Role of Divalent Cations in Plasmid DNA Adsorption to Natural

Jul 6, 2007 - The adsorption kinetics of supercoiled and linear plasmid DNA onto a natural organic matter (NOM)-coated silica surface are acquired usi...
0 downloads 7 Views 157KB Size
Download PDF
Environ. Sci. Technol. 2007, 41, 5370-5375

Role of Divalent Cations in Plasmid DNA Adsorption to Natural Organic Matter-Coated Silica Surface T H A N H H . N G U Y E N * ,† A N D KAI LOON CHEN‡ Department of Civil and Environmental Engineering, University of Illinois at Urbana-Champaign, 3230 Newmark Laboratory, Urbana, Illinois 61801, and Department of Chemical Engineering, Environmental Engineering Program, Yale University, New Haven, Connecticut 06520-8286

The adsorption kinetics of supercoiled and linear plasmid DNA onto a natural organic matter (NOM)-coated silica surface are acquired using a quartz crystal microbalance with dissipation monitoring (QCM-D) in the presence of common divalent electrolytes CaCl2 and MgCl2. The adsorption kinetics of both DNA are noticeably higher in the presence of CaCl2 compared to MgCl2. We hypothesize that specific bridging between the DNA phosphate groups and NOM carboxyl functional groups in the presence of Ca2+ cations may lead to more efficient attachment than in the presence of Mg2+ cations, which are only likely to allow for charge neutralization. The influence of background Na+ cations on the adsorption kinetics in the presence of CaCl2 is found to be insignificant, while the presence of Na+ leads to slower attachment kinetics in MgCl2. Rinsing the DNA layer adsorbed in the presence of CaCl2 with a solution of low NaCl concentration followed by deionized water does not result in observable detachment, indicating irreversibility of DNA adsorption. Instead, softening of the DNA layer adsorbed in the presence of CaCl2 with background Na+ occurs with the rinses due to the increase in electrostatic repulsion between the phosphate functional groups along the DNA backbone. In the case of the DNA layer adsorbed in the presence of CaCl2 without background Na+, softening of the layer does not occur with the rinses.

Introduction Extracellular nucleic acids are found in both aquatic and soil/sediment environments in the concentration range of µg per L of water and µg per g of soil, respectively (1-8). Some live bacteria release extracellular DNA actively, while others do so only upon cell lysis (9). Even though extracellular DNA are degraded in soils by deoxyribonuclease (DNase), sufficient evidence has shown that DNA adsorption to soil particles provides physical protection against degradation (10-13). Moreover, adsorbed DNA retain their ability to transform competent bacteria through natural transformation (14-17). It has been suggested that the occurrence of lateral gene transfer can result in microbial diversity (1519). The competent or naturally transformable bacteria are * Corresponding author phone: 217-244-5965; fax: 217-333-6968; e-mail: [email protected]. † University of Illinois at Urbana-Champaign. ‡ Yale University. 5370

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 41, NO. 15, 2007

able to develop a regulated physiological ability to take up extracellular DNA and subsequently integrate the acquired DNA into their genome. More than 40 bacteria have been identified to develop competence (15-17). In addition, it has been proposed that extracellular DNA are an unaccounted source of phosphorus in aquatic environments (5, 20). Experimental studies conducted with the objective of estimating the degree of DNA adsorption to clay and sand sediments under different solution conditions have mostly been based on the batch or column reactor technique (11, 21-26). For example, Ogram et al. (23) showed that the smaller DNA fragments (i.e., 2.69 kbp) adsorbed to soils to a greater degree compared to the larger DNA fragments of 11.19 and 23 kbp. Poly et al. (21) employed cryoscanning electron micrographs to demonstrate that plasmid DNA can bridge clay platelets together. Cai et al. (13) used Fourier transform infrared (FTIR) spectroscopy, circular dichroism, and fluorescence to show that salmon sperm DNA retain their B-form conformation upon binding to montmorillonite and organic clay, but change to Z-form conformation upon binding to kaolinite and other inorganic clay. Adsorption experiments to determine the initial kinetics of DNA adsorption onto silica surfaces using a quartz crystal microbalance with dissipation (QCM-D) were reported for the first time by Nguyen and Elimelech (27). Most surfaces in the environment are covered with natural organic matter (NOM). Therefore, studies on adsorption of extracellular DNA to NOM-coated surfaces would help us understand the fate, transport, and bioavailability of extracellular DNA in natural aquatic systems. Early studies demonstrated that DNA adsorbed to humic acid were protected against degradation by DNase and that the adsorbed DNA retained their ability to transform competent bacteria (12, 28). Nguyen and Elimelech (29) found that the kinetics of DNA adsorption to NOM-coated silica surfaces depend on ionic strength and that DNA adsorption is significant at moderately high ionic strength (e.g., 300 mM). However, until now, no systematic study on the role of divalent cations in the adsorption of DNA onto NOM-coated surfaces has been conducted. In this paper, we compare the DNA adsorption rates on Suwannee River NOM-coated silica surfaces in the presence of two common divalent electrolytes (CaCl2 and MgCl2) by using a QCM-D. We also investigate the possibility of DNA detachment and the change in conformation of the adsorbed DNA by exposing the DNA layer to solutions of low ionic strengths. This investigation includes both supercoiled and linear plasmid DNA.

Materials and Methods Plasmid DNA Preparation. E. coli XL1 blue strains with the ampicillin-resistant plasmid vector pGEM-T Easy (3015 bp) were used as a source of plasmid DNA. The protocols for the extraction and purification of plasmid DNA were reported in ref 29. Briefly, 5 L of bacteria were grown overnight in LB broth to obtain a cell concentration of 109/mL, determined though measurement of the optical density (SmartSpect 3000, Biorad Laboratory, CA). Qiagen Endofree Plasmid Giga kits (Qiagen Inc., CA) were used to extract and purify the supercoiled plasmid DNA. Half of the purified supercoiled plasmid DNA were then digested with the Nsi I enzyme (New England Biolabs, Beverly, MA) for plasmid DNA linearization. The digested linear plasmid DNA were purified with the Qiagen Plasmid Column to remove residual enzymes. Both the supercoiled and linear plasmid DNA were dissolved in Endofree DI water and stored at -20 °C before use. Using a NanoDrop ND-1000 spectrophotometer (Nanodrop Tech10.1021/es070425m CCC: $37.00

 2007 American Chemical Society Published on Web 07/06/2007

nologies, DE), the final supercoiled and linear DNA stock solutions were determined to have concentrations of 1500 and 1200 mg/L, respectively. The complete digestion for the linear DNA stock solution as well as the absence of chromosome DNA and protein contamination for both stock solutions were verified using the UV spectrophotometer and agarose gel electrophoresis, as reported in refs 27 and 29. Plasmid DNA Diffusion Coefficients. Dynamic light scattering (DLS) measurements were conducted to estimate the diffusion coefficients of the supercoiled and linear plasmid DNA in the presence of different solution chemistries. A multidetector light scattering unit (ALV-5000, Langen, Germany) was employed to carry out the DLS measurements. It utilizes a Nd:vanadate laser (Verdi V2, Coherent, Santa Clara, CA) with a wavelength of 532 nm, and the DLS measurements were conducted at a scattering angle of 90°. More details on the unit and experimental protocol are provided in our earlier publications (27, 29). For each solution chemistry, 20 DLS measurements were conducted on each DNA sample. For each measurement, the decay of the autocorrelation function was fitted with a biexponential function, and the slow relaxation time obtained from the fitting procedure was used to derive a value of the diffusion coefficient of the DNA (30, 31). The mean and standard deviation of the diffusion coefficients obtained from the 20 measurements for each solution composition were calculated. Because of possible dependence of the obtained diffusion coefficient on the scattering angle (30, 31), the average diffusion coefficients derived at a single angle in this study are only approximations of the true DNA translational diffusion coefficients. Quartz Crystal Microbalance with Dissipation Monitoring (QCM-D). In this study, we used the QCM-D D300 system (Q-Sense AB, Gothenburg, Sweden) to determine the DNA adsorption rate, adsorption reversibility, and the viscoelastic properties of the adsorbed DNA layer. This system employs an ultrasensitive AT-cut silica-coated quartz crystal sensor with a fundamental resonant frequency f0 ≈ 5 MHz housed in a radial stagnation point flow cell. The flow cell has welldefined geometry and hydrodynamic characteristics. This technique allows for the real-time monitoring of mass adsorption with no required labeling. The piezoelectric quartz crystal oscillates laterally with an amplitude of 1-2 nm when a voltage is applied to the electrodes affixed to the quartz crystal. As deposition (adsorption) occurs on the crystal surface, it leads to a shift in the vibrational frequency of the crystal. In addition to monitoring the frequency shift to determine the mass of DNA adsorbed on the crystal, information on the adsorbed DNA structure can be extracted by simultaneous monitoring of the energy of dissipation. In the DNA adsorption experiments, the variations of frequency (∆f) and dissipation (∆D) were monitored at three overtones (n ) 3, 5, and 7). The adsorption rate was taken to be indicated by the initial rate of change in frequency at the third overtone (∆f(3)). As vibration of the crystal takes place, the sum of all energy losses in the system per oscillation cycle is expressed as dissipation factors. For example, a soft film loses more energy during oscillation and therefore has a higher dissipation than a stiff film. The Voigt model presented in ref 32 was used to derive the viscoelastic properties of the DNA adsorbed layer. Specifically, experimental data were collected at two or three overtones (i.e., n ) 3,5; 3,7; 3,5,7), and assumed values of solution density (1000 kg/m3), solution viscosity (10-3 Pa s), and density of the adsorbed layer (1030 kg/m3) were employed in the Voigt model. The fitting parameters are the viscosity, shear modulus, and thickness of the adsorbed layer. More details on the experimental and data analysis technique can be found elsewhere (33). DNA Adsorption onto NOM-Coated Silica Surface. The quartz crystals were supplied by Q-Sense AB (batch 051031).

Before each experiment, the crystals were soaked in 2% Hellmanex II (Hellma GmbH & Co. KG, Mu¨llheim, Germany) cleaning solution for at least 2 h, rinsed thoroughly with deionized (DI) water, dried with ultrahigh-purity N2, and treated in an ozone/UV chamber for 30 min. The cleaned crystal was mounted in a temperature-controlled (25 °C) chamber of the QCM-D system. All the test solutions were fed into the chamber using a syringe pump (Kd Scientific Inc., Holliston, MA) operating in a withdrawal mode at a rate of 0.1 mL/min. For each adsorption experiment, the quartz crystal in the chamber was first pre-coated with a layer of Suwannee River NOM. This was done by equilibrating the crystal with a HEPES buffer made from 10 mM N-(2-hydroxyethyl)piperazine-N′2-ethanesulfonic acid and 100 mM NaCl which had been filtered through a 0.22 µm cellulose acetate membrane. The equilibration process usually took at least 30 min to achieve a reasonably stable baseline. After the equilibration period, the crystal was coated with a layer of poly-L-lysine (PLL with molecular weight e150 kDa) by flowing 2 mL of PLL hydrobromide solution prepared in the HEPES buffer at a concentration of 0.1 g/L through the sensor chamber (34, 35). At a pH of 5.8, the PLL layer was positively charged. This layer was rinsed with 2 mL of HEPES buffer for 20 min, followed by 2 mL of 10 mM NaCl for an additional 20 min. This layer of PLL was then coated with a layer of NOM by flowing 2 mL of the pre-prepared NOM solution, comprising 21.83 mg/L TOC and 10 mM NaCl, through the sensor chamber. This NOM layer was subsequently rinsed with 2 mL of 10 mM NaCl solution, followed by 2 mL of DNA-free electrolyte solution of the same ionic composition to be used in the subsequent DNA adsorption experiment. Finally, DNA adsorption experiments were conducted by flowing 4 mL of plasmid DNA solution that had been diluted in an electrolyte solution at a concentration of 120 mg/L over the NOM-coated silica surface (step 1), followed by rinsing the DNA layer with 2 mL of the same electrolyte solution in the absence of DNA (step 2). The studied electrolyte solutions were 1 mM Ca2+ with or without a background electrolyte of 7 mM Na+, and 1 mM Mg2+ with or without a background electrolyte of 7 mM Na+. For the adsorption experiments conducted in electrolyte solutions containing background Na+, the adsorbed DNA layer was then rinsed with 2 mL of 10 mM NaCl solution (step 3), followed by 2 mL of 1 mM NaCl solution (step 4), and finally 2 mL of DI water (step 5). For the experiments with electrolyte solutions containing no background Na+, step 3 was omitted and steps 4 and 5 were performed. The adsorption kinetics of DNA onto the NOM layer were determined in step 1, while adsorption reversibility and structural changes of plasmid DNA layers as a function of solution composition were observed in steps 3-5. All experiments were conducted in duplicate at an ambient pH of 5.8 and a flow rate of 0.1 mL/min. The DNA adsorption rate is represented by the initial rate of frequency shift as a function of time. The adsorption rate for each solution composition employed is then presented as attachment efficiency R, which is the normalization of the adsorption rate measured for that solution chemistry by the favorable adsorption rate of DNA onto a PLL layer. The favorable adsorption rates of 16.0 Hz/min for supercoiled plasmid DNA, and 17.0 Hz/min for linear DNA, were published in the previous work of Nguyen and Elimelech (29). Details on the preparation of the electrolytes and NOM solution can be found in the Supporting Information.

Results and Discussion Plasmid Diffusion Coefficients. Figure 1a and b presents the diffusion coefficients, estimated from DLS measurements, for supercoiled and linear plasmid DNA in the divalent VOL. 41, NO. 15, 2007 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

5371

FIGURE 1. Diffusion coefficients of supercoiled and linear plasmid DNA in (a) 1 mM Ca2+ and Mg2+ solutions with 7 mM Na+, and (b) 1 mM Ca2+ and Mg2+ solutions with no Na+. All solutions were prepared at ambient pH of 5.8. The presented data are averages and standard deviations of 20 measurements for each DNA sample.

electrolyte solutions in the presence and absence of background NaCl (7 mM), respectively. Our literature review revealed no previous study reporting DLS measurements for diffusion coefficients of plasmid DNA molecules in the presence of divalent cations. Fishman and Patterson (36) reported diffusion coefficients for supercoiled and nicked plasmid DNA with sizes varying from 2 to 10 kbp in solutions containing 100 mM Na+. These diffusion coefficients ranged from 2.5 × 10-8 cm2/s (for a 10 kbp plasmid) to 5.4 × 10-8 cm2/s (for a 2 kbp plasmid). In another study, Liu et al. (37) reported a similar range of diffusion coefficients for 2.3 kbp and 1.5 kbp plasmid DNA (4.5 × 10-8 cm2/s to 6.3 × 10-8 cm2/s in 100-200 mM Na+). Hence, the diffusion coefficients we obtained in this study fall reasonably close to the ones reported in the literature. Plasmid DNA Adsorption Kinetics. The attachment efficiencies R of plasmid DNA to the NOM-coated silica surface in the presence of 1 mM Ca2+ are shown in Figure 2a. In the solutions of 1 mM Ca2+ with and without background Na+, the attachment efficiencies are statistically similar 0.50 ( 0.10 vs 0.40 ( 0.10 for supercoiled DNA, and 0.040 ( 0.10 vs 0.30 ( 0.07 for linear DNA). Among the alkaline earth metal ions, Ca2+ is the only cation with which the DNA phosphate groups can form inner sphere complexes (38, 39). A recent molecular dynamic simulation by Kanilichev and Kirkpatrick (40) suggests that the carboxyl groups of NOM can also form inner sphere complexes with Ca2+. Because both NOM carboxyl groups and DNA phosphate groups are able to form inner sphere complexes with Ca2+, we hypothesize that Ca2+ may form bridges between those two groups. On the other hand, NOM carboxyl groups do not form inner sphere complexes with Na+, and Na+ can only screen the charges of NOM macromolecules (40). At the same time, Na+ can also screen the negative charges along the DNA backbone. Thus, based on our earlier hypothesis, we postulate that the specific bridging between the DNA and NOM controls the adsorption kinetics in 1 mM Ca2+ both in the presence 5372

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 41, NO. 15, 2007

FIGURE 2. Attachment efficiencies of supercoiled and linear plasmid DNA on NOM-coated silica surfaces at ambient pH of 5.8 in solutions of (a) 1 mM Ca2+ with and without 7 mM Na+ and (b) 1 mM Mg2+ with and without 7 mM Na+. Shown in the graphs are average values of at least 2 replicate measurements. Error bars indicate standard deviations. The plasmid concentration employed in the experiments was 120 mg/L. The experiments were all conducted at 25 °C. and absence of background Na+. The presence of Na+ is unlikely to exert significant influence over the formation of calcium bridges between DNA phosphate and NOM carboxyl groups. Figure 2b presents the attachment efficiencies of plasmid DNA to the NOM-coated silica surface in the presence of 1 mM Mg2+ with and without background Na+. The attachment efficiencies are statistically lower in the presence of background Na+ for both DNA (0.01 ( 0.001 vs 0.09 ( 0.05 for supercoiled DNA, and 0.97 ( 0.01 vs 0.25 ( 0.02 for linear DNA). Unlike in the presence of Ca2+, the phosphate groups along the DNA backbone form weaker outer sphere complexes with Mg2+ (38, 39). Also, the NOM carboxyl groups are not able to form inner sphere complexes with Mg2+ (40). Therefore, we hypothesize that Mg2+ is unlikely to form bridges between DNA phosphate groups and NOM carboxyl groups. Instead, the presence of Mg2+ leads to charge neutralization of the DNA phosphate groups and NOM carboxyl groups, thus reducing the energy barrier between the DNA macromolecules and NOM layer. In the presence of background Na+, Na+ may compete with Mg2+ for the DNA phosphate groups, leading to a reduced degree of charge neutralization by Mg2+. Similarly, Na+ may also compete with Mg2+ for the carboxyl groups of NOM. As a result, the energy barrier between DNA molecules and the NOM layer increases in the presence of background Na+, leading to a lower adsorption rate compared to the rate in the absence of background Na+, as shown in Figure 2b. Comparing Figure 2a and b, an important observation is that the adsorption rates are higher in the presence of Ca2+ than in Mg2+. This behavior is consistent with our hypothesis that calcium bridging between the DNA macromolecules and NOM layer gives rise to more favorable adsorption than in the presence of Mg2+ which only allows for charge neutral-

FIGURE 3. Frequency shifts normalized by the third harmonic number and their associated dissipation shifts as a function of time for supercoiled plasmid DNA adsorption at ambient pH of 5.8 and in the presence of Ca2+. The plasmid DNA concentration employed in the experiments was 120 mg/L and temperature was maintained at 25 °C. (a) The plasmid DNA adsorption took place in 1 mM CaCl2 and 7 mM NaCl (step 1). The adsorbed plasmid DNA layer was washed with 1 mM CaCl2 and 7 mM NaCl solution (step 2), 10 mM NaCl solution (step 3), 1 mM NaCl solution (step 4), and DI water (step 5). (b) The plasmid DNA adsorption took place in 1 mM CaCl2 (step 1). The adsorbed plasmid DNA layer was washed with 1 mM CaCl2 solution (step 2), 1 mM NaCl solution (step 4), and DI water (step 5). ization. Similar observation and explanation were reported in ref 41 on adhesion forces experienced between NOM macromolecules in the presence of Ca2+ and Mg2+ measured through the employment of the atomic force microscope (AFM). However, a more comprehensive study has to be conducted to verify our hypothesis that Ca2+ bridges DNA and NOM. Is DNA Adsorption Reversible? To study the reversibility of DNA adsorption, the adsorbed DNA layer was exposed to solutions of lower ionic strengths. Specifically, the DNA layers formed in the presence of 1 mM Ca2+ and 1 mM Mg2+, both with background electrolyte of 7 mM Na+, were rinsed sequentially with 10 mM Na+ solution (step 3), 1 mM Na+ solution (step 4), and DI water (step 5). The DNA layers formed in the presence of 1 mM Ca2+ and 1 mM Mg2+ without the background electrolyte of NaCl were rinsed sequentially with 1 mM Na+ solution (step 4) and DI water (step 5). The occurrence of DNA detachment from the NOM layer will be indicated by an increase in frequency response and a simultaneous decrease in the dissipation response. To ensure that the changes in frequency and dissipation response are due to changes in the DNA layer and not changes of the bulk solution, we conducted independent control tests in which we rinsed the NOM layers sequentially with 1 mM Ca2+ or Mg2+ solutions with background NaCl (7 mM), 10 mM NaCl,

FIGURE 4. Frequency shifts normalized by the third harmonic number and their associated dissipation shifts as a function of time for supercoiled plasmid DNA adsorption at ambient pH of 5.8 and in the presence of Mg2+. The plasmid DNA concentration employed in the experiments was 120 mg/L and temperature was maintained at 25 °C. (a) The plasmid DNA adsorption took place in 1 mM MgCl2 and 7 mM NaCl (step 1). The adsorbed plasmid DNA layer was washed with 1 mM MgCl2 and 7 mM NaCl solution (step 2), 10 mM NaCl solution (step 3), 1 mM NaCl solution (step 4), and DI water (step 5). (b) The plasmid DNA adsorption took place in 1 mM MgCl2 (step 1). The adsorbed plasmid DNA layer was washed with 1 mM MgCl2 solution (step 2), 1 mM NaCl solution (step 4), and DI water (step 5). and finally 1 mM NaCl. The magnitudes of the frequency and dissipation shifts were comparable with the ones of baseline fluctuation. This observation verified that the changes of bulk solution did not result in significant shifts in the frequency and dissipation response. During the rinsing of the supercoiled DNA layers formed in 1 mM Ca2+ and Mg2+ with background Na+ with 10 mM Na+ solution (step 3), either small or insignificant frequency shifts were observed (Figures 3a and 4a). During the rinsing of the layers with 1 mM Na+ and then DI water, small increases in frequency of about 0.4 Hz (step 4) and 1.8 Hz (step 5) were observed for all cases. These changes in frequency in steps 3-5 were associated with increases in dissipation, suggesting that no DNA detached from the NOM layer. Instead, softening of the DNA layers occurred. This irreversibility of adsorption and softening behavior was also observed during the rinsing of the linear DNA layers formed in 1 mM Ca2+ and Mg2+ with background Na+ (Figures 1Sa and 2Sa in the Supporting Information). The softening behavior will be discussed in greater detail in the following section. The absence of DNA detachment was consistent with previous results reported by Crecchio et al. (12), who observed no desorption of DNA when the DNA adsorbed to clay-humic acid complexes were washed with DI water. VOL. 41, NO. 15, 2007 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

5373

FIGURE 5. (a) Thicknesses and (b) shear viscosities of the supercoiled and linear plasmid DNA layers formed in the presence of 1 mM Ca2+ and 7 mM Na+ in different ionic compositions at ambient pH of 5.8. (c) Thicknesses and (d) shear viscosities of the supercoiled and linear plasmid DNA layers formed in the presence of 1 mM Ca2+ and no Na+ in different ionic compositions at ambient pH of 5.8. Shown in the graphs are representative data for at least 2 replicate measurements. The plasmid DNA concentration employed in the experiments was 120 mg/L and temperature was maintained at 25 °C. No detachment from the supercoiled DNA layers formed in the presence of 1 mM Ca2+ and Mg2+ with no background Na+ was also observed, as shown in Figures 3b and 4b. Similarly, we observed that the adsorption of the linear DNA in 1 mM Ca2+ and Mg2+ was irreversible (Figures 1Sb and 2Sb in the Supporting Information). These results imply that the DNA adsorbed in the presence of both 1 mM Ca2+ and Mg2+, regardless of the presence of 7 mM NaCl, is stable and not likely to be washed off when exposed to solutions of low salt concentrations, which can take place, for instance, in the events of rain and floods. Irreversible adsorption of plasmid DNA to NOM also explains why it is difficult to recover adsorbed extracellular DNA from soil samples using mild treatments, as reported in several studies on extraction and detection of extracellular DNA from soils (6, 42, 43). Conformation Changes of Adsorbed Plasmid DNA. Figure 5 shows the changes in the thickness and viscosity of the supercoiled and linear plasmid DNA layers formed in the presence of 1 mM Ca2+ either with or without background Na+ as the rinsing with the different solutions was performed. For both supercoiled and linear plasmid DNA layers formed in 1 mM Ca+ with background Na+, the changes in the frequency and dissipation response during the final two rinses with 1 mM NaCl solution and DI water described in the previous section reflect the increase in the thickness and decrease in the viscosity of the DNA layers, as obtained with the employment of the Voigt model (Figure 5a and b). In other words, softening of the layers occur during the rinses. In comparison, the DNA layers formed in 1 mM Ca+ without Na+ background exhibited a different behavior. The thickness and viscosity of the DNA adsorbed layers did not change significantly with the rinses (Figure 5c and d, and Table 1S in the Supporting Information). When the DNA layers are formed in the presence of both Ca2+ and background Na+, Na+ may compete with Ca2+ for the phosphate groups along the DNA backbone. Therefore, during the subsequent rinsing with 1 mM Na+ solution (step 5374

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 41, NO. 15, 2007

4) and DI water (step 5), it is possible that the decrease in the Na+ concentration in the bulk solutions may result in the decrease in charge shielding of the phosphate groups which are not complexed with Ca2+ and thus greater electrostatic repulsion between the subunits of the DNA macromolecules. This in turn will lead to softer and less compact DNA layers. For the layers formed in 1 mM Ca+ without Na+ background, the exchange of Ca2+ by Na+ may be insignificant since the inner sphere complexes formed between the DNA phosphate groups and Ca2+ are stronger compared to the outer sphere complexes formed in the presence of Mg2+. This is reflected by the DNA layer thickness which remains constant during the rinses with 1 mM NaCl solution and DI water. In addition, for the DNA layers formed with or without background Na+, the linear plasmid DNA layers are thicker with lower viscosity compared to the supercoiled DNA layers. This is likely because linear DNA molecules are able to extend themselves into the bulk solution more than the supercoiled DNA molecules. The changes in thickness and viscosity of the DNA layer formed in 1 mM Mg2+ either with or without Na+ are shown in Figure 3S (Supporting Information). The changes due to the rinses with 1 mM NaCl solution and DI water are not observable here due to the fact that there was considerably less DNA adsorption in the presence of Mg2+ than in Ca2+. In summary, we hypothesize that DNA adsorption to the NOM-coated silica surface in the presence of 1 mM Ca2+ is mainly controlled by specific bridging between the DNA phosphate groups and NOM carboxyl groups. On the other hand, charge neutralization is likely to be the major adsorption mechanism in the presence of 1 mM Mg2+. Exposing the DNA layer pre-adsorbed in the presence of Ca2+ with background Na+ to solutions of lower ionic strengths does not lead to DNA detachment, but results in a softer DNA layer. Rinsing the DNA layer pre-adsorbed in the presence of Ca2+ with no background Na+ with solutions of lower ionic strengths does not result in DNA desorption or softening of the layer.

Acknowledgments Funding was provided by the Yale Institute for Biospheric Studies and the National Science Foundation (BES 0228911). We thank Professor Menachem Elimelech for his guidance during the study. We also thank Professor Paul Van Tassel for letting us use the QCM-D and Professor Barbara Kazmierczak for providing the bacterial strain.

Supporting Information Available Frequency shifts normalized by the third harmonic number and their associated dissipation shifts as a function of time for linear plasmid DNA adsorption for linear plasmid DNA adsorption in the presence of Ca2+ (Figure 1S) and Mg2+ (Figure 2S); plots of estimated thicknesses and shear viscosities of the adsorbed plasmid DNA layers formed in the presence of 1 mM Mg2+ in different ionic compositions (Figure 3S); tabulated data for shear viscosity and effective thickness as obtained using the Voigt-based model (Table 1S); salt and NOM solution preparation. This material is available free of charge via the Internet at http://pubs.acs.org.

Literature Cited (1) Torsvik, V. L.; Goksoyr, J. Determination of bacterial DNA in soil. Soil Biol. Biochem. 1978, 10 (1), 7-12. (2) Holmhans, O; Sutcliff, Wh.; Sharp, J. Measurement of deoxyribonucleic acid in ocean and its ecological significance. Limnol. Oceanogr. 1968, 13 (3), 507. (3) Corinaldesi, C.; Danovaro, R.; Dell’Anno, A. Simultaneous recovery of extracellular and intracellular DNA suitable for molecular studies from marine sediments. Appl. Environ. Microbiol. 2005, 71 (1), 46-50.

(4) Dell’Anno, A.; Bompadre, S.; Danovaro, R. Quantification base composition, and fate of extracellular DNA in marine sediments. Limnol. Oceanogr. 2002, 47 (3), 899-905. (5) Dell’Anno, A.; Corinaldesi, C. Degradation and turnover of extracellular DNA in marine sediments: Ecological and methodological considerations. Appl. Environ. Microbiol. 2004, 70 (7), 4384-4386. (6) Niemeyer, J.; Gessler, F. Determination of free DNA in soils. J. Plant Nutr. 2002, 165 (2), 121-124. (7) Frostegard, A.; Courtois, S.; Ramisse, V.; Clerc, S.; Bernillon, D.; Le Gall, F.; Jeannin, P.; Nesme, X.; Simonet, P. Quantification of bias related to the extraction of DNA directly from soils. Appl. Environ. Microbiol. 1999, 65 (12), 5409-5420. (8) Agnelli, A.; Ascher, J.; Corti, G.; Ceccherini, M. T.; Nannipieri, P.; Pietramellara, G. Distribution of microbial communities in a forest soil profile investigated by microbial biomass, soil respiration and dgge of total and extracellular DNA. Soil Biol. Biochem. 2004, 36 (5), 859-868. (9) Madigan, M. T.; Martinko, J. M.; Parker, J. Brock Biology of Microorganisms, 10th ed.; Prentice Hall: Upper Saddle River, NJ, 2003. (10) Demaneche, S.; Jocteur-Monrozier, L.; Quiquampoix, H.; Simonet, P. Evaluation of biological and physical protection against nuclease degradation of clay-bound plasmid DNA. Appl. Environ. Microbiol. 2001, 67 (1), 293-299. (11) Khanna, M.; Stotzky, G. Transformation of Bacillus subtilis by DNA bound on montmorillonite and effect of dnase on the transforming ability of bound DNA. Appl. Environ. Microbiol. 1992, 58 (6), 1930-1939. (12) Crecchio, C.; Ruggiero, P.; Curci, M.; Colombo, C.; Palumbo, G.; Stotzky, G. Binding of DNA from Bacillus subtilis on montmorillonite-humic acids-aluminum or iron hydroxypolymers: Effects on transformation and protection against dnase. Soil Sci. Soc. Am. J. 2005, 69 (3), 834-841. (13) Cai, P.; Huang, Q. Y.; Zhang, X. W. Interactions of DNA with clay minerals and soil colloidal particles and protection against degradation by dnase. Environ. Sci. Technol. 2006, 40 (9), 29712976. (14) Demaneche, S.; Kay, E.; Gourbiere, F.; Simonet, P. Natural transformation of Pseudomonas fluorescens and Agrobacterium tumefaciens in soil. Appl. Environ. Microbiol. 2001, 67 (6), 26172621. (15) Paget, E.; Simonet, P. On the track of natural transformation in soil. FEMS Microbiol. Ecol. 1994, 15 (1-2), 109-117. (16) Lorenz, M. G.; Wackernagel, W. Bacterial gene-transfer by natural genetic-transformation in the environment. Microbiol. Rev. 1994, 58 (3), 563-602. (17) Thomas, C. M.; Nielsen, K. M. Mechanisms of, and barriers to, horizontal gene transfer between bacteria. Nat. Rev. Microbiol. 2005, 3 (9), 711-721. (18) Ochman, H.; Lawrence, J. G.; Groisman, E. A. Lateral gene transfer and the nature of bacterial innovation. Nature 2000, 405 (6784), 299-304. (19) Droge, M.; Puhler, A.; Selbitschka, W. Horizontal gene transfer among bacteria in terrestrial and aquatic habitats as assessed by microcosm and field studies. Biol. Fertil. Soils 1999, 29 (3), 221-245. (20) Dell’Anno, A.; Danovaro, R. Extracellular DNA plays a key role in deep-sea ecosystem functioning. Science 2005, 309 (5744), 2179-2179. (21) Poly, F.; Chenu, C.; Simonet, P.; Rouiller, J.; Monrozier, L. J. Differences between linear chromosomal and supercoiled plasmid DNA in their mechanisms and extent of adsorption on clay minerals. Langmuir 2000, 16 (3), 1233-1238. (22) Ogram, A.; Sayler, G. S.; Gustin, D.; Lewis, R. J. DNA adsorption to soils and sediments. Environ. Sci. Technol. 1988, 22 (8), 982984. (23) Ogram, A. V.; Mathot, M. L.; Harsh, J. B.; Boyle, J.; Pettigrew, C. A., Jr. Effects of DNA polymer length on its adsorption to soils. Appl. Environ. Microbiol. 1994, 60 (2), 393-6. (24) Pietramellara, G.; Franchi, M.; Gallori, E.; Nannipieri, P. Effect of molecular characteristics of DNA on its adsorption and

(25)

(26)

(27)

(28)

(29)

(30)

(31)

(32)

(33)

(34)

(35)

(36)

(37)

(38)

(39)

(40)

(41)

(42)

(43)

binding on homoionic montmorillonite and kaolinite. Biol. Fertil. Soils 2001, 33 (5), 402-409. Blum, S. A. E.; Lorenz, M. G.; Wackernagel, W. Mechanism of retarded DNA degradation and prokaryotic origin of DNases in nonsterile soils. Syst. Appl. Microbiol. 1997, 20 (4), 513-521. Romanowski, G.; Lorenz, M. G.; Wackernagel, W. Plasmid DNA in a groundwater aquifer microcosm-adsorption, dnaase resistance and natural genetic-transformation of Bacillus subtilis. Mol. Ecol. 1993, 2 (3), 171-181. Nguyen, T. H.; Elimelech, M. Plasmid DNA adsorption on silica: Kinetics and conformational changes in monovalent and divalent salts. Biomacromolecules 2007, 8 (1), 24-32. Crecchio, C.; Stotzky, G. Binding of DNA on humic acids: Effect on transformation of Bacillus subtilis and resistance to dnase. Soil Biol. Biochem. 1998, 30 (8-9), 1061-1067. Nguyen, T. H.; Elimelech, M. Adsorption of plasmid DNA to a natural organic matter-coated silica surface: Kinetics, conformation, and reversibility. Langmuir 2007, 23 (6), 3273-3279. Langowski, J. Salt effects on internal motions of superhelical and linear puc8 DNA - dynamic light-scattering-studies. Biophys. Chem. 1987, 27 (3), 263-271. Langowski, J.; Giesen, U. Configurational and dynamic properties of different length superhelical DNAs measured by dynamic light-scattering. Biophys. Chem. 1989, 34 (1), 9-18. Voinova, M. V.; Rodahl, M.; Jonson, M.; Kasemo, B. Viscoelastic acoustic response of layered polymer films at fluid-solid interfaces: Continuum mechanics approach. Phys. Scr. 1999, 59 (5), 391-396. Rodahl, M.; Hook, F.; Krozer, A.; Brzezinski, P.; Kasemo, B. Quartz-crystal microbalance setup for frequency and q-factor measurements in gaseous and liquid environments. Rev. Sci. Instrum. 1995, 66 (7), 3924-3930. de Kerchove, A. J.; Elimelech, M. Formation of polysaccharide gel layers in the presence of Ca2+ and K+ ions: Measurements and mechanisms. Biomacromolecules 2007, 8 (1), 113-121. de Kerchove, A. J.; Elimelech, M. Structural growth and viscoelastic properties of adsorbed alginate layers in monovalent and divalent salts. Macromolecules 2006, 39 (19), 6558-6564. Fishman, D. M.; Patterson, G. D. Light scattering studies of supercoiled and nicked DNA. Biopolymers 1996, 38 (4), 535552. Liu, H.; Gapinski, J.; Skibinska, L.; Patkowski, A.; Pecora, R. Effect of electrostatic interactions on the dynamics of semiflexible monodisperse DNA fragments. J. Chem. Phys. 2000, 113 (14), 6001-6010. Minasov, G.; Tereshko, V.; Egli, M. Atomic-resolution crystal structures of b-DNA reveal specific influences of divalent metal ions on conformation and packing. J. Mol. Biol. 1999, 291 (1), 83-99. Kankia, B. I. Interaction of alkaline-earth metal ions with calf thymus DNA. Volume and compressibility effects in diluted aqueous solutions. Biophys. Chem. 2000, 84 (3), 227-237. Kalinichev, A. G.; Kirkpatrick, R. J. Molecular dynamics simulation of cationic complexation with natural organic matter. Eur. J. Soil Sci. 2007, in press. Li, Q. L.; Elimelech, M. Organic fouling and chemical cleaning of nanofiltration membranes: Measurements and mechanisms. Environ. Sci. Technol. 2004, 38 (17), 4683-4693. England, L. S.; Vincent, M. L.; Trevors, J. T.; Holmes, S. B. Extraction detection and persistence of extracellular DNA in forest litter microcosms. Mol. Cell. Probes 2004, 18 (5), 313319. Ogram, A.; Sayler, G. S.; Barkay, T. The extraction and purification of microbial DNA from sediments. J. Microbiol. Meth. 1987, 7 (2-3), 57-66.

Received for review February 19, 2007. Revised manuscript received May 16, 2007. Accepted May 18, 2007. ES070425M

VOL. 41, NO. 15, 2007 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

5375