Environ. Sci. Technol. 1904, 18, 967-972
(15) Cunningham,P. T.; Johnson, S. A. Science (Washington, D.C.) 1976, 191, 77-79. (16) Delumyea, R.; Macias, E. S.; Cobourn, W. G. Atmos. Environ. 1979, 13, 1337-1338. (17) Cobourn,W. G.; Djukic-Husar,J.; Husar,R. B. J , Geophys. Res. 1980,85, 4487-4494. (18) Cobourn, W. G.; Husar, R. B. Atmos. Environ. 1982,16, 1441-1450. (19) McMurry, P. H.; Rader, D. J.; Stith, J. L. Atmos. Environ. 1981,15, 2315-2327. (20) McMurry, P. H.; Wilson, J. C. Atmos. Environ. 1982, 16, 121-134. (21) Cantrell, B. K.; Whitby, K. T. Atmos. Environ. 1978,12, 323-333. (22) Husar, R. B.; Patterson, D. E.; Husar, J. D.; Gillani, N. V.; Wilson, W. E., Jr. Atmos. Enuiron. 1978, 12, 549-568. (23) White, W. H. Nature (London)1976,264, 735-736. (24) Pierson, W. R.; Brachaczek, W. W.; Truex, T. J.; Butler, J. W.; Korniski, T. J. Ann. N.Y. Acad. Sei. 1980, 338, 145-173. (25) Weiss, R. E.; Waggoner, A. P.; Charlson, R. J.; Ahlquist, N. C. Science (Washington, D.C.)1977,195, 979-981. (26) Samson, P. J.; Ragland, K. W. J. Geophys. Res. 1977,16, . . 1101-1106. (27) Hidy, G. M.; Mueller, P. K.; Tong, E. Y. Atmos. Environ. 1978,12,735-152. (28) Vukovich, F. M. Atmos. Enuiron. 1979, 13, 255-265. (29) King, W. J.; Vukovich, F. M. Atmos. Environ. 1982, 16, 1171-1181. (30) Wolff, G. T.; Kelly, N. A.; Ferman, M. A. Science (Washington, D.C.) 1981,211, 703-705. (31) Wolff, G. T.; Kelly, N. A,; Ferman, M. A. Water, Air, Soil Pollut. 1982, 18, 65-81.
author, page number) and prepayment, check or money order for $7.50 for photocopy ($9.50 foreign) or $6.00 for microfiche ($7.00 foreign),are required. Registry No* "2, 7446-09-5; '3, 1002815-6; H2S049 7664-93-9; (NH4&304,7783-20-2; ammonium, 14798-03-9.
Literature Cited Calvert,J. G.; Stockwell,W. R. Enuiron. Sci. Technol. 1983, 17,482A-443A.
Beilke, S.; Gravenhorst, G. Atmos. Environ. 1978, 12, 231-239.
Hegg, D. A.; Hobbs, P. V. Atmos. Environ. 1978, 12, 241-253.
Robbins, R. C.; Cadle, R. D. J. Phys. Chem. 1958, 62, 469-471.
Cadle, R. D.; Robbins, R. C. Discuss. Faraday SOC.1961, 30,155-161.
Huntzicker, J. J.; Cary, R. A.; Ling, C.-S. Environ. Sci. Technol. 1980, 14, 819-824.
McMurry, P. H.; Takano, H.; Anderson, G. R. Environ. Sei. Technol. 1983, 17, 347-352.
Camp, D. C.; Stevens,R. K.; Cobourn, W. G.; Husar, R. B.; Collins, J. F.; Huntzicker,J. J.; Husar, J. D.; Jaklevic, J. M.; McKenzie, R. L.; Tanner, R. L.; Tesch, J. W. Atmos. Environ. 1982, 16, 911-916.
Huntzicker,J. J.; Hoffman, R. S.; Ling, C.-S. Atmos. Environ. 1978, 12, 83-88.
Jaklevic, J. M.; Loo, B. W.; Fujita, T. Y. Enuiron. Sei. Technol. 1981,15, 687-690.
Tang, 1. N.; Munkelwitz, H. R.; Davis, J. G. J. Aerosol Sei. 1978,9, 505-511.
Charlson, R. J.; Vanderpol, A. H.; Covert, D. S.; Waggoner, A. P.; Ahlquist, N. C. Science (Washington,D.C.) 1974,184, 156-158.
Charlson, R. J.; Vanderpol, A. H.; Covert, D. S.; Waggoner, A. P.; Ahlquist, N. C. Atmos. Environ. 1974,8,1257-1267. Vanderpol. A. H.; Carsey, F. D.; Covert, D. S.; Charlson, R. J.; Waggoner, A. P. Science (Washington, D.C.) 1975, 190, 510.
Received for review January 23,1984. Accepted June 14,1984. This research was supported in part by US.Environmental Protection Agency Grant R804750 and by Cooperative Agreement CR807654. The paper has not been subjected to EPA$ peer review and therefore does not necessarily reflect the views of EPA. Thus, no official endorsement should be inferred.
Acute Toxicity Screening of Water Pollutants Using a Bacterial Electrode Elalne J. Dorward and 6. George Barisas" Department of Chemistry, Colorado State University, ~
Fort Collins, Colorado 80523 ~~
Escherichia coli electrodes were used in an instrumental bioassay of the acute toxicity of substances in water. The method involves potentiometric measurement of C02 production by E. coli cells immobilized a t the surface of a C02-sensing electrode. The net rate of COz production by the bacteria reflects the complex series of biochemical reactions which constitute the respiratory processes of the cells. The inhibition of any part of the respiratory process by some pollutant will result in a measurable decrease in bacterial COz production. The E. coli electrode is able to measure the acute toxicity of a broad range of substances, including metals, anions, gases, and organic compounds. Dose-effect curves obtained with the E. coli electrode are compared with results reported for the Beckman Microtox bioassay and for rainbow trout 96-h LC50values. Acute toxicity values measured with the E. coli electrode for cadmium, lead, copper, cyanide, and arsenite are comparable to those obtained with the 1Bmin Microtox bioassay. Introduction Water quality criteria for aquatic environments are based primarily on bioassays or toxicity tests. Acute toxicity tests are the first steps in determining, for a particular species, acceptable levels of a given chemical 0013-936X/84/0918-0967$01 SO10
substance. Many problems are associated with commonly used techniques for measuring acute toxicity. High cost, long experiment times, and requirements for specialized laboratories and much laboratory space are among the important problems. Quantitation of biological responses is another difficulty; the inherent variability of biological systems makes it difficult to measure chemical toxicity by purely biological means. To measure acute toxicity effectively, a system must provide a simple, sensitive, and rapid measurement of physiological parameters which are indicative of overall organism viability. Such parameters might be associated with a major metabolic process controlled by interdependent enzyme systems. Metabolic processes can be effectively monitored by electroanalytical techniques. For example, bioselective sensors, which use intact, living bacterial cells in place of isolated enzymes a t the surface of a membrane electrode ( I ) have been developed as extensions of enzyme electrodes. The measurement of L-glutamine in aqueous solutions and in human serum by a highly selective and sensitive potentiometric bacterial membrane electrode has been reported (1). Potentiometric bacterial membrane electrodes have been employed to measure L-aspartate (2),L-histidine (3),and L-arginine (4)
0 1984 American Chemical Society
Environ. Sci. Technol., Vol. 18, No. 12, 1984 967
by using an ",-sensing electrode. L-Cysteine has been measured by a bacterial electrode sensitive to HzS (5). Bacterial membrane electrodes have been developed to determine NO3- by using an ",-sensing electrode (6). The determination of ammonia under strongly alkaline conditions was accomplished by using immobilized nitrifying bacteria and an oxygen electrode (7). Karube et al. have reported two amperometric microbial electrode systems for screening mutagenic substances (8,9).Recently, Kobos and co-workers have reported the microbiological assay of several antibiotics using a potentiometric COz sensor (IO). The method is based upon inhibition of COz production in a suspension of bacteria after incubation with antibiotic. This article reports the development of a rapid, inexpensive, and quantitative instrumental bioassay for preliminary measurements of the acute toxicity of pollutants in water. The instrumental bioassay developed for this purpose is based on the Escherichia coli electrode. COz production by E. coli cells immobilized at the surface of a COz-sensingelectrode is measured potentiometrically. The production of COz by the bacteria is a result of the complex series of reactions which constitutes the respiratory process of the cells. The inhibition of any of these reactions by some pollutant results in a measurable decrease in COz production by the bacteria. The respiratory process presents many biochemical targets for toxic substances, permitting the E. coli electrode to measure the acute toxicities of a broad range of pollutants.
Materials Growth medium for E. coli wa prepared from 1.6 g of KZHPO4,1.6 g of KHZPO4,1.0 g of NaC1,2.5 g of glucose, 4.0 g of (NHJZSO4,0.7 g of MgSO,, and 0.5 g of sodium citrate dissolved in 1L of distilled water and brought to pH 7 with 1M K2HPO4. For growing large cultures, the medium was sterilized in 250-mL Erlenmeyer flasks with cotton plugs for 15 rnin at 120 OC. Piperazine-N,"-bis(2-ethanesulfonic acid) (PIPES) was purchased from U S . Biochemical Corp. PIPES buffers were prepared by using 1 M NaOH to dissolve and neutralize the free acid in distilled water. The following chemicals were used to prepare toxicant solutions: lead acetate, silver nitrate, cadmium acetate, sodium arsenite, p(ch1oromercuri)benzoic acid, phenol, calcium hypochlorite, ammonium hydroxide, sodium nitrite, and cupric chloride. The internal electrolyte for the Orion COz electrode was 0.01 M NaHCO, in 0.1 M NaC1. Calibration standards for the C02 electrode were prepared with NaHCOS. All compounds used were of analytical reagent grade except for PIPES and p(ch1oromercuri)benzoic acid, which were of U S . Biochemical Corp. buffer grade and Sigma Chemical Co. A grade, respectively. Double distilled water was used to prepare all solutions. Experimental Section Electrode Construction. Initial construction of the E . coli electrode followed procedures similar to those reported in the literature (2,4) and was based on the Orion Model 95-02 carbon dioxide electrode. Potentials were measured on an Orion Model 701A digital pH/mV meter in conjunctionwith a Varian A-25 strip-chart recorder. All experiments were conducted at 37 OC, and solutions were kept in a 37 "C water bath. To prepare an electrode, an aliquot of E. coli suspension was filtered onto a 13-mm diameter Nuclepore membrane filter of 0.45-pm pore size. The layer of cells on the Nuclepore membrane was placed in the bottom of the lower half of the outer body of the Orion COz electrode with the 968
Environ. Sci. Technol., Vol. 18, No. 12, 1984
cell layer inward. The gas-permeablemembrane was then placed on top of the cells, followed by the rubber O-ring. The electrode outer bodies were assembled and filled with internal electrolyte, and the inner body was placed inside the outer body (12). The assembled E. coli electrodes were conditioned in a beaker containing 0.05 M phosphate buffer or 0.05 M PIPES buffer for 10 min before using. Escherichia coli bacteria were chosen as the test bacterium since they have high levels of COz production, reproduce rapidly, and are easy to handle and grow in large quantities. E. coli R.F. Hill Strain B 23226 was purchased from the American Type Culture Collection, Rockville, MD. The bacteria were grown and maintained in the chemically defined medium described previously. The bacteria were grown in 250-mL Erlenmeyer flasks with side arms in a shaker-incubator at 37 OC. Growth of the E. coli populations was monitored by optical density of the cultures of 530 nm. The bacteria were harvested at the maximum stage of growth by centrifugation at 6000g for 15 min. The bacteria were washed 3 times with cold 0.05 M phosphate buffer, suspended in sterile buffer, and stored at 4 "C in 100-mL volumetric flasks. Cells were used within 40 h of harvest. Immobilization of the bacterial cells required a membrane that would allow diffusion of nutrient and toxicant molecules to the bacterial cells. Most of the bacterial electrodes reported to date have used a dialysis membrane for this purpose (2,643); however, advantages of using a sterilizing membrane filter have also been reported (3). E. coli electrodes were initially constructed by using both types of membranes for cell immobilization. Although comparable response times were obtained by using both membrane types, the sterilizing filters have greater stiffness and strength, making them easier to use. Also, filtering the bacteria directly onto the membrane helped in attaining a reproducible number of E. coli on each membrane. Nuclepore membrane filters with 13-mm diameter and 0.45-pm pore size were used to construct the E. coli electrodes used in this research. The E. coli electrode response to glucose-enriched buffer was measured for various sizes of bacterial populations immobilized. Varying volumes of E. coli suspension of known density were filtered onto 13-mm Nuclepore membranes. The average number of E. coli cells per milliliter of clean E. coli suspension was 4 X lo8 as determined by a viable count procedure (12). With 7 mL of cell suspension or 3 X lo9E. coli cells, approximately 3 min is required for the maximum steady-state level of C02 production to be obtained. An immobilized E. coli population of 3 X log cells reaches a steady state of approximately 0.9 mM COz at the gas-sensing electrode surface. Response times cited in the literature for bacterial electrodes range from 3 to 20 rnin (2, 4, 13, 14). The way electrodes are conditioned before use influences response time and linear range. Electrodes were conditioned by soaking the assembled electrodes in 0.05 M PIPES or 0.05 M phosphate buffer for approximately 10 min before use. Conditioning the bacterial electrodes before use stabilized the response of the immobilized cells to the glucose-enriched buffers. Experimental Procedure. In the E. coli electrode, the gas-permeable membrane separates the bacterial layer from the electrode internal electrolyte. COz produced by the immobilized cells causes an increase in the concentration of COz in the bacterial layer until a steady-state concentration of COz is reached, the production of COZ by the cells then being equal to its diffusion out of the bacterial layer. The COz present in the bacterial layer diffuses
I .o
j;
-
'
I
1
I
I
t
I
P.
0.6
E
n
0" 0.4
pF
0.2
0.0 0.0L 0.0
I
1
1
4.0 TIME (MINI
I
5
e .o
Flgure 1. Line (-) shows the E . co/i electrode signal in giucose-enriched phosphate buffer. Borken line (---) shows the change in COP production upon addition of 70 mM cyanide in glucose-enriched phosphate buffer. Arrow indicates introduction of cyanide. Curves were drawn from a strip-chart recorder trace. Individual points are not shown.
through the gas-permeable membrane and equilibrates with the electrode internal electrolyte. The resulting change in pH of the internal electrolyte is measured by the glass electrode. Hence, the apparent COz concentration measured by the carbon dioxide electrode is proportional to the COz production of the immobilized E. coli layer. Experiments were conducted at pH 7 in 0.05 M phosphate buffer or 0.05 M PIPES. Phosphate buffer was used in most experiments, except those involving toxicants precipitated by phosphate. In such cases, PIPES, which combines low metal-binding constants with a pK, of 6.8 (15), was used. Experiments involving the E. coli electrodes were performed in buffers containing 0.014 M glucose. Glucose served as the energy source for the immobilized cells. When the E. coli electrode is transferred from plain buffer to glucose-enriched buffer, an increased rate of COz production by the immobilized cells is immediately observed. Figure 1shows the increase in COz production to a steady-state level which occurs when an E. coli electrode is placed in glucose-enriched buffer. The protocol for each experiment involved adding 25 mL of glucose-enriched buffer to a 50-mL beaker containing a magnetic stir bar. The beaker was placed in a 37 OC water bath and the electrode tip immersed in the buffer. The increase in COz production by the immobilized E. coli was monitored on the pH /mV meter. When a steadystate level of COz was reached, a known quantity of substance (toxicant, nutrient, or blank) in glocuse-enriched buffer was added to the beaker. Response of the E. coli electrode to the substance was then monitored for 5-30 min. A fresh electrode was prepared for each experiment. Data Analysis. Data analysis for the E. coli electrode method of measuring acute toxicity involves two stages. In the first stage the electrical signal produced by the electrode is recorded and converted to a concentration of COP The second part involves using these raw data, COz concentration measured as a function of time, to estimate the toxicity of a test substance at its known concentration. The parameter used to measure the immobilized bacterial response to a toxicant is the change from the initial steady state of C 0 2production Pito some final steady state of COZ production Pp Figure 2 illustrates idealized re-
IO
TIME (MINI
1
Figure 2. Idealized change in COP production by the immobilized E. coli in response to applied toxicants. Electrode responses to substances that inhiblt COPproduction with decreasing speed are indicated by curves (-), (- -), and ( - . e -), respectlvely.
-
sponse curves such as might be obtained for toxic substances causing different rates of respiratory inhibition. The high initial rate of C 0 2 production is indicated by the curve to the left of the arrow. The arrow indicates the introduction of toxicant. Upon introduction of toxicant, the COz production is observed to fall exponentially to a lower final steady-state value. The solid line shows the change in COz production by the immobilized E. coli in response to a very rapidly acting toxicant where the rate of decrease in measured COz concentration is limited only by diffusion of COz out of the cell layer. We find experimentally that electrode response is generally slower than this limiting case and is dependent on toxicant concentration and that the change in eledrode response with time is exponential. The exponential kinetic response of the E. coli electrode when exposed to a substance that inhibits cellular respiration can be written as C ( t ) = A[Pf (Pi- Pf)exp(-kt)] (1) where C is the concentration of COz at the gas-sensing surface, k is the apparent first-order rate constant, t is the time, and A is a constant dependent upon electrode size and construction. This equation describes how the concentration of COz at the surface of the electrode-sensing element changes in an exponential manner from Pi to PP When it is impractical to measure Pf directly, due to the often slow electrode response, this equation is used to calculate Pp This is accomplished by evaluating k by using the Guggenheim method (16) and then plotting C ( t ) vs. exp(-kt) to yield a straight line with slope equal to A(Pi - Pf)and y intercept equal to A(Pf). The E. coli electrode response to a toxicant is expressed as the percent inhibition (% I) of initial COz production. Percent inhibition is defined by Pi - Pf I=100 (2)
+
pi
where % I is the percent inhibition of cellular respiration, Pi is the initial steady-state level of COz production, and Pf is the final steady-state level of COz production. Figure 1illustrates raw data. The line is the response of an E. coli electrode upon being placed in glucose-enriched phosphate buffer. The broken line is the response caused by the addition of 70 mM cyanide. The difference in the apparent COz production of the two electrodes is clear: the electrode exposed to cyanide rapidly approaches Environ. Sci. Technol., Voi. 18, No. 12, 1984
969
a steady-state level of C02production markedly lower than the initial C02 production level. This change in level of C02 production is used to calculate the cyanide-induced inhibition of respiration in the immobilized E. coli cells by using the equations presented previously. For the example given here, 70 mM cyanide causes 81% I of immobilized E. coli carbon dioxide production.
Results COPElectrode Measurement. Measurements of the 0.05 M phosphate and PIPES buffers with the Orion C02 electrode show that, at pH 7, the amount of atmospheric COP in the phosphate and PIPES buffers is below the detection limit of the C02electrode. Adding 1.2 mL of 1.0 M NaHCO, to the glucose-enriched buffer raises the measured C02 concentration to 0.95 mM, which is comparable to the CO2 production of an immobilized E. coli population. Adding small amounts of glucose-enriched buffer to such a C02-containingsolution causes no change in apparent C02concentration. Likewise, small amounts of the buffers containing test substances generally cause no change in the C02 electrode signal. However, some of the test substances do cause interference problems. Substances such as dissolved gases, organic acids, etc., which are soluble in silicone rubber, are able to diffuse through the electrode membrane (17). If such a substance participates in an acid-base equilibrium, it can cause interference by shifting the internal electrolyte pH. Sodium hypochlorite, sodium nitrite, phenol, and sodium arsenite cause interferences with the COPelectrode. Such interference causes a change in the apparent C02concentration of the sample which depends on the level of the interfering substance but which is approximately independent of the actual C02concentration. Correction for such interference can therefore be accomplished by comparing the response of the COPelectrode to a C02-containingbuffer with and without the interfering substance. The difference in apparent C02concentrations can then be used to correct COP levels determined for the bacterial electrode in solutions containing the same level of interfering substance. Only a single bacterial electrode is thus required for toxicity measurements, even when the toxicant interferes with the C02 sensor. Toxicity Measurements. Substances were chosen for study based on their importance in water quality monitoring and on the availability of acute toxicity information. The E. coli electrode was evaluated with the following substances: cadmium, copper, lead, zinc, silver, chlorine, ammonia, nitrite, cyanide, phenol, 2,4-dichlorophenoxyacetic acid, arsenite, p-(chloromercuri)benzoate, and the commercial wood treatment formulation CCA which contains mainly o-arsenic acid, copper(I1) oxide, and chromium(II1) oxide. Representative dose-effect curves for cyanide and cadmium are presented as examples of results obtained with the E. coli electrode. The dose in these experiments is the concentration of substance present in the buffer system in which the electrode tip is immersed. The effect being measured is the percent inhibition of COPproduction by the immobilized cells due to exposure to a particular concentration of some toxicant. Each point on the doseeffect curves is the average of 3 or 4 measurements. Each measurement is made with a different bacterial population and is independent of other measurements for the same concentration of toxicant. The dose-effect curves for cyanide are shown in Figures 3 and 4. Cyanide exhibited the widest concentration range over which toxic responses could be measured. Measurable inhibition occurs with concentrations as low 20 pM, and 970
Environ. Sci. Technol., Vol. 18, No. 12, 1984
80.0 c
60.0 U
zt 40.0
20.0
00 00
08
04
CCN-3 (mM)
Flgure 3. Respiratory inhibitkm of immobRized E.coli by concentrations of cyanide less than 1 mM. Error bars are 45.0% I or twice the average SEM for the experimental points shown. 100 0
I
I
I
i
1
1
60.0 U
z
Lk
I I'
40,Y 011
I
20 0
001 00
I
1
I
60.0
120 0
i I
CCN-3 (mM)
Figure 4. Respiratory inhibitkm of knmobked E . cdi by concentrations of cyanide greater than 1 mM. Error bars are 450% I or twice the average SEM for the experimental points shown.
a concentration of 142 mM is necessary for the maximum inhibition of nearly 100%. The cyanide dose-effect relationship is described by two curves. An effect plateau, a region where inhibition does not change with increasing dose, is reached between 700 pM and 7 mM, where the percent inhibition remains at approximately 45%. The rapid, dramatic change in the rate of C02 production by the immobilized E. coli is a typical response to exposure to a substance that directly inhibits respiration. Cyanide is a potent and rapidly acting chemical asphyxiant that prevents tissue utilization of O2by inhibiting cytochrome oxidase (18). The dose-effect curve obtained for cadmium is shown in Figure 5. The percent respiratory inhibition increases rapidly with increasing concentration, approaching a plateau of 84% inhibition at 180 pM. The range of response
100.0
9
Table I. Comparison between Rainbow Trout LC50Values, Microtox ECroValues, and E . coli Electrode 40% I Values for Various Toxic Substances
substance phenol HCN Cd2+ AsO< Pb2+ cu2+
rainbow trout LCm, ppma 5.0 0.7
Microtox E. coli electrode 40% I, ppmC EC5o9 PPmb 28.0 1209.0 4.0 20.0
3.5
0.006 13.3 1.2
26.0 0.6
0.3 1.5 0.13
0.4
0.8
1.1
OLCs0 values are obtained from 48- or 96-h exposures. References are as follows: phenol (20),HCN (21),Cd2+(22),As02- (23), Pb2+(24),Cu2+(25). bEC50values were obtained from 15-min exposures at the 15 "C test temperature. All values were obtained in the Microtox reagent designed for making these measurements. Values obtained from ref 26. e40% I values were obtained by interpolation of dose-effect curves.
0.0 0.0
80 0 L C d 3 (UM)
160 0
Flgure 5. Respiratory lnhlbition of lmmoblllzed E . co// by cadmium. Error bars are f5.0% I which equals the average SEM for the experimental points shown.
is large; the percent inhibition of 1.5 pM cadmium is 14%. The toxic action of cadmium on E. coli is attributed to competition with magnesium in cellular processes (19). In solutions containing low concentrations of magnesium, the addition of cadmium causes decreased C02production. In the presence of high concentrations of magnesium, the toxicity of cadmium is markedly lowered. The dose-effect curves for phenol, arsenite, p-(chloromercuri)benzoate, copper, and lead exhibit characteristics similar to those shown for cyanide and cadmium. All of the dose-effect curves show that E. coli C02 production is progressively inhibited as the concentration of toxicant increases. In most cases after an initial rapid increase in percent inhibition with increasing toxicant concentration, a plateau is reached where the percent inhibition changes only slightly as toxicant concentration increases. The measurable range of toxicant concentration varies with the substance being tested. Minimum concentrations of these toxicants causing 40% inhibition of bacterial respiration are reported in Table I. E. coli electrode response to solutions containing arsenite, p-(chloromercuri)benzoate,cyanide, cadmium, and phenol is rapid. Moreover, respiratory inhibition can be measured over wide ranges of toxicant concentrations. Because lead and copper ions exert their toxic effect at the membrane level (19) rather than directly on cellular respiration or metabolism, E. coli electrode response to these materials is slow. Some time is required after addition of lead or copper for a measurable change in immobilized cell respiration to occur. At low concentrations, the kinetics of toxic action become so slow that a final steady-state level of C02 production is difficult to estimate. Because of the slow response of the E. coli electrode to low concentrations of lead and copper, it was difficult to measure percent inhibitions of less than 35%. In addition to the s u w c e s discussed previously, the E. coli electrode wa used to study the acute toxicity of nitrite, hypochlorit&'?Ag, Zn, and 2,4-dichlorophenoxyacetic acid. Nitrite and Zn were not toxic to the immobilized E. coli. Even at high concentrations, no significant inhibition of cellular respiration was observed. Silver was essentially insoluble in the buffer systems used.
The acute toxicity of the commercial wood treatment formulation CCA was also examined. Experiments were conducted with this substance exactly as with the single compounds. The dose-effect curve for CCA is similar to those obtained for the calibration substances. The maximum percent inhibition is 75% I at a dose of 0.0042 vol 5% CCA in glucose-enriched PIPES. Thirty-five percent inhibition was measured with a dose of 1.6 X vol % CCA in glucose-enriched PIPES. The E. coli electrode response to additioh of growth medium was also measured. An aliquot of growth medium was added to the glucose-enrichedbuffers in which the E. coli electrode tip was immersed. No immediate effect was observed, since the glucose concentration in the buffer is already in excess of the amount necessary for maximal bacterial respiration. Under these conditions increased C02 production could only be the result of an increase in the E. coli population which would become apparent only after approximately 17 min, the length of time required for E. coli replication. The longevity of the E. coli electrode was also studied. E. coli electrode response to glucose-enriched phosphate buffer was measured every day for a period of 41 days. The electrode was stored for the 6-week period in plain 0.05 M phosphate buffer at 5 "C. The level of apparent C02 production remains fairly constant for about 1week. Then the level of C02production declines steadily, leveling off after 21/2 weeks, where it remains stable for another 4 weeks. At the end of this period, the bacterial respiration produces a COPconcentration at the electrode surface of 0.4 mM. The lower end of the linear range for the C02 electrode is 0.1 mM; so even after 6 weeks, the electrode is still functional. Longevities of bacterial electrodes reported in the literature vary from 4 days to 40 days ( 3 , 4 , 13, 14).
Discussion Another microbial test that has been used to evaluate acute toxicity is the Beckman Microtox bioassay. Both the bacterial electrode system and the Microtox system are based on monitoring inhibition of total metabolic processes. The methods are thus readily comparable. The Microtox system uses a luminescent bacteria that closely resembles Photo bacterium phosphoreum. Light production in these microorganisms is an expression of the total metabolic process of the cells and is a reflection of cell viability (27). Inhibition of any one of the many enzymes involved in overall energy metabolism may cause a change in the rate of light production. The Microtox system has been compared to 24-96-h fish bioassays for acute toxicity Envlron. Scl. Technol., Vol. 18,
No. 12, 1984 971
(28). Microtox data are generally presented in the form of an EC, value (the concentration of toxicant causing a 50% reduction in light production) obtained for a specified length of time, usually 5, 15, or 30 min. Fish bioassays are widely used to evaluate acute toxicity of pollutants. Many different species of fish have been used for acute toxicity studies; one of the most common species is rainbow trout (Salmogairdneri). The trout 96-h LC, (the median lethal concentration) has become one of the most generally employed means of evaluating and setting standards for toxic materials introduced into aquatic systems. Results obtained with the E. coli electrode for acute toxicity are compared to Table I with other methods for measuring the acute toxicity of pollutants in water. Inhibition levels of 40% were chosen for comparison with other methods. Table I contains toxicity data comparing rainbow trout 96-h LC50 values, E. coli electrode 40% I values, and Microtox 15-min EC50 values. The microbial test values measure the intensity of effect from a range of doses on populations of prokaryotic cells. The two microbial toxicity assays yield comparable results for most substances, the largest discrepancy between the two techniques being the Microtox EC50 of 28 ppm of phenol and the E. coli electrode 40% I of 1207 ppm. For cadmium, copper, lead, arsenite, and cyanide, the toxicity measurements obtained by the two microbial tests are comparable. Data analysis for the two microbial systems is quite similar. In both cases percent inhibitions of metabolic functions are measured, light production in the Microtox system and COz production in the E. coli electrode system. The trout LC50 values are less than either the Microtox ECS0or the E. coli electrode 40% I values for most of the substances compared in Table I. In summary, the E. coli electrode system for measuring the acute toxicity of substances in water is feasible to build and to use, costs of construction and use are minimal, and electrode application in the laboratory is easy to implement. Electrode responses relate the toxicity of substances in water to bacterial respiration rates. The E. coli electrode system yields dose-effect relationships with acceptable precision for a toxicity test ( < E % relative standard deviation for each measurement). However, for the 12 substances evaluated by using the E. coli electrode, only seven yielded dose-effect relationships which have provided useful toxicological information. Interferences and lack of sensitivity for slow-acting toxicants at low concentrations were the main limitations on the applicability of the method. Perhaps most importantly, our studies of the E. coli electrode system for measuring acute toxicity have provided a useful foundation to guide the development of other instrumental toxicity bioassays. Registry No. Cd, 7440-43-9;Pb, 7439-92-1;Cu, 7440-50-8; HCN, 57-12-5;As03, 15502-74-6;Zn, 7440-66-6; Ag, 7440-22-4; chlorine, 7782-50-5;ammonia, 7664-41-7;nitrite, 14797-65-0; phenol, 108-95-2;2,4-dichlorophenoxyacetic acid, 94-75-7;p (chloromercuri)benzoate, 59-85-8.
Literature Cited (1) Rechnitiz, G. A.; Reichel, T. L.; Kobos, R. K.; Meyerhoff,
M.E.Science (Washington, D.C.) 1978,199,440-441.
972
Environ. Scl. Technol., Vol. 18, No. 12, 1984
(2) Kobos, R. K.; Rechnitz, G. A. Anal. Lett. 1977,10,751-758. (3) Walters, R.; Moriarity, B. E.;Buck, R. P.Anal. Chem. 1980, 52, 1680-1684. (4) Arnold, M. A.; Rechnitz, G. A. Anal. Chem. 1980, 52, 1170-1 174. (5) Jensen, M. A,; Rechnitz, G. A. Anal. Chim. Acta 1978,101, 125-130. (6) Kobos, R. K.; Rice, D. J.; Flourney, D. S. Anal. Chem. 1979, 51 1122-1125. (7) Hikumo, M.; Kubo, T.; Yasudo, T.; Karube, I.; Suzuki, S. Anal. Chem. 1980,52,1020-1024. (8) Karube, I.; Matsunaga, T.; Nakahara, T.; Suzuki, S.; Kada, T. Anal. Chem. 1981,53, 1024-1026. (9) Karube, I.; Nakahara, T.; Matsunaga, T.; Suzuki, S. Anal. Chem. 1982,54,1725-1727. (10) Simpson, D. L.; Kobos, R. K. Anal. Chem. 1983, 55, 1974-1977. (11) Orion Research Incorporated “Instruction Manual Carbon Dioxide Electrode Model 95-02”;Orion Research Incorporated: Cambridge, MA, 1978. (12) Colwell, R. R.; et al. “Marine and Estuarine Microbiology Laboratory Manual”; University Park Press: Baltimore, MD, 1975;pp 25-26. (13) Rechnitz, G. A.; Kobos, R. K.; Reichel, S. J.; Gebaur, C. R. Anal. Chim. Acta 1977,94,357-365. (14) Arnold, M. A. Am. Lab. (Fairfield,Conn.) 1983,15,34-40. (15) Good, N. E.;et al. Biochemistry 1966,58 467-477. (16) Laidler, K. T. “Chemical Kinetics”; McGraw-Hill: Hightstown, NJ, 1965;pp 14-15. (17) Kobos, R. K.; Park, S. J.; Meyerhoff, M. E. Anal. Chem. 1982,54,1976-1984. (18) Lehninger, A. L. “Biochemistry”,2nd ed.; Worth Publishers: New York, 1975;p 497. (19) Block, S. S. ”Disinfection, Sterilization, and Preservation”, 2nd ed.; Lea & Febiger: Philadelphia, PA, 1977;pp 408-409. (20) Office of Water Regulations and Standards “Ambient Water Quality Criteria for Phenol”; US. Environmental Protection Agency: Washington, DC, 1980;EPA-44015-80-066, pp 17-18. (21) Office of Water Regulations and Standards “Ambient Water Quality Criteria for Cyanide”; U S . Environmental Protection Agency: Washington, DC, 1970;EPA-44015-80-037, p 21. (22) Office of Water Regulations and Standards “Ambient Water Quality Criteria for Cadmium”; U.S. Environmental Protection Agency: Washington, DC, 1980;EPA-44015-80-025, pp 46-48. (23) Office of Water Regulations and Standards “Ambient Water Quality Criteria for Arsenic”; US.Environmental Protection Agency: Washington, DC, 1980;EPA-44015-80-021, p 13. (24) Davies, P. H. Water Res. 1976,10, 199-210. (25) Schneider, B. A., Ed. “Toxicology Handbook on Mammalian and Aquatic Data: Toxicology Data”; U.S. Environmental Protection Agency: Beltsville, MD, 1979;Book I, p 23. (26) Beckman Instruments “Microtox Application Notes”; Beckman Instruments, Inc.: Carlsbad, CA, 1981;No. M102. (27) Beckman Instruments “Microtox Model 2055 Toxicity Analyzer System”; Beckman Instruments, Inc.: Carlsbad, CA, 1978. (28) Bulich, A. A.; Greene, M. W.; Isenberg, D. L. ASTM Spec. Tech. Publ. 1981, STP 737,338-347. Received for review February 2,1984, Accepted August 6,1984. This work was supported in part by the State-Sponsored Organized Research Program of the Colorado Commission on Higher Education.
