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FEMS Microbiology Ecology 54 (2005) 123–130
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Root cap influences root colonisation by Pseudomonas fluorescens
SBW25 on maize
Sonia N. Humphris a,*, A. Glyn Bengough a, Bryan S. Griffiths a, Ken Kilham b,
Sheena Rodger c, Vicky Stubbs a, Tracy A. Valentine a, Iain M. Young c
a
Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, UK
School of Biological Sciences, University of Aberdeen, Aberdeen AB24 2TZ, UK
Scottish Informatics Mathematics Biology and Statistics (SIMBIOS) Centre, University of Abertay, Bell Street, Dundee DD1 1HG, UK
b
c
Received 13 August 2004; received in revised form 28 January 2005; accepted 11 March 2005
First published online 1 April 2005
Abstract
We investigated the influence of root border cells on the colonisation of seedling Zea mays roots by Pseudomonas fluorescens
SBW25 in sandy loam soil packed at two dry bulk densities. Numbers of colony forming units (CFU) were counted on sequential
sections of root for intact and decapped inoculated roots grown in loose (1.0 mg m 3) and compacted (1.3 mg m 3) soil. After two
days of root growth, the numbers of P. fluorescens (CFU cm 1) were highest on the section of root just below the seed with progressively fewer bacteria near the tip, irrespective of density. The decapped roots had significantly more colonies of P. fluorescens
at the tip compared with the intact roots: approximately 100-fold more in the loose and 30-fold more in the compact soil. In addition, confocal images of the root tips grown in agar showed that P. fluorescens could only be detected on the tips of the decapped
roots. These results indicated that border cells, and their associated mucilage, prevented complete colonization of the root tip by the
biocontrol agent P. fluorescens, possibly by acting as a disposable surface or sheath around the cap.
2005 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved.
Keywords: Zea mays; Border cell; Pseudomonas fluorescens; Biocontrol; Root cap; Confocal microscopy; Rhizosphere
1. Introduction
Root border cells, and their associated root cap mucilage, are thought to have roles in a number of processes
including aiding root penetration into compacted soils
[1,2], regulating root cap mitosis [3] and in establishing
rhizosphere communities [4]. Recently, research has focused on the role of border cells in protecting plant
health by controlling the growth of microorganisms in
the rhizosphere [5]. Border cells synthesise and export
a diverse range of low molecular weight proteins [6]
*
Corresponding author. Tel.: +44 1382 562731; fax: +44 1382
568502/562426.
E-mail address: [email protected] (S.N. Humphris).
and supply a readily available source of carbon into
the extracellular environment [7], which could influence
the activity of microbial communities in the rhizosphere.
In this way, border cells potentially have the capacity to
promote the growth of beneficial microorganisms and
repel or inhibit the growth of pathogenic organisms
[4,5,8]. The quantity and quality of exudates produced
by border cells depends on the plant genotype. The
microbial communities present in the rhizosphere will
be influenced by the ability of these organisms to respond to and utilise particular compounds released from
border cells [9].
Mechanisms by which border cells protect plant
health have been suggested to include: acting as a decoy
to attract fungal infection away from the vulnerable root
0168-6496/$22.00 2005 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved.
doi:10.1016/j.femsec.2005.03.005
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S.N. Humphris et al. / FEMS Microbiology Ecology 54 (2005) 123–130
tip, thereby leaving the tip free from infection when the
border cells detach [10,11]; attracting and then inducing
a transient quiescent state in root knot nematodes,
allowing time for the root tip to grow past the area of
danger [12]; and producing mucilage that repels pathogenic bacteria [11]. Border cell production has also been
positively correlated with arbuscular mycorrhizal (AM)
colonisation. AM fungi mainly benefit plant growth by
improving immobile nutrient uptake from soil. Plants
producing large numbers of border cells tend to show
greater mycorrhizal root colonisation [13,14].
One group of beneficial rhizosphere microorganisms,
that have received significant attention, is the plant
growth promoting rhizobacteria (PGPR). Some PGPR
can suppress the growth of pathogenic organisms directly by producing either antibiotic compounds [15],
or siderophores that allow effective iron capture from
the rhizosphere, depriving pathogenic organisms of this
necessary element [16]. They can also indirectly promote
plant growth by inducing systemic resistance in the plant
[17,18]. Many PGPR belong to the fluorescent pseudomonads, including Pseudomonas fluorescens, which have
been used successfully to control soil-borne plant pathogens [19]. Fluorescent pseudomonads are also capable of
exploiting a variety of compounds exuded by roots, have
a fast growth rate and are motile, [20,21]. Despite this,
biological control of plant pathogens in the field is often
inconsistent, partly due to poor root colonisation
[19,20], leading to decreased biocontrol activity [22].
Successful biological control requires the density of the
introduced biocontrol bacteria to be at the sufficient level, time, and location favourable for potential pathogenic attack irrespective of environmental conditions
[23]. Soil physical conditions determine the size ranges
of soil pores present and whether they are water-filled.
This is relevant both for organism movement, and root
growth rate (controlled by soil strength and matric potential). These factors will determine the impact of bulk
soil density on the spread and location of biocontrol
bacteria.
The aim of this paper was to determine the influence
of border cells, and their associated mucilage, on the
growth and distribution of P. fluorescens SBW25, when
grown on Zea mays roots, in loose and compacted soil.
Root colonisation by P. fluorescens was determined in
intact and decapped roots at two soil bulk densities.
2. Materials and methods
2.1. Seed sterilisation
Caryopses of Z. mays KX0141 (KWS, Germany)
were surfaced sterilised by soaking in a 10-ppm solution
of oxytetracycline for 20 h at room temperature and
then in 0.1% (w/v) silver nitrate solution for 10 min.
Once drained, the seeds were rinsed in a 0.5 % sodium
chloride solution to precipitate the remaining silver ions
and finally, washed with vigorous shaking in three
changes of sterile distilled water (SDW) [24]. The seeds
were germinated between sheets of moist, sterile filter
paper (Whatman No.1) in 14 cm diameter Petri dishes,
sealed with laboratory film and incubated at 25 C at
an angle of 40 to the vertical.
2.2. Root treatment
After approximately 44 h incubation, when the roots
were between 2.5–3 cm long, the root caps were removed
from 20 seedlings under a stereomicroscope in a flow
cabinet, using a new sterile scalpel blade to lever off
the cap at the junction between the translucent cap
and root. The seedlings were returned to the Petri dishes,
sealed, replaced in the incubator at 25 C at an angle of
40 to the vertical for 4 h, to allow time for the wound to
heal. Seedlings with their cap removed are denoted as
decapped and roots without their caps removed as
intact.
2.3. Root border cell counts
A further 15 roots were decapped and returned to the
Petri dishes along with 15 intact roots, the plates were
sealed and replaced in the incubator at 25 C at an angle
of 40o to the vertical. Border cell counts were performed
in triplicate on 5 intact and 5 decapped roots after 4, 24
and 48 h. Each seedling was placed over a micro-centrifuge tube containing 500 ll of SDW so that the root tip
and approximately 10 mm of root penetrated the water.
The tips were left to hydrate for 20 min, with agitation
of the water using a micropipette after 10 min. Fifty
microlitres of cell suspension was pipetted into a counting dish, stained with 3 ll of Toluidine blue (0.1% in
0.1 M phosphate buffer) and mixed with 500 ll of
SDW. The cells were allowed to settle for 5 min before
counting with a dissecting microscope, as previously described [25].
2.4. Plant growth medium
PVC tubing (220 mm h · 21 mm d), which had been
sawn in half longitudinally, was sterilised by soaking
in 1% bleach overnight. The tubes were rinsed thoroughly in SDW and left to dry in a laminar airflow cabinet. Once dry, the original halves were sealed back
together using insulating tape with plastic lids taped to
one end to provide a base. Ten tubes were packed with
sterile (autoclaved twice, 1 day apart at 121 C for
20 min at 1 atm.) sandy loam soil (sand 56%, silt 36%,
clay 8%) sieved to <2 mm at a dry bulk density of
1 mg m 3 with a gravimetric water content of 22%. A
further 10 tubes were packed at a dry bulk density of
S.N. Humphris et al. / FEMS Microbiology Ecology 54 (2005) 123–130
1.3 mg m 3 with a gravimetric water content of 20%,
this ensured the matric potential would be approximately 10 kPa at both bulk densities. All tubes were
sealed with laboratory film after packing and left at
12 C until the seedlings were ready to be planted.
2.5. Preparation of P. fluorescens expressing GFP
Transformation was carried out using a mating technique between the GFP expressing Escherichia coli s17-1
lambda pir PSM1696 (containing the plasmid pUTkmgfp) and P. fluorescens SBW25. P. fluorescens was
maintained on Pseudomonas selective agar (PSA) plus rifamycin (100 lg ml 1) at 28 C. The E. coli was maintained on LB plus ampicillin and kanamycin
(100 lg ml 1 of each) at 37 C. P. fluorescens and E. coli
were grown up separately in 5 ml of LB overnight and
then 1.5 ml of each culture was spun at 10000g for 5 min
and the supernatant discarded. The resulting pellet was
washed 3 times with 1 ml of 1/10 LB and finally re-suspended in 1 ml 1/10 LB. P. fluorescens (700 ll) and E. coli
(300 ll) were mixed, centrifuged at 10000g for 5 min, the
supernatant discarded and the pellet re-suspended in
1 ml 1/10 LB. Fifty microlitres of the mixed cells was applied to a 0.2 lm 13 mm filter on Tryptone Soya Agar and
incubated for 6 h at 28 C. The filter was removed and
transferred to a microcentrifuge tube containing 1 ml of
1/10 LB, vortexed and a dilution series prepared. One
hundred microlitres of the dilution series was plated onto
PSA plus rifamycin and kanamycin (100 lg ml 1 of each)
and ferrous sulphate (45 mM) and incubated at 28 C.
The resulting transcongjugant were screened for GFP
and cultures from a single positive colony used to inoculate roots. The transcongjugant were checked for stability
following the method in Bailey et al. [26].
2.6. Inoculation of roots with P. fluorescens
Under sterile conditions, the length of 10 decapped
and 10 intact roots was measured and then dipped in
an overnight culture of P. fluorescence SBW25, that
had been previously grown at 25 C (concentration of
approximately 4.4 · 108 CFU ml 1). The roots were left
for 2 min in the P. fluorescens culture and then allowed
to dry on a sterile Petri dish for 5 min in a laminar airflow cabinet. The tubes were removed from the 12 C
incubator and a small hole was made in the soil, using
a sterile mounted needle (1 mm diameter), into which
the roots inoculated with bacteria were planted. The
tubes were sealed with laboratory film and incubated
at 25 C for 44 h. There were five replicates of both
the decapped and intact roots planted at each soil bulk
density. In addition, 5 replicates of both decapped and
intact roots, inoculated with P. fluorescens and 5 replicates, which had not been inoculated, were harvested
immediately (i.e. they were not planted in the soil).
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2.7. Root sampling
After 44 h growth, the maize seedlings were carefully
removed from the tubes by cutting through the electrical
tape sealing the two halves of tube. The roots were shaken gently and any soil adhering tightly to the roots was
included in the harvest. The primary root of each seedling was measured and divided into 4 sections. The
lower 10 mm were removed for assessment of bacteria
on the root tip and referred to as tip section. The top
30 mm (the average length of roots when planted) was
removed to represent the zone of inoculation and referred to as seed section. The remaining root section
was cut in half, giving a section that had been closest
to the tip and the other closest to the seed. The two sections were denoted as next-to-tip and next-to-seed,
respectively (Fig. 1). The four root sections were
chopped into small pieces and placed into sterile micro-centrifuge tubes containing 100 ll of sterile 1/4
strength Ringers (2.25 g l 1 NaCl, 0.105 g l 1 KCl,
0.12 g l 1 CaCl2 Æ 2H2O, 0.05 g l 1 NaHCO3), and
0.1 g of a sterile mixture of coarse and fine grinding
sand. The root pieces were homogenised by grinding
with a micro-pestle and then, after adding 900 ll of 1/4
strength Ringers, the samples were vortexed for 20 s.
A 10-fold dilution series was prepared in sterile 96-well
microtitre plates for each root section down to 10 6
using 1/4 strength Ringers. Ten microlitres of each dilution was pipetted onto plates of Luria Bertani medium
(LB) plus supplements (0.05 mg ml 1 kanamycin,
0.05 mg ml 1 cyclohexamide and 0.125 mg ml 1 ferrous
sulphate). The visualisation of GFP in P. fluorescens is
difficult due to the fluorescent nature of the siderophores
that these bacteria produce. To overcome this problem
iron was added to the growth media in the form of ferrous sulphate. A dilution series was also prepared for
roots sampled immediately before planting and the sterile soil used to pack the tubes. The dilution series was repeated 3 times for all samples and each series was plated
out in triplicate. The LB plates were incubated at 25 C
for 2 days and number of colonies counted.
2.8. Statistical analysis
Mean counts for each dilution series were log
(base10) transformed to normalise the data before using
ANOVA and a general analysis of variance. All data
was analysed using Genstat 6th Edition statistical software (Lawes Agricultural Trust).
2.9. In situ imaging of roots grown on agar
A potential artefact of removing roots from the soil
system is that bacterial cells colonising the root tip mucilage can be left behind in the soil. To determine whether
this was the case and to image the distribution of
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Fig. 1. Colony counts of P. fluorescens on the 4 root sections: tip; next-to-tip; next-to-seed and seed in loose and compacted soil. Results are
expressed as log transformed colony forming units (CFU) per cm of root. Decapped roots have diagonal shading and intact roots are solid colour.
Bars represent standard error of the mean, n = 5. Picture shows sections of an intact root grown in loose soil.
P. fluorescens on the root tips, both intact and decapped
roots inoculated with bacteria were grown on agar. Under sterile conditions, the lengths of 10 intact roots and
10 decapped roots were measured and then dipped in an
overnight culture of P. fluorescens for 2 min and left to
dry on a sterile Petri dish for 5 min. The roots were then
sandwiched between 2 layers of 2 mm thick 2% agar
technical in 14 cm diameter Petri dishes, sealed with laboratory film and incubated at an angle of 40 to the vertical and left to grow at 25 C for 24 h, after which time
the root lengths were measured. They were then transferred to a fridge at 4 C for 24 h to slow root growth.
This allowed observation of bacterial colonies in situ with
a Bio-Rad MRC 1000 confocal laser-scanning microscope (Bio-Rad, Hercules CA) attached to an Optiphot
II microscope (Nikon, Tokyo) using either 10·, (10/
0.25 160/- ELWD) or 40· (40/0.55 160/0-2.5 ELWD)
lenses. Excitation: 100 mW Argon laser at 488 nm, Emission filter: 522DF 32 nm. The roots were then removed
from the agar, mounted in H2O under a coverslip, and
viewed using 60· (Plan 60 /1.4 Oil 160/0.17) lens.
pacted soil (Table 1). There were no significant
differences in the elongation rates of the intact and decapped roots grown on agar (Table 1). The three time
points for the border cell counts were chosen to represent the numbers of border cells on the maize roots during the P. fluorescens soil experiment. For example, 4 h
represented when the roots inoculated with bacteria
were planted in the soil of the growing tubes and 48 h
represented, the border cell counts at the end of the
experiment, before the P. fluorescens was harvested.
Border cell counts revealed significantly more
(P < 0.05) border cells on the intact roots compared
with the decapped roots at all 3 time points (Table 2).
Table 1
Elongation rates for intact and decapped roots inoculated with
P. fluorescens and grown in loose (1.0 mg m 3) or compact
(1.3 mg m 3) soil for 44 h at 25 C and sandwiched on agar for 24 h
at 25 C and 24 h at 4 C
Elongation rate (mm h 1)
Treatment
1.0 mg m
1.3 mg m
Agar
3
3
soil
soil
Intact
Decapped
2.1 ± 0.07
1.6 ± 0.09
2.0 ± 0.05
0.9 ± 0.11
0.6 ± 0.03
1.9 ± 0.13
3. Results
Mean ± standard error of the mean, n = 5.
3.1. Root growth and border cell counts
Table 2
Border cell counts for intact and decapped roots 4, 24 and 48 h after
decapping
At both soil densities, intact roots inoculated with P.
fluorescens grew significantly faster (P < 0.05) than the
decapped roots inoculated with P. fluorescens. There
was no significant difference in root growth of the decapped roots between the soil densities, but the growth
of the intact roots in the loose soil was significantly faster (P < 0.05) than the intact roots grown in the com-
Time (h)
4
24
48
Border cells per root
Intact
Decapped
1927 ± 130
4100 ± 590
1745 ± 226
775 ± 103
325 ± 102
309 ± 124
Mean ± standard error of the mean, n = 5.
S.N. Humphris et al. / FEMS Microbiology Ecology 54 (2005) 123–130
127
Fig. 2. Confocal images of the root tips of: uninoculated intact (a) and decapped (b) controls at 10· magnifications. Inoculated intact (c) and
decapped (d) roots at 10· magnification. Inoculated intact (e) and decapped (f) roots at 40· magnification. Bars represent 100 lm.
3.2. Density of P. fluorescens along the root sections
There was no significant difference in the inoculation
density of P. fluorescens between the whole and dec-
Fig. 3. Confocal image of bacterial colonies on the root tip of an
inoculated decapped root at 60· magnification. Bar represents 10 lm.
apped roots immediately after being inoculated, having
3 · 105 CFU ml 1 and 2.7 · 105 CFU ml 1, respectively.
The data analysis of the colony counts revealed that
there was a significant interaction (P < 0.001) between
the treatments and root sections, indicating that the distribution of bacteria between the root sections was affected by the treatment (Fig. 1). There were significant
differences in the number of bacteria on the majority
of different root sections, with most bacteria on the seed
section of the root and progressively fewer bacteria near
the tip. The numbers of bacteria (CFU cm 1) on all 4
sections of the intact roots grown in loose soil were significantly different from each other (P < 0.05). There
was no significant difference (P > 0.05) between the P.
fluorescens density on the seed and next-to-seed sections
for the other 3 treatments and the density on the nextto-seed and next-to-tip for the decapped roots grown
in loose soil.
Analysis of the P. fluorescens densities between the
treatments within an individual section, demonstrated
that there were no significant differences in numbers of
bacteria on the seed sections after 44 h growth in the
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S.N. Humphris et al. / FEMS Microbiology Ecology 54 (2005) 123–130
soil. At the next-to-seed section, the intact and decapped
roots grown in the loose soil had significantly fewer colonies (P < 0.01 and <0.05 respectively) of P. fluorescens
than the intact and decapped roots grown in the compacted soil. The intact roots grown in the loose soil
had significantly less (P < 0.01) P. fluorescens than the
other 3 treatments at both the next-to-tip and tip section. In fact, the tip section of the intact roots grown
in both the loose and compacted soil had significantly
fewer colonies (P < 0.05) of P. fluorescens than the 2
decapped root treatments.
No bacterial colonies were observed on the sterile soil
that was used to pack the growing tubes or the sampled
uninoculated roots.
3.3. Distribution of P. fluorescens along the root sections
Comparison of the confocal images of the decapped
roots inoculated and grown for 1 day on agar (Fig.
2(d)) with the uninoculated decapped roots (Fig. 2(b)),
taken at the same gain levels, revealed that the increased
brightness caused by the GFP genes within the P. fluorescens can be clearly seen on the decapped inoculated
root. P. fluorescens is covering the very tip of the inoculated decapped root (arrow 1, Fig. 2(d)), the newly exposed re-growing cap (arrow 2, Fig. 2(d)) and streaks
just behind where the root cap was excised (arrow 3,
Fig. 2(d)). Comparable decapped and non-decapped
images at 10· magnifications showed a background level
of auto fluorescence from the root. However at 40·
magnification (Fig. 2(f)), the auto fluorescence becomes
less visible due to the difference in gain used for these
images and the thinner optical sections used. Only in
the inoculated decapped root is fluorescence from the
bacteria visible. The only fluorescence visible at the
higher magnification was on the tips of the decapped
roots inoculated with bacteria (Fig. 2(f)). By removing
the inoculated decapped roots from the agar and viewing at 60 · magnification individual bacterial colonies
could be identified (Fig. 3). This confirmed that the
areas of increased brightness were indeed caused by
the P. fluorescens. Removing the roots from between
the 2 layers of agar however, potentially disturbs the
bacterial distribution, therefore only the roots left in situ
could be used to visualise the distribution of P. fluorescens. The small areas of brighter fluorescence on the
uninoculated decapped root (Fig. 2(b)) are likely to be
dead or dying border cells, which have remained after
de-capping. The comparison of the uninoculated intact
roots (Fig. 2(a)) with the inoculated intact roots (Fig.
2(c)) reveals very little or no differences between the 2
treatments. No fluorescence could be seen at the higher
magnification for either treatment (Fig. 2(e)). Again, it is
likely that the small areas of brighter fluorescence on
both the control and inoculated intact root are dead
or dying border cells.
The fast elongation rate (>30 lm min 1) of the roots
incubated at 25 C for 48 h resulted in confocal images
that were blurred. However, comparison of these images
with the images of the roots grown at 25 C for 24 h and
then 4 C for 24 h, demonstrated that the distribution of
P. fluorescens remained the same (observations not
shown), with the slower rate of growth at 4 C allowing
rather sharper images.
4. Discussion
There were significant differences between the density
(CFU cm 1) of bacteria on the different root sections,
with the highest density of P. fluorescens located on
the seed section of the root and progressively fewer bacteria near the tip. This is presumably because the bacteria have to move to and colonise the rhizosphere and
rhizoplane around the new root tissue, resulting in fewer
bacteria adjacent to young tissue. The distribution of
bacteria between root sections was significantly affected
by decapping and by soil density during the 2 days of
growth in the soil. Decapping of a maize root leads to
the stimulation of cell division in the quiescent centre
and results in the regeneration of a new cap, usually in
3–4 days [27]. The results of the border cell counts show
that the regeneration of the root cap was not complete
on the decapped roots after 2 days, indicating that there
was no border cell production on the decapped roots to
prevent the colonisation of P. fluorescens at the very tip
of the root. Therefore it can be assumed that the increased colonisation of the decapped root tip is associated with decreased production of border cells and
mucilage from the root tip.
Differences in Pseudomonas densities at the tip and
next-to-tip section can be related to both root cap and
soil density treatments. The intact roots have significantly fewer bacteria at the tip section compared to
the decapped roots in both loose and compact soil. In
addition, in situ images of the root tips grown in agar,
clearly show that no P. fluorescens can be detected on
or around the tips of the intact roots (Fig. 2(c) and
(e)). This mirrors what was seen when the roots were
grown in soil and shows that the lower density of P. fluorescens on the intact roots was not simply due to bacteria on the root tip being ‘‘left behind’’ in the soil
when the root was sampled. The lower colony counts
on the tip section of the intact roots are probably caused
by the abundant border cells and mucilage associated
with the root cap preventing complete colonisation of
the intact root tip by P. fluorescens. Border cells and
mucilage may form a disposable surface around the root
tip that prevents colonisation by the bacteria.
At the next-to-seed section, the density of P. fluorescens was significantly greater on the roots grown in the
compact soil compared with the roots grown in the loose
S.N. Humphris et al. / FEMS Microbiology Ecology 54 (2005) 123–130
soil. As soil density increases, there are also increases in
root diameter [28], resulting in a larger surface area
available, per unit root length, for colonisation by P. fluorescens. This increase in surface area however is not enough to be solely responsible for the large differences
(e.g. 100-fold) in numbers of P. fluorescens. Nor can it
account for the larger numbers of P. fluorescens on the
tip and next-to-tip section of the decapped roots grown
in the loose soil. Compaction may affect bacterial distribution by decreasing root elongation rate, physically
abrading the root surface, changing border cell and
mucilage production or by causing changes to exudation
patterns.
P. fluorescens populations located on the seed section
were greater than on other root sections and did not differ significantly between soil density and decapping
treatment. This is because the seed section represents
the tissue inoculated initially and on which the bacteria
have had most time to multiply. During the early stages
of growth, (first 10 days) of P. fluorescens coated seedlings of various crops, the highest concentrations of bacteria are located on the area of root just below the seed,
with a rapid reduction in bacterial density down the root
length to the tip [29–31]. Motility is thought to be play
an essential role in the colonisation of roots by Pseudomonas [32], but more specifically motility based on chemotaxis towards exudate compounds [33]. Areas such as
the root hair zone, root tip and emergence of lateral
roots produce exudates that differ in quantity and quality [34], which undoubtedly will influence the distribution of Pseudomonas. Indeed these exudation patterns
change with both location on the root, plant age [35]
and increases in root diameter caused by increased compaction [36]. During the early stages of root growth, a
high concentration of exudates is found in the zone closest to the seed, probably due to seed storage compounds
[37], which may in part explain the high numbers of P.
fluorescens at the seed section. It is likely that distribution of P. fluorescens along the root would continue to
change with time in accordance with exudate patterns
and soil physical conditions. For example, Gamalero
et al. [38] found that colonisation of tomato roots by
P. fluorescens (seed inoculated) changed in time, with a
significant reduction in bacterial density at the root tip
of 7-day-old roots compared with 3-day-old roots.
Poor colonisation of bacteria down the length of
roots is often linked to the fast growth rate of roots,
which overcomes bacterial growth [39]. However, while
the intact roots were significantly longer (P < 0.05) than
the decapped roots after 2 days growth in soil (but not
before planting), root length was not a statistically significant factor in the bacterial distribution. The elongation rates of the intact and decapped roots when grown
on the agar were very similar and not significantly different from each other, and yet the decapped root tip was
clearly colonised by the bacteria (Fig. 2). Decapping a
129
root eliminates the gravitropic response but does not alter the growth in length [40]. The decapped roots generally deviated from the vertical much more than the
intact roots (unpublished observation). These facts indicate that the soil density was responsible for the differences seen in the lengths of the intact and decapped
roots during this experiment.
This experiment was conducted in sterile soil, so the
influence of other microorganisms on the growth and
distribution of P. fluorescens is unknown. In addition,
to confer biocontrol of pathogens it is not known
whether the P. fluorescens must colonise the entire root
tip, as root tip protection by the border cells and protection immediately behind the root tip by the P. fluorescens may be adequate. In conclusion, while border
cells may protect plant health by preventing immediate
infection of the root tip by pathogens [5], in this experiment they also prevented complete colonisation of the
root tip by the biocontrol agent P. fluorescens. Colony
numbers of P. fluorescens on the intact root tips were
not as great as the colonies on the decapped root tips,
which had no border cells to prevent the distribution
of P. fluorescens. Increased soil density caused a significant increase in the colony numbers of P. fluorescens on
the tip of the intact roots due to a combination of
changes in root elongation rate and border cell and
mucilage production. We conclude that the production
of border cells and mucilage may prevent root tip colonisation by acting as a disposable surface or sheath
around the cap.
Acknowledgements
We thank Drs. Oliver Knox and Dominic Standing
for help and advice on the Pseudomonas culture, and
Dr. D. Cooke for helpful discussions. We acknowledge
funding from the Biotechnology and Biological Research Council. The Scottish Crop Research Institute receives grant-in-aid from the Scottish Executive Rural
Affairs Department.
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