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* Department of Pharmaceutical Chemistry, University of Kansas, Lawrence, Kansas 66047; Higuchi Bioscience Center, University of Kansas, Lawrence Kansas 66047; and Valentis Inc., Burlingame, California 94010
Correspondence: Address reprint requests to C. Russell Middaugh, Dept. of Pharmaceutical Chemistry, University of Kansas, 2095 Constant Ave., Lawrence, KS 66047. Tel.: 785-864-5813; Fax: 785-864-5814; E-mail: Middaugh{at}ku.edu.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL SECTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL SECTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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; Xu et al., 1999; Radler et al., 1998). These analyses suggest that mesophases are formed in which DNA is enclosed by lipid bilayers in either lamellar or inverted hexagonal phases that are stacked to form higher order structures. The state of the DNA within these mesophases is not, however, clearly defined.
Recently, spectroscopic characterization of CLDCs using Raman, FTIR, UV/visible, fluorescence, and CD spectroscopies has been used to evaluate the structure of the individual components (Deng et al., 2000; Choosakoonkriang et al., 2001a,b; Braun et al., 2001). Vibrational spectroscopy suggests that the DNA maintains a B form conformation and is dehydrated at the lipid interface (Hirsch-Lerner and Barenholz, 1999; Choosakoonkriang et al., 2001a). This work also reveals, as expected, that primary lipid/DNA interactions occur between the negatively charged phosphates of the DNA backbone and the positively charged lipid headgoups. Changes in lipid methylene vibrations also indicate that the apolar region of the lipid bilayer becomes further fluidized when DNA is bound (Davies et al., 1990; Choosakoonkriang et al., 2001a; Mendelsohn et al., 1989, 1991). These techniques are limited, however, by a lack of sensitivity, requiring formation of complexes at high concentrations, a condition under which colloidal stability is compromised. CLDCs form lyotropic mesophases that are known to have a strong dependence on concentration, making extrapolation to lower, more biologically and clinically relevant conditions difficult (Kabanov et al., 1998). In contrast, CD spectroscopy can be used to measure the long-range structure of DNA directly at concentrations from 10- to 100-fold lower than those used in vibrational spectroscopy. CD is particularly sensitive to the precise nature of the helical state of the DNA. The weak chirality of lipids allows the signals from the DNA in CLDCs to be clearly resolved with little or no interference from the lipid component. Initial CD studies of cationic lipid/DNA systems have been interpreted to indicate that DNA assumes a C form conformation in direct contradiction to the FTIR results (Patil and Rhodes, 2000; Simberg et al., 2001; Akao et al., 1996; Zuidam et al., 1999; Choosakoonkriang et al., 2001a).
The size and complexity of CLDCs have the potential to introduce significant artifacts into their CD spectra. In general, the particles formed upon complexation are typically greater than 
120 nm diameter and display a heterogeneous size distribution. These large sizes and the heterogeneous nature of the complexes including a concentrated and uneven distribution of chromophores could give rise to significant absorption flattening effects (Wallace and Teeters, 1987; Mao and Wallace, 1984; Rodger and Ismail, 2000; Maestre and Reich, 1980; Glaeser and Jap, 1985). Furthermore, differential light scattering can occur when the particle dimensions are greater than one-quarter the wavelength of the incident light (Bustamante et al., 1983; Keller and Bustamante, 1986a,b) and the refractive indices for left and right circularly polarized light differ. In this work, we consider the role or lack thereof of such contributions to the CD spectra of CLDCs. After analysis of these effects, we examine the basis of the altered CD spectra of the DNA within CLDCs in terms of complementary FTIR and Raman analyses, model complexes, and finally molecular dynamics simulations. The effects of lipids in both the gel and liquid crystalline state as well as modification of these properties by incorporation of nonbilayer forming (helper) lipids (Hafez et al., 2001; Koltover et al., 1998) are also addressed. We find that the evidence for C form DNA in the complexes unconvincing, but rather that irregular changes in base/base interactions offer a better explanation for the observed spectra.
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EXPERIMENTAL SECTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL SECTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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229 bp), poly(dGdC) · poly(dGdC) (724 bp), and poly(dAdT) · poly(dAdT) (3.2 kbp) ODNs were acquired from Amersham Pharmacia Biotech (Piscataway, NJ). Purified single-stranded ODNs (20 bases) of poly(dA), poly(dT), poly(dC), poly(dG), poly(dAdT), and poly(dCdG) were synthesized by Operon Technologies (Alameda, CA). All polynucleotides were judged to be better than 98% pure as determined by mass spectrometry.
Preparation of CLDC
Liposomes were prepared by drying a lipid containing chloroform solution in a glass vial with a stream of nitrogen. The thin film formed was dried for a minimum of 2.5 h under vacuum desiccation. The film was hydrated with 10 mM Tris buffer at pH 7.4 for 30 min at a temperature above the phase transition temperature of the highest Tc component lipid. Uniform single lamellar liposomes were formed by extrusion 10 times through a 100-nm pore size polycarbonate filter (Whatman, Clifton, NJ). Plasmids and duplex ODNs were prepared by diluting a stock solution to 100–200 µg/ml. The concentrations of these solutions were determined using the UV absorbance at 260 nm (A260) and a molar absorptivity coefficient of 6490 Lxmole-1xcm-1. The concentration of single-stranded ODNs was obtained from the UV absorbance using the ssDNA molar absorptivity coefficients provided by Operon. Duplexes were formed by annealing at 5°C below the transition temperature (provided by Operon) of the corresponding double-stranded polynucleotide and cooling at 0.5°C/min.
Liposomes were stepwise diluted to concentrations appropriate for equal volume mixing with DNA at each charge ratio. Complexation was conducted by rapid addition with 30–60 s of mixing in small volumes of 0.4–1.2 ml. The order of addition was selected to avoid passing through charge neutrality. All charge ratios indicated in the text are given as the ratio of positive to negative charge. Note, however, that molar and charge ratios are essentially equivalent in these systems.
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METHODS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL SECTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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FTIR spectroscopy
FTIR spectra were obtained with a Magna-IR 560 spectrometer (Nicolet, Madison, WI) equipped with a mercury-cadmium-telluride detector using an attenuated total reflectance method in which the sample solution was placed directly in a ZnSe trough element of effective pathlength 12 µm (Thermal A.R.K., SpectraTech, Shelton, CT). Spectra were obtained under dry air purge by coaddition of 256 interferograms. The interferograms were apodized using the Happ-Genzel function, with no zero filling, to give a final resolution of 4 cm-1. The association band of water at 2200 cm-1 was used as a reference for subtraction of water. Baseline corrections (1804-904 cm-1) and seven-point Satvisky-Golay smoothing were applied to the spectra. Peak positions were assigned by an algorithm in Omnic software (Nicolet).
Raman spectroscopy
Raman spectra were acquired with a Raman 2000 instrument (Chromex, Albuquerque, NM). Samples were excited at 785 nm with 180 mW of power measured at the source. Raman scattering was collected by a thermoelectrically cooled charge-coupled-device of 1024x256 pixels. The frequency axis was calibrated with 4-acetamidophenol before acquiring sample spectra. Samples were measured at room temperature in a quartz cuvette as the accumulated average of 18 exposures of 100 s at a DNA concentration of 2–3 mg/ml. Baseline subtraction and data analysis was performed with Grams 32 software (Galactic Industries, Salem, NH).
Molecular dynamics simulation
Molecular dynamics simulation of DNA liposome interactions was performed with the force field and the potential parameters from the latest versions of CHARMM (all hydrogen) for nucleic acids (Foloppe and MacKerell, 1998; Foloppe and MacKerell, 1999) and lipids (Schlenkrich et al., 1996). The initial configuration of the system was constructed in several stages. A lipid monolayer was generated by placing 64 protonated 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) molecules on an 8 x 8 grid. Random chain conformations were selected from a library distributed with CHARMM version 26. The positions of the lipids were optimized first by rigid-body translations and rotations of whole molecules and then by energy minimization of the system periodic boundary conditions in the monolayer (xy) plane. The size of the square cell in the xy plane was kept fixed at 74 Å, leading to an area of 85.6 Å2 per lipid, which was found to be optimal for simulations (Feller and Pastor, 1999). To maintain planarity, harmonic constraints of the form k(z-z0)2 were placed on the nitrogen atoms of DMPC, k = 25 kcalxmole-1xÅ-1 and z0 = 0, to maintain average planarity of the monolayer. The B-DNA decamer of sequence C-T-C-T-C-G-A-G-A-G coordinates were taken from 196D.pdb (Goodsell et al., 1995). Initially, the DNA was placed 10.0 Å above the monolayer, oriented along the x axis. To search for optimal DNA:liposome interactions, we performed a systematic search in which the position of the DNA center of geometry was moved relative to the lipid center by -4, 0, and 4 Å along the x and y; and rotations by 0, 60, and 120° around x; -30, -15, 0, 15, and 30° around y; and 0, 30, 60, and 90° around z were applied. At each of the 540 starting points, a 20-ps molecular dynamics simulation was performed, using Langevin dynamics with a friction coefficient of 50 ps-1, a bath temperature of 300 K, and a time step of 2 fs. This was followed by 5000 steps of geometry optimization using the Adopted Basis Newton-Raphson algorithm. A nonbonded cutoff of 12 Å was employed. Screening of electrostatic interactions was introduced by using a dielectric constant equal to 1/R, with R being the interatomic distance (Brooks et al., 1983). Calculations for a typical grid point took 0.5 h on an SGI Origin 2400 supercomputer. The optimized structures were sorted by energy and lowest energy structures analyzed graphically using WaveLab ViewerPro.
Dynamic light scattering
Particle sizing was performed with a Brookhaven (Holtsville, NY) instrument and 9000AT autocorrelator configured with a 50-mW HeNe laser operating at 532 nm (JDS Uniphase) and an EMI 9863 photomultplier tube (PMT) mounted on a BI-200M goniometer. Measurements were taken at a 90° angle from the incident light. The autocorrelation function was analyzed using the method of cumulants (Koppel, 1972), and the resultant mean translation diffusion coefficients were converted to mean hydrodynamic radii by the Stokes-Einstein equation. Each sample was measured three times with the mean and standard deviation reported.
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RESULTS |
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; Johnson, 1994, 1996). The CD of plasmid DNA is significantly perturbed when complexed with two commonly employed cationic liposomes, DOTAP and DDAB. Examples of spectra at various charge ratios of DDAB or DOTAP in complexes with DNA are shown in Fig. 1, A and B. The characteristic features of the canonical B form, a positive band at 275 nm, negative signal at 245 nm, and crossover point near 258 nm, (Johnson, 1994; Ivanov et al., 1973) are altered. Changes include red shifting of both bands and a significant reduction in intensity of the 275-nm peak. The presence of the cationic lipids also appears to induce some degree of differential light scattering at charge ratios near or greater than neutrality as indicated by the presence of spectral "tails". This signal at extended wavelengths outside of absorbance bands is characteristic of the dispersive part of differential light scattering (Keller and Bustamante, 1986a). Spectra that appear flattened, at charge ratios between 1.25 and 4 with DDAB and 1.25–1.75 for DOTAP complexes, formed visible precipitates and were excluded from further analysis.
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75% of the uncomplexed plasmid. The observed red shifts correlate with the intensity changes in that the peak position shifts of both bands are greatest near a 1:1 lipid to DNA molar ratio. In CLDCs of DDAB, the peak positions shift up 3.3 and 8.6 nm for the 245 and 275 nm bands, respectively. When DOTAP is employed, the largest shifts were 2.6 nm for the 245-nm band and 7.6 nm for the 275-nm peak. The lack of spectral changes at high charge ratios suggests the formation of stable structures, but their final values remain significantly different from that seen in unliganded DNA.
Potential contributions of absorption flattening and differential scattering to CD spectra of CLDCs
In an attempt to unambiguously assign these altered spectra to a specific DNA conformation(s), we investigated the potential contribution of the two most common CD spectral artifacts. Because absorption flattening is a purely absorptive phenomenon, it can be investigated independently of CD contributions (Mao and Wallace, 1984). OD spectra of various DOTAP CLDCs show only small shifts in absorption maxima (
0.4 nm) in contrast to the 3- and 8-nm shifts seen in the CD bands (not illustrated). Furthermore, examination of CD spectra over a range of pathlengths and CLDC concentrations reveals little difference in concentration normalized data (data not shown). Both of these results are inconsistent with major contributions of absorption flattening to the CLDC CD spectra.
Differential scattering can distort CD spectra by anisotropically scattering light away from PMT detector. By moving the sample close to the detector, however, a collection angle of >70° can be achieved. This can partially correct for dispersive light scattering by catching much of the differentially scattered light (Rodger and Ismail, 2000). In Fig. 2, the CD spectra of a 1:1 complex of DDAB and DNA show how spectra differ at various distances between the sample and PMT. As expected, the spectrum of the plasmid itself is unaffected by its distance from the PMT. The CLDC spectra, however, do show some distance dependence with the displacement of the spectrum from baseline larger at greater distances. It therefore appears that a greater collection angle (72°) is sufficient to correct for the majority of the differential scattering, but that this has little effect on the large decreases seen in the DNA peak at 275 nm in the CLDCs. Thus, differential scattering appears unable to account for the majority of the CLDC-induced spectral changes.
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; Thomas and Peticolas, 1983a,b; Nishimura and Tsuboi, 1986). Spectra from both methods (which differ in their selection rules) are shown in Fig. 3 and Fig. 4 at lipid/DNA charge ratios that exhibit maximal perturbation in their complementary CD spectra. It should be noted, that the vibrational methods require concentrations that are 10–20x the CD measurements. We find, however, that the size and 
-potential of the complexes (as measured by dynamic and phase analysis light scattering, respectively) do not change over this concentration range. Furthermore, CD analysis conducted at FTIR and Raman concentrations using short pathlength cells produce spectra very similar to those seen at the lower concentrations (not illustrated). Several conformationally sensitive vibrations are present in the FTIR and Raman spectra of all lipid/DNA complexes. The most informative FTIR vibrations are the guanine/thymine carbonyl stretch (1715 cm-1), thymine aromatic amine stretch (1328 cm-1), symmetric and asymmetric phosphate stretches (1090 and 1223 cm-1, respectively), and a strongly coupled sugar-phosphodiester signal (970 cm-1) (Pichler et al., 1999; Colthup et al., 1975). The 1715, 1328, 1223, 1090, and 970 cm-1 bands are traditionally taken as markers of the B form and upon complexation their positions remain consistent with the presence of this form. This suggests that the conformation of the DNA in CLDCs remains in the B form. The helix-sensitive Raman signals include the symmetric phosphate stretch (1095 cm-1), asymmetric phosphodiester (835 cm-1), pyrimidine ring (785 cm-1), guanine ring breathing (683 cm-1), thymine aromatic amine stretch (1375 cm-1), and guanine N7 stretch (1489 cm-1) modes (Serban et al., 2002). The regions from 600 to 700 cm-1 and 800 to 900 cm-1 are uniquely sensitive to the base-sugar conformation. In particular, B and C conformations are easily discerned by the position of the phosphodiester band, which at 875 cm-1 is indicative of C form, and when at 835 cm-1, as in the unliganded plasmid, demonstrative of B form (Erfurth et al., 1972; Nishimura and Tsuboi, 1986; Peticolas et al., 1987). In both complexes, this region gives rise to additional peaks at 823 and 850 cm-1. Due to their greater intensity than the 785 cm-1 band and splitting analogous to the 1095 cm-1 phosphate stretching, we believe these additional peaks reflect lipid/DNA interactions rather than a new nucleotide geometry. The 683 cm-1 peak of guanine is shifted to lower frequency (5 cm-1) in both CLDCs. The shifted DDAB and DOTAP CLDC peaks at 681 and 676 cm-1, respectively, however, remain indicative of the C2' endo-anti nucleotide conformation (Peticolas et al., 1987). Furthermore, the 1375 cm-1 thymine bands remain at 1374 cm-1 upon complexation, suggesting the C2' endo-anti conformation (Nishimura and Tsuboi, 1986). This C2'-endo sugar pucker and anti sugar-base bond angle are the established nucleotide geometry of B form. There are three additional base vibrations altered in complexes, including adenine N6 amine (1422 cm-1), guanine ring N7 (1488 cm-1), and adenine/guanine ring N3 signals (1578 cm-1) (Peticolas and Evertsz, 1992; Peticolas et al., 1987; Serban et al., 2002). These three prominent peaks in the plasmid are reduced or absent in the spectra of both complexes. A new peak also appears in the CLDC spectra at 1465 cm-1, which can be assigned to the guanine ring N7 vibration in the hydrogen bonded state (Peticolas et al., 1987).
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,b). The poly(dGdC) · poly(dGdC) titration in Fig. 5 B produces spectra representative of this state at charge ratios from 3 to 5. The PSI-like effects are opposite to those seen in the plasmid spectra inasmuch as they involve large increases in peak intensity.
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) and their effect on the CD of CLDCs examined. Helper lipids were incorporated at a 1:1 molar ratio to the cationic host lipid, a ratio at which they are known to enhance transfection efficiency (Felgner et al., 1994). Addition of helper lipids to CLDCs results in minor CD spectral differences from the spectra of equivalent charge ratio complexes composed solely of the cationic lipid (data not shown). |
DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL SECTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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C change in secondary structure (Akao et al., 1996; Zuidam et al., 1999; Patil and Rhodes, 2000; Zhang et al., 2002). This is based on the observation that increasing cationic lipid concentrations cause the long wavelength (275 nm) peak to decrease in height whereas the negative band (245 nm) maintains its intensity. Although a loss of 275 nm CD intensity compared to that of B form is, in fact, characteristic of C form, a 245-nm band intensity approximately two-thirds that of the B form is thought to be more characteristic of the C state (Bokma et al., 1987). The reduced 275-nm band and intensification of the 245-nm band we observe, however, more closely resemble cation-induced spectral changes that have been attributed to a noncooperative increase in DNA winding angle (from 10.410.2 bp/turn) that maintains the B form geometry (Bokma et al., 1987). In the latter case, a collapse of the 275-nm peak with little alteration of the 245-nm band is seen (Bokma et al., 1987; Bloomfield et al., 1999). It is clear from our studies that although dispersive light scattering perturbs band intensities, neither absorption flattening nor differential scattering is sufficient to account for these changes.
FTIR suggests that the sugar-base conformation of the plasmid in CLDCs maintains the C2'-endo-anti geometry. This nucleotide conformation excludes many secondary structures, but the B and C forms share this arrangement. Both are members of a class of structural states that have two nucleotide conformations (BI and BII), which differ in their phosphate orientations (van Dam and Levitt, 2000). C form is generally considered to arise when the BII geometry makes up 
45% of the nucleotide population (van Dam and Levitt, 2000). The change from B to C forms is therefore noncooperative, unlike most ODN conformational changes. Nevertheless, IR spectra are still capable of distinguishing the two forms inasmuch as many vibrational states undergo changes in BC transformations. Signals from the G/T carbonyl, phosphodiester and asymmetric phosphate vibrations reportedly shift to 1710, 960, and 1231 cm-1, respectively, in C form (Adam et al., 1987; Taillandier and Liquier, 1992; Pichler et al., 1999; Loprete and Hartman, 1989). In this work, the 970 cm-1 band is unchanged, whereas the 1715 cm-1 and 1223 cm-1 shift but to frequencies inconsistent with C form. The relative intensity of the symmetric phosphate vibration either weakly (1089 cm-1) or strongly (1109 cm-1) hydrogen-bonded has been shown to correlate with the BI and BII conformations, respectively (Pichler et al., 1999). Although an increase in the 1109 cm-1 intensity is apparent in Raman CLDC spectra, this is probably due to electrostatic interactions between the lipid and DNA phosphate, because the 1109 cm-1 band is absent in FTIR spectra. This suggests that BI nucleotides remain the predominant conformation in DNA complexed with cationic lipid. One cannot exclude some change in the relative distribution of the BI and BII forms, however.
Recently a deoxyoligonucleotide duplex (CCCCGGGG) exhibiting an intermediate B/A conformation has been described (Trantirek et al., 2000). The conformation of this duplex has a B-type sugar phosphate backbone (deoxyribose sugar pucker). The bases, however, are shifted from the helix center to the periphery with a wide minor groove appearing at the midpoint of the ODN, an arrangement characteristic of the A form. This intermediate state results from the tendency of guanosine-guanosine stacking to adopt an A-like conformation even within an overall B-DNA helix (Stefl et al., 2001). This polymorphism of local base stacking interactions while maintaining a B-form backbone geometry is consistent with the altered CD spectra occurring in CLDCs and the diverse structures seen at longer times in the molecular dynamics generated images (Fig. 6).
An alternative hypothesis to explain the large changes induced in the CD spectrum of DNA by cationic lipids involves the formation of some type of supramolecular structure whose spectral contribution subtracts from the native DNA spectrum (Zuidam et al., 1999). Such structure is usually thought to arise from long-range interactions primarily within DNA molecules but not necessarily in a condensed state. Studies of linear ODNs ranging from 20 bp to 4000 basepairs, however, show no obvious correlation between basepair length and the spectral intensities of the DNA within CLDCs. This suggests that the CD spectra of cationic lipid/DNA complexes cannot be directly explained by the presence of such structures.
Some hints about the origin of the altered nature of the DNA spectra may be obtained from FTIR and Raman spectroscopies, which provide evidence of altered hydrogen bonding in the bases. Shifts in the FTIR vibrations of the guanine carbonyl and imidazole nitrogen (N7) upon complexation have both been interpreted as reflecting altered hydrogen bonding (Choosakoonkriang et al., 2001a). In Raman spectra, a similar response in the guanine imidazole nitrogen is observed upon complexation. The guanine N7 (1488 cm-1) disappears, whereas a prominent peak at 1465 cm-1, indicative of the hydrogen bonded state, appears (Peticolas et al., 1987). The reduced intensity of the adenine exocyclic amine N6 (1422 cm-1) and ring N3 (1578 cm-1) observed in complexes has been associated with hydrogen bonding (Peticolas et al., 1987). These sites on purine nucleotides are each accessible from the major groove except for the adenine/guanine ring N3, which can be contacted through the minor groove.
Studies of the binding of a cationic amine to DNA produce CD spectra similar to those induced by CLDCs (Chen et al., 1981). In these investigations, calf thymus DNA was modified by covalent binding of butyl amine (BuA) to minor groove guanine exocyclic amines (Chen et al., 1981; Maibenco et al., 1989). Attachment at 12–15% of the bases produced CD spectra with loss of 90% of the 275-nm peak intensity with little change in 245-nm band values (Chen et al., 1981). These CD spectral changes were interpreted in terms of the presence of a variant B form with increased winding angle (Fish et al., 1983; Chen et al., 1983). Interestingly, titration of the adducts' ionization state demonstrates reversibility of the 275-nm intensity change, implying that the charged state of the BuA is critical. Another alternative suggested by the authors is that direct perturbation of the bases by the positively charged bound amine may be involved in the CD spectral changes. The presence of the positively charged headgroup in the cationic lipids studied here could play a corresponding role. Whatever explanation is correct, a significant disruption of the planar interaction between the bases would seem to provide the most direct explanation for the decrease in rotational strength seen here.
Although lack of structural parameters for a suitable cationic lipid prevented building an ideal model, we used a positively charged form of DMPC to mimic a cationic lipid surface and performed a molecular dynamics study of the interaction of this lipid with a 10-bp duplex of DNA. Striking alterations in DNA structure were observed in this model of DNA/lipid interaction. The changes in helicity and base stacking clearly observed in Fig. 6 are undoubtedly excessive due to lack of hydration and the potential conformational mobility of the DNA and DMPC components, which would be expected to be reduced by the DNAs sandwiching between lipid bilayers in actual CLDCs. Nevertheless, it seems clear that lipid/DNA interactions extend beyond direct electrostatic coupling between the lipid headgroup and phosphate backbone to interactions between headgroups and DNA bases. This, in turn, produces alterations in base stacking interactions that would be expected to reduce the intensity of CD signals arising from electrostatic and magnetic coupling between bases. The BuA adduct model demonstrates that a small number of modified sites can have a large effect on CD spectra, implying that limited structural changes could be responsible for the large CD changes seen upon cationic lipid and DNA interaction. In the case of actual plasmid complexes, it seems that disruptive interactions could be limited by both accessibility and competing torsional forces in the supercoiled DNA.
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ACKNOWLEDGEMENTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL SECTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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FOOTNOTES |
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Submitted on August 7, 2002; accepted for publication September 26, 2002.
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