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* Department of Pharmaceutical Chemistry, University of Kansas, Lawrence, Kansas 66047; Department of Biochemistry and Molecular Biology, University of Kansas Medical Center, Kansas City, Kansas 66160; Dendritic Nanotechnologies, Mt. Pleasant, Michigan 48858; and Valentis Inc., Burlingame, California 94010
Correspondence: Address reprint requests to C. Russell Middaugh, Dept. of Pharmaceutical Chemistry, 2095 Constant Ave., Lawrence, KS 66047. Tel.: 785-864-5813; Fax: 785-864-5814; Email: middaugh{at}ku.edu.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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; Felgner et al., 1994, 1987; Huang et al., 2002, 1998; Kukowska-Latallo et al., 1996; Murphy et al., 1998; Philip et al., 1993; Zhu et al., 1993). In concert with these efforts, attempts have been made to establish structure/function relationships between polycationic delivery systems and their biological activity (Braun et al., 2003; Choosakoonkriang et al., 2003a,b, 2001; Dunlap et al., 1997; Golan et al., 1999; Koltover et al., 1998; Lee et al., 2001; Lin et al., 2003; Lobo et al., 2001, 2003, 2002; Radler et al., 1997; Wiethoff et al., 2002, 2003, 2004, 2001). Surprisingly, however, the assembly of polycationic gene delivery complexes has rarely been studied except under equilibrium and near-equilibrium conditions (Lai and van Zanten, 2001, 2002; Tecle et al., 2003). Here we explore temporal aspects of DNA/polymer assembly, employing stopped-flow circular dichroism (CD) and extrinsic fluorescence.
At a minimum, assembly of DNA/polycationic complexes can be viewed to consist of at least two sequential events. First, a spontaneous, electrostatically mediated binding occurs between positively charged amines on polycations with the negatively charged phosphates of the DNA. This produces local and eventually long-range charge neutralization resulting in collapse of the DNA and polycation into a more condensed phase. Cationic lipid/DNA complexes (CLDCs) undergo condensation to a locally ordered phase characterized by stacked arrays of lipid bilayer lamellae between which DNA is sandwiched (Radler et al., 1997). In contrast, polyamine polymers such as polyamidoamine dendrimer/DNA complexes (PDDCs) appears to condense to rod-like or toroidal structures with an underlying square or hexagonal unit cell depending on solution conditions and the chemistry of the individual polycations (Dunlap et al., 1997; Evans et al., 2003; Golan et al., 1999).
The spectroscopic properties of CLDCs and PDDCs at equilibrium have been studied previously. The CD spectra of these complexes are altered significantly from that of unliganded DNA, generally displaying red-shifted peaks of decreased rotational strength characteristics of B-form oligonucleotides (Akao et al., 1996; Braun et al., 2003; Zuidam et al., 1999). The fluorescence of many dyes that bind DNA is altered either through displacement from their binding sites by polycations or by dehydration of the DNA upon formation of complexes (Bhattacharya and Mandal, 1998; Birchall et al., 1999; Eastman et al., 1997; Gershon et al., 1993; Harvie et al., 1998; McKenzie et al., 2000; Wong et al., 2003). By extending these two techniques into the time domain in this study, we attempt to resolve the binding event from the condensation/reorganization process.
To this end, we have examined the formation kinetics of both CLDCs (consisting of dimethyldioctadecylammonium bromide (DDAB) or 1, 2-dioleoyl- 3-trimethylammonium-propane (DOTAP)) and a series of PDDCs (consisting of generation 2, 4, 7, or 9 polyamidoamine (PAMAM) dendrimer) by stopped-flow circular dichroism and fluorescence spectroscopy. The chemical structures of the cationic lipids and a representative PAMAM dendrimer are shown on the left side of Fig. 1. Cationic lipid/DNA complexes are formed from single unilamellar liposomes of 
100 nm in diameter whereas the various generations of PAMAM dendrimer utilized to form PDDCs vary considerably in size. An illustration of the relative sizes of each cation is represented by a series of circles in Fig. 1. The kinetics of formation of complexes from each of these cationic vehicles was studied at five temperatures from 10 to 50°C to derive the activation energy from the temperature dependence of each kinetic phase. The rates derived from exponential fits to the resultant data as well as the associated activation energies (Ea) are discussed in terms of the associated physical events of the complexation process.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Preparation of complexes
Liposomes were prepared by placing a chloroform solution containing the cationic lipid into a glass vial and evaporating the solvent by use of a stream of nitrogen gas. The lipid film formed was dried for 2.5 h in a desiccator under vacuum. The lipid was then hydrated with 10 mM MOPS buffer, pH 7.4, by vortexing. The resultant liposomes were allowed to equilibrate for 30 min and were then extruded 11 times through a 100-nm pore polycarbonate membrane (Whatman, Clifton, NJ) to form small unilamellar vesicles. Both the hydration and extrusion of lipids were carried out above their relevant phase transition temperatures (i.e., room temperature for DOTAP and 55°C for DDAB). The final size of the liposomes used in all experiments was 105 ± 17 nm as measured by dynamic light scattering.
The dendritic polymers were received at high concentration (>10% w/w) in methyl alcohol. An aliquot of the stock solution was placed into a glass vial and the solvent was removed by evaporation using a stream of dry nitrogen. Removal of the solvent was assured by placing the vial containing the dendrimer under vacuum until a stable weight was achieved. The dendrimer was then hydrated in 10 mM MOPS buffer, pH 7.4, by vortex mixing before analysis. The plasmid was diluted into 10 mM MOPS buffer (pH 7.4) from a stock solution to an appropriate concentration for further analysis. The concentration of DNA solutions was determined using their ultraviolet absorbance at 260 nm (A260) and a molar extinction coefficient of 0.02 (mg–1 cm–1 ml). Complexes were prepared by stopped-flow mixing from two independent solutions at the time of data collection. The size of the resultant complexes are 245, 105, 172, and 258 nm (±29 nm) for G2, 4, 7, and 9 dendrimers, respectively, as determined by DLS.
Stopped-flow circular dichroism spectroscopy
Kinetic data were acquired with a stopped-flow CD, PiStar-180 instrument (Applied Photophysics, Surrey, UK). The instrument's optical system was configured with a 75-W xenon lamp, circular light polarizer, and end-mounted photomultiplier. The stopped-flow system was configured with a two-syringe (2.5-ml) kinetic sample handling unit, a 20-µl flow cell (with a 10-mm pathlength), and 2.5-ml stop syringe. Temperature control was achieved through use of a Polyscience digital circulating water bath (Niles, IL). Individual solutions containing pMB 290 (40 µg ml–1) and either DDAB (58 µg ml–1), DOTAP (65 µg ml–1) or dendrimer (20 µg ml–1) were dispensed into the drive syringes. Formation of 0.75 charge ratio (positive/negative; from here on in the text, all ratios will be of the form +/–) complexes was induced by mixing equal volumes of the two components by stopped-flow addition to the flow cell. Acquisition of CD data was performed at a wavelength of 275 nm over a time frame of up to 30 s with slit widths of 3–10 nm. Before kinetic runs, the flow cell was purged with five 50 µl injections of the solutions to be analyzed. Kinetic data were then acquired as the average of 10 separate 50 µl injections.
Stopped-flow fluorescence spectroscopy
Kinetic fluorescence data were acquired with a PiStar-180 stopped-flow fluorimeter (Applied Photophysics). The optical system employed a 75-W xenon lamp, optical cut-off filters appropriate for the dye used, and an emission photomultiplier mounted 90° from the excitation light path. The kinetic sampling unit was identical to that described above. Temperature control was achieved through use of a Polyscience digital circulating water bath. The plasmid DNA (20 µg ml–1) was associated with the DNA binding dye Hoechst 33258 (excitation 352 nm, emission 462 nm) at a ratio of 1 dye molecule per 150 basepairs (bp) or with YOYO-1 (excitation 491 nm, emission 509 nm) at a ratio of 1 dye molecule per 50 basepairs. Individual solutions containing the cationic vehicles DDAB (233 µg ml–1), DOTAP (129 µg ml–1), or dendrimers (40 µg ml–1) were transferred with fluorescently labeled DNA to the appropriate instrument drive syringe. Although polyamidoamine dendrimers have been reported to spontaneously fluoresce the fluorescence is highly pH dependent and can be essentially neglected above a pH of 6 (Wang and Imae, 2004). Complexes of charge ratio 3 (+/–) were formed for DOTAP CLDCs and all PDDCs whereas DDAB CLDCs were formed at a charge ratio of 6 (+/–) to avoid the colloidal instability present with the latter liposomes (Braun et al., 2003).
Kinetic data were acquired as the average of five 50 µl injections. Instrument parameters for Hoechst-33258-labeled DNA consisted of excitation at 352 nm, a slit width of 0.5 nm, and a long pass 395-nm optical filter (Oriel, Stratford, CT). The instrument parameters for YOYO-1-labeled DNA employed excitation at 470 nm, a slit width of 0.3 nm, and a long pass 495-nm optical filter. Data were collected at a rate of 800 points over a 2 s interval.
Data analysis was performed using Origin 7 software (Northampton, IL). Kinetic data were fit by a nonlinear least-squares method to a biexponential relationship (Eq. 1) with the exception of experiments with HOEC G2/DNA complexes at high temperatures (40 and 50°C) that were best fit to a monoexponential form:
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Calculated rate constants were plotted according to a linear form of the Arrhenius equation (Eq. 2):
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A linear least-squares fit was performed and the activation energy for each kinetic event was estimated from the slope.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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; Braun et al., 2003; Choosakoonkriang et al., 2003a). The most prominent change in the CD spectrum as the DNA is complexed is a reduction (
30%) in the intensity of the characteristic B-form ellipticity peak at 275 nm. The maximal spectral change for the cationic reagents employed here occurs when the cationic charge is slightly less than theoretical charge neutrality (0.75) (Braun et al., 2003). In fact, our stopped-flow investigations of DNA complexing to both cationic lipids (DDAB and DOTAP) and PAMAM dendrimers (generations 2, 4, 7, and 9) provided little useful kinetic data. Kinetic plots of the intensity of the 275 nm B-form peak of DNA show only a sharp downward spike centered at 40–50 ms (Fig. 2). This abrupt spectral change, however, occurs in controls consisting of DNA mixed with buffer. Because this time-dependent spectral feature appeared in the absence of any cationic ligand, we believe it arises from either a shear-induced alteration of base stacking within the DNA upon injection into the flow cell (Crook et al., 1996; Levy et al., 2000) or residual Schlieren optical effects from the mixing process itself. We further conclude that the reduction in rotational strength expected from formation of nonviral complexes (and seen in complexes removed from the mixing chamber) occurs within the 40–50 ms time frame in which shear-induced effects predominate. A comparison of agarose gel electrophoretic banding patterns before and after stopped-flow analysis shows no discernible difference suggesting that any alteration in the CD is a temporary effect and does not reflect fragmentation or loss of supercoiling in the plasmid (data not shown).
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). This property combined with the sensitivity of its fluorescence to environmental polarity makes HOEC an effective probe of the kinetics of binding of cationic lipids to DNA. Some degree of dye displacement is expected with PAMAM dendrimers based on studies that find displacement of 50% of the dye by another polyamine, polyethyleneimine (PEI) (Wiethoff et al., 2003). We found, however, that the fraction of HOEC that remains bound is sufficient to assess PAMAM dendrimer association as well. Representative kinetic traces of the change in relative fluorescence intensity during the first 2 s of the binding of DNA to dendrimers and cationic lipids are shown in Fig. 3, A–D. Kinetic traces of DNA diluted with 10 mM MOPS buffer (pH 7.4) and the formation of G2 dendrimer/DNA complexes are shown in Fig. 3, A and B, respectively. The similarity of the DNA alone and G2 dendrimer/DNA complex traces suggests that G2 is unable to displace HOEC at temperatures 
30°C. At the higher temperatures examined (40 and 50°C), the slope of the initial phase is negative, indicating a loss of relative fluorescence intensity consistent with displacement of the Hoechst 33258 dye. Shear-induced perturbation is again apparent under 1:1 DNA dilution conditions (Fig. 3 A) as reduced fluorescence intensity. Nevertheless, good biexponential fits could be achieved by neglecting the data at 40 ms. The kinetics of G4 dendrimer/DNA complex formation (Fig. 3 C) is typical of PDDCs formed from the higher generations of PAMAM dendrimers. The sharp reduction in fluorescence intensity observed during G4/DNA complex formation presumably reflects a displacement of the HOEC from the minor groove of the DNA. In contrast, an initial increase in fluorescence intensity concomitant with formation of DOTAP/DNA CLDCs suggests that HOEC remains in the minor groove and experiences a more apolar environment during association with the liposome. A second kinetic phase, however, manifests a reduction in fluorescence intensity perhaps related to a rearrangement of the complex inducing dye displacement. The G4/DNA and DOTAP/DNA complexes were chosen for further investigation due to their higher transfection efficiency (Bielinska et al., 1999; Braun et al., 2005; Kukowska-Latallo et al., 1996; Wiethoff et al., 2004).
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40°C, an initial loss in fluorescence intensity and a rate similar to that seen with higher generation PDDCs is apparent. This may indicate displacement of HOEC under these conditions. This initial rate was fit to a single exponential and reported as such. In fitting the CLDCs kinetic data, the initial rate (k1) corresponding to apparent association appears to occur too quickly to be measured at elevated temperature (40–50°C for both DDAB and DOTAP CLDCs) and therefore is not presented in the table.
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G7 > G4. Although the activation energy for release of HOEC from the kinetics of G2 PDDC formation is based on only two points, we calculated an approximate Ea value of 13 kcal mol–1 (data not shown). This G2/DNA activation energy has only limited utility, however, because we cannot unambiguously assign it to either the fastest (k1) or slower (k2) rate process.
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; Wiethoff et al., 2003; Wong et al., 2001; Zaric et al., 2004). Intercalating dyes (e.g., ethidium bromide) have seen extensive use as probes to measure the conformational perturbation (condensation) of DNA when bound to a variety of nonviral gene delivery agents (Bhattacharya and Mandal, 1998; Choosakoonkriang et al., 2003a; Eastman et al., 1997; Gershon et al., 1993). Representative kinetic traces of changes in the fluorescence of YOYO-1 DNA bound dye upon formation of PDDCs (G4/DNA) and CLDCs (DOTAP/DNA) are illustrated in Fig. 6. A kinetic trace of YOYO-labeled DNA undergoing dilution shows a rapid loss of fluorescence intensity related to shear-induced alteration of DNA structure (Fig. 6 A). The rapid initial increase in fluorescence intensity followed by an eventual decrease during G4/DNA PDDC formation (Fig. 6 B) suggests the early phase may be related to DNA-dendrimer binding. It has been suggested previously that polyamine binding results in dehydration of the DNA, which could explain the increase in fluorescence intensity (Choosakoonkriang et al., 2003a; Wiethoff et al., 2003). The later kinetic phase, characterized by a reduction in fluorescence intensity, probably reflects rearrangement to a more compact structure and quenching of dye fluorescence or exclusion of the dye from its intercalated binding site. The kinetics of formation G2/DNA PDDCs show the initial (k1) phase but no decrease in fluorescence intensity over the 2 s time frame examined. This again suggests that the G2 dendrimer is incapable of displacing YOYO-1 (data not shown). The representative CLDC kinetic trace (Fig. 6 C) shows a brief period of reduced fluorescence intensity (<40 ms), again presumably related to shear effects, followed by an increase in fluorescence intensity that was fit to a biexponential rate equation. The kinetic phase characterized by an increasing fluorescence signal should reflect lipid-DNA binding.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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; Braun et al., 2003, 2005; Wiethoff et al., 2003). On this basis, the kinetics of formation of cationic lipid/DNA complexes composed of DDAB or DOTAP and PAMAM dendrimer/DNA complexes (G2, 4, 7, and 9) were explored by monitoring spectral changes in a stopped-flow mode. Circular dichroism was used to directly evaluate conformational changes within DNA itself whereas changes in extrinsic dye fluorescence were employed to investigate both binding processes and condensation of the DNA within complexes.
Stopped-flow CD spectroscopy
Stopped-flow CD kinetic traces of the assembly of both CLDCs and PDDCs were dominated by a sharp decrease in the ellipticity at the wavelength of the major CD maximum (275 nm), with the change maximal at 40–50 ms. Nearly identical kinetic traces were observed when DNA was simply diluted. As discussed above, this sharp reduction in rotational strength is probably related to a shear-induced perturbation of the base-stacking interactions of the plasmid. This explanation is consistent with observations from the plasmid processing literature in which shear-induced tertiary structural changes have been reported for plasmid DNA (Anchordoquy et al., 1998; Lengsfeld and Anchordoquy, 2002; Levy et al., 1999, 2000). Agarose gel electrophoresis of the plasmid before and after being subjected to stopped-flow shear confirmed the extent of supercoiling remained constant suggesting the perturbation is transient. Unfortunately, the predicted stopped-flow CD changes associated with the formation of both CLDCs and PDDCs seen in equilibrium experiments occurred within the 100 ms time period over which this shear-induced effect predominates. Therefore, we can only conclude that the early binding events that produce the conformation changes reflected in the loss of rotational strength occur within 100 ms. A similar sub-100-ms time for binding of a fluorescent peptide has been found previously in a stopped-flow fluorescence study of cationic peptide/DNA assembly (Tecle et al., 2003).
Stopped-flow fluorescence spectroscopy
High-affinity DNA binding fluorescent dyes have made important contributions to the brief history of the study of nonviral gene delivery (Bhattacharya and Mandal, 1998; Birchall et al., 1999; Chen et al., 2000; Gershon et al., 1993; Harvie et al., 1998; Richter-Egger et al., 2001). We combined the use of two such probes with different binding modes to investigate the formation kinetics of both CLDCs and PDDCs. For this purpose, Hoechst 33258, a minor groove binding dye, and YOYO-1, a bisintercalating dye were employed. The dye/basepair (dye/bp) ratio used to label the DNA was based on preliminary studies to assure a homogenous binding mode for each dye. Hoechst 33258 was associated with DNA at a 1:150 dye/bp ratio to promote preferential binding to AT-rich regions and minimize the potential for dye displacement (Wiethoff et al., 2003). YOYO-1 was associated with DNA at a 1:50 dye/bp ratio at which the binding mode is exclusively bisintercalation (Larsson et al., 1994; Rye et al., 1992) and no energy transfer or self-quenching occurs (Cosa et al., 2001).
In contrast to the CD kinetic studies, the stopped-flow induced shear is only modestly reflected in the extrinsic fluorescence changes. When DNA labeled with either dye was diluted by stopped flow, the fluorescence undergoes a rapid change in intensity followed by a slower recovery of the fluorescence intensity. The magnitude of this shear-induced fluorescence change is small but significant with respect to the changes occurring during formation of CLDCs and PDDCs. The presence of the cationic reagents affects the rate of DNA recovery to its equilibrium state. Therefore, this shear-induced effect cannot be deconvoluted from the complexation kinetics and must be considered in the interpretation of the data described below.
Hoechst 33258 as a probe of nonviral gene delivery complex formation
The kinetics of formation of PDDCs and CLDCs differ in that the initial kinetic phase (k1) for PDDCs undergoes a rapid biexponential loss of fluorescence intensity whereas CLDCs induce increasing intensity followed by a modest decrease in fluorescence. This result agrees with the observation that HOEC remains substantially bound within CLDCs but is displaced by polyamines such as PEI (Wiethoff et al., 2003). In the case of PDDC formation, both the k1 and k2 phases are associated with a rapid loss of fluorescence intensity commonly assigned to displacement of HOEC from its DNA binding site (Wiethoff et al., 2003). This occurs due to the 30-fold difference in quantum yield between HOEC bound to DNA and free in solution (Cosa et al., 2001). In a related manner, the intercalating dye (ethidium bromide) is displaced from DNA during formation of PDDCs (Braun et al., 2005). The modest reduction in fluorescence intensity observed in the second phase of CLDC formation seems to be due to HOEC displacement. The smaller apparent dye displacement seen with CLDCs compared to PDDCs is consistent with the extent of HOEC displacement observed in equilibrium studies (Wiethoff et al., 2003). The initial phase of CLDC formation is characterized by a small increase in fluorescence intensity that presumably reflects the initial binding of the lipids to DNA. This results in a reduction in the polarity of the environment of the bound dye and a consequent increase in fluorescence intensity. This increase is modest and therefore the contribution of the recovery of the DNA equilibrium state may influence this rate. With regard to the binding event, it has been argued on the basis of a number of Fourier transform infrared (FTIR) studies that DNA is dehydrated as a consequence of the formation of electrostatically assembled nonviral gene delivery systems (Choosakoonkriang et al., 2003a, 2001; Wiethoff et al., 2003). Furthermore, it has been previously shown that a significant blue shift in the HOEC fluorescence peak in CLDCs is accompanied by a >25% increase in intensity suggesting these spectral changes are directly related to dehydration of the DNA site where HOEC is bound (Wiethoff et al., 2003). The CD studies reported here and in a previous report utilizing a fluorescent peptide show that the initial binding event occurs in <100 ms. This implies that the event related to dye dehydration consists of at least two steps involving both a binding and release of water through a reorganization step (Tecle et al., 2003).
YOYO-1 as a probe of nonviral gene delivery complex formation
The response of YOYO-1 to the formation of both PDDCs and CLDCs includes an initial phase marked by an increase in fluorescence intensity. This early phase is presumably due to the binding/dehydration event. The appearance of this early binding phase at the low dye/bp ratio of 1:50 suggests the dye remained predominately bound (Cosa et al., 2001; Larsson et al., 1994). The fact that PDDCs show this early phase in contrast to the results obtained with HOEC suggests dehydration occurs with polyamine polymers as well (Choosakoonkriang et al., 2003a; Wiethoff et al., 2003). The assembly of higher generation (G4, 7, and 9) DNA/dendrimer complexes includes an additional kinetic phase (k2) characterized by a decrease in fluorescence intensity. Although it is tempting to assign this later phase to dye displacement, it is also possible fluorescence quenching upon condensation of the complexes may account for the fluorescence loss. Although both mechanisms appear feasible, we favor the quenching hypothesis because the shortest measured fluorescence lifetime remained much longer (by a factor of 5) when DNA containing YOYO-1 was condensed with PEI than the lifetime of the dye free in solution (Krishnamoorthy et al., 2002; Wong et al., 2001). The series of dendrimers used in these studies have also been reported to condense DNA into small particles with a toroidal structure (Bielinska et al., 1997; Braun et al., 2005). It should be noted that quenching does not require overall collapse into small dense particles but can also be accomplished by localized condensation near dye binding sites. Therefore, we assume the second kinetic phase of PDDC formation provides the best measure of the condensation event. Neither G2/DNA PDDCs nor CLDCs show a reduction in fluorescence intensity within the 2 s period examined. This suggests that the degree of condensation is less for these polycations than for higher generation PAMAM dendrimers. The finding that generation size correlates with an increased ability to displace fluorescent dyes has been reported previously with the intercalating dye ethidium bromide (Braun et al., 2005; Chen et al., 2000).
Activation energies
Based on assignment of each kinetic phase to particular molecular events, we can apply this interpretation to the related activation energies. Using HOEC as a probe, only the first kinetic phase (k1) of CLDCs appears representative of the binding/dehydration event. The results with YOYO were used to calculate rates (k1) for the binding/dehydration event of both cationic lipids and dendrimers. The higher generation PDDCs (G4, 7, and 9) have activation energies that follow the trend of G4 > G7 > G9 and values ranging from 15.7 to 11.1 kcal mol–1. The remaining PAMAM dendrimer (G2) has a binding activation energy (6.5 kcal mol–1) nearly half that of the higher generations. In addition, the slope of the latter phase (k2) changes from positive in the case of the G2 dendrimer to negative with the higher generation complexes. These differences suggest a different binding mode for G2, a conclusion drawn in previous equilibrium fluorescence dye studies (Chen et al., 2000). The CLDCs show similar activation energies (DDAB 11.2 kcal mol–1 and DOTAP 10.3 kcal mol–1) to those of the higher generation PDDCs. The activation energies of binding for CLDCs derived from HOEC (DDAB 3.0 kcal mol–1 and DOTAP 13.7 kcal mol–1) do not correspond well with those derived from the YOYO studies. The low HOEC activation energy for DDAB binding is inconsistent with the values derived for DOTAP CLDCs or those from YOYO studies. This discrepancy could arise from differing degrees of DNA dehydration upon binding between the two lipids. Both fluorescence and FTIR studies at equilibrium, however, suggest the degree of dehydration of DDAB and DOTAP is similar (Choosakoonkriang et al., 2001; Wiethoff et al., 2003). It thus seems more likely that differences in the intrinsic nature of the oligonucleotide/polycation interaction between the lipids account for the activation energy discrepancy. DDAB and DOTAP differ in both the size of their headgroups and the degree of saturation of their hydrophobic tails. It has been observed in thermal stability and FTIR studies that the larger headgroup of DOTAP has a greater destabilizing effect on DNA structure and a greater degree of interaction with the nucleotide bases (Lobo et al., 2002). Although DDAB is in a gel phase and DOTAP a more fluid liquid crystalline bilayer state in the temperature range (10–30°C) from which the activation energies were determined, it seems more likely that headgroup chemistry dominates the binding event given the surface location of these moieties (Lobo et al., 2002). That a similar trend is not apparent in activation energies derived from YOYO fluorescence suggests DDAB may have a preferential interaction with HOEC or its minor groove binding site.
The secondary condensation event was best observed during PDDC formation. Activation energies of all the polycations were determined from the HOEC fluorescence studies. The activation energies for PDDCs follow the trend G9 > G7 > G4, which suggests the activation energy barrier is higher for larger PAMAM dendrimers. This same trend is observed for the activation energies determined by YOYO monitored condensation. The only significant difference between the HOEC and YOYO studies is that the average activation energy is 4 kcal mol–1 more for YOYO. These higher activation energies suggest that a greater degree of condensation is necessary to quench YOYO fluorescence than to displace the HOEC dye. The displacement of HOEC by G2 PAMAM dendrimers at 40 and 50°C suggests that HOEC binding is weaker at elevated temperature.
In the case of CLDCs, no kinetic event related to YOYO quenching (condensation) was observed. Although as argued earlier it seems likely only a small fraction of HOEC is displaced by CLDCs, the trend in activation energies of DDAB >> DOTAP suggests a difference in the structure of the CLDCs formed. Hoechst 33258 and FTIR studies do show differences between the two lipids, which may be related to differential condensation. Equilibrium Hoechst 33258 studies confirm DOTAP dehydrates the DNA to a greater extent than DDAB. At high dye/DNA (1:25) loading, however, DDAB complexes of high charge ratio (>2) have a higher fluorescence intensity (Wiethoff et al., 2003). This suggests DOTAP complexes lose intensity through dye displacement during condensation whereas the dye within DDAB complexes remains bound. Additionally, the frequency of both the asymmetric phosphate stretching band (1223 cm–1) and the base carbonyl stretching vibration (1715 cm–1) increases more when complexed to DOTAP than DDAB (Choosakoonkriang et al., 2001). Together, these observations suggest that DOTAP may condense DNA to a greater degree than DDAB and that is reflected in a stronger interaction. The difference in headgroup between the two lipids, however, may also contribute to these effects. Because the condensation event involves structural rearrangement of the complexes, the difference in activation energies between DDAB (27 kcal mol–1) and DOTAP (8 kcal mol–1) may arise from the difference in their bilayer properties. A number of studies show that DDAB CLDCs are significantly larger than DOTAP complexes suggesting a reduced degree of compaction in the former (Braun et al., 2003; Choosakoonkriang et al., 2001; Lobo et al., 2001).
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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, 2005; Choosakoonkriang et al., 2003a; Lobo et al., 2001; Wiethoff et al., 2002, 2003, 2004). Transfection studies of both PDDCs and CLDCs indicate that complexes composed of moderate (G4–G7) generation PAMAM dendrimers or DOTAP liposomes provide the greatest biological activity (Braun et al., 2005; Kukowska-Latallo et al., 1996; Wiethoff et al., 2004). The activation energies of complex formation obtained here also show no obvious correlation with transfection efficiency. The kinetic behavior of dendrimer mediated complexation may best be considered in terms of two different classes of the polycations; the smaller G2 dendrimer and the larger G4–G9 molecules. For the series of higher generation dendrimers, the binding event follows the trend G4 > G7 > G9, whereas the opposite order was observed in the activation energies reflecting condensation events. Thus, we suggest a balance between a more energetically favorable condensation reaction and less favorable binding may prove beneficial to the competing needs of cellular uptake and dissociation of complexes within the cytoplasm. Submitted on October 28, 2004; accepted for publication March 25, 2005.
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REFERENCES |
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Bielinska, A. U., J. F. Kukowska-Latallo, and J. R. Baker, Jr. 1997. The interaction of plasmid DNA with polyamidoamine dendrimers: mechanism of complex formation and analysis of alterations induced in nuclease sensitivity and transcriptional activity of the complexed DNA. Biochim. Biophys. Acta. 1353:180–190.[Medline]
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Braun, C. S., G. S. Jas, S. Choosakoonkriang, G. S. Koe, J. G. Smith, and C. R. Middaugh. 2003. The structure of DNA within cationic lipid/DNA complexes. Biophys. J. 84:1114–1123.
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) and DOTAP/DNA complexes (
). The kinetic plots were acquired at 20°C for 2 s as the average of 10 individual injections.
