Virginia Water Resources Research Center

Rapid Assays for Microbial Degradation

of 2-Chlorophenol

I. S. Pal ·.Murphy \. C. Carter .. W. Drew

Bulletin 128

Bulletin 128 November 1980

Rapid Assays for

Microbial Degradation

of 2-Chlorophenol

H. S. Pal T. Murphy

A. C. Carter S. W. Drew

Department of Chemical Engineering Virginia Polytechnic Institute and State University

Blacksburg, Virginia 24061

The work upon which this report is based was supported in part by funds provided by the United States Department of the Interior,

Office of Water Research and Technology, as authorized by the Water Research and Development Act of 1978 (P.L. 95-467).

Project B-088-V A VPl-VWRRC-BULL 128

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A publication of Virginia Water Resources Research Center

Virginia Polytechnic Institute and State University Blacksburg, Virginia 24060

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Contents of this publication do not necessarily reflect the views and policies of the United States Department of the Interior,

Office of Water Research and Technology, nor does mention of trade names or commercial products constitute their endorsement

or recommendation for use by the United States government.

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Additional copies of this publication, while the supply lasts, may be obtained from

the Virginia Water Resources Research Center. Single copies are provided free to persons

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TABLE OF CONTENTS

List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . iv

List of Tables . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . iv

Acknowl~dgments . ....................... . ................ v

Abstract ................................................ 1

Introduction ............................................. 3

Literature Review . ........................................ 4

Materials and Methods . ..................................... 6 I. Growth and Isolation of Microorqanisms ................... 6

A. Isolates ......................................... 6 B. Mixed Culture .................................... 6 C. Continuous Culture ................................ 6 D. Experimental I nocula .............................. 7

II. Media ............................................. 7 A. Medium A ....................................... 7 B. Medium 8 ....................................... 7

II I. Determination of 2-CP and Phenol ....................... 8 A. Gas Chromatography ............................... 8 B. Distillation ...................................... 8

IV. Determination of Glucose and Cellobiose .................. 8 V. Determination of Oxygen Consumption Rate ............... 9

A. Isolates ......................................... 9 B. Batch ........................................... 9

VI. Determination of Chloride Ion .......................... 9 VI I. Pure-Culture Microbial Degradation of 2-CP ................ 9

VII I. Preparation of Isolates RS 10 and RS 17 for Testing the Differential Plate Count Technique ...................... 1 O

IX. Preparation of Undefined Mixed Batch Cultures for 2-CP Shock Studies ...................................... 10

Results and Discussion .................................... 12 I. Microbial Degradation of 2-CP by Pure Cultures ............ 12

11. Microbial Degradation of 2-CP by Mixed Populations ........ 12 111. Changes in Microbial Populations Exposed to 2-CP .......... 13

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IV. System Response to Shock Load Doses of 2-CP ............ 17

Conclusions . ............................................ 19

Literature Cited . ................... . ..................... 21

Figure . ................................................ 23

Tables .. ............................................... 25

FIGURE

1. Effect of Pulse Addition of 2-CP at Two Levels to Undefined Mixed Cultures ............................... 24

TABLES

1. Growth and Chlorophenol Metabolism of Microbial Isolates in the Presence of 20 mg/I 2-CP ........................... '26

2. Influence of 2-CP on the Oxygen Respiration Rate of Acclimated and Unacclimated Undefined Mixed Cultures ....... 27

3. Cometabolism of 2-CP by an Acclimated Mixed Culture ........ 28

4. Enumeration of 2-CP Degrading Microorganisms in Defined Mixtures ...................................... 30

5. Number of Cells Able to Grow (Resistant) and Able to Grow and Degrade 2-CP in Acclimated and Unacclimated Cultures .... 31

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ACKNOWLEDGMENTS

The authors would like to acknowledge the following: the Koppers Co., Inc. and the American Iron and Steel Institute for their helpful sugges­tions during the early part of this study; the Department of Chemical Engineering at Virginia Polytechnic Institute and State University for partial financial support of H.S. Pal; T.W. Johnson of the Virginia Water Resources Research Center for his assistance with the administration of this project; and finally, the Office of Water Research and Technology for their support of this study.

Special acknowledgment is made to the following who generously gave their time to a critical review of the manuscript: T. Al Austin, Iowa State Water Resources Research Institute; R. R. Colwell and Roy D. Sjoblad, Department of Microbiology, University of Maryland; and Makram T. Suidan, Department of Civil Engineering, Georgia Institute of Techno­logy. Acknowledgment is made also to Sandra P. Milliken for editorial processing and typesetting and to Gretchen S. Bingman for layout com­position.

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ABSTRACT

Until recently it has been impossible to characterize the microbial sub­populations of complex ecosystems on the basis of their metabolic char­acteristics. This study presents two simple procedures for estimating the size of microbial subpopulations and their ability to degrade 2-chloro­phenol. The assay techniques were used to study shifts in microbial sub­populations identified as sensitive or resistant to the toxic effects of 2-chlorophenol. The studies also show the effect of cosubstrate metabolism on degradation of 2-chlorophenol by pure cultures.

Adaptation or acclimation to a normally toxic environment results in a shift in microbial subpopulations. Once the subpopulations were estab­lished in acclimated cultures, their response to the pulsed addition of 2-chlorophenol was much less than was the response of a paired unaccli­mated culture. The screening procedure indicated that increased resis­tance to 2-chlorophenol paralleled the increased 2-chlorophenol degra­ding capability of the microbial subpopulations. The assays make possible rapid and accurate estimation of 2-chlorophenol resistance and degrada­tion in microbial populations.

Key Words: Indicator Organisms, Chlorophenol, Most-Probable-Number, Culture Acclimation, Microbial Subpopulations.

INTRODUCTION

Chlorinated monophenolic compounds in water present serious health hazards, not only because of their inherent toxicity but also because they react with other substances to form compounds of still greater toxicity. The presence of chlorinated phenolic materials in surface waters can re­sult from chlorinated phenolic pesticides and biocides on agricultural lands, industrial spills, and inadequate secondary treatment of industrial wastes. Increased public concern over water pollution by these impor­tant compounds has led to their control by the United States Environ­mental Protection Agency. The 1977 standard for best practicable con­trol technology currently available (BPCTCA 1977) is set at 0.1 µg of phenol per gram of discharge, with limitation in 1983 by best available control technology economically achievable (BACTEA 1983) expected to be 0.02 µg of phenolic compound per gram of discharge (EPA, 410/ 174014a, 1977). Currently, acclimated and stabilized biological systems can reduce phenol concentrations to approximately 0.1 µg of phenol per gram of discharge.

Clearly, the maintenance of water quality will require a more complete understanding than we presently have of biological processes involved in the removal of chlorinated phenolic materials from wastewaters. Fur­ther, more precise measures of the sensitivity of chlorophenol must be de­veloped if proposed regulations are to be met. This work focused on the development of a rapid method for determining the number of micro­organisms in wastewaters capable of degrading chlorinated phenolic compounds.

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LITE RA TUR E REVIEW

Microbial metabolism of halogenated aromatic compounds has been ex­tensively studied in recent years. Microbial metabolism of halogen-sub­stituted benzoic acid [Johnson et al., 1972; Milne et al., 1968; Horvath and Flathmen, 1976], mono- and di-substituted chlorophenoxyacetic acids [Kirkland, 1967; Fryer and Kirkland, 1970; Hurle and Radlemacher, 1970], and chlorine-substituted phenols [Watanabe, 1977; Broecker and Zahn, 1977] have been reported. In each case, the toxicity of the halo­genated aromatic compound to the microbial system was evident, al ­though the degree of toxic response and extent of degradation seemed to depend heavily on microbial system characteristics. It is clear that in spite of their toxic nature, chlorinated phenolic compounds are microbially degraded in nature. It seems, however, that these compounds seldom serve as primary carbon sources but rather are usually cometabolized with more easily metabolized carbon sources [Horvath, 1972; Broecker and Zahn, 1977].

Microbial populations in natural environments are highly hetrogeneous. Simple systems such as fast-moving streams typically have low population densities and rather broad and fluctuating distribution of microbial spe­cies. More complex systems such as activated sludge facilities may have very high population densities and fairly stable distribution of microbial types. It is probable that the toxicity to chlorinated phenols is different between free moving streams and activated sludge systems. This idea is supported by Broecker and Zahn [ 1977] who studied response of simple receiving water and activated sludge systems to pulsed or stepped addition of 3,5-dichlorophenol. Their work showed that the response of the two systems was inherently similar at very low doses. However, as the dose of 3,5-dichlorophenol increased, toxic response of the receiving waters was more severe .than that of activated sludge. Although these authors did not quantify the viable microbial population or subpopulations, the size and composition of microbial populations are presumably important parameters in toxic pressure.

Broecker and Zahn [ 1977] also noted that the magnitude of toxic re­sponse and rate of recovery from pulsed shock loading was very much a function of the recent history of adaptation to the chlorinated phenol. Systems maintained in the absence of 3,5-dichlorophenol for more than three days were much more sensitive to subsequent shock loading of the chlorophenol than was the control system. Since their test system was a

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continuous reactor with high cell retention characteristics, it is tempting to speculate that acclimation to a particular toxic substrate or loss of acclimation involved a change in the distribut ion of microbial subpopu­lations or at least a shift in the phenotypic expression of a subpopulation's metabolic potential. Alteration of a microbial population in response to the introduction of recalcitrant hydrocarbons has been demonstrated [Walker and Colwell, 1973; Hood et al., 1975].

This study examines the problem of evaluating mixed microbial systems which degrade 2-chlorophenol (2-CP). We have developed two assay sys­tems by which we are able to quantify the concentration of microbial species (as discrete colony-forming units) that are capable of degrading 2-CP. We have also addressed optimization of 2-CP metabolism as a func­tion of cosubstrate type and concentration. The data show that adaptive metabolic response of populations in natural and contrived wastewaters exposed to 2-CP during prolonged contact in mixed cultures is the result of population changes.

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MATERIALS AND METHODS

I. Growth and Isolation of Microorganisms

Sources of microorganisms for this study were activated sludge from the Roanoke sewage treatment plant, Roanoke, Virginia, and a soil sample collected from an area used for storing creosote-coated lumber.

A. Isolates

One ml aliquots of activated sludge or 0.2 ml from 5 percent (w/v) moist soil suspensions in sterile water were inoculated to 100 ml portions of nutrient broth containing 200 mg/I phenol. After incubation for two days at 28°C in a rotary shaker at 250 rpm, the cultures were plated on nutri ­ent agar containing 100 mg/I phenol (this medium was also used for main­taining the isolated cultures). All single-colony isolates were purified by restreaking on the above medium and inspected by microscopic exami­nation for homogeneous morphology.

B. Mixed Culture

The undefined mixed culture was established by inoculating 1 liter of nutrient broth (Difeo Laboratories, Detroit, Michigan) containing 50 mg phenol with two soil samples and one activated sludge sample, as de­scribed above. The complex inoculum was incubated at 30°C with agi­tation at 220 rpm with a two-inch throw on a New Brunswick Psychro­therm shaker (New Brunswick Science Co., New Brunswick, New Jersey). After 24 hours, the entire volume was harvested by centrifugation at 12,000 x g for 20 minutes and washed four times with equal volumes of sterile 0.1 M potassium phosphate buffer at pH 7.0. Finally, the cells were resusupended in phosphate buffer to form a thin cell paste.

C. Continuous Culture

The cell paste from the mixed culture was used as inoculum for continu­ous culture on medium A (described below) supplemented with 200 mg/I 2-CP (Eastman Organic Chemicals, Rochester, New York). The condi­tions for continuous cultivation were 500 ml operating volume, a dilu­tion rate of 0.035h-1 , automatic pH control at pH 7.0 with 0.1 M NaOH and 0.1 M HCI, an air-flow rate of 500 ml/minute, agitation by turbine impeller at 400 rpm, and temperature control at 30°C. The 2-CP con-

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centration in the feed was gradually raised over an eighteen-week period from 20 mg/I to 120 mg/I to force culture acclimation. The continous culture was maintained at steady state cell concentration with 120 mg 2-CP/1 of influent medium for two weeks at which time the cells were harvested by centrifugation at 12,000 x g for 20 minutes and stored in discrete aliquots in liquid nitrogen.

D. Experimental I nocula

The acclimated mixed culture stored in liquid nitrogen served as the ino­culum for studies on acclimated mixed cultures. As needed, frozen ali­quots were recovered and inoculated into mixed cultures or parallel liquid media containing 50 ml of medium A supplemented with 200 mg/I yeast extract and either 80 mg/I 2-CP for acclimated culture or 800 mg/I glu­cose for unacclimated culture. Both cultures were grown in 250 ml Erlen­meyer flasks at 3D°C with agitation (250 rpm) to a minimum of threefold increase in cell mass, determined by turbidity using a Klett-Summerson Photo-electric Colorimeter, and transferred to fresh medium. The res­pective cultures served as acclimated and unacclimated inocula for experi­ments.

II. Media

A. Medium A

Medium A had the following composition: Na2 HP04 , 0.2 g; N H4 Cl, 0.12 g;CuS04 • 5H 2 0,0.001 g; FeS04 • 7H 2 0,0.006g;MgS04 • 7H 2 0, 0.1 g; ZnS04 • 7H 2 0, 0.003 g; tap water, 200 ml; deionized water to 1 liter. Adjustments to pH were made with 0.1 N HCI or 0.1 N H3 P04 .

The pH before autoclaving was 7.5 unless otherwise noted. Phenol and 2-CP were added to the autoclaved, cooled medium, without steri I ization, in the final concentration indicated. Glucose, cellobiose, yeast extract, and casamino acids were added to autoclaved, cooled medium as filter­sterilized solutions. Solid medium was prepared by the addition of 18 g/I bacto agar (Difeo Laboratories, Detroit, Michigan) before autoclaving.

B. Medium B

Medium B had the following compos1t1on: (NH 4 ) 2 HP04 , 0.1 g; (NH 4 )2 S04 , 0.25 g; CuS04 • 5H 2 0, 0.001 g; FeS04 • 7H 2 0, 0.006y; MgS04 • 7H 2 0. 0.05 g; ZnS04 • 7H 2 0, 0.002 g; casamino acids, 0.1 g;

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yeast extract, 0.1 g; tap water, 200 ml; bromothymol blue or brom­cresol purple, 0.04 g; a·nd deionized water to 1 liter final volume. Yeast extract, casamino acids, and 2-CP were added to autoclaved, cooled med­ium as described above. The pH of medium 8 was adjusted to 7.0 as de­cribed for medium A. Sol id medium was prepared by addition of 18 g/I bacto agar (Difeo Laboratories, Detroit, Michigan) before autoclaving.

111. Determination of 2-CP and Phenol

A. Gas Chromatography

Liquid cultures were freed of the microbial cells by centrifugation and 1-2 µI portions of the supernatants were used to determine 2-CP, and phenol by gas chromatography with a Perkin-Elmer GC model 3920, equipped with a Poropak Q column, hydrogen flame ionization detector and Autolab digital intergrator, model 6300. The column temperature was 180°C and 2,4,6-tribromophenol was used as an internal standard.

B. Distillation

In some samples, 2-CP and phenol were determined according to the direct condensation of 4-aminoantipyrine with .aromatics and subse­quent oxidation to colored products (red). Three ml samples were com­bined with 0.1 ml of 5 percent K3 Fe(CN )6 in 0.1 M glycine buffer at pH 9. 7 and 0.5 ml of 5 percent 4-am inoantipyrine in 0.1 M glycine buffer at pH 9.7 and mixed thoroughly. Absorbance was read at 505 mm after 10 minutes. The assay was accurate to 0.5 mg/I± 0.25 2-CP or phenol. The liquid media, containing no pH indicator dyes, did not interfere with the test. Loss of 2-CP due to adsorption onto solids was noted only in the case of high concentrations of yeast extract (1.2 g/I); however, the distillation aminoantipyrine method allowed quantitative recovery and assay of ad­sorbed 2-CP. All assays were performed with a minimum of three repli­cates.

IV. Determination of Glucose and Cellobiose

Concentrations of glucose and cellobiose were measured by the Somogyi titration method for reducing sugars [Somogyi, 1952].

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V. Determination of Oxygen Consumption Rate

Dissolved oxygen was measured with an Instrumentation Laboratories dissolved oxygen meter and probe (Model 531 ).

A. Isolates

The consumption rate of oxygen was measured in completely filled, closed bottles containing oxygen-saturated fresh medium and fitted with a gas-tight stopper. The linear decline in oxygen concentration was meas­ured with an oxygen-specific electrode mounted through the gas-tight stopper. Consumption rate was calculated from the slope of the plot of dissolved oxygen concentration versus time.

B. Batch

Oxygen uptak~ rate in the batch reactor study was determined by moni­toring the initial rate of decrease in dissolved oxygen concentration after momentary interruption of the air supply to the reactor. Dissolved oxy­gen tension was never allowed to drop below 2 mg/I du ring the experi­ments.

VI. Determination of Chloride Ion

Chloride was measured by the method of Bergmann and Sonik [ 1957] and confirmed by a chloride-specific electrode.

VI I. Pure-Culture Microbial Degradation of 2-CP

Sixteen isolates, obtained from wastewater and soil, were screened to determine their ability to degrade 2-CP in 20 ml portions of liquid me­dium A containing 200 mg/I yeast extract and 20 mg/12-CP. The medium did not contain a pH indicator. The medium was inoculated with 0.4 ml portions of washed cell-suspensions of the respective isolates, resuspended to identical optical densities, and incubated at 3Cf C. Growth was assessed a~ter 21 hours incubation in medium A with yeast extract and 2-CP by relative turbidity. At that time five replicate samples were taken from each flask and assayed for residual 2-CP content.

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VI 11. Preparation of Isolates RS 10 and RS 17 for Testing the Differential Plate Count Technique

The pure cultures of RS 10 and RS 17 were grown for 18 hours in me­d iu m A supplemented with 20 mg/I of 2-CP and 200 mg/I of yeast extract. Cells from the two cultures were recovered by centrifugation, washed two times in 0.1 M potassium phosphate buffer at pH 7 .0, and finally resuspended in phosphate buffer. The cell concentration of each inocu­lum was determined by direct microscopic count in a hemocytometer. Appropriate volumes of the two cell suspensions were mixed so that in two final mixtures the ratios of 2-CP-degrading to 2-CP-resistant, but nondegrading, cells were 5: 1 and 1 :5 respectively. These inocula sources were serially diluted and placed on trypticase soy agar plates (for total viable counts) and plates of medium B containing 40 mg/I of 2-CP. The inoculated plates were incubated at 30°C for 36 hours and counted. RS 10 is a rod-shaped bacterium; RS 17 is a yeast.

IX. Preparation of Undefined Mixed Batch Cultures for 2-CP Shock Studies

Acclimated and u nacclimated mixed cultures were developed as described above and characterized by a modification of the most-probable-number technique. The 2-CP degrading and resistance characteristics of the accli­mated and unacclimated cultures are given in Table 5.

Four stirred batch reactors, each containing two liters of medium B with­out pH indicator but with 1.5 g/I glucose, were inoculated with accli­mated or u nacclimated cells and allowed to attain steady state rates of oxygen consumption. Two reactors were inoculated with identical masses of unacclimated culture equivalent to 0.7 g dry weight/I. The other reac­tors were inoculated with acclimated culture to a final cell mass of 1.04 g dry cell weight/I. 2-CP was added to the reactors after 4 hours incubation and the oxygen uptake rate monitored for both acclimated and unaccli­mated cultures at dose levels of 20 and 100 mg of 2-CP per liter. All four cultures had achieved steady state oxygen uptake by four hours post­jnocu latior;i. No significant increase in dry cell weight was observed during 1:his stabilization period. Oxygen uptake rates were determined by momentarily interrupting the flow of oxygen to each reactor and moni­toring the linear decrease in dissolved oxygen concentration for several minutes. The initial rate of dissolved oxygen reduction was taken as the oxygen uptake rate in each case. Air sparging was resumed in each case

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as the dissolved oxygen concentration approached 30 percent of the satu ­ration value.

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RESULTS AND DISCUSSION

I. Microbial Degradation of 2-CP by Pure Cultures

The isolates listed in Table 1 with RS-prefixes were obtained from a municipal activated sludge treatment plant; those designated by CP-pre­fixes were obtained from soil which had been consistently exposed to phenolic compounds for several years. Table 1 shows that a range of growth sensitivity to the presence of 2-CP exists and that in some cases (CP 12 and CP 13) cultures, normally able to grow in the presence of phe­nolic materials, are completely inhibited by the presence of 20 mg/12-CP. It is also apparent that extensive microbial growth in the presence of 2-CP is unnecessary for significant removal of this material (RS 17). Two iso­lates (RS 17 and CP 6A) from this study were selected as potential 2-CP degraders and carried forward to chlorophenol metabolism studies with pure and mixed, but defined, cultures.

11. Microbial Degradation of 2-CP by Mixed Populations

An undefined population of cells including at least one yeast and several bacteria were grown in continuous culture in the presence of 2-CP for eighteen weeks. The stable culture (constant dry cell weight) was the source of acclimated mixed culture inocula for subsequent study. Unac­climated inocula were prepared by growinq out the acclimated culture inocula in medium containing yeast extract and glucose, but no 2-CP. The relative populations shifted during growth in the absence of 2-CP, but no loss of morphological type was observed.

Batch cultivation of the acclimated and u naccl i mated cultures in medium A supplemented with 40 mg/I 2-CP for 86 hours at 30°C showed that the acclimated and unacclimated cultures consumed 70 percent and 20 per­cent of the 2-CP respectively.

The effect of elevated levels of 2-CP (100 mg/I) on acclimated and unaccli­mated mixed culture oxygen uptake rate is shown in Table2. The oxygen uptake rates of the u nacclimated culture were more sensitive to in hibi­tion by high concentrations of 2-CP than were the oxygen uptake rates of the equivalent acclimated culture. This observation is consistent with the literature [ Broecker and Zahn, 1977].

During preliminary studies on differences between acclimated and unac-

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climated cultures a difference was noticed in the rates of growth of the respective cultures in the presence of identical concentrations of 2-CP depending on the presence of additional carbon sources. Table 3 shows the affect of various alternative carbon sources on the metabolism of 2-CP by the acclimated mixed culture. Efficient metabolism of 2-CP re­quired the presence of an alternative substrate although some degradation occurred without cometabolism.This observation suggests that truecome­tabolism is not necessary for 2-CP metabolism. However, no net growth occurred in the absence of an alternative substrate. Thedisti llation method for assay of 2-CP and phenol used in this study allowed complete recovery of adsorbed 2-CP and phenol.

The addition of 200 mg/I yeast extract results in somewhat more rapid metabolism of phenol and 2-CP (Table 3). However, the extent of metab­olism of the two phenolic compounds is not dramatic. Excessive levels of yeast extract (1,800 mg/I) result in less extensive metabolism of both 2-CP and phenol. Medium supplemented with 1,800 mg/I yeast extract supported higher levels of culture growth as indicated by increased opti­cal density.

Phenol and glucose in the presence of 200 mg/I yeast extract support roughly the same level of 2-CP metabolism, indicating a lack of specificity for cosubstrate. Preliminary studies showed the presence of cellobiose and 200 mg/I yeast extract supported a more rapid rate of 2-CP metabo­lism during the early hours of incubation. Table3showsthatglucosewas consumed early in the fermentation while cellobiose was not utilized until the latter hours.

Although carbon cosubstrate availability may be a factor inefficient meta­bolism in the presence of cellobiose, the difference in early rates of meta­bolism shown in Table 3 suggests that 2-CP metabolism is favored by slower rates of growth. Free glucose was not detectable in the flasks con­taining cellobiose and yeast at any time during the fermentation. No di­auxic phenomena were observed with any of the cosubstrates.

111. Changes in Microbial Populations Exposed to 2-CP

The ability of wastewater treatment systems to remove biologically toxic materials depends on the recent history of the system.System acclimation may involve changes in the relative abundance of microbial subpopula­tions and/or changes in metabolic activities which are independent of sub-

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population shifts. Detection of subpopulation shifts requires the ability to quantify individual microbial types in mixed cultures. In the case of 2-CP removal, this might conventionally involve plating of serially diluted wastewater samples on total-count agar plates, screening of each pure iso­late in a medium containing 2-CP, and determining the residual 2-CP in the incubated cultures by gas chromatography or spectrophotometric methods. Routine determination of residual 2-CP in each culture is, at

··best, impractical.

Ralston and Vela [ 1974] have suggested the use of a liquid medium to screen wastewater microoganisms, previously isolated on trypticase soy agar (TSA) plates, to detect phenol-degrading microbes. The method works because several metabolic products of phenol degradation by micro­organisms are acidic. The decline in the pH of the liquid medium is visi­bly indicated by the dye bromothymol blue.Cultures that exhibit a change in color from the original blue to yellowish-green are scored as phenol­degrading microbes. These workers invariably observed that a decline in pH corresponded to the degradation of phenol. Th is procedure, however, requires testing of a large number of isolates to obtain statistically relia­ble data. Isolation of microorganisms on TSA plates would require a mini­mum of four days. Our preliminary studies on this system show that some phenol degrading isolates fail to show a positive test within 24 to 48 hours in Ralston and Vela [ 1974] medium ( 13) if the initial phenol con­centration is greater than 400 mq/I, unless a large inoculum is added or the medium is supplemented with growth-promoting cofactors. Con­versely, at low concentrations of phenol (200 mg/I), partial deqradation of phenol by some isolates goes undetected because of the high buffering capacity of the medium.

As a result of these preliminary experiments, we decided to attempt devel­opment of an agar plate assay based on the technique described above for phenol. The agar plate technique would allow more direct assessment of 2-CP degradinq microorganisms than the analoqous liquid culture assay. The procedure relies on formation of acids from incomplete metabolism of the phenolic moiety of 2-CP.

A liquid medium composed only of sewage waters and 40 mg/I of added 2-CP was used to study chlorophenol biodegradation by isolates RS 17 and CP 6A. These cultures cou Id degrade 2-CP to less than 0.5 mg/I within 100 hours. However, when this liquid medium was supplemented with chlorophenol, agar and bromothymol blue to prepare agar plates, neither

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of the isolates showed any decoloration in the purple plates. The buff­fering capacity of sewage water medium was determined by titration with HCI and found to be quite high (0.17 m eq H+/ml of medium required for pH indicator endpoint). The buffering capacity masked the minute quantities of acidic metabolic products of phenol. This medium was, therefore, unsuitable for the visual detection of the acidity attendent with 2-CP biodegradation in the agar plates.

Based upon the information gathered from the preceding experiments, a new liquid medium (medium B) was formulated. Medium 8, solidified by agar, was designed with very weak buffer capacity (0.03 m eq H+/ml of medium for pH indicator endpoint) and did allow detection of acidic metabolities of 2-CP. Preliminary studies utilized bromothymol blue as the pH indicator but resulted in an occasional false-positive scoring for 2-CP metabolism in the presence of yeast extract. The color transition with bromothymol blue ranged from light blue at pH 7.5 to yellow at pH 6.0. Bromcresol purple (pH end point for yellow color formation, 5.2) was ultimately substituted for bromothymol blue to increase the pH range. The substitution of bromcresol purple eliminated false-positive scoring because of formation from sources other than 2-CP. The presence of yeast extract and vitamin-free casamino acids in medium B allowed detectable growth of all the isolates listed in Table 1. Only those colonies surrounded by a yellow halo were scored as positive for 2-CP degradation. A test of this procedure with six isolates, RS 2A, RS 10, CP 4, RS 9, RS 17, and CP 6A, showed that only the last three gave positive results.

A check of the ability of the agar plate assay to accurately determine total counts and specific counts of 2-CP-degrading microorganisms was accomplished with a mixture of RS 17, a 2-CP degrading yeast, and RS 10, an unidentified bacterial rod capable of growing in the presence of 2-CP but incapable of metabolizing the chlorophenol. The results shown in Table 4 indicate that 2-CP degrading microorganisms may be detected and enumerated directly by this differential plate count method. The agar plate method utilizing medium B accurately determines the percent­age of microorganisms capable of degrading 2-CP but cannot replace total count techniques on richer media. Similiar results were obtained with mixtures of up to five different isolates of varying capabilities for degradation of 2-CP. This assay system has not been tested in undefined systems.

Although the agar plate method succeeded in identifying organisms capa-

15

ble of metabolizing 2-CP in a defined mixed culture, the assay wash ighly sensitive to incubation time and temperature. Incubation periods of less than 24 hours did not allow sufficient shift in the color of the pH indica­tor to guarantee positive scoring. Incubation periods in excess of 36 hours resulted in spreading and fading of the pH color zone asa result of acid diffusion and led to false-positive scoring. In all cases color develop­ment was slight.

A liquid medium assay was developed utilizing medium B supplemented with 2-CP. Serial dilution of the inoculum allowed determination of the most-probable-number of 2-CP-degrading microorganisms. The basic method involves inoculation of a series of test tubes containing 2-CP and bromothymol blue in medium 8. Measurement of culture turbidity after 18 hours is scored as an index of the ability of pure or undefined mixed cultures to grow in the presence of 2-CP. (Resistant cultures could be distinguished from chlorophenol-sensitive cultures by their ability to grow at higher concentrations of 2-CP.) Chlorophenol-degrading cultures were tentatively scored by their ability to cause a pH shift as indicated by pH indicator color shift after 36 hours. Tentative designation of chloro­phenol-degrading cultures was confirmed by analysis of released chloride ion and chromatographic determination of residual 2-CP. In all cases, formation of acid in the liquid medium as indicated by the pH indicator color shift coincided with confirmed metabolism of 2-CP as expected. Confirmed 2-CP-degrading microorganisms (RS 17 and CP 6A) did not give false-positive tests in the liquid medium in the absence of 2-CP or as a result of the presence of yeast extract and casamino acids. The liquid culture analogue of the agar plate technique disallowed the possi­bility of false-positive scoring resulting from the cross diffusion of the pH indicator dye while allowing an estimation of the number of chloro­phenol-degrading cultures.

A 10 percent-by-volume inoculum from the continuous culture vessel con­taining the acclimated, undefined mixed culture was transferred to asec­ond continuous culture setup utilizing medium A without 2-CP but with 65 mg/I glucose. The glucose-containing chemostat was allowed to reach steady state cell population (100 hours). Samples were withdrawn from the acclimated and unacclimated cultures and serially diluted for inocu­lation into the most-probable-number assay tu bes.

The results shown in Table 5 indicate a large difference between the acclimated and unacclimated cultures in the number of organisms per

16

ml capable of degrading 2-CP. The total plate counts shown in Table 5 from trypticase soy agar indicate that the number of 2-CP degrading and resistant microorganisms in the acclimated culture constitute roughly 62 percent of the total population. This latter comparison may be some­what inaccurate since DiGeron imo et al. [ 1978] suggest that the total plate count technique may overestimate the number of microorganisms present compared to general most-probable-number procedures. How­ever, one must conclude that within experi.mental system constraints, subpopulation profiles correlate well with acclimation to the presence·of 2-CP and suggest, as hypothesized, a mechanism for acclimation.

IV. System Response to Shock Load Doses of 2-CP

The response of batch-mixed cultures to shock load doses of 2-CP were studied in the acclimated and u nacclimated cultures defined above by the most-probable-number assay. Figure 1 presents the data of this experi­ment. After attaining steady state rates of oxygen uptake, the addition of 2-CP at 20 mg/I resulted in a rapid initial loss of respiration rate in both cultures. The loss of respiration activity by the unacclimated cul­ture was not regained and continued to decrease for several minutes after addition of the toxic compound, eventually stabilizing at roughly 20 per­cent of the initial oxygen uptake rate. The initial drop in dissolved oxy­gen uptake by the acclimated culture was followed by a rapid increase in oxygen uptake rate resulting in higher activity than observed prior to addition of 2-CP. The peak rise in respiration rate by the acclimated cul­ture was followed by approach to a steady state rate of oxygen uptake at approximately the same level of activity as observed prior to addition of 2-CP. Although both the acclimated and unacclimated cultures were affected by addition of 2-CP, the effect was not directly proportional to the differences in relative proportions of subpopulations, as is indicated by the data of Table 5.

Figure 1 also shows the effect of addition of 2-CP to a final concentration of 100 mg/I. The results show that this concentration of 2-CP is highly toxic to the unacclimated culture and results in a rapid and long-lasting loss of oxygen uptake capability. The slight recovery of oxygen uptake capability near the end of the experiment may indicate some degree of recovery from toxic response. Addition of 2-CP to a final concentration of200 mg/I in the acclimated culture also resulted in a loss of respiration but to ·a much 1.esser extent than observed with the u nacclimated culture. Althoygh respiration activity by the acclimated culture continued to drop

17

slowly after pulse addition of 100 mg/I 2-CP, the final respiration rate was approximately 40 percent of the initial steady state oxygen uptake rate. Clearly, the acclimated culture containing a large subpopulation of 2-CP degrading microorganisms is less sensitive to pulsed addition of 2-CP than is a system containing a small subpopulation of degrading micro­organisms. This observation, although certainly not unexpected, does support the conclusion that the modified most-probable-number proce­dure developed in this study accurately predicts the ability of a culture to withstand pulsed addition of a toxic compound. The data of Figure 1 compared to those of Table 5 suggest that resistance of a culture to sudden exposure to toxic levels of 2-CP is not simply a linear function of the number of 2-CP degrading microorganisms present. It is tempting to speculate that surface adsorption may play a role in this disproportion­ate response.

18

CONCLUSIONS

This study has shown that standard agar and liquid culture techniques for evaluation of pure and mixed cultures can be modified to allow for an estimate of the resistance to and degradation of 2-CP. The techniques developed here shou Id be usefu I in predicting the ability of waste treat­ment facilities to cope with pulsed loads of 2-CP and may be extended to other toxic compounds. The most powerfu I assay in th is regard is the modified most-probable-number indicator assay, which by differential reading at 18 and 36 hours allows correlation of 2-CP resistant and 2-CP degrading subpopulations of a culture. Th is assay confirms the expected changes in microbial subpopulations that occur during acclimation of cultures to 2-CP.

The specific conclusions based on experimental data follow:

1. metabolism of 2-CP can occur in an undefined mixed culture in the absence of a carbon cosubstrate;

2. metabolism of 2-CP in the absence of cosubstrate occurs without growth of the culture;

3. the presence of cosu bstrates may increase the rate and extent of 2-CP metabolism;

4. cellobiose allows more complete-metabolism of 2-CP when pres­ent as a cosubstrate than does glucose, possible because of longer availability (slower metabolism);

5. the oxygen uptake rate of an unacclimated culture is, as expected, more severely inhibited by 2-CP than is the oxygen uptake rate of an acclimated culture;

6. the modified differential plate count method developed in this paper estimates both the resistance to the toxic response and degradation of 2-CP in mixed cultures and suggests that differ­ences exist between the two classifications, i.e. apparently, resis­tance to 2-CP can occur without concomitant degradation;

7. the modified most-probable-number indicator assay developed here can allow relative measure of 2-CP sensitive and 2-CP resis-

19

tant members of the total population in undefined mixed cul­tures. The technique enables one to estimate the size of the 2-CP degrading subpopulation; and

8. undefined cultures that contain a large subpopulation of 2-CP degrading microorganisms, as defined by the modified most-prob­able-number technique, are, as expected, more resistant to toxic levels of 2-CP than cultures containing a small subpopulation of 2-CP degrading microbes. We conclude that the modified most­probable-number technique is a convenient, rapid, and accurate method for prediction of both resistance to and degradation of 2-CP by mixed, undefined cultures.

20

LITERATURE CITED

Bergmann, J. G. and J. Sanik, 1957. "Determination of Trace Amounts of Chlorine in Naphtha." Analytical Chemistry 29:241-45.

Broecker, B. and R. Zahn, 1977. "The Performance of Activated Sludge Plants Compared with the Results of Various Bacterial Toxicity Tests­A Study with 3,5-Dichlorophenol." Water Resources 11 :165-72.

U. S. Environmental Protection Agency, 1977. Development Document for Effluent Limitations: Guidelines for Petroleum Refineries. EPA 410/ 1-7 4-014-a.

Di Geronimo, M. J., M. Nikaido, and M. Alexander, 1978. "Most-Probable­Number Technique for the Enumeration of Aromatic Degraders in Natu­ral Environments." Microbial Ecology 4:263-66.

Fryer, J. D. and K. Kirkland, 1970. "Field Experiments to Investigate Long-Term Effects of Repeated Application of MGPA, Tri-allate, Sima­zine and Linuron." Weed Research 10: 133-58.

Hood, M. A. et al., 1975. "Microbial Indicators of Oil-Rich Salt Marsh Sediments." Applied Microbiology 30:982-87.

Horvath, R. S., 1972. "Microbial Co-Metabolism and the Degradation of Organic Compounds in Nature." Bacteriological Review 36: 146-55.

Horvath, R. S. and P. Flath man, 1976. "Co-Metabolism of F luoroben­zoates by Natural Microbial Population." Applied Environmental Micro­biology 31 (6) :889-91.

Hurle, K. and B. Rademacher, 1970. "Untersuchungen uber den Einfluss Langjahrig Wiederholter Anwendung Von DNOC und 2.4.-D auf ihren Abbau i m Boden." Weed Research 10: 159-64.

Johnston, H. W., and G. G. Briggs, and M. Alexander, 1972. "Metabolism of 3-Chlorobenzoic by a Pseudomonad." Soil Biology and Biochemistry 4: 187-90.

Kirkland, K., 1967. "Inactivation of MCPA in Soil." Weed Research 7: 364-67.

21

Milne, G. W. A., P. Goldmand, and J. L. Holtzman, 1968. "The Metabo­lism of 2-Flurorbenzoic Acid; 11, Studies with L802 ."Journal of Biologi­cal Chemistry 243:5374-76.

Ralston, J. R. and G. R. Vela, 1974. "A Medium for Detecting Phenol­Degrading Bacteria." Journal of Applied Bacteriology 37:347-51.

Somogyi, M., 1952. "Notes on Sugar Determination." Journal of Biologi­cal Chemistry 159: 19.

Walker, J. D. and R. R. Colwell, 1973. "Microbial Ecology of Petroleum Utilization in Chesapeake Bay." In APl/EFA/USCS Conference on Pre­vention and Control of Oil Spills Proceedings, p. 685-91. American Petro­leum Institute, Washington, D. C.

Watanabe, I., 1977 "Pentachlorophenol-Decomposing and PCP-Tolerant Bacteria in Field Soil Treated with PCP." Soil Biological Biochemistry 9:99-103.

22

FIGURE

w f­c::( 0:::

w ::..: c::( f­a...

120

100

:::> 4 z: w (.!:)

>­x 0

LL 0 20 z: 0

f­

c:o I

FIGURE 1 Effect of Pulse Addition of 2-CP

at Two Levels to Undefined Mixed Cultures*

SYMBOL CULTURE INITIAL 2-CP INITIAL OUR (mg/1-1) (mg/1- 1/min- 1)

0 ACCLIMATED 20 8.7

• ACCLIMATED 100 8.2

6. UNACCL I MATED 20 6.1

• UNACCLIMATED 100 6.7

ADDITION OF 2-CP

z 0l--1..~..A::,;IU~IL.t~...&......A.A....iL.IL__.~~:::::::;~=-i__~~__i____J

0 2 3 4 5 6

TIME (hrs)

*The acclimated culture was prepared by long-term growth in the presence of 2-CP; approximately 33 percent of the population was capable of rapid degradation of 2-CP (Table 5). The unacclimated culture was prepared by long-term growth of the mixed, accUmated culture in the absence of 2-CP but in the presence of glucose; less tpan 0.01 percent of the population could rapidly degrade 2-CP (Table 5). Toxicity is measured as inhibition of oxygen uptake rate (OUR).

24

TABLES

TABLE 1 Growth and Chlorophenol Metabolism of Microbial Isolates

in the Presence of 20 mg/I 2-CP

Isolate Identification Growth, Residual Number Klett Units 2-Cp mg/I* ,t

RS 2 0 20 RS 2A 120 18 RS 4 134 18 RS 7 6 20 RS 8A 118 17 RS 9 154 16 RS 10 162 19 RS 128 2 20 RS 17 16 14 RS 19 42 17 RS 20 4 18 RS 22 4 20 CP 1 96 16 CP 4 146 18 CP 6A 122 12 CP 12 0 20 CP 13 8 20

*Isolates were inoculated into medium A with 200 mg/I yeast extract and 20 mg/I 2-CP from nutrient broth containing 10 mg/I phenol after dilution to identical optical density. Relative growth and residual 2-CP were determined after 21 hours in cu bati on at 30°C. Growth was determined by turbidity with a Klett-Sommerson colorimeter. Data were the averages of five replicate samples from each culture. tResidual 2-CP was determined by gas chromatography.

26

Time, (hrs)

0.0 0.25 0.5 1.0 2.0 3.0

TABLE 2 Influence of 2-CP on the Oxygen Respiration Rate of

Acclimated and Unacclimated Undefined Mixed Cultures*

Acclimated

0 7

23 38 42 40

Oxygen Uptake Rate % Inhibition

Un acclimated

0 34 56 84 86 82

*Percent inhibition in respiration rate compared to respective controls growing in the absence of 2-CP. Acclimated and unacclimated inocula were taken from rapidly growing cultures (12 hours old) and added to medium A with 200 mg/I yeast extract and 10 mg/I glucose and 100 mg/I 2-CP.

27

N

TA

BL

E 3

co

C

om

eta

bo

lism

of

2-C

P b

y a

n A

cclim

ate

d M

ixe

d C

ult

ure

Co

nce

ntr

atio

n o

f 2-

CP

*, P

heno

l, G

luco

se,

or

Cel

lobi

ose t,

mg/

I T

ime

Aft

er

Ino

cula

tio

n (

hrs)

A

dd

itio

ns

to

Sal

ts M

ed

ium

A

pre

-in

ocu

latio

n

5 1

2

20

2

6

40

5

3

76

9

8

12

5

18

0

2-C

P

40

40

3

9

39

3

9

39

3

9

38

37

32

29

2-C

P

41

41

36

36

36

32

31

2

5

22

19

12

P

heno

l 27

2

6

26

25

25

2

5

22

0

0 0

0

2-C

P

41

39

37

34

3

3

31

27

2

0

19

15

10

Phe

nol

26

26

25

24

24

10

0

0 0

0 0

Yea

st E

xtra

ct (

200

mg/

I)

2-C

P

39

36

34

34

34

34

2

9

26

2

6

25

20

P

heno

l 27

27

27

27

27

27

27

26

2

2

18

0 Y

east

Ext

ract

( 1

,80

0 m

g/I)

2-C

P

40

3

6

31

31

31

27

27

22

2

2

21

14

Yea

st E

xtra

ct (

200

mg/

I)

2-C

P

40

33

33

30

30

29

2

8

27

26

16

16

G

luco

se (

65 m

g/I)

65

4

2

24

0 0

0 0

0 0

0 0

Yea

st E

xtra

ct (

200

mg

/I)

(co

nti

nu

ed

)

I\.)

co

TA

BL

E 3

(co

nti

nu

ed

)

Con

cent

rati

on o

f 2-

CP

*, P

heno

l, G

luco

se,

or

Cel

lobi

oset

, m

g/I

Tim

e A

fter

Ino

cula

tion

(hr

s)

Add

itio

ns t

o

Sal

ts M

ediu

m A

pr

e-in

ocul

atio

n 5

12

20

26

4

0

2-C

P 4

0

34

32

2

7

23

18

C

ello

bios

e (1

15 m

g/I)

11

5 10

8 10

0 72

70

4

6

Yea

st E

xtra

ct (

200

mg/

I)

*Det

erm

ined

by

the

amin

oant

ipyr

ine

met

hod

and

gas

chro

mat

ogra

phy

of

dist

illa

tes.

tD

ata

repr

esen

t th

e av

erag

e o

f 4

repl

icat

es.

53

1

6

38

76

9

8

12

9 11

0

125

18

0

4 <

2

0 0

w

0 T

AB

LE

4

En

um

era

tion

of

2-C

P D

egra

ding

Mic

roor

gani

sms

in D

efin

ed

Mix

ture

s

Co

ntr

ive

d

% M

icro

bia

l T

ype

T

ota

l V

iab

le

Co

un

ts o

n M

ed

ium

B P

late

st,:f

:

2-C

P

Non

-2-C

P

Co

un

ts o

n T

SA

* ,:f:

T

ota

l C

ou

nts

Deg

rade

rs

Deg

rade

rs

Pla

tes

cfu

/ml

(RS

17)

(R

S 1

0)

cfu

/ml

x 10

6 x

106

80

20

6.

4 5.

5 (1

00%

)

20

80

5.

2 4

.4

(100

%)

*Try

pic

ase

soy

aga

r (T

SA

).

tPer

cent

age

on 2

-CP

-deg

radi

ng m

icro

orga

nism

s ar

e ca

lcu

late

d o

n t

he

bas

is o

f to

tal

cou

nts

on

Me

diu

m B

. :f:

Cfu

: co

lon

y fo

rmin

g u

nits

aft

er

36

hou

rs a

t 3

0°C

.

2-C

P-D

egra

der

cfu

/ml

x 10

6

4.7

(85%

)

0.6

(14%

)

TABLE 5 Number of Cells Able to Grow (Resistant) and Able to Grow and Degrade

2-CP in Acclimated and Unacclimated Cultures*

Culture Condition

Acclimated

Unacclimated

Number of cells/ml MPN Count

Resistant

4.5 ± 0.3 x 106

7.2 ± 0.5 x 104

Degradingt

3.9 ± 0.5 x 106

1.2 ± 0.8 x 102

TSA Plate Count

1.2 ± 0.4 x 107

8.0 ± 1.2 x 106

* Most-probable-number (MPN) tubes were incubated for 28 hours at 30°C with shaking. Counts were corrected for growth due to carry -over of nutrients by incuba­tion in the absence of added carbon sources in Medium B. MPN values were obtained from tables for five tubes inoculated with successive tenfold dilutions. t Only those dilutions which allow both growth and color shift were scored as de­grading. Color change was not observed without growth (turbidity).

31

The Virginia Water Resources Research Center is a federal-state partnership agency attempting to find solutions to the state's water resources problems through careful research and analysis. Established at Virginia Polytechnic In­stitute and State University under provisions of the Water Research and Devel­opment Act of 1978 (P.L. 95467), the Center serves five primary functions:

• It studies the state's water and related land-use problems, includ­ing their ecological , political, economic, institutional, legal, and social implications.

• It sponsors and administers research investigations of these prob­lems.

• It collects and disseminates information about water resources and water resources research.

• It provides training opportunities in research for future water scientists enrolled at the state's colleges and universities.

• It provides other public services to the state in a wide variety of forms .

More information on programs and activities may be obtained by contacting the Center at the address below.

Virginia Water Resources Research Center Virginia Polytechnic Institute and State University

617 North Main Street Blacksburg, Virginia 24060

Phone (703) 961-5624