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Microstructure study of liposomes decorated by hydrophobic magnetic nanoparticles
Chemistry and Physics of Lipids 165 (2012) 563–570
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Chemistry and Physics of Lipids journal homepage: www.elsevier.com/locate/chemphyslip
Microstructure study of liposomes decorated by hydrophobic magnetic nanoparticles Dan Qiu, Xueqin An ∗ , Zhiyun Chen, Xingyuan Ma School of Chemistry and Molecular Engineering, East China University of Science and Technology, 130 Meilong Road, Shanghai 200237, China
a r t i c l e
i n f o
Article history: Received 21 March 2012 Received in revised form 11 June 2012 Accepted 11 June 2012 Available online xxx Keywords: Magnetoliposomes Hydrophobic magnetic nanoparticles Lipid bilayer Morphology Microstructure
a b s t r a c t Magnetoliposomes, consisting of liposomes and magnetic nanoparticles (MNPs), have been tailored as very promising delivery vehicles in biotechnology and biomedicine applications. In this paper, liposomes with hydrophobic MNPs were prepared. The hydrophobic MNPs were successfully embedded in the lipid bilayer, which was proved by the results obtained from transmission electron microscope, atomic force microscope, differential scanning calorimetry and steady state fluorescence measurements. Moreover, systematic researches were carried out to investigate the effects of hydrophobic MNPs concentration on the morphology and microstructure of liposomes. The results show that the lipid bilayer was saturated with the hydrophobic MNPs when the mass ratio of MNPs to lipid reached 0.002. © 2012 Elsevier Ireland Ltd. All rights reserved.
1. Introduction Liposomes are artificially prepared spherically shaped objects consisting of a lipid bilayer surrounding an aqueous core. The cell membrane-like structure provides great capabilities of encapsulation, delivery and in vitro triggered release of a controlled dose of inclusions at a specific time and location (Park et al., 2006; Yatvin et al., 1978). The functional diversity of liposome renders them one of the most extensively developed drug delivery systems (Peer et al., 2007). In order to establish a drug delivery system that ensures stability, long-circulating and controllable release in vivo (Allen and Cullis, 2004), numerous studies have been carried out on liposomes decorated with nanoparticles (Al-Jamal and Kostarelos, 2007, 2011; An et al., 2010; Elersic et al., 2012; Mohanraj et al., 2010; Paasonen et al., 2010; Weng et al., 2008), polymers (Torchilin et al., 1994), coordination complexes (Kutsenko et al., 2011), antibodies (Ahmad et al., 1993; Lukyanov et al., 2004) etc. Among them, the designs of hybrid liposomes containing functional nanoparticles as nanoscale therapeutics in applications of imaging (Al-Jamal and Kostarelos, 2007; Martina et al., 2005; Mulder et al., 2006), biosensing (Zhao et al., 2006), hyperthermia treatment (Pradhan et al., 2007; Volodkin et al., 2009; Wu et al., 2008a) and controlledrelease (Chen et al., 2010; Viroonchatapan et al., 1997) have been intensively investigated.
∗ Corresponding author. Tel.: +86 21 64250804; fax: +86 21 64250804. E-mail address: [email protected] (X. An). 0009-3084/$ – see front matter © 2012 Elsevier Ireland Ltd. All rights reserved. http://dx.doi.org/10.1016/j.chemphyslip.2012.06.004
Magnetoliposomes (MLs) were the first multifunctional hybrid liposome/nanoparticle assembly (Bangham et al., 1958; Hamaguchi et al., 2003). Being an essential constitute of MLs, one of the critical functions of MNPs is to guide MLs to the target site in vivo (Hamaguchi et al., 2003; Kullberg et al., 2005). Moreover, MNPs make classic liposomes have several novel functions (Babincov et al., 1999; Hodenius et al., 2002; Shinkai et al., 1995; Skouras et al., 2011; Viroonchatapan et al., 1997) mentioned above. Regarding the structure, there have been two types of MLs: hydrophilic nanoparticles encapsulated in the core water of liposomes (MLs-W) (Martina et al., 2005; Pradhan et al., 2009; Skouras et al., 2011; Soenen et al., 2009; Viroonchatapan et al., 1997; Wijaya and Hamad-Schifferli, 2007), and hydrophobic nanoparticles embedded into the lipid bilayer (MLs-M) (Amstad et al., 2011; Chen et al., 2010; Nappini et al., 2011). MLs with the former structure are the major targets of numerous studies, while the latter are scarce. Although fewer attentions had been paid to MLs-M, some advantages of the structure in applications have already been noticed. High sensitivity and responsiveness of MLs-M for triggered drug release in low or high frequency magnetic fields were presented (Amstad et al., 2011; Nappini et al., 2011). The embedment of hydrophobic MNPs has ability to stabilize liposomes and suppress spontaneous leakage of cargo in core water (Chen et al., 2010). Except for these applications, few researches were carried out on the microstructure of MLs-M, especially the influence of hydrophobic MNPs on membrane structure. As the structures are of fundamental concern for applications, the aims of this work were to prepare magnetoliposomes enclosing hydrophobic MNPs into their membrane (Fig. 1), and to
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the residue was re-dispersed in a known volume of chloroform by ultrasound for 1 h. To obtain the individual MNPs, the synthesized nanoparticles were isolated from the microemulsion using ethanol. The MNPs were washed three times with ethanol and double distilled water, respectively.
2.3. Preparation of liposomes
Fig. 1. Schematic model for magnetolioposomes with hydrophobic MNPs in lipid bilayer.
characterize the influence of the MNPs on microstructure of liposomes. To realize the goals, hydrophobic ferriferous oxide nanoparticles dispersed in chloroform and liposomes with MNPs embedded in the bilayer were successfully prepared. Transmission electron microscope (TEM), atomic force microscope (AFM), differential scanning calorimetry (DSC) and fluorescent spectroscopy were applied to identify the presence of the hydrophobic MNPs in liposomes. Moreover, the effect of hydrophobic MNPs concentration on liposomes’ morphology, phase transition temperature, membrane fluidity and free volume were investigated in detail. These results may provide useful information not only for the MLs-M applications, but also for the development of liposomes encapsulating hydrophobic gold (Kojima et al., 2008; Park et al., 2006), silver (Bothun, 2008) nanoparticles, quantum (Al-Jamal et al., 2008) etc.
Blank liposomes (BLs) were prepared by the thin-film hydration method (Bangham et al., 1965). Briefly, chloroform–methanol (2:1, v/v) solution of lecithin was evaporated in a rotating round bottom flask, forming a phospholipid film on the flask walls. Then double distilled water was added into the flask and shaken for 30 min. The liposome suspension was prepared in a 60 ◦ C water bath for 1 h. After warm water bath preparation, the suspension was stirred for another 30 min, and left to cool to the room temperature. For MLsM, hydrophobic MNPs in chloroform were added into the lecithin chloroform–methanol solution as a component of the film formed on the flask walls. As for MLs-W, the phospholipid film was reconstructed with double distilled water in which MNPs were dispersed. Then the MLs suspension was treated by the same process as that used for the blank liposomes. The final lipid concentration in liposomes or magnetoliposomes suspension was 2 mg/mL.
2.4. Transmission electron microscope The samples for TEM measurement were obtained by dispersing a small drop of the suspension onto a copper grid pre-coated with amorphous carbon. Then the copper grid was dried in vacuum for 1 day before observation. The samples were characterized by H-7650 TEM (Hitachi High-Technologies, Tokyo, Japan) using an accelerating voltage of 80 kV.
2. Materials and methods 2.1. Materials Sodium bis (2-ethylhexyl) sulfosuccinate (AOT) from Sigma was dried under vacuum over P2 O5 until constant weight. Soybean lecithin (phosphatidylcholine 90%, iodine value 32, GengBen Biotechnology Shanghai Co., Ltd.) and the following commercially available chemicals were used without further purification: 1,6Diphenyl-1,3,5-hexatriene (DPH, Sigma), pyrene (Sigma), iron (II) sulfate heptahydrate (FeSO4 ·7H2 O, analytical reagent grades (AR), Sinopharm Chemical Reagent Co., Ltd (SCRC)), iron (III) chloride hexahydrate (FeCl3 ·6H2 O, AR, SCRC), sodium hydroxide (NaOH, AR, SCRC), 2,2,4-trimethylpentane (C8 H18 , AR, SCRC), chloroform (CHCl3 , AR, SCRC), and absolute methanol (CH4 O, AR, SCRC). 2.2. Synthesis of the magnetic nanoparticles Ferriferous oxide nanoparticles (Fe3 O4 , NPs) were synthesized with the microemulsion method (Liz et al., 1994; Wu et al., 2008b; Zhou et al., 2001). Iron (II) sulfate and iron (III) chloride (1.2:1, mol/mol) were dissolved in double distilled water. The solution was deoxygenated by bubbling N2 for about half an hour. The water/AOT/isooctane microemulsion of a fixed molar ratio (10:1) of water to AOT was prepared by mixing the aqueous iron solution prepared previously with 9 wt.% AOT isooctane solution. The microemulsion was heated to 78 ◦ C under N2 atmosphere. Then an aqueous NaOH solution was added dropwisely and the mixture was continuously stirred for 1 h. When the reaction was completed, the solvent of the mixture was removed by rotary evaporation, and
2.5. Atomic force microscope observation The lipid bilayer for AFM observation was prepared with the vesicle fusion method (Egawa and Furusawa, 1999; Kaasgaard et al., 2001). Liposomes prepared previously were sonicated to obtain the unilamellar liposomes. The freshly cleaved mica was buried in liposomes suspension for 5 h at 25 ◦ C. Then, the mica was washed ten times with double distilled water at the same temperature. All of the as-prepared films for AFM observation were dried in vacuum at room temperature to eliminate the last trace of solvent. The AFM topography images were obtained in the tapping mode by an AFM (AJ-III, Aijian Nanotechnology Inc., China) with triangularly shaped silicon cantilever with spring constant 48 N/m (Mikro Masch Co., Russia). A resonance frequency in the range of 240–400 kHz was used. The resonance peaks typically at 330 kHz in the frequency response of the cantilever were chosen for the tapping mode oscillation. The AFM images were obtained with a maximum scan range 20 m × 20 m and scanning frequencies were usually in the range between 0.6 and 2.5 Hz per line. The measurements were carried out in air at room temperature (Han et al., 2006).
2.6. Steady state fluorescence measurements DPH was applied as the membrane fluorescent probe in the steady-state anisotropy fluorescence measurement. Fluorescence anisotropy was measured using a fluorescence spectrometer (FLS920, Edinburgh Instruments, Edinburgh, UK) with polarization
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accessory unit. Steady-state fluorescence anisotropy, r, was calculated as: r=
I|| − gI⊥ I|| + 2gI⊥
Table 1 The anisotropy of DPH loaded in liposome membranes in 298 K and 318 K. T/K
Liposome type
Anisotropy of DPH (r)
298
BLs MLs-W MLs-M
0.0736 ± 0.00169 0.0761 ± 0.00138 0.0566 ± 0.00205
318
BLs MLs-W MLs-M
0.0427 ± 0.00092 0.0432 ± 0.00168 0.0405 ± 0.00175
in which g=
I⊥ I||
I⊥ and I|| are the perpendicular and parallel components of fluorescence observed corresponding to the plane of polarization of the excitation beam, respectively (Tang et al., 1998). DPH tetrahydrofuran solution was added into the lecithin chloroform–methanol solution when forming the phospholipid film. All samples with DPH were excited at 360 nm, and the emission light was observed at 430 nm. During the experiment, the turbidity of all samples was lower than 0.088 (absorbance units) and a 395 nm optical filter was applied to filtrate the stray light. The dwell time was 0.1 s and the scan speed was 1.0 nm/step. All samples were stirred for the whole time during the experiment by a magnetic bar equipped in the spectrometer cuvette at 15 and 45 ◦ C, respectively. Pyrene, another hydrophobic fluorescent probe, was added to the lecithin chloroform–methanol solution when forming the phospholipid film at the concentration 1 mol% of the lipid. The steady state fluorescence measurements were carried out at 15 and 45 ◦ C, respectively. Liposomes with pyrene were excited at 335 nm, the emission light was observed at 373 nm. Moreover, 350 nm optical filter was applied to filtrate the stray light. The dwell time was 0.1 s and the scan speed was 1.0 nm/step. Samples were stirred by a magnetic bar in the spectrometer cuvette through the fluorescence measurement. 2.7. Differential scanning calorimetry The calorimetric experiments were performed using a MicroDSC III (Setaram, Caluire, France). The samples were placed in a 1 mL sample cell in the calorimeter while double distilled water placed in another 1 mL reference cell was used as blank. The cells were scanned at a rate of 1 ◦ C/min from 4 to 60 ◦ C. The DSC thermograms were obtained and the phase transition temperature was characterized by the endothermic peak determined using SETSOFT 2000 software (Setaram Inc, France). 3. Results and discussion 3.1. Location of hydrophobic magnetic nanoparticles in liposomes As MNPs coated with AOT were applied as a component of liposomes, the exact location of the hydrophobic MNPs in liposomes was identified by the combination of TEM, AFM and fluorescence anisotropy spectroscopy. The MNPs shown in Fig. 2(a) are spherical in shape with the diameter of about 6 nm. The MNPs coated with AOT can be dispersed stably in chloroform after centrifugation at 12,000 rpm for 30 min. The stabilization of MNPs coated with AOT dispersed in chloroform was better than that of MNPs without the AOT coating, because of the formation of the surface layer on MNPs. TEM was also applied to characterize the morphology of liposomes with hydrophobic MNPs (Fig. 2(b)). Blank liposomes without heavymetal staining were invisible in TEM image, because lipid cannot deflect an electron beam sufficiently (Hall et al., 2007). For the liposomes with hydrophobic MNPs without staining, some dark dots arranging in circle shape were shown in Fig. 2(b). These dark dots, similar in size and morphology to MNPs in Fig. 2(a), were contributed by the hydrophobic MNPs. The circle arrangement of MNPs is mostly different from the random scatter arrangement in
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Fig. 2(a). The difference could be putatively attributed to the sphere shape of liposomes in which the MNPs were embedded. Hence the TEM results confirmed that the hydrophobic MNPs were successfully attached to liposomes. On the other hand, the morphology and size of hydrophobic MNPs-loaded liposomes can be evaluated indirectly by the arrangement of the embedded MNPs. In the TEM photograph, the morphology of MLs-M is spherical in shape and their average size is about 100 nm. The location of hydrophobic MNPs in liposomes was still ambiguous from the TEM results. Given the hydrophobicity of the MNPs surfaces and their strong preference for binding into a hydrophobic environment, the hydrophobic MNPs may be embedded into the membrane resulting in the potential surface topography changes of the lipid bilayer. AFM is a powerful technique for studying membrane surface features, and was applied to confirm the location of the MNPs. The lipid bilayer on mica plate was obtained by vesicle fusion method (Egawa and Furusawa, 1999; Kaasgaard et al., 2001). For blank lipid bilayer, the AFM topographic image was smooth and featureless as shown in Fig. 3(a). Furthermore, the thickness of the layer on mica plate revealed by the depth of the holes (cross section) was about 5 nm, which is similar to the reported thickness of lipid bilayer (Rosenberg et al., 1997). Nevertheless, in AFM image of the membrane with hydrophobic MNPs (Fig. 3(b)), apparent thicker and thinner membrane domains can be observed. Fig. 3(c) shows the smooth bilayer morphology of MLs-W because the hydrophilic MNPs were encapsulated in inner aqueous core. Considering the surface properties of membranes with hydrophilic and hydrophobic MNPs, the height fluctuation in Fig. 3(b) might be related to lipid bilayer distortion, attributed due to the embedment of hydrophobic MNPs in the lipid bilayer, which resulted in the potential changes of membrane’s surface. According to the TEM results, the diameter of MNPs in this work was about 6 nm which was bigger than the thickness of a single lipid bilayer (∼5 nm). Therefore lipid bilayer with embedded hydrophobic MNPs was about 0.7 nm thicker than that without hydrophobic MNPs. The difference, h, as illustrated in Fig. 3(b) led to the surface fluctuation when the membrane was unsaturated with the MNPs. A similar result concerning the membrane distortion induced by embedded hydrophobic nanoparticles was observed using the small angle neutron scattering method by Reimhult et al. (Amstad et al., 2011). These results confirmed that the hydrophobic MNPs were embedded into the lipid bilayer of liposomes. In addition to the morphology method above, the fluorescence anisotropy was applied to further verify the location of hydrophobic MNPs in liposomes. When a hydrophobic fluorescence probe is introduced into lipid bilayer, its rotation rate is restricted by membrane fluidity. The fluidity, which is sensitive to microstructure changes or additives within the membrane (Lande et al., 1995), is mostly revealed by the steady state fluorescence anisotropy of the hydrophobic probe (Tang et al., 1998; Van der Heide et al., 1996). In this case, diphenylhexatriene (DPH) was applied as the fluorescence probe. Generally, the relatively more flexible structure with better membrane fluidity gives rise to the higher rate of DPH’s rotational motion with lower anisotropy. As shown in Table 1, the DPH
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Fig. 2. TEM images of magnetic nanoparticles (a) and liposomes with hydrophobic MNPs (b). The inset shows enlarged view of the magnetoliposomes. The mass ratio of MNPs to lipid in magnetoliposomes was 0.002.
anisotropy for MLs-W is similar to that for BLs at a certain temperature, because the immediate environment of DPH in MLs-W was not influenced by the hydrophilic MNPs which were located in the inner aqueous core of liposomes. However, the anisotropy value of DPH in MLs-M is lower than that of BLs and MLs-W at same temperatures. Combined with the results above, that was because the hydrophobic MNPs were encapsulated within the lipid bilayer and disrupted the original regular arrangement of lipid molecules. Hence the membrane structure of MLs-M was less compact, and the DPH molecules could rotate easily among its surroundings, which gave rise to much lower anisotropy value. 3.2. Effects of MNPs concentration on the morphology and microstructure of liposomes The morphology of lipid bilayer with various hydrophobic MNPs concentration was investigated by AFM. The membrane morphology changed by varying the mass ratio of MNPs to lipid was shown in Fig. 4. The distortion of MLs-M membrane was observed, which could be due to the difference between the MNPs diameter and the
lipid bilayer thickness. With the increase of hydrophobic MNPs concentration, the thicker (light) domains related to the MNPs-loaded lipid bilayer became larger, whereas the thinner (dark) domains referring to lipid bilayer without MNPs became smaller. When the mass ratio of MNPs to lipid reached 0.002, a uniform and featureless bilayer surface could be detected again (Fig. 4(d)). This was because the lipid bilayer was saturated with hydrophobic MNPs. The calculated number of MNPs was about 4.26 × 1014 g−1 of lecithin at the saturation concentration. Tm describing the transition temperature of membrane from gel phase to liquid-crystalline phase can be characterized by the endothermic peak in DSC thermograms. Changes of Tm are sensitive to the additives in membrane and their subsequent effects on the alkyl chain organization. The Tm change depending on the MNPs concentration was shown in Fig. 5. The increase of MNPs concentration induced a reduction of Tm when the MNPs/lipid mass ratio was below 0.002 (Fig. 6). The decrease of Tm suggested that the inclusion of hydrophobic MNPs within the membrane can lead to bilayer disordering (Babincov et al., 1999), which was also proved by the DPH anisotropy method (Table 1). However, the Tm was almost constant
Fig. 3. AFM amplitude images for lipid bilayer of blank liposomes (a), MLs-M (b) and MLs-W (c). The mass ratios of MNPs to lipid in MLs-M or MLs-W were 0.001. The holes in (a) were about 5 nm depth. The inset in each figure shows the schematic diagram of lipid bilayer.
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Fig. 4. AFM amplitude images of the MLs membrane with different concentrations of MNPs. The mass ratios of MNPs to lipid are: (a) 0.0005; (b) 0.0010; (c) 0.0015; (d) 0.0020. The inset in each image shows the schematic diagram of the hydrophobic MNPs-loaded lipid bilayer.
Fig. 5. The thermograms of MLs-M as a function of hydrophobic MNPs concentration determined by differential scanning calorimetry (DSC).
Fig. 6. The phase transition temperature of lipid bilayer as a function of the MNPs concentration in hydrophobic MNPs-loaded liposomes.
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Fig. 8. Changes in fluorescence intensity ratio of pyrene (I1 /I3 ) in hydrophobic MNPs-embedded MLs as a function of increasing concentration of MNPs in gel phase (䊉) and liquid-crystalline phase ().
Fig. 7. The DPH anisotropy as a function of the MNPs concentration in hydrophobic MNPs-loaded liposomes at 15 ◦ C (䊉) and 45 ◦ C ().
when MNPs concentration was beyond 0.002. The result indicated that the lipid bilayer was saturated with MNPs at the concentration, which was consistent with AFM study in this work. The influence of the hydrophobic MNPs concentration on membrane fluidity was revealed by DPH anisotropy change at 15 and 45 ◦ C, respectively. As seen in Fig. 7, the DPH anisotropy in the membrane at 15 ◦ C was lower than that at 45 ◦ C, because the lipid bilayer was in an ordered gel phase with lower fluidity at lower temperature and in a disordered liquid-crystalline phase with higher fluidity at higher temperature. According to Fig. 7, the membrane fluidity in gel phase was improved by adding hydrophobic MNPs, which indicated that the MNPs inclusion disturbed the original rigid structure of lipid bilayer at temperature of 15 ◦ C. When the MNPs/lipid mass ratio was larger than 0.002, the anisotropy value of DPH was almost constant because of the saturation of hydrophobic MNPs in lipid bilayer. This result confirmed the existence of the saturation concentration of hydrophobic MNPs in membrane mentioned above. For MLs-M in liquid-crystalline phase at 45 ◦ C, the membrane fluidity was affected weakly by increasing MNPs concentration. The results could be attributed to the flexible structure of membrane in liquid-crystalline phase, so the disturbing effect of hydrophobic MNPs on membrane structure is insignificant. It suggest that, to some extent, adding hydrophobic MNPs into the membrane has the same effect as rising temperature, which could result in loosening of the membrane structure. This result provides the probability to control the membrane permeabilization by encapsulating hydrophobic MNPs at lower temperature. To gain further insight into the effect of hydrophobic MNPs concentration on membrane structure, pyrene which is located in the lipid bilayer was applied as fluorescence probe (Kutsenko et al., 2011; Shrivastava and Chattopadhyay, 2007). The intensity ratio (I1 /I3 ) of the first (373 nm) to the third (384 nm) peak in fluorescence emission spectrum of pyrene is an indicator of lipid bilayer polarity. Generally, low intensity ratio is indicative of low polarity in lipid bilayer. Fig. 8 shows that the membrane polarity decreases with the increasing MNPs concentration in both gel (15 ◦ C) and liquid-crystalline (45 ◦ C) phase. However, there was a slight decrease of membrane polarity when the MNPs/lipid mass ratio was beyond 0.002. It can be rationalized in terms of the
microstructure change within the nonpolar domain of lipid bilayer. As described above, the hydrophobic MNPs were coated with a layer of AOT, which is a surfactant having a similar structure as lipid. When embedded with the hydrophobic MNPs, lipid membrane could be divided into two pieces of bilayer (Fig. 9). The new bilayer was composed of a lipid layer and an AOT layer. The nonpolar domain was enlarged by the division, which facilitates the inclusion of more pyrene. It was reported that the intensity ratio (I1 /I3 ) of pyrene in membrane medium also depends on the probe concentration in nonpolar or polar areas (Ioffe and Gorbenko, 2005). More inclusion of pyrene in the expanded nonpolar area induces the decrease of polarity. When the MNPs/lipid mass ratio was beyond 0.002, the slight intensity ratio decrease could also be due to the saturation of MNPs in lipid bilayer. In addition, the polarity change in membrane of gel phase was more pronounced than that of liquid-crystalline phase, because water penetration into non-polar membrane region was increased in the relatively uncondensed structure in liquid-crystalline phase. This influence led to the increase in apparent polarity, which weakened the effect of hydrophobic MNPs inclusion on the nonpolar domain. Although the values of I1 /I3 decreased as the temperature increased from 15 ◦ C to 45 ◦ C, this change did not reflect a decrease of polarity. Because water, ethanol, and cyclohexane show a similar decrease of I1 /I3 when temperature increased (Szczupak et al., 2010; Zana et al., 1997). Pyrene is capable of forming a transient excited state complex, known as “excimer”, composed of an excited molecule and
Fig. 9. Schematic diagram of nonpolar area in the blank liposomes and the MNPsloaded liposomes.
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4. Conclusions
Fig. 10. Pyrene excimer (480 nm)/monomer (389 nm) fluorescence intensity ratio in hydrophobic MNPs-embedded MLs as a function of increasing concentration of MNPs in gel phase (䊉) and liquid-crystalline phase ().
a ground-state one (Ioffe and Gorbenko, 2005). The excimer-tomonomer intensity ratio (Iex /Iem ) is determined by measuring fluorescence intensity at the excimer (480 nm) and the monomer (389 nm) peaks, which is informative when detecting the structure change in membrane. In case of 1 mol% of pyrene incorporated into liposomes, the free volume model was applied to account for the pyrene excimerization in membranes (Ioffe and Gorbenko, 2005; L’Heureux and Fragata, 1989). In this model, the lateral displacement of hydrocarbon chains gives rise to appearance of voids where the pyrene excimer forms. Once the void was formed, it will be closed by the moving adjacent chains. If the movement is hindered, the final void, known as free volume, was enlarged. Hence, the higher Iex /Iem reflects the larger free volume in lipid bilayer. As can be seen from Fig. 10, the MNPs inclusion resulted in enlarged free volume in membrane when the mass ratio of MNPs to lipid was lower than 0.002 in both gel and liquid-crystalline phases. According to the DPH anisotropy results, the hydrophobic MNPs disrupted the rigid structure of the membrane. Therefore, the uncondensed structure gave rise to the appearance of more voids in membrane. Furthermore, as the hydrophobic MNPs are rigid additives, their existence in membrane prevented the adjacent lipid molecules to fill the void because of the thermal motion. The hindrance effect provided larger volume for pyrene excimer formation. However, the sustained presence of MNPs beyond 0.002 had different effect on the free volume. Consistent with the results above, the saturation concentration of MNPs in lipid bilayer was detected again. Beyond the concentration threshold, Iex /Iem of pyrene in lipid bilayer at two different temperatures followed two different trends: a plateau reached in gel phase (15 ◦ C), whereas the Iex /Iem decreased in liquid-crystalline phase (45 ◦ C). The unembedded AOT-coated MNPs were dispersed in aqueous water, when the membrane was saturated with MNPs. The AOT molecular was physically absorbed on the surface of MNPs. Since the solvent was changed from the hydrophobic organic solvents to water during the preparation process of MLs-M, the reversibly adsorbed AOT dissociated from the MNPs surface and diffused into the water (Amstad et al., 2009, 2011). Because of the rigid structure of membrane in gel phase, the influence of extra AOT molecule on membrane’s structure was weak. While in the liquid-crystalline phase, the looser structure enabled the extra AOT’s hydrophobic alkyl chains insert into the membrane. The insertion of alkyl chains occupied the existed voids, which reduced the existent free volume and gave rise to a decreasing Iex /Iem value beyond the concentration threshold.
Hydrophobic MNPs coated with AOT were synthesized and embedded into the bilayer of liposomes. The influence of MNPs on the membrane morphology was revealed by TEM and AFM methods. The results showed that the MLs-M were spherical in shape with the diameter of about 100 nm and the lipid bilayer could be distorted to accommodate hydrophobic MNPs, which led to the fluctuations of membrane surface. The effect of hydrophobic MNPs on the microstructure of liposomes was investigated by the DSC and steady state fluorescence measurements. The hydrophobic MNPs in the lipid bilayer not only improved membrane fluidity but also enlarged the nonpolar domain and the free volume in membrane. The lipid phase transition temperature decreased, when the hydrophobic MNPs were embedded in the lipid membrane. Moreover, the concentration of hydrophobic MNPs played an important role in membrane microstructure changes. Several results showed that the lipid bilayer was saturated with the hydrophobic MNPs when the mass ratio of MNPs to lipid reached 0.002. Based on these results, the encapsulation of hydrophobic nanoparticles in the membrane can be applied to modulate the membrane phase behavior and to control the membrane permeabilization. Further studies are needed to develop the application of the MLs-M as a drug delivery system. Acknowledgments This work is financially supported by the National Nature Science Foundation of China (20673059, 20573056 and 21073063), and Nature Science Keystone Foundation of Shanghai (08jc1408100), the National High-Tech Research and Development Plan of China (“863” plan, No. 2011AA06A107) and the Fundamental Research Funds for the Central Universities, China (No. WK0913002). References Ahmad, I., et al., 1993. Antibody-targeted delivery of doxorubicin entrapped in sterically stabilized liposomes can eradicate lung cancer in mice. Cancer Research 53, 1484–1488. Al-Jamal, W.T., et al., 2008. Lipid-quantum dot bilayer vesicles enhance tumor cell uptake and retention in vitro and in vivo. ACS Nano 2, 408–418. Al-Jamal, W.T., Kostarelos, K., 2007. Liposome–nanoparticle hybrids for multimodal diagnostic and therapeutic applications. Nanomedicine 2, 85–98. Al-Jamal, W.T., Kostarelos, K., 2011. Liposomes: from a clinically established drug delivery system to a nanoparticle platform for theranostic nanomedicine. Accounts of Chemical Research 44, 1094–1104. Allen, T.M., Cullis, P.R., 2004. Drug delivery systems: entering the mainstream. Science 303, 1818–1822. Amstad, E., Gillich, T., Bilecka, I., Textor, M., Reimhult, E., 2009. Ultrastable iron oxide nanoparticle colloidal suspensions using dispersants with catechol-derived anchor groups. Nano Letters 9 (12), 4042–4048. Amstad, E., et al., 2011. Triggered release from liposomes through magnetic actuation of iron oxide nanoparticle containing membranes. Nano Letters 11, 1664–1670. An, X., et al., 2010. Photoinduced drug release from thermosensitive AuNPs–liposome using a AuNPs-switch. Chemical Communications 46, 7202–7204. Babincov, M., et al., 1999. Laser triggered drug release from magnetoliposomes. Journal of Magnetism and Magnetic Materials 194, 163–166. Bangham, A.D., et al., 1958. An apparatus for microelectrophoresis of small particles. Nature 182, 642–644. Bangham, A.D., et al., 1965. Diffusion of univalent ions across the lamellae of swollen phospholipids. Journal of Molecular Biology 13, 238–252. Bothun, G., 2008. Hydrophobic silver nanoparticles trapped in lipid bilayers: size distribution, bilayer phase behavior, and optical properties. Journal of Nanobiotechnology 6, 13–23. Chen, Y., et al., 2010. Controlled release from bilayer-decorated magnetoliposomes via electromagnetic heating. ACS Nano 4, 3215–3221. Egawa, H., Furusawa, K., 1999. Liposome adhesion on mica surface studied by atomic force microscopy. Langmuir 15, 1660–1666. Elersic, K., et al., 2012. Electric-field controlled liposome formation with embedded superparamagnetic iron oxide nanoparticles. Chemistry and Physics of Lipids 165, 120–124.
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Contents lists available at SciVerse ScienceDirect
Chemistry and Physics of Lipids journal homepage: www.elsevier.com/locate/chemphyslip
Microstructure study of liposomes decorated by hydrophobic magnetic nanoparticles Dan Qiu, Xueqin An ∗ , Zhiyun Chen, Xingyuan Ma School of Chemistry and Molecular Engineering, East China University of Science and Technology, 130 Meilong Road, Shanghai 200237, China
a r t i c l e
i n f o
Article history: Received 21 March 2012 Received in revised form 11 June 2012 Accepted 11 June 2012 Available online xxx Keywords: Magnetoliposomes Hydrophobic magnetic nanoparticles Lipid bilayer Morphology Microstructure
a b s t r a c t Magnetoliposomes, consisting of liposomes and magnetic nanoparticles (MNPs), have been tailored as very promising delivery vehicles in biotechnology and biomedicine applications. In this paper, liposomes with hydrophobic MNPs were prepared. The hydrophobic MNPs were successfully embedded in the lipid bilayer, which was proved by the results obtained from transmission electron microscope, atomic force microscope, differential scanning calorimetry and steady state fluorescence measurements. Moreover, systematic researches were carried out to investigate the effects of hydrophobic MNPs concentration on the morphology and microstructure of liposomes. The results show that the lipid bilayer was saturated with the hydrophobic MNPs when the mass ratio of MNPs to lipid reached 0.002. © 2012 Elsevier Ireland Ltd. All rights reserved.
1. Introduction Liposomes are artificially prepared spherically shaped objects consisting of a lipid bilayer surrounding an aqueous core. The cell membrane-like structure provides great capabilities of encapsulation, delivery and in vitro triggered release of a controlled dose of inclusions at a specific time and location (Park et al., 2006; Yatvin et al., 1978). The functional diversity of liposome renders them one of the most extensively developed drug delivery systems (Peer et al., 2007). In order to establish a drug delivery system that ensures stability, long-circulating and controllable release in vivo (Allen and Cullis, 2004), numerous studies have been carried out on liposomes decorated with nanoparticles (Al-Jamal and Kostarelos, 2007, 2011; An et al., 2010; Elersic et al., 2012; Mohanraj et al., 2010; Paasonen et al., 2010; Weng et al., 2008), polymers (Torchilin et al., 1994), coordination complexes (Kutsenko et al., 2011), antibodies (Ahmad et al., 1993; Lukyanov et al., 2004) etc. Among them, the designs of hybrid liposomes containing functional nanoparticles as nanoscale therapeutics in applications of imaging (Al-Jamal and Kostarelos, 2007; Martina et al., 2005; Mulder et al., 2006), biosensing (Zhao et al., 2006), hyperthermia treatment (Pradhan et al., 2007; Volodkin et al., 2009; Wu et al., 2008a) and controlledrelease (Chen et al., 2010; Viroonchatapan et al., 1997) have been intensively investigated.
∗ Corresponding author. Tel.: +86 21 64250804; fax: +86 21 64250804. E-mail address: [email protected] (X. An). 0009-3084/$ – see front matter © 2012 Elsevier Ireland Ltd. All rights reserved. http://dx.doi.org/10.1016/j.chemphyslip.2012.06.004
Magnetoliposomes (MLs) were the first multifunctional hybrid liposome/nanoparticle assembly (Bangham et al., 1958; Hamaguchi et al., 2003). Being an essential constitute of MLs, one of the critical functions of MNPs is to guide MLs to the target site in vivo (Hamaguchi et al., 2003; Kullberg et al., 2005). Moreover, MNPs make classic liposomes have several novel functions (Babincov et al., 1999; Hodenius et al., 2002; Shinkai et al., 1995; Skouras et al., 2011; Viroonchatapan et al., 1997) mentioned above. Regarding the structure, there have been two types of MLs: hydrophilic nanoparticles encapsulated in the core water of liposomes (MLs-W) (Martina et al., 2005; Pradhan et al., 2009; Skouras et al., 2011; Soenen et al., 2009; Viroonchatapan et al., 1997; Wijaya and Hamad-Schifferli, 2007), and hydrophobic nanoparticles embedded into the lipid bilayer (MLs-M) (Amstad et al., 2011; Chen et al., 2010; Nappini et al., 2011). MLs with the former structure are the major targets of numerous studies, while the latter are scarce. Although fewer attentions had been paid to MLs-M, some advantages of the structure in applications have already been noticed. High sensitivity and responsiveness of MLs-M for triggered drug release in low or high frequency magnetic fields were presented (Amstad et al., 2011; Nappini et al., 2011). The embedment of hydrophobic MNPs has ability to stabilize liposomes and suppress spontaneous leakage of cargo in core water (Chen et al., 2010). Except for these applications, few researches were carried out on the microstructure of MLs-M, especially the influence of hydrophobic MNPs on membrane structure. As the structures are of fundamental concern for applications, the aims of this work were to prepare magnetoliposomes enclosing hydrophobic MNPs into their membrane (Fig. 1), and to
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the residue was re-dispersed in a known volume of chloroform by ultrasound for 1 h. To obtain the individual MNPs, the synthesized nanoparticles were isolated from the microemulsion using ethanol. The MNPs were washed three times with ethanol and double distilled water, respectively.
2.3. Preparation of liposomes
Fig. 1. Schematic model for magnetolioposomes with hydrophobic MNPs in lipid bilayer.
characterize the influence of the MNPs on microstructure of liposomes. To realize the goals, hydrophobic ferriferous oxide nanoparticles dispersed in chloroform and liposomes with MNPs embedded in the bilayer were successfully prepared. Transmission electron microscope (TEM), atomic force microscope (AFM), differential scanning calorimetry (DSC) and fluorescent spectroscopy were applied to identify the presence of the hydrophobic MNPs in liposomes. Moreover, the effect of hydrophobic MNPs concentration on liposomes’ morphology, phase transition temperature, membrane fluidity and free volume were investigated in detail. These results may provide useful information not only for the MLs-M applications, but also for the development of liposomes encapsulating hydrophobic gold (Kojima et al., 2008; Park et al., 2006), silver (Bothun, 2008) nanoparticles, quantum (Al-Jamal et al., 2008) etc.
Blank liposomes (BLs) were prepared by the thin-film hydration method (Bangham et al., 1965). Briefly, chloroform–methanol (2:1, v/v) solution of lecithin was evaporated in a rotating round bottom flask, forming a phospholipid film on the flask walls. Then double distilled water was added into the flask and shaken for 30 min. The liposome suspension was prepared in a 60 ◦ C water bath for 1 h. After warm water bath preparation, the suspension was stirred for another 30 min, and left to cool to the room temperature. For MLsM, hydrophobic MNPs in chloroform were added into the lecithin chloroform–methanol solution as a component of the film formed on the flask walls. As for MLs-W, the phospholipid film was reconstructed with double distilled water in which MNPs were dispersed. Then the MLs suspension was treated by the same process as that used for the blank liposomes. The final lipid concentration in liposomes or magnetoliposomes suspension was 2 mg/mL.
2.4. Transmission electron microscope The samples for TEM measurement were obtained by dispersing a small drop of the suspension onto a copper grid pre-coated with amorphous carbon. Then the copper grid was dried in vacuum for 1 day before observation. The samples were characterized by H-7650 TEM (Hitachi High-Technologies, Tokyo, Japan) using an accelerating voltage of 80 kV.
2. Materials and methods 2.1. Materials Sodium bis (2-ethylhexyl) sulfosuccinate (AOT) from Sigma was dried under vacuum over P2 O5 until constant weight. Soybean lecithin (phosphatidylcholine 90%, iodine value 32, GengBen Biotechnology Shanghai Co., Ltd.) and the following commercially available chemicals were used without further purification: 1,6Diphenyl-1,3,5-hexatriene (DPH, Sigma), pyrene (Sigma), iron (II) sulfate heptahydrate (FeSO4 ·7H2 O, analytical reagent grades (AR), Sinopharm Chemical Reagent Co., Ltd (SCRC)), iron (III) chloride hexahydrate (FeCl3 ·6H2 O, AR, SCRC), sodium hydroxide (NaOH, AR, SCRC), 2,2,4-trimethylpentane (C8 H18 , AR, SCRC), chloroform (CHCl3 , AR, SCRC), and absolute methanol (CH4 O, AR, SCRC). 2.2. Synthesis of the magnetic nanoparticles Ferriferous oxide nanoparticles (Fe3 O4 , NPs) were synthesized with the microemulsion method (Liz et al., 1994; Wu et al., 2008b; Zhou et al., 2001). Iron (II) sulfate and iron (III) chloride (1.2:1, mol/mol) were dissolved in double distilled water. The solution was deoxygenated by bubbling N2 for about half an hour. The water/AOT/isooctane microemulsion of a fixed molar ratio (10:1) of water to AOT was prepared by mixing the aqueous iron solution prepared previously with 9 wt.% AOT isooctane solution. The microemulsion was heated to 78 ◦ C under N2 atmosphere. Then an aqueous NaOH solution was added dropwisely and the mixture was continuously stirred for 1 h. When the reaction was completed, the solvent of the mixture was removed by rotary evaporation, and
2.5. Atomic force microscope observation The lipid bilayer for AFM observation was prepared with the vesicle fusion method (Egawa and Furusawa, 1999; Kaasgaard et al., 2001). Liposomes prepared previously were sonicated to obtain the unilamellar liposomes. The freshly cleaved mica was buried in liposomes suspension for 5 h at 25 ◦ C. Then, the mica was washed ten times with double distilled water at the same temperature. All of the as-prepared films for AFM observation were dried in vacuum at room temperature to eliminate the last trace of solvent. The AFM topography images were obtained in the tapping mode by an AFM (AJ-III, Aijian Nanotechnology Inc., China) with triangularly shaped silicon cantilever with spring constant 48 N/m (Mikro Masch Co., Russia). A resonance frequency in the range of 240–400 kHz was used. The resonance peaks typically at 330 kHz in the frequency response of the cantilever were chosen for the tapping mode oscillation. The AFM images were obtained with a maximum scan range 20 m × 20 m and scanning frequencies were usually in the range between 0.6 and 2.5 Hz per line. The measurements were carried out in air at room temperature (Han et al., 2006).
2.6. Steady state fluorescence measurements DPH was applied as the membrane fluorescent probe in the steady-state anisotropy fluorescence measurement. Fluorescence anisotropy was measured using a fluorescence spectrometer (FLS920, Edinburgh Instruments, Edinburgh, UK) with polarization
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accessory unit. Steady-state fluorescence anisotropy, r, was calculated as: r=
I|| − gI⊥ I|| + 2gI⊥
Table 1 The anisotropy of DPH loaded in liposome membranes in 298 K and 318 K. T/K
Liposome type
Anisotropy of DPH (r)
298
BLs MLs-W MLs-M
0.0736 ± 0.00169 0.0761 ± 0.00138 0.0566 ± 0.00205
318
BLs MLs-W MLs-M
0.0427 ± 0.00092 0.0432 ± 0.00168 0.0405 ± 0.00175
in which g=
I⊥ I||
I⊥ and I|| are the perpendicular and parallel components of fluorescence observed corresponding to the plane of polarization of the excitation beam, respectively (Tang et al., 1998). DPH tetrahydrofuran solution was added into the lecithin chloroform–methanol solution when forming the phospholipid film. All samples with DPH were excited at 360 nm, and the emission light was observed at 430 nm. During the experiment, the turbidity of all samples was lower than 0.088 (absorbance units) and a 395 nm optical filter was applied to filtrate the stray light. The dwell time was 0.1 s and the scan speed was 1.0 nm/step. All samples were stirred for the whole time during the experiment by a magnetic bar equipped in the spectrometer cuvette at 15 and 45 ◦ C, respectively. Pyrene, another hydrophobic fluorescent probe, was added to the lecithin chloroform–methanol solution when forming the phospholipid film at the concentration 1 mol% of the lipid. The steady state fluorescence measurements were carried out at 15 and 45 ◦ C, respectively. Liposomes with pyrene were excited at 335 nm, the emission light was observed at 373 nm. Moreover, 350 nm optical filter was applied to filtrate the stray light. The dwell time was 0.1 s and the scan speed was 1.0 nm/step. Samples were stirred by a magnetic bar in the spectrometer cuvette through the fluorescence measurement. 2.7. Differential scanning calorimetry The calorimetric experiments were performed using a MicroDSC III (Setaram, Caluire, France). The samples were placed in a 1 mL sample cell in the calorimeter while double distilled water placed in another 1 mL reference cell was used as blank. The cells were scanned at a rate of 1 ◦ C/min from 4 to 60 ◦ C. The DSC thermograms were obtained and the phase transition temperature was characterized by the endothermic peak determined using SETSOFT 2000 software (Setaram Inc, France). 3. Results and discussion 3.1. Location of hydrophobic magnetic nanoparticles in liposomes As MNPs coated with AOT were applied as a component of liposomes, the exact location of the hydrophobic MNPs in liposomes was identified by the combination of TEM, AFM and fluorescence anisotropy spectroscopy. The MNPs shown in Fig. 2(a) are spherical in shape with the diameter of about 6 nm. The MNPs coated with AOT can be dispersed stably in chloroform after centrifugation at 12,000 rpm for 30 min. The stabilization of MNPs coated with AOT dispersed in chloroform was better than that of MNPs without the AOT coating, because of the formation of the surface layer on MNPs. TEM was also applied to characterize the morphology of liposomes with hydrophobic MNPs (Fig. 2(b)). Blank liposomes without heavymetal staining were invisible in TEM image, because lipid cannot deflect an electron beam sufficiently (Hall et al., 2007). For the liposomes with hydrophobic MNPs without staining, some dark dots arranging in circle shape were shown in Fig. 2(b). These dark dots, similar in size and morphology to MNPs in Fig. 2(a), were contributed by the hydrophobic MNPs. The circle arrangement of MNPs is mostly different from the random scatter arrangement in
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Fig. 2(a). The difference could be putatively attributed to the sphere shape of liposomes in which the MNPs were embedded. Hence the TEM results confirmed that the hydrophobic MNPs were successfully attached to liposomes. On the other hand, the morphology and size of hydrophobic MNPs-loaded liposomes can be evaluated indirectly by the arrangement of the embedded MNPs. In the TEM photograph, the morphology of MLs-M is spherical in shape and their average size is about 100 nm. The location of hydrophobic MNPs in liposomes was still ambiguous from the TEM results. Given the hydrophobicity of the MNPs surfaces and their strong preference for binding into a hydrophobic environment, the hydrophobic MNPs may be embedded into the membrane resulting in the potential surface topography changes of the lipid bilayer. AFM is a powerful technique for studying membrane surface features, and was applied to confirm the location of the MNPs. The lipid bilayer on mica plate was obtained by vesicle fusion method (Egawa and Furusawa, 1999; Kaasgaard et al., 2001). For blank lipid bilayer, the AFM topographic image was smooth and featureless as shown in Fig. 3(a). Furthermore, the thickness of the layer on mica plate revealed by the depth of the holes (cross section) was about 5 nm, which is similar to the reported thickness of lipid bilayer (Rosenberg et al., 1997). Nevertheless, in AFM image of the membrane with hydrophobic MNPs (Fig. 3(b)), apparent thicker and thinner membrane domains can be observed. Fig. 3(c) shows the smooth bilayer morphology of MLs-W because the hydrophilic MNPs were encapsulated in inner aqueous core. Considering the surface properties of membranes with hydrophilic and hydrophobic MNPs, the height fluctuation in Fig. 3(b) might be related to lipid bilayer distortion, attributed due to the embedment of hydrophobic MNPs in the lipid bilayer, which resulted in the potential changes of membrane’s surface. According to the TEM results, the diameter of MNPs in this work was about 6 nm which was bigger than the thickness of a single lipid bilayer (∼5 nm). Therefore lipid bilayer with embedded hydrophobic MNPs was about 0.7 nm thicker than that without hydrophobic MNPs. The difference, h, as illustrated in Fig. 3(b) led to the surface fluctuation when the membrane was unsaturated with the MNPs. A similar result concerning the membrane distortion induced by embedded hydrophobic nanoparticles was observed using the small angle neutron scattering method by Reimhult et al. (Amstad et al., 2011). These results confirmed that the hydrophobic MNPs were embedded into the lipid bilayer of liposomes. In addition to the morphology method above, the fluorescence anisotropy was applied to further verify the location of hydrophobic MNPs in liposomes. When a hydrophobic fluorescence probe is introduced into lipid bilayer, its rotation rate is restricted by membrane fluidity. The fluidity, which is sensitive to microstructure changes or additives within the membrane (Lande et al., 1995), is mostly revealed by the steady state fluorescence anisotropy of the hydrophobic probe (Tang et al., 1998; Van der Heide et al., 1996). In this case, diphenylhexatriene (DPH) was applied as the fluorescence probe. Generally, the relatively more flexible structure with better membrane fluidity gives rise to the higher rate of DPH’s rotational motion with lower anisotropy. As shown in Table 1, the DPH
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Fig. 2. TEM images of magnetic nanoparticles (a) and liposomes with hydrophobic MNPs (b). The inset shows enlarged view of the magnetoliposomes. The mass ratio of MNPs to lipid in magnetoliposomes was 0.002.
anisotropy for MLs-W is similar to that for BLs at a certain temperature, because the immediate environment of DPH in MLs-W was not influenced by the hydrophilic MNPs which were located in the inner aqueous core of liposomes. However, the anisotropy value of DPH in MLs-M is lower than that of BLs and MLs-W at same temperatures. Combined with the results above, that was because the hydrophobic MNPs were encapsulated within the lipid bilayer and disrupted the original regular arrangement of lipid molecules. Hence the membrane structure of MLs-M was less compact, and the DPH molecules could rotate easily among its surroundings, which gave rise to much lower anisotropy value. 3.2. Effects of MNPs concentration on the morphology and microstructure of liposomes The morphology of lipid bilayer with various hydrophobic MNPs concentration was investigated by AFM. The membrane morphology changed by varying the mass ratio of MNPs to lipid was shown in Fig. 4. The distortion of MLs-M membrane was observed, which could be due to the difference between the MNPs diameter and the
lipid bilayer thickness. With the increase of hydrophobic MNPs concentration, the thicker (light) domains related to the MNPs-loaded lipid bilayer became larger, whereas the thinner (dark) domains referring to lipid bilayer without MNPs became smaller. When the mass ratio of MNPs to lipid reached 0.002, a uniform and featureless bilayer surface could be detected again (Fig. 4(d)). This was because the lipid bilayer was saturated with hydrophobic MNPs. The calculated number of MNPs was about 4.26 × 1014 g−1 of lecithin at the saturation concentration. Tm describing the transition temperature of membrane from gel phase to liquid-crystalline phase can be characterized by the endothermic peak in DSC thermograms. Changes of Tm are sensitive to the additives in membrane and their subsequent effects on the alkyl chain organization. The Tm change depending on the MNPs concentration was shown in Fig. 5. The increase of MNPs concentration induced a reduction of Tm when the MNPs/lipid mass ratio was below 0.002 (Fig. 6). The decrease of Tm suggested that the inclusion of hydrophobic MNPs within the membrane can lead to bilayer disordering (Babincov et al., 1999), which was also proved by the DPH anisotropy method (Table 1). However, the Tm was almost constant
Fig. 3. AFM amplitude images for lipid bilayer of blank liposomes (a), MLs-M (b) and MLs-W (c). The mass ratios of MNPs to lipid in MLs-M or MLs-W were 0.001. The holes in (a) were about 5 nm depth. The inset in each figure shows the schematic diagram of lipid bilayer.
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Fig. 4. AFM amplitude images of the MLs membrane with different concentrations of MNPs. The mass ratios of MNPs to lipid are: (a) 0.0005; (b) 0.0010; (c) 0.0015; (d) 0.0020. The inset in each image shows the schematic diagram of the hydrophobic MNPs-loaded lipid bilayer.
Fig. 5. The thermograms of MLs-M as a function of hydrophobic MNPs concentration determined by differential scanning calorimetry (DSC).
Fig. 6. The phase transition temperature of lipid bilayer as a function of the MNPs concentration in hydrophobic MNPs-loaded liposomes.
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Fig. 8. Changes in fluorescence intensity ratio of pyrene (I1 /I3 ) in hydrophobic MNPs-embedded MLs as a function of increasing concentration of MNPs in gel phase (䊉) and liquid-crystalline phase ().
Fig. 7. The DPH anisotropy as a function of the MNPs concentration in hydrophobic MNPs-loaded liposomes at 15 ◦ C (䊉) and 45 ◦ C ().
when MNPs concentration was beyond 0.002. The result indicated that the lipid bilayer was saturated with MNPs at the concentration, which was consistent with AFM study in this work. The influence of the hydrophobic MNPs concentration on membrane fluidity was revealed by DPH anisotropy change at 15 and 45 ◦ C, respectively. As seen in Fig. 7, the DPH anisotropy in the membrane at 15 ◦ C was lower than that at 45 ◦ C, because the lipid bilayer was in an ordered gel phase with lower fluidity at lower temperature and in a disordered liquid-crystalline phase with higher fluidity at higher temperature. According to Fig. 7, the membrane fluidity in gel phase was improved by adding hydrophobic MNPs, which indicated that the MNPs inclusion disturbed the original rigid structure of lipid bilayer at temperature of 15 ◦ C. When the MNPs/lipid mass ratio was larger than 0.002, the anisotropy value of DPH was almost constant because of the saturation of hydrophobic MNPs in lipid bilayer. This result confirmed the existence of the saturation concentration of hydrophobic MNPs in membrane mentioned above. For MLs-M in liquid-crystalline phase at 45 ◦ C, the membrane fluidity was affected weakly by increasing MNPs concentration. The results could be attributed to the flexible structure of membrane in liquid-crystalline phase, so the disturbing effect of hydrophobic MNPs on membrane structure is insignificant. It suggest that, to some extent, adding hydrophobic MNPs into the membrane has the same effect as rising temperature, which could result in loosening of the membrane structure. This result provides the probability to control the membrane permeabilization by encapsulating hydrophobic MNPs at lower temperature. To gain further insight into the effect of hydrophobic MNPs concentration on membrane structure, pyrene which is located in the lipid bilayer was applied as fluorescence probe (Kutsenko et al., 2011; Shrivastava and Chattopadhyay, 2007). The intensity ratio (I1 /I3 ) of the first (373 nm) to the third (384 nm) peak in fluorescence emission spectrum of pyrene is an indicator of lipid bilayer polarity. Generally, low intensity ratio is indicative of low polarity in lipid bilayer. Fig. 8 shows that the membrane polarity decreases with the increasing MNPs concentration in both gel (15 ◦ C) and liquid-crystalline (45 ◦ C) phase. However, there was a slight decrease of membrane polarity when the MNPs/lipid mass ratio was beyond 0.002. It can be rationalized in terms of the
microstructure change within the nonpolar domain of lipid bilayer. As described above, the hydrophobic MNPs were coated with a layer of AOT, which is a surfactant having a similar structure as lipid. When embedded with the hydrophobic MNPs, lipid membrane could be divided into two pieces of bilayer (Fig. 9). The new bilayer was composed of a lipid layer and an AOT layer. The nonpolar domain was enlarged by the division, which facilitates the inclusion of more pyrene. It was reported that the intensity ratio (I1 /I3 ) of pyrene in membrane medium also depends on the probe concentration in nonpolar or polar areas (Ioffe and Gorbenko, 2005). More inclusion of pyrene in the expanded nonpolar area induces the decrease of polarity. When the MNPs/lipid mass ratio was beyond 0.002, the slight intensity ratio decrease could also be due to the saturation of MNPs in lipid bilayer. In addition, the polarity change in membrane of gel phase was more pronounced than that of liquid-crystalline phase, because water penetration into non-polar membrane region was increased in the relatively uncondensed structure in liquid-crystalline phase. This influence led to the increase in apparent polarity, which weakened the effect of hydrophobic MNPs inclusion on the nonpolar domain. Although the values of I1 /I3 decreased as the temperature increased from 15 ◦ C to 45 ◦ C, this change did not reflect a decrease of polarity. Because water, ethanol, and cyclohexane show a similar decrease of I1 /I3 when temperature increased (Szczupak et al., 2010; Zana et al., 1997). Pyrene is capable of forming a transient excited state complex, known as “excimer”, composed of an excited molecule and
Fig. 9. Schematic diagram of nonpolar area in the blank liposomes and the MNPsloaded liposomes.
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4. Conclusions
Fig. 10. Pyrene excimer (480 nm)/monomer (389 nm) fluorescence intensity ratio in hydrophobic MNPs-embedded MLs as a function of increasing concentration of MNPs in gel phase (䊉) and liquid-crystalline phase ().
a ground-state one (Ioffe and Gorbenko, 2005). The excimer-tomonomer intensity ratio (Iex /Iem ) is determined by measuring fluorescence intensity at the excimer (480 nm) and the monomer (389 nm) peaks, which is informative when detecting the structure change in membrane. In case of 1 mol% of pyrene incorporated into liposomes, the free volume model was applied to account for the pyrene excimerization in membranes (Ioffe and Gorbenko, 2005; L’Heureux and Fragata, 1989). In this model, the lateral displacement of hydrocarbon chains gives rise to appearance of voids where the pyrene excimer forms. Once the void was formed, it will be closed by the moving adjacent chains. If the movement is hindered, the final void, known as free volume, was enlarged. Hence, the higher Iex /Iem reflects the larger free volume in lipid bilayer. As can be seen from Fig. 10, the MNPs inclusion resulted in enlarged free volume in membrane when the mass ratio of MNPs to lipid was lower than 0.002 in both gel and liquid-crystalline phases. According to the DPH anisotropy results, the hydrophobic MNPs disrupted the rigid structure of the membrane. Therefore, the uncondensed structure gave rise to the appearance of more voids in membrane. Furthermore, as the hydrophobic MNPs are rigid additives, their existence in membrane prevented the adjacent lipid molecules to fill the void because of the thermal motion. The hindrance effect provided larger volume for pyrene excimer formation. However, the sustained presence of MNPs beyond 0.002 had different effect on the free volume. Consistent with the results above, the saturation concentration of MNPs in lipid bilayer was detected again. Beyond the concentration threshold, Iex /Iem of pyrene in lipid bilayer at two different temperatures followed two different trends: a plateau reached in gel phase (15 ◦ C), whereas the Iex /Iem decreased in liquid-crystalline phase (45 ◦ C). The unembedded AOT-coated MNPs were dispersed in aqueous water, when the membrane was saturated with MNPs. The AOT molecular was physically absorbed on the surface of MNPs. Since the solvent was changed from the hydrophobic organic solvents to water during the preparation process of MLs-M, the reversibly adsorbed AOT dissociated from the MNPs surface and diffused into the water (Amstad et al., 2009, 2011). Because of the rigid structure of membrane in gel phase, the influence of extra AOT molecule on membrane’s structure was weak. While in the liquid-crystalline phase, the looser structure enabled the extra AOT’s hydrophobic alkyl chains insert into the membrane. The insertion of alkyl chains occupied the existed voids, which reduced the existent free volume and gave rise to a decreasing Iex /Iem value beyond the concentration threshold.
Hydrophobic MNPs coated with AOT were synthesized and embedded into the bilayer of liposomes. The influence of MNPs on the membrane morphology was revealed by TEM and AFM methods. The results showed that the MLs-M were spherical in shape with the diameter of about 100 nm and the lipid bilayer could be distorted to accommodate hydrophobic MNPs, which led to the fluctuations of membrane surface. The effect of hydrophobic MNPs on the microstructure of liposomes was investigated by the DSC and steady state fluorescence measurements. The hydrophobic MNPs in the lipid bilayer not only improved membrane fluidity but also enlarged the nonpolar domain and the free volume in membrane. The lipid phase transition temperature decreased, when the hydrophobic MNPs were embedded in the lipid membrane. Moreover, the concentration of hydrophobic MNPs played an important role in membrane microstructure changes. Several results showed that the lipid bilayer was saturated with the hydrophobic MNPs when the mass ratio of MNPs to lipid reached 0.002. Based on these results, the encapsulation of hydrophobic nanoparticles in the membrane can be applied to modulate the membrane phase behavior and to control the membrane permeabilization. Further studies are needed to develop the application of the MLs-M as a drug delivery system. Acknowledgments This work is financially supported by the National Nature Science Foundation of China (20673059, 20573056 and 21073063), and Nature Science Keystone Foundation of Shanghai (08jc1408100), the National High-Tech Research and Development Plan of China (“863” plan, No. 2011AA06A107) and the Fundamental Research Funds for the Central Universities, China (No. WK0913002). References Ahmad, I., et al., 1993. Antibody-targeted delivery of doxorubicin entrapped in sterically stabilized liposomes can eradicate lung cancer in mice. Cancer Research 53, 1484–1488. Al-Jamal, W.T., et al., 2008. Lipid-quantum dot bilayer vesicles enhance tumor cell uptake and retention in vitro and in vivo. ACS Nano 2, 408–418. Al-Jamal, W.T., Kostarelos, K., 2007. Liposome–nanoparticle hybrids for multimodal diagnostic and therapeutic applications. Nanomedicine 2, 85–98. Al-Jamal, W.T., Kostarelos, K., 2011. Liposomes: from a clinically established drug delivery system to a nanoparticle platform for theranostic nanomedicine. Accounts of Chemical Research 44, 1094–1104. Allen, T.M., Cullis, P.R., 2004. Drug delivery systems: entering the mainstream. Science 303, 1818–1822. Amstad, E., Gillich, T., Bilecka, I., Textor, M., Reimhult, E., 2009. Ultrastable iron oxide nanoparticle colloidal suspensions using dispersants with catechol-derived anchor groups. Nano Letters 9 (12), 4042–4048. Amstad, E., et al., 2011. Triggered release from liposomes through magnetic actuation of iron oxide nanoparticle containing membranes. Nano Letters 11, 1664–1670. An, X., et al., 2010. Photoinduced drug release from thermosensitive AuNPs–liposome using a AuNPs-switch. Chemical Communications 46, 7202–7204. Babincov, M., et al., 1999. Laser triggered drug release from magnetoliposomes. Journal of Magnetism and Magnetic Materials 194, 163–166. Bangham, A.D., et al., 1958. An apparatus for microelectrophoresis of small particles. Nature 182, 642–644. Bangham, A.D., et al., 1965. Diffusion of univalent ions across the lamellae of swollen phospholipids. Journal of Molecular Biology 13, 238–252. Bothun, G., 2008. Hydrophobic silver nanoparticles trapped in lipid bilayers: size distribution, bilayer phase behavior, and optical properties. Journal of Nanobiotechnology 6, 13–23. Chen, Y., et al., 2010. Controlled release from bilayer-decorated magnetoliposomes via electromagnetic heating. ACS Nano 4, 3215–3221. Egawa, H., Furusawa, K., 1999. Liposome adhesion on mica surface studied by atomic force microscopy. Langmuir 15, 1660–1666. Elersic, K., et al., 2012. Electric-field controlled liposome formation with embedded superparamagnetic iron oxide nanoparticles. Chemistry and Physics of Lipids 165, 120–124.
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