Increasing Surface Charge Density Induces Interdigitation in Vesicles

May 13, 2005 - of the phosphocholine headgroup, resulting in tighter packing of the acyl chains, which increases the thermal energy required for trans...
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Langmuir 2005, 21, 5707-5715

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Increasing Surface Charge Density Induces Interdigitation in Vesicles of Cationic Amphiphile and Phosphatidylcholine Samppa J. Ryha¨nen, Juha-Matti I. Alakoskela, and Paavo K. J. Kinnunen*,† Helsinki Biophysics and Biomembrane Group, Institute of Biomedicine/Biochemistry, Biomedicum, Post Office Box 63 (Haartmaninkatu 8), FIN-00014 University of Helsinki, Helsinki, Finland Received February 4, 2005. In Final Form: April 15, 2005 Binary vesicles of cationic lipid dihexadecyldimethylammoniumbromide (DHAB) and 1,2-dimyristoylsn-glycero-3-phosphocholine (DMPC) were examined by differential scanning calorimetry, fluorescence spectroscopy, and Fourier transform infrared spectroscopy. DHAB/DMPC vesicles demonstrate a complex dependence of the main-transition temperature (Tm) on their mole proportion of DHAB, with a maximum of 42 °C at XDHAB ) 0.4. An increase of Tm at XDHAB < 0.4 is explained by reorientation of P--N+ dipoles of the phosphocholine headgroup, resulting in tighter packing of the acyl chains, which increases the thermal energy required for trans f gauche isomerization. At XDHAB > 0.4, Coulombic repulsion between the cationic DHAB headgroups expands the bilayer evident as a decrease in Tm until a plateau of approximately 28 °C at 0.7 e XDHAB g 0.9 is reached, followed by an increment of Tm to approximately 30 °C at XDHAB > 0.9. The quenching of DPH-PC fluorescence emission and the decrease in the ratio of peak height intensities of symmetric and antisymmetric -CH2- stretching modes suggest an interdigitated phase to form at XDHAB > 0.6. Interdigitation allows the membrane to accommodate the augmented Coulombic repulsion between DHAB headgroups because of increasing cationic surface charge density while simultaneously causing tighter packing of the acyl chains evident first as a plateau at 0.7 e XDHAB g 0.9 and subsequently as an increase in Tm at XDHAB > 0.9. Screening of the membrane charges by NaCl abolishes the quenching of DPH emission and decreases Tm, thus revealing electrostatic repulsion as the driving force for interdigitation.

Introduction The completion of “the genome project” has opened the possibility for making gene therapy a practical utility. Accordingly, efficient and reliable means for introducing foreign genetic material into eukaryotic cells are needed to fully utilize the vast knowledge gathered about our genes. However, we are still far from having reached this goal. Viruses were initially considered as a natural first choice for this purpose, but their use is associated with some inherent problems, such as immunogenicity and a complicated manufacturing process. Recent reports1,2 have also raised questions about the safety of viral gene therapy and highlight the need for synthetic gene transfection systems. This is why cationic amphiphiles have received considerable attention in colloid and surface chemistry as well as molecular biology because their potential usage as nonviral transfection vectors is being investigated.3,4 More specifically, the latter types of vectors are complexes comprising of DNA and cationic lipids. These so-called “lipoplexes” have become very popular, and liposomal transfection, “lipofection”,4 is at present considered to be the most promising candidate for safe and reproducible * To whom correspondence should be addressed. Telephone: 3589-191 25400. Fax: 358-9-191 25444. E-mail: paavo.kinnunen@ helsinki.fi. † MEMPHYS, Center for Biomembrane Physics. (1) Alesci, S.; Ramsey, W. J.; Bornstein, S. R.; Chrousos, G. P.; Hornsby, P. J.; Benvenga, S.; Trimarchi, F.; Ehrhart-Bornstein, M. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 7484-7489. (2) Hacein-Bey-Abina, S.; von Kalle, C.; Schmidt, M.; Le Deist, F.; Wulffraat, N.; McIntyre, E.; Radford, I.; Villeval, J. L.; Fraser, C. C.; Cavazzana-Calvo, M.; Fischer, A. N. Engl. J. Med. 2003, 348, 255-256. (3) Zuhorn, I. S.; Hoekstra, D. J. Membr. Biol. 2002, 189, 167-179. (4) Felgner, P. L.; Gadek, T. R.; Holm, M.; Roman, R.; Chan, H. W.; Wenz, M.; Northrop, J. P.; Ringold, G. M.; Danielsen, M. Proc. Natl. Acad. Sci. U.S.A. 1987, 84, 7413-7417.

gene transfer into eukaryotic cells both in vitro and in vivo. The complexes are easy to prepare, practically nonimmunogenic, and nonpathogenic. Furthermore, cationic liposomes are able to complex and, more importantly, convey also other anionic biopolymers into cells. It seems possible to develop lipoplex technology also for applications involving anti-sense oligonucleotides5 and small inhibitory RNAs (siRNA)6-8 to fight cancer, hypercholesterolemia, and viral infections, for instance. Intense research efforts have been aimed to better understand the structure-function relationship correlating the biophysical properties of the lipoplexes to their transfection efficiency. Despite considerable progress made so far, optimization of lipofection for most applications remains empirical. The process leading to successful lipofection is thought to proceed through “barriers” that the lipoplex has to cross before the expression of the desired protein.9 The first prerequisite for lipofection is the electrostatically driven complex formation by DNA and cationic liposomes, which provides protection of the nucleic acid from, e.g., degrading enzymes by condensing it10 and enveloping it by a lipid layer. We have previously reported (5) Meyer, O.; Kirpotin, D.; Hong, K.; Sternberg, B.; Park, J. W.; Woodle, M. C.; Papahadjopoulos, D. J. Biol. Chem. 1998, 273, 1562115627. (6) Saksela, K. Trends Microbiol. 2003, 11, 345-347. (7) Davidson, B. L. N. Engl. J. Med. 2003, 349, 2357-2359. (8) Soutschek, J.; Akinc, A.; Bramlage, B.; Charisse, K.; Constien, R.; Donoghue, M.; Elbashir, S.; Geick, A.; Hadwiger, P.; Harborth, J.; John, M.; Kesavan, V.; Lavine, G.; Pandey, R. K.; Racie, T.; Rajeev, K. G.; Rohl, I.; Toudjarska, I.; Wang, G.; Wuschko, S.; Bumcrot, D.; Koteliansky, V.; Limmer, S.; Manoharan, M.; Vornlocher, H. P. Nature 2004, 432, 173-178. (9) Zabner, J.; Fasbender, A. J.; Moninger, T.; Poellinger, K. A.; Welsh, M. J. J. Biol. Chem. 1995, 270, 18997-19007. (10) Bloomfield, V. A. Curr. Opin. Struct. Biol. 1996, 6, 334-341.

10.1021/la0503303 CCC: $30.25 © 2005 American Chemical Society Published on Web 05/13/2005

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that surface charge density represents an important determinant both for the condensation of DNA and for efficient transfection.11 The latter was later confirmed by Lin and co-workers,12 and these authors further suggested membrane charge density to be an universal determinant of transfection behavior. Complexation with cationic liposomes also promotes internalization of nucleic acids by the cells, most likely via nonspecific charge-mediated endocytosis.3,9,13,14 After endocytosis, lipoplexes end up into endosomes and further in lysosomes, where low pH and nucleases would eventually destroy DNA. Accordingly, escape of the DNA from lipoplexes into the cytoplasmic space is required while the complex is still in the endosomal compartment and is likely to require neutralization of the positive charges in the lipoplex by anionic phospholipids in cellular membranes.15 Several recent reports have addressed the importance of endosomal escape, and it seems that lipoplexes containing lipids disrupting the endosomal membrane, e.g., by promoting the formation of the inverted hexagonal phase, have superior transfection efficiency.16-18 Thus, the mechanism of DNA release is closely related to the 3D phase behavior and the packing parameter19 of the lipids in the lipoplexes. The lack of detailed knowledge on the molecular level organization changing as a function of the surface charge density motivated a more comprehensive study on DHAB/ DMPC vesicles. In brief, we studied the impact of surface charge density on the organization of mixed cationic surfactant and phospholipid vesicles and their phase behavior by differential scanning calorimetry (DSC), fluorescence spectroscopy, and Fourier transform infrared spectroscopy (FTIR).

Ryha¨ nen et al. Preparation of Vesicles. Vesicles were prepared by mixing appropriate amounts of the lipid stock solutions in dry chloroform to obtain the desired compositions. Subsequently, the solvent was removed by evaporation under a stream of nitrogen. For removal of residual solvent, the samples were further maintained under high vacuum for at least 2 h. The resulting dry lipid films were then hydrated with 5 mM Hepes and 0.1 mM EDTA at pH 7.4 and thereafter incubated for 30 min at approximately 60 °C, i.e., above the temperatures of the transition endotherms of the lipid components and their mixtures. When indicated, the given concentrations of NaCl were present in the buffer used for hydration. DSC. After hydration, vesicles were vortexed and immediately loaded into the calorimeter cuvette (final concentration of 1 mM). A VP-DSC microcalorimeter (Microcal Inc., Northampton, MA) was operated at a heating rate of 0.5 °C/min. The instrument was interfaced to a 486 PC, and the data were analyzed using the routines of the software provided by the instrument manufacturer. Fluorescence Spectroscopy and Static Light Scattering. The fluorescence spectra, anisotropy, and static light scattering data were collected using Varian Cary Eclipse equipped with a Peltier thermostated cuvette compartment containing four cuvettes and a temperature probe immersed in a cuvette filled with Millipore water. For fluorescence experiments, DPH-PC was added to vesicles to yield a 1:500 molar ratio of the probe to lipid (XDPH-PC ) 0.002). After hydration, vesicles were vortexed and diluted in 5 mM Hepes and 0.1 mM EDTA at pH 7.4 to a final concentration of 50 µM. When indicated, NaCl was included in the buffer used for dilution of the vesicles. Changes in membrane acyl chain order were assessed by measuring the steady-state anisotropy r for DPH-PC.20,21 In brief, the fluorophore was excited at 345 nm, while anisotropy values were recorded at 450 nm with polarizers in the excitation and emission pathways. Corresponding band-passes were 5 and 10 nm. Anisotropy is defined by22

Materials and Methods Materials. 1,2-Dimyristoyl-sn-glycero-3-phosphocholine (DMPC), N-(2-hydroxyethyl)piperazine-N′-2-ethanesulfonic acid (Hepes), and ethylenediaminetetraacetic acid (EDTA) were from Sigma. 2-(3-(Diphenylhexatrienyl)propanoyl)-1-hexadecanoyl-snglycero-3-phosphocholine (DPH-PC) was from Molecular Probes (Eugene, OR); sodium chloride was from J. T. Baker (Deventer, Holland); and dihexadecyldimethylammoniumbromide (DHAB) was from Fluka. The purity of the above lipids was checked by thin-layer chromatography on silicic acid coated plates (Merck, Darmstadt, Germany) using chloroform/methanol/water (65:25: 4, by volume) as a solvent system. Examination of the plates after iodine staining or, when appropriate, by fluorescence illumination revealed no impurities. Concentrations of DMPC and DHAB were determined gravimetrically using a highprecision electrobalance (Cahn, Cerritos, CA), while the concentration of DPH-PC was determined by absorption at 355 nm ( ) 80 000 cm-1 M-1). Freshly deionized filtered water (Milli RO/Milli Q, Millipore Inc., Jaffrey, NH) was used in all experiments. (11) Ryha¨nen, S. J.; Sa¨ily, M. J.; Paukku, T.; Borocci, S.; Mancini, G.; Holopainen, J. M.; Kinnunen, P. K. J. Biophys. J. 2003, 84, 578587. (12) Lin, A. J.; Slack, N. L.; Ahmad, A.; George, C. X.; Samuel, C. E.; Safinya, C. R. Biophys. J. 2003, 84, 3307-3316. (13) Wrobel, I.; Collins, D. Biochim. Biophys. Acta 1995, 1235, 296304. (14) Hui, S. W.; Langner, M.; Zhao, Y. L.; Ross, P.; Hurley, E.; Chan, K. Biophys. J. 1996, 71, 590-599. (15) Kinnunen, P. K.; Ryto¨maa, M.; Koiv, A.; Lehtonen, J.; Mustonen, P.; Aro, A. Chem. Phys. Lipids 1993, 66, 75-85. (16) Smisterova, J.; Wagenaar, A.; Stuart, M. C.; Polushkin, E.; ten Brinke, G.; Hulst, R.; Engberts, J. B.; Hoekstra, D. J. Biol. Chem. 2001, 276, 47615-47622. (17) Koltover, I.; Salditt, T.; Ra¨dler, J. O.; Safinya, C. R. Science 1998, 281, 78-81. (18) Fielden, M. L.; Perrin, C.; Kremer, A.; Bergsma, M.; Stuart, M. C.; Camilleri, P.; Engberts, J. B. F. N. Eur. J. Biochem. 2001, 268, 1269-1279. (19) Israelachvili, J. N.; Marcelja, S.; Horn, R. G. Q. Rev. Biophys. 1980, 13, 121-200.

r)

I| - I⊥ I| + 2I⊥

where I| and I⊥ are the intensities of fluorescence emission recorded with an emission polarizer oriented parallel and perpendicular, respectively, to the direction of the vertically polarized excitation. G factor, the ratio of sensitivities of the instrument for vertically and horizontally polarized light, was determined before every measurement and was used to correct the anisotropy values.22 Emission spectra were collected in the range of 370-550 nm as above except without using polarizers in the optical pathways. Static light scattering for DHAB/DMPC/ DPH-PC liposomes was measured at an angle of 90°; the incident wavelength was set as 550 nm; and the integrated intensity of scattered light from a range of 540 to 560 nm was calculated. All fluorescence and light scattering measurements were done from at least three separate samples, and the results shown represent the averages of the measured values (( SD). FTIR. Appropriate amounts of DMPC and DHAB were mixed and dispersed into pure Millipore water to reach a total lipid concentration of 10 mg/mL. Subsequently, the solution was incubated for 30 min at 60 °C and vigorously vortexed every 10 min after which the solution was sonicated in a bath-type sonicator for 20 min. Approximately 1.5 mL of the resulting vesicle solution was loaded on a zinc selenide attenuated total reflectance (ATR) crystal that was temperature-controlled by an external circulating water bath (ThermoHaake, Karlsruhe, Germany). Vesicles were allowed to deposit on the crystal surface, and the temperature to stabilize the sample was kept at each defined temperature for approximately 15 min before recording spectra. The latter was collected by a Bruker EQUINOX 55 spectrometer (Bruker, Karlsruhe, Germany) using a mercury-cadmium(20) Lakowicz, J. R.; Prendergast, F. G.; Hogen, D. Biochemistry 1979, 18, 508-519. (21) Lakowicz, J. R.; Prendergast, F. G.; Hogen, D. Biochemistry 1979, 18, 520-527. (22) Lakowicz, J. R. In Principles of Fluorescence Spectroscopy; Kluwer Academic/Plenum Publishers: New York, 1999; p 698.

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Figure 1. DSC traces of DHAB/DMPC vesicles with indicated mole fractions XDHAB of the cationic lipid. The concentration was 1 mM in 5 mM Hepes and 0.1 mM EDTA at pH 7.4. The heating rate was 0.5 °C/min. The calibration bar represents 5 mJ/°C. telluride detector. The sample compartment was purged with dry air generated by an adsorber (Zander, Essen, Germany). A total of 1024 interferograms were co-added to yield each spectrum, and data were analyzed using dedicated software (OPUS) provided by the instrument manufacturer. The resolution of the FTIR spectra collected was 4 cm-1, and the frequencies of the IR bands were determined by a routine of OPUS software utilizing second derivative.

Results Thermal Phase Behavior of DHAB/DMPC Vesicles. DSC and fluorescence anisotropy r for DPH-PC20,21 were employed to determine the thermal phase behavior and estimate changes in the acyl chain order in the vesicles, respectively. In keeping with DSC data recorded earlier with a slightly different experimental setup,11 a sigmoidal dependency of Tm on XDHAB is evident in the thermograms, with a maximum of Tm at XDHAB ) 0.4 and a local minimum at XDHAB ) 0.9 (Figure 1). To better illustrate the effect of increasing charge density, the data gathered on thermal phase behavior of the binary DHAB/DMPC vesicles is compiled as a function of XDHAB (Figure 2). In the fluorescent probe used, the fluorescent DPH moiety is attached via a propionyl spacer to the sn-2 position of the glycerol backbone of the phospholipid, thus restricting its orientation to mainly align with the long axis of phospholipids. We recorded r for DHAB/DMPC vesicles with varying XDHAB and as a function of the temperature in the temperature range of 10-50 °C. In the course of the main-phase transition, the acyl chains become more mobile, evident as a decrease in r centered at Tm. Minima for the first derivatives dr/dT thus yield an approximation for Tm as a function of XDHAB. However, if the first derivative of the raw data was ambiguously defined, we utilized a visually acceptable polynomial fit for analysis. The obtained boundary of the phase diagram yields values for Tm that are in good

Figure 2. (A) Temperatures that exhibit the minima of first derivatives of DPH-PC fluorescence anisotropy versus temperature scans of DHAB/DMPC/DPH-PC vesicles (9) and maintransition temperatures (Tm) for DHAB/DMPC vesicles (O) plotted as a function of XDHAB. (B) Widths of the main-transition endotherms at the half-height (∆T1/2) for DHAB/DMPC vesicles plotted as a function of XDHAB. Both Tm and ∆T1/2 values were obtained from DSC data shown in Figure 1.

agreement with those determined by DSC (Figure 2A). Accordingly, Tm of neat DMPC vesicles is 23.2 °C determined by DPH-PC fluorescence and 23.7 °C determined by DSC. As XDHAB is increased, Tm progressively rises to a maximum of 42.7 °C (42.1 °C by DSC) at XDHAB ) 0.4. At XDHAB > 0.4, the value for Tm decreases until at XDHAB g 0.7 a plateau at approximately 28 °C is reached. Neat DHAB vesicles demonstrate somewhat higher Tm (Tm ) 33.8 °C from DPH fluorescence and 29.4 °C by DSC) compared to vesicles with XDHAB ) 0.9, resulting in a sigmoidal dependence of Tm on XDHAB. DSC demonstrates that also the cooperativity of transitions, evident as the width of the transition endotherms, changes as the surface charge density is varied. To illustrate this better, the widths of the endotherms at the half-height ∆T1/223 are shown as a function of XDHAB (Figure 2B). These data reveal two maxima in ∆T1/2, at XDHAB ) 0.1 and 0.6 (∆T1/2 ) 5.21 and 3.42 °C, respectively), indicating low cooperativity of the transition for these mixtures and a local minimum at XDHAB ) 0.4 with ∆T1/2 ) 1.07 °C. At XDHAB g 0.7, ∆T1/2 is low (between 0.43 and (23) McElhaney, R. N. Chem. Phys. Lipids 1982, 30, 229-259.

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0.71 °C), indicating higher cooperativity than in the mixture vesicles with a lower molar proportion of DHAB but still markedly higher values than for neat DMPC vesicles (∆T1/2 ) 0.17 °C), which exhibit the highest cooperativity. However, it should be noted that for the samples demonstrating marked phase separation, i.e., at XDHAB ) 0.6, 0.7, and 0.8, the ∆T1/2 is measured at the half-height of the highest endothermic peak and the wide shoulder at the higher temperatures is omitted (Figure 1). The overall cooperativity of chain melting in these samples is thus overestimated. Fluorescence Spectroscopy of DHAB/DMPC/DPHPC Vesicles. Changes in the environment of the DPHPC affect both the emission energy as well as the quantum yield (maximal emission intensity, Imax) of this fluorescent probe. To obtain a more detailed view on the molecular level changes in DHAB/DMPC liposomes in the different thermally induced phases, we recorded emission spectra for DPH-PC as a function of XDHAB and at selected temperatures (20, 35, and 48 °C) to cross phase boundaries revealed by DSC (Figure 3). XDHAB has a dramatic effect on the Imax (Figure 3A). When a mole fraction of DHAB is increased in DMPC vesicles, Imax decreases slowly until XDHAB ) 0.2. At 0.3 e XDHAB e 0.5, significant quenching is observed with a local minimum centered at XDHAB ) 0.4. After this minimum, Imax increases rapidly and reaches a local maximum at XDHAB ) 0.6. At XDHAB > 0.6, a nearly linear decrease in Imax as a function of XDHAB is evident that continues until XDHAB ) 1.0. It should be noted that the counterion of the cationic DHAB, bromide, is a collisional quencher and could thus contribute to the diminished fluorescence intensity. However, the concentration of Br- is small in the fluorescence experiments (maximally 50 µM at XDHAB ) 1.0), which makes this possibility unlikely. The wavelengths for the center of mass for the DPH emission peaks (λc) were calculated from the collected spectra and are depicted as a function of XDHAB (Figure 3B). DPH fluorescence shifts to shorter wavelengths when the surrounding environment becomes more hydrophobic, e.g., when less water penetrates into the hydrocarbon phase of the lipid bilayer.24 This is readily evident from our data. DPH in the fluid, disordered state vesicles at 48 °C exhibit λc in the range of 446-450 nm, in contrast to vesicles in the gel state at 20 °C with λc ≈ 441-445 nm. This is observed also for vesicles with 0.2 < XDHAB < 0.6 at 35 °C (i.e., in the gel state) having λc at approximately 441-446 nm. Interestingly, a shift in the emission maximum to shorter wavelengths, which is independent from the thermal phase behavior determined by fluorescence anisotropy and DSC (Figure 2), is seen at 0.1 < XDHAB e 0.3 and XDHAB ) 1 for both gel (at 20 °C) and fluid (at 50 °C) state vesicles (Figure 3B), most likely reflecting changes in the environment of DPH-PC because of composition-dependent reorganization of the DHAB/ DMPC vesicles. The steady-state anisotropy data for DPH are plotted as a function of XDHAB at 20, 35, and 50 °C in Figure 3C. At 20 °C, all vesicle compositions of this study are below their Tm (Figures 1 and 2) and are therefore in the gel state, in keeping with the high values for r. Similarly, at 50 °C, all vesicles are in the fluid state with low values for r. However, a local maximum in r at XDHAB ) 0.4 is observed, indicating tighter packing of the acyl chains. At XDHAB > 0.8 and at both 35 and 48 °C, an increase in r is (24) Lakowicz, J. R. Principles of fluorescence spectroscopy. In Solvent Effects on Emission Spectra; Lakowicz, J. R., Ed.; Kluwer Academic/ Plenum Publishers: New York, 1999; pp 185-210.

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Figure 3. (A) Maximal emission intensity of DPH-PC (Imax), (B) spectral center of the mass calculated from the emission spectra of DPH-PC (λc), and (C) emission anisotropy (r) recorded for DHAB/DMPC/DPH-PC vesicles depicted as a function of XDHAB. The temperature was 20 (9), 35 (b), and 48 °C (2). The total lipid concentration was 50 µM in 5 mM Hepes and 0.1 mM EDTA at pH 7.4.

evident, whereas at 20 °C, a decrement in r is observed for these vesicles. At 35 °C, composition-dependent changes in the phase state of the vesicles are evident. More specifically, at this temperature, vesicles with XDHAB ) 0.3, 0.4, and 0.5 are in the gel state, whereas vesicles with XDHAB ) 0.2 and 0.6 are in the main-transition regime and all other compositions are in the fluid state (Figures 1 and 2), thus revealing pronounced changes in r as a

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Figure 4. Maximal emission intensity of DPH-PC (Imax). DHAB/DMPC/DPH-PC vesicles shown as a function of the temperature. XDHAB was 0.4 (9) and 1.0 (O). The buffer was 5 mM Hepes and 0.1 mM EDTA at pH 7.4, and the total lipid concentration was 50 µM.

function of XDHAB. Importantly, because we did not determine the lifetime of the probe, these changes in r should in strict sense be taken as tentative only. However, in light of the perfect alignment of the anisotropy data with all other measurements on the system at hand, we may conclude r in this case to reflect changes in acyl chain order. It is expected that Imax decreases with temperature and especially when Tm is exceeded because of enhanced collisions of the fluorophore with water upon increasing average intermolecular areas. However, for vesicles with XDHAB from 0.2 to 0.4, Imax is lowest at 35 °C and increases at 48 °C (Figure 3A). Imax as a function of the temperature for vesicles with XDHAB ) 0.4 and 1.0 reveals different behavior of these two vesicle compositions demonstrating quenching of DPH emission (Figure 4). For XDHAB ) 0.4, Imax first decreases as the temperature is elevated but suddenly increases at T ≈ 42 °C, i.e., at Tm of these vesicles. This is contrasted by vesicles with XDHAB ) 1.0, which exhibit steadily decreasing Imax as a function of the temperature without discontinuities at Tm. To evaluate the role of surface electrostatics to DPH fluorescence, the same data were recorded for neat DHAB vesicles also in the presence of varying [NaCl] (Figure 5). Electrostatic screening upon [NaCl] increasing from 0 to 50 mM enhances Imax drastically at all temperatures, after which Imax decreases and reaches a plateau at [NaCl] ) 75-150 mM (Figure 5A). When [NaCl] is increased further to 200 mM, Imax decreases significantly at T ) 20 and 35 °C, while at 48 °C, it remains at approximately on the level measured at [NaCl] ) 150 mM. In these experiments, macroscopic aggregation of the vesicles because of the added salt was observed. To approximate the impact of aggregation on the Imax values, static light scattering was measured (Figure 5B). The effect of salt on Tm values of the DHAB vesicles was further assessed by analysis of r versus T scans similarly to the data presented earlier (Figure 2A) and depicted as a function of [NaCl] (Figure

Figure 5. (A) Maximal emission intensity of DPH-PC (Imax), (B) integrated scattered intensity (SI), and (C) temperatures that exhibit the minima of first derivatives of DPH-PC fluorescence anisotropy versus temperature scans recorded for DHAB/DPH-PC vesicles depicted as a function of [NaCl]. In A and B, temperatures were 20 (9), 35 (b), and 48 °C (2). The total lipid concentration was 50 µM in 5 mM Hepes and 0.1 mM EDTA at pH 7.4 with indicated [NaCl].

5C). For vesicles below Tm, i.e., at 20 °C, markedly increased scattering because of salt is observed until [NaCl] ) 150 mM after which intensity of scattered light significantly diminishes at [NaCl] ) 200 mM. As the temperature exceeds Tm, scattering diminishes rapidly and for fluid-state vesicles the intensity of scattered light is practically independent from [NaCl]. As [NaCl] is increased to 50 mM, the value for Tm diminishes to approximately 28.4 °C, subsequently reaching a minimum

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cence spectroscopy (Figures 1 and 2). Accordingly, when temperature is increased from 20 to 48 °C, a shift in the wavenumber of υsCH2 absorption from e2851 to g2853 cm-1 is observed for all vesicle compositions (Figure 6A). At 35 °C, the vesicles with XDHAB ) 0.2 and 0.4 are expected to be in the gel state based on DSC and fluorescence data, while other compositions are in the fluid state or the maintransition regime (XDHAB ) 0.6). In keeping with this, at 35 °C, a significant shift to lower wavenumbers is evident for the vesicles with XDHAB ) 0.2 and 0.4. However, also at XDHAB ) 1.0, a similar shift to lower wavenumbers was observed that was not expected from the fluorescence and DSC measurements. At XDHAB ) 0.6 and at 35 and 48 °C, a marked shift to higher wavenumbers was recorded and was absent in the data recorded for gel-state vesicles at 20 °C. Very similar data were obtained also for the υasCH2 mode (Figure 6B), with the characteristic shift to higher wavenumbers accompanying the main phase transition. In contrast to the data for the υsCH2 mode, the shift to the higher wavenumbers evident in the vesicles with XDHAB ) 0.6 was detected in all temperatures. Discussion

Figure 6. Wavenumber shift in -CH2- (A) symmetric and (B) asymmetric stretch IR absorption band as a function of XDHAB. The temperature was 20 (9), 35 (b), and 48 °C (2). Lipids were dissolved in pure Millipore water, and the total lipid concentration was 10 mg/mL.

(approximately 27.5 °C) at [NaCl] ) 150 mM. Upon further increase in [NaCl], the transition temperature increases, and at [NaCl] ) 200 mM, Tm is approximately 31 °C. FTIR of DHAB/DMPC Vesicles. To obtain a more detailed view of the phase behavior of DHAB/DMPC vesicles, we recorded FTIR spectra for six different vesicle compositions, viz. XDHAB ) 0.0, 0.2, 0.4, 0.6, 0.8, and 1.0. Shifts in the wavenumbers of the -CH2- symmetric and antisymmetric stretching modes (υsCH2 and υasCH2; parts A and B of Figure 6, respectively) were employed to monitor the trans f gauche isomerization of the lipid acyl chains and to detect the main phase transition. Because there are fewer overlapping absorption bands in the vicinity of the υsCH2 mode at ∼2850 cm-1, the latter is generally thought to more specifically reflect the lipid-phase behavior. Instead, other vibrational modes from methyl groups overlap with the υasCH2 mode at ∼2920 cm-1 in some phase states, making changes in these bends more difficult to interpret.25 The FTIR data on -CH2- vibrational modes are in keeping with the phase behavior of DHAB/DMPC vesicles determined by DSC and fluores(25) Lewis, R. N. A. H.; McElhaney, R. N. Infrared spectroscopy of biomolecules. In Fourier Transform Infrared Spectroscopy in the Study of Hydrated Lipids and Lipid Bilayer Membranes; Mantsch, H. H., Chapman, D., Eds.; Wiley-Liss, Inc.: Hoboken, NJ, 1996; pp 159-202.

The Tm of the binary system of cationic lipid DHAB and zwitterionic phospholipid DMPC reaches a maximum at XDHAB ) 0.4 (Figures 1 and 2). This somewhat counterintuitive behavior is readily explained by the reorientation of P--N+ dipoles of the phosphocholine headgroups. More specifically, positively charged headgroups of the cationic lipids pair with the anionic phosphates of P--N+ dipoles, and the Coulombic repulsion between the cationic headgroup and choline moiety reorients the P--N+ dipole from a parallel to a more vertical orientation. This electrostatically driven reorganization in the headgroup level has been demonstrated previously by NMR,26 Langmuir balance,27-29 DSC,11,30 and molecular dynamics simulations.31 Together with a reduced hydration of the headgroups because of charge neutralization of the phosphate,32,33 the reorientation of the phosphocholine moiety reduces its average area in the bilayer, allowing augmented chain-chain interactions evident as higher Tm. As previously reported in the DSC study by Silvius,30 pairing of DHAB with the PC headgroup also increases the cooperativity of the transition indicated by the sharp endotherms and, accordingly, local minimum in ∆T1/2 (Figure 2B). Importantly, in this same composition range, the area occupied by the lipids in monolayers has a minimum, providing direct evidence for the proposed mechanism.28 Ideally, one would expect Tm to reach a maximum at a 1:1 DHAB/phospholipid stoichiometry, i.e., when all cationic headgroups are paired with anionic phosphates of P--N+ dipoles. However, the bulky PC headgroup is likely to represent a sterical hindrance to the lateral organization of the membrane, and thus, the most efficient pairing of charges is achieved with a slight excess of DMPC, i.e., at 2:3 DHAB/DMPC. (26) Scherer, P. G.; Seelig, J. Biochemistry 1989, 28, 7720-7728. (27) Zantl, R.; Baicu, L.; Artzner, F.; Sprenger, I.; Rapp, G.; Ra¨dler, J. O. J. Phys. Chem. B 1999, 103, 10300-10310. (28) Sa¨ily, V. M. J.; Ryha¨nen, S. J.; Holopainen, J. M.; Borocci, S.; Mancini, G.; Kinnunen, P. K. J. Biophys. J. 2001, 81, 2135-2143. (29) Sa¨ily, V. M. J.; Alakoskela, J.; Ryha¨nen, S. J.; Karttunen, M.; Kinnunen, P. K. J. Langmuir 2003, 19, 8956-8963. (30) Silvius, J. R. Biochim. Biophys. Acta 1991, 1070, 51-59. (31) Bandyopadhyay, S.; Tarek, M.; Klein, M. L. J. Phys. Chem. B 1999, 103, 10075-10080. (32) Zuidam, N. J.; Barenholz, Y. Biochim. Biophys. Acta 1997, 1329, 211-222. (33) Hirsch-Lerner, D.; Barenholz, Y. Biochim. Biophys. Acta 1999, 1461, 47-57.

Interdigitated Phase in Vesicles of Cationic Amphiphile and PC

Fluorescence anisotropy of DPH-PC (Figure 3C) and the analysis of the symmetric and antisymmetric stretching modes for -CH2- (Figure 6), which are known to be sensitive to the thermal phase behavior are in accordance with the above results. The electrostatic reorganization in the headgroup region is evident also from the data on the values of λc for DPH-PC emission (Figure 3B). At 20 and 48 °C, where there are no thermally induced phase transitions interfering with the interpretation of the data, a shift to shorter wavelengths is observed at 0.2 e XDHAB e 0.4. The average access of water molecules in the hydrocarbon region of the membrane accommodating the fluorophore of DPH-PC34 should become more restricted as the headgroup region is condensed because of the reorientation of the P--N+ dipoles. Accordingly, as the average environment of the DPH becomes more hydrophobic, less energy is lost during the excited-state lifetime to solvent interactions and higher energy emission is evident as a shift to shorter wavelengths.24 After reaching a maximum at XDHAB ) 0.4, the value for Tm decreases gradually until at XDHAB > 0.6 a plateau and at XDHAB > 0.9 an increase are observed. A decrease of Tm is readily explained by the classical Gouy-Chapman theory because augmenting Coulombic repulsion between the DHAB headgroups results in an expansion of the bilayer and thus also reduced chain-chain interactions and lower Tm. However, why does the decrease in Tm stop at XDHAB > 0.6 and at XDHAB > 0.9 in which an increase is evident despite the increasing net charge density? A larger headgroup area compared to the area occupied by the acyl chains is known to favor either the micellar or interdigitated phase in the amphiphile aggregates.19,35 To this end, Haas and co-workers concluded from their X-ray diffraction data on DHAB and similar surfactants the thickness of the bilayer to be generally smaller than the length of the acyl chains.36 However, they did not provide a plausible interpretation for this observation. Similarly, Pohle et al. reported X-ray data suggesting a headgroup interdigitated phase in vesicles composed of the cationic lipid DMTAP with a high surface charge density.37 Interdigitation would provide a rational explanation also for our data as well as those of Silvius,30 demonstrating counterintuitive phase behavior at high surface charge densities. More specifically, interdigitation allows us to maximize the distance between the cationic headgroups of DHAB, while simultaneously resulting in a tighter packing of the acyl chains in the hydrophobic region. The suggested reorganizations at the critical mole proportions in DHAB/DMPC vesicles are schematically illustrated in Figure 7. Neutron scattering experiments have indicated the fluorophore containing sn-2 chain of DPH-PC to fluctuate between a conformation with the rodlike probe aligning the acyl chains and a conformation in which the probe resides in the surface, parallel to the plane of the bilayer.38 Because of the proximity of the fluorophore to the charged lipid headgroups in the latter conformation, significant reduction in the quantum yield can be expected. To this end, the quantum yield of DPH-PC has been shown to be (34) Kaiser, R. D.; London, E. Biochemistry 1998, 37, 8180-8190. (35) Pascher, I.; Lundmark, M.; Nyholm, P. G.; Sundell, S. Biochim. Biophys. Acta 1992, 1113, 339-373. (36) Haas, S.; Hoffmann, H.; Thunig, C.; Hoinkis, E. Colloid Polym. Sci. 1999, 277, 856-867. (37) Pohle, W.; Selle, C.; Gauger, D. R.; Zantl, R.; Artzner, F.; Ra¨dler, J. O. Phys. Chem. Chem. Phys. 2000, 2, 4642-4650. (38) Pebay-Peyroula, E.; Dufourc, E. J.; Szabo, A. G. Biophys. Chem. 1994, 53, 45-56.

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Figure 7. Schematic illustration of the DHAB/DMPC/DPHPC bilayer at XDHAB ) 0, 0.4, and 1.0 representing the suggested impact of increasing surface charge density on the molecular organization of the membrane. Red circles represent positive charges, and blue circles represent negative charges in the headgroups. The DPH moiety of the DPH-PC is depicted by green ovals.

sensitive to interdigitation of the bilayer39 because in the interdigitated bilayer the contained fluorophore becomes exposed to the aqueous phase and the vicinal headgroups in both possible conformations (Figure 7). Accordingly, the decrease in DPH-PC fluorescence observed at XDHAB > 0.6 (Figure 3A) would be in keeping with interdigitation. Furthermore, a nearly linear decrease in Imax suggests interdigitation to proceed as a first-order process, i.e., to involve a two-phase region. The formation of this phase would therefore be driven by an increasing surface charge density at XDHAB > 0.6 to a fully chain interdigitated lipid membrane at XDHAB ) 1.0. Interestingly, in the DSC traces recorded for vesicles with XDHAB ) 0.7 and 0.8, a sharp peak appears in the wide endotherm at a temperature very close to main endotherms at XDHAB ) 0.9 and 1.0. This phase separation together with the gradual decrease of DPH emission intensity implies that there are interdigitated domains in the DHAB/DMPC vesicles at XDHAB > 0.6, whose size increases until the percolation threshold, with the entire membrane becoming interdigitated at XDHAB ≈ 0.9. Notably, a decrease in fluorescence is not (39) Nambi, P.; Rowe, E. S.; McIntosh, T. J. Biochemistry 1988, 27, 9175-9182.

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Figure 8. Ratio of integrated peak intensities of symmetric and antisymmetric -CH2- stretching modes in DHAB/DMPC vesicles as a function of XDHAB. The temperature was 20 (9), 35 (b), and 48 °C (2). Lipids were dissolved in pure Millipore water, and the total lipid concentration was 10 mg/mL.

due to augmented light scattering because the latter does not change significantly as a function of XDHAB (data not shown). The minimum in Imax for DPH-PC at XDHAB ) 0.4 is compatible with tight association of the cationic headgroups of DHAB with the negatively charged phosphate moieties of DMPC. More specifically, the high Tm and r (r ≈ 0.35) as well as low ∆T1/2 suggest a very tightly packed acyl chain region of the bilayer at XDHAB ) 0.4. The relatively bulky DPH moiety of DPH-PC does not easily fit between tightly packed acyl chains, and thus, the conformation in which the DPH moiety is aligned parallel to the plane of the bilayer is favored (Figure 7). In comparison to normal fatty acid chains, this partitioning of DPH into the interface is also promoted by the polarization of the aromatic fluorophore. The measured diminished fluorescence is thus emitted by the relatively small population of DPH moieties located in the bilayer. To this end, the Imax values are significantly increased in vesicles with XDHAB from 0.3 to 0.5 at T > Tm, when more loose acyl chain packing allows the fraction of DPH moieties accommodated in the hydrocarbon phase of the bilayer to increase (Figure 4). This behavior is in contrast with the samples suggested to be in the interdigitated phase, i.e., with XDHAB > 0.6, that do not show changes in Imax at Tm but instead reveal a steady decrease of Imax as a function of the temperature (Figure 4). From their Raman spectroscopic study on interdigitated DMPC bilayers, O’Leary and Levin40 observed interdigitation to result in a decrease of the intensity ratio of the symmetric and antisymmetric methylene stretching modes. We calculated this ratio from our FTIR data (Figure 8). In accordance with fluorescence spectroscopy data suggesting interdigitation, there is an almost linear decrease in the I(υsCH2)/I(υasCH2) ratio at all temperatures when XDHAB > 0.6. The decrease in the I(υsCH2)/I(υasCH2) ratio accompanying the interdigitation could result from broadening and a concomitant decrease in the intensity of symmetric -CH2- stretching mode because of augmented interactions between neighboring acyl chains in the interdigitated phase. (40) O’Leary, T. J.; Levin, I. W. Biochim. Biophys. Acta 1984, 776, 185-189.

Ryha¨ nen et al.

Because interdigitation is assumed to be driven by an electrostatic repulsion between the cationic headgroups of DHAB, it was of interest to study how electrostatic screening by NaCl affects the data (Figure 5). As expected, adding salt to the vesicle solution increased Imax, suggesting the loss of interdigitation. Somewhat surprisingly at [NaCl] > 150 mM and at 20 and 35 °C, Imax again decreased. This behavior is, however, explained by aggregation of DHAB vesicles in the high salt concentration evident from the light scattering data (Figure 5B). Accordingly, because the electrostatic screening decreases Coulombic repulsion between the cationic charges of DHAB within the bilayer, evident as a loss of interdigitation, the repulsion between vesicles decreases, resulting in vesicle aggregation. At relatively low [NaCl], aggregation is evident as augmented light scattering for gel-state vesicles, whereas at [NaCl] > 150 mM, the aggregates are large, with precipitation evident as a decrease in both Imax and scattered intensity. The fluid-state vesicles have a smaller tendency for aggregation as suggested by low scattering values throughout the NaCl concentration range studied. To this end, also the Imax measured at [NaCl] > 150 mM for fluid DHAB vesicles (at 48 °C) is of the same magnitude as the values measured in lower [NaCl]. In keeping with loss of interdigitation and diminished chain-chain interactions, the Tm values determined from DPH-PC anisotropy versus temperature scans decrease at [NaCl] from 50 to 150 mM (Figure 5C). At [NaCl] ) 200 mM, Tm again increases in keeping with the GouyChapman theory because the efficient electrostatic screening reduces the effective headgroup area of DHAB with augmented acyl chain packing. In our previous study on DHAB/DMPC vesicles complexed with plasmid DNA, we demonstrated the surface charge density to be an important determinant for the transfection efficiency of these complexes in cultured cells.11 The impact of cationic charge density on lipofection was subsequently confirmed by Lin and co-workers.12 These authors utilized several cationic amphiphiles with varying valences and concluded cationic charge density to represent an universal determinant for efficient lipofection with lamellar systems. Interestingly, both interdigitation and high transfection efficiency are evident in the approximately same charge density range, i.e., at XDHAB > 0.5. However, transfection experiments were conducted at the physiological ionic strength ([NaCl] ) 150 mM), which is enough to prevent interdigitation (Figure 5A). Our preliminary experiments further suggest that also charge neutralization by DNA abolishes interdigitation (data not shown). However, the observed association between the interdigitated phase and efficient transfection could be due to enhanced endosomal escape. It is generally assumed that especially lamellar lipofection complexes enter the cells via endocytosis after which escape from the endosomes is required9,12 and is thought to be driven by formation of structures with negative curvature in the endosomal environment, such as the inverted HII phase.17,41 However, it has been shown that cationic gemini surfactants exhibiting vesicle-micelle transition at endosomal pH are efficient in transfection.18 In combination with our present findings, the report by Fielden and co-workers suggests that also amphiphiles with a propensity to form surfaces with positive curvature are able to disrupt the endosomal membrane and promote the transfer of plasmid DNA into cytoplasm. The data presented in this study provide evidence that a fundamental change in the phase behavior of mixed (41) Hafez, I. M.; Maurer, N.; Cullis, P. R. Gene Ther. 2001, 8, 11881196.

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cationic lipid and phosphatidylcholine vesicles takes place as the surface charge density is increased. Formation of interdigitated bilayers has been previously indicated for cationic liposomes of P-O-ethyl phosphocholines.42-45 However, the bulky headgroup of P-O-ethyl phosphocholines makes these lipids exceptional among the cationic amphiphiles, and the large surface area occupied by the headgroup could induce interdigitation even in the absence of a net positive charge. In contrast, DHAB has a relatively small headgroup that is unlikely to induce interdigitation as such without strong Coulombic repulsion between headgroups. This notion raises the intriguing possibility that formation of the interdigitated phase could be a general property of cationic liposomes providing that they have sufficient charge density and are in a low ionic strength medium. Strikingly, DHAB vesicles seem to be interdigitated also in the fluid state in contrast to the previous reports on the interdigitated phase because the Imax of the DPH-PC is unaffected at T > Tm (Figure 4).

reorientation at low contents of cationic lipid in phosphocholine matrixes. More importantly, our findings suggest the formation of the interdigitated phase in DHAB/ DMPC vesicles with XDHAB > 0.6 that is induced by Coulombic repulsion as the positive surface charge density is increased. The interdigitation is abolished by electrostatic screening when NaCl is added, thus indicating that Coulombic repulsion is indeed the driving force for interdigitation. Interestingly, DHAB vesicles seem to remain interdigitated also in the fluid state because Imax of DPH-PC is unaltered at T > Tm. Furthermore, DSC experiments demonstrate phase separation in DHAB/ DMPC vesicles, which together with a gradually decreasing Imax of DPH-PC and an I(υsCH2)/I(υasCH2) ratio suggest that there are interdigitated domains at XDHAB > 0.6. These domains are likely to grow with increasing surface charge density and eventually merge to form a totally interdigitated membrane.

Conclusions The present results further strengthen the evidence gathered for electrostatic phosphocholine headgroup

Acknowledgment. The authors thank Kristiina So¨derholm for technical assistance and Prof. Erik Goormaghtigh for critical reading of the manuscript. S. J. R. acknowledges grants from Finnish Medical Society Duodecim, Finnish Medical Foundation, and Farmos Research and Science Foundation. S. J. R. and J.-M. I. A. are supported by Helsinki Biomedical Graduate School. HBBG is supported by the Finnish Academy.

(42) Lewis, R. N.; Winter, I.; Kriechbaum, M.; Lohner, K.; McElhaney, R. N. Biophys. J. 2001, 80, 1329-1342. (43) Winter, I.; Pabst, G.; Rappolt, M.; Lohner, K. Chem. Phys. Lipids 2001, 112, 137-150. (44) Koynova, R.; MacDonald, R. C. Biophys. J. 2003, 85, 24492465. (45) Koynova, R.; MacDonald, R. C. Nano Lett. 2004, 4, 1475-1479.

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