentially steady with slight fluctuations
for over a week (not shown), suggesting that
although seed and root exudates are released
constantly by the plant, the amount of nutrients present in them determines a maximal
population size per weight unit of plant
tissue.
The experiments presented here may not
342
be a direct reflection of the ‘ real world ’, but
they offer an interesting perspective of what
happens during the processes ordinarily used
to study root colonization in the lab. The
establishment of P. putida on plant roots
appears as a very fast and dynamic process in
which three stages can be defined : i) an
‘ attraction phase ’ of bacterial movement
probably resulting from chemotactic responses to the presence of the root ; ii) a rapid
‘ settlement phase ’, in which bacteria grow,
divide and aggregate to form microcolonies
around the root ; and iii) a ‘ residence phase ’,
where the established population reaches the
maximal size relative to root weight and its
further growth couples with plant growth, so
that the bacterial numbers relative to root
weight remain constant afterwards. In the
experiments presented here, starting from
inoculated seedlings (a protocol routinely
used in colonization studies), this maximum
is attained as soon as 48 h after seed inoculation. The dynamic nature of the early
colonization stages suggests they may be
substantial for the later events, and so their
study could answer key questions with respect
to the mechanisms of mutualistic plant–
Pseudomonas interactions.
Microbiology 148, February 2002
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Microbiology Comment
Acknowledgements
This work was supported by grant 99178
from the US–Spain Joint Commission for
Scientific and Technological Co-operation.
Manuel Espinosa-Urgel1, Roberto Kolter2
and Juan-Luis Ramos1
1
Department of Plant Biochemistry and
Molecular and Cellular Biology, Estacio! n
Experimental del Zaidı! n, CSIC, Profesor
Albareda 1, Granada 18008, Spain.
2
Department of Microbiology and Molecular
Genetics, Harvard Medical School, 200
Longwood Ave, Boston, MA 02115, USA.
Author for correspondence : Manuel
Espinosa-Urgel. Tel : j34 958 121011. Fax :
j34 958 129600.
e-mail : mespinos!eez.csic.es
1. Bashan, Y. (1986). Migration of the rhizosphere
bacteria Azospirillum brasilense and Pseudomonas
fluorescens towards wheat roots in the soil. J Gen
Microbiol 132, 3407–3414.
2. Chin-A-Woeng, T. F., Bloemberg, G. V., Mulders,
I. H., Dekkers, L. C. & Lugtenberg, B. J. (2000). Root
colonization by phenazine-1-carboxamide-producing
bacterium Pseudomonas chlororaphis PCL1391 is
essential for biocontrol of tomato foot and root rot. Mol
Plant Microbe Interact 13, 1340–1345.
3. Espinosa-Urgel M. & Ramos, J. L. (2001). Expression
of a Pseudomonas putida aminotransferase involved in
lysine catabolism is induced in the rhizosphere. Appl
Environ Microbiol 67, 5219–5224.
4. Espinosa-Urgel, M., Salido, A. & Ramos, J. L. (2000).
Genetic analysis of functions involved in adhesion of
Pseudomonas putida to seeds. J Bacteriol 182,
2363–2369.
5. Franklin, F. C. H., Bagdasarian, M., Bagdasarian,
M. M. & Timmis, K. N. (1981). Molecular and functional
analysis of the TOL plasmid pWW0 from Pseudomonas
putida and cloning of genes for the entire regulated
aromatic ring meta-cleavage pathway. Proc Natl Acad
Sci U S A 78, 7458–7462.
6. Molina, L., Ramos, C., Duque, E., Ronchel, M. C.,
Garcı! a, J. M., Wyke, L. & Ramos, J. L. (2000). Survival
of Pseudomonas putida KT2440 in soil and in the
rhizosphere of plants under greenhouse and
environmental conditions. Soil Biol Biochem 32, 315–321.
7. Turnbull, G. A., Morgan, J. A. W., Whipps, J. M. &
Saunders, J. R. (2001). The role of motility in the in vitro
attachment of Pseudomonas putida PaW8 to wheat
roots. FEMS Microbiol Ecol 35, 57–65.
8. Turnbull, G. A., Morgan, J. A. W., Whipps, J. M. &
Saunders, J. R. (2001). The role of motility in the
survival and spread of Pseudomonas fluorescens in soil
and in the attachment and colonization of wheat roots.
FEMS Microbiol Ecol 36, 21–31.
9. Vande Broek, A. & Vanderleyden, J. (1995). The role
of bacterial motility, chemotaxis, and attachment in
bacteria-plant interactions. Mol Plant-Microbe Interact
8, 800–810.
10. Vande Broek, A., Lambrecht, M. & Vanderleyden, J.
(1998). Bacterial chemotactic motility is important for
the initiation of wheat root colonization by Azospirillum
brasilense. Microbiology 144, 2599–2606.
New chlamydial lineages
from freshwater samples
Chlamydiae are important obligate intracellular bacteria, causing a variety of diseases
in vertebrates, including humans. Small sub-
unit ribosomal RNA gene (16S rDNA) sequence analyses have allowed the identification of new chlamydial lineages ; at present
four families, for a total of six genera and 13
species, have been described (6, 11, 15).
Among the new lineages the family Parachlamydiaceae that includes endosymbionts
of amoebae of the group Acanthamoeba –
Hartmannella is of interest. Parachlamydiae
have been identified in various strains of
amoebae isolated from soil, water conduit
systems, sewage sludge, corneal\contact lens
samples and nasal mucosa of healthy individuals (1, 9, 11), and there is evidence for
their implication in human respiratory infections (2, 4). 16S rDNA analyses showed
that this group of chlamydiae is very rich,
probably including several species and genera
(10). Simkania negevensis is another emerging
chlamydia, isolated initially as a cell culture
contaminant and successively associated with
human respiratory infection in Israel, Great
Britain and North America (8, 12). The
diversity within chlamydiae is likely to be
more important : using PCR, new 16S rDNA
phylotypes have been detected in clinical
samples (4, 14), and sequence databases contain over 100 partial 16S rDNA sequences
apparently belonging to distinct chlamydial
species. Increasing diversity within Chlamydiales is also shown by results reported by
Bowman et al. (3), who found 16S rDNA
sequences from sediments of Antarctic marine
salinity lakes (Burton-46 ; 999 bp) and a basin
(Taynaya-24 ; 995 bp). The sequence Burton46 seems to belong to an organism related to
parachlamydiae and Waddlia chondrophila,
while the sequence Taynaya-24 probably
represents a new lineage distantly related to
the other chlamydiae.
Herein we report for the first time the
presence of a Simkania-related sequence in an
environmental sample, as well as the presence
of another sequence representing a novel
chlamydial lineage distant from the other
chlamydiae described until now. These sequences were named cvE6 (GenBank accession no. AF448722) and cvE9 (GenBank accession no. AF448723), and were detected in
freshwater samples from two independent
ponds, one located in France, the other in
Italy.
Total DNA was extracted from 1 ml
aliquots of subsurface shore samples by the
classic phenol-chloroform method after proteinase K digestion. Chlamydial 16S rDNA
was searched for by using a pan-chlamydia
primer set amplifying almost all the gene
(nucleotide sequences for forward and reverse
primers were 5h-CGT GGA TGA GGC ATG
C(A\G)A GTC G-3h and 5h-GTC ATC
(A\G)GC C(T\C)(T\C) ACC TT(A\C\G)
(C\G)(A\G)C (A\G)(T\C)(T\C) TCT-3h,
positions 35–1481, Parachlamydia acanthamoebae 16S rDNA numbering). Manipulations were performed according to recommended guidelines and included negative
Microbiology 148, February 2002
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controls starting from the DNA extraction step. Both strands of an inner portion
(" 1100 bp) of the PCR products were sequenced (three repetitions) using a series of
inner primers, and the partial 16S rDNA
sequences were aligned with all chlamydial
16S rDNA sequences available in the dataset.
The sequences cvE6 and cvE9 showed
90n1 % similarity with each other, and values
of 87n9 and 92n7 %, respectively, with that of
S. negevensis. Sequence similarities of cvE6
and cvE9 with the other chlamydiae were
85n7 % and 86n4 % for W. chondrophila,
86n9–89n5 % and 89n1–91 % for Parachlamydiaceae, and 84–85 % and 86n5–87 % for Chlamydiaceae. In phylogenetic reconstructions
our sequences cluster with S. negevensis
showing high bootstrap values (Fig. 1a).
Phylogenetic analysis was also performed on
a 770 bp portion common to all chlamydial
16S rDNA sequences, including Antarctic
sequences (Fig. 1b). The sequence Burton-46
showed similarity values of 89n4–92n2 % to
those of parachlamydiae, and 88n7 % to that
of W. chondrophila. However, the branching
pattern was not clearly defined. Even if based
on partial sequence analysis only, it seems
clear that our sequence cvE9 was from an
organism specifically related to S. negevensis
(92n7 % similarity, 100 % bootstrap), representing either a new member of the Simkaniaceae or an organism of a new, closely
related lineage. This is the first report, to our
knowledge, on a Simkania-like organism from
environmental samples. On the contrary, the
sequence cvE6 seems to be distantly related to
the Antarctic sequence Taynaya-24 (87n9 %
similarity), and both seem to form new
independent lineages, perhaps distantly related to Simkaniaceae. Amplification with S.
negevensis-specific primers (12) was negative
in both our samples.
The presence of a Simkania-related organism in an environmental habitat might be
surprising. S. negevensis is at present the only
known member of the family Simkaniaceae.
Its natural host is unknown, and there is
evidence that human infections are widespread (8). Recently Kahane et al. (13) demonstrated S. negevensis being able in vitro to
infect and to multiply within Acanthamoeba.
It seems then conceivable that our sequence
cvE9 was from a protistan endosymbiont.
In addition, at least one member from the
Chlamydiaceae, Chlamydophila pneumoniae,
may also grow in vitro within acanthamoebae
(5). Intra-amoebal growth does not seem to be
restricted to parachlamydiae, and amoebae
and perhaps other protists (e.g. ciliates) may
be natural or occasional hosts of chlamydiae,
as it occurs for a large variety of bacteria.
Similarly, the most plausible explanation for
the origin of our second sequence, cvE6, as
well as of Antarctic sequences (3), is that they
are from chlamydial endosymbionts of protists. In addition, a rich mosaic of chloroplast
and eukaryotic gene homologues has been
343
Microbiology Comment
(a)
Neochlamydia hartmannellae
UWC22
100
50
0.05
UWE1
corvenA4
100 89
100
100
81
83
0.05
corvenA4
96
UWE1
UWF25
Parachlamydia acanthamoebae
Waddlia chondrophila
cvE6
Simkania negevenis
cvE9
50
Neochlamydia hartmannellae
UWC22
97
99 TUME1
81
97
UWE25
Parachlamydia acanthamoebae
Waddlia chondrophila
50
Burton-46
53
Taynaya-24
97
cvE6
78
Chlamydophila pneumoniae TW-183
Chlamydophila psittaci NJ1
100
Simkania negevenis
100
cvE9
Chlamydia trachomatis Har-13
Chlamydophila pneumoniae TW-183
100
vadinBE97
70
100
(b)
73
veCb1
Pirellula staleyi
Chlamydophila psittaci NJ1
Chlamydia trachomatis Har-13
vadinBE97
..................................................................................................................................................................................................................
Fig. 1. Phylogenetic reconstructions for partial 16S rDNA sequences using the neighbour-joining
method (Jukes & Cantor option), generated by Molecular Evolutionary Genetics Analysis (MEGA).
Topology stability was evaluated by bootstrapping (200 replicates), only values of 50 % or more are
indicated. Sequences cvE6 and cvE9 are indicated in bold. Uncultured bacterium vadinBE97 (vadin
lineage), verrucomicrobial strain VeCb1 (Verrucomicrobia), and Pirellula staleyi (Planctomycetales)
sequences were used as outgroups. Bars indicate estimated genetic distance. Nucleotide positions are P.
acanthamoebae 16S rDNA numbering. (a) Phylogenetic tree based on a 1053 bp fragment (nt positions
230–1284). (b) Phylogenetic tree based on a 770 bp fragment (nt positions 512–1284) to include also 16S
rDNA sequences Burton-46 and Taynaya-24 from Antarctic samples.
discovered in the sequenced genomes of Chlamydiaceae (16), and the 23S rDNA of S.
negevensis possesses an unspliced group I
intron that is closely related to introns found
in chloroplasts of some green algae (Chlamydomonas spp.) and in mitochondria of
Acanthamoeba (7). Such a genetic mosaicism
has led to the supposition that lateral gene
transfer events may have occurred in the
ancestors of present-day chlamydiae living
probably as endosymbionts of protists.
Searches in environmental habitats might
lead to the discovery of new chlamydial
lineages and\or new natural hosts other than
acanthamoebae. For example, the recently
discovered Neochlamydia hartmannellae, belonging to Parachlamydiaceae, is able to infect
both Hartmannella and Dictyostelium, but
not Acanthamoeba (11). This implies that
other eukaryotes may harbour chlamydial
organisms. Chlamydia-like organisms have
been morphologically described in some invertebrates as hydras and mussels, but confirmation by phylogenetic analyses is lacking.
It may be interesting also to evaluate the
protist host range of chlamydiae by experimental infections of various types of protists
other than amoebae, including ciliates and
algae. In this way it has already been shown
that both the respiratory pathogens C. pneumoniae and S. negevensis may survive and
multiply within Acanthamoeba (5, 13). They
might then utilize amoebae as a reservoir\
vector, like parachlamydiae or legionellae.
The ability to use intra-amoebal growth may
be considered as a preadaptation for successful infection of mammalian cells, including phagocytic ones.
Previous reports on new chlamydial organisms were from clinical samples (2, 4, 8,
12, 14, 15), and environmental isolates concerned exclusively new phylotypes of parachlamydiae (9–11). Our data and those of
Bowman et al. (3) indicated that new distinct
chlamydial lineages exist in natural environments as different as Antarctic marine salinity
lake sediments and European freshwater
ponds. All the chlamydiae described until
now are able to infect vertebrates and to
induce diseases : it may be of interest to
estimate their prevalence and diversity in
various natural habitats and in this way to
identify possible sources of infections for both
known and new chlamydiae.
Daniele Corsaro,1 Danielle Venditti2 and
Marcello Valassina3
1
Laboratoire de Virologie, Centre Hospitalier
Universitaire de Nancy, France.
2
TREDI De! partement Recherche, Technopo# le
de Nancy-Brabois, France.
3
Department of Molecular Biology,
Microbiology Section, University of Siena,
Italy.
Author for correspondence : Danielle
Venditti. Tel : j33 3 83 44 12 10. Fax : j33 3
83 44 02 61.
e-mail : tredi.recherche!wanadoo.fr
344
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