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(both supplied by Paul De Ley, University of California,. Riverside, CA ..... 61, 2950^2957. [6] Ryder, M.H. and Rovira, A.D. (1993) Biological control of take-all.
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Nematode-enhanced microbial colonization of the wheat rhizosphere O.G.G. Knox, K. Killham, C.E. Mullins, M.J. Wilson



School of Biological Sciences, University of Aberdeen, Cruickshank Building, St Machar Drive, Aberdeen AB24 3UU, UK Received 28 April 2003; received in revised form 11 June 2003; accepted 25 June 2003 First published online 17 July 2003

Abstract The mechanisms by which seed-applied bacteria colonize the rhizosphere in the absence of percolating water are poorly understood. Without mass flow, transport of bacteria by growing roots or soil animals, particularly nematodes may be important. We used a sandbased microcosm system to investigate the ability of three species of nematodes (Caenorhabditis elegans, Acrobeloides thornei and a Cruznema sp.) to promote rhizosphere colonization by four strains of beneficial rhizobacteria. In nearly all cases, rhizosphere colonization was substantially increased by the presence of nematodes, irrespective of bacterial or nematode species. Our results suggest that nematodes are important vectors for bacteria rhizosphere colonization in the absence of percolating water. 3 2003 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. Keywords : Biocontrol ; Nematodes; Rhizosphere Colonization; Seed-Applied Bacteria ; Microsomas

1. Introduction The use of seed-applied bene¢cial soil bacteria as biofertilizers, biocontrol products or for bioremediation is an area of intense study [1^3]. The rhizosphere favors bacterial growth and survival, and is the area of soil where biofertilizers [5] and biofungicides [6,7] are targeted. It has also been suggested as a site that can enhance activity of bioremediating bacteria [8] (rhizoremediation). Thus, seed application of bene¢cial bacteria is a key strategy for introducing bacteria to soil [7,9]. The ability of di¡erent species and strains of bacteria to colonize the rhizosphere under di¡erent soil moisture conditions varies greatly [10]. Under conditions of mass £ow, transport of bacteria by water is the main mechanism of colonization for seed-applied bacteria [2,11]. In soils at or close to ¢eld capacity, bacterial motility in response to chemotaxis is thought to be important for movement into root channels or into contact with transporting agents, although motility is not generally considered a method of colonization in itself [11]. Under drier conditions, root colonization is

* Corresponding author. Tel. : +44 (1224) 272845; Fax : +44 (1224) 272703. E-mail address : [email protected] (M.J. Wilson).

likely to involve either adherence to the growing root tip and mucigel utilization, adherence to extending fungal mycelia, or movement by soil animals. It is well known that earthworms can transport bacteria [12^15]. Furthermore, it is known that they can aid root colonization by bacteria [16]. However, there are few published studies on the in£uence of nematodes in root colonization. Nematodes are invariably the most abundant soil animals in agricultural soils, and may occur in extremely high numbers ( s 100 g31 soil) [17,18]. While nematodes move much shorter distances than earthworms, their ubiquitous nature means that bacteria, introduced on a coated seed, are much more likely to come into contact with nematodes [19]. The presence of nematodes in the rhizosphere has been shown to increase bacterial growth, stimulated by the products of incomplete digestion released by nematodes accompanied by increased nitrogen mineralization [20,21]. In addition, presence of nematodes in model soils enhances bacterial activity by distributing bacterial colonies over the organic substrate [22]. There has been some work demonstrating that nematodes can act as vectors of plant pathogenic bacteria [23] or rhizobium [24,25]. These studies found that the nematodes distributed cells more evenly over the root surface and thus bene¢ted the plants, and it is therefore surprising that this area of research has not been further developed.

0378-1097 / 03 / $22.00 3 2003 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. doi:10.1016/S0378-1097(03)00517-2

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In order to investigate the potential for nematodes to act as vectors for improved colonization of root surfaces by seed-applied bacteria, we undertook a simple sandbased microcosm study using a combination of motile Gram-negative and Gram-positive bacteria, with or without free-living bacterial-feeding nematodes of the families Rhabditae and Cephalobidae. The bacteria selected had been previously studied in rhizosphere colonization experiments and were known to remain culturable. We used colonization of the wheat rhizosphere in the absence of percolating water as a model system.

2. Materials and methods 2.1. Organisms Winter wheat (Triticum aestivum var. Savannah) (Advanta Seeds, UK) was selected for use in the trial as a good representative of presently used winter wheat varieties. Nematode species used were the Cephalobid Acrobeloides thornei DWF1109, a small nematode, the Rhabditid Cruznema sp. Rosario strain, a much larger nematode (both supplied by Paul De Ley, University of California, Riverside, CA, USA) and the laboratory standard Rhabditid nematode Caenorhabditis elegans N2 (Christina Lagido, University of Aberdeen, UK). Three Gram-negative pseudomonads were selected from two species. Pseudomonas corrugata 2140, isolated from Australian soil, had been reported as a potential biocontrol agent for take-all in wheat and a good rhizosphere colonizer [6]. P. £uorescens 10585 (NCIMB, Aberdeen, UK) has demonstrated rhizosphere colonization capabilities under matric potentials of 3500 kPa [5] and had been previously used in soil studies [4,26^30]. P. £uorescens SBW25 (CEH, Oxford, UK) is a sugar beet phytosphere isolate that has been characterized as a colonizer of roots and shoots for several plants in ¢eld and laboratory trials [31^34]. The Gram-positive isolate Bacillus subtilis MBI600 (Becker Underwood, Littlehampton, UK) is sold commercially as a biocontrol agent. All pseudomonad strains were chromosomally marked with Kanr genes. Introduced antibiotic genes were used as a selectable marker for recovery of all bacterial species. The B. subtilis was marked with the plasmid pSB340 [35,36] carrying chloramphenicol resistance. All bacterial strains were routinely cultured in Luria Bertani broth (LB) or LB agar containing the appropriate antibiotics. 2.2. Growth and preparation of nematodes Petri dishes of Nematode Growth Medium [37] were inoculated with E. coli HB101 1 day prior to inoculation with the nematodes C. elegans, A. thornei and Cruznema sp. and grown at 25‡C for 7 days. Nematodes were then maintained at 17‡C until required, when they were transferred to a Baermann funnel apparatus [38] for extraction

into water. Nematodes were counted using a dissecting microscope and added to microcosms at a rate of 10 g31 sand (A. thornei and Cruznema) or 5 g31 sand (C. elegans). 2.3. Surface sterilization, coating and germination of wheat seeds Wheat seeds were sterilized using an oxytetracycline/silver nitrate method [39]. For seed coating, test bacteria were grown at 25‡C with shaking at 200 rpm for 20 h after which 6 ml of culture was centrifuged, washed in 1 ml of 1/4 strength Ringer’s solution (Oxoid), and then re-centrifuged. The resulting bacterial pellet was suspended in sterile 1% (w/v) high viscosity Carboxy Methyl Cellulose in 1/4 strength Ringer’s solution [9], applied to 4 g of surface-sterilized wheat seeds, air-dried for 2 h prior to being germinated on moist ¢lter paper in the dark at 25‡C for 24 h. 2.4. Preparation of microcosms Microcosms were prepared in root trainers (60 mm deep, with a square top of 36U36 mm) containing 37.5 g sharp sand (99, 94 and 13% less than 500, 355, and 180 Wm, respectively, CHAP Construction, Scotland, UK) that had been washed and autoclaved then ovendried at 105‡C. The sand was prepared at water contents of 0.104 and 0.208 g g31 dry sand. These correspond to matric potentials of 32.6 and 32.0 kPa, respectively, as determined by the ¢lter-paper method [40]. Microcosms were established by adding the required amount of water (containing the appropriate numbers of nematodes) to the surface and leaving for 16 h to equilibrate before germinated seeds were planted at 1 cm depth. Microcosms were sealed in clear polythene bags and incubated in a high light Phytotron for 14 days with 12-h periods of light and dark at 13.5 and 15‡C, respectively. 2.5. Experimental designs We did three colonization experiments. In the ¢rst, microcosms were established at 0.104 and 0.208 g g31 dry sand, containing no nematodes, A. thornei or Cruznema sp. Wheat seeds were coated with all of four test bacteria and four germinated seeds were planted out for each investigated treatment, resulting in a total of 96 microcosms. The second experiment was similar, but included an additional nematode treatment (C. elegans) and thus comprised 128 microcosms. The third experiment used microcosms established only at 0.208 g g31 dry sand and used all three nematode species. In addition, a dead nematode treatment was included as a further control. This consisted of a mixture of equal numbers of the three nematode species, killed by heating to 65‡C for 2 h and applied to give 10 dead nematodes g31 sand. The third experiment thus comprised a total of 80 microcosms.

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Fig. 1. Transformed mean log10 numbers of cfu per mg of root for the three colonization experiments (A, C and E, respectively). Nematode treatments were; no nematodes (¢rst bar), Cruznema sp. (second bar in A and C; third bar in E), A. thornei (third bar in A and C; fourth bar in E), C. elegans (fourth bar in C; ¢fth bar in E) and dead nematodes (second bar in E). Interactions between nematodes and bacterial species are shown in B, D and F for trials 1, 2 and 3, respectively. Nematode treatments are indicated on the x-axis and bacterial treatments are shaded according to; P. £uorescens 10586 (¢rst bars), P. £uorescens SBW25 (second bars), B. subtilis (third bars) and P. corrugata 2140 (fourth bars). LSD = least signi¢cant di¡erence P = 0.05.

2.6. Plant harvesting, root recovery and bacterial enumeration

plates. Colonies were counted and the primary roots were oven-dried at 105‡C for 48 h and weighed.

Root systems were recovered from sand microcosms and the shoots and seeds removed. The root system was then divided into primary roots, which were measured. These were then placed into 1/4 strength Ringer’s, vortexed for 5 s and left to stand for 2 h. The resulting bacterial suspension was serially diluted and four replicate 10-Wl aliquots of each dilution grown on selective agar

2.7. Nematode recovery from microcosms and investigation into nematode distribution within the rhizosphere In the ¢rst two experiments, one microcosm of each treatment was transferred to a Baermann funnel for 16 h and extracted nematodes were then counted. In the third

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experiment nematodes were recovered in this manner from all microcosms. To establish nematode distribution throughout the sand microcosm 50-ml centrifuge tubes were cut into 2-cm sections. Tubes were reassembled with masking tape, covered in aluminum foil and 50 g of sterile oven-dried sand added. A. thornei, Cruznema and C. elegans were added as previously described with all microcosms established at 0.208 g g31 dry sand. P. £uorescens 10586 lux coated seeds were prepared, germinated and planted as previously described. Microcosms were incubated for 2 weeks in a high light Phytotron. After this time, the tubes were dismantled into sections, the sand and roots in each section weighed, and nematodes extracted using a centrifugal £otation method [38] and counted. 2.8. Statistical analysis All data were log10-transformed to stabilize the variance and compared using an analysis of variance (ANOVA) General Linear Model approach (Minitab release 12.21). When ANOVA showed signi¢cant treatment e¡ects, individual means were compared using Tukey’s pairwise comparison.

3. Results 3.1. Bacterial recovery from primary roots In the ¢rst experiment, ANOVA showed signi¢cant effects of bacterial seed coat and nematode treatment, but not water content on numbers of bacterial cfu mg31 root. Numbers of cfu mg31 root for the various nematode treatments and interactions between bacterial seed coat and the nematode treatment are shown in Fig. 1A,B. The presence of nematodes increased bacterial colonization, and this

was particularly true in microcosms inoculated with Cruznema sp. (Fig. 1A). The e¡ects of bacterial and nematode species interacted signi¢cantly. This is largely a result of the high level of colonization of P. £uorescens 10586 in the absence of nematodes, whereas all other bacteria colonized less e¡ectively (Fig. 1B). In addition, presence of A. thornei signi¢cantly reduced colonization by this bacterium, while this nematode signi¢cantly (P 6 0.05) increased colonization by P. £uorescens SBW25 and P. corrugata 2140. In the second experiment, there were signi¢cant (P 6 0.05) e¡ects for bacterial seed coat, nematode treatment and water content on colonization. Addition of all three nematode species to microcosms increased colonization at a highly signi¢cant (P 6 0.001) level (Fig. 1C). Unlike the ¢rst experiment, all nematode species signi¢cantly increased colonization by all bacterial seed coats (Fig. 1D). Despite the small di¡erence in matric potential, colonization of bacteria was signi¢cantly (P 6 0.001) greater in microcosms that established a water content of 0.208 than 0.104 g g31 (mean log10 cfu mg31 = 2.59 R 0.05 S.D. and 2.28 R 0.05 S.D., respectively). In the third experiment, signi¢cant (P 6 0.05) e¡ects for seed coat, and nematode species were once again observed and the two factors interacted. Trends were generally similar to the second experiment. An important observation was that addition of dead nematodes did not signi¢cantly increase bacterial colonization above control levels (Fig. 1E). In this experiment, no nematode treatment signi¢cantly increased colonization of B. subtilis, and only C. elegans signi¢cantly increased colonization by P. £uorescens SBW25 (Fig. 1F). 3.2. Nematode recovery form microcosms Nematode recovery from microcosms after 2 weeks of plant growth was low, fewer than three nematodes g31 sand recovered in all experiments, and variable regardless

Fig. 2. Number of nematodes recovered g31 sand from experimental microcosm set up in trial 1 (E), trial 2 (l) and trial 3 (F). Nematode names abbreviated to Cruz (Cruznema sp.), A. th (A. thornei) and C. ele (C. elegans). Bacterial seed coat abbreviated to 2140 (P. corrugata 2140), MBI600 (B. subtilis MBI600), 10586 (P. £uorescens 10586) and SBW25 (P. £uorescens SBW25).

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Fig. 3. The number of nematodes recovered from di¡erent depths of triplicate microcosms, expressed as a percentage of the number of nematodes originally added. Nematode treatment identi¢cation; Cruznema sp. (E), A. thornei (l), C. elegans (F). Error bars represent the standard deviation of the means.

of nematode species and seed coat. Survival of A. thornei tended to be greater than the other two nematode species (Fig. 2). 3.3. Nematode location in the wheat rhizosphere Recovery of nematodes using the centrifugal £otation method again returned low counts compared with numbers added to the sand-based microcosms. Typical recovery never exceeded 5% of nematodes added for each sampled depth (Fig. 3). Numbers of Cruznema and C. elegans were lower in samples taken from the lower fractions of the microcosms, with virtually none being recovered in the 6^8-cm section. Recovered numbers of A. thornei showed a di¡erent pattern, with greatest numbers being recovered in both the 0^2- and 6^8-cm sections, and much lower numbers in intermediate depth sections.

4. Discussion In the model system we used, no further additions of water were made to the microcosms after establishment, preventing bacterial movement by or in percolating water. Analysis of the results indicates that all of our test bacteria were able to colonize and grow throughout a developing wheat rhizosphere, but to di¡erent extents. Di¡erences in colonization probably resulted from di¡erences in bacterial species ability to adhere to the root tip, utilize plant exudates and/or a lack of motility [41,42]. However, addition of live nematodes to our system substantially increased the degree of colonization for all test bacteria. When the ubiquitous nature of bacteria and nematodes in soils is considered, along with the fact that all nematodes share an environment with bacteria at one time in their life [19], it seems highly likely that nematodes will have a bene¢cial impact on rhizosphere colonization by seed-applied bacteria. The mechanisms by which nema-

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todes increased bacterial colonization are not ascertainable from the data presented here, but probably a combination of movement of the bacteria on the nematodes cuticle [17], passage of viable bacteria through the nematodes digestive system [23] or enhancing bacterial growth as a result of nematode excretion [20] are probably all involved. Di¡erences in colonization observed between experiments might have resulted from di¡erent rates of nematode mortality. This in turn would also explain why the recovered numbers of nematodes di¡ered at the end of each experiment (Fig. 2). Reasons for changes in mortality between experiments and treatments are not clear, but reductions in nematode numbers from the initial inoculum were recorded in every experiment possibly due to either the low pH of the washed sterile sand (4.8 R 0.2), or starvation. Poor nematode survival in sand microcosms has been reported previously. Decreases in nematode numbers to about 60% of the initial inoculum in sand microcosms [43] and in storage at pH 4^6 [44] have been reported for entomopathogenic nematodes. Our third experiment showed that the increased bacterial colonization could not have resulted solely from increased availability of substrate and nutrients from dead nematodes. Microcosms to which dead nematode treatments had been added had levels of bacterial colonization similar to control microcosms identifying the importance of live nematodes. Whatever the reasons for the decrease in nematode numbers within the microcosms, the overall e¡ect on bacterial colonization of adding live nematodes to the system was to increase it. Analysis of abundance of the nematodes throughout the wheat rhizosphere indicated di¡erences in survival rate and location of each species. Previous studies involving C. elegans demonstrated that sand in the size range of 0.125^1.25 mm did not a¡ect nematode distribution [18], although in those systems, the sand was kept moist. Water is important for nematode movement, although the presence of A. thornei and not Cruznema or C. elegans in the lower fractions of the recovered sections suggests that nematode size was probably a more important factor in determining depth of nematode movement. Attraction of nematodes to active bacterial colonies has been demonstrated [18,19,45] and activity of P. £uorescens 10586 was probably highest near the root tip, where increased root exudation occurs [8]. If Cruznema and C. elegans had been unable to locate this food source, due to either an inability to keep up with or get to the root tip, then these nematodes would likely starve and be recovered in lower numbers, as was observed to be the case. However, in trials 1 and 3 the smaller A. thornei was less successful at increasing colonization than the larger Cruznema sp. or C. elegans. These observations might imply that the larger nematodes carry more bacteria on their cuticle and increase colonization in this manner, although this was not investigated here. Despite the observed reduction in nematode numbers during experiments, our results suggest that these animals

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play a useful role as vectors for bacteria colonization of the rhizosphere in the absence of percolating water.

Acknowledgements We thank the BBSRC and the BIRE initiative for funding this work, and Mark Bailey, Tracey Timms-Wilson (CEH, Oxford, UK) and Paul DeLey (University of California, Riverside, CA, USA) for provision of cultures and technical assistance.

References [1] Bashan, Y. (1998) Inoculants of plant growth-promoting bacteria for use in agriculture. Biotechnol. Adv. 16, 729^770. [2] Parke, J.L. (1991) Root colonization by indigenous and introduced microorganisms. In: The Rhizosphere and Plant Growth (Glaser, D.L. and Cregan, P.B., Eds.), pp 33^42. Kluwer, Dordrecht. [3] Weller, D.M. and Thomashow, L.S. (1994) Current challenges in introducing bene¢cial microorganisms into the rhizosphere. In: Molecular Ecology of Rhizosphere Micro-Organisms (O’Gara, F, Dowling, D.N. and Boesten, B., Eds.), pp. 1^18. VCH, Weinheim. [4] Rattray, E.A.S., Prosser, J.I., Glover, L.A. and Killham, K. (1992) Matric potential in relation to survival and activity of a genetically modi¢ed microbial inoculum in soil. Soil Biol. Biochem. 24, 421^425. [5] Rattray, E.A.S., Prosser, J.I., Glover, L.A. and Killham, K. (1995) Characterisation of rhizosphere colonization by luminescent Enterobacter cloacae at population and single-cell levels. Appl. Environ. Microbiol. 61, 2950^2957. [6] Ryder, M.H. and Rovira, A.D. (1993) Biological control of take-all of glasshouse-grown wheat using strains of Pseudomonas corrugata isolated from wheat ¢eld soil. Soil Biol. Biochem. 25, 311^320. [7] Weller, D.M. (1988) Biological control of soil-borne pathogens in the rhizosphere with bacteria. Annu. Rev. Phytopathol. 26, 379^407. [8] Killham, K. (1994) Soil Ecology. Cambridge University Press, Cambridge. [9] Ciccillp, F., Fiore, A., Bevivino, A., Dalmastri, C., Tabacchioni, S. and Chiarini, I. (2002) E¡ects of two di¡erent application methods of Burkholderia ambifaria MCI 7 on plant growth and rhizosphere bacterial diversity. Environ. Microbiol. 4, 238^245. [10] Rattray, E.A.S., Tyrrel, J.A., Prosser, J.I., Glover, L.A. and Killham, K. (1993) E¡ect of soil bulk density and temperature on wheat rhizosphere colonisation by lux-marked Pseudomonas £uorescens. Eur. J. Soil Biol. 29, 73^82. [11] Gammack, S.M., Paterson, E., Kemp, J.S., Cresser, M.S. and Killham, K. (1992) Factors a¡ecting the movement of microorganisms in soils. In: Soil Biochemistry, Vol. 7 (Stotzky, G. and Bollag, J.M., Eds.), pp. 263^305. Marcel Dekker, New York. [12] Fredrickson, J.K., Bentjen, S.A., Bolton Jr., H., Li, S.W. and van Voris, P. (1989) Fate of Tn5 mutants of root growth-inhibiting Pseudomonas sp. in intact soil-core microcosms. Can. J. Microbiol. 35, 867^873. [13] Madsen, E. and Alexander, M. (1982) Transport of Rhizobium and Pseudomonas through soil. J. Soil Sci. Soc. Am. 42, 557^560. [14] Schmidt, O., Doube, B.M., Ryder, M.H. and Killham, K. (1997) Population dynamics of Pseudomonas corrugata 2140R Lux 8 in earthworm food and in earthworm casts. Soil Biol. Biochem. 29, 523^528. [15] Thorpe, I., Killham, K., Prosser, J.I. and Glover, L.A. (1993) Novel method for the study of the population dynamics of a genetically modi¢ed micro-organism in the gut of the earthworm Lumbricus terrestris. Biol. Fertil. Soils 15, 55^59.

[16] Stephens, P.M., Davoren, C.W., Ryder, M.J. and Doube, B.M. (1993) In£uence of the lumbricid earthworm Aporrectodea trapezoides on the colonization of wheat roots by Pseudomonas-Corrugata strain 2140R in soil. Soil Biol. Biochem. 25, 1719^1724. [17] Poinar Jr., G.O. (1983) The Natural History of Nematodes. PrenticeHall, Englewood Cli¡s, NJ. [18] Young, I.M., Gri⁄ths, B.S., Robertson, W.M. and McNicol, J.W. (1998) Nematode (Caenorhabditis elegans) movement in sand as affected by particle size, moisture and the presence of bacteria (Escherichia coli). Eur. J. Soil Sci. 49, 237^241. [19] Poinar Jr., G.O. and Hansen, E.D. (1986) Associations between Nematodes and Bacteria. Helminthol. Abstr. Ser. B 55, 61^79. [20] Anderson, R.V., Gould, W.D., Woods, L.E., Cambardella, C., Ingham, R.E. and Coleman, D.C. (1983) Organic and inorganic nitrogenous losses by microbivorous nematodes in soil. Oikos 40, 75^80. [21] Ferris, H., Venette, R.C., van der Meulen, H.R. and Lau, S.S. (1998) Nitrogen mineralization by bacterial-feeding nematodes: veri¢cation and measurement. Plant Soil 203, 159^171. [22] Whitford, W.G., Freckman, D.W., Santos, P.F., Elkins, N.A. and Parker, L.W. (1982) The role of nematodes in desert ecosystems. In: Nematodes in Soil Ecosystems (Freckman, D.W. and Wallwork, J.A., Eds.), pp. 98^115. University of Texas Press, Austin, TX. [23] Wasilewska, L. and Webster, J.M. (1975) Free living nematodes as disease factors of man and his crops. Int. J. Environ. Stud. 7, 204^ 210. [24] Jatala, P., Jenson, H.J. and Russel, S.A. (1974) Pristiounchus Iheritieri as a carrier of Rhizobium japonicum. J. Nematol. 6, 130^131. [25] Cayrol, J.C., Couderc, C. and Evrard, I. (1977) Studies of the relationships between free-living soil nematodes and nodule bacteria of leguminous plants. Rev. Zool. Agric. Path. Veg. 76, 77^89. [26] Amin-Hanjani, S., Meikle, A., Glover, L.A., Prosser, J.I. and Killham, K. (1993) Plasmid and chromosomal encoded luminescence marker systems for detection of Pseudomonas £uorescens in soil. Mol. Ecol. 2, 47^54. [27] Kemp, J.S., Patterson, E., Gammack, S.M., Cresser, M.S. and Killham, K. (1992) Leaching of genetically modi¢ed Pseudomonas £uorescens through organic soil : In£uence of temperature, soil pH and roots. Biol. Fertil. Soils 13, 214^218. [28] Meikle, A., Killham, K., Prosser, J.I. and Glover, L.A. (1992) Luminometric measurement of population activity of genetically modi¢ed Pseudomonas £uorescens in the soil. FEMS Microbiol. Lett. 99, 217^ 220. [29] Meikle, A., Glover, L.A., Killham, K. and Prosser, J.I. (1994) Potential luminescence as an indicator of activation of genetically-modi¢ed Pseudomonas £uorescens in liquid culture and in soil. Soil Biol. Biochem. 26, 747^755. [30] Yeomans, C., Porteous, F., Paterson, E., Meharg, A.A. and Killham, K. (1999) Assessment of lux-marked Pseudomonas £uorescens for reporting on organic carbon compounds. FEMS Microbiol. Lett. 176, 79^83. [31] Bailey, M.J., Lilley, A.K., Thompson, I.P., Rainey, P.B. and Ellis, R.J. (1995) Site directed chromosomal marking of a £uorescent pseudomonad isolate from the phytosphere of sugar beet; stability and potential for marker gene transfer. Mol. Ecol. 4, 755^763. [32] De Leij, F.A.A.M., Sutton, E.J., Whipps, J.M., Fenlon, J.S. and Lynch, J.M. (1995) Impact of ¢eld release of genetically modi¢ed Pseudomonas £uorescens on indigenous microbial populations of wheat. Appl. Environ. Microbiol. 61, 3443^3453. [33] De Leij, F.A.A.M., Thomas, C.E., Bailey, M.J., Whipps, J. and Lynch, J.M. (1998) E¡ect of insertion site and metabolic load on the environmental ¢tness of genetically modi¢ed Pseudomonas £uorescens isolate. Appl. Environ. Microbiol. 64, 2634^2638. [34] Thompson, I.P., Lilley, A.K., Ellis, R.J., Bramwell, P.A. and Bailey, M.J. (1995) Survival, colonization and dispersal of a genetically modi¢ed Pseudomonas £uorescens (SBW25) in the phytosphere of ¢eld grown sugar beet. Biotechnology 13, 1493^1497. [35] Hill, P.J., Hall, L., Vinicombe, D.A., Soper, C.J., Setlow, P., Waites,

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[36]

[37]

[38]

[39] [40]

W.M., Denyer, S. and Stewart, G.S.A.B. (1994) Bioluminescence and spores as biological indicators of inimical processes. J. Appl. Bacteriol. Symp. Suppl. 76, 129^134. Knox, O.G.G., Killham, K. and Leifert, C. (2002) The reliability of marker/reporter genes in biocontrol studies with B. subtilis. Biocontrol Sci. Tech. 12, 637^641. Epstein, H.F. and Shakes, D.C. (1995) Caenorhabditis elegans : Modern Biological Analysis of an Organism ^ Methods in Cell Biology, Vol. 48. Academic Press, San Diego, CA. Hooper, D.J. (1986) Extraction of free-living stages from soil. In: J.F. Laboratory Methods for Work with Plant and Soil Nematodes (Southey, J.F., Ed.), pp. 5^31. HMSO, London. Speakman, J.B. and Kruger, W. (1983) A comparison of methods to surface sterilise wheat seeds. Trans. Br. Mycol. Soc. 80, 374^376. Deka, R.N., Wairiu, M., Mtakwa, P.W., Mullins, C.E., Veenendaal, E.M. and Townend, J. (1995) Use and accuracy of the ¢lter paper technique for measurement of soil matric potential. Eur. J. Soil Sci. 46, 233^238.

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[41] Bashan, Y. and Levanoy, H. (1989) Wheat root tips for passive vertical transfer of Azospirillum brasilense. J. Gen. Microbiol. 135, 2899^2909. [42] Trevors, J.T., VanElsas, J.D., VanOverbeek, L.S. and Starodub, M.E. (1990) Transport of genetically engineered Pseudomonas £uorescens strain through a soil microcosm. Appl. Environ. Microbiol. 56, 401^408. [43] Csontos, A.S. (2002) Lateral movement of the entomopathogenic nematodes Steinernema glaseri and Heterorhabditis bacteriophora in sand at di¡erent temperatures in response to host seeking. Biocontrol Sci. Tech. 12, 137^139. [44] Strauch, O., Niemann, I., Neumann, A., Schmidt, A.J., Peters, A. and Ehlers, R.U. (2000) Storage and formulation of the entomopathogenic nematodes Heterorhabditis indica and H. bacteriophora. Biocontrol 45, 483^500. [45] Ward, S. (1973) Chemotaxis by the nematode Caenorhabditis elegans : identi¢cation of attractants and analysis of the response by use of mutants. Proc. Natl. Acad. Sci. USA 70, 817^821.

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