Colonization pattern of primary tomato roots by Pseudomonas fluorescens A6RI characterized by dilution plating, flow cytometry, fluorescence, confocal and scanning electron microscopy

FEMS Microbiology Ecology 48 (2004) 79–87 www.fems-microbiology.org

Colonization pattern of primary tomato roots by Pseudomonas fluorescens A6RI characterized by dilution plating, flow cytometry, fluorescence, confocal and scanning electron microscopy Elisa Gamalero

a,*

, Guido Lingua a, Flavia Giusy Capr
ı a, Anna Fusconi b, Graziella Berta a, Philippe Lemanceau c

a

Department of Science and Advanced Technology, Universit
a del Piemonte Orientale ‘‘Amedeo Avogadro’’, Alessandria 15100, Italy b Department of Plant Biology, Universit
a di Torino, Torino 10125, Italy c UMR INRA/Universite de Bourgogne, ‘Microbiologie et Geochimie des Sols’, INRA-CMSE, BP 86510, Dijon cedex 21065, France Received 25 August 2003; received in revised form 1 December 2003; accepted 19 December 2003 First published online 30 January 2004

Abstract Early colonization of primary tomato roots, grown in vitro, by Pseudomonas fluorescens A6RI, introduced by seed bacterization, was monitored for 7 days in three different root zones (zone A, apex + elongation + young hairy zone; zone B, hairy zone; zone C, old hairy zone + collar). Bacterial quantification was assessed by enumeration of (i) colony forming units (cfu) after dilution plating and of (ii) total bacterial cells by flow cytometry. Bacterial distribution and organization in the root zones were analyzed by fluorescence, confocal and scanning electron microscopy. For all sampling dates and zones, the densities of total bacterial cells were significantly higher than those of the cfu. The kinetics of cfu densities varied according to the root zone. Their density decreased with time in zone A, while no variation with time was recorded in zones B and C. Densities of total bacterial cells did not show any significant temporal variation for any of the root zones. Microscopic analyses allowed the characterization of the distribution and organizational patterns of the bacterial cells according to time and space. In 3-day-old plants, bacteria were mostly present as single cells and were evenly distributed in the two root zones analyzed (A and B). In 5- and 7-day-old plants, distribution and organization differed according to the root zone. In zone A, only few single cells were observed, whereas zones B and C were mostly covered by cells localized between epidermal root cells and organized in pairs and strings, respectively. Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. Keywords: Pseudomonas fluorescens; Root colonization; Dilution-plating; Flow cytometry; Immunolocalization; Electronic scanning microscopy

1. Introduction Fluorescent pseudomonads are considered as potential biocontrol agents of soilborne diseases [1]. Although successful biocontrol has been reported even in semicommercial conditions, positive effects are often inconsistent [1,2]. This lack of reliability has been at least partly ascribed to impaired root colonization by the introduced biocontrol strain [3]. Indeed, a clear relationship has been *

Corresponding author. Tel.: +39-0-131-283838; fax: +39-0-131254410. E-mail address: [email protected] (E. Gamalero).

established between suppression of the wheat root disease take-all and that of fusarium wilts by different strains of fluorescent pseudomonads and the densities of these bacteria in the rhizosphere of the corresponding host plant [4,5]. There is then a need for better knowledge of bacterial traits promoting rhizosphere competence of fluorescent pseudomonads. Various studies have been dedicated to the identification of these traits [6–10]. In these studies, microbial quantifications were performed, at different dates, on the whole root system or on root fractions without taking in account the spatial variation related to the morpho-physiological characteristics of the different root zones. However, variations in these

0168-6496/$22.00 Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. doi:10.1016/j.femsec.2003.12.012

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characteristics might affect the root exudates as mirrored by differences in microbial communities along the roots [11,12]. Therefore, root colonization by introduced bacterial strains is expected to differ according to the different root zones. The localization of bacteria on the root is also likely to play an important role in pathogen suppression. Indeed, where bacterial antagonism is mediated by secondary metabolites, such as antibiotics and siderophores [13,14], its expression would require the presence of the biocontrol agent where the pathogen is active and penetrates the root. When disease suppression results from triggering the defense reactions of the host-plant, this requirement does not stand [15,16]. However, since the induced systemic resistance has been at least partly ascribed to membrane properties of the bacterial elicitor [17], a close contact of the bacteria with the root cells of the host-plant is thought to be needed. Attention has been given to patterns of root colonization by various fluorescent pseudomonad strains in the rhizosphere of different host-plants [18–22]. In most studies, bacterial densities were quantified by dilution plating [23], in others the distribution patterns were characterized by electronic [19] or confocal [18,21,22] microscopical observations. However, a combination of bacterial quantification and colonization pattern description has rarely been achieved [20,24,25] and, even in these studies, data were not related to morphologically different root zones. The aim of the present work was to study the spatiotemporal colonization of primary tomato roots by an introduced model strain of Pseudomonas fluorescens (A6RI). Densities of this model strain in different root zones were assessed both by dilution plating and flow cytometry, and distribution and organization of the bacterial cells were characterized by fluorescence, confocal and scanning electron microscopic observations.

2. Materials and methods 2.1. Bacterial inoculant Pseudomonas fluorescens A6RI is a rifampicin resistant mutant of the strain CFBP2392 [26], which was shown to suppress damping-off [27], root rot (Berta et al., unpublished data), and to promote plant growth [28]. Bacterial cultures, grown in Luria–Bertani broth medium [29] supplemented with rifampicin (100 lg ml
1 ), were stored at )80 °C in 50% glycerol. P. fluorescens A6RI inoculant was produced on solid King B medium [30] plates at 28 °C for 48 h. Bacteria were scraped from the medium and suspended in 0.1 M MgSO4  7H2 O, pelletted by centrifugation (5000 rpm, 20 min), washed twice and suspended in 0.1 M MgSO4  7H2 O. The bacterial density of the suspension

was determined using a calibration curve assessed by turbidity (k ¼ 600 nm) and adjusted to 107 colony forming units per ml (cfu ml
1 ). 2.2. Plant growth conditions and root sampling Tomato seeds (Lycopersicon esculentum Mill. cv. Early Mech) were surface sterilized by gentle shaking for 3 min in a solution of 5% sodium hypochlorite. The sterilized seeds were then washed six times for 5 min and four times for 20 min in sterile demineralized water. The seeds were placed in Petri dishes on moist sterile filter paper and incubated in the dark at 24 °C for 5 days. Sterile germinated tomato seeds were bacterized by dipping them for 20 min in a suspension containing 107 cfu ml
1 of P. fluorescens A6RI. Bacterized germinated seeds were aseptically placed on a 15-cm Petri dish containing agarized (1.1%) modified Long Ashton nutrient solution [31]. The Petri dishes were placed at an angle of 45° and incubated in a growth chamber (16/8 h light dark photoperiod and 24°/20 °C thermoperiod, 150 lE m
2 s
1 ) with 60% of relative humidity. The lower half of the Petri dishes was wrapped with aluminium foil in order to prevent the light to reach the roots. Primary roots were gently dragged from the soft agar and sampled 3, 5 and 7 days after bacterial inoculation on six plants per date in order to study their colonization by bacteria adhering to the roots. For bacterial enumerations, and electron scanning microscopic observations, primary tomato roots were aseptically cut in 1-cm long pieces from the tip to the upper part of the root, meaning that the last piece could be less than 1 cm long. These root pieces were then classified according to morphological observations made with a Leica stereo microscope into three zones including: (i) the apex, the elongation and the young hairy zones (zone A), (ii) the hairy zone (zone B) and (iii) the old hairy and the collar zones (zone C). For fluorescent and confocal laser scanning microscopic observations, the whole primary roots were sampled and the root zones were identified, before processing the roots. 2.3. Enumeration of bacterial colony forming units and cells Bacterial enumerations were performed both by dilution plating and by flow cytometry. For each date and plant replicate, root pieces belonging to a given root zone were pooled, suspended and vortexed for 10 min in 3 ml of 0.1 M MgSO4  7H2 O in order to remove adhering bacteria. Dilution plating and flow cytometry analyses were performed on the same suspension for each sampling date and plant replicate. The suspensions obtained were serially diluted and plated on solid KB supplemented with rifampicin

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(100 lg ml
1 ). After incubation for 48 h at 25 °C, the numbers of cfu were determined. Five hundred microliters of each suspension were added to 1 ml of ethanol and maintained at 4 °C. Bacteria in these suspensions were pelletted by centrifugation (6000 rpm, 20 min) to eliminate ethanol and then resuspended in 0.1 M MgSO4  7H2 O to a density calculated to be 105 cfu ml
1 according to the previous cfu density measurements. These suspensions were stained with 50 lg ml
1 propidium iodide and finally analyzed with a PAS flow cytometer (Partec GmbH, M€ unster, Germany). Excitation wavelength at 488 nm was provided by an argon laser. Red light emission (k > 610 nm) side and forward scatter were measured. All the parameters were acquired with logarithmic amplification and analyzed using the Flow Max software (Partec). The volume count option of the instrument was used, in order to measure the bacteria densities. Since the size of the root zones varied with plant replicates and age, bacterial densities were expressed per centimeter of root to allow comparisons. Density values of cfu and cells were logarithmically transformed before analysis because bacterial populations approximate an exponential normal distribution [32]. Transformed values were submitted to analysis of variance and then FisherÕs least significant test (P ¼ 0:05) using Statview statistic package (SAS Institute Inc., Cary, NC).

and sealed with transparent nail polish to prevent dehydration of the roots. The distribution and organization of P. fluorescens A6RI cells along the root were examined in situ by microscopic observations. Observations with epifluorescence microscopy were performed with an Axioscope (Zeiss-Oberkochen, Germany) equipped with FT510 and FT395 filters with excitation wavelengths of 450– 490 and 365 nm, respectively. Oil immersion lenses 40 and 100 were used and micrographs were taken on a Kodak Ektachrome P1600 color reversal film with a MC 80 microscope camera. Observations with CLSM were performed with a Zeiss LSM 510 laser scanning microscope that is composed of an Axiovert 100 M inverted microscope equipped with two photomultipliers for fluorescence and two lasers (Ar 15 mW and HeNe 0.5 mW) having excitation wavelengths of 458–488 and 543 nm, respectively. Observations were performed using the 488 nm Ar 15 mW laser. In addition to the fluorescent light, the microscope could simultaneously record the light transmitted through the specimen by a third photomultiplier. Oil immersion lenses Plan neofluor 40 (1.3 NA), 63 (1.4 NA) and 100 (1.4 NA) were used. The CLSM was digitally controlled and the image analyzed with a PentiumTM PC and the 32-bit Windows NT 4.0 operating system using a software package (LSM 510 software version 2.01SP 2) provided by Zeiss.

2.4. Fluorescent and confocal laser scanning microscopy

2.5. Electron scanning microscopy

Bacterial distribution on primary tomato roots was studied by indirect immunolocalization using fluorescent and confocal laser scanning microscopy (CLSM). Primary antibodies against P. fluorescens A6RI were raised in a rabbit immunized with heat-inactived (100 °C for 2 h) cells of A6RI. For this, heat-inactivated cells were mixed with an equal volume of incomplete FreundÕs adjuvant. The serum was raised by six sub-cutaneous injections of the suspension given at 3-week intervals. The final booster was given 9 days before recovery of the serum. The whole primary root of each replicate and date was fixed in ethanol–acetic acid 3:1 for 1 h at room temperature. The roots were then placed on separate slides previously treated with acetone for 15 min. The root samples were then incubated, for 45 min, in 100 ll of primary antibodies diluted (1:80) in sterile PBS supplemented with 1.5% BSA and 0.1% sodium azide. The roots were washed three times with this modified PBS buffer. The root samples were then incubated in 100 ll FITC conjugated goat anti-rabbit IgG secondary antibody (Chemicon International, Inc., Temecula, CA) diluted (1:400) in the sterile modified PBS. Incubation was performed in a moist chamber for 1 h at room temperature in the dark. The root samples were finally washed three times with this buffer, mounted in water

One-centimeter root pieces were fixed in 3% glutaraldehyde in distilled water for 3 h and rinsed three times in distilled water. The root pieces were then processed with the PATOTO method [33] consisting of oxidization of the vicinal diols with 1% periodic acid for 15 min, complexing of the resulting aldehyde groups with 1% thiocarbohydrazide (TCH) for 1 h. TCH in turn was coupled to osmium tetroxyde (1%, for 2 h). The latter two treatments were repeated twice. The process was carried at room temperature, all chemicals were dissolved in distilled water and the samples were thoroughly washed with distilled water after every step. The samples were then dehydrated in an ethanol series, transferred to propilene oxyde, air-dried, mounted on aluminium stubs with ScotchTM double-sided tape, coated with gold in a sputtering Hummer II (Technics, Springfield, VA) and examined in a Cambridge S360 Scanning Electron Microscope.

3. Results 3.1. Primary root growth and differentiation The average length of primary root increased with time. This increase was more important from the 3rd

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Table 1 Average length (mm  SE) of the primary roots and of the root zones B (hairy zone) and C (collar zone), 3, 5 and 7 days after bacterial inoculation

3 days 5 days 7 days

19.00  2.82 46.75  7.92 49.50  3.80

Zone B 9.00  2.71 32.63  7.45 32.75  3.20

Due to the sampling procedure, the root zone A (apex + elongation + young hairy zone) was 10-mm long for the three dates.

7 days a

9

Zone C – 4.13  0.66 6.75  1.44

5 days

a a a a

a

log cells/cm

Primary root

3 days

10

a

a

8 7 6 5

A

B

(a)

C

root zone 10 3 days

5 days

7 days

9

log cfu/cm

and the 5th day than from the 5th and 7th day (Table 1). The average size of the different root zones is given in Table 1. Due to the sampling procedure, zone A (apex + elongation + young hairy zone) was always 10mm long. Morphological observations of the primary root showed that the starting point of the hairy zone became more distant from the root tip as the plant became older (1.27  0.13, 1.50  0.20 and 2.00  0.41 mm, for the 3rd, 5th and 7th day, respectively). In contrast, the beginning of the differentiated zone (expressed as appearance of the first xylem element) was closer to the root tip with time (1.74  0.22, 1.50  0.20 and 0.74  0.12 mm, for the 3rd, 5th and 7th day, respectively). Therefore, within zone A, the average size of apex + elongation zone decreased as the plants grew older. The size of zone B (hairy zone) increased between the 3rd and 5th day. Zone C (collar zone) was only observed in 5- and 7-day-old plants.

8 a

7

a

a a 6

b

a

a

b

5

3.2. Temporal and spatial variations of bacterial densities For all sampling dates and root zones, densities of bacterial cells assessed by flow cytometry were always significantly higher (in a range of 2 log orders) than cfu densities assessed by dilution plating. Temporal changes in bacterial densities according to the root zones are shown in Fig. 1. Cell densities did not vary significantly within each root zone (Fig. 1(a)). In contrast, cfu densities significantly decreased with time in zone A, while no significant differences were recorded in zones B and C (Fig. 1(b)). Consequently, the ratio cfu vs cell densities expressed as percentage decreased significantly between the 3rd and the 5th day in zone A (1.10, 0.68, respectively). Spatial changes in bacterial densities were most obvious on 7-day-old plants. Total cell densities (log per cm) were significantly higher in zone C (9.08) than in zones B (8.25) and A (7.81), between which no significant differences were recorded. A similar pattern was observed for the cfu, their densities (log per cm) being not significantly different in zones A (5.43) and B (5.71) but significantly higher in zone C (6.82). The ratio cfu vs cells expressed as percentage was significantly higher in zone C (0.61) than in zone B (0.29), indicating that in

A (b)

B

C

root zone

Fig. 1. Comparison of bacterial densities 3, 5 and 7 days after inoculation in the different root zones. Bacterial densities were assessed by flow cytometry (a) and dilution plating (b). For a given root zone, bars with the same letter are not significantly different (P < 0:05).

7-day-old plants the proportion of culturable bacteria was significantly higher in root zone C than in zone B. 3.3. Localization and organization of bacterial cells along the root Just after bacterial inoculation, bacteria were randomly distributed as single cells along the whole primary root. Three days after microbial inoculation, bacterial cells were distributed both in apex + elongation zone (Fig. 2(a)) and in zone B (Fig. 2(b)). Five days after inoculation, bacterial cells were mostly observed in zones B and C; they were organized in pairs and located closely to the longitudinal junctions between epidermis cell walls (Figs. 3(b) and (c)); few bacterial cells were detected in the apex + elongation zone (Fig. 3(a)). Seven days after inoculation, hardly any bacteria were found

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Fig. 2. Scanning electron and fluorescence microscopy images of primary roots colonized by P. fluorescens A6RI, 3 days after inoculation. Scanning electron microscopy (a) and fluorescence microscopy (b) images show bacterial cells in zones A and B, respectively. Bacterial cells are indicated by arrows.

directly in the apex (Fig. 4(a)), while numerous bacterial cells were present in zones B and C forming strings that followed the longitudinal root cell walls (Fig. 4(b)). SEM observations further showed numerous bacterial cells in zone C (Fig. 4(c)). Numerous bacterial cells were detected on the root hair for all sampling dates. They were homogeneously distributed and were clustered (Figs. 5(b) and (c)) or not (Fig. 5(a)).

4. Discussion Root colonization by biocontrol agents and more specifically by fluorescent pseudomonads is critical for the expression of their beneficial effects. Consequently, numerous studies, aimed at improving our knowledge of

Fig. 3. Scanning electron and fluorescence microscopy images of primary roots colonized by P. fluorescens A6RI, 5 days after inoculation. Fluorescence microscopy images (a, b) show bacterial cells in zones A and B, respectively. Scanning electron microscopy image (c) shows bacterial cells on the collar C. Bacterial cells are indicated by arrows.

bacterial traits promoting rhizosphere competence, have been performed over the last years [6–10]. Studies on survival kinetics of introduced bacteria have often been based on the quantification of culturable bacteria on the whole root system or on portions of the root system sampled at random. However, the survival kinetics of introduced bacteria is a dynamic process resulting both from root growth and differentiation, and from bacterial division and death. Therefore,

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Fig. 4. Confocal laser scanning and scanning electron microscopy (CLSM) of primary roots colonized by P. fluorescens A6RI, 7 days after inoculation. CLSM images show a root apex (green is related to the sample autofluorescence) with no bacterial cell (a) and numerous bacterial cells evenly distributed in zone C (b). Photo A was obtained using a LP 505 filter, giving red color, and a HFT 488–543 dichroic mirror, and photo B resulted from a vertical projection of a z-stack. Scanning electron microscopy image (c) shows numerous bacterial cells evenly distributed in zone C. Bacterial cells are indicated by arrows.

Fig. 5. Fluorescence, confocal laser scanning microscopy and scanning electron microscopy images of root hairs colonized by P. fluorescens A6RI, 3, 5 and 7 days after inoculation. Fluorescence microscopy image shows single bacterial cells along a root hair of 3-days-old tomato plant (a). CLSM image shows the presence of bacterial cells in cluster randomly distributed along root hairs of 5-days-old tomato plant (b). Photo B was obtained using a LP 505 filter giving red color and a HFT 488–543 dichroic mirror. Scanning electron microscopy image (c) shows bacterial cells in clusters. Bacterial cells are indicated by arrows.

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bacterial distribution along a root system is expected to vary according to space and time. Densities of P. fluorescens A6RI were quantified by cfu (dilution plating) and cell (flow cytometry) enumerations. Enumeration of cfu provided information on the culturable bacteria, whereas numeration by flow cytometry provided information on the total cell number without discrimination between alive and dead cells. The densities of cfu were always considerable lower than those of total bacterial cells. Possible bias, due to differences in the dispersion of the cells when suspending them in water prior to plating and when storing them in ethanol prior flow cytometry analysis, cannot be ruled out. However, such possible bias would only account partly to the very significant differences recorded. Hence, a significant proportion of bacterial cells present in the rhizosphere was not culturable. The non-culturable state has been proposed as being a survival strategy [34], although this has been subject to controversy [6,35]. In agreement with previous works [20,36,37], the demonstration, made in the present study, that most bacteria in the rhizosphere are non-culturable, stresses the importance of taking this physiological state in account when assessing the autoecology of fluorescent pseudomonads introduced into the rhizosphere. A combination of bacterial quantification and these microscopic observations gave us more insight on the colonization pattern of P. fluorescens A6RI. Data obtained indicate that the evolution of the spatial pattern of root colonization was related to the differentiation of the root zones. This pattern changed as the primary root became older and as the root zones were more clearly differentiated. In 3-day-old plants, the bacterial cells were evenly distributed along the root, but for the older root differences were seen between the root zones, these differences being the most significant in 7-day-old plants. Within zone A (apex + elongation + young hairy zones), the length of the apex + elongation zone decreased with time, this decrease being related both to reduced activity and physiological changes of this zone [38]. These changes are associated with a significant decrease in the densities of culturable bacteria whereas the densities of total cells did not change, in such way that the proportion of culturable cells decreased significantly in this zone. As the plants grew older, the differentiation zone became longer and closer to the root tip, while the hairy zone, mostly included in zone B, became located further from the root tip. In contrast to zone A, the proportion of culturable bacteria in zone B did not decrease with time suggesting that their activity did not decrease either. This hypothesis was supported by microscopic observations showing that the cells were mostly in pairs in 5-day-old plants and in strings in 7-day-old plants, which we take to mean that the bacteria were dividing. These dividing cells were mostly present closely to the

85

junction between root epidermal cells. Although, the organization of the cells by pairs could be seen by fluorescent microscopy, their presence and localization were especially clear with the higher magnification and three dimension view obtained by SEM. Zone C, including the old hairy zone, was only present in the oldest primary roots (5 and 7 days old). In this zone, the densities of total and culturable bacteria increased with time although not significantly. The cells were organized in strings, following the longitudinal epidermal cell walls of the root, indicating that they were dividing and then active. As a consequence of these divisions, in 7-day-old plants, the densities of both total and culturable bacteria were significantly higher than on the other root zones. The description of the proliferation of individual cells that developed into strings or microcolonies at specific and favorable sites, such as the junction between epidermal cells, the branching point and the root collar, is consistent with the report of ChinA-Woeng et al. [19]. The cell division was further illustrated by the calculation of the total cells (log) per root which significantly increased between the 3rd (8.44) and the 7th (9.12) days. In contrast with studies [18,19,22] made with other introduced strains, root hairs, in zones B and C, were showed to be colonized by clusters of bacteria as indicated by microscopic observations. The gradient of colonization along the root by P. fluorescens A6RI is consistent with previous reports quantifying culturable bacteria [23] and active bacteria by the use of lux reporter gene [24,39–41]. However, in these studies root zones were not differentiated by morphology. Variations in the bacterial densities along the root have been assumed to be related to patterns of exudate concentrations [42], and the proportion of culturable bacteria along the root is expected to be affected by the concentration and composition of root exudates. Indeed, environmental stresses including nutrient limitations would account for variations in bacterial physiology leading to a non-culturable state [43]. Data on root exudation concern mostly the total root system [44]. However, studies on microbial communities characterized with respect to their growth rates and carbon utilization along the root of various plant species has recently given some insight about the variations of carbon flow of root exudates with root location and time [11,12,45,46]. As an example, Maloney et al. [45] have showed that, along tomato root, the ratio copiotrophic vs oligotrophic microorganisms was low at the root tip (0.50–0.68) and the highest at the root base (1.20–1.28), suggesting that these variations may reflect differences in root exudation. Indeed, the collar is known to be a location of high exudation [47]. This would then account for the high density of culturable bacteria and for their division recorded in zone C in this study. This observation

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is consistent with that of S€ oderberg and B a ath [48] who showed with thymidine and leucine incorporating techniques that bacteria present in the collar zone had the highest metabolic activity. A decrease of activity from the collar to the apical zone using lux AB-tagged pseudomonads has also been reported by Kragelund et al. [39]. In the same way, the preferential location of bacteria at the junction between epidermal cells and the branching point in the hairy zone and their intensive divisions at these sites recorded in here could be related to the facts that these spots are known to be major exudation sites [49]. In conclusion, P. fluorescens A6RI showed a precise spatial-temporal pattern in the early colonization of primary tomato roots. The characterization of this spatiotemporal pattern was made possible, thanks to a combination of different approaches including dilution-plating, flow cytometry and microscopic observations. Further research is currently being done in soil conditions in the presence of the indigenous microflora which is likely to affect the exudation and colonization pattern. The colonization pattern of P. fluorescens A6RI is also going to be compared to that of pathogenic fungi for which this fluorescent pseudomonad is used as a biocontrol agent.

Acknowledgements The authors are grateful to K. Klein for correcting the English text and to Stefania Aimo for the technical support. This work was supported by the INCO-DC Program ERBIC18CT970180.

References [1] Weller, D.M. (1998) Biological control of soilborne pathogens in the rhizosphere with bacteria. Ann. Rev. Phytopathol. 26, 379– 407. [2] Lemanceau, P. and Alabouvette, C. (1993) Suppression of fusarium-wilts by fluorescent pseudomonads: mechanisms and applications. Biocontrol Sci. Technol. 3, 219–234. [3] Schippers, B., Bakker, A.W. and Bakker, P.A.H.M. (1987) Interactions of deleterious and beneficial rhizosphere microorganisms and the effect on cropping practices. Annu. Rev. Phytopathol. 25, 339–358. [4] Bull, C.T., Weller, D.M. and Thomashow, L.S. (1991) Relationship between root colonization and suppression of Gaeumannomyces graminis var. tritici by Pseudomonas fluorescens strain 2-79. Phytopathology 83, 63–68. [5] Raaijmakers, J.M., Leeman, M., van Oorschot, M.M.P., van der Sluis, I., Schippers, B. and Bakker, P.A.H.M. (1995) Dose– response relationships in biological control of fusarium wilt of radish by Pseudomonas spp. Phytopathology 85, 1075–1081. [6] van Overbeek, L.S., Eberl, L., Givskov, M., Molin, S. and van Elsas, J.D. (1995) Survival of, and induced stress resistance in, carbon-starved Pseudomonas fluorescens cells residing in soil. Appl. Environ. Microbiol. 61, 4202–4208. [7] Rainey, P.B. (1999) Adaptation of Pseudomonas fluorescens to the plant rhizosphere. Environ. Microbiol. 1, 243–257.

[8] Mirleau, P., Delorme, S., Philippot, L., Meyer, J., Mazurier, S. and Lemanceau, P. (2000) Fitness in soil and rhizosphere of Pseudomonas fluorescens C7R12 compared with a C7R12 mutant affected in pyoverdine synthesis and uptake. FEMS Microbiol. Ecol. 34, 35–44. [9] Mirleau, P., Philippot, L., Corberand, T. and Lemanceau, P. (2001) Involvement of nitrate reductase and pyoverdine in competitiveness of Pseudomonas fluorescens strain C7R12 in soil. Appl. Environ. Microbiol. 67, 2627–2635. [10] Lugtenberg, B.J.J., Dekkers, L. and Bloemberg, G.V. (2001) Molecular determinants of rhizosphere colonization by Pseudomonas. Ann. Rev. Phytopathol. 39, 461–490. [11] Folman, L.B., Postma, J. and van Veen, J.A. (2001) Ecophysiological characterization of rhizosphere bacterial communities at different root locations and plant developmental stages of cucumber grown on rockwool. Microb. Ecol. 42, 586–597. [12] Baudoin, E., Benizri, E. and Guckert, A. (2002) Impact of growth stage on the bacterial community structure along maize roots, as determined by metabolic and genetic fingerprinting. Appl. Soil Ecol. 19, 135–145. [13] Loper, J.E. and Buyer, J.S. (1991) Siderophores in microbial interactions on plant surfaces. Mol. Plant–Microbe Interact. 4, 5–13. [14] Cook, R.J., Thomashow, L.S., Weller, D.M., Fujimoto, D., Mazzola, M., Bangera, G. and Kim, D.-S. (1995) Molecular mechanisms of defense by rhizobacteria against root disease. Proc. Natl. Acad. Sci. USA 92, 4197–4201. [15] Duijff, B.J., Pouhair, D., Olivain, C., Alabouvette, C. and Lemanceau, P. (1998) Implication of systemic induced resistance in the suppression of fusarium wilt of tomato by Pseudomonas fluorescens WCS417r and nonpathogenic Fusarium oxysporum Fo47. Eur. J. Plant Pathol. 104, 903–910. [16] van Loon, L.C., Bakker, P.A.H.M. and Pieterse, C.M.J (1998) Systemic resistance induced by rhizosphere bacteria. Ann. Rev. Phytopathol. 36, 453–483. [17] Leeman, M., Van Pelt, J.A., Den Ouden, F.M., Heinsbroek, M., Bakker, P.A.H.M. and Schippers, B. (1995) Induction of systemic resistance against fusarium wlt of radish by lipopolysaccharides of Pseudomonas fluorescens. Phytopathology 85, 1021–1027. [18] Hansen, M., Kragelund, L., Nybroe, O. and Sørensen, J. (1997) Early colonization of barley roots by Pseudomonas fluorescens studied by immunofluorescence technique and confocal laser scanning microscopy. FEMS Microbiol. Ecol. 23, 353–360. [19] Chin-A-Woeng, T.F.C., de Priester, W., van der Bij, A. and Lugtenberg, B.J.J. (1997) Description of the colonization of a gnotobiotic tomato rhizosphere by Pseudomonas fluorescens biocontrol strain WCS365, using scanning electron microscopy. Mol. Plant-Microbe Interact 10, 79–86. [20] Normander, B., Hendriksen, N.B. and Nybroe, O. (1999) Green fluorescent protein marked Pseudomonas fluorescens: localization, viability and activity in the natural barley rhizosphere. Appl. Environ. Microbiol. 65, 4646–4651. [21] Bloemberg, G.V., Wijfjes, A.H., Lamers, G.E., Stuurman, N. and Lugtenberg, B.J. (2000) Simultaneous imaging of Pseudomonas fluorescens WCS365 populations expressing three different autofluorescent proteins in the rhizosphere: new perspectives for studying microbial communities. Mol. Plant-Microbe Interact 13, 1170–1176. [22] L€ ubeck, P.S., Hansen, M. and Sørensen, J. (2000) Simultaneous detection of the establishment of seed-inoculated Pseudomonas fluorescens strain DR54 and native soil bacteria on sugar beet root surface using fluorescence antibody and in situ hybridisation techniques. FEMS Microbiol. Ecol. 33, 11–19. [23] Simons, M., van der Bij, A.J., Brand, I., de Weger, L.A., Wijffelman, C.A. and Lugtenberg, B.J.J (1996) Gnotobiotic system for studying rhizosphere colonization by plant growth promoting Pseudomonas bacteria. Mol. Plant-Microbe Interact 9, 600–607.

E. Gamalero et al. / FEMS Microbiology Ecology 48 (2004) 79–87 [24] Rattray, E.A.S., Prosser, J.I., Glover, L.A. and Killham, K. (1995) Characterization of rhizosphere colonization by luminescent Enterobacter cloacae at the population and single-cell levels. Appl. Environ. Microbiol. 61, 2950–2957. [25] Kragelund, L. and Nybroe, O. (1996) Competition between Pseudomonas fluorescens Ag1 and Alcaligenes eutrophus JMP134 (pJP4) during colonization of barley roots. FEMS Microbiol. Ecol. 20, 41–51. [26] Eparvier, A., Lemanceau, P. and Alabouvette, C. (1991) Population dynamics of non-pathogenic Fusarium and fluorescent Pseudomonas strains in rockwool, a substratum for soilless culture. FEMS Microbiol. Ecol. 86, 177–184. [27] Lemanceau, P. and Samson, R. (1983). Relations entre quelques caracteristiques in vitro de 10 Pseudomonas fluorescents et leur effet sur la croissance du haricot (Phaseolus vulgaris). In: Les antagonismes microbiens, 24
eme colloque de la SFP, Bordeaux, p. 360. [28] Gamalero, E., Martinotti, M.G., Trotta, A., Lemanceau, P. and Berta, G. (2002) Morphogenetic modifications induced by Pseudomonas fluorescens A6RI and Glomus mosseae BEG12 in the root system of tomato differ according to plant growth conditions. New Phytol. 155, 293–300. [29] Miller, J.H. (1972) Experiments in Molecular Genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. [30] King, E.O., Ward, M.K. and Raney, D.E. (1954) Two simple media for the demonstration of pyocianin and fluorescin. J. Lab. Clin. Med. 44, 301–307. [31] Trotta, A., Varese, G.C., Gnavi, E., Fusconi, E., SampoÕ, S. and Berta, G. (1996) Interaction between the soil borne pathogen Phytophthora parasitica var. parasitica and the arbuscular mycorrhizal fungus Glomus mosseae in tomato plants. Plant Soil 185, 199–209. [32] Loper, J.E., Haack, C. and Schroth, M.N. (1985) Population dynamics of soil pseudomonads in the rhizosphere of potato (Solanum tuberosum L.). Appl. Environ. Microbiol. 49, 416– 422. [33] Jeffree, C.E. and Read, N.D. (1991) Ambient- and low-temperature scanning electron microscopy. In: Electron Microscopy of Plant Cells (Hall, J.L. and Hawes, C., Eds.). Academic Press, London, San Diego. [34] Oliver, J.D. (1993) Formation of viable but nonculturable cells. In: Starvation in Bacteria (Kjelleberg, S., Ed.), pp. 239–272. Plenum Press, New York. [35] Mascher, F., Hase, C., Moenne-Loccoz, Y. and Defago, G. (2000) The viable-but-nonculturable state induced by abiotic stress in the biocontrol agent Pseudomonas fluorescens CHA0 does not promote strain persistence in soil. Appl. Environ. Microbiol. 66, 1662–1667.

87

[36] Binnerup, S.J., Jensen, D.F., Thordal-Christensen, H. and Sørensen, J. (1993) Detection of viable, but non-culturable Pseudomonas fluorescens DF57 in soil using a microcolony epifluorescence technique. FEMS Microbiol. Ecol. 12, 97–105. [37] Troxler, J., Zala, M., Natsch, A., Mo€enne-Loccoz, Y., Keel, C. and Defago, G. (1997) Autoecology of the biocontrol strain Pseudomonas fluorescens CHA0 in the rhizosphere and inside roots at later stage of plant developement. FEMS Microbiol. Ecol. 24, 119–130. [38] Fusconi, A., Gnavi, E., Trotta, A. and Berta, G. (1999) Apical meristems of tomato roots and their modifications induced by arbuscular mycorrhizal and soilborne pathogenic fungi. New Phytol. 142, 505–516. [39] Kragelund, L., Hosbond, C. and Nybroe, O. (1997) Distribution of metabolic activity and phosphate starvation response of luxtagged Pseudomonas fluorescens reporter bacteria in the barley rhizosphere. Appl. Environ. Microbiol. 63, 4920–4928. [40] Roberts, D.P., Kobayashi, D.Y., Dery, P.D. and Short, N.M. (1999) An image analysis method for determination of spatial colonization patterns of bacteria in plant rhizosphere. Appl. Microbiol. Biotechnol. 51, 653–658. [41] Ma, W., Zalec, K. and Glick, B.R. (2001) Biological activity and colonization pattern of the bioluminescence-labelled plant growthpromoting bacterium Kluyvera ascorbata SUD165/26. FEMS Microbiol. Ecol. 35, 137–144. [42] van Vuurde, J.W.L. and Schippers, B. (1980) Bacterial colonization of seminal wheat roots. Soil Biol. Biochem. 12, 559–565. [43] McDougald, D., Rice, S.A., Weichart, D. and Kjelleberg, S. (1997) Nonculturability: adaptation or debilitation? FEMS Microbiol. Ecol. 25, 1–9. [44] Whipps, J.M. (1987) Carbon loss from the roots of tomato and pea seedlings in soil. Plant Soil 103, 95–100. [45] Maloney, P.E., van Bruggen, A.H.C. and Hu, S. (1997) Bacterial community structure in relation to the carbon environments in lettuce and tomato rhizospheres and in bulk soil. Microb. Ecol. 34, 109–117. [46] Semenov, A.M., van Bruggen, A.H.C. and Zelenev, V.V. (1999) Moving waves of bacterial populations and total organic carbon along roots of wheat. Microb. Ecol. 37, 116–128. [47] Grayston, S.J., Vaughan, D. and Jones, D. (1996) Rhizosphere carbon flow in trees, in comparison with annual plants: the importance of root exudation and its impact on microbial activity and nutrient availability. Appl. Soil Ecol. 5, 29–56. [48] S€ oderberg, K.H. and B a ath, E. (1998) Bacterial activity along a young barley root measured by the thymidine and leucine incorporation techniques. Soil Biol. Biochem. 30, 1259–1268. [49] Curl, E.A. and Truelove, B. (1986) The Rhizosphere. Springer, New York. 288 pp.