Bacterial populations were isolated from the soil-root interface and root-free regions of Agropyron ... ical, and genetic heterogeneity within a community (1) and.
APPLIED AND ENVIRONMENTAL MICROBIOLOGY, OCt. 1986, p. 765-770
Vol. 52, No. 4
0099-2240/86/100765-06$02.00/0 Copyright © 1986, American Society for Microbiology
Bacterial Physiological Diversity in the Rhizosphere of Range Plants in Response to Retorted Shale Stress W. C.
Departments of Microbiologyl
METZGER,'* D.
A. KLEIN,' AND E. F. REDENTE2 and Range Science,2 Colorado State University, Fort Collins, Colorado 80523 Received 21 February 1986/Accepted 24 June 1986
Bacterial populations were isolated from the soil-root interface and root-free regions of Agropyron smithii Rydb. and Atriplex canescens (Pursh) Nutt. grown in soil, retorted shale, or soil over shale. Bacteria isolated from retorted shale exhibited a wider range of tolerance to alkalinity and salinity and decreased growth on amino acid substrates compared with bacteria from soil and soil-over-shale environments. Exoenzyme production was only slightly affected by growth medium treatment. Viable bacterial populations were higher in the rhizosphere and rhizoplane of plants grown in retorted shale than in plants grown in soil or soil over shale. In addition, a greater number of physiological groups of rhizosphere bacteria was observed in retorted shale compared with soil alone. Two patterns of community similarity were observed in comparisons of bacteria from soil over shale with those from soil and retorted-shale environments. Root-associated populations from soil over shale had a higher proportion of physiological groups in common with those from the soil control than with those from the retorted-shale treatment. However, in non-rhizosphere populations, bacterial groups from soil over shale more closely resembled the physiological groups from retorted shale.
Diversity measurements have recently received increased emphasis in microbial ecological studies, especially as a tool to assist in evaluation of microbial community functional responses to environmental stress (2). Diversity measurements can provide information about taxonomic, physiological, and genetic heterogeneity within a community (1) and also allow community responses to be evaluated on a series of levels. In this study, the influence of retorted shale on the physiological diversity of bacterial isolates from rhizosphere (RS), rhizoplane (RP), and non-rhizosphere (non-RS) bacterial populations associated with selected reclamation plant species was investigated. The physiological unit characters were chosen to provide information related to bacterial functioning and community structure at the soil-root interface which might be responsive to the presence of retorted shale. Retorted shale, produced as a byproduct of pyrolytic decomposition of organic material in oil shale, is characterized as being extremely alkaline and highly saline (17), and in addition this material can contain hazardous trace elements (J. P. Fox, Ph.D. thesis, University of California, Berkeley, 1980). Plant growth can occur on retorted shale wastes or on soil placed over retorted shale; however, the presence of this material has been associated with negative effects on belowground process development (10). Sorensen et al. (26) found reduced enzyme activity in microbial communities collected from revegetated surface soils that directly overlay retorted shale. Decreased microbial activity and populations have also been observed in soil microcosms amended with retorted shale (9). In addition, lower bacterial taxonomic diversity has been associated with the initial colonization of surface horizons of retorted shale in field lysimeter studies compared with adjacent soil (16). However, little information is available on the responses of root-associated microorganisms to the presence of retorted shale. The objectives of this study were twofold: (i) to assess some root-associated, in addition to non-RS, bacterial re*
sponses to retorted shale and (ii) to evaluate physiological diversity responses of bacteria in a plant-soil system exposed to retorted-shale stress. MATERIALS AND METHODS Plant growth materials. Surface soil material (0- to 15-cm depth) was obtained from an undisturbed site in a big sagebrush (Artemisia tridentata) plant community in the Piceance Basin of northwest Colorado (15). Native soils of this region have been mapped as a soil complex that consisted of a Rentsac channery loam (loamy-skeletal, mixed, frigid Lithic Ustic Torriorthents), a Piceance fine sandy loam (fine-loamy, mixed, Borollic Camborthids), and a Yamac loam (fine-loamy, mixed Borollic Cambrothids) (27). Soil was passed through a 6-mm-mesh screen and stored in plastic bags at 6°C until ready for use in the greenhouse pot experiment. Retorted oil shale was obtained from the Laramie Energy Technology Center, Laramie, Wyo. The oil shale was mined at Anvil Points, Colo., and retorted via a simulated vertical modified in situ batch process. The retorted shale was leached with approximately 3 pore volumes of water in an attempt to reduce its initially high electrical conductivity after it was received from the Energy Technology Center. Rosana western wheatgrass (Agropyron smithii), and fourwing saltbush (Atriplex canescens) were each grown in a soil control, retorted shale, or 8.5 cm of soil over 8.5 cm of retorted shale plant growth medium treatment. In addition, each plant growth medium received a 2.5-cm surface layer of a vermiculite-perlite-composted pine bark mixture which served as a germination bed. A split-block experimental design with four blocks was used to contain all possible combinations of plant species and plant growth medium treatments. Soil, retorted shale, and surface layer germination materials were added to 20-cm-deep plastic pots which had been previously surface sterilized in a solution of Wescodyne. The seeds were surface sterilized for 3 min in a 10% (vol/vol) sodium hypochlorite solution before they were planted. All treatments were fertilized with the equivalent of 110 kg of
Corresponding author. 765
766
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METZGER ET AL.
TABLE 1. Chemical characteristics of the soil control and leached, retorted shale before plant growth Characteristica
Sample
pHb
EC (mmho
cm-l)c
Soil control 7.4 Leached shale 12.6
0.4 15.4
Cation content (meg literl)c:
SARC
Anion content (meg literl)c:
HCO3 CO3 OH Cl S04 0 0.3 NDe 0.38 6.8 1.0 0.6 0.1 5.6 0 6.6 54.9 1.1 5.0 66.8 0.9 0.1 47.0 13.2 0 Ca
Mg
Na
K
Nitrate N
P
K
% Organic
(qg g-l)d (>Lg g-l)d (pLg g-l)d
8 2
2 19
45 442
matter
1.1 0.2
Abbreviations: EC, electrical conductivity; SAR, sodium adsorption ratio. b Paste. c Saturation extract. d NH4HCO3-diethyltriaminepentaacetic acid extract. ' ND, Not determined. a
nitrogen ha-' and 90 kg of phosphorus ha-1. One-half of the nitrogen, as ammonium nitrate, was applied 1 week after emergence; the balance was applied 1.5 months later. Phosphorus, as triple super phosphate, was mixed into the soil and shale materials prior to seeding. The western wheatgrass and fourwing saltbush stands were thinned to 10 and 4 plants, respectively, per pot after emergence. Pots were watered from above to near field capacity and maintained under natural light in a greenhouse. Plants were harvested for microbial population analyses after a growth period of 3 months. Enumeration of viable aerobic heterotrophic bacteria. One plant of each species was randomly selected per block from each of the three plant growth medium treatments. The surface germination layer was removed and discarded, and the root systems were gently extracted from the soil or shale. Material that adhered to the roots was designated as the RS fraction, whereas soil collected from the root-free regions of the pot was designated non-RS soil. The RS fraction was separated into RS soil and the root surface or RP. Bacterial isolates were obtained from RS soil and the RP with a modification (18) of the technique described by Louw and Webley (11). Enumerations of viable RS, RP, and non-RS bacteria were estimated by dilution and spread plate techniques (31) on material obtained from soil, retorted shale, and the soil layer from the soil-over-shale treatment. Dilutions of plant growth media were plated in triplicate on sodium caseinate agar at pH 6.8 (24). Plates were incubated at 25°C in the dark and counted after a 2-week incubation. Characterization of bacterial isolates. Aerobic heterotrophic bacteria were classified by a numerical taxonomic approach (23). A total of 383 well-isolated RS, RP, and non-RS bacterial colonies were randomly obtained from the original viable enumeration samples. Bacterial colonies were maintained on replicate sodium caseinate agar plates. Bacterial isolates were characterized for growth at pH 10 on sodium caseinate medium; growth on sodium caseinate, pH 7, plus addition of 1.0, 2.5, 5.0, 7.5, or 10.0% (wt/vol) sodium chloride; exoenzyme production in petri dishes (amylase [31], proteolysis [31], Tween 80 esterase [19], gelatinase [6], chitinase [21], cellulase in a 1.0% cellulose-azure-modified Pettersson medium [22] without an additional carbon source); and growth on 0.1% glycine and methionine as sole carbon sources (3). Isolate responses were coded 1 for a positive and 0 for a negative response to these tests. Bacterial isolates were clustered with CLUSTAN (30), a cluster analysis computer package. A simple matching coefficient was used as the similarity coefficient. Clusters were formed by means of average linkage, which is equivalent to the unweighted pair-group method of Sokal and Michener (25). A cluster
generated from CLUSTAN in this study was synonymous with a bacterial cluster, i.e., a group of bacterial isolates functionally related to each other at some preselected level of similarity. Bacterial diversity analysis. Bacterial diversity was calculated by the rarefaction method. Isolates were pooled from samples collected from western wheatgrass and fourwing saltbush. Simberloff (20) developed a computer program that calculated the expected number of groups present in a subsample of individuals drawn from the total population. The raw data, i.e., the total number of bacterial clusters and the number of isolates per cluster, needed for the rarefaction program were obtained from the cluster analysis program. Bacterial isolates were clustered at an 85% level of similarity. This level was selected by following procedures similar to those given by Mills and Wassel (12). Bacterial community similarity. Community similarity was determined from the pooled bacterial sample with the Sorensen index (13). Pairwise comparisons of all possible plant growth medium treatment combinations were made in the RS, RP, and non-RS bacterial communities. The binary unit character data from each treatment of the pairwise comparison were reclustered by the above-noted procedures. Values of the index ranged from zero (no common clusters) to one (all clusters in common). Statistical analyses. Log-transformed bacterial enumeration data were analyzed by analysis-of-variance techniques. Significant effects (i.e., F statistic at P < 0.05) of the plant growth medium were further evaluated by the leastsignificant-difference method with a 95% confidence interval. Bacterial physiological data were evaluated for effects of plant growth medium on unit character occurrence by the Kruskal-Wallis one-way analysis-of-variance test. RESULTS Characterization of plant growth media. Results from selected chemical analyses of the plant growth media used TABLE 2. Viable bacterial populations in the plant root environment of range plants grown in soil, retorted shale, or soil over shale
Mean bacterial populationa
Growth medium treatment
RS
RP
Non-RS
Soil control Retorted shale Soil over shale
8.2 (b) 9.4 (a) 8.5 (b)
8.6 (b) 9.6 (a) 8.8 (b)
6.3 (a) 6.6 (a) 6.6 (a)
g-1
a RS and non-RS, log CFU of dry soil; RP, log CFU g-1 of dry root. Means with different letters within a bacterial population are significantly
different (P < 0.05).
VOL. 52, 1986
RETORTED SHALE AND RHIZOSPHERE BACTERIAL DIVERSITY
767
A 100 w 80 z W
60
0 0
40
2 20
O
Glycine Proteolysis Esterase UNIT CHARACTER
100 -
B
a
a
W 80 C.) z
ob
w
cr 60 -
C.)
a
o 40
-
a2 20
-
bb
b
0
a iu ; b ;. ag 0~~~~~~~~~~o
b. b
0
M
---
I t
pH 10
*X
L.
A
-~~~~~~..
2.5%NaCI Methionine Glycine Proteolysis Esterase
0
a
Glaotinase Amylose Chitinose
UNIT CHARACTER
C 100
-
0
80
z B w
a
z 60
-
o 40
-
a ob
b b
0
b' 0
0
at 20
b
TE
0
pH 10
a
0
al I.
2.5%NaCI Methionine Glycine
0
a a a
Proteolysis Esterose Gelotinose Amylose Chitinase CHARACTER
UNIT
FIG. 1. Percent occurrence of selected unit characters used to assess bacterial diversity as affected by plant growth media: (A) RS, (B) RP, (C) non-RS. Means with different letters within a unit character were significantly different (P < 0.05). are in Table 1. The soil control was nonsaline and nonsodic with a near-neutral pH. The predominant cations extracted from the soil solution were the basic cations of calcium and magnesium, reflected in the relatively low sodium adsorption ratio. High alkalinity and sodicity characterized the leached retorted shale, with sodium and potassium as the predominant cations in solution. The retorted-shale plant growth medium has a low organic-matter content. Bacterial enumeration. Distinct differences in microbial population responses to the presence of retorted shale were
evident (Table 2). RS and RP viable bacterial populations significantly elevated in the retorted-shale plant growth medium treatment compared with the soil control and soilover-shale treatments. Retorted shale had no effect on the numbers of non-RS bacteria. Similar bacterial populations were found in the soil and soil-over-shale treatments in the RS, RP, and non-RS environments. Characterization of bacterial clusters. The percent occurrence of unit characters in the different bacterial clusters showed distinct effects of the plant growth media used (Fig.
were
768
APPL. ENVIRON. MICROBIOL.
METZGER ET AL.
TABLE 3. Percent growth of bacterial isolates in the plant root environment of plants grown in soil, retorted shale, and soil over retorted shale at various NaCl concentrations' % Growth at an NaCI concn (wt/vol) of:
Growth medium treatment
1%
2.5%
5%
7.5%
10%
Soil control Retorted shale Soil over shale
78 97 84
49 91 48
11 27 14
0 8 2
0 1 0
aSodium caseinate, pH 7.
1). Amino acid utilization was lower and tolerance to salinity and alkalinity was higher in isolates obtained from the retorted-shale growth medium than in those from the soil control. Exoenzyme activity of selected hydrolases in bacteria from retorted shale was variable. Significantly more root-associated (RS and RP) bacteria from the retorted-shale treatment than from the soil control exhibited proteolytic activity. A higher number of RP isolates from the soil control possessed gelatinase activity compared with RP isolates from the retorted-shale treatment. The percent occurrence of the other tested hydrolases in all three bacterial populations were similar in the soil control and retorted-shale treatments. Generally, hydrolase activities of bacteria in the soil-over-shale treatment were not significantly different from those of bacteria from the soil control. Chitinase activity was found in only a few RP clusters isolated from the soil control and soil-over-shale treatments and in one nonRS cluster isolated from the soil-over-shale treatment. No cellulase activity was observed. The tolerance of isolates to various levels of NaCl addition to the bacterial medium was also affected by the plant growth medium (Table 3). Consistently more salt-tolerant RS, RP, and non-RS bacteria were isolated from retorted shale for each NaCl addition level compared with the soil and soil-over-shale treatments. The most salt-tolerant bacteria were also found in retorted shale, where some isolates were capable of growth with 10% added NaCl. The maximum tolerances of isolates from soil and soil over shale were 5.0 and 7.5%, respectively. Bacterial diversity. The raw data obtained from the cluster analysis indicated that most bacterial isolates were distributed in a few major clusters (Table 4). The rarefaction curves TABLE 4. Distribution of isolates among bacterial clusters from range plants grown in soil, retorted shale, or soil over shale Population source
Distribution of isolates among clustersa
RS Soil ......... Retorted shale ......... Soil over shale .........
13, 9, 7, 3, 2 (3), 1 16, 12, 5, 3, 2 (3), 1 (4) 12, 10, 5, 4 (2), 2 (4), 1 (2)
RP Soil ......... Retorted shale ......... Soil over shale .........
19, 11, 5, 2, 1 (4) 16, 12, 4 (3), 3 (2), 1 (2) 28, 4 (2), 3, 2 (3), 1 (2)
Non-RS Soil ......... Retorted shale ......... Soil over shale .........
9, 6, 5 (2), 3, 2, 1 (3) 15, 10, 7, 4 (2), 2 (2), 1 (2) 9, 6 (2), 3 (2), 2 (3), 1 (5)
showed that the most distinct effect of retorted shale on bacterial diversity was found in RS populations (Fig. 2A). The presence of retorted shale, alone or under soil, resulted in higher diversity in these two treatments compared with RS bacteria from plants grown in soil. In comparison, the plant growth medium had no effect on physiological diversity of bacterial isolates from the RP (Fig. 2B). Diversity in non-RS bacterial populations was highest in the soil-overshale treatment compared with retorted shale and the soil control (Fig. 2C); diversity in the latter two treatments was similar. Bacterial community similarity. The values obtained for the Sorensen index among comparisons in the RS and
12.0 -
]A.
8.0-
-
,'
Retorted shale -
Soil control I-
4.0 'I
a: 0.0
LLJ C/) :D
F-
0 H cr 0 11J z-
m
LuJ D z 15.0-
C. Soil over shale
12.0
A....
Soil control _-
9.06.0-
-V Retorted shale
,.'~'
-
',
3.0'-.f If _
a Numbers outside parentheses represent the number of isolates in the bacterial cluster. Numbers in parentheses represent the number of bacterial clusters of that size.
.>Soil over shale-----
:_:~
. _< /
_
0.0
.
o0.0
. I.
20.0 30.0 40.0
.
-
.
56.0 66.0
NUMBER OF ISOLATES FIG. 2. Rarefaction curves of physiological diversity in bacterial populations as affected by plant growth media: (A) RS, (B) RP, (C) non-RS. Each curve is bracketed by a band 1.96 standard deviations wide which constitutes approximately 95% confidence limits.
VOL. 52, 1986
RETORTED SHALE AND RHIZOSPHERE BACTERIAL DIVERSITY
769
TABLE 5. Measurements of bacterial community similarity obtained from pairwise comparisons of plant growth media from RS, RP, and non-RS communities No. of unique clusters (isolates)
Growth medium comparison A
B
A
B
No. of common clusters
RS Shale Shale Soil over shale
Soil over shale Soil Soil
5 (8) 5 (22) 4 (8)
6 (13) 4 (11) 3 (9)
7 5 7
0.56 0.53 0.67
Shale Shale Soil over shale
Soil over shale Soil Soil
6 (19) 8 (32) 4 (7)
8 (33) 6 (28) 3 (4)
3 2 7
0.30 0.22 0.67
Non-RS Shale Shale Soil over shale
Soil over shale Soil Soil
3 (13) 4 (9) 6 (8)
5 (8) 4 (9) 2 (4)
7 6 6
0.64 0.60 0.60
Sorensen index
RP
non-RS were similar (Table 5). The number of common clusters were equal to or greater than the number of unique clusters in all RS and non-RS comparisons. Values of the Sorensen index did vary considerably among comparisons in the RP. The number of unique clusters was greater than the number of common clusters when RP bacterial clusters from soil or soil over shale were compared with the retorted-shale plant growth medium. However, when RP bacterial clusters from the soil were compared with clusters from soil over shale, a cluster distribution similar to that found in the RS and non-RS was found, which resulted in a high index value. In addition, two patterns of similarity were observed among the different bacterial communities. Soil over shale and soil was the most similar pair in RS and RP communities, whereas in non-RS bacterial populations, the most similar pair was soil over shale and shale. DISCUSSION Plants and their associated soil microbial communities respond to environmental stress in many subtle and interactive ways. Environmental stress, whether natural or man made, can alter the structure of a microbial community and consequently has been shown to affect microbial diversity (1, 2, 28, 29). Results from this study showed that an applied stress (retorted shale) had a differential effect on the parameters of microbial physiological diversity that were measured. Physiological functioning of the bacterial isolates also suggested differential effects of the plant growth media. Whereas many of the bacterial populations from retorted shale were tolerant of pH 10 with added NaCl, bacterial populations from the soil and soil-over-shale treatments were restricted to a narrower range of alkalinity and salinity. In addition, bacteria from the retorted-shale plant growth medium had a decreased amino acid utilization potential compared with isolates from soil. In contrast, the presence of retorted shale had only a slight effect on hydrolytic exoenzyme activity. Amino acid content in plant root exudates has been shown to be suppressed in plants exposed to salt-induced osmotic stress (14). Hence, the retorted-shale environment may favor organisms that do not utilize amino acids. It has also been shown that hydrolases were less inhibited than- other enzyme classes by soil salinity (5). Results from this study showed that retorted shale had a differential effect on the number of bacterial clusters associ-
ated with each bacterial population. Because of the inconsistent effect of retorted shale (soil over shale versus retorted shale) in the non-RS, it was difficult to identify precisely the factor(s) that contributed to the observed responses. However, when comparisons were made between the soil control and retorted-shale treatments, the only difference in bacterial cluster number was found in the RS. This may be a result of environmentally induced changes in root exudation patterns. Generally, the amount of materials released from roots increases with plant stress (4, 7, 8). A more diverse population could be supported if more microbial substrates were released by plant roots grown in retorted shale. Based on the results of this study, it appeared that non-RS, and possibly also RP, populations were less influenced than RS populations by possible retorted-shale-related changes in root exudation patterns. As discussed by Curl and Truelove (4), RP bacteria are often closely associated with polysaccharide-containing mucigels, which may be utilized directly by these bacteria. Mucigel itself would be a more consistent carbon and energy source for microbial functioning. In addition, as noted by these researchers, the mucigel represents an environment of high root exudate accumulation. Based on these concepts, RP microorganism might be less influenced by variations in exudate composition and release rates. It should be emphasized that treatments with which similar diversities occurred were not necessarily composed of the same bacterial groups. It might be expected that distinct microbial groups would develop in retorted shale and in soil as a result of inherent differences in the chemical characteristics of these two plant growth media. In contrast, a high degree of bacterial similarity might be expected between pairwise comparisons of soil over shale with soil. Initially, the soil layer from the soil-over-shale treatment was the same material as the soil control. These two plant growth media would share a common pool of organisms. This was the case for RS and RP populations, in which the highest Sorensen index value was associated with the comparison of soil over shale with soil. However, among non-RS comparisons, bacterial populations from soil over shale were more similar to populations from retorted shale than to the soil control. This suggested that bacterial populations in non-RS soils may be more susceptible to the effects of retorted shale in the plant growth medium compared with RS and RP populations. Direct contact with retorted shale may not be a
770
METZGER ET AL.
necessary prerequisite to alter the functional status in the non-RS. Bacterial diversity in soil microbial communities in response to environmentally induced stress is influenced by two interactive factors. The physiochemical properties of the environment have a more direct effect on the functioning and community structure of non-RS populations than on populations in the root zone region. The response of bacteria at the soil-root interface to stress also depends on the physiological response of the plant to that stress. ACKNOWLEDGMENTS This work was supported by the U.S. Department of Energy under contract no. DE-AS02-76EV04018. We thank Kathleen Smoke and Joseph Mankowski for their excellent technical assistance. LITERATURE CITED 1. Atlas, R. M. 1984. Diversity of microbial communities, p. 1-47. In K. C. Marshall (ed.), Advances in microbial ecology, vol. 7. Plenum Publishing Corp., New York. 2. Altas, R. M. 1984. Use of microbial diversity measurements to assess environmental stress, p. 540-545. In M. J. Klug and C. A. Reddy (ed.), Current perspectives in microbial ecology. American Society for Microbiology, Washington, D.C. 3. Colwell, R. R., and W. J. Wiebe. 1970. Core characteristics for use in classifying aerobic, heterotrophic bacteria by numerical taxonomy. Bull. Georgia Acad. Sci. 28:165-185. 4. Curl, E. A., and B. Truelove. 1986. The rhizosphere. Adv. Ser. Agric. Sci. 15:9-54. 5. Frankenberger, W. T., Jr., and F. T. Bingham. 1982. Influence of salinity on soil enzyme activities. Soil Sci. Soc. Am. J. 46:1173-1177. 6. Frazier, W. C. 1926. A method for detection of changes in gelatin due to bacteria. J. Infect. Dis. 39:302-309. 7. Hale, M. G., and L. D. Moore. 1979. Factors affecting root exudation II: 1970-1978. Adv. Agron. 31:93-124. 8. Hale, M. G., L. D. Moore, and G. J. Griffin. 1978. Root exudates and exudation, p. 163-204. In Y. R. Dommergues and S. V. Krupa (ed.), Interactions between non-pathogenic soil microorganisms and plants. Elsevier Publishing Co., Amsterdam. 9. Hersman, L. E., and D. A. Klein. 1979. Retorted oil shale effects on soil microbiological characteristics. J. Environ. Qual. 8:520-524. 10. Klein, D. A., D. L. Sorensen, and E. F. Redente. 1985. Soil enzymes: a predictor of reclamation potential and progress, p. 141-171. In R. L. Tate III and D. A. Klein (ed.), Soil reclamation processes: microbial analyses and applications. Marcel Dekker, Inc., New York. 11. Louw, H. A., and D. M. Webley. 1959. The bacteriology of the root region of the oat plant grown under controlled pot culture conditions. J. Appl. Bacteriol. 22:216-226. 12. Mills, A. L., and R. A. Wassel. 1980. Aspects of diversity measurement for microbial communities. Appl. Environ. Microbiol. 40:578-586. 13. Pielou, E. C. 1977. Mathematical ecology. John Wiley & Sons, Inc., New York.
APPL. ENVIRON. MICROBIOL. 14. Polenenko, D. R., E. B. Dumbroff, and C. I. Mayfield. 1983. Microbial responses to salt-induced osmotic stress. III. Effects of stress on metabolites in the roots, shoots and rhizosphere of barley. Plant Soil 73:211-225. 15. Redente, E. F., C. B. Mount, and W. J. Ruzzo. 1982. Vegetation composition and production as affected by soil thickness over retorted oil shale. Reclam. & Reveg. Res. 1:109-122. 16. Rogers, J. E., V. M. McNair, S. W. Li, T. R. Garland, and R. E. Wildung. 1982. Microbial colonization of retorted shale in field and laboratory studies. PNL-4311. Pacific Northwest Laboratory, Richland, Wash. 17. Schmehl, W. R., and B. D. McCaslin. 1973. Some properties of spent oil shale significant to plant growth, p. 27-43. In R. J. Hutnik and G. Davis (ed.), Ecology and reclamation of devastated land, vol. 1. Gordon and Breach, New York. 18. Sherwood, J. E., and D. A. Klein. 1981. Antibiotic-resistant Arthrobacter sp. and Pseudomonas sp. responses in the rhizosphere of blue grama after herbage removal. Plant Soil 69:91-96. 19. Sierra, G. 1957. A simple method for the detection of lipolytic activity of microorganisms and some observations on the influence of the contact between cells and fatty substrates. Antonie van Leeuwenhoek J. Microbiol. Serol. 23:15-23. 20. Simberloff, D. 1978. Use of rarefaction and related methods in ecology, p. 150-165. In K. L. Dickson, J. Cairns, Jr., and R. J. Livingston (ed.), Biological data in water pollution assessment: quantitative and statistical analyses. ASTM STP652. American Society for Testing and Materials, Philadelphia. 21. Skujins, J. J., H. J. Potgieter, and M. Alexander. 1965. Dissolution of fungal cell walls by a streptomycete chitinase and ,B-(1 3) glucanase. Arch. Biochem. Biophys. 111:358-364. 22. Smith, R. E. 1977. Rapid tube test for detecting fungal cellulase production. Appl. Environ. Microbiol. 33:980-981. 23. Sneath, P. H. A. 1957. Application of computers to bacterial taxonomy. J. Gen. Microbiol. 17:210-226. 24. Society of American Bacteriologists. 1957. Manual of microbiological methods, p. 43. McGraw-Hill Book Co., New York. 25. Sokal, R. R., and C. D. Michener. 1958. A statistical method for evaluating systematic relationships. Univ. Kans. Sci. Bull. 38:1409-1438. 26. Sorensen, D. L., D. A. Klein, W. J. Ruzzo, and L. E. Hersman. 1981. Enzyme activities in revegetated surface soil overlying spent Paraho process oil shale. J. Environ. Qual. 10:369-371. 27. Stark, J. M., and E. F. Redente. 1985. Soil-plant diversity relationships on a disturbed site in northwestern Colorado. Soil Sci. Soc. Am. J. 49:1028-1034. 28. Tate, R. L., III, and A. L. Mills. 1983. Cropping and the diversity and function of bacteria in Pahokee muck. Soil Biol. Biochem. 15:175-179. 29. Wassel, R. A., and A. L. Mills. 1983. Changes in water and sediment bacterial community structure in a lake receiving acid mine drainage. Microb. Ecol. 9:155-169. 30. Wishart, D. 1978. CLUSTAN user manual, 3rd ed. Program Library Unit, Edinburgh University, Edinburgh. 31. Wollum, A. G., II. 1982. Cultural methods for soil microorganisms, p.781-802. In A. L. Page, R. H. Miller, and D. R. Keeney (ed.), Methods of soil analysis, part 2: chemical and microbiological properties, 2nd ed. American Society of Agronomy, Inc., Madison. Wis.
