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Enumeration of wax-degrading microorganisms in water repellent soils using a miniaturised Most-Probable-Number method.

Introduction

Water repellency is mostly associated with sandy soils and affects more than 5 million hectares of agricultural land in western and southern Australia (Tate et al. 1989). Repellency is caused by hydrophobic substances on the surfaces of the sand grains (DeBano et al. 1970; Roberts and Carbon 1972; Franco et al. 1995), and plant waxes and their biodegradation products are the main contributors to the phenomenon (Roberts and Carbon 1972; McGhie and Posner 1980, 1981; Tate et al. 1989; Franco et al. 1995; Doerr et al. 2000). Analysis of organic materials responsible for water repellency indicated that the main components are branched and unbranched [C.sub.16] to [C.sub.36] fatty acids, and their esters, alkanes, alcohols, and sterols (Ma'shum et al. 1988; Franco et al. 2000a). Roper (1998, 2004) isolated wax-degrading bacteria from soils (including water repellent soils) and showed that these bacteria produce biosurfactants which appear to be responsible for their potential to degrade waxes that cause water repellency. The capacity for bioremediation of water repellent soils by wax-degrading bacteria is likely to be related to the size of the populations of wax-degrading bacteria. Therefore, the development of any management options to remediate water repellent soils requires an understanding of the seasonal dynamics of populations of wax-degrading bacteria, natural or introduced, and the factors that regulate their size and activity.

To our knowledge, no methods for measuring populations of wax-degrading bacteria in natural soils exist. Previously, agar plates with wool wax on the surface were used to select and grow wax-degrading bacteria from soils (Roper 2004), but the distribution of the wax on the plate was generally uneven and unsuitable for reliable and reproducible enumeration of populations of these bacteria. A Most Probable Number (MPN) method offers the opportunity to measure the size of a particular microbial population using a specific characteristic (Alexander 1982, Gupta and Roper 1994). The MPN technique is based on a determination of the presence or absence of microorganisms in several replicates of consecutive dilutions of soil and requires that the population being measured brings about some readily recognisable characteristic or transformation in the medium (Alexander 1982). In this case it combines both growth and the ability to perform a function (wax-degradation) by the population. This paper reports on the development of such a MPN method designed specifically to select and count wax-degrading microorganisms in soils based on their ability to metabolise waxes.

Materials and methods

Measurement of water repellency

Water repellency of soil was measured using the MED test (King 1981). Droplets of aqueous ethanol in the range of 0-5 M are placed on the surface of soils dried (105[degrees]C for 48h) and cooled to room temperature (20[degrees]C) and the entry time is recorded. The repellency of soils is determined by the molarity of the ethanol drop (MED) that enters the soil within 10s. Wetting soils have a MED of 0 and as the soils become more repellent the MED value increases. Severely repellent soils have a MED [greater than or equal to] 4.

Most-Probable-Number Method

The MPN method was adapted from a "Sheen Screen' method developed by Brown and Braddock (1990). Tissue culture plates (100mm square with 25 wells) were used. To each well a 2-mL aqueous solution of sterile mineral salts (g/L: NaN[O.sub.3], 2: [K.sub.2]HP[O.sub.4], 1; K[H.sub.2]P[O.sub.4], 0.5: KCl, 0.1: MgS[O.sub.4].7[H.sub.2]0, 0.5: Ca[Cl.sub.2], 0.01: FeS[O.sub.4].7[H.sub.2]0, 0.01: Na EDTA, 0.0015 (Akit et al. 1981) + yeast extract, 3) was added using a sell:refilling dispensing syringe (Socorex). Decimal dilutions (up to [10.sup.-6]) of soil in 0.1% saline containing Tween 80 (0.01% v/v; Sigma) to disperse the microorganisms were prepared; 0.1 mL of each dilution was added to 5 replicate wells in a column on each plate. One column of 5 wells was left uninoculated as a control and the remaining 4 columns were inoculated with 4 dilutions of a single sample (Fig. 1). Coconut oil or hydrocarbon (5 [micro]L) containing carotene as a colourant was added to each well to form a sheen. The plates were incubated at 30 C and observed daily: 30C was the preferred temperature for growth of wax-degrading bacteria isolated from soils and other sources by Roper (20(14). The plates were read at 5 days and wells were scored as positive when oil/hydrocarbon was emulsified due to surfactant production (indicated by disruption of the sheen) (Fig. 1). The MPN of wax-degrading bacteria was determined using a table of Most Probable Numbers (Alexander 1982) or an Excel based computer programme (R Correll et al., CSIRO, unpublished).

[FIGURE 1 OMITTED]

Selection and preparation of oil/rocarbon

Coconut oil was chosen as the carbon source for the wax-degrading bacteria. Analysis of coconut oil by the suppliers (Meadow Lea Foods, Sydney, Australia) indicated that it contained [C.sub.8] to [C.sub.20] chains including fatty acids, with [C.sub.12] [C.sub.14], [C.sub.16], and [C.sub.18] predominant. In addition it had a melting point of 25 C, making it suitable to form a sheen at the incubation temperature of 30[degrees]C. A few drops of [beta]-carotene (Meadow Lea Foods, Australia) were added to colour the coconut oil (20 mL) to a bright yellow. The mixture was sterilised by autoclaving at 100 kPa (121[degrees]C) for 20 mm prior to use. The composition of coconut oil was unchanged by autoclaving since the boiling point of all of its fatty acid components was >200[degrees]C (Noller 1966).

Hexadecane ([C.sub.16] hydrocarbon) was tested as an alternative source of carbon in the MPN assay. With a melting point of 18.5[degrees]C it was also suitable for producing a sheen in the wells. Hexadecane was sterilised by filtration through a 0.22-[micro]m Millex filter (Millipore, Massachusetts, USA). Autoclaved [beta]-carotene was used to colour the hexadecane as for the coconut oil.

Growth of/emulsification by pure cultures of wax-degrading bacteria in mineral salts solution with coconut oil or hexadecane as a carbon source

Fourteen isolates from a collection of soil bacteria, selected on the basis of their ability to grow on wax as a sole carbon source (Roper 2004), were tested for their ability to emulsify coconut oil and/or hexadecane. The cultures were grown on glucose mineral salts (GMS) agar, which contained mineral salts (Akit et al. 1981) and yeast extract (3 g/L), glucose (20 g/L) as a carbon source, and agar (GIBCO BRL) (15 g/L). A single colony from each culture was suspended in 9 mL of the same mineral salts solution, and 50 [micro]L of each bacterial suspension was added to 5 wells of tissue culture plates containing 2 mL mineral salts with yeast extract as used in the MPN determinations. Either coconut oil or hexadecane was added as described above. The ability of the cultures to grow and/or emulsify coconut oil or hexadecane was observed alter 5 days.

Comparison of the MPN method with agar plate counts of known wax-degrading bacteria

The efficiency of the MPN method was tested using pure cultures by comparing the MPN with counts determined by agar plating. Isolates from our collection of wax-degrading bacteria were chosen for the comparisons based on strong growth and significant biosurfactant production on hexadecane (Roper 2004). Comparisons of the 2 methods were done with either (i) pure culture suspensions alone or (ii) sterile soils inoculated with pure cultures.

Evaluation using pure culture suspensions

Four isolates (66b, 731, 73ww, 83wwl) all belonging to the genus Rhodococcus were used for this comparison. Cultures of each isolate were grown in a 250-mL Erlenmeyer flask containing 100 mL of mineral salts solution (Akit et al. 1981) with yeast extract (3 g/L) and glucose (20 g/L). The cultures were incubated for 4 days at 30[degrees]C on a rotary shaker (New Brnuswick Innova 4300), after which counts by MPN (as described above) and plate counting were done. The same dilution series was used for both the MPN and the plate counts.

Plate counts were prepared by aseptically dispensing, in triplicate, 50-[micro]L aliquots of each dilution onto GMS agar. The plates were incubated at 30[degrees]C for 3-5 days after which counts were made. Counts are presented as the average of the 3 replicates.

Evaluation using pure cultures inoculated into soil

Three isolates (36a, Nocardia sp.; 77ww 1, Mycobacterium sp.; 66b, Rhodococcus sp.) were used for this comparison. Cultures of each isolate were grown m shake culture as described in the previous section; 5 mL of broth culture was added to 15 g of sterile (autoclaved at 100 kPa (121[degrees]) for 60 min) non-wetting soil (grey sand, pH(Ca[Cl.sub.2]) 5.0, soil classification Ucl.2: Northcote el al. 1975) in a tube and mixed thoroughly. The tube was closed with a sterile cotton wool plug and the inoculated soils were left to dry at room temperature for several weeks. Samples (0.1 g) from each tube were then suspended in 9.9 mL of saline solution (0.1%), mixed on a vortex mixer for 2 min and diluted in a 10-fold dilution series to [10.sup.-9]. The inoculants that were recovered were enumerated using plate counting and MPN methods as above,

MPN of wax-degrading bacteria in natural soils containing mixed populations of microorganisms

Reproducibility of MPN method (25-well plates)

The reproducibility of the MPN method was tested by multiple MPN determinations of subsamples of soils collected from 3 different suburban locations in Perth (31[degrees] 57' S, 115[degrees] 51' E), Western Australia.

Soil 1. Pale-coloured siliceous sand (Ucl.21; Northcote et al. 1975) with black organic matter, collected from underneath a layer of pine/eucalypt bark chips in a suburban garden. The soil was well watered and maintained in a moist condition. It had received granulated garden fertiliser (Cresco, CSBP. Perth, WA, Australia (N : P : K : S 12:3.4:10:11.8 + trace elements)) at the recommended rate of 50 g/[m.sup.2] about 3 weeks prior to collection. This soil was moist on collection but when dried it was moderately repellent with a MED 3. The pH in water was 6.5.

Soil 2. Grey-brown loamy sand (Uc4.2) collected from a bushland nature reserve. Soil was dry on collection but wettable (MED 0). The pH in water was 4.7.

Soil 3. Pale-coloured siliceous sand (Ucl.21) with black organic matter, collected from underneath a stand of eucalypt trees. The soil was dry when collected and severely non-wetting (MED 4.2). The pH in water was 5.8.

Each soil was subsampled into 3 parts (A, B, C). For each subsample 10 g soil was added to 90 mL 0.1% saline and serial dilutions prepared to [10.sup-8]; 0.1 mL of each dilution was added to 5 wells in a column on a MPN plate. Each MPN plate was replicated 5 times for each soil. After 5 days incubation at 30[degrees]C, MPN counts were determined and reproducibility was assessed using 'coefficient of variation' (CV) calculations.

Miniaturised MPN method (96-well plates)

In order to evaluate the sensitivity and accuracy of the MPN method, 25-well plates were compared with 96-well plates as a means of testing a larger number of samples with smaller dilution steps. In the 96-well plates each dilution was inoculated into 8 wells, whereas only 5 wells were inoculated for each dilution in the 25-well plates. The 96-well plates were Sarstedt microtest 96 'U' well with lid, sterile and individually wrapped, well volume 350[micro]L To each well (except the second column), 200[micro]L of sterile mineral salts (Akit et al. 1981) with yeast extract (3 g/L) was added using a multichannel electronic pipettor (50 1200[micro]L). Column I was a nil control. Into the second column 250[micro]L of soil suspension (prepared as in the previous section rising the 3 natural soils) was added to each well. Serial dilutions (1 in 5) were then prepared by transferring 50 [micro]L from this column to the next and so on through to the 12th column using the multichannel pipettor. (The contents of each well were mixed by pipetting liquid tip and down 2-3 times.) Liquid (50 [micro]L) was removed from column 12 wells to leave the same volume as the others. Coconut oil (~ 15 [micro]L) containing carotene as a colourant was added aseptically to each well to form a sheen. The plates were incubated at 30[degrees]C and observed daily. The plates were read at 5 days and wells were scored as positive when oil was emulsified. MPN counts were compared with those obtained using the 25-well plates.

Statistical analyses

The results were analysed using the statistical package GENSTAT (version 7.1, Lawes Agricultural Trust, Rothamsted Experimental Station, UK).

Results

Growth of/emulsification by pure cultures of wax-degrading bacteria in mineral salts solution with coconut oil or hexadecane as a carbon source

Growth and emulsification by bacterial isolates inoculated into tissue culture wells containing coconut oil or hexadecane is shown in Table 1. All isolates grew on coconut oil or hexadecane. All 14 isolates emulsified the fatty acids in coconut oil, but only 9 emulsified hexadecane. Subsequently, further evaluations of the MPN method are reported for coconut oil only.

Comparison of the MPN method with agar plate counts of known wax-degrading bacteria

Evaluation using pure culture suspensions

Comparison of counts of pure cultures, by MPN and plating, are shown in Fig. 2 and are represented as [log.sub.10] of the numbers counted. Counts determined by plating and by MPN were similar for each culture. A 1-tailed t-test indicated there was no statistical difference between the 2 counting procedures (P = 0.364).

[FIGURE 2 OMITTED]

Evaluation using pure cultures inoculated into soil

Counts ([log.sub.10]) of bacteria, recovered from sterile soil samples inoculated with pure cultures of known wax-degrading bacteria, are shown in Fig. 3. Counts are represented as averages of 4 replicates (agar plate counting) and 5 replicates (MPN). There was strong agreement between counts determined by the 2 methods. An ANOVA of the data indicated no significant difference between estimates from the MPN method and those from the plate counting procedure (P = 0.640). Within each counting procedure, variation was low [CV range 2-4% (MPN) and 1-2.5% (plate counts)].

[FIGURE 3 OMITTED]

MPN of wax-degrading bacteria in natural soils containing mixed populations of microorganisms

Reproducibility of MPN method

MPN counts of wax-degrading bacteria in soil samples from the 3 sites are presented in Table 2. Numbers of wax-degrading bacteria ranged from [10.sup.3] in Soil 2 up to [10.sup.5] in Soil 1. The results reveal significant differences in the population size of wax-degrading bacteria among each of the 3 soils (ANOVA, P < 0.001). Variation (CV) within each subsample (n = 5) of each soil was low (Soil 1,2-4%; Soil 2, 2-11%; Soil 3, 5-16%) (Table 2). Generally, variation among the 5 replicate analyses within each subsample was < 10%, except for 2 replicates. Similarly the CV between the 3 subsamples of each soil collected was low (Soil 1, 1%; Soil 2, 6%; Soil 3, 1.7%) (Table 2), indicating that the MPN method is consistent and reproducible.

Miniaturised MPN method (96-well plates)

MPN counts determined using 96-well plates are compared with counts on 25-well plates in Table 3. Because of their smaller well-size, observing emulsification in the 96-well plates was more difficult than in the 25-well plates. Despite this, an ANOVA revealed that there was no significant difference between results from the 2 MPN procedures (P = 0.492).

Discussion

A Most Probable Number (MPN) method allows the enumeration of specific groups of microorganisms whose abundance cannot be estimated by conventional plating techniques (Alexander 1982). Unlike the plating techniques using agar media, the MPN method proposed in this study relies on the expression of a specific characteristic that selects for the organisms being counted. Therefore this method is more relevant than plating techniques because it combines the expression of function (in this case wax-degradation) with the ability of the organisms to grow.

Agar plates with wool wax on the surface were used in earlier studies to select for and isolate wax-degrading bacteria (Roper 2004). However, this system could not be used to estimate population sizes because wool wax was solid at the incubation temperature (30[degrees]C), and hence emulsification (a direct measure of wax-degradation) could not be observed. [An incubation temperature of 30[degrees]C was chosen because it was the preferred temperature for growth of wax-degrading bacteria isolated from soils and other sources by Roper (2004).] Growth on wool wax agar plates could not confirm the ability of isolates to perform the function of wax-degradation and therefore counts were meaningless, particularly if opportunistic groups utilising the products of wax degradation were present. Furthermore, the solid wax was unevenly distributed on the surface of the medium resulting in poor reproducibility. The MPN method, oil the other hand. used liquid medium which ensured uniform distribution of the wax substrates. Estimates of numbers of wax-degrading bacteria, determined by their ability to perform the wax-degrading function (emulsification). meant the measurements of population size were truly based on the capability to perform the function rather than on incidental growth on the media.

In order to select for and count potential wax-degrading bacteria in a liquid based system, it was necessary to choose carbon sources with components implicated as causative agents of hydrophobicity in sands. These are branched and unbranched [C.sub.16] to [C.sub.36] fatty acids, and their esters, alkanes, alcohols, and sterols (Ma'shum et al. 1988: Franco et al. 2000a). Coconut oil contained some of the shorter chains known to be responsible for water repellency in soils and was liquid above 25[degrees]C. It formed a sheen on the liquid surface and emulsification of the oil could readily be observed. Hexadecane (with a melting point 18.5[degrees]C) was also considered suitable for forming a sheen.

Pure cultures of known wax-degrading bacteria were used to test the potential of the MPN method. Results presented in this paper showed that all isolates of wax-degrading bacteria grew on coconut oil and hexadecane. However, some isolates did not emulsify hexadecane, whereas all of them emulsified coconut oil (Table 1) and this resulted in slightly lower MPN estimates using hexadecane compared with coconut oil as a C source (data not shown). On this basis coconut oil was chosen for all subsequent evaluations of the MPN method and emulsification of the oil was the criterion for a positive test (Fig. 1). Despite the differences between the 2 carbon sources, MPN estimates using hexadecane were not significantly lower than those estimates using coconut oil (1-tailed t-test, P = 0.466; data not shown).

MPN estimates (using coconut oil) were compared with plate counts using pure cultures of known wax-degrading bacteria. In pure culture suspensions, MPN counts compared favourably to plate counts (Fig. 2). In a separate experiment using sterile soils into which pure cultures of known wax-degrading bacteria had been added some months earlier. MPN counts were statistically similar to plate counts (ANOVA, P = 0.640) (Fig. 3), indicating that the MPN method is an accurate measure of size of the wax-degrading population. Extensive replication of measurements both in the soils containing pure cultures and in the 3 soils collected from different natural habitats indicated that, based on measurements of variation (CV), the method was reproducible for a range of different soil types (Fig. 3, Table 2). Based on its accuracy and reproducibility the MPN method was considered suitable for the determination of the size of populations of wax-degrading bacteria in a range of soils.

It has been suggested that by increasing the number of replicate wells per dilution and/or decreasing the dilution magnitude (i.e. 5-fold instead of 10-fold dilutions) it is possible to increase the sensitivity and accuracy of MPN estimates (Darbyshire et al. 1974; Alexander 1982). For this a 96-well plate system was compared to the 25-well plate system. Comparisons between counts of wax-degrading bacteria in the 3 soils collected from natural habitats made on the 96- and 25-well plates showed similar trends (Table 3) (ANOVA. P = 0.492). While there was potential to increase the sensitivity of the MPN measurement using 96-well plates, there was some difficulty in observing emulsification in the much smaller wells on these plates and hence a 25-well system was preferred.

MPN counts of the 3 natural soils showed significant differences in the size of the populations of wax-degrading bacteria (P < 0.001). The 2 non-wetting soils (Soil 1 and Soil 3, both Ucl.21; Northcote et al. 1975) with coatings of wax on their particle surfaces (Franco el al. 1995) contained significantly more wax-degrading bacteria than the naturally wettable Soil 2. In the water repellent Soil 1, which had been maintained in a moist condition in a suburban garden for ~12 months, thus favouring microbial activity, the numbers of wax-degrading bacteria were higher than in the dryland water repellent Soil 3 by an order of magnitude. Soil 1 had also received granular fertiliser about 3 weeks before sampling. Franco et al. (2000b) demonstrated that fertilisers stimulate indigenous wax-degrading microorganisms to reduce water repellency and this may further explain the larger numbers of wax-degrading bacteria counted in this soil as well as the lowered level of repellency (MED 3) compared with the similar soil type ('Soil 3', MED 4.2).

The MPN method suggested in this paper is a simple, accurate, rapid, and cost-effective means of measuring wax degradation. Emulsification of coconut oil is clear and easy to identify allowing positive and negative wells to be readily distinguished (Fig. 1). Thus the proposed method is useful for monitoring populations of wax-degrading bacteria in soils and therefore is likely to be useful for researching the potential for biological remediation and the conditions that favour it. Water repellent soils contain populations of wax-degrading bacteria (Roper 2004) with the ability to produce biosurfactant molecules that facilitate their use of hydrophobic compounds (Lang and Philp 1998: Roper 2004). However, conditions do not always favour their activity, especially when repellency delays 'wetting up' of the soil soon after opening rains when soil temperatures are still high enough to support microbial activity. Measuring populations of wax-degrading bacteria early in the season may identify soils which have the potential to degrade waxes and could benefit from short-term remediation measures, e.g. fertiliser application (Franco et al. 2000b) or the application of soil wetting agents (Wallis and Home 1992). Measuring water repellency (MED) gives only a snapshot of repellency at any particular time, but no indication of the capacity for biological remediation. From a longer term perspective, monitoring populations of wax-degrading bacteria and water repellence (MED) at different times of the year and under different agricultural managements may provide clues for the development of new management systems that promote populations of this group of bacteria and minimise the risk of water repellency in agricultural soils.
Table 1. Growth of (G+/-) and emulsification by (E+/-) cultures of
known wax-degrading bacteria inoculated into tissue culture wells
containing coconut oil or hexadecane

Details of identification are given in Roper (2004)

Culture Identification Coconut oil Hexadecane

8a Streptomyces sp. G+ E+ G+ E+
36a Nocardia sp. G+ E+ G+ E+
66b Rhodococcus sp. G+ E+ G+ E+
71d Achromobacter G+ E+ G+ E-
72.1a Unidentified G+ E+ G+ E-
73a Rhodococcus sp. G+ E+ G+ E+
73ww Rhodococcus sp. G+ E+ G+ E+
74b Mycobacterium sp. G+ E+ G+ E+
77ww1 Mycobacterium sp. G+ E+ G+ E+
83ww1 Rhodococcus sp. G+ E+ G+ E+
83ww2 Unidentified G+ E+ G+ E-
85b Rhodococcus sp. G+ E+ G+ E+
91ww1 Ochrobactrum sp. G+ E+ G+ E-
109a Unidentified G+ E+ G+ E-

Table 2. Reproducibility of the Most Probable Number (MPN)
method--MPN of wax-degrading bacteria in natural soils containing
mixed populations of microorganisms

 Reproducibility within subsamples (n = 5)

 Mean subsample
 counts Mean s.d.
Subsample ([10.sup.3]/g [log.sub.10] [log.sub.10] CV%
 soil) counts counts

 Soil 1(A)

A 101.4 4.97 0.20 3.97
C 114.6 5.05 0.13 2.61

 Soil 2

A 1.29 3.03 0.33 10.98
B 0.59 2.75 0.14 5.24
C 1.22 3.08 0.07 2.13

 Soil 3

A 5.66 3.70 0.22 5.93
B 14.66 3.75 0.61 16.16
C 8.29 3.82 0.31 8.04

 s.e.
Subsample [log.sub.10]
 counts

 Soil 1(A)

A 0.09
C 0.06

 Soil 2

A 0.15
B 0.06
C 0.03

 Soil 3

A 0.10
B 0.27
C 0.14

Reproducibility between subsamples (n = 2, Soil 1;
n = 3, Soils 2 & 3)

 Sample mean Mean s.d.
([10.sub.3/g soil] [log.sub.10] counts [log.sub.10] counts

 Soil 1

 108.0 5.01 0.05

 Soil 2

 1.03 2.95 0.18

 Soil 3

 9.53 3.76 0.06

 Sample mean s.e.
([10.sub.3/g soil] CV% [log.sub.10] counts

 Soil 1

 108.0 1.07 0.05

 Soil 2

 1.03 6.06 0.10

 Soil 3

 9.53 1.65 0.04

(A) Subsample B not determined.

Table 3. Comparison of Most Probable Number (MPN) estimates
of wax-degrading bacteria using 96- and 25-well plates in natural
soils containing mixed populations of microorganisms

ANOVA: there was no statistical difference between estimates
determined by the 2 MPN procedures (P = 0.492)

 Sample mean (A) Mean s.e.
([10.sup.3]/g soil) [log.sub.10] counts [log.sub.10] counts

 96-well 25-well 96-well 25-well 96-well 25-well

 Soil 1

 154.91 108.0 5.10 5.01 0.21 0.05

 Soil 2

 0.32 1.03 2.49 2.95 0.08 0.10

 Soil 3

 1.50 9.53 3.17 3.76 0.05 0.04

(A) Average of 10 replicates (soil 1) and 15 replicates
(soils 2 and 3) (25-well plates); average of 3 replicates
(96-well plates).


Acknowledgments

This work was supported by GRDC. The authors are grateful to Anne McMurdo for technical assistance. Funding for VVSRG was provided by CSIRO Land and Water.

References

Akit J, Cooper DG, Manninen KI, Zajic JE (1981) Investigation of potential biosurfactant production among phytopathogenic Corynebacteria and related soil microbes. Current Microbiology 6, 145-150.

Alexander M (1982) Most probable number method for microbial populations. In "Methods of soil analysis. Part 2. Chemical and microbiological properties'. Agronomy Monograph No. 9, 2nd edn (Eds AL Page, RH Miller, DR Keency) pp. 815 820. (ASA-SSSA: Madison, WI)

Brown EJ, Braddock JF (1990) Sheen screen, a miniaturised Most-Probable-Number method for enumeration of oil-degrading microorganisms. Applied and Environmental Microbiology 56, 3895-3896.

Darbyshire JF, Wheatley RE, Greaves MP, Inkson RHE (1974) A rapid micromethod for estimating bacterial and protozoan populations in soil. Revue d'Ecologie et de Biologie du Sol 11, 465-475.

DeBano LF, Mann LD, Hamilton DA (1970) Translocation of hydrophobic substances into soil by burning organic litter. Soil Science Society of America Proceedings 34, 130-133.

Doerr SH, Shakesby RA, Walsh RPD (2000) Soil water repellency: its causes, characteristics and hydro-geomorphological significance. Earth-Science Reviews 51. 33-65. doi: 10.1016/S0012-8252 (00)00011-8

Franco CMM, Clarke PJ, Tate ME, Oades JM (2000a) Hydrophobic properties and chemical characterisation of natural water repellent materials in Australian sands. Journal of Hydrology 231-232, 47-58. doi: 10.1016/S0022-1694(00)00182-7

Franco CMM, Michelsen PP. Oades JM (2000h) Amelioration of water repellency: application of slow-release fertilisers to stimulate microbial breakdown of waxes. Journal of Hydrology 231-232, 342-351. doi: 10.1016/S0022-1694(00)00206-7

Franco CMM, Tate ME, Oades JM (1995) Studies on non-wetting sands. The role of intrinsic particulate organic matter in the development of water-repellency in non-wetting sands. Australian Journal of Soil Research 33, 253-263.

Gupta VVSR, Roper MM (1994) A Most-Probable-Number method to enumerate the populations of cellulolytic bacteria and fungi in soils. In 'Soil biota management in sustainable farming systems'. Poster papers. (Ed. CE Pankhurst) pp. 115-117. (CSIRO: Australia)

King PM (1981) Comparison of methods for measuring severity of water repellence of sandy soils and assessment of some factors that affect its measurement. Australian Journal of Soil Research 19, 275-285.

Lang S, Philp JC (1998) Surface-active lipids in rhodococci. Antonie van Leeuwenhoek 74, 59-70. doi: 10.1023/A: 1001799711799

Ma'shum M, Tate ME, Jones GP, Oades JM (1988) Extraction and characterization of water-repellent materials from Australian soils. Journal of Soil Science 39, 99-110.

McGhie DA, Posner AM (1980) Water repellence of a heavy-textured Western Australian surface soil. Australian Journal of Soil Research 18, 309-323.

McGhie DA, Posner AM (1981) The effect of plant top material on the water repellence of fired sands and water repellent soils. Austrulian Journal of Agricultural Research 32, 609-620. doi: 10.1071/AR9810609

Noller CR (1966) 'Textbook of organic chemistry.' (Saunders: Philadelphia, London)

Northcote KH, Hubble GD, Isbell RF, Thompson CH, Benenay E (1975) 'A description of Australian soils.' (CSIRO: Australia)

Roberts FJ, Carbon BA (1972) Water repellence in sandy soils of south-western Australia. II Some chemical characteristics of the hydrophobic skins. Australian Journal of Soil Research 10. 354-2.

Roper MM (1998) Sorting out sandy soils. Microbiology Australia 19, 6-7.

Roper MM (2004) The isolation and characterisation of bacteria with the potential to degrade waxes that cause water repellency in sandy soils. Australian Journal of Soil Research 42, 427-434. doi: 10.1071/SR03153

Tale ME, Oades JM, Ma'shum M (1989) Non-wetting soils, natural and induced: overview and future developments. In 'The Theory and Practice of Soil Management for Sustainable Agriculture. A Workshop of the Wheat Research Council'. pp. 70-77. (AGPS: Canberra, ACT)

Wallis MG, Home DJ (1992) Soil water repellency. Advances in Soil Science 20, 91-146.

Manuscript received 30 July 2004, accepted 3 December 2004

Margaret M. Roper (A,C) and V. V. S. R. Gupta (B)

(A) CSIRO Plant Industry, Private Bag No. 5, Wembley, WA 6913, Australia.

(B) CSIRO Land and Water, Private Bag No. 2, Glen Osmond, SA 5064, Australia.

(C) Corresponding author. Email: Margaret.Roper@csiro.au
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Author:Roper, Margaret M.; Gupta, V.V.S.R.
Publication:Australian Journal of Soil Research
Geographic Code:8AUST
Date:Mar 1, 2005
Words:5072
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