stability of manipulated plasmid dna in aquatic environments

7
Stability of Manipulated Plasmid DNA in Aq uatic E nvi r on men ts Abdiel J. Alvarez, Gladys M. Yumet, Carlos L. Santiago, and Gary A. Toranzos* Department of Biology, P.O. Box 23360, College of Natural Sciences, University of Puerto Rico, Rio Piedras, Puerto Rico 00931-3360 Very little is known about the stability, behavior, and probability of perpetuation of genetically engineered DNA within a microbial community. The prevalence of genetically engineered microor- ganisms (and hence their DNA) in the environment as a result of deliberate or accidental release has been an increasingly controversial issue during the past few years. In the present study genetically manipulated plasmid DNA was seeded into different types of waters to determine its fate over time in the presence of naturally occurring bacterial populations. The seeded DNA was rapidly degraded in surface and marine waters. Up to 10 pg/mL was degraded in a few hours. The DNA was not degraded in water that had no detectable DNAse-producing microflora. No uptake of the DNA by the resident microflora was observed even where the DNA remained undegraded up to 7 days. We can thus extrapolate that, in some aquatic environments, the genetically engineered DNA will be degraded in a very short time and therefore the concerns of uptake of novel genes by the resident microflora may be overstated. 0 7996 by John Wiley & Sons, Inc. I NTRODU CT I 0 N The release (and possible persistence) of genetically engineered microorganisms and the stability of geneti- cally engineered DNA (GED) in the environment has been an increasingly controversial issue in the past few years (Bentjen et al., 1989; Zeph and Stotzky, 1989). At the present time, very little is known about the stability, behavior, and probability of perpetuation of GED in the environment. There is, also little informa- tion on the probability that if the recombinant DNA molecule “survives” and is perpetuated, adverse con- sequences could be manifested within the resident mi- croflora. Information on how naked DNA, and more specifically GED, behaves in the environment is thus needed to assess any potential hazards associated with GED and to predict any environmental consequence * To whom correspondence should be addressed. that might arise from the accidental or deliberate re- lease of genetically engineered microorganisms (GEMS) into the environment (Tiedje et al., 1989). Attention has to be paid to the potential horizontal transfer of exogenous genetic determinants from engi- neered bacteria to the natural microbial population. When dealing with genetically engineered DNA the concerns about the uptake of the nucleic acids via transformation should be considered. Transformation as a means of gene transfer occurs in many bacterial genera (Lorenz and Wackernagel, 1994). The probabil- ity of the transfer of chimeric plasmid DNA by transfor- mation is dependent on the lysis of host cells, “sur- vival” of the DNA, and the likelihood that this GED would be taken up by, and survive, intracellular degra- dation by indigenous/endogenous DNAses of the recip- ients. Conditions required for transformation (tempera- ture and/or organic solvents) are very unlikely to be found in the environment (Cohen et al., 1972; Cosloy and Oishi, 1973). Even under these conditions, trans- Environmental Toxicology and Water Quality: An International Journal, Vol. 11 (1996) 129-135 0 1996 by John Wiley & Sons, Inc. CCC 1053-47251961021 29-07 129

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Stability of Manipulated Plasmid DNA in Aq uat ic E nvi r on men ts

Abdiel J. Alvarez, Gladys M. Yumet, Carlos L. Santiago, and Gary A. Toranzos*

Department of Biology, P.O. Box 23360, College of Natural Sciences, University of Puerto Rico, Rio Piedras, Puerto Rico 00931-3360

Very little is known about the stability, behavior, and probability of perpetuation of genetically engineered DNA within a microbial community. The prevalence of genetically engineered microor- ganisms (and hence their DNA) in the environment as a result of deliberate or accidental release has been an increasingly controversial issue during the past few years. In the present study genetically manipulated plasmid DNA was seeded into different types of waters to determine its fate over time in the presence of naturally occurring bacterial populations. The seeded DNA was rapidly degraded in surface and marine waters. Up to 10 pg/mL was degraded in a few hours. The DNA was not degraded in water that had no detectable DNAse-producing microflora. No uptake of the DNA by the resident microflora was observed even where the DNA remained undegraded up to 7 days. We can thus extrapolate that, in some aquatic environments, the genetically engineered DNA will be degraded in a very short time and therefore the concerns of uptake of novel genes by the resident microflora may be overstated. 0 7996 by John Wiley & Sons, Inc.

I NTRODU CT I 0 N

The release (and possible persistence) of genetically engineered microorganisms and the stability of geneti- cally engineered DNA (GED) in the environment has been an increasingly controversial issue in the past few years (Bentjen et al., 1989; Zeph and Stotzky, 1989). At the present time, very little is known about the stability, behavior, and probability of perpetuation of GED in the environment. There is, also little informa- tion on the probability that if the recombinant DNA molecule “survives” and is perpetuated, adverse con- sequences could be manifested within the resident mi- croflora. Information on how naked DNA, and more specifically GED, behaves in the environment is thus needed to assess any potential hazards associated with GED and to predict any environmental consequence

* To whom correspondence should be addressed.

that might arise from the accidental or deliberate re- lease of genetically engineered microorganisms (GEMS) into the environment (Tiedje et al., 1989).

Attention has to be paid to the potential horizontal transfer of exogenous genetic determinants from engi- neered bacteria to the natural microbial population. When dealing with genetically engineered DNA the concerns about the uptake of the nucleic acids via transformation should be considered. Transformation as a means of gene transfer occurs in many bacterial genera (Lorenz and Wackernagel, 1994). The probabil- ity of the transfer of chimeric plasmid DNA by transfor- mation is dependent on the lysis of host cells, “sur- vival” of the DNA, and the likelihood that this GED would be taken up by, and survive, intracellular degra- dation by indigenous/endogenous DNAses of the recip- ients. Conditions required for transformation (tempera- ture and/or organic solvents) are very unlikely to be found in the environment (Cohen et al., 1972; Cosloy and Oishi, 1973). Even under these conditions, trans-

Environmental Toxicology and Water Quality: An International Journal, Vol. 11 (1996) 129-135 0 1996 by John Wiley & Sons, Inc. CCC 1053-47251961021 29-07

129

130 ALVAREZ ET AL.

formation has a very low probability ( of occurring in noncompetent hosts. It thus would seem most im- probable that GED could be a potential hazard in this manner, but the possibility, however small, remains (Gealt et al., 1985; Saye et al., 1987).

Two factors that are likely to control the actual up- take of GED from GEMs in the environment are the competence of the recipient cells and the availability of naked DNA for uptake (Olson et al., 1990). Cell competence may be under internal and external con- trols, but in aquatic environments external induction of competence may be unlikely because of dilution of possible competence factors, the opposite of what seems to happen on soils due to the occurrence of protected microniches (Stotzky, 1989) or microbial bio- films (Lisle and Rose, 1994). Internal competence and possibly subsequent expression is mediated by a solu- ble, proteinaceous factor that can induce the generation of specific active sites along the cell membrane for DNA uptake (Istock, 1989). The second factor, GED in the environment, needs to be characterized in terms of availability, stability, and behavior. The sources of extracellular DNA in the environment include natural lysis of dead cells, the release of DNA from cells due to lytic phage infections (Redfield, 1988), and possibly accidental release of free DNA or microorganisms from research or industrial laboratories. A possible abun- dance of extracellular DNAses present in the environ- ment would argue that naked DNA would be rapidly degraded (Greaves and Wilson, 1970), although naked DNA has been detected in environments where DNAses were shown to be stable (DeFlaun et al., 1986).

In order to answer some of the questions on the behavior of genetically engineered DNA in the environ- ment, we seeded naked GED into four different non- sterile aquatic systems to measure its stability over time. Microcosms are a useful tool for risk assessment studies of the release of GEMs to the environment (Cairns and Pratt, 1986; Trevors, 1988), and in this case allowed for a thorough study of DNA and its interactions with the resident microflora. We estimated the length of time manipulated DNA remains unde- graded in the environment, demonstrated the presence of DNAses in the systems, and determined if naked DNA could transform the resident microflora of aquatic systems.

MATERIALS AND METHODS

Genetically Engineered DNA

As a model of genetically engineered DNA, we used the pWTAla5’ plasmid (a kind gift of Karen Sprague, University of Oregon). This is a 4.8 kilobase plasmid

that contains a 437 base pair (bp) insert (coding for tRNA, Ala) obtained from Bombix mori (Larson et al., 1983). Tbe plasmid was cloned in Escherichia coli DH1 and purified by conventional techniques (Sambrook et al., 1989).

Aquatic Systems

Four different types of water were used for this study, namely distilled, tap, marine, and river. The distilled and tap waters were obtained from the laboratory. The tap water was dechlorinated by the addition of sodium thiosulfate. The marine sample was obtained from a public bathing beach and the river water from a sewage- impacted site in Puerto Rico. These water samples were seeded with 10 or 2 pg/mL of pWTAla5’ and incubated at room temperature (23-25°C) in the dark to avoid algal growth. Aliquots were taken at increasing time intervals and stored at -20°C until analyzed.

Bacteriological Analyses

Samples from each water source were used for the determination of fecal coliforms, total coliforms, and total bacterial count. These analyses were done using the membrane filtration technique as outlined in Stan- dard Methods for the Examination of Water and Wastewater (American Public Health Association, 1985). The media used were as follows: mEndo, mFC, and plate count agar for total coliforms, fecal coliforms, and total plate count, respectively. All media were ob- tained from Difco Laboratories (Detroit, MI).

The plasmid pWTAlaS’, a derivative of pBR322, codes for resistance to ampicillin and tetracycline. These antibiotic resistance markers were used in order to determine if the resident microflora had been trans- formed or not by the genetically engineered DNA. Aliquots were filtered and membranes placed onto antibiotic-free Luria Bertani (LB) medium (Sambrook et al., 1989) as well as LB medium containing ampicillin (50 pg/mL) and tetracycline (15 pg/mL). Bacterial col- onies growing in both media were replica-plated onto nitrocellulose membranes and analyzed by the colony hybridization technique (Grunstein and Honess, 1975) in order to determine if the antibiotic resistance had been acquired by transformation and to differentiate GEMs from background resistant flora.

Nuclease Test

Nuclease (DNAse) presence was determined using DNA agar, as described by Lachica et al. (1971), with some modifications. To describe the process briefly; after an overnight growth of the bacteria on Plate Count Agar (Difco), a replica was made onto the DNA agar

GENETICALLY MANIPULATED DNA 131

and left growing overnight. After incubation, toluidine blue was poured onto the plate and colonies that devel- oped a pink halo were considered positive for the pro- duction of nucleases. Pure cultures of Staphylococcus aureus and E . coli were cultured in the same manner as positive and negative controls, respectively.

Dot Hybridization

Dilutions from the various time aliquots of each water source were blotted onto nitrocellulose membranes (S & S, Kneene, New Hampshire) with the help of slot-blot or dot-blot manifolds (Bio-Dot SF, Bio Rad, Richmond, CA). The DNA was immobilized in a vac- uum oven (SOOC for 2 h). Hybridization conditions were as follows: prehybridization was done with 5x Den- hardt's solution, 5x sodium saline citrate (SSC), 0.1% SDS, 100 pg/mL salmon sperm DNA, at 45°C. Radiola- beling of the DNA probes (437 bp plasmid insert) was done using the Random Primer Extension Labeling System NEP-103 (DuPont Co., Wilmington, DE) in the presence of high specific activity 32P-dCTP (3000 Ci/ mmole, Amersham, Arlington Heights, IL). The la- beled probe (specific activity, 8 x lo8 CPMlpg) was added to a final concentration of 1 X lo6 CPM/mL. Posthydridization washes were done under high strin- gency conditions (Sambrook et al., 1989). Autoradiog- raphy was done by overnight (or longer, if necessary) exposure to Kodak X-Omat AR film at -70°C.

TABLE 1. Bacterial densities in the aquatic systems seeded with GED

Total Water Viable Total Fecal DNAse-Producing Source Counta Coliformb Coliformb BacteriaC

- Distilled 4 x lo2 0 0 Tap I x lo2 0 0 + River 3 x lo6 2 x lo2 20 + Marine 9 x lo2 4 x 10' 3 +

a CFUlrnL (mean values from three trials). CFU/100 mL (mean values from three trials). + indicates presence of DNAse-producing bacteria. No quantia-

tion was attempted.

fecal coliforms were detected only in the river and ocean samples (at concentrations of 200 and 20 CFUl 100 mL; and 40 and 3 CFU/100 mL, respectively). There was no detectable incorporation of GED by the resident microflora as indicated by a lack of bacterial growth on antibiotic containing plates. This was further demonstrated by colony hybridization of baceria from the systems grown on antibiotic-free LB medium. The negative results of the colony hybridization (data not shown) reduce the possibility that there may have been transformation but no expression of the genes in ques- tion. Extracellular DNAses were present in all but the

Bacterial Transformations

TO determine the bioactivity (i.e., ability of the gene to be taken UP and expressed by bacterial cells) of the naked genetically engineered DNA, bacterial transfor- mation studies were carried out. Commercially ob- tained E . coti HB 101 subcloning efficiency competent

distilled water (Table I); however, no quantification of the DNAse-producing bacteria was attempted in this study. In all cases positive DNA agar plates gave a reaction covering most of the plate.

Dot Hybridization cells (BRL, Gaithersburg, MD) were used as plasmid hosts. The transformation procedure was conducted as described by the supplier. Aliquots of competent cells (2 x 108/mL) were mixed with sample dilutions from the marine and tap water systems. After a 30 min incubation in ice, cells were heat shocked for 20 s at 42"C, followed by a 1 h incubation at 37°C prior to plating. The experimental reaction was diluted, and cells spread on selective media and incubated at 37°C for expression. Transformation efficiency [colony forming units (CFU)/pg] was determined for each source sampled.

RESULTS

Bacterial Counts and Nuclease Tests

Figure 1 shows the fate (over time) of genetically engi- neered DNA in the presence of naturally occurring bacterial populations by means of dot-blot analysis. The concentration of seeded genetically engineered DNA (10 pg/mL in each system) remained constant up to five days in distilled water; on the other hand, GED seeded into tap water begins to decline at day 5. Both of these sets of results are in sharp contrast to the seeded river and marine water systems. Barely any DNA is detectable after only 24 h in the river and marine waters.

In order to follow more closely the degradation rate of seeded genetically engineered DNA, shorter time points were taken; distilled and tap water were sampled daily for up to 7 days, and the river and marine water systems were sampled every 6 h for up to 40 h (Fig.

Table I shows total viable counts for the four aquatic sources, which ranged from lo6 CFU/mL (river water) to lo2 CFU/mL (the other three systems). Total and

2). In this set of experiments, all water sources were seeded with 2.0 pg GED/mL. Similar results to the first set of experiments were observed. Plasmid degra-

132 ALVAREZ ET AL.

Fig. 1. Dot-blot analyses of aliquots obtained daily (up to 5 days) from GED seeded (10 pg/mL) aquatic systems. Autoradiographs show the results from tap, distilled, river, and marine water. Lanes 1-6 corresponds to sampling time in days. Negative (A-DNA) and positive (pWTAla 5 ’ ) controls are in columns 7 and 8 , respectively. Rows A-C are subsequent tenfold dilutions of each sample.

dation is visible after 6 days in tap water and the geneti- cally engineered DNA remained undegraded in the dis- tilled water microcosm for up to 7 days, at which time a slight decrease in the concentration was observed. On the other hand, in the marine water system the seeded DNA was fully degraded after 12 h, and in the river water all the DNA was degraded after 18 h, as indicated by the absence of hybridization signal on the autoradiography. In addition, these samples were subjected to amplification using the polymerase chain reaction using specific primers (data no shown). No amplification was observed, indicating that the absence of a hybridization signal was in fact due to the complete degradation of the target DNA.

Transformation Resu I ts

Table I1 shows bacterial transformation experiments from various aliquots of the marine and tap water sys- tems (to compare two systems with rapid vs slow degra- dation rates). Samples analyzed correspond to 0, 1, and 7 days. The seeded DNA present in the marine water microcosm is shown to be bioactive (i.e., able

to transform and be expressed) only during the first day, which confirms the rapid degradation of the geneti- cally engineered DNA once exposed to highly contami- nated environments. However, GED present in the tap water system was determined to be bioactive up to 7 days.

DISCUSSION

The potential for transformation to occur in aquatic environments has not been studied extensively. An important aspect of gene transfer by transformation in the environment is the stability of extracellular DNA in natural bacterial habitats. For this reason, the fate of seeded genetically engineered DNA into four different nonsterile aquatic systems was monitored over a period of 7 days.

Data presented here indicate that extracellular DNA turns over rapidly in marine and river water systems. However, the rate of plasmid degradation varied among the systems. The rate was lowest in distilled water, and higher in tap, river, and marine water sys-

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134 ALVAREZ ET AL.

TABLE II. Transformation of competent E. coli HB101 cells by genetically engineered DNA seeded in marine and tap water

Incubation Time (Days) Marine WateP Tap Watera

0 1 7

2.4 x 105 7.4 x 105 Ob 1.8 x 105 Ob 5.4 x 103

a Transformation efficiency (CFU/pg of DNA). Ampicillin- resistant transformants were scored.

< I transformant/pg of DNA.

tems (7 days, 6 days, 18 h, and less than 12 h, respec- tively). Confirming these results were those from the bacterial transformation assays. Data show that trans- forming activity of the plasmid DNA was detectable in tap water (slow-degrading potential) for up to 7 days. However, biological activity in the marine water was detected only during the first day. These results confirm the rapid degradation of the genetically engineered DNA upon being exposed to environments containing high bacterial concentrations and DNAses.

The degradation of DNA may be directly related to the presence of extracellular nucleases (Aardema et al., 1983; Stotzky, 1989). It remains to be determined if the enzymes responsible for the DNA degradation are endonucleases or exonucleases. The exclusive presence of exonucleases in a system may lead to the slow degradation of a plasmid, while they could be easily and rapidly degraded in the presence of endo- and exonucleases. This may be underlying the slow degradation of the DNA in the tap water. The seeded plasmid was mostly in the covalently closed circular conformation (as observed by gel electrophoresis) when seeded; therefore, the presence of an endonucle- ase would be necessary to nick the plasmid before complete degradation by the exonucleases could be achieved. Plausible explanations for the degradation could include as yet unknown abiotic factors (Awong et al., 1990; Romanowski et al., 1992). More likely the degradation may be due to lysis of bacteria1 cells, which would result in the release of intracellular nucleases.

These results agree with previous results in aquatic systems (Awong et al., 1990; DeFlaun and Paul, 1989; Paul et al., 1989). DeFlaun and Paul (1989) showed that intact plasmid DNA introduced into natural estuarine water was not detectable after 8 h. In contrast, a rela- tively long persistance of plasmid DNA in soil has been demonstrated before (Greaveas and Wilson, 1970; Khanna and Stotzky, 1992; Romanowski et al., 1992). These latter investigations demonstrated that DNA ad- sorption to minerals and other surfaces contributed to its protection against nucleolytic degradation.

The probability of the transfer of genetic information between GEMs and the indigenous organisms in the environment is one of the primary concerns related to the possible future release of GEMs. A very rapid turnover time for extracellular genetically engineered DNA does not favor transformation in the river and marine environments, the systems most likely to be receptors of genetically manipulated DNA. It would seem very unlikely that treated tap water would be the recipient of an accidental release.

In conclusion, there was a direct relation between the presence of DNAses and the degradation of geneti- cally engineered DNA in various aquatic systems. In some aquatic environments extracellular DNA will be degraded in a very short time, thus uptake of geneti- cally engineered DNA by resident microflora may be minimal.

This study was supported, in part, by RIM1 No. M3289B91, INDUNIV No. 9020539303 and G-91-02, and WRRI NO. 14-08-OOOlG1611.

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