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To better understand the influence of environmental conditions on the adsorption of extracellular chromosomal DNA and its availability for natural transformation, the amount and conformation of adsorbed DNA were monitored under different conditions in parallel with transformation assays using the soil bacterium Azotobacter vinelandii. DNA adsorption was monitored using the technique of quartz crystal microbalance with dissipation (QCM-D). Both silica and natural organic matter (NOM) surfaces were evaluated in solutions containing either 100 mM NaCl or 1 mM CaCl2. The QCM-D data suggest that DNA adsorbed to silica surfaces has a more compact and rigid conformation in Ca2+ solution than in Na+ solution and that the reverse is true when DNA is adsorbed to NOM surfaces. While the amounts of DNA adsorbed on a silica surface were similar for Ca2+ and Na+ solutions, the amount of DNA adsorbed on an NOM-coated surface was higher in Ca2+ solution than in Na+ solution. Transformation frequencies for dissolved DNA and DNA adsorbed to silica and to NOM were 6 × 10−5, 5 × 10−5, and 2.5 × 10−4, respectively. For NOM-coated surfaces, transformation frequencies from individual experiments were 2- to 50-fold higher in the presence of Ca2+ than in the presence of Na+. The results suggest that groundwater hardness (i.e., Ca2+ concentration) will affect the amount of extracellular DNA adsorbed to the soil surface but that neither adsorption nor changes in the conformation of the adsorbed DNA will have a strong effect on the frequency of natural transformation of A. vinelandii.
Horizontal gene transfer contributes to microbial evolution and provides mechanisms for the spread of both antimicrobial resistance genes and genetically engineered DNA. While most studies on horizontal gene transfer focus on conjugation, recent reviews on extracellular DNA (16, 27) document the need to consider also natural transformation. The amount of extracellular DNA in the soil is on the order of hundreds of ng/g of dry soil (27). Extracellular DNA adsorbs to many common soil constituents, including sand, clay, and natural organic matter (NOM) (16, 27), and once adsorbed may persist for days or years (16).
As first reported by Lorenz et al. (18), adsorbed DNA is available for natural transformation and therefore represents a potential environmental reservoir facilitating horizontal gene transfer. DNA adsorbed on sand surfaces has been successfully transferred to both Gram-positive and Gram-negative soil bacteria, including Bacillus subtilis (18), Pseudomonas stutzeri (19), and Acinetobacter calcoaceticus (4). Other studies have focused on transformation with DNA adsorbed to other surfaces (for example, clay minerals , humic acids , and intact soils [25, 31]). In most previous studies, adsorbed DNA transformed at a lower frequency than dissolved DNA (see, for example, references 4 and 11). However, higher transformation frequencies for adsorbed DNA than for dissolved DNA have been reported in two studies (18, 19). In addition, Demaneche et al. were unable to detect Pseudomonas fluorescens transformants in a variety of liquid media despite successful transformations in sterile soil columns (8).
Several studies have evaluated the influence of nutrients (24, 34) and soil (8, 11, 23, 31) on transformation efficiency. Less information is available on the influence of adsorbed DNA conformation on transformation efficiency. Pietramellara et al. speculated that the decrease in transformation rates they observed upon repeated wetting and drying cycles of adsorbed DNA was due to conformational changes (28). Cai et al. also speculated that differences in the conformation of adsorbed DNA could be responsible for lower transformation efficiencies for DNA bound to kaolinte and inorganic clays (2), based on their previous work characterizing adsorption to different surfaces (3). Detailed characterizations of the conformation of adsorbed DNA only recently became feasible, and the influence of the conformation of adsorbed DNA on transformation frequencies has not, to our knowledge, been systematically investigated.
Characterization of the mass and conformation of DNA adsorbed on different surfaces can be accomplished using quartz crystal microbalance with dissipation (QCM-D) (7, 22). QCM-D measurements are based on the shift in frequency (ΔF) and the decay in vibrating energy (ΔD) that occur as molecules adsorb to piezoelectric sensors (29, 33). Viscosity, elastic shear modulus, and effective thickness of the adsorbed material can be determined by fitting the frequency and dissipation data to the viscoelastic Voigt model (7). Based on Nguyen and colleagues' previous work with plasmid DNA adsorption on silica and NOM-coated surfaces, increasing electrolyte concentrations and the presence of divalent cations favor DNA adsorption (20-22). In addition, inner sphere complexation by Ca2+ with DNA phosphate backbone allows the formation of DNA-adsorbed layers that are more compact than the DNA-adsorbed layer formed by charge shielding in solution with a high Na+ concentration (20-22).
The objective of this study was to investigate both the adsorption of chromosomal DNA to representative soil particle surfaces and the effect of such adsorption on natural transformation. We used QCM-D to characterize the conformation of adsorbed DNA on silica and NOM surfaces under two different solution chemistries (100 mM Na+ and 1 mM Ca2+). The influences of adsorption and of differences in the conformation of the adsorbed DNA on transformation frequencies were tested in a common soil bacterium, Azotobacter vinelandii. A. vinelandii is naturally competent (26) but had not been previously reported to be transformed with adsorbed DNA.
Wild-type A. vinelandii strain DJ was the source of chromosomal DNA. Strain DJ77, which carries a 128-bp deletion in the nifH gene that prevents nitrogen fixation in the presence of molybdenum (12), was used as the recipient. The strains were cultured in modified Burk's (B) medium (32) at 30°C and 170-rpm shaking, with addition of 0.013 mol/liter ammonium acetate (BN medium) for growth of Nif− cells (13).
Chromosomal DNA was purified from cultures grown to an optical density at 560 nm (OD560) between 0.8 and 1.0 using standard procedures (30). Chromosomal DNA was dissolved into endotoxin-free water (catalog number 100371-028; VWR) and stored at −20°C. The concentration of DNA was measured by UV spectroscopy, and the size was monitored by agarose gel electrophoresis (30).
HEPES and poly-l-lysine (PLL) solutions were prepared as previously described (21). The resistivity of the MilliQ water used in QCM-D solutions was at least 17.5 MΩ. All solutions used in QCM-D tests were filtered with 0.22-μm-pore-size cellulose acetate membranes and stored at 4°C. Before use, the solutions were sonicated for at least 20 min to remove air bubbles and allowed to warm up to room temperature. Morpholinepropanesulfonic acid (MOPS) salt transformation buffers were prepared in MilliQ water and adjusted to pH 7.2 with HCl and NaOH. DNA stocks were warmed to room temperature immediately before being diluted in electrolyte solution to a final concentration of 20 mg/liter for adsorption and QCM-D tests.
Florida peat humic acid reference (International Humic Substances Society, St. Paul, MN) was used to prepare a soil NOM solution as described previously for Suwannee River NOM (20, 21). Briefly, 56.2 mg of NOM was dissolved in 100 ml of MilliQ water and stirred for 24 h before being filtered through a 0.22-μm-pore-size cellulose acetate filter. Total organic matter (TOC) for this NOM solution was 46 mg/liter as measured by a TOC-Phoenix 8000a. Na+, Ca2+, Mg2+, and Al3+ contents in this NOM solution were on the order of 0.01 ppb as measured by inductively coupled plasma optical emission spectrometry (ICP-OES; Microanalysis Laboratory, University of Illinois). The working NOM solution was prepared by diluting the filtrate to 4.6 mg of TOC/liter with NaCl solution, for a final concentration of 10 mM NaCl, and filtering through a cellulose acetate filter again.
Silica beads of 1.6-μm diameter were purchased from Bangs Laboratory, Inc. Uncoated and NOM-coated silica beads were prepared as described previously (17). The silica beads were washed three times with deionized (DI) water. Preparation of NOM-coated beads was a two-step process. First, 1.2 × 108 of the clean silica beads were mixed with 1 ml of PLL solution for 24 h and washed three times with DI water. Then the resulting PLL-coated beads were mixed with 1 ml of NOM solution for 24 h and again washed three times with DI water. For DNA adsorption, 1-ml aliquots of 20 mg/liter DNA in the desired solution were mixed with 100 μl of uncoated or NOM-coated silica bead suspension (1.2 × 108 particles/ml) in 1.5-ml microcentrifuge tubes. The tubes were rotated for 24 h in the dark, followed by centrifugation to separate the supernatant for quantification of the remaining dissolved DNA. The beads were then rinsed with the MOPS buffer twice before being resuspended at 1.2 × 108 particles/ml. The amount of DNA adsorbed on the beads was calculated by mass balance from the dissolved DNA concentrations measured before and after adsorption. No dissolved DNA was detectable in the wash solution.
Electrophoretic mobilities of uncoated (silica), PLL-coated, and NOM-coated beads as well as of the DNA and competent cells were determined using a Zetasizer Nano ZS90 (Malver Instruments, Southborough, MA). Uncoated, PLL-coated, and NOM-coated silica beads were suspended as 1.2 × 108 particles/ml in 100 mM Na+ or 1 mM Ca2+ solution. Zeta potentials were calculated from the electrophoretic mobility measurements using the Smoluchowski equation.
QCM-D has been used to determine viscoelastic properties of macromolecules (e.g., DNA and protein), as described in our previous work (20-22). The QCM-D technique is based on piezoelectricity of a thin quartz crystal (29, 33). When an electric current is going through the crystal, the crystal deforms and vibrates. As more mass adsorbs on the surface of the crystal sensor, the frequency of vibration decreases; the shift of vibration frequency (ΔF) is proportional to the mass adsorbed. When the voltage is turned off, the vibration of the crystal with the adsorbed layer dies off. If the adsorbed layer is a soft layer of macromolecules, the energy dissipated (dissipation, ΔD) during the dying-off time is larger than when the adsorbed layer is rigid. The mass adsorbed and some structural properties of the adsorbed layers can thus be obtained simultaneously and in real time from ΔD and ΔF. n is the overtone number.
A D300 QCM-D system (Q-sense AB, Gothenburg, Sweden) and AT-cut, silica-coated crystal sensors (sensor QSX 303, batch 070112-2; Q-Sense AB) mounted in a radial stagnant-point flow cell were used. The sensors were washed and dried before experiments and rinsed after experiments. All solutions were added to the flow cell using a syringe pump operated in suction mode at 0.1 ml/min.
For adsorption experiments on a silica surface, the flow cell was equilibrated first with filtered MilliQ water and then with the desired electrolyte solution, followed by addition of 2 to 4 ml of DNA in the same electrolyte solution. Frequency and dissipation signals were recorded automatically by QSoft301. For NOM-coated silica surface experiments, there were eight steps: MilliQ water equilibration, HEPES buffer equilibration, PLL coating, HEPES equilibration, 10 mM NaCl equilibration, NOM coating, electrolyte solution equilibration, and DNA adsorption. Each step was held until ΔD and ΔF stabilized. The QCM-D experiments were performed at 100 and 200 mM Na+ and 1 mM Ca2+ as prior work (20-22) suggested that these conditions would result in similar adsorption rates but different adsorbed DNA conformations.
To confirm that our experiments were, as intended, testing the adsorption of DNA to NOM rather than to the intermediate layer of PLL, we tested DNA adsorption separately on a PLL-coated surface and on a surface coated with PLL and then NOM in 1 mM NaCl solution.
Selective experiments were also conducted to investigate desorption (i.e., adsorption reversibility) of DNA. For these experiments, immediate adsorption steps were followed by rinsing with DNA-free electrolyte solutions at the same salt concentrations as for the DNA adsorption solutions. No changes in F and D signals were observed for the rinsing step. Thus, DNA adsorption is irreversible. This observation is consistent with previous studies on DNA adsorption and reversibility (20-22).
The thickness and viscosity of the PLL and NOM coatings and the adsorbed DNA layer were estimated by fitting the change in frequency and dissipation of at least two different overtones to the Voigt model, as described by Voinova et al. (33). The fitting curves were similar among different combination of overtones (i.e., n = 3 and 5; n = 3 and 7; n = 5 and 7; n = 3, 5, and 7). The fitting parameters were forced to be between the following minimal and maximal values, respectively: shear viscosity, 0.001 and 0.01 kg/ms; shear modulus, 1,000 and 3 × 106 Pa; and thickness, 0.1 and 10 nm. Fitting was conducted separately for the PLL layer, the NOM layer, and the DNA layer, with the frequency and dissipation change reset to zero at the beginning of each layer.
To induce competence, DJ77 was grown on BN plates without molybdenum at 30°C for 2 days, inoculated into BN liquid, and grown at 30°C with shaking (170 rpm) for 18 to 20 h (D. R. Dean, personal communication). Existing transformation protocols for A. vinelandii were not appropriate for this work because they used buffers containing divalent cations, typically 1 mM Ca2+ or Mg2+. However, we observed detectable transformation frequencies with cation-free MOPS buffer at pH 7.2 (data not shown), as well as after washing competent cells twice to remove Ca2+ and Mg2+ present in the growth medium; so cations were omitted from the base transformation buffer, and washed cells were used in all experiments presented here. Transformation mixtures contained 200 μl of competent cell suspension, 200 μl of MOPS buffer with the specified salt additions, and various volumes of dissolved or adsorbed chromosomal DNA (target mass of 2 μg of DNA). Transformations were carried out for 20 to 35 min at room temperature.
Transformation mixtures were then diluted with sterile phosphate buffer (0.16 M KH2PO4 and 0.51 M K2HPO4 dissolved in DI water) and spread onto B and BN plates. The plates were incubated at 30°C for 3 to 5 days. Transformation frequencies were calculated by dividing the number of transformants (colonies on B plates) by the number of viable cells (colonies on BN plates) in the transformation mixture.
For statistical analysis of transformation frequencies, Bartlett's test was first used to test homogeneity of variances. No significant difference was found between the variances of transformation frequencies of dissolved DNA and DNA adsorbed on silica surfaces, but transformation frequencies of DNA adsorbed on NOM surfaces had different variances. A two-sample t test was therefore used to compare the transformation frequencies from dissolved DNA and DNA adsorbed on silica, while an unequal variance t test was used for comparison of transformation frequencies of DNA adsorbed on NOM surfaces. A P value of 0.05 was considered significant.
According to the classic double-layer theory, in electrolyte solution the ion layer adjacent to a charged solid surface is immobile, and this is called the compact layer, while the counterions outside the compact layer are mobile. The electrostatic potential at the boundary between the compact layer and its outside layer is defined as the zeta potential and provides surface charge characteristics (10). The zeta potentials for individual transformation assay components and for beads from each step in the bead-coating procedure are shown in Table Table1.1. The silica beads, the chromosomal DNA, and the competent cells all had negative zeta potentials in both electrolyte solutions. The zeta potential results from the coated beads supported sufficient surface coverage as the PLL coating resulted in a positive zeta potential of 6.4 and 19.2 mV, and the NOM coating restored a negative zeta potential of −31.2 and −23.4 mV (Table (Table11).
A representative DNA adsorption experiment on a silica surface is shown in Fig. Fig.11 a. The decrease in frequency and increase in dissipation in step C are due to DNA adsorption. For comparison among experiments, the shift in dissipation (ΔD3) is plotted versus the shift in frequency (ΔF3) for the DNA adsorption step (Fig. (Fig.1b).1b). The slope of ΔD3/ΔF3 decreased from 100 mM Na+ to 200 mM Na+ and was lowest for experiments in 1 mM Ca2+. Previous studies have shown that a higher ΔD/ΔF ratio indicates a more liquid-like adsorbed layer with lower viscosity because more liquid-like layers damp more rapidly and thus have a higher ΔD (7, 20-22). Results from the Voigt model, which combines data from multiple overtones, were consistent with ΔD/ΔF analysis. On silica surfaces, the viscosity of adsorbed DNA in the presence of 100 mM Na+ was 1.24 × 10−3 Pa·s, lower than the 1.44 × 10−3 Pa·s in 200 mM Na+ and the 1.45 × 10−3 Pa·s in 1 mM Ca2+ (Fig. (Fig.22 ).
We first describe the results for the DNA adsorption experiments on PLL-coated surface and on NOM-coated surface in 1 mM NaCl solution. DNA adsorption on the PLL surface resulted in a drop of more than 10 Hz in vibration frequency. In contrast, no measurable frequency shift was observed for the NOM layer, indicating no detectable adsorption. These results showed that the NOM was completely covering the PLL layer; otherwise, the attractive interaction between the positively charged PLL layer and the negatively charged DNA molecules would have produced a significant frequency change.
The conformation of DNA adsorbed to NOM was investigated using the same approach described above for a silica surface, except that creating the NOM coating added several steps. Adsorption of DNA on the NOM surface occurred in step H (Fig. (Fig.33 a); the adsorption data are also plotted for different solution chemistries in Fig. Fig.3b.3b. In contrast to the results with a silica surface, the DNA layer adsorbed on NOM surfaces had the highest slope of ΔD3/ΔF3 in the presence of 1 mM Ca2+, followed by 200 mM Na+ and then 100 mM Na+.
A much higher shift in frequency and dissipation was observed when DNA was adsorbed to an NOM-coated surface in 1 mM Ca2+ than to this surface in Na+. To control for the difference in mass adsorbed, we conducted another Ca2+ experiment with a lower concentration of DNA. The ΔD3/ΔF3 ratio was still higher than that in Na+ solution (Fig. (Fig.3b).3b). Thus, the observed higher ratio of ΔD3/ΔF3 was not solely due to greater DNA adsorption in the presence of Ca2+.
Results from the Voigt model were again consistent with the ΔD3/ΔF3 analysis (Fig. (Fig.2).2). On NOM surfaces, the viscosity of adsorbed DNA in 100 mM Na+ solution was the highest, i.e., 2.39 × 10−3 Pa·s, with values of 1.84 × 10−3 Pa·s in 200 mM Na+ solution and 1.30 × 10−3 Pa·s in 1 mM Ca2+ solution. The viscosity of the layer formed by a lower DNA concentration in 1 mM Ca2+ solution was slightly higher, 1.46 × 10−3 Pa·s, but still less than that of the DNA layers formed in the presence of Na+.
For the transformation assays, we used DNA adsorbed on uncoated silica beads and NOM-coated silica beads. The amount of DNA adsorbed on silica and NOM-coated beads was between 4.4 to 7.0 μg/ml. High background of the NOM in 100 mM Na+ solution prevented accurate measurement of the DNA adsorbed under those conditions. In general, more DNA adsorbed in the 1 mM Ca2+ solution than in 200 mM Na+. The amount of the beads added to different transformation assays was therefore adjusted to maintain a consistent amount of adsorbed DNA.
To investigate the effects of cations on natural transformation, A. vinelandii transformations with DNA dissolved in MOPS buffer amended with nothing, 200 mM Na+, 100 mM Na+, or 1 mM Ca2+ were performed (Fig. (Fig.4).4). In the presence of 200 mM Na+, the viable cell counts were 10 to 100 times lower than in the presence of 100 mM Na+ or 1 mM Ca2+. Because of this toxicity, we have omitted the results of these transformations. DNA dissolved in MOPS buffer transformed A. vinelandii at an average frequency of 2 × 10−5. The addition of either 100 mM Na+ or 1 mM Ca2+ slightly increased transformation frequencies, to around 6 × 10−5 and 7 × 10−5, respectively. Based on these results we concluded that comparisons between these two salt concentrations were valid.
Adsorbed DNA did not significantly influence transformation frequencies compared to frequencies with dissolved DNA (Fig. (Fig.4).4). Furthermore, despite differences in the conformation of the silica-adsorbed DNA in the presence of Na+ and Ca2+, its transformation frequencies in the presence of 100 mM Na+ and 1 mM Ca2+ were not significantly different. For NOM surfaces, the adsorbed DNA also gave transformation frequencies similar to those of dissolved DNA, but in this case the transformation frequencies were higher in 1 mM Ca2+ than in 100 mM Na+, and the difference was confirmed to be significant by an unpaired t test. Although the concentration of DNA in our transformation assays was selected such that DNA should not be limiting, we wanted to test whether this difference was due to lower DNA adsorption in the presence of Na+, so we repeated the Ca2+ transformations with less DNA. Decreasing the amount of DNA still resulted in a transformation frequency that was higher than in the presence of 100 mM Na+ (Fig. (Fig.4)4) and not significantly different from the previous Ca2+ tests, based on a two-sample t test (95% confidence level). For DNA adsorbed to NOM, the transformation efficiency was consistently higher in the presence of Ca2+ than in Na+.
Extracellular DNA adsorption to soil surfaces is of concern because adsorption protects the DNA from degradation and increases the potential for horizontal gene transfer. Several studies have confirmed the availability of adsorbed DNA for transformation, although usually at a reduced transformation frequency (4-6, 11, 15, 23, 25, 28). Notable exceptions include the work of Lorenz and Wackernagel (18, 19), where transformation frequencies were higher for DNA adsorbed to sand than for dissolved DNA. Our results showed that adsorbed chromosomal DNA is also able to transform A. vinelandii. Strikingly, in the present work the transformation frequencies were similar for dissolved and adsorbed DNA. There are several possible explanations for this result, including the presence of excess DNA, full availability of adsorbed DNA, or effects of the particles on cells. For A. vinelandii, transformation frequencies level off around 1 μg of donor DNA (9); our experiments were conducted with 2 μg. The presence of excess DNA could theoretically obscure any decrease in availability of DNA when it is adsorbed. However, this explanation appears less likely, based on prior reports. One-microgram DNA saturating concentrations have also been reported for B. subtilis (1). Despite the use of more than 10 times excess DNA, decreased transformation frequencies have been reported for B. subtilis with adsorbed DNA (6, 15). The similar transformation frequencies we observed for dissolved and adsorbed DNA could also suggest that adsorbed DNA is fully available to A. vinelandii, in which case the discrepancy with prior reports could be attributed to the use of different model organisms. Alternatively, adsorption of cells may serve to increase the effective concentration of cells and DNA and compensate for reduced availability of adsorbed DNA.
The conformation of adsorbed chromosomal DNA on a silica surface is increasingly compact and rigid in the presence of 200 mM Na+, 100 mM Na+, and 1 mM Ca2+, as demonstrated here by the QCM-D results. The results on silica surfaces are consistent with those previously observed for plasmid DNA adsorption on both silica and NOM-coated surfaces (20, 21). The presence of Na+ allows charge shielding and the compression of the electrostatic double layers surrounding the negatively charged DNA molecules (10), and this compression increased as the concentration of Na+ increased from 100 to 200 mM. In Ca2+ solutions, however, Ca2+ forms inner-sphere complexation with both DNA phosphate groups and NOM carboxylate groups and thus allows the formation of cation bridging between DNA and NOM molecules and within different regions of a DNA molecule (14, 20). Compared to charge shielding by Na+, inner-sphere complexation by Ca2+ is expected to form a more rigid adsorbed DNA layer, as observed on a silica surface.
Surprisingly, on NOM surfaces the trend was reversed; the DNA-adsorbed layer was more compact and rigid in the presence of a 100 mM Na+ solution than in a 200 mM Na+ solution. The compactness and rigidity of the DNA-adsorbed layer in a 1 mM Ca2+ solution were lower than in Na+ solutions. Although no previous data are available for the specific combination of DNA, NOM, and Ca2+ tested here, there is some precedent for this observation, as the adsorbed alginate layer formed in 100 mM Na+ solution is less fluidic and more rigid than the layer formed in 1 mM Ca2+ (7). de Kerchove and Elimelech (7) suggested that Ca2+ complexation with carboxylate groups of alginate allows the formation of gel-like adsorbed alginate layers and that water molecules are trapped within these layers. Similarly, complexation of Ca2+ with both DNA phosphate groups and the carboxylic groups on NOM surfaces may allow the formation of a gel-like and fluidic DNA adsorbed layer.
In most cases, changes in the conformation of adsorbed DNA had no effect on the transformation frequency. The exception was on NOM surfaces, where transformation frequencies were consistently higher in the presence of Ca2+. The higher transformation frequency with NOM and Ca2+ was probably not due to the change in viscosity of the adsorbed DNA layer because the change in viscosity for NOM-adsorbed DNA layers was less than that observed for silica, where the conformational changes had no effect on transformation frequency. Because the experiments with NOM and Ca2+ had the most DNA adsorption, we also considered the possibility that the higher transformation frequency in the presence of Ca2+ was an artifact of increased DNA concentration, but lowering the concentration of DNA did not reduce the transformation efficiency. However, even in the experiments with a lower concentration of DNA, the NOM/Ca2+ experiments had a higher DNA density on individual beads, so density effects could explain the slightly higher transformation frequency observed in the presence of Ca2+. Alternatively, it could be due to a three-way interaction between the cells, Ca2+, and NOM.
In the broader context of the fate of extracellular DNA in the soil and particularly with respect to its contributions to horizontal gene transfer, this study adds A. vinelandii to the list of microorganisms that can be transformed by adsorbed DNA. Furthermore, when considered in the context of prior reports of transformation frequencies that varied over 2 to 4 orders of magnitude (18), these results suggest that the solution chemistry, the type of surface, and the conformation of the adsorbed DNA will not have a strong influence on the transformation frequency and can be neglected in modeling the transformation. However, the solution chemistry and the type of surface will influence the amount of DNA adsorbed and, thus, indirectly the transformation frequency, particularly for low concentrations of extracellular DNA.
This research work was supported by the USDA (grant 2008-35102-19143) and WaterCAMPWS, a Science and Technology Center of Advanced Materials for the Purification of Water with Systems under National Science Foundation agreement number CTS-0120978.
We thank Dennis Dean for providing A. vinelandii strains and protocols and anonymous reviewers for constructive comments.
Experiments, data interpretation, and manuscript preparation were conducted by N. Lu with supervision by T. H. Nguyen and J. L. Zilles.
Published ahead of print on 7 May 2010.