Adsorption Kinetics and Reversibility of Linear Plasmid DNA on Silica

Apr 7, 2010 - Baltimore, Maryland 21218, and Department of Chemical Engineering, Environmental .... University of Illinois at Urbana-Champaign...
0 downloads 0 Views 856KB Size
Download PDF
Biomacromolecules 2010, 11, 1225–1230

1225

Adsorption Kinetics and Reversibility of Linear Plasmid DNA on Silica Surfaces: Influence of Alkaline Earth and Transition Metal Ions Thanh H. Nguyen,*,† Kai Loon Chen,‡ and Menachem Elimelech§ Department of Civil and Environmental Engineering, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, Department of Geography and Environmental Engineering, Johns Hopkins University, Baltimore, Maryland 21218, and Department of Chemical Engineering, Environmental Engineering Program, Yale University, New Haven, Connecticut 06520 Received December 14, 2009; Revised Manuscript Received March 19, 2010

A quartz crystal microbalance with dissipation monitoring is used to study the adsorption of linear plasmid DNA on silica surfaces and silica surfaces coated with poly-L-lysine (PLL) in solutions containing either alkaline earth (calcium and magnesium) or transition (cobalt, copper, and zinc) metals. Our results show that electrostatic attraction alone does not fully explain the significantly higher adsorption rate of DNA on the positively charged PLL layer in Cu2+ solution compared to solutions containing Ca2+, Mg2+, Co2+, or Zn2+. Diffusion coefficients measured by dynamic light scattering reveal that the compactness of plasmid DNA molecules is greater in solutions containing Cu2+ compared to that of DNA in other divalent electrolyte solutions. When the adsorption rate of plasmid DNA on silica is normalized to the corresponding adsorption rate on PLL-coated surfaces at the same solution condition, the attachment (adsorption) efficiencies are about 0.01 for Ca2+ or Mg2, but close to unity for Co2+, Cu2+, or Zn2+. Results from viscoelastic modeling of adsorbed DNA layers suggest that the DNA layer formed in Cu2+ solutions is thicker and more viscous compared to that formed in Co2+ solutions. This study demonstrates that plasmid DNA has a strong affinity to Cu2+, which results in a more compact conformation of DNA molecules compared to the case with the other divalent cations investigated.

Introduction DNA interaction with metal ions, such as alkaline and alkaline earth cations, plays a critical role in biology.1 Adequate supply of these metals allows biological cells to function properly, while an excess of these metals may lead to various diseases, such as Wilson’s disease.1 Toxicity of transition and heavy metals may be due to their strong complexation with DNA molecules.2,3 A number of studies have been designed to investigate the mechanisms of DNA interaction with metal ions.1,2,4-9 While alkaline metals such as Na+ and K+ do not bind to DNA molecules, Mg2+ and Ca2+ cations can bind to the DNA backbone via outer sphere and inner sphere complexation, respectively.2,8 Compared to Ca2+ and transition metals, Mg2+ has a more tightly held hydration sphere which prevents it from binding directly to DNA phosphate backbone.2,8,10-12 Similar to Ca2+, transition metal ions, such as Co2+, Cu2+, and Zn2+, bind to the DNA phosphate backbone by inner sphere complexation.13-15 In solutions with Cu2+ concentrations higher than 10 mM, Cu2+ ions can also bind directly to DNA bases and disrupt the base-pair hydrogen bonds between two DNA strands.14 DNA immobilization and adsorption on surfaces have direct applications for biosensing,16-18 DNA purification,19,20 and nanofabrication.21 A better understanding of DNA adsorption behavior will also allow for the prediction of environmental fate and transport of extracellular DNA.22 Studies on DNA adsorption range from simple batch experiments with soils to spectroscopic investigations with well-defined surfaces.4,16,23-33 * To whom correspondence should be addressed. Phone: 217-244-5965. Fax: 217-333-6968. E-mail: [email protected]. † University of Illinois at Urbana-Champaign. ‡ Johns Hopkins University. § Yale University.

Results from some studies employing Fourier transform infrared (FTIR) spectroscopy have suggested that DNA compactness changes upon adsorption to clay surfaces.30,31 The role of monovalent and divalent salts on DNA adsorption to silica and natural organic matter (NOM)-coated silica surfaces has also been studied using a quartz crystal microbalance with dissipation monitoring (QCM-D), which allows for real-time monitoring of DNA adsorption and conformational changes of adsorbed DNA molecules.23-25,29 For example, the hybridization of single-stranded DNA adsorbed on polycationic film was detected and studied using the QCM-D technique.29 Atomic force microscopy studies suggest that DNA bind strongly to mica surfaces in the presence of transition metals, such as Co2+, Ni2+, and Zn2+, but weakly to mica surfaces in the presence of other divalent cations, such as Ca2+ and Mg2+.34-37 Charge neutralization of both the mica surface and the DNA molecules by Ni2+ has been implicated to explain the strong adsorption of DNA on mica surfaces.35 More recent studies using long-period X-ray standing waves technique6,38 suggested that, in solution with low concentration of ZnCl2 (i.e., 50 µM), Zn2+ cations react with weakly charged hydroxyl groups on silica surface. As a result, the repulsion between negatively charged synthetic RNA molecules and silica surface is reduced to allow RNA adsorption on the hydroxyl-terminated silica surfaces.6,38 Previous studies on DNA adsorption were mostly performed using a limited range of solution chemistry. Studies to quantify DNA adsorption rates in solutions containing alkaline earth or transition metals and to determine the relationship between the compactness of adsorbed DNA layers and cation type are lacking. The objective of this paper is to investigate the role of alkaline earth cations (Ca2+ and Mg2+) and transition divalent metal

10.1021/bm901427n  2010 American Chemical Society Published on Web 04/07/2010

1226

Biomacromolecules, Vol. 11, No. 5, 2010

Nguyen et al.

cations (Co2+, Cu2+, and Zn2+) on the adsorption behavior of linear plasmid DNA on bare and PLL-modified silica surfaces. Two complementary techniques, QCM-D and dynamic light scattering (DLS), are used to probe the change in compactness of DNA molecules in solution and DNA adsorbed on silica surfaces. Reversibility of DNA adsorption is studied by exposing the adsorbed DNA molecules to solutions of low ionic strength or solutions containing a strong chelating agent, ethylenediaminetetraacetic acid (EDTA).

Materials and Methods Preparation of Linear Plasmid DNA. The ampicillin-resistant plasmid vector pGEM-Teasy (3015 bp) was used for this study. The protocol for plasmid DNA extraction and purification from E. coli XL1 blue strains was described in our previous publications.23-25 Briefly, we used a Qiagen EndoFree Plasmid Giga kit (Qiagen Inc. CA) to extract plasmid DNA from an E. coli culture that was grown overnight. After purification, as recommended by the manufacturer, the plasmid DNA was precipitated with isopropanol and washed with 70% ethanol, and the precipitated DNA was dissolved in 20 mL of RNase-free and endotoxin-free water from American Bioanalytical (Natick, MA). This 20 mL DNA solution was further digested with enzyme Nsi I (New England Biolabs, Inc., Beverly, MA) to obtain linear plasmid DNA. The completion of digestion was verified using agarose gel. The linearized DNA solution was purified using the Qiagen-tip 10000s following the protocol recommended by the manufacturer. The purified linear plasmid DNA was precipitated with isopropanol, subsequently washed with 70% ethanol and then dissolved in 20 mL of RNase-free and endotoxin-free water. The final DNA solution was further divided into 200 µL aliquots and stored at -20 °C until use. We used gel electrophoresis to check the integrity of the plasmid DNA and UV spectroscopy (260 nm wavelength) to measure its concentration.25 Dynamic Light Scattering (DLS) Measurement of Plasmid DNA Diffusion Coefficients. The diffusion coefficients of linear plasmid DNA prepared in different solution chemistries were determined by DLS. The DLS measurements were conducted using a multidetector light scattering unit (ALV-5000, Langen, Germany), which utilizes a Nd:vanadate laser (Verdi V2, Coherent, Santa Clara, CA) with a wavelength of 532 nm. Details on the unit, sample preparation, and experimental procedure for DLS measurements are provided in our earlier publications.23-25 Briefly, 20 DLS measurements were conducted for each solution composition at a scattering angle of 90°. The decay of the autocorrelation function obtained from each DLS measurement was fitted with a biexponential function. This fitting procedure yields a slow relaxation time that is subsequently used to derive the diffusion coefficient of the DNA.23,39,40 Due to possible dependence of the diffusion coefficient on DLS scattering angle,39,40 the derived diffusion coefficients at a fixed angle (90°) can be viewed as approximations of the DNA translational diffusion coefficients. Electrophoretic Mobility of Plasmid DNA. We used a Zetasizer Nano ZS (Malvern Instruments, Southborough, MA) to measure electrophoretic mobilities of plasmid DNA in solution chemistries similar to those employed for the adsorption studies. The solution chemistries were 10 mM NaCl, 1 mM CaCl2, 1 mM MgCl2, 1 mM CoCl2, 1 mM CuCl2, and 1 mM ZnCl2. Plasmid DNA concentrations used for the electrophoretic mobility measurements were 220 mg/L, and the temperature was 25 °C. All solutions were unbuffered with a solution pH of 5.6. Quartz Crystal Microbalance with Dissipation Monitoring (QCM-D) Measurements. A QCM-D D300 system (Q-Sense AB, Sweden) connected to a syringe pump operating in withdrawing mode at 0.1 mL/min flow rate was used to study plasmid DNA adsorption. Quartz sensors coated with SiO2 were used for all measurements. For adsorption experiments performed on PLL-coated silica surfaces, the SiO2 surface was precoated with a layer of PLL by flowing 2 mL of a PLL hydrobromide solution (0.1 g/L of PLL hydrobrimide in HEPES

Figure 1. Diffusion coefficients of linear plasmid DNA in monovalent and divalent electrolyte solutions in the absence and presence of 1 mM EDTA. All measurements are conducted at an electrolyte concentration of 1 mM and unadjusted pH of 5.6. Plasmid concentration employed for the measurements is 220 mg/L and the temperature is 25 °C.

buffer) through the sensor chamber.23-25,41 This PLL adsorption step was followed by 40 min rinsing with HEPES buffer. The PLL layer was then equilibrated for 20 min with the same electrolyte solution to be used for the subsequent DNA adsorption experiment. For adsorption experiments on silica surfaces, the system was left to equilibrate for at least 20 min with the same electrolyte solution to be used for the subsequent DNA adsorption experiment. Before each QCM-D experiment, the sensors were soaked in 2% Hellmanex II solution (Hellma GmbH & Co KG, Germany) for 2 h, then rinsed with deionized (DI) water, dried with ultra-high-purity nitrogen, and treated for 30 min in a UV/O3 chamber. The sensors for adsorption on silica surface were only used 5 times. The DNA adsorption rate was estimated from the QCM-D measurements using the initial slope of the change in frequency monitored at the third overtone, f(3), versus time curve, as described in our previous studies.23-25,42 Reversibility of DNA Adsorption. The adsorption step of DNA on the silica surface was followed by three rinsing steps to study adsorption reversibility. The adsorbed DNA layer was rinsed with a divalent electrolyte solution similar to that used during adsorption (step 2), 1 mM NaCl solution (step 3), and DI water (step 4). For the two adsorption experiments performed in solutions containing 1 mM Co2+ and Cu2+, 1 mM EDTA solution at unadjusted pH 5.6 was used to rinse the adsorbed DNA layers to better understand the role of divalent cations in adsorption kinetics and reversibility. We conducted control experiments and verified that the shifts in frequency and dissipation signals when DNA layers formed in 1 mM Co2+ or 1 mM Cu2+ were exposed to 1 mM EDTA solutions were much higher than the signal shifts due to buffer effects.43 Specifically, when the solution in contact with a bare QCM-D sensor was switched from 1 mM Cu2+ to 1 mM EDTA, the frequency shift for the third overtone and the dissipation shift were 8.9 Hz and 0.2 × 10-6, respectively. This shift is much smaller than the shift observed when the adsorbed DNA layer was exposed to 1 mM EDTA (i.e., the frequency and dissipation shifts were 61 Hz and 25 × 10-6, respectively). For the case of Co2+, when the divalent electrolyte solution was switched to 1 mM EDTA solution, the corresponding frequency and dissipation shifts were 1 Hz and 0.03 × 10-6 without the DNA adsorbed layer and 12 Hz and 0.27 × 10-6 with the DNA adsorbed layer.

Results and Discussion Influence of Divalent Cations on Plasmid DNA Diffusion Coefficients. Figure 1 shows the diffusion coefficients of linear plasmid DNA in solutions containing 1 mM of monovalent and divalent cations, both in the absence and presence of 1 mM

Linear Plasmid DNA on Silica Surfaces

EDTA. In the absence of EDTA, the diffusion coefficient is smallest in the presence of NaCl. At such low ionic strength conditions (1 mM), charge screening effects are insignificant. The negatively charged functional groups along the DNA molecules repel each other, causing the DNA to take an extended conformation and thus resulting in a small diffusion coefficient. The diffusion coefficients in the presence of divalent cations are generally larger. Divalent cations bind specifically to the DNA phosphate backbone, resulting in charge neutralization and hence a more compact conformation of the DNA molecules. This binding effect seems to be especially strong in the presence of Cu2+ ions, as reflected by the high diffusion coefficient. Results from different spectroscopic studies suggest that depending on Cu2+ concentration, Cu2+ could bind strongly to both the phosphate backbone and bases of DNA moleules.9,14,44-46 In solution containing more than 10 mM Cu2+, the Cu2+ ions bind to the DNA bases, resulting in breakage of some of the hydrogen bonds and a change in DNA molecule compactness.14 The binding mechanisms of Cu2+ to DNA phosphate backbone and bases depend on DNA sequence and Cu2+ concentrations.14 Hackl et al.45 and Andrushchenko et al.14 studied Cu2+ binding to DNA over a wide range of concentrations (10-4-10-1 M) by infrared (IR) and vibrational circular dichroism (VCD) spectroscopy. At intermediate and high Cu2+ concentrations (>10-2 M), the increase in the intensity of DNA absorption band in IR spectra was suggested to be a result of Cu2+-induced compaction of the DNA molecules due to Cu2+ binding to DNA bases.45 In another study, VCD spectra showed evidence of Cu2+ binding to DNA bases at a Cu2+ concentration of 0.4 M.14 The same study reported that the low concentration of Cu2+ used in our study (1 mM) does not result in structural distortion due to Cu2+ binding to DNA bases.14 Similar to the observations in the IR and VCD spectroscopy studies,14,45 Raman spectroscopy also showed that among the 10 divalent cations studied, Cu2+ has the strongest affinity to bind to the phosphate backbone.44 In solutions containing both Cu2+ and EDTA, the diffusion coefficient of plasmid DNA is significantly lower than in the absence of EDTA. This observation suggests that EDTA acts as an effective chelating agent to prevent Cu2+ from binding to DNA molecules and thus allow the DNA molecules to have a less compact conformation. However, in contrast to our expectation, the presence of EDTA did not change the diffusion coefficients of plasmid DNA in solutions containing either Co2+ or Zn2+. It is likely that the binding of Co2+ and Zn2+ to the phosphate backbone of plasmid DNA molecules does not produce a substantial change in compactness of plasmid DNA in the first place, as is evident from the diffusion coefficients that are only slightly larger than that in Na+. Note that the metal-ligand stability constants (log K) for EDTA with Co2+, Cu2+, and Zn2+ are 16.21, 18.8, and 16.5, respectively.47 Thus, the effect of EDTA in complexing metal ions is more pronounced in the Cu2+-DNA solution than in Co2+- or Zn2+-DNA solution. Electrophoretic Mobilities of Plasmid DNA. Plasmid DNA molecules have the most and the least negative values of electrophoretic mobility in solutions of 10 mM NaCl and 1 mM CuCl2, respectively (Figure 2). Similar electrophoretic mobilities are observed in 1 mM of the other divalent salt solutions (CaCl2, MgCl2, CoCl2, or ZnCl2). While the divalent cations have the ability to bind to DNA phosphate backbone and neutralize the negative charges of DNA molecules, monovalent cations (i.e., Na+) can only screen the charges. As a result, plasmid DNA molecules in monovalent electrolyte solutions are more negatively charged than in divalent electrolytes at the present experimental conditions.

Biomacromolecules, Vol. 11, No. 5, 2010

1227

Figure 2. Average electrophoretic mobilities of plasmid DNA in monovalent and divalent electrolyte solutions at unadjusted pH 5.6. Shown are the averages of six measurements for each solution. Error bars indicate standard deviations. Plasmid concentration employed for the measurements is 220 mg/L and the temperature is 25 °C.

Adsorption Kinetics of Plasmid DNA on PLL-Coated Silica Surfaces. We first measured plasmid DNA adsorption kinetics on silica surfaces coated with PLL. At pH 5.6 (unadjusted pH), the PLL molecules are positively charged while the DNA molecules are negatively charged. The interactions between the DNA and PLL layer at this pH are attractive due to attractive electrostatic and van der Waals interactions. As shown in Figure 3a, the adsorption rates are comparable in solutions containing Ca2+, Mg2+, Co2+, and Zn2+. However, in solutions containing Cu2+, the adsorption rate increased 4-fold. This dramatic increase is likely related to the marked change in DNA compactness due to strong binding of Cu2+ to the DNA phosphate backbone, which is consistent with the findings of other Raman, IR, and VCD spectroscopy studies.14,44,45 The DLS data suggests a significant size reduction of the DNA molecules in Cu2+ solution. As the DNA molecules in Cu2+ become more compact, the rate of DNA adsorption onto the positively charged PLL layer increases due to higher convectivediffusive transport toward the PLL-coated silica surface. Adsorption Kinetics of Plasmid DNA on Bare Silica Surfaces. Under the solution conditions tested (pH 5.6), both silica surfaces and DNA are negatively charged, thereby resulting in electrostatic repulsion. The attachment (adsorption) efficiency of DNA on silica (Figure 3b) is obtained by normalizing the actual plasmid adsorption rate on bare silica surfaces (i.e., in the presence of repulsive interactions) to the corresponding adsorption rate on PLL-coated silica surfaces (i.e., in the presence of attractive interactions) obtained at the same solution chemistry.48 This normalization eliminates the effect of different rates of DNA transport to silica surface due to changes in compactness and allows for the isolation of the influence of solution chemistry on DNA adsorption kinetics. For plasmid DNA in solutions containing Ca2+ or Mg2+, the attachment efficiencies are around 0.01. In solutions containing Co2+, Cu2+, or Zn2+, the attachment efficiencies approach the maximum value of 1.0. A surface reaction followed by adsorption has been proposed for polynucleotide adsorption to silica surfaces in solutions containing zinc.6,38 The silanol groups on silica surfaces can react with Zn2+ according to the following reaction:

≡SiOH + M2+ + H2O ) ≡SiOM+ + H3O+

(1)

1228

Biomacromolecules, Vol. 11, No. 5, 2010

Figure 3. (a) Adsorption rates of plasmid DNA on silica surfaces coated with PLL in divalent electrolytes at unadjusted pH 5.6. (b) Attachment (adsorption) efficiencies of plasmid DNA on bare silica surfaces. Shown in (a) and (b) are average values of at least two replicate measurements. Error bars indicate standard deviations. Plasmid concentration employed for the experiments is 220 mg/L and the temperature is 25 °C. Attachment efficiency was calculated by dividing the average adsorption rate on silica surface by the average adsorption rate on PLL-coated surface obtained at the same solution chemistry.

where M represents a metal (including Zn). This reaction reduces the overall negative charge of the silica surface and, as a result, promotes the adsorption of polynucleotides. The log association constants (pK ) -log K) of eq 1 are on the order of 7.3 for Ca2+, 8.1 for Mg2+, and 5.5 for Cu2+ ions.49 Lower association constants for Mg2+ and Ca2+ compared to Cu2+ suggest a weaker affinity of Mg2+ or Ca2+ to react with the silanol groups, thus, resulting in the silica surfaces to be more negatively charged in Mg2+ or Ca2+ compared to the ones in Cu2+ solutions. In addition, Cu2+ has a higher affinity to DNA phosphate backbone compared to Ca2+ or Mg2+ and is more efficient in neutralizing the charges on DNA molecules as shown by the electrophoretic mobility data (Figure 2). The combination of these two factors leads to a higher attachment efficiency in Cu2+ solutions. Strong binding of Cu2+ to the phosphate backbone has also been shown in previous experimental and theoretical studies.13-15 It is possible that the energy barrier between DNA molecules and silica surface vanishes in Cu2+ solutions, giving rise to the observed maximum attachment efficiency. Experimental data for pK values of cation reaction with amorphous SiO2 have been reported only for Ca2+ (7.3), Mg2+ (8.1), Cu2+ (5.5), Cd2+ (6.1), and Pb2+ (5.1).49 Cheng et al.6 and Libera et al.38 suggested that the pK value for the reaction between silanol groups and Zn2+ is slightly higher than that for Cu2+. As a result, Zn2+ are expected to bind to the silanol groups quite strongly (while not as strong as Cu2+) to reduce the

Nguyen et al.

Figure 4. Normalized frequency shifts and associated dissipation shifts as a function of time at unadjusted pH 5.6. Plasmid DNA is first adsorbed on the silica surface in 1 mM divalent electrolyte solution (step 1). The adsorbed plasmid layer is then rinsed with 1 mM of the same divalent electrolyte solution (step 2), 1 mM NaCl (step 3), and DI water (step 4). Plasmid concentration employed for the experiments is 220 mg/L and the temperature is 25 °C.

negative charge of the silica surface. The higher reactivity of Zn2+ and Cu2+ with silica surfaces compared to Ca2+ and Mg2+ leads to higher attachment efficiency for the three transitional metals. Detachment (Reversibility) of Adsorbed DNA. The reversibility of DNA adsorption was studied for the cases where DNA adsorbs to silica in the presence of transition metals (Figures 4 and 5). Detachment of adsorbed DNA is assessed by monitoring both the frequency and dissipation responses with the QCM-D when the adsorbed DNA layers were rinsed sequentially with 1 mM of the corresponding divalent electrolyte solution (step 2), 1 mM NaCl (step 3), and DI water (step 4), as shown in Figure 4, or with 1 mM EDTA solution, as shown in Figure 5. For adsorbed DNA layers formed in the presence of Co2+ or Zn2+, no change in frequency signals but a small increase in dissipation signals was observed during the rinsing steps (Figure 4). These observations suggest that no detachment occurred and the adsorbed DNA layers become slightly less compact when the layers were rinsed with low ionic strength solutions. When plasmid DNA adsorbed on the silica surface in the presence of Cu2+, a substantial decrease in frequency and an increase in dissipation were observed (step 1; Figure 4). These observations indicate the formation of a DNA adsorbed layer. However, the adsorbed DNA layer appeared to be unstable with slightly increased frequency and decreased dissipation. When the adsorbed layer was rinsed with Cu2+ solution (step 2), the frequency continued to increase while the dissipation continued

Linear Plasmid DNA on Silica Surfaces

Figure 5. Normalized frequency shifts and associated dissipation shifts as a function of time at unadjusted pH 5.6. Plasmid DNA is first adsorbed on the silica surface in 1 mM (a) CuCl2 and (b) CoCl2 solution. The adsorbed plasmid layer is then rinsed with 1 mM EDTA. Plasmid concentration employed for the experiments is 220 mg/L and the temperature is 25 °C.

to decrease. These changes in frequency and dissipation were probably due to some DNA detachment from the surface. When the rinsing solutions were switched to 1 mM NaCl (step 3) followed by DI water (step 4), the frequency became stable and the dissipation continued to increase, showing that the adsorbed DNA layer became less compact. Note that detachment during the first rinse with divalent cation solution was not observed for the adsorbed DNA layers formed in the presence of Co2+ or Zn2+. It is possible that in the presence of Cu2+, multiple layers of adsorbed DNA were formed but only the top layers were removed during the first rinsing step. A separate set of experiments was conducted using a solution containing 1 mM EDTA at pH 5.6 as the rinsing solution (Figure 5). For the DNA layer formed in Cu2+, rinsing with EDTA solution led to sharp increases in both frequency and dissipation followed by a gradual decrease in frequency and continued increase in dissipation. When the adsorbed DNA layer formed in the presence of Co2+ was washed with EDTA, an immediate increase in frequency and decrease in dissipation followed by a slow decrease in frequency and increase in dissipation was observed. When the data were fitted using the Voigt model,23-25,41,48 the following trends were found. In the presence of Cu2+, a DNA layer of 27 nm thickness with viscosity of 3.3 × 10-3 Pa s and shear modulus of 3.4 × 105 Pa was formed. Within 2.5 min of EDTA rinsing, the thickness decreased to 20 nm, while the viscosity and shear modulus of the adsorbed DNA layer decreased to 1.5 × 10-3 Pa s and 1.1 × 105 Pa, respectively. Within the next 14 min, the thickness increased to 47 nm, while the viscosity and shear modulus further decreased to 1.48 × 10-3 Pa s and 0.62 × 105 Pa, respectively. The initial decrease of thickness and viscosity of the DNA layer during rinsing with EDTA solution suggested partial detachment of the adsorbed

Biomacromolecules, Vol. 11, No. 5, 2010

1229

DNA. The subsequent increase of thickness and decrease of viscosity indicated a more fluidic adsorbed DNA layer. As a strong chelating agent, EDTA removed Cu2+ bound to DNA molecules, causing the DNA molecules to become less compact and allowing water molecules to hydrate the adsorbed DNA layer. For the case of Co2+, the initial DNA layer has a thickness of 10 nm, viscosity of 1.6 × 10-3 Pa s, and shear modulus of 0.9 × 105 Pa. The changes in frequency and dissipation signals observed within the first minute of EDTA rinsing were too small for us to draw any conclusions about any changes in DNA layer properties. From 55.6 to 73.5 min, the DNA layer increased in thickness to 43 nm, while the viscosity and shear modulus decreased to 1.2 × 10-3 Pa s and 0.11 × 105 Pa, respectively. Compared to the DNA layer formed in Cu2+ solution, the layer formed in Co2+ solution is much thinner (10 vs 27 nm) and less viscous (1.6 × 10-3 vs 3.3 × 10-3 Pa s). For adsorbed DNA layers formed in either Co2+ or Cu2+ solution, rinsing with EDTA led to thicker and more fluidic DNA layers. Similar to the case of Cu2+, EDTA removed Co2+ bound to DNA phosphate backbone to allow water molecules to be incorporated into the DNA adsorbed layer. The metal stability constant (log K) for Cu2+ and EDTA is 2 orders of magnitude higher than that for Co2+ and EDTA (18.8 vs 16.2).47 At the same time, Cu2+ also has higher affinity to DNA than Co2+.44 Because EDTA, however, is a much stronger chelating agent compared to DNA, it is likely that the chelating effects of EDTA will overwhelm the one for DNA, thus, resulting in a greater degree of Cu2+ removal from DNA compared to Co2+. As expected, the change in viscosity of the DNA adsorbed layer is more pronounced for DNA layer formed in Cu2+ than in that formed in Co2+ solution (3.3 × 10-3 to 1.48 × 10-3 Pa s vs 1.6 × 10-3 to 1.2 × 10-3 Pa s).

Conclusion Using complementary techniques (QCM-D and DLS), we show that among the five divalent cations studied (Ca2+, Mg2+, Co2+, Cu2+, and Zn2+), the presence of transition metal cations Co2+, Cu2+, or Zn2+ resulted in stronger adsorption of plasmid DNA to silica and smaller diffusion coefficient of plasmid DNA in solution. This observation is attributed to the high binding affinity of transition metal cations to the phosphate backbone of plasmid DNA molecules and silanol groups on silica surfaces. When an EDTA solution was used to rinse the adsorbed layer formed in Cu2+ solution, only partial detachment was observed and the adsorbed layer became thicker and less viscous. Adsorption rate of plasmid DNA on silica surfaces is in agreement with the association constants for the reactions between divalent cations and silanol groups. The presence of transition metals with higher association constants allows for higher adsorption rates than in Ca2+ or Mg2+. We suggest that DNA adsorption involves two steps: reaction of metal cations with silica surfaces to reduce negative charge of the silica surfaces and subsequent adsorption of plasmid DNA on the silica surfaces. This mechanism proposed previously for Zn2+ and a synthetic nucleic acid appears to be consistent with our experimental results for five divalent cations. Acknowledgment. Funding was provided by the Yale Institute for Biospheric Studies and the National Science Foundation (CTS-0120978) and USDA (Grant 2008-3510219143).

References and Notes (1) Anastassopoulou, J.; Theophanides, T. Crit. ReV. Oncol./Hematol. 2002, 42, 79–91.

1230

Biomacromolecules, Vol. 11, No. 5, 2010

(2) Anastassopoulou, J. J. Mol. Struct. 2003, 19–26. (3) Kukushkin, V. Y.; Oskarsson, A.; Elding, L. I.; Farrell, N.; Vicente, J.; Chicote, M. T.; Kauffman, G. B.; Houghten, R. A.; Likins, R. E.; Posson, P. L.; Ray, R. K.; Tkachuk, V. M.; Vorobiov-Desiatovsky, N. V.; Hill, G. S.; Irwin, M. J.; Levy, C. J.; Rendina, L. M.; Puddephatt, R. J.; Romeo, R.; Monsu’scolaro, L.; Ruffo, F.; De Renzi, A.; Panunzi, A.; Byers, P. K.; Canty, A. J.; Jin, H.; Kruis, D.; Markies, B. A.; Boersma, J.; van Koten, G.; Nazeeruddin, K.; Kalyanasundaram, R.; Gratzel, M.; Bessel, C. A.; Leising, R. A.; Szczepura, L. F.; Perez, W. J.; Huyhn, M. H. V.; Takeuchi, K. J.; Poli, R.; Krueger, S. T.; Mattamana, S. P.; Jacobsen, C. J. H.; Klinke, K. K.; Hyldtoft, J.; Villadsen, J.; Abernethy, C. D.; Botommley, F.; Chen, J.; Kemp, M. F.; Mallais, T. C.; Womiloju, O. O.; Morris, R. J.; Shaw, S. L.; Jefferis, J. M.; Storhoff, J. J.; Goedde, D. M.; Hakanson, M. Transition metal complexes and precursors. In Inorganic Syntheses; Darensbourg, M. Y., Ed.; John Wiley & Sons, Inc.: New York, 1998; Vol. 32, pp 141-228. (4) Bin, X. M.; Kraatz, H. B. Analyst 2009, 134, 1309–1313. (5) Burda, J. V.; Sponer, J.; Leszczynski, J.; Hobza, P. J. Phys. Chem. B 1997, 101, 9670–9677. (6) Cheng, H.; Zhang, K.; Libera, J. A.; de la Cruz, M. O.; Bedzyk, M. J. Biophys. J. 2006, 90, 1164–1174. (7) Di Felice, R.; Calzolari, A.; Zhang, H. Nanotechnology 2004, 15, 1256– 1263. (8) Egli, M. Chem. Biol. 2002, 9, 277–286. (9) Sorokin, V. A. Biofizika 1994, 39, 993–1003. (10) Lu, X. Q.; Zhu, K. M.; Zhang, M.; Liu, H. D.; Kang, J. W. J. Biochem. Biophys. Methods 2002, 52, 189–200. (11) Metcalfe, C.; Thomas, J. A. Chem. Soc. ReV. 2003, 32, 215–224. (12) Petrov, A. S.; Funseth-Smotzer, J.; Pack, G. R. Int. J. Quantum Chem. 2005, 102, 645–655. (13) Andrushchenko, V.; Bour, P. J. Phys. Chem. B 2009, 113, 283–291. (14) Andrushchenko, V.; van de Sande, J. H.; Wieser, H. Biopolymers 2003, 72, 374–390. (15) Rulisek, L.; Sponer, J. J. Phys. Chem. B 2003, 107, 1913–1923. (16) Stoliar, P.; Bystrenova, E.; Quiroga, S. D.; Annibale, P.; Facchini, M.; Spijkman, M.; Setayesh, S.; de Leeuw, D.; Biscarini, F. Biosens. Bioelectron. 2009, 24, 2935–2938. (17) Lee, D. Y.; Choi, W. S.; Kafi, A. K. M.; Park, S. H.; Kwon, Y. S. J. Nanosci. Nanotechnol. 2008, 4553–4556. (18) Mairal, T.; Ozalp, V. C.; Sanchez, P. L.; Mir, M.; Katakis, I.; O’Sullivan, C. K. Anal. Bioanal. Chem. 2008, 390, 989–1007. (19) Poeckh, T.; Lopez, S.; Fuller, A. O.; Solomon, M. J.; Larson, R. G. Anal. Biochem. 2008, 373, 253–262. (20) Latulippe, D. R.; Zydney, A. L. Biotechnol. Bioeng. 2008, 99, 390– 398. (21) Kaufmann, R.; Averbukh, I.; Naaman, R.; Daube, S. S. Langmuir 2008, 24, 927–931. (22) Pietramellara, G.; Ascher, J.; Borgogni, F.; Ceccherini, M. T.; Guerri, G.; Nannipieri, P. Biol. Fertil. Soils 2009, 45, 219–235. (23) Nguyen, T. H.; Chen, K. L. EnViron. Sci. Technol. 2007, 41, 5370– 5375.

Nguyen et al. (24) Nguyen, T. H.; Elimelech, M. Biomacromolecules 2007, 8, 24–32. (25) Nguyen, T. H.; Elimelech, M. Langmuir 2007, 23, 3273–3279. (26) Rawle, R. J.; Selassie, C. R. D.; Johal, M. S. Langmuir 2007, 23, 9563–9566. (27) Douarche, C.; Cortes, R.; de Villeneuve, C. H.; Roser, S. J.; Braslau, A. J. Chem. Phys. 2008, 128, 15. (28) Douarche, C.; Cortes, R.; Roser, S. J.; Sikorav, J. L.; Braslau, A. J. Phys. Chem. B 2008, 112, 13676–13679. (29) Yang, A. Y.; Rawle, R. J.; Selassie, C. R. D.; Johal, M. S. Biomacromolecules 2008, 9, 3416–3421. (30) Cai, P.; Huang, Q. Y.; Li, M.; Liang, W. Colloids Surf., B 2008, 62, 299–306. (31) Cai, P.; Huang, Q. Y.; Zhang, X. W. EnViron. Sci. Technol. 2006, 40, 2971–2976. (32) Cai, P.; Huang, Q. Y.; Zhu, J.; Jiang, D. H.; Zhou, X. Y.; Rong, X. M.; Liang, W. Colloids Surf., B 2007, 54, 53–59. (33) Pietramellara, G.; Ascher, J.; Ceccherini, M. T.; Nannipieri, P.; Wenderoth, D. Biol. Fertil. Soils 2007, 43, 731–739. (34) Hansma, H. G.; Laney, D. E. Biophys. J. 1996, 70, 1933–1939. (35) Pastre, D.; Pietrement, O.; Fusil, P.; Landousy, F.; Jeusset, J.; David, M. O.; Hamon, C.; Le Cam, E.; Zozime, A. Biophys. J. 2003, 85, 2507–2518. (36) Thomson, N. H.; Kasas, S.; Smith, B.; Hansma, H. G.; Hansma, P. K. Langmuir 1996, 12, 5905–5908. (37) Pietrement, O.; Pastre, D.; Fusil, S.; Jeusset, J.; David, M. O.; Landousy, F.; Hamon, L.; Zozime, A.; Le Cam, E. Langmuir 2003, 19, 2536–2539. (38) Libera, J. A.; Cheng, H.; de la Cruz, M. O.; Bedzyk, M. J. J. Phys. Chem. B 2005, 109, 23001–23007. (39) Langowski, J. Biophys. Chem. 1987, 27, 263–271. (40) Langowski, J.; Giesen, U. Biophys. Chem. 1989, 34, 9–18. (41) de Kerchove, A. J.; Elimelech, M. Macromolecules 2006, 39, 6558– 6564. (42) Yuan, B. L.; Pham, M.; Nguyen, T. H. EnViron. Sci. Technol. 2008, 42, 7628–7633. (43) Martin, S. J.; Granstaff, V. E.; Frye, G. C. Anal. Chem. 1991, 63, 2272–2281. (44) Duguid, J.; Bloomfield, V. A.; Benevides, J.; Thomas, G. J. Biophys. J. 1993, 65, 1916–1928. (45) Hackl, E. V.; Kornilova, S. V.; Blagoi, Y. P. Int. J. Biol. Macromol. 2005, 35, 175–191. (46) Richard, H.; Schreibe, J.; Daune, M. Biopolymers 1973, 12, 1–10. (47) Martell, A. E. Smith, R. M. NIST critically selected stability constants of metal complexes database; National Institute of Standards and Technology (NIST): Gaithersburg, MD, 2001. (48) de Kerchove, A. J.; Elimelech, M. Appl. EnViron. Microbiol. 2007, 73, 5227–5234. (49) Stumm, W.; Hohl, H.; Dalang, F. Croat. Chem. Acta 1976, 48, 491–504.

BM901427N