Real time monitoring of lipoplex molar mass, size and density

Journal of Controlled Release 82 (2002) 149–158 www.elsevier.com / locate / jconrel

Real time monitoring of lipoplex molar mass, size and density Eva Lai a , John H. van Zanten b , * a

b

Department of Chemical Engineering, Johns Hopkins University, Baltimore, MD 21218, USA Chemical Engineering Department, North Carolina State University, Raleigh, NC 27695 -7905, USA Received 22 December 2001; accepted 13 April 2002

Abstract Time-resolved multiangle laser light scattering (TR-MALLS) is used to monitor the temporal variation of DNA / cationic liposome lipoplex molar masses and geometric sizes throughout the complexation process. The measured molar masses and geometric sizes are in turn used to estimate lipoplex density. The DNA / cationic lipid charge ratio is found to be the primary factor governing lipoplex formation kinetics and the final lipoplex molar mass, geometric size and density. Charge ratios near unity lead to a growing kinetic regime in which initially formed primary lipoplexes undergo further aggregation eventually forming large molar mass lipoplexes of high density, while charge ratios very far from unity yield low molar mass lipoplexes of lower density. It is also noted that solvent composition can play a significant role in the lipoplex formation process with lipoplexes formed in ion-containing media being larger and denser than those formed in dextrose solution.  2002 Elsevier Science B.V. All rights reserved. Keywords: Cationic liposome; Lipoplex; Molar mass; Nonviral gene delivery vector; Light scattering

1. Introduction It has been more than a decade since Felgner et al. [1] demonstrated that cationic lipids could be used to condense and deliver DNA to cells in vitro. Since this initial study, many cationic liposome preparations have been investigated for their suitability as DNA delivery vectors. Most of these studies have focused on synthesizing novel cationic lipids [2,3], determining DNA-cationic lipid lipoplex microstructures [4–9], increasing the stability of the lipoplexes [10–13] or investigating the gene transfer efficiency *Corresponding author. Tel.: 11-919-515-2520; fax: 11-919515-3465. E-mail address: john [email protected] (J.H. van Zanten). ]

of various cationic liposome formulations [14–17]. While a plethora of studies have been published on cationic liposome-based gene delivery vectors, very few discuss the formation of DNA / cationic liposome lipoplexes. In fact, Safinya and coworkers have noted that DNA condensation by cationic agents is an important process that must be understood and is a crucial factor in the development of non-viral gene delivery vectors [18]. Despite this observation, clinical trials employing cationic lipids for gene delivery have started even though lipoplex formation and its underlying mechanisms are unresolved. This lack of physical understanding has greatly hindered the development of all non-viral gene delivery vectors as other than a few exceptional cases, very few investigators have made sufficient efforts to separate and

0168-3659 / 02 / $ – see front matter  2002 Elsevier Science B.V. All rights reserved. PII: S0168-3659( 02 )00104-9

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quantitatively evaluate the influence of DNA complex geometric and composition factors on gene delivery efficacy. A major contributing factor to this situation is the lack of a rapid, noninvasive technique for ascertaining lipoplex size, mass and structure. This report details a method for determining these important physicochemical parameters. Several mechanisms for DNA / cationic liposome lipoplex formation have been proposed. One study suggests cationic liposomes cluster with DNA and the subsequent fusion of the lipid membranes induces DNA to collapse [4]. This DNA collapse leads to condensed structures, with the DNA encapsulated within the fused lipid bilayers. For multilamellar complexes, it has been suggested that the complex growth starts with one DNA-coated ‘template’ liposome [19]. These studies rely on images obtained from metal shadowing and cryo-electron microscopy, respectively. Thus, the possibility of artifacts introduced by chemical fixation and freezing-induced water crystals may alter the interpretation of the true mechanism. Small angle X-ray scattering measurements of DNA / cationic liposome lipoplexes indicate that either lamellar or hexagonal ordering may exist at small length scales depending on the lipid composition and DNA / lipid mass ratio [6–8]. DNA complex formation kinetics provide insight into and an understanding of the parameters that stabilize the final complexes. Because complex dissociation is believed to be a prerequisite step in the transfection process, the complex stability is a key parameter. The limited number of DNA / cationic liposome lipoplex formation studies are primarily focused on DNA and cationic lipid interactions [20,21] and less so on the actual lipoplex growth process. This study is a first attempt to fill this gap in the literature by focusing on the formation process from initial mixing to the final DNA / cationic liposome lipoplex. The lipoplex formation process is probed non-invasively in bulk solution with timeresolved multi-angle laser light scattering (TRMALLS), in which the temporal evolution of the lipoplex molar mass and geometric size is measured [22]. Although the lipoplexes are prepared and characterized under more dilute conditions than typically used for clinical studies (i.e. 1–10 mg / ml vs. 100–1000 mg / ml), significant insight regarding their size and structure can be obtained. The primary

finding of this study is that the DNA / cationic lipid charge ratio is the major factor governing lipoplex formation kinetics and final lipoplex molar mass, geometric size and density. Final lipoplex molar masses can be in excess of 10 9 g / mol and their densities can be on the order of 0.01–0.30 g / cm 3 , depending on the DNA / cationic lipid charge ratio with charge ratios near unity yielding the largest lipoplex molar masses and densities.

2. Material and methods

2.1. Materials Dextrose (5 wt%) was purchased from Baxter Healthcare Corp. (Deerfield, IL) and prefiltered with 0.02-mm Anotop Whatman filters (Clifton, NJ) prior to use. Careful filtration is necessary to minimize dust contamination that is deleterious to light scattering measurements. DMEM / F-12 and fetal bovine serum (FBS) purchased from Gibco Life Technologies (Gaithersburg, MD) were purified and sterilized with a 0.22-mm low protein-binding cellulose acetate membrane filter purchased from Corning, Inc. (Corning, NY). Two monovalent cationic lipids, 1-[2-(9-(Z)-octadecenoyloxy)ethyl]-2-(8-(Z)heptadecenyl)-3-(hydroxyethyl)imidazolinium chloride (DOTIM) and 1,2-dimyristoyl-sn-glycero-3ethylphosphocholine (EDMPC), as well as two neutral lipids, cholesterol and diphytanoyl phosphatidylethanolamine (DipPE), were considered. Cationic liposomes extruded to a nominal diameter of 100 nm were a generous gift from Valentis, Inc. (Burlingame, CA). Nitric oxide synthase (NOS) plasmids (8.1 kb) were also a gift from Valentis, Inc. Green fluorescent protein (GFP) plasmids (4.7 kb) were purchased from Clontech Laboratories, Inc. (Palo Alto, CA), amplified in DH5a competent cells according to the manufacturer’s instructions (Gibco Life Technologies, Gaithersburg, MD), and purified using a Qiagen Plasmid Maxi Kit (Valencia, CA). The nucleic acid concentration and purity were measured by UV absorption at 260 and 280 nm. DNA verification by electrophoresis on 1% agarose gel confirmed plasmid size and indicated that more than 90% of the plasmids were supercoiled.

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2.2. Time resolved multiangle laser light scattering measurements TR-MALLS measurements were carried out with a DAWN DSP Laser Photometer (Wyatt Technology Inc., Santa Barbara, CA). Lipoplex formation reactions were conducted in 20-ml glass scintillation vials, which were carefully cleaned, rinsed with filtered deionized water, and oven dried to eliminate contaminants including dust and ions. Before each experiment, samples were checked for clarity by measuring the solvent background scattering at very low angles and were discarded if the background noise was too high. Lipoplex formation was initiated by adding cationic liposome dispersion to an existing DNA solution to a final volume of 5 ml. The solution was gently mixed to avoid bubble formation for a couple of seconds (approximately 2 s) prior to measurements. The DNA and liposome concentrations are indicated in the plots. All experiments were conducted at room temperature, approximately 23 8C, and repeated several times. The Rayleigh–Gans–Debye approximation was used to interpret the measured light scattering spectra. The method provides the light scattering-average lipoplex molar mass and geometric size. The analysis is greatly simplified in that the measured refractive index increments for DNA and cationic liposomes are nearly equal to one another. It should also be noted that owing to the very dilute solutions considered here, interparticle (lipoplex–lipoplex) interactions can be neglected. In the absence of interparticle interactions, the excess scattering or Rayleigh ratio is given by [22] Ru 5 K9c 2SC c SC MSC,LS PSC,LS (q) 1 K9c 2L c L ML,LS PL,LS (q) 2

1 K9c DNA c DNA MDNA,LS PDNA,LS (q)

where n s is the solvent refractive index, lo is the wavelength of the incident / scattered light and u is the scattering angle. The intraparticle interference factor contains the geometric size information accessible to this technique, namely the light scattering root mean square radius or radius of gyration. Owing to the near equality of the DNA and cationic liposome refractive index increments, the molar mass measured here is the weight average value while the geometric size is the z-average of the mean square radius or radius of gyration. The subscript ‘SC’ denotes a supramolecular complex or, in the system considered here, the DNA / cationic liposome lipoplex, the subscript ‘L’ represents the free cationic liposomes in solution and ‘DNA’ denotes free DNA in solution. The scattering contribution of the free plasmid DNA can be neglected in most cases since the DNA plasmid scattering contribution is insignificant and equivalent to background noise in comparison to the light scattering signal from the much larger molar mass cationic liposomes and lipoplexes. However, it should be noted that this might not be true when a large excess of DNA is present. This leads to a simplified description of the excess Rayleigh ratio Ru 5 K9c 2SC c SC MSC,LS PSC,LS (q) 1 K9c 2L c L ML,LS PL,LS (q)

(2)

Thus, the measured or apparent molar mass will reflect contributions of both the lipoplexes and free liposomes. Since the liposome molar masses are on the order of 10 7 g / mol, apparent molar masses$10 8 are indicative of the true lipoplex molar mass values. A much more detailed discussion of the method can be found in an earlier report on DNA / poly-L-lysine polyplex formation [22].

(1)

where Ru denotes the excess Rayleigh ratio, K9 is an optical constant, c is the mass concentration, c is the refractive index increment, MLS is the light scattering average molar mass and PLS (q) is the light scattering average intraparticle interference factor. The scattering wave vector, q, is given by 4p n s q 5 ]] sin(u / 2) lo

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3. Results and discussion

3.1. Lipoplex formation kinetics depend on charge ratio The formation of DNA / cationic liposome lipoplexes may be kinetically controlled [23]. An understanding of this process may provide insight for designing future generations of synthetic vectors.

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The formation kinetics are directly monitored in terms of the apparent lipoplex molar mass and geometric size as determined by TR-MALLS. Lipoplex formation kinetics were studied as a function of liposome formulation, DNA / cationic lipid charge ratio and cationic liposome and DNA concentration. The lipoplex formation kinetics were found to be primarily influenced by the DNA / cationic lipid charge ratio regardless of liposome formulation or DNA and cationic liposome concentration with two distinct regimes being noted. First, a growing kinetic regime wherein the initially formed primary lipoplexes further aggregate into ever larger lipoplexes that eventually reach a final, stable state. Second, an essentially kinetically static regime in which the initially formed primary lipoplexes are stable and do not undergo further aggregation and growth (Fig. 1). The growing kinetic regime was observed for DNA / cationic lipid charge ratios near unity. This regime is characterized by an increase in the lipoplex apparent molar mass and geometric size for several minutes following the initiation of complexation. This indicates that for charge ratios near unity, the initial primary DNA complexes must be unstable and undergo further aggregation. This secondary aggregation process is confirmed by the observation of the growing apparent molar mass and geometric size.

The aggregation process is complete when the apparent molar mass and geometric size values become essentially constant. As noted, the growing kinetic regime is completed within a few minutes following the addition of the cationic liposome dispersion to the DNA solution. This time scale is much shorter than that previously observed for DNA / poly-L-lysine polyplex formation [22]. The second type of lipoplex formation kinetics is observed at excess DNA or cationic liposome concentration. Under these conditions, TR-MALLS measurements indicate that the apparent lipoplex molar mass and geometric size rapidly reach steady-state values. Therefore, the lipoplex formation process is essentially instantaneous on the time scales accessible to the measurement technique and the primary lipoplexes are very stable against further aggregation. This was also previously observed for DNA / poly-L-lysine polyplex formation [22]. In order to capture the very rapid complexation kinetics at the same solution concentrations, a stop flow apparatus would be required to mix the two solutions. As it is not a trivial matter to construct a multiangle laser light scattering stop flow instrument, the best one could hope to do would be to use a 908 light scattering stop flow instrument [24]. Unfortunately, one requires multiple angles in order to extract quantitative molar mass and geometric size information. A possible solution to this dilemma would require one to consider very dilute DNA / cationic liposome solutions in the approach outlined here and elsewhere [22]. However, this scenario is limited by the low signal-to-noise ratios, which may prove impossible to overcome with existing instruments.

3.2. Initial lipoplex formation rate exhibits firstorder DNA concentration dependence

Fig. 1. The temporal behavior of the apparent molar mass for DNA complexing with DOTIM / cholesterol cationic liposomes is illustrated here. The DNA / cationic lipid charge ratios are as indicated in the plot. The total DNA concentration is kept fixed at 1.08 mg / ml. The DNA plasmid size is 8.1 kb. Similar trends are observed for the temporal behavior of the apparent geometric size.

The temporal evolution of the scattered light wavevector dependence for lipoplexes formed from NOS plasmids and DOTIM / cholesterol liposomes at a charge ratio of unity is shown in Fig. 2. It is readily apparent that the scattered intensity and wavevector variation increases with increasing time. The increasing scattered intensity corresponds to the lipoplex molar mass growth that occurs as the lipoplexes incorporate multiple plasmid copies. The observation that the wavevector variation also increases with

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Fig. 2. The temporal evolution of the scattered light wavevector dependence for DNA / DOTIM / cholesterol lipoplexes is shown. The times of aggregation plotted are (•) 01 min, (h) 0.2 min, (m) 0.3 min, (♦) 1 min, (.) 2 min, (^) 3 min, (s) 4 min, and (j) 5 min. The charge ratio is unity and the DNA concentration is fixed at 1.08 mg / ml. The DNA plasmid size is 8.1 kb.

time is indicative of the concurrent increase in the lipoplex geometric size. Eventually the light scattering signal plateaus indicating that the lipoplex formation process has come to steady-state. As the charge ratio deviates significantly from unity, the intensity does not vary significantly from the value found following the initial mixing, thereby indicating that the primary lipoplexes formed for charge ratios far from unity are stable against further aggregation. The same trends are observed for all liposome formulations and DNA concentrations considered in this study. A qualitative measure of the lipoplex formation rate can be gained by examining the time dependence of the scattered intensity at one scattering angle or wavevector. Initial intensity changes [dI / dt] t →0 at the 908 scattering angle were determined for NOS plasmid / DOTIM / cholesterol lipoplexes formed at various charge ratios and DNA concentrations. The results are shown in Fig. 3 and indicate that the initial lipoplex formation rates increase linearly with DNA (cationic liposome) concentration and are very dependent on charge ratio. As expected, the lipoplex formation rate reaches its maximum value at charge ratios near unity. Interestingly, as the charge ratio exceeds unity (i.e. an excess of DNA), the slowest lipoplex formation rate is observed. This lipoplex formation rate difference is primarily a

153

Fig. 3. The initial intensity change [dI / dt] t →0 at the 908 scattering angle were determined for DNA / DOTIM / cholesterol lipoplexes formed at various charge ratios and DNA concentrations. These are plotted as a function of the total DNA mass dissolved in 5 ml solution. The DNA / cationic lipid charge ratios are as indicated in the plot. The DNA plasmid size is 8.1 kb. Each line is labeled with its respective charge ratio. It is apparent that the lipoplex formation rate by DOTIM / cholesterol liposomes increases linearly with DNA concentration. The solid lines are linear fit through the kinetic rate data. Similar results were found for the other cationic liposome formulations considered in this report.

result of the fact that under conditions of excess DNA at fixed DNA concentration a smaller number of primary complexes are formed. When the cationic liposome mass is in excess, a greater number of primary complexes are formed. The primary complex aggregation rate is controlled by the number of primary complexes as well as the interparticle interactions occurring between them. At excess DNA, a relatively small number of negatively charged primary complexes are formed, which in turn aggregate very slowly if at all. On the other hand at excess cationic liposome concentration, the largest number of complexes are formed yet they are positively charged and therefore also resistant to aggregation. The aggregation rate is enhanced for DNA / cationic lipid charge ratios near unity in that a fairly large number of weakly charged to neutral complexes are formed that undergo further rapid aggregation.

3.3. Charge ratio influences final lipoplex properties Four different cationic liposome formulations were

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investigated as lipoplex formation agents at eight different DNA / cationic lipid charge ratios. Interestingly, at all the charge ratios considered, all four cationic liposome formulations exhibit similar lipoplex formation kinetics when combined with DNA solutions (Fig. 1). The final lipoplex molar mass and geometric size are most dependent on the DNA / cationic lipid charge ratio. To better illustrate this observation, the final values of the apparent molar mass and geometric size were determined for each DNA / cationic liposome pair considered here and are plotted as a function of the DNA / cationic lipid charge ratio in Figs. 4 and 5. The spread in the data reflects the fact that the DNA concentration was varied from 1.08 to 10.8 mg / ml (five concentrations were considered for each formulation) as well as the fact that the cationic liposomes considered here were not all the same mean size. In fact, the EDMPC / DipPE and DOTIM / DipPE liposomes exhibited significantly larger molar masses than the EDMPC / cholesterol and DOTIM / cholesterol liposomes. In addition, the spread in the measured geometric sizes is larger than that observed for the measured molar masses. This reflects the greater difficulty in determining the geometric size owing to the lipoplex

Fig. 4. The final apparent molar masses of lipoplexes formed from four different cationic liposome formulations (DOTIM / cholesterol (squares), DOTIM / DipPE (circles), EDMPC / cholesterol (diamonds), and EDMPC / DipPE (triangles)) at eight charge ratios and five DNA concentrations. The DNA concentration varies from 1.08 to 10.8 mg / ml with the smallest symbols corresponding to the lowest concentration and the symbol size increasing with increasing DNA concentration. The DNA plasmid size is 8.1 kb.

Fig. 5. The final apparent radii of gyration of lipoplexes formed from four different cationic liposome formulations (DOTIM / cholesterol (squares), DOTIM / DipPE (circles), EDMPC / cholesterol (diamonds), and EDMPC / DipPE (triangles)) at eight charge ratios and five DNA concentrations. The DNA concentration varies from 1.08 to 10.8 mg / ml with the smallest symbols corresponding to the lowest concentration and the symbol size increasing with increasing DNA concentration. The DNA plasmid size is 8.1 kb.

geometric sizes considered here. This is particularly true when the apparent lipoplex radius of gyration is #30 nm as the angular variation of the scattered intensity is very low and the determination of the small slopes is fraught with error. The strong qualitative agreement between the results found for the four different liposome formulations indicates that the DNA / cationic lipid charge ratio is the key parameter for understanding lipoplex formation regardless of cationic liposome formulation, liposome size and DNA or lipid concentration. It should be noted that similar molar mass and geometric size trends have been observed for DNA–poly-L-lysine polyplexes [22]. It is readily apparent that the lipoplex molar mass and geometric size exhibit maxima at a charge ratio near unity for each of the cationic liposome formulations and DNA concentrations considered. Below or above this charge ratio, the apparent molar mass and geometric size both decrease. At a charge ratio near unity, essentially every lipoplex may be neutral and therefore can undergo further aggregation. For charge ratios sufficiently greater than unity, the resulting lipoplexes may bear a net negative charge and are therefore stable against further aggregation.

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For DNA / cationic lipid charge ratios sufficiently smaller than unity, a net positive charge is expected and decreased apparent lipoplex molar masses and geometric sizes are observed. However, for the case of excess cationic liposomes, the measured apparent lipoplex molar masses and radii of gyration are strongly influenced by the presence of a significant amount of free cationic liposomes owing to their relatively large molar mass when compared to the free DNA molar mass. Under these conditions, it is most likely that both the lipoplex molar masses and radii of gyration are underestimated with the latter being especially underestimated since it represents a higher order moment of the size distribution. Therefore, an accurate estimate of the lipoplex molar mass and radii of gyration cannot be determined by MALLS for fairly small DNA / cationic lipid charge ratios. It should be noted that lipoplex systems are very different from polyplex systems in that the polyplexes are oftentimes formed from relatively low molar mass polyelectrolytes whose scattering is negligible in comparison with the free DNA and polyplexes that are present [22]. A similar trend has been observed for the hydrodynamic radius via dynamic light scattering [25,26], right angle light scattering turbidity [27,28] and has been predicted theoretically [29]. Thus, it appears that the physics which underlie DNA complexation by large cationic macroions, whether polymers or cationic liposomes, are essentially the same. Current research efforts in our lab are focused on developing combined fractionation-characterization schemes capable of resolving DNA complexes, DNA and free condensing agents such as cationic liposomes or polyelectrolytes in order to further explore these observations [30,31]. The unique ability of MALLS to determine the lipoplex molar mass as well as the geometric size allows a direct estimate of the lipoplex density to be made. The lipoplex density can provide a quantitative measure of the lipoplex structure in that the degree of DNA / lipid packing can be estimated [22]. Assuming solid sphere geometry, the lipoplex density is calculated as molar mass Lipoplex density 5 ]]]] 4 5 3/2 3 ] ] pR g 3 3

SD

(3)

where R g is the apparent lipoplex radius of gyration.

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The (5 / 3)3 / 2 factor arises from the fact that for a solid sphere R g 5(3 / 5)1 / 2 R where R is the sphere radius. As a point of reference, a solid sphere with a density of 1 g / cm 3 and radius of gyration of 100 nm would have a molar mass of approximately 1.23 10 10 g / mol. Since all four liposome formulations exhibited similar behavior, only the calculated density values for the DOTIM / cholesterol lipoplexes are tabulated in Table 1 along with the radii of gyration and molar masses. Once again, the large spread in lipoplex density values at low DNA / cationic lipid ratios (#0.5) is due to the difficulty of accurately determining the lipoplex radius of gyration under these conditions. This measurement error is further compounded by the fact that the radius of gyration enters the lipoplex density cubed. As noted previously, at low charge ratios both the lipoplex molar mass and geometric size are underestimated by MALLS with the latter quantity being underestimated to a larger degree. Therefore, the lipoplex densities are over-estimated under these conditions. After taking all of these facts into account, it is apparent that the lipoplex density maximum occurs near a charge ratio of unity just as observed for DNA / poly-L-lysine polyplexes [22]. The data presented in Table 1 also indicate that the DNA concentration has very little if any influence on the lipoplex properties for DNA concentrations ranging from 1.08 to 10.8 mg / ml. This was also observed to be the case for the other three liposome formulations considered.

3.4. Influence of solvent composition on lipoplex formation The solvent composition also affects the lipoplex formation process. Lipoplexes prepared in DMEM / F-12 media (a commonly used cell growth and transfection medium) exhibit larger lipoplex apparent molar masses and geometric sizes compared to the values found for lipoplexes prepared in 5 wt% dextrose at the same DNA / cationic lipid charge ratio and DNA concentration (Fig. 6). These higher apparent molar masses and geometric sizes are believed to be due to enhanced primary lipoplex aggregation promoted by the components present in the media such as divalent cations (Ca 21 , Mg 21 ), amino acids, and proteins. This solvent effect must be accounted for when lipoplex physicochemical

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Table 1 DOTIM / cholesterol a lipoplex structural properties DNA / lipid charge ratio

[DNA] (mg / ml)

Rg (nm)

Molar mass (g / mol)

Density (g / cm 3 )

1:4 1:4 1:4 1:4 1:4

1.08 2.16 3.24 5.40 10.8

43.2 49.9 40.9 47.5 56.7

5.93310 7 8.22310 7 7.17310 7 8.69310 7 9.48310 7

0.136 0.122 0.194 0.150 0.096

1:3 1:3 1:3 1:3 1:3

1.08 2.16 3.24 5.40 10.8

44.0 44.0 45.5 58.8 64.6

8.35310 7 9.53310 7 1.05310 8 1.24310 8 1.40310 8

0.181 0.206 0.206 0.112 0.096

1:2 1:2 1:2 1:2 1:2

1.08 2.16 3.24 5.40 10.8

51.9 47.2 58.6 60.7 71.1

1.49310 8 1.56310 8 2.28310 8 2.08310 8 2.30310 8

0.196 0.273 0.209 0.172 0.118

2:3 2:3 2:3 2:3 2:3

1.08 2.16 3.24 5.40 10.8

89.3 96.4 75.5 86.0 80.7

5.06310 8 6.16310 8 4.76310 8 5.28310 8 4.56310 8

0.131 0.127 0.204 0.153 0.160

1:1 1:1 1:1 1:1 1:1

1.08 2.16 3.24 5.40 10.8

93.0 92.2 87.5 98.9 97.0

6.43310 8 6.92310 8 5.90310 8 9.36310 8 8.95310 8

0.147 0.163 0.162 0.178 0.181

2:1 2:1 2:1 2:1 2:1

1.08 2.16 3.24 5.40 10.8

63.8 68.8 64.8 70.6 81.0

2.39310 8 1.90310 8 1.81310 8 2.41310 8 5.64310 8

0.170 0.107 0.123 0.126 0.195

4:1 4:1 4:1 4:1 4:1

1.08 2.16 3.24 5.40 10.8

56.4 64.3 59.3 59.6 69.9

5.79310 7 9.03310 7 7.08310 7 9.03310 7 1.13310 8

0.060 0.062 0.063 0.079 0.061

a

Similar tables for the other three formulations may be obtained from the corresponding author.

properties are used in the interpretation of in vitro transfection studies. As a means of ascertaining the origin of the enhanced aggregation, the influence of media composition on the cationic liposomes used in this study

Fig. 6. Lipoplexes prepared in DMEM / F12 media generally have larger apparent molar mass values when compared with those prepared in 5 wt% dextrose. This trend occurs at all charge ratios studied. The above data are for DNA / DOTIM / cholesterol lipoplexes. The total DNA concentration is fixed at 10 mg / ml. The DNA plasmid size is 4.7 kb. Similar trends are observed for the apparent geometric size.

was investigated. Further dilution in 5 wt% dextrose media had no effect on any of the cationic liposomes considered here which is not surprising in that the cationic liposomes themselves are extruded in 5 wt% dextrose media. However, when the DMEM / F-12 media was used to dilute the cationic liposomes, aggregation was observed. Liposomes incubated in similar media have also been observed to aggregate with no significant change in their surface charge [32]. In particular, serum has been found to interact strongly with positively charged liposomes compared to neutral and negatively charged liposomes and this interaction could lead to lipid oxidation, thereby causing the liposomes to become unstable [33]. An intensity plot at 908 scattering angle indicates that DOTIM / DipPE liposomes diluted in DMEM / F-12 media aggregate, while the same liposomes diluted in 5 wt% dextrose do not (Fig. 7). The DOTIM / DipPE liposomes exhibit extensive aggregation as indicated by the observed maximum in the scattered intensity that is a signature of colloidal aggregation processes [34]. However, sometimes changes due to dilution in media are subtle as illustrated for the case of DOTIM / cholesterol liposomes also shown in Fig. 7. Not surprisingly, DOTIM / DipPE lipoplexes pre-

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Fig. 7. The 908 scattered intensity is plotted to illustrate the changes that can occur when cationic liposomes are dispersed in DMEM / F12 media. DOTIM / DipPE (DD) liposomes exhibit large changes, while the other cationic liposome formulations considered here (DOTIM / cholesterol, EDMPC / cholesterol, and EDMPC / DipPE) exhibit only slight changes as illustrated here for the case of DOTIM / cholesterol (DC) liposomes.

157

ic regimes were observed. For charge ratios near unity, increasing lipoplex apparent molar masses and geometric sizes were observed for several minutes. This indicates for charge ratios near unity, initially formed primary lipoplexes were unstable and subsequently aggregated to form larger DNA complexes until reaching an apparently stable, final state. For charge ratios far from unity, the initially formed primary lipoplexes were stable and did not undergo further aggregation. Essentially the same phenomena were previously observed for DNA-poly-L-lysine polyplexes [22]. Thus, it appears that the physics which underlie DNA complexation by large cationic macroions, whether polymers or cationic liposomes, are essentially the same. The influence of solvent composition on complexation was also studied. DNA complexes prepared by diluting DNA and cationic liposomes separately in DMEM / F-12 media prior to mixing tend to form particles with higher apparent molar masses and geometric sizes compared to those prepared in 5 wt% dextrose. Thus, the components found in the DMEM / F-12 media, such as divalent salts and amino acids, enhance lipoplex aggregation.

pared in DMEM / F-12 media are much larger than lipoplexes derived from DOTIM / cholesterol liposomes in the same media.

Acknowledgements

4. Conclusions

The authors would like to acknowledge the support of Valentis, Inc. and the National Science Foundation via grant number CTS-9702413. In addition, the authors thank James D. Arroway for critically reading the manuscript.

The formation of DNA / cationic liposome lipoplexes was probed non-invasively using time-resolved multi-angle laser light scattering (TR-MALLS). The formation kinetics were monitored in terms of the lipoplex apparent molar mass and geometric size as determined absolutely by TR-MALLS. Lipoplexes were prepared from four different cationic liposome formulations. The effect of DNA / cationic lipid charge ratio and DNA concentration on the lipoplex formation kinetics, final molar mass and geometric size was investigated in detail. It was observed that the lipoplex formation kinetics and final lipoplex molar masses, geometric sizes and densities were primarily determined by the DNA / cationic charge ratio with the DNA concentration playing little or no role at all. Depending on the DNA / cationic lipid charge ratio, two distinctly different formation kinet-

References [1] P.L. Felgner, T.R. Gadek, M. Holm, R. Roman, H.W. Chan, M. Wenz, J.P. Northrop, G.M. Ringold, M. Danielsen, Lipofection—a highly efficient, lipid-mediated DNA-transfection procedure, Proc. Natl. Acad. Sci. USA 84 (1987) 7413–7417. [2] J.H. Felgner, R. Kumar, C.N. Sridhar, C.J. Wheeler, Y.J. Tsai, R. Border, P. Ramsey, M. Martin, P.L. Felgner, Enhanced gene delivery and mechanism studies with a novel series of cationic lipid formulations, J. Biol. Chem. 269 (1994) 2550–2561. [3] C.J. Wheeler, P.L. Felgner, Y.J. Tsai, J. Marshall, L. Sukhu, S.G. Doh, J. Hartikka, J. Nietupski, M. Manthorpe, M. Nichols, M. Plewe, X.W. Liang, J. Norman, A. Smith, S.H. Cheng, A novel cationic lipid greatly enhances plasmid DNA

158

[4]

[5]

[6]

[7]

[8]

[9]

[10]

[11]

[12]

[13]

[14]

[15]

[16]

[17]

E. Lai, J.H. van Zanten / Journal of Controlled Release 82 (2002) 149 – 158 delivery and expression in mouse lung, Proc. Natl. Acad. Sci. USA 93 (1996) 11454–11459. H. Gershon, R. Ghirlando, S.B. Guttman, A. Minsky, Mode of formation and structural features of DNA cationic liposome complexes used for transfection, Biochemistry 32 (1993) 7143–7151. J. Gustafsson, G. Arvidson, G. Karlsson, M. Almgren, Complexes between cationic liposomes and DNA visualized by cryo-TEM, Biochim. Biophys. Acta 1235 (1995) 305– 312. I. Koltover, T. Salditt, J.O. Radler, C.R. Safinya, An inverted hexagonal phase of cationic liposome–DNA complexes related to DNA release and delivery, Science 281 (1998) 78–81. D.D. Lasic, H. Strey, M.C.A. Stuart, R. Podgornik, P.M. Fredick, The structure of DNA–liposome complexes, J. Am. Chem. Soc. 119 (1997) 832–833. J.O. Radler, I. Koltover, T. Salditt, C.R. Safinya, Structure of DNA–cationic liposome complexes: DNA intercalation in multilamellar membranes in distinct interhelical packing regimes, Science 275 (1997) 810–814. B. Sternberg, K. Hong, W. Zheng, D. Papahadjopoulos, Ultrastructural characterization of cationic liposome–DNA complexes showing enhanced stability in serum and high transfection activity in vivo, Biochim. Biophys. Acta 1375 (1998) 23–35. H.E.J. Hofland, L. Shephard, S.M. Sullivan, Formation of stable cationic lipid / DNA complexes for gene transfer, Proc. Natl. Acad. Sci. USA 93 (1996) 7305–7309. T.J. Anchordoquy, J.F. Carpenter, D.J. Kroll, Maintenance of transfection rates and physical characterization of lipid / DNA complexes after freeze-drying and rehydration, Arch. Biochem. Biophys. 348 (1997) 199–206. S.D. Allison, T.J. Anchordoquy, Mechanisms of protection of cationic lipid–DNA complexes during lyophilization, J. Pharm. Sci. 89 (2000) 682–691. B. Li, S. Li, Y. Tan, D.B. Stolz, S.C. Watkins, L.H. Block, L. Huang, Lyophilization of cationic lipid–protamine–DNA (LPD) complexes, J. Pharm. Sci. 89 (2000) 355–364. N. Ballas, N. Zakai, I. Sela, A. Loyter, Liposomes bearing a quaternary ammonium detergent as an efficient vehicle for functional transfer of TMV-RNA into plant-protoplasts, Biochim. Biophys. Acta 939 (1988) 8–18. X. Zhou, A.L. Klibanov, L. Huang, Lipophilic polylysines mediate efficient DNA transfection in mammalian-cells, Biochim. Biophys. Acta 1065 (1991) 8–14. K. Hong, W.W. Zheng, A. Baker, D. Papahadjopoulos, Stabilization of cationic liposome–plasmid DNA complexes by polyamines and poly(ethylene glycol)–phospholipid conjugates for efficient in vivo gene delivery, FEBS Lett. 400 (1997) 233–237. P.C. Ross, M.L. Hensen, R. Supabphol, S.W. Hui, Multilamellar cationic liposomes are efficient vectors for in vitro gene transfer in serum, J. Liposome Res. 8 (1998) 499–520.

[18] C.R. Safinya, I. Koltover, J. Raedler, DNA at membrane surfaces: an experimental overview, Curr. Opin. Colloid Interface Sci. 3 (1998) 69–77. [19] S. Huebner, B.J. Battersby, R. Grimm, G. Cevc, Lipid–DNA complex formation: Reorganization and rupture of lipid vesicles in the presence of DNA as observed by cryoelectron microscopy, Biophys. J. 76 (1999) 3158–3166. [20] M.T. Kennedy, E.V. Pozharski, V.A. Rakhmanova, R.C. MacDonald, Factors governing the assembly of cationic phospholipid–DNA complexes, Biophys. J. 78 (2000) 1620– 1633. [21] I.S. Kikuchi, A.M. Carmona-Riberio, Interactions between DNA and synthetic cationic liposomes, J. Phys. Chem. B 104 (2000) 2829–2835. [22] E. Lai, J.H. van Zanten, Monitoring DNA / poly-L-lysine polyplex formation with time-resolved multiangle laser light scattering, Biophys. J. 80 (2001) 864–873. [23] V.A. Bloomfield, Condensation of DNA by multivalent cations—considerations on mechanism, Biopolymers 31 (1991) 1471–1481. [24] D. Porschke, Dynamics of DNA condensation, Biochemistry 23 (1984) 4821–4828. [25] Y. Xu, S.W. Hui, P. Frederik, F.C. Szoka, Physicochemical characterization and purification of cationic lipoplexes, Biophys. J. 77 (1999) 341–353. [26] E. Tomlinson, A.P. Rolland, Controllable gene therapy— Pharmaceutics of non-viral gene delivery systems, J. Controlled Release 39 (1996) 357–372. [27] N.J. Zuidam, Y. Barenholz, Characterization of DNA–lipid complexes commonly used for gene delivery, Int. J. Pharm. 183 (1999) 43–46. [28] N.J. Zuidam, Y. Barenholz, Electrostatic and structural properties of complexes involving plasmid DNA and cationic lipids commonly used for gene delivery, Biochim. Biophys. Acta 1368 (1998) 115–128. [29] R. Bruinsma, Electrostatics of DNA–cationic lipid complexes: isoelectric instability, Eur. Phys. J. B4 (1998) 75–88. [30] B.A. Korgel, J.H. van Zanten, H.G. Monbouquette, Vesicle size distributions measured by flow field-flow fractionation coupled with multiangle light scattering, Biophys. J. 74 (1998) 3264–3272. [31] H. Lee, S.K.R. Williams, S.D. Allison, T.J. Anchordoquy, Analysis of self-assembled cationic lipid–DNA gene carrier complexes using flow field-flow fractionation and light scattering, Anal. Chem. 73 (2001) 837–843. [32] K.K. Son, D.H. Patel, A. Park, Zeta potential of transfection complexes formed in serum-free medium call predict in vitro gene transfer efficiency of transfection reagent, Biochim. Biophys. Acta 1466 (2000) 11–14. [33] M. Foradada, M.D. Pujol, J. Bermudez, J. Estelrich, Chemical degradation of liposomes by serum components detected by NMR, Chem. Phys. Lipids 104 (2000) 133–148. [34] J.H. van Zanten, M. Elimelech, J. Colloid. Interf. Sci. 154 (1992) 1–7.