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Biotechnol. Prog. 1990, 6, 76-81
Genetic Engineering Approach to Toxic Waste Management: Case Study for Organophosphate Waste Treatment Steven J. Coppella,t*sNeslihan DelaCrtqt Gregory F. Payne,*pt9S Burton M. Pogell,s9s Marilyn K. Speedie,$Jeffrey S. Karns,l Edward M. Sybert,"and Michael A. Connorll Chemical and Biochemical Engineering, University of Maryland Baltimore County, Baltimore, Maryland 21228, Maryland Biotechnology Institute and Biomedicinal Chemistry, University of Maryland at Baltimore, Baltimore, Maryland 21201, Beltsville Agricultural Research Center, United States Department of Agriculture, Beltsville, Maryland 20705, and Engineering Research Center, University of Maryland, College Park, Maryland 20742
Currently, there has been limited use of genetic engineering for waste treatment. In this work, we are developing a procedure for the in situ treatment of toxic organophosphate wastes using the enzyme parathion hydrolase. Since this strategy is based on the use of an enzyme and not viable microorganisms, recombinant DNA technology could be used without the problems associated with releasing genetically altered microorganisms into the environment. The gene coding for parathion hydrolase was cloned into a Streptomyces liuidans, and this transformed bacterium was observed to express and excrete this enzyme. Subsequently, fermentation conditions were developed to enhance enzyme production, and this fermentation was scaled-up to the pilot scale. T h e cell-free culture fluid (i.e., a nonpurified enzyme solution) was observed to be capable of effectively hydrolyzing organophosphate compounds under laboratory and simulated in situ conditions.
Introduction Despite its potential impact, genetic engineering has currently had little effect on how toxic wastes are treated. Here, we report on a strategy in which a recombinant microorganism is used to overproduce an enzyme that can then be used to detoxify organophosphate-containing wastes. When a waste is treated with an enzyme(s), rather than a microbe, problems associated with the release of genetically engineered organisms into the environment are eliminated. Thus, enzyme-basedtreatment strategies are environmentally more acceptable and may therefore be the first application of genetic engineering to solve in situ waste problems. However, there are several technical prerequisites that must be met for an enzyme-based treatment approach to be appropriate. First, since an enzyme can catalyze only a single reaction, or a small number of closely related reactions, it is essential that the enzyme-catalyzed step be useful. If the product of the reaction is less toxic, or more readily degraded than the reactant, an enzymebased strategy could be effective. Second, since enzymatic reactions often require cosubstrates, these cosubstrates must be readily available. Water- or oxygenrequiring enzymes (e.g., hydrolases or oxygenases, respectively) are the most obvious candidates for waste applications, while enzymes that require cell-generated cosubstrates (e.g., ATP or NAD(P)H) would be less practical. A third prerequisite is that the enzyme must have an appropriate catalytic activity and affinity for the reactant as compared to alternative treatment methods. Because of their high substrate affinities, enzymes are typically well suited for treating dilute wastes. Finally, Chemical and Biochemical Engineering. Maryland Biotechnology Institute. 8 Biomedicinal Chemistry, University of Maryland a t Baltimore. Beltsville Agricultural Research Center. '1 Engineering Research Center, University of Maryland, College Park.
the enzyme must have sufficient stability to be effective under the required operating conditions. Enzymes are often labile to extremes in pH, temperatures above ambient, and organic solvents. In this work, we are examining the potential of an enzyme-based strategy for the treatment of organophosphate wastes. Specifically, we are examining the use of an enzyme to detoxify an aqueous waste stream containing the insecticide coumaphos. This waste stream is generated from a tick eradication program in which cattle are dipped in a solution containing coumaphos. Vats containing this solution are recharged annually, and the waste solutions must be detoxified. Figure 1 shows the strategy that was developed in previous work (1). The coumaphos was first hydrolyzed in a biological step, and the chlorferon product was later fragmented by UV/ ozonation. The resulting fragments were then susceptible to mineralization by endogenous soil organisms. Biological hydrolysis is required since coumaphos is not fragmented by UV/ozonation. In addition, it appears that coumaphos can be dehalogenated under normal dip-vat conditions to form potasan. It would be desirable if potasan could also be hydrolyzed in this biological step. With respect to the biological hydrolysis, Pseudomonas (2, 3), Flavobacteria ( 4 ) , and other uncharacterized bacteria (5)capable of hydrolyzing organophosphates have been isolated from the environment. However, the uncontained use of viable coumaphos-hydrolyzing organisms is inappropriate because such populations could spread into and detoxify coumaphos in the vats, which are being used for insect control (1). Thus, an enzyme-based strategy was considered for this application. In this paper, we will summarize our results on the production and use of the organophosphate-degrading enzyme, parathion hydrolase.
Enzyme Biosynthesis Initial efforts to produce the parathion hydrolase enzyme using the above-mentioned bacterial isolates were plagued
8756-7938/90/3006-0076$02.50/00 1990 American Chemical Society and American Institute of Chemical Engineers
Biotechnol. Prog., 1990,Vol. 6,NO. 1
77 UV-Ozonation
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Figure 1. Strategy for treating coumaphos-containing waste cattle dip. Biological hydrolysis of this organophosphate is followed by UV ozonolysis. Also, coumaphos dechlorination has been reported to occur in dip vats leading to the production of potasan (25). (Adapted from Kearney et al. (I).) by problems. In addition to low levels of production, the enzyme produced by these isolates was either retained within the cells or associated with the cell membrane (5-9). Thus, if enzyme was to be produced using these isolates, cell disruption, or membrane solubilization steps would be required for enzyme recovery. Because of these problems, and because the opd gene coding for this enzyme ( 9 , 10, 11, 12) is known, genetic engineering approaches have been considered for the production of parathion hydrolase (7, 8). I t should be noted that the required genetic information (Le., the opd gene in this case) must be available before recombinant technology can be applied. The purpose for using genetic engineering is to clone the opd gene into a host cell that is easily cultured and is capable of producing large amounts of the enzyme. Cloning for Organophosphate Degradation. Currently, Escherichia coli is the most commonly used bacterial host for genetic engineering applications. However, in this work we chose to use the bacterium S t r e p tomyces liuidans as the recombinant host because Streptomyces are able to excrete foreign proteins outside the cell. Excretion is a major advantage for this application because it eliminates the need for cell disruption operations and therefore greatly simplifies any production process (for review of the use of Streptomyces for genetic engineering applications, see Crawford (13)). The major drawback to using Streptomyces as a host is that, at the present time, production of foreign proteins by Streptomyces is generally less than production by E. coli. For transfer of the opd gene, the Streptomyces plasmid pIJ702 was used (14). As shown in Figure 2, Steiert et al. (15) cloned the opd gene into this plasmid and transformed the Streptomyces lividans host with the modified plasmid, pRYE1. Transformed cells were observed to produce and excrete the parathion hydrolase enzyme (15). Parathion hydrolase production by this transformant was also observed to be stable-even in the absence
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Figure 2. Schematic of the genetics in this project. The parathion hydrolysis gene (opd) was cloned into the pIJ702 plasmid to produce a new plasmid, pRYE1. Streptomyces liuidans was transformed with thispRYEl plasmid. (Adaptedfrom Steiert et al. (15).) of selective pressure (16). This agrees with other studies that have indicated that the expression of foreign proteins with this host-vector combination is stable (17,18). Optimization of Parathion Hydrolase Production. Subsequent studies with this recombinant focused on the development of fermentation conditions that are optimal for parathion hydrolase production. In this work, parathion hydrolase is measured in terms of activity units where 1 unit of activity is capable of hydrolyzing 1pmol of parathion/min at room temperature and at pH 8.5. We have observed that 1 unit of activity is equivalent to about 1 pg of enzyme. Initial fermentation studies (15) in which this recombinant was cultured in 500-mL (150-mL liquid volume) flasks indicated that enzyme production was low, with levels reaching only about 0.6 unit/mL. To optimize enzyme production, three approaches were taken: (i) culture the cells in fermentors instead of flasks; (ii) improve the fermentation medium; and (iii) develop improved fermentor operating strategies (i.e., nutrient feeding vs batch
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0
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20
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Hours Figure 3. Pilot scale production of parathion hydrolase. Cells were cultured in a 30-L (25-L liquid volume) fermentor. Aeration and agitation were set at 25 L of air/min and 200 rpm, respectively. Tryptone and glucose, the only two nutrients used in the medium, were fed intermittantly throughout the experiment. (Further details of the fermentation conditions can be found in Payne
et al. (16).)
cultivation). These changes led to a 25-fold increase in enzyme production (16). To obtain larger quantities of enzyme for treatment, this fermentation process will need to be scaled-up from the 1-L bench scale. Initial efforts to produce parathion hydrolase on a pilot scale were conducted in a 30-L (25-L liquid volume) fermentor. Figure 3 shows that, in this pilot scale fermentor, 11units/mL of extracellular enzyme was obtained after 40 h. This is comparable to the activity of enzyme produced over the same period in our bench scale systems. A final point that is very important for the application of this enzyme should be noted. Currently, we envision using this enzyme by adding the cell-free culture fluid directly to a waste stream. Thus, we do not propose to purify this enzyme prior to use. However, a possible problem with this strategy occurs if the culture produces extracellular proteases. If significant protease activities were present in our culture filtrates, then these fiitrates would need to be treated to either remove or inactivate the proteases or to partially purify the parathion hydrolase enzyme. Such steps appear to be unnecessary for our system since protease activity could not be detected in the extracellular culture fluid.
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Enzyme-Based Treatment
ificity (5, 20, 21) and are therefore capable of hydrolyzing many related organophosphate compounds. Kinetics. Typically enzyme-catalyzed reactions follow Michaelis-Menten or saturation kinetics, which are described by the equation
The use of enzymes to hydrolyze organophosphates has been considered by other workers. Enzymes responsible for catalyzing this hydrolytic step (referred to as either organophosphorous phosphotriesterase or parathion hydrolase) have been identified and characterized (5, 19-21). However, for enzyme-based treatment strategies to be effective, the enzyme must be capable of effectively catalyzing the detoxification reaction and must be stable enough to remain active for a sufficient period of time to achieve detoxification. In this work, we examined the kinetics and the stability of a cell-free enzyme solution obtained directly from the fermentation. These tests were conducted with use of parathion as the organophosphate reactant. It should be noted that many parathion hydrolase enzymes studied have a broad substrate spec-
In this equation, V and V,,, are the reaction and maximum reaction "velocities" in units per milligram of enzyme, S is the reactant concentration in millimoles per liter, and K , is the half-saturation constant. As can be seen from Figure 4, parathion hydrolysis by this unpurified, cell-free culture fluid appears to follow saturation kinetics. The K , determined from this study is approximately 0.02 mmol/L. Other researchers have observed K,'s for parathion hydrolase to be of this same order of magnitude although considerable variabilities exist (5, 19-22). These discrepancies could be due to the existence of several, different parathion hydrolase enzymes
Biotechnoi. Prog., 1990,Vol. 6, No. 1
(5); the existence of different multimeric forms of the same enzyme (21);the fact that small additions of hydrophobic compounds (e.g., low molecular weight organics or detergents), which can significantly affect the behavior of the enzyme, are often added to reaction mixtures in these kinetic studies (15, 20); and the low solubility of typical organophosphate substrates that limit accurate quantitative study of intrinsic reaction kinetics. The low K , observed in this study indicates that the crude enzyme preparations used in this study retain significant affinities for organophosphate reactants. Although we have not compared the kinetics of enzyme hydrolysis with the alternative method of hydrolyzing organophosphates using alkaline conditions, Munnecke (20) reported that enzyme-catalyzed parathion hydrolysis was orders of magnitude more rapid than chemical hydrolysis with 0.1 N NaOH. However, the performance of any enzyme-based treatment system will be strongly dependent on the amount of enzyme used. Thus, for a complete comparison of these technologies, the cost of enzyme production and the amount of enzyme required for treatment must be determined. Temperature Dependence of Hydrolysis Reaction. Because enzymes are rather labile, they can be easily inactivated at high temperatures. However, at lower temperatures, the rate of enzyme-catalyzed reactions follows a simple Arrhenius temperature dependence. Arrhenius dependence is described quantitatively by
where Eactis the activation energy of the enzyme-catalyzed reaction, R is the gas constant, T is temperature, and A is a constant. Figure 5 shows that the enzymecatalyzed hydrolysis of parathion follows this Arrhenius dependence with an activation energy of 3.6 kcal/mol. Brown (21) also reported a low activation energy for organophosphate hydrolysis by a parathion hydrolase enzyme. Thermal Stability of the Hydrolase Enzyme. Although increasing temperatures lead to enhanced organophosphate degradation, it is important to recognize that the parathion hydrolase enzyme is labile and undergoes a temperature-dependent inactivation. This inactivation is often considered to be a first-order process in which the active enzyme (E) reacts to form an inactive enzyme (E*) by the reaction
E+E* (3) In a batch process, the rate of disappearance of enzyme activity is described by the equation d[E]/dt = -k[E] (4) where k is the first-order rate constant for inactivation. In the biological literature, enzyme inactivation is reported not in terms of the first-order rate constant but rather in terms of the enzyme's half-life. The half-life (tl,& is related to the first-order rate constant by
t l l z = (In 2)/k (5) The first-order rate constant for enzyme inactivation is also observed to follow Arrhenius temperature dependence such that
k = k,e-pa 'IRT (6) Eact I in eq 6 is the activation energy for enzyme inactivation and is different from the previously described activation energy for enzyme-catalyzed organophosphate hydrolysis. Figure 6 shows that parathion hydrolase inactivation follows this Arrhenius dependency with an acti-
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Figure 5. Arrhenius plot of enzyme-catalyzedparathion hydrolysis. Data from separate experiments are indicated by different symbols. The activation energy for enzyme-catalyzedhydrolysis was calculated from this graph to be 3.6 kcal/mol. L. . . . . .
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vation energy of 23 kcal/mol. By comparison of these two activation energies, it can be seen that although the enzyme-catalyzed reaction rate has only a weak dependence on temperature, the enzyme stability is considerably reduced at higher temperatures. Since treatment operations are often conducted a t ambient temperatures, the effectiveness of an enzyme-based treatment strategy may be strongly dependent upon the ambient temperature. Simulation of Enzyme Stability. To predict the longterm performance of a reaction system that employs parathion hydrolase, eqs 2, 4, and 6 were used to simulate the enzyme performance over time (23,241. For this simulation, the volumetric rate of organophosphate hydrolysis is given by ?. = VmaX[El (7) where [E] represents the concentration of the active form of the parathion hydrolase. These simulations are shown in Figure 7 . The results are normalized to an initial rate that would be observed if the enzyme were allowed to react at 25 "C. In general, Figure 7 shows that the volumetric reaction rate is reduced as the enzyme is inactivated. However, at 4 "C this inactivation would be small with over 90% of the initial activity being retained even after 80 h. At the highest temperature (51 "C), Figure 7 shows that this enzyme is rapidly inactivated with less than 10% of its initial activity remaining after only 7 h. Although somewhat extreme, this high-temperature case is signif-
80
Biotechnol. Prog., 1990, Vol. 6,No. 1
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Figure 7. Simulation of enzyme activity over time for three temperatures. These simulations are normalized to an initial activity that would be observed at 25 O C . At the high temperatures, the enzyme activity rapidly decays due to thermal inactivation of the enzyme. icant since our initial application of this enzyme is for the detoxification of a coumaphos waste, which results from an insect control program on the Texas-Mexico border (1,251. Thus, for improved performance, it may be desirable to use this enzyme a t cooler times during the evening. Finally, it should be mentioned that this simulation only accounts for thermal effects on the enzyme. The enzyme activity and stability may also be affected by additional physical, chemical, and biological conditions that exist at each specific treatment site. For liquid-phase waste streams, it may be desirable to immobilize the enzyme on, or within, a solid support. Immobilization is advantageous because immobilized enzymes can be retained within a treatment vessel, and thus, continuous-flow treatment strategies can be considered. Another possible benefit is that enzyme activity and stability may be improved by immobilization. This is a particularly interesting possibility for the parathion hydrolase enzyme studied here. In nature, this enzyme is associated with the hydrophobic environment of the cell membrane. Thus, by immobilizing the enzyme in a hydrophobic environment, it may be possible to better simulate the natural environment of this enzyme with the possible result that enzymatic properties are improved. Application of the Parathion Hydrolase Enzyme to Coumaphos Waste. To test the enzyme-based treatment strategy under conditions that closely simulate actual conditions, 25 p L of a cell-free enzyme solution was added to 5 mL of a waste cattle-dip solution. This waste was obtained from cattle dip that had actually been used in the field. As shown in Figure 8, the coumaphos in this waste cattle dip was hydrolyzed over the 60-h period with nearly stoichiometric quantities of chlorferon being produced. Shelton and Karns (25)observed that, under conditions observed in the dip vats, coumaphos can be reductively dechlorinated to potasan. Figure 8 also shows that potasan that is present in the waste dip solution is degraded with nearly stoichiometric quantities of its hydrolysis product, 4-methylumbelliferone, being produced. These results are very encouraging with respect to the use of this enzymebased system under field conditions. It is also interesting to note that over the course of the experiment the rate of hydrolysis of coumaphos was reduced, and this observed reduction is similar to those predicted in Figure 7 . Figure 7 suggests that, after 24 h at 25 O C , the enzyme activity would have been reduced to 60% of its initial value. The enzymatic activity observed between 8 and 24 h was approximately 65% of the ini-
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Figure 8. Waste cattle-dip detoxification by cell-free parathion hydrolase. Coumaphos (closed triangles) and the dechlorinated potasan (closed circles) were shown to disappear,while their hydrolysis products chlorferon (open triangles) and the dechlorinated 4-methylumbelliferone(open circles) were shown to appear during treatment. Treatment was conducted by adding 25 pL of cell-free enzyme to 5 mL of the waste cattle dip and mixing at 28 “C. (The analytical techniques are described by Shelton and Karns (25).) tial activity. After 57 h, Figure 7 suggests that only 20% of the initial enzyme activity would be retained. This compares to the rate observed between 24 and 57 h, which was 15% of the initial value. These small discrepancies between observed and predicted inactivation could be due to differences in temperature (i.e., the dip solution was treated at 28 “ C ) or the difficulties in working with the slightly soluble coumaphos substrate. Nevertheless, the predicted inactivation of the parathion hydrolase enzyme is in good agreement with the observations in Figure 8. Further work will be required to better quantify the various phenomena involved in enzyme inactivation.
Conclusions Until the release of genetically engineered organisms is accepted as being safe, there are few possible strategies for using genetic engineering technology to treat toxic wastes. The strategy considered in this work is the use of an enzyme, rather than viable microbes, for the in situ detoxification of organophosphate-containing wastes. In this paper, we have summarized our work on the production of parathion hydrolase by a recombinant Streptomyces liuidans. This host cell is particularly useful since it excretes the enzyme, and excretion greatly facilitates enzyme recovery by eliminating the need for cell disruption operation. After the gene coding for the parathion hydrolase was cloned, enzyme production by this recombinant was improved with use of traditional industrial microbiological techniques, and this process was scaled to the pilot level. In addition to biological production, the properties and performance of the enzyme are described. Standard kinetic equations were used to describe the activity and thermal instability of the enzyme. With these equations, it was shown that long-term high-temperature applications may be limited by the enzyme’s instability. Our results also demonstrate that the parathion hydrolase enzyme is able to hydrolyze the organophosphate compounds, coumaphos and potasan, under conditions that closely simulate those expected in the field. A final, nontechnical, point is that because of the diverse range of skills required, the success of this, or related
Biotechnol. Prog., 1990, Vol. 6, No. 1
projects, hinges on strong interdisciplinary collaborations. We believe such collaborations are critical if biotechnology is to be used to solve real waste problems.
Acknowledgments Financial support for this work was provided by the University of Maryland’s Biotechnology Institute. Also, we thank E. R. Squibb and Sons and New Brunswick Scientific for their equipment and materials support. Literature Cited (1) Kearney, P. C.; Karns, J. S.; Muldoon, M. T.; Ruth, J. M.
J . Agric. Food Chem. 1986,34,702. (2) Munnecke, D. M.; Hsieh, D. P. H. Appl. Microbiol. 1974, 28,212. (3) Daughton, C. G.; Hsieh, D. P. H. Appl. Enuiron. Microbiol. 1977, 34, 175. (4) Sethunathan, N.; Yoshida, T. Can. J. Microbiol. 1973, 19, 873. (5) Mulbry, W. W.; Karns, J. S. Appl. Enuiron. Microbiol. 1989, 55, 289. (6) Munnecke, D. M.; Fischer, H. F. Eur. J. Appl. Microbiol. 1979,8, 103. (7) Serdar, C. M.; Gibson, D. T. Biotechnology 1985, 3, 567. (8) Kearney, P. C.; Karns, J. S.; Mulbry, W. W. Engineering soil microorganisms for pesticide degradation. In Pesticide Science and Biotechnology; Greenhalgh, R., Roberts, T. R., Eds.; Blackwell Scientific: London, 1985, pp 591-596. (9) McDaniel, C. S.; Harper, L. L.; Wild, J. R. J . Bacteriol. 1988, 170, 2306.
81
(10) Mulbry, W. W.; Karns, J. S.; Kearney, P. C.; Nelson, J. D.; McDaniel, C. S.; Wild, J. R. Appl. Enuiron. Microbiol. 1986,51, 926. (11) Chaudhry, G. R.; Ali, A. N.; Wheeler, W. B. Appl. Enuiron. Microbiol. 1988, 54, 288. (12) Harper, L. L.; McDaniel, C. S.; Miller, C. E.; Wild, J. R. Appl. Enuiron. Microbiol. 1988, 54, 2586. (13) Crawford, D. L. Biotech. Adu. 1988, 6, 183. (14) Katz, E.; Thompson, C. J.; Hopwood, D. A. J. Gen. Microbiol. 1983, 129, 2703. (15) Steiert, J. S.; Pogell, B. M.; Speedie, M. K.; Laredo, J. Biotechnology 1989, 7, 65. (16) Payne, G. F.; DelaCruz, N.; Coppella, S. J. Submitted for publication, 1989. (17) Ghangas, G. S.; Wilson, D. B. Appl. Enuiron. Microbiol. .. 1987, 53,-1470. (18) Bertrand, J.-L.; Morosoli, R.; Shareck, F.; Kluepfel, D. Biotechnol. Bioeng. 1989, 33, 791. (19) Munnecke, D. M.; Hsieh, D. H. Appl. Enuiron. Microbiol. 1976, 31, 63. (20) Munnecke, D. M. Appl. Enuiron. Microbiol. 1976, 32, 7. (21) Brown, K. A. Soil Biol. Biochem. 1980,12, 105. (22) Munnecke, D. M. Appl. Enuiron. Microbiol. 1977,33,503. (23) Yang, S.-T.; Okos, M. R. Biotechnol. Bioeng. 1989, 33, 873. (24) Peterson, R. S.; Hill, C. G.; Amundson, C. H. Biotechnol. Bioeng. 1989, 34, 429. (25) Shelton, D. R.; Karns, J. S. J. Agric. Food Chem. 1988, 36, 831.
Accepted January 2, 1990.
