Factors affecting the survival of a FLavo bacterium species in non-planted and rhizosphere soil

Soil Bid. Bioclwm. Vol. 26, No. 7. pp. 849-X59, 1994

0038-0717(94)EOOO2-H

Copyright @> 1994 Elsevier Science Ltd Printed in Great Britain.All rights reserved

0038-0717/94$7.00+ 0.00

FACTORS AFFECTING THE SURVIVAL OF A FLA VOBACTERIUM SPECIES IN NON-PLANTED AND RHIZOSPHERE SOIL JANEL. MAWDSLEY*and RICHARDG. BURNS~ Research School of Biosciences, Biological Laboratory, University of Kent, Canterbury, Kent CT2 7NJ, England (Accepted

I7 December

1993)

Summary-Factors alkting the survival of a Gram-negative soil Flavobacterium sp. (P25) were investigated in small-scale laboratory microcosms. Two terms for expressing P25 persistence are used: survivalAescribing the number of P25 reisolated from soil at a certain time point; and survival rare-expressed as the time in days taken for a 90% (t90), 99% (t99) or 99.9% (t99.9) decline in numbers of P25. Survival rates of P25 increased in rhizosphere as opposed to bulk soil and were increased further by applying the inoculum directly to the planted seedling (t99 = 21 d) rather than homogenizing it through the soil prior to planting (t99 = 10.5 d). Survival was increased by using greater inoculum densities, for example following inoculation at I. I x IO9P25 gg ’ soil, numbers surviving at day 40 were l2.5-fold greater than that achieved with I.1 x IO4P25 g-‘. In addition, survival rates of P25 were increased by applying the inoculant to soil at moisture holding capacities (MHC) between 40 and 50% (t99.9 = 12.2d) in comparison to either wetter (e.g. 100% MHC, t99.9 = 3.5 d) or drier (e.g. 10% MHC, t99.9 = I d) soils. Carbon amendments (i.e. galactose, maltose, sorbitol) significantly increased survival of P25 in nonplanted soil throughout the 35 d study whereas in rhizosphere soil an increase was only recorded for the first l4d.

INTRODUCTION

In recent years microbial inoculants have been used for soil bioremediation (Have1 and Reineke, 1992), plant growth promotion (Seong et al., 1992) and biological control (Maurhofer et al., 1992). However, although microorganisms have shown great promise in in vitro screens, the expression of their beneficial properties in the natural environment is often unpredictable (van Elsas and Heijnen, 1990). The fundamental reasons for the disparity between laboratory data and field data are poorly understood (Hozore and Alexander, 1991) which means that each putative inoculant requires detailed studies of its behaviour in soil and its effect on soil processes and populations before it can be released into the environment. The solution to the problem of predicting inoculant behaviour is considered to be the use of a wide variety of tests involving sterile and non-sterile laboratory microcosms (Pickup et al., 1991). Much debate has focused on the relevance and design of microcosm experiments (Burns, 1988) but some have been shown to be good predictors of survival and activity in the field (Bolton et al., 1991; Teuben and Verhoef, 1992; Thompson et al., 1992). *Present address: AFRC Institute of Grassland and Environmental Research, Plas Gogerddan, Dyfed SY23 3EB, Wales. TAuthor for correspondence.

Aberystwyth,

The introduction of bacteria to soil is often followed by a gradual or, in some cases, a rapid decline in numbers. In the case of the biocontrol bacterium Bacillus thuringiensis this does not effect its efficacy, as ingestion and cell lysis releases the insecticidal protein (Beringer et al., 1989). However, for the majority of inoculants, expression of their beneficial properties is dependant upon their survival and growth in soil (Heijnen et al., 1993). Soil is a highly heterogenous environment (van Elsas and Heijnen, 1990) and many abiotic and biotic factors will affect soil inoculants (England et al., 1993). Abiotic factors which have been reported to affect inoculant survival include: method of inoculation (Ordentlich et al., 1987), inoculum density (Postma et al., 1990a), soil type (van Elsas et al., 1989), temperature (Vandenhove et al., 1991), carbon availability (Acea et al., 1988; Wessendorf and Lingens, 1989) and soil water content (Postma et al., 1989). Biotic factors include: physiological state of the inoculant (Vandenhove ef al., 1991), presence of plant roots and the plant age at inoculation (Bashan, 1986; Thompson et al., 1992), competitiveness of the inoculant species (Milus and Rothrock, 1993) and susceptibility of the inoculant to predation (Heijnen et al., 1988). We report the effects of soil moisture, carbon availability and method and density of inoculation on the survival in soil of an inoculant Flavobacterium species.

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JANE L. MAWDSLEY and RICHARD G. BURNS

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MATERIALS AND METHODS

Microorganism

A triple antibiotic resistant (kanamycin, rifampicin and streptomycin) Ffavobuctrium strain, P2.5,was used in all studies. This species, the wild type of which most closely resembles Fiavobactetium baiustinum, was originally isolated by Thompson et ai. (1990). P25 inocuium was grown in nutrient broth at 25°C for 16 h on an orbital shaker (Gallenkamp, U.K.) at 180 rev min-‘. Cells were harvested by centrifugation (15 min, 3500 rev min-’ MSE Centaur 2 Bench Centrifuge), washed twice in sterile phosphate buffer (PI-I ‘7.2)to remove nutrients and resuspended to give an optimal density (600 nm) of 1 (ca 1 x 10’ P25 ml-‘). Plant growth substrata and wheat plants

The sand used in ~otobiotic studies was a low iron type with a mesh size of 40-100 : 1SO-420 pm (Fisons S/0360/63, Loughborough, UK.). Before use, sand was acid washed to remove trace impurities. This involved rinsing once in 1 M HCI and five times in distilled water before drying at 105°C for 24 h. In some studies experiments were carried out in both sterile and non-sterile conditions in order to establish the influence of competition from the indigenous soil microflora on the survival of P25. Sand was sterilized by autoclaving twice at 120°C for 30 min with a 72 h period between sterilizations. A calcareous grassland soil, a silty loam of the Coombe series (12% sand, 55% silt, 33% clay, pII 6.9) with no history of cultivation or agricultural treatment, was used in all studies. Soil was collected from a depth of 15-30 cm (so that only small amounts of roots were present in the soil) and large stones removed before the soil was transported back to the laboratory where it was stored at 4°C in sealed black plastic bags. Before use, the soil was sieved (2.88 mm) to remove the majority of remaining stones. Subsamples of the soil were used to determine field moisture content (0.21-0.26ml g-r depending on sample collection date), soil moisture holding capacity (MHC, 0.60 ml g-‘) and moisture release characteristics, i.e. matric potentials. Winter wheat (Triticum aestivum var. Avalon) ws used in all experiments. Seeds were surface sterilized using the method of Maplestone and Campbell (1989) which involved agitation in 2.5% (w/v) AgNO, for 4 min, washing with an equal volume of 2.5% NaCl to precipitate any remaining AgNOX and rinsing four times in sterile distilled water. Sterilized seeds were germinated on l/l0 strength tryptone soya agar plates at 25°C for 3 days (i.e. until three radicles and the coleoptiie had emerged) to ensure that only viable, sterile seeds were used. Laboratory microcosms: assembiy and sampling tech niques

The apparatus used to monitor P25 population dynamics consisted of a 50 cm3 boiling tube containing

log (dry wt) of soil or sand. In gnotobiotic studies the sterilized sand tubes were plugged with nonabsorbent cotton wool and capped with aluminium foil; non-sterile soil tubes were left unplugged. In rhizosphere studies, tubes were planted with a single, surface-sterilized wheat seedling at a depth of i cm. In those treatments containing P25, 1 ml of washed cells was applied directly to the seedling immediately after planting and before covering with soil. In non-planted soil the inoculum was added directly to the surface. In studies investigating the effect of carbon amendments on P25 survival, P25 was added as I ml of a suspension in 0.5% (w~v) gaiactose, sorbitoi or maltose; this resulted in a final sugar concentration of 0.5 mg g-’ soil. Unless otherwise stated, the soil or sand was maintained at 60% MHC (-20.6 kPa). In sterile sand tubes this was achieved using sterile i/2 strength Hoagiund’s solution which also provided the plants with the necessary nutrient supply. In non-sterile soil distilled water was used. All tubes were kept in a plant growth chamber with a light intensity of 4200 ix, a light dark cycle of 16-8 h and a constant temperature of 20°C. Rhizosphere counts were determined by adding a 10 ml aliquot of sterile phosphate buffer (pH 7.2) to each boiling tube and mixing the entire contents (sand plus roots or soil plus roots) for 1 min on a vortex mixer before the washed root was removed. The remaining soil suspension was used to prepare a IO-fold serial dilution in phosphate buffer. Using the method of Miles and Misra (1938) dilutions were plated onto nutrient agar supplemented with P25 selective antibiotics kanamycin (50 fig ml-‘), rifampic~n (100 gg ml-‘) and streptomycin (250 pg ml-‘) and cycloheximide (50 pg ml-‘). The drops were allowed to dry at room temperature (20 & 2‘C) and the plates were kept at 25°C for 48 h. Rhizopiane counts were obtained by transferring the washed root to a universal containing 10 ml sterile phosphate buffer and macerating using a Polytron homogenizer, speed, 5, 2 x 30 s (Polytron Kinematica AG, Littau, Lucerne, Switzerland). The resulting suspension was diluted and plated as for the rhizosphere samples. Sampling of microcosms was more frequent during the early stages of each experiment than at later stages in order to detect any rapid changes in P25 numbers following inoculation. On each occasion three replicate tubes were sampled and at least eight drops plated per dilution. Ail data are presented as the means of diiutions from replicate tubes and are shown with bars representing the standard error of the mean. The significance of the difference between two treatment means was assessed using Student’s t-test. In order to allow comparisons between the rates of survival of bacteria in different treatments, survival rates of inocula were expressed as the time in days taken for a 90% (t90). 99% (t99) or 99.9% (t99.9) decline in numbers (Thompson et ai., 1990, 1992).

Rhizosphere

survival

RESULTS

Effect of inoculation method on Flavobacterium P25 survival in non -planted and rhizosphere sand and soil

The introduction of FIavobacterium P25 (9.1 x to the surface of non-planted sand was followed by a rapid decline in numbers so that 72 h after inoculation no viable cells were detected. However, P25 added to sand containing developing wheat plants showed a decrease in numbers at day 1 (5.38 x lo4 and 2.53 x IO5 P25 gg’ in homogenized and seedling inoculated samples, respectively) followed by an increase to between 1 x IO’ and 1 x 10’ at day 7; thereafter numbers were constant. The two methods of inoculation were compared but, following the initial increase in numbers between day 1 and day 7 (which was significantly faster when the inoculum was applied directly to the seedling rather than being homogenized throughout the sand prior to planting), there was no difference between them. Similarly, with rhizoplane populations, an increase in numbers to a maximum of between 1 x 10’ and 1 x 1O’cfu gg’ root was measured following inoculation but there was no overall differences in survival between the two treatments. Studies using non-sterile soil gave quite different results. After the first 7 days, P25 survival in 106g-‘)

14

7

14

of a bacterial

inoculant

851

the rhizosphere of developing wheat plants was significantly (P < 0.01) greater than in non-planted soil (Fig. 1). Prior to this, the soil could not be described accurately as rhizosphere because the wheat roots had not grown sufficiently. However, as these experiments were predominantly concerned with final survival rather than initial survival, the transition from non-rhizosphere to rhizosphere was not considered. The application of P25 directly to the planted seedling [hereafter P25 (seedling)] or to the soil surface [hereafter P25 (soil surface)] also led to increased survival rates in relation to those recorded following homogenization of the inoculum in the soil prior to planting [hereafter P25 (homogenized)] (Fig. 1). In non-planted soil experiments P25 (homogenized) declined below quantifiable limits (i.e. < 1 x lO’g_’ soil) within 4 days; whereas P25 (soil surface) was recorded at a density of 2.26 x 10’ gg ’ at the end of the 35-day study. Similarly, in planted soil experiments, P25 (seedling) numbers were always significantly (P < 0.005) greater than P25 (homogenized). The method of inoculation also affected the survival of P25 on the rhizoplane (Fig. 2). P25 (seedling) was recorded in significantly (P < 0.005) higher numbers on days 1,7,21, and 35 and P25 (homogenized). Perhaps more important though, is that P25

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28

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Time after inoculation (days) Fig. 1. Effect of inoculation method on Fluuobacferium P25 survival in non-planted soil and in the wheat rhizosphere. P25 added directly to the planted seedling (0). P25 homogenized throughout the soil prior to planting (m), P25 added to the surface of non-planted soil (0) P25 homogenized throughout non-planted soil (0). BQL = below quantifiable limits (i.e. < 1 x IO’cfu g-’ soil). Bars represent k standard error. SBBx17-o

JANEL. MAWDSLEY and RICHARDG. BURNS

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0 7

14

21

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35

Time after inoculation (days) Fig. 2. Effect of inoculation method on FlauobacteriumP25 survival on the rhizoplane of wheat grown

in non-sterile soil. P25 added directly to the planted seedling (0). P25 homogenized throughout the soil prior to planting (M). Bars represent + standard error.

(seedling) counts were constant over the course of the experiment. In contrast, the rhizoplane counts obtained from the P25 (homogenized) treatment showed a much greater degree of variation. In subsequent experiments the P25 (seedling) method of inoculation was used because of the higher survival rates and the lower degree of variability. Efect of inoculum density on Flavobacterium P2.5 survival

wheat with Flavobacbetween 8.7 x 10) and 8.7 x lo8 cfu g-’ sand was followed, in all cases, by a decline in numbers after 24 h. However, between day 1 and day 7 numbers increased, the extent of which was inversely related to the initial density (Fig. 3). From day 7 onwards there was no relationship between the numbers of bacteria applied and those detected, although at some time points there was a significant difference in the numbers of P25 between different treatments. Similar changes in numbers were recorded on the rhizoplane with the lowest initial P25 concentrations again showing the greatest increases. In most cases the introduction of P25 to non-sterile soil at densities between 1.1 x IO4 and 1.1 x 109g-’ was followed by an increase in numbers and then a general decline, the rate of which was initially rapid but decreased as numbers became constant (Fig. 4). Inoculation

of gnotobiotic

terium P25 at a range of concentrations

Although the survival rates were lower for the higher inoculum densities (e.g. t99.9 was 16 d for 1 x 10’ vs 41 d for I x 10”) the final numbers were still greater in the treatments receiving the higher inoculum density. At the final sampling date, P25 was not detected in the rhizospheres of plants inoculated at a density of 1 x lo6 or below, Similar relationships were recorded between inoculum density and survival of rhizoplane populations. Again, although survival rate increased with decreasing inoculum size (e.g. t99.9 = 21 d for 1 x IO9inoculum vs >54d for I x 106), P25 survival was significantly (P < 0.05) greater in the treatments receiving the higher inoculum densities. Efect of soil water content on JIavobacterium P25 survival Flavobacterium P25 survival was influenced by soil water content. The first study, which investigated survival in soils maintained at MHCs of between 45 and 100% (-73 and -0 kPa), showed that P25 was highly sensitive to increases in water content (Fig. 5). For example, survival was significantly (P < 0.001) greater at 45% MHC after 7 d and 50% MHC (- 52 kPa) after 11d than in soil held at any of the higher MHCs. The second phase of the study showed that inoculant survival was reduced when soils were dried. Survival of P25 inoculated into soils which had been

Rhizosphere survival of a bacterial inoculant dried from field wetness (40% MHC; - 100 kPa) to MHCs of 1I, 19.9 or 30.67% was less than in soils at 40 and 45% MHC (Fig. 6); in addition, survival rates decreased as MHCs were reduced, e.g. t99.9 = 12.2 d at 45% MHC compared with 1 d at 1I % MHC. Effect of carbon amendments on Flavobacterium P25 survival and total bacterial populations The effect of soil amendment with galactose, maltose or sorbitol (compounds identified as promoting growth of P25 in vitrodata not presented) on the survival of P25 and the numbers of total bacteria in both non-planted and rhizosphere soil was investigated. All three sugars, but particularly sorbitol, increased the survival of P25 in non-planted soil (Fig. 7). In amended rhizosphere soil, significant (P < 0.001) increases in the survival of P25 were recorded over the first 7 days of the study but, as the plant developed, these became less pronounced and by day 35 survival in the three treatments was not significantly different from that in non-amended soils (Fig. 8). The addition of a readily-metabolizable carbon source to non-planted soil produced the expected microbial flush so that within 24 h of amendment populations of indigenous bacteria were significantly greater (5.6 x IO9 vs 2.1 x 108g-’ soil) than in non-amended soil. This difference was maintained over the 35 d study. In rhizosphere soil the response of the indigenous bacteria to carbon amend-

853

ments was less marked and only sorbitol a significant increase in numbers.

gave rise to

DISCUSSION

Increased growth rates and survival rates of microorganisms in the rhizosphere (as opposed to nonplanted soil) are reported frequently and bacteria are considered to show the greatest response (Curl and Truelove, 1986). Flavobacterium P25 survival in both sand and soil rhizospheres was significantly greater than in non-planted sand or soil. In unplanted sand P25 fell rapidly (<4d) below quantifiable limits but in rhizosphere sand P25 proliferated, indicating the ability of the inoculant to utilize some of the many carbon and energy sources contained in root exudates and secretions. In non-sterile soil a decline in the numbers of P25 was recorded following inoculation but survival rate and level of survival was greater in rhizosphere than in non-planted soil. The increase in readily-assimilable organic matter in the rhizosphere results in rapid microbial growth in comparison to the bulk soil (Johannson, 1992). In addition to stimulating indigenous microbial populations, the increase in available nutrients has also been demonstrated to promote survival of introduced bacteria (Thompson ef al., 1992; Heijnen et al., 1993). However, not all bacterial species behave in the same way. Yeung et al. (1989) investigated the growth of

11 ZlO _ 8 L i

9-

a8z 't7m CO6ES%'e ::3 E 12l21

01 Time after inoculation (days)

Fig. 3. Effect of inoculum density on FluoobacferiumP25 survival in the rhizosphere of wheat grown in sterilized sand. P25 inoculated at 8.7 x IO* (H). 8.7 x IO’(A), 8.7 x IO6 (@), 8.7 x IO5 (0). 8.7 x IO4 (A), or 8.7 x IO3 (0) cfu g-’ sand. Bars represent + standard error.

JANEL. MAWDSLEY and RICHARDG. BURNS

II,

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I

1;

2;

iI

Time after inoculation (days) Fig. 4. Effect of inoculum density on survival of Fluvobacterium P25 in the rhizosphere of wheat grown in non-sterile soil. P25 inoculated at 1.1 x lo9 (m), 1.1 x IO* (A), 1.1 x 10’ (a), I.1 x IO6 (O), 1.1 x 10’ (A), 1.1 x lo4 (0) cfu gg’ soil. BQL = below quantifiable limits (i.e. < 1 x 10Zcfu g-’ soil). Bars represent k standard error.

six Pseudomonas aeruginosa and P. putida strains in both non-planted and tomato rhizosphere soil. With one exception, inoculant survival was less in rhizosphere than in non-rhizosphere soil. In addition to the influence of plant roots, our results show that the method of inoculation affects inoculant survival in soil but not in sterile sand. In sand treatments, although up to day 7 the survival of P25 in rhizosphere and rhizoplane samples was greater following introduction directly to the seedling than following sand inoculation, over the remainder of the study there was no difference between the two treatments. In sterile sand, where competition has been eliminated but nutrient levels are minimal, the P25 (homogenized) survived (5.38 x IO4gg’) for the short period (~3 d) until contact was made with the developing root. However, in non-planted soil, P25 (homogenized) rapidly decreased in numbers and fell below quantifiable limits within 4 days whereas P25 (soil surface) was still detected 35 days after inoculation. Thus the survival of P25 is much reduced when mixed throughout the soil in comparison with surface application. It seems unlikely that this reduction in survival on incorporation is a response to nutrient limitation because the higher densities per unit area of P25 on the soil surface would be expected to become starved more rapidly. It is more likely that distributing P25 throughout the soil brings the inoc-

ulant into contact with a greater number of indigenous microbial competitors and predators. Many studies (e.g. Heijnen et al., 1988, 1992; Postma et al., 1990b; Foster and Dormaar, 1991) have shown that the spatial distribution of bacteria in relation to clay particles and microaggregates and their position within micropores can affect the degree of predation. In addition, it is possible that oxygen diffusion may be restricted and that this reduced the survival of the obligately aerobic P25 when mixed throughout the soil. The contrast between the two application methods were less marked in rhizosphere soil, although counts recorded for P25 (seedling) were still significantly greater than P25 (homogenized). On the rhizoplane, at most sampling dates, densities of P25 (homogenized) were lower than those of P25 (seedling) and showed a greater degree of variability: a disadvantage if relatively constant population densities are required to achieve beneficial effects (Iswandi et al., 1987). The method of introducing the inoculant to plant and soil can be crucial in determining its success (Ordentlich et al., 1987). However, when evaluating different methods it is important to consider whether they can be adapted to large scale use and their compatibility with agricultural practices. Possible delivery systems for inoculants include seed coating (McQuilken et al., 1990) synthetic carriers (van Elsas

Rhizosphere survival of a bacterial inoculant ef al., 1992) and in-furrow spraying (Zablotowicz et al., 1991). P25 inoculum density was not correlated with survival in the rhizosphere of gnotobiotic wheat. These results correspond to results obtained in other gnotobiotic studies. For example, Bennett and Lynch (1981) investigated the effect of inoculum density on the growth of three bacterial species (Pseudomonas sp., Mycoplana sp. and Curtobacterium sp.) in the rhizosphere of barley (Hordeum vuigare). The bacteria were inoculated at densities of ca 1 x 103, 1 x lo5 and 1 x IO’cfu gg’ root but, in all cases, final (day 8) population sizes were ca 5 x 10’. They concluded that each plant could support a maximum population, a concept they termed ‘colonization potential’, and that this was independent of inoculum density. Similar results were obtained by Postma et al. (1990a) following introduction Rhizobium leguminosarum br. trifolii into non-planted sterilized soil. These authors reported that final population sizes of ca 5 x 10’ after 30 d (loamy sand) and 2.5 x 10’ after 14d (silt loam) were achieved and maintained over the 50 d study regardless of the inoculum density and concluded that colonization potential reflects the capacity of the soil with regard to ‘habitable space’ (i.e. the surface area suitable for bacterial colonization), matric potential and the presence of appropriate substrates. However, in our work using non-sterile soil, the

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survival of P25 both in the rhizosphere and on the rhizoplane was affected by inoculum density. Similar results were obtained by Dupler and Baker (1984) following introduction of P. putida at 1 x 10’ and 1 x IO’cfu gg’ soil, and by Wessendorf and Lingens (1989) using P. Juorescens at 1 x IO’, 1 x IO’ and 1 x 109cfu gg’ soil. In both cases, survival was greatest at the highest inoculum density. In a study by Hebbar et al. (1992) that was comparable in methodology to ours, P. cepacia was introduced to seeds and populations monitored in both rhizosphere and rhizoplane samples. Higher numbers of the inoculant were recorded following introduction at 7 x lo5 per seed than at 7 x IO3 or 6 x 10’. However, this difference was only significant over the first 15 days of the study; at subsequent sampling dates (up to 60 days) survival was independent of inoculum density. However, not all studies have shown inoculum density to have a significant effect on survival (e.g. Bashan, 1986; Iswandi et al., 1987; Ordenthch et al., 1987; Elliott-Juhnke et al., 1989) indicating the need for individual studies of each putative inoculant. Survival of P25 was greatest when applied to soils with a MHC of between 40 and 50% (- 100 to -52 kPa). As water availability is decreased the dimensions and number of water filled pores and water films decrease (Griffin, 1981). Consequently, soil bacteria which are restricted to such films, will be

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Time after inoculation (days) Fig. 5. Effect of soil water content on FluuobucteriumP25 survival at 45% (W), 50% (A), 65% (O), 75% (n), 85% (A) and 100% (0) MHC. BQL = below quantifiable limits (i.e. < 1 x IO’cfu g-’ soil). Bars

represent f standard error.

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JANE L. MAWDSLEYand RICHARDG. BURNS

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Time after

inoculation

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52

(days)

Fig. 6. Effect of soil water content on FlavobacferiumP25 survival at 45% (m), 40% (A), 30% (a), 20% (O), and 11% (A) MHC. BQL = below quantifiable limits (i.e. ~1 x 102cfu g-’ soil). Bars rep-

resent + standard error. limited in both growth (due to a reduction in the area from which nutrients can be utilized) and movement (Griffin and Quail, 1968). These factors undoubtedly contribute to the poor performance of inoculants in dry soils (Rattray et al., 1992). Conversely, soil saturation will also affect the survival of obligately aerobic inoculants such as P25. The literature suggests inoculants will react in different ways to changes in soil water content. Postma et al. (1989) investigated survival of R. leguminosarum bv. trifolii in a loamy sand and a silt loam at different water contents. In both soils greatest inoculant survival occurred at the lowest initial moisture contents. In contrast, Rattray et al. (1992) showed that survival (and microbial activity) of Escherichiu coli in a sand loam soil held at - 5, - 64 and - 1.5 MPa decreased with decreasing water content. A similar relationship was reported by Dupler and Baker (1984) following introduction of P. putidu into soil at -0.3, -2, -15 and -100 bars (-33, -200, - 1.5 and - 10 MPa). Within 3 days of inoculation the bacteria added to the soil at - 100 bars had fallen below quantifiable limits. However, when soil at - 15 bars was slowly dried to - 100 bars over a period of 5 weeks, > 5 x 10’ P. putidu g-l soil were recorded. This suggests that some inoculants survive soil dehydration if this takes place gradually. Various theories have been presented to explain the different abilities of bacteria to withstand desiccation. Studies by Chen and Alexander (1973) suggest that

bacteria with a high internal osmotic tension are better able to survive conditions of water stress whilst certain bacterial species may undergo morphological adaptation in order to enhance their survival. For example, Labeda et al. (1976) have shown that Arthrobacter globiformis assumes a cocco-bacillus state when dried and that this is resistant to soil drying. Exopolysaccharide (EPS) production may be an important factor determining the ability of bacteria to survive desiccating conditions (Roberson and Firestone, 1992). The ability of an inoculant species to utilize one or more of the many compounds produced by plant roots would give it a competitive advantage over the indigenous population. However, it seems unlikely that stimulation of one particular species would occur unless the inoculant is chosen especially for its capacity to metabohse a recalcitrant root exudate (van Elsas, 1992). Alternatively, inoculant survival rates may be increased by incorporating a specific substrate in the delivery formulation (Klein and Casida, 1967; Lynch, 1989). In our studies, the addition of simple carbohydrates to planted and non-planted soil enhanced not only the survival of P25 but also increased the indigenous bacterial population. In non-planted soil the carbon amendments confirmed that carbon limitation was at least partly responsible for decreases in P25 numbers following inoculation. This is reinforced by the rhizosphere data which shows that the in-

Rhizosphere survival of a bacterial inocculant

;

1;

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2;

3;

Time after inwuiatian [days] Fig. 7, ESect of galaetose (a), Fkzv~buctrium

maltose (A) and sorbitol I(*) amendments

on the survival of

P25 in non-planted soil in comparison to non-amended soil f+)” Bars represent rfrstandard

error,

Fig. 8, Etrect of gatactose (m), maltose (A) and sorbitol (e) amendments on the survival of Flavobacterium P25 in rkzosphere soii in comparison to non-amended soil {f )+Bars represent + standard error.

JANE L. MAWDSLEYand RICHARDCi. BURNS

858 creased

numbers of P25 in carbon amended treatments only occurred during the first 14 days. It is probable that the excess carbon has been depleted and that root secretions have increased thereby decreasing the difference in substrate concentrations between amended and non-amended soils. Acknowledgement-This research was dentship from the Natural Environment

funded by a stuRestarch Council.

REFERENCES

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