Chemistry and Physics of Lipids 177 (2014) 8–18

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Chemistry and Physics of Lipids journal homepage: www.elsevier.com/locate/chemphyslip

Review

Novel methods for liposome preparation Yogita P. Patil, Sameer Jadhav ∗ Department of Chemical Engineering, Indian Institute of Technology Bombay, Powai, Mumbai 400 076, India

a r t i c l e

i n f o

Article history: Received 15 July 2013 Received in revised form 24 October 2013 Accepted 30 October 2013 Available online 9 November 2013 Keywords: Unilamellar Multilamellar Microfluidic Electroformation Supercritical fluid

a b s t r a c t Liposomes are bilayer vesicles which have found use, among other applications, as drug delivery vehicles. Conventional techniques for liposome preparation and size reduction remain popular as these are simple to implement and do not require sophisticated equipment. However, issues related to scale-up for industrial production and scale-down for point-of-care applications have motivated improvements to conventional processes and have also led to the development of novel routes to liposome formation. In this article, these modified and new methods for liposome preparation have been reviewed and classified with the objective of updating the reader to recent developments in liposome production technology. © 2013 Elsevier Ireland Ltd. All rights reserved.

Contents 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Energetics and kinetics of liposomes formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Liposome characterization methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conventional methods for preparing giant uni-lamellar vesicles (GUVs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conventional methods for preparing multilamellar vesicles (MLVs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conventional methods for preparing small and large unilamellar vesicles (SUVs and LUVs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Microfluidic methods for liposome formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Supercritical fluids for liposome formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Modified electroformation methods for preparation of giant vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Size reduction of MLVs and GUVs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Other novel methods for preparing liposomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions and perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction Liposomes are spherical vesicles having an aqueous core enclosed by one or more phospholipid bilayers or lamellae. Liposomes are most frequently classified on the basis of their size (small, large and giant vesicles), number of bilayers (uni-, oligoand multi-lamellar) (Vemuri and Rhodes, 1995; Vuillemard, 1991) and phospholipid charge (neutral, anionic or cationic) (Storm and Crommelin, 1998). Recently, liposomes have also been categorized with respect to their function such as conventional, stealth,

∗ Corresponding author. Tel.: +91 22 2576 7219; fax: +91 22 2572 6895. E-mail addresses: [email protected], [email protected] (S. Jadhav). 0009-3084/$ – see front matter © 2013 Elsevier Ireland Ltd. All rights reserved. http://dx.doi.org/10.1016/j.chemphyslip.2013.10.011

8 9 9 10 10 10 11 13 13 13 14 15 16 16

ligand-targeted, long-release, and triggered-release (Sharma and Sharma, 1997; Storm and Crommelin, 1998). Multi-functional liposomes possessing a combination of these features have also been reported (Kale and Torchilin, 2010; Perche and Torchilin, 2013; Xiang et al., 2013). Even though liposomes were first reported close to half a century ago (Bangham et al., 1965), it was three decades after their discovery that the first liposomal drug Ambisome® entered the market (Davidson et al., 1994; Hann and Prentice, 2001). Since then, the list of liposomal drugs has continued to increase (Allen and Cullis, 2013). Over the years, different recipes for liposome preparation at the laboratory scale have been developed and optimized. Liposome preparation techniques may be divided into (a) bulk methods, where liposomes are obtained by transfer of phospholipids from

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an organic phase into an aqueous phase, and (b) film methods, in which lipid films are first deposited on a substrate and subsequently hydrated to give liposomes. The preparation methods have also been classified based on mean size, polydispersity and lamellarity of liposomes obtained, because control over these parameters remains a challenge with almost all preparation methods. This problem is exacerbated when moving from the laboratory to industrial scale (Mozafari, 2005; Riaz, 1996; Wagner and Vorauer-Uhl, 2011). For drug delivery applications the desirable size of liposomes ranges between 50 and 200 nm (Harashima et al., 1994; Woodle, 1995). Therefore, reduction of size and lamellarity of liposomes is typically carried out by subjecting them to homogenization, sonication, extrusion or freeze–thaw cycles (Brandl et al., 1990; Furukawa, 2010; Hope et al., 1985, 1986; Johnson et al., 1971). Several exhaustive reviews have been published describing and comparing conventional methods for liposome preparation (Akbarzadeh et al., 2013; Dua et al., 2012; Mansoori, 2012; Mozafari, 2005; Riaz, 1996; Sharma and Sharma, 1997; Shashi et al., 2012; Storm and Crommelin, 1998; Wagner and VorauerUhl, 2011). The present review covers these methods only briefly and further details may be found in cited literature. The review also does not cover the vast literature on synthetic modifications to phospholipids that are aimed at avoiding liposome detection and clearance by the immune system, targeting of liposomes to afflicted sites and controlled drug release. The focus of this review is on novel methods of liposome preparation that have become possible due to recent developments in technologies enabling fabrication and pattern formation on materials at the micro- and nano-scale. The review also discusses innovative strategies that have been used to overcome the limitations faced in conventional methods of liposome preparation. However, one should bear in mind that the new liposome preparation methods discussed in this review have only been demonstrated at the laboratory scale and remain to be developed for industrial use. Prior to describing the liposome preparation techniques we mention some commonly used liposome characterization methods and briefly discuss the forces and mechanisms governing liposome formation.

2. Energetics and kinetics of liposomes formation Phospholipids have a hydrophilic head group and two long hydrophobic tails which make them poorly soluble in water unless they self-assemble into bilayers. A finite patch of the phospholipid bilayer has an energy associated with its edge where the hydrophobic tails are exposed to water and is proportional to the perimeter of the patch (Fig. 1A). This energy may be minimized by eliminating the edge if the bilayer patch closes to form a spherical vesicle. However, there is also an energy penalty involved with bending the bilayer into a sphere which is proportional to the inverse of the square of the sphere radius. As the bilayer rearranges from a flat disc into a sphere, the total energy of the system first increases due to contributions from bending energy of the bilayer. Subsequently, the total energy decreases as the edges meet and disappear. During the process of bending of a bilayer into a spherical vesicle (Fig. 1A), the patch may grow in size due to addition of phospholipid molecules and other bilayer fragments. It is also possible that the hydrodynamic and other destabilizing forces may lead to fragmentation of the bilayer, which may result in the formation of smaller liposomes. Phospholipid molecules self-assemble into a stack of bilayers on a substrate when the organic solvent is removed by evaporation (Lasic, 1993). On hydration, the bilayer stacks separate out very slowly and if the bilayer edges are allowed to merge at a faster rate, multi-lamellar vesicles (MLVs) are formed (Fig. 1B). Increasing the rate of bilayer separation by application of electric fields or decreasing the rate of bilayer merging by suppressing

Fig. 1. Schematic illustrating phospholipid self-assembly involved in liposome formation. The smallest liposome is formed when the edge energy first exceeds the bending energy (A). The relative kinetics of bilayer folding due to hydrodynamic forces and bilayer separation under hydration forces dictates the size and lamellarity of vesicles formed (B). Schematic diagram not to scale.

hydrodynamic flow, one can obtain uni-lameller vesicles (Angelova and Dimitrov, 1986; Reeves and Dowben, 1969). 3. Liposome characterization methods Since liposome function is strongly dependent on properties such as liposome size, shape, lamellarity and surface charge (Hunt et al., 1979; Juliano and Stamp, 1975; Park et al., 1992), accurate estimation of these properties is vital. Electron microscopic techniques not only enable visualization of liposomes to study their morphology and lamellarity but also facilitate accurate estimation of size of individual liposomes (Egerdie and Singer, 1982; Larrabee et al., 1978). However, electron microscopy facilities are not portable and the techniques are time consuming, requiring a high level of skill to execute intricate sample preparation protocols, and therefore not amenable for routine measurements (Almgren et al., 2000; Egelhaaf et al., 1996). In contrast, dynamic light scattering (DLS) (Miyamoto and Stoeckenius, 1971), also known as photon correlation spectroscopy (PCS), allows quick estimation of the size distribution of liposome populations in the sub-micron range. A limitation of DLS is that it does not provide reliable estimates of size distributions of highly polydisperse liposome populations (Filipe

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et al., 2010). Atomic force microscopy (AFM), which relies on fast scanning by a nanometer sized probe over a sample fixed on glass or mica surface (Wagner, 1998), has also been employed to measure liposome size (Jass et al., 2003). However, the shape of the fragile liposomes may get altered upon interaction with the mica surface and or the AFM probe. Size exclusion chromatography (SEC) may also be used to fractionate liposome populations by resolving them on the basis of their hydrodynamic size (Grabielle-Madelmont et al., 2003). However, it has been reported that higher affinity of lipids for the column material may lead to rupture of liposomes, thereby compromising SEC measurements (Lundahl et al., 1999). Another more recently developed technique is the Asymmetric Field Flow Fractionation (AFFF) which has been increasingly used in estimation of liposome size distribution. This technique, which depends on application of an electric/magnetic/gravitation/centrifugal force field in a direction perpendicular to the flow of a colloidal suspension, is able to accurately measure size distribution of highly polydisperse samples (Korgel et al., 1998). However, the high cost of the instrument limits widespread adoption of AFFF technique (Moon et al., 1998). For estimating size distribution of liposomes in the micron range, light microscopy has been extensively used (Akashi et al., 1996; Jiskoot et al., 1986). Another technique for measuring size distribution of micron sized-liposome is the coulter counter (Khalil et al., 1996; Payne et al., 1986a) in which liposome size is estimated from the change in electrical impedance (found to be proportional to volume of vesicle) as each liposome passes through an orifice. Estimation of liposome lamellarity is often accomplished by 31 P NMR (Frohlich et al., 2001) in which the degree of lamellarity is determined from the signal ratio before and after Mn2+ addition (Mayer et al., 1986). Though frequently used, this technique has been found to be quite sensitive to the Mn2+ concentration, type of liposome and buffer concentration (Frohlich et al., 2001). Other techniques for lamellarity determination include small angle Xray scattering (SAXS) (Jousma et al., 1987; Skalko et al., 1998) and electron microscopy (Hauser, 1984; Hope et al., 1986). Stability of liposome suspensions is highly dependent on surface charge or zeta potential (du Plessis et al., 1996; Woodle, 1993) which may be estimated from their electrophoretic mobility measured using laser Doppler velocimetry (Cevc, 1993). Before discussing novel techniques of liposome preparation that have been recently reported, we briefly describe the conventional methods that have been extensively used.

4. Conventional methods for preparing giant uni-lamellar vesicles (GUVs) A. Gentle hydration of a phospholipid film. The method involves deposition of phospholipids, from a solution in an organic solvent such as chloroform or ethanol, onto a substrate. The film consisting of stacked phospholipid bilayers is subsequently hydrated over a couple of days in the absence of hydrodynamic flow to obtain an aqueous suspension of GUVs (Reeves and Dowben, 1969). Though a significant fraction of the vesicle population also comprises of MLVs (Kuroiwa et al., 2009; Reeves and Dowben, 1969), the gentle hydration method (also known as the “natural swelling” method) is preferred for the obtaining GUVs of charged phospholipids (Akashi et al., 1996). B. Electro-formation. In this method, the phospholipid film is deposited on electrodes and subsequently hydrated for a couple of hours in the presence of an electric field (Angelova and Dimitrov, 1986). Though GUVs are formed by the application of both alternating and direct current, DC fields are not preferred as they lead to bubbling due to electrolysis of water (Angelova and Dimitrov, 1986; Dimitrov and Angelova, 1988). It has been reported that 80%

of the electroformed vesicle population is unilamellar and free from defects (Rodriguez et al., 2005). C. Coalescence of small vesicles. Other routes to obtain GUVs have also been reported but are not in frequent use. For instance, spontaneous coalescence of small vesicles that are stored in suspension for several days, results in the formation of GUVs. Fusion of LUVs may be induced by various means including subjecting them to freeze–thaw cycles in concentrated solutions of electrolytes (Oku and Macdonald, 1983), using oppositely charged phospholipids (Bailey and Cullis, 1997), addition of poly(ethylene glycol) (Massenburg and Lentz, 1993) or fusogenic peptides (Haluska et al., 2006), and incorporating divalent cations into negatively charged phospholipids (Ohki and Arnold, 2000). 5. Conventional methods for preparing multilamellar vesicles (MLVs) A. Hydration of a phospholipid film under hydrodynamic flow. MLVs are formed when a dry phospholipid film of stacked bilayers, deposited on a substrate, is rehydrated under strong hydrodynamic flows for a couple of hours (Bangham et al., 1967). The resulting MLV suspension contains vesicles that are heterogeneous in size and lamellarity. B. Solvent spherule method. Vigorous mixing of an organic phase containing phospholipids and an aqueous phase for ∼1 hr under low vacuum yields an oil-in-water emulsion containing small spherules of lipid-containing solvents. The organic phase is subsequently removed by evaporation which results in the conversion of the spherules into MLVs having a narrow size distribution (Chowhan et al., 1972; Kim et al., 1985). C. Hydration of proliposomes. Proliposomes are stable, dry, freeflowing granular composites containing a phospholipid and a carrier/drug, and form MLVs when dispersed in an aqueous phase. Proliposomes may be formed by drying an organic solution of phospholipids and a carrier/drug resulting in particles comprising of crystalline carrier/drug at the core encapsulated by a phospholipid shell. The removal of the organic phase can be carried out in a rotary vacuum evaporator (Payne et al., 1986b), fluidized bed (Chen and Alli, 1987) or a spray dryer (Alves and Santana, 2004; Chen et al., 1997). 6. Conventional methods for preparing small and large unilamellar vesicles (SUVs and LUVs) A. Reverse phase evaporation. Similar to the solvent spherule method used for preparation of MLVs, reverse phase evaporation also involves hydration of phospholipids dissolved in an organic phase by addition of water with vigorous mixing. In contrast, to the solvent spherule method, a water-in-oil emulsion is formed in this case and evaporation of the organic phase results in an aqueous suspension containing LUVs (Deamer and Bangham, 1976; Szoka Jr and Papahadjopoulos, 1978) as well as MLVs (Gruner et al., 1985). Lower concentrations of phospholipids in the aqueous suspension yield a higher fraction of LUVs compared to MLVs (Pidgeon et al., 1987). B. Injection of organic solvent with dissolved phospholipids into an aqueous phase. When an organic solvent (ethanol or ether), containing dissolved phospholipids, is injected into an aqueous buffer, spontaneous formation of SUVs is observed (Batzri and Korn, 1973). When ethanol is injected, it dissolves in water and dilution of ethanol below a critical concentration forces the dissolved phospholipids to self-assemble in the aqueous phase and form SUVs (Lasic, 1995). In contrast, on injecting an ether solution of phospholipids into water, SUV formation results from the evaporation of ether (Deamer and Bangham, 1976). It has also been shown that

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increasing phospholipid concentration in the organic solvent leads to an increase in the liposome size, polydispersity and lamellarity (Kremer et al., 1977). C. Detergent dialysis. Detergent dialysis, commonly employed to prepare LUVs, was first proposed with the objective of reconstituting integral membrane proteins into phospholipid bilayers (Helenius et al., 1977; Kagawa and Racker, 1971). In this method, phospholipids are solubilized by detergent micelles in an aqueous phase. Subsequent removal of the detergent by dialysis, results in coalescence of the phospholipid rich micelles and formation of LUVs (Alpes et al., 1986; Milsmann et al., 1978). Chromatographic techniques have also been employed for the removal of detergents (Enoch and Strittmatter, 1979; Philippot et al., 1985). D. Reduction of size and lamellarity of MLVs. Since spontaneous formation of MLVs is easily attained, techniques such as the French press, sonication, homogenization and membrane extrusion are often used for conversion of MLVs into SUVs and LUVs. In sonication, ultrasonic waves disrupt MLVs to form vesicles having size ∼30–50 nm (Huang, 1969). However, low captured volumes of resulting polydisperse SUVs as well as degradation of phospholipid and encapsulated drug, limit the use of sonication route for size reduction (Parente and Lentz, 1984). The French press technique involves extrusion of MLV suspensions at high pressures through a small orifice resulting in SUVs (Barenholz and Amselem, 1979; Hamilton Jr et al., 1980). Liposomes produced by French press are larger than those obtained by sonication of MLVs. Homogenization is extensively used for reduction of liposome size and lamellarity for batches prepared at the industrial scale. During homogenization, the liposome suspension is continuously pumped through an orifice and collides with a stainless steel wall in the homogenizer system at very high pressures to downsize liposomes (Brandl et al., 1990). The major drawback of this method is the use of very high operating pressure. The most frequently used method for conversion of MLVs to SUVs and LUVs is membrane extrusion. In this method, a suspension of MLVs is extruded several times through uniform cylindrical pores of a track-etched polycarbonate membrane yielding smaller vesicles (Hope et al., 1985; MacDonald et al., 1991). The mean size of vesicles obtained by extrusion decreases with increase in the trans-membrane pressure as well as the number of extrusion cycles (Frisken et al., 2000; Hunter and Frisken, 1998; Mayer et al., 1986; Patty and Frisken, 2003; Popa et al., 2008). However, the minimum attainable size is governed by the size of membrane pore (Frisken et al., 2000; Hunter and Frisken, 1998; Mayer et al., 1986; Patty and Frisken, 2003; Popa et al., 2008).

7. Microfluidic methods for liposome formation Microfluidics involves fluid flow in channels having crosssectional dimensions, typically in the range of 5–500 ␮m. In the last decade, several novel microfluidics-based techniques have been developed to produce liposomes. The features of microfluidic systems that can be used to advantage in liposome production include ability to accurately dispense nanoliter volumes, precise control over the position of the interface, diffusion-dominated axial mixing and continuous mode of operation at low volumes. A critical review has recently compared the benefits and limitations of various microfluidic techniques for liposome production on the basis of stability, encapsulation efficiency, mean size and polydispersity of liposome populations obtained (Van Swaay and Demello, 2013). A. Micro hydrodynamic focussing (MHF) for generating small vesicles. MHF-mediated liposome formation was first proposed by Jahn and coworkers (Jahn et al., 2004). It is the most extensively studied microfluidic technique that yields monodisperse populations of SUVs and LUVs with exquisite control over liposome size. In this method, an aqueous buffer flows along two opposite walls

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Fig. 2. Liposome formation by micro-hydrodynamic focusing. Diffusive mixing between phospholipid solution in alcohol and an aqueous buffer is facilitated by flow in a microchannel. Dissolution of water and alcohol into each other causes reduction in concentration of the good solvent (alcohol) resulting in self-assembly of phospholipids into bilayers which eventually close into liposomes. Schematic diagram not to scale (Jahn et al., 2004).

of a rectangular channel, while a phospholipid solution in isopropyl alcohol flows between the aqueous layers along the axis of the channel (Fig. 2). Counter diffusion of water and isopropyl alcohol leads to regions with low alcohol fraction where phospholipids are forced to self-assemble into bilayers which eventually close into liposomes. The mean radius of liposomes was found to decrease asymptotically from 140 to 40 nm when the aqueous buffer to phospholipid containing alcohol flow rate ratio (FRR) was increased from 5:1 to 50:1 after which the liposome size remained largely unchanged (Jahn et al., 2010). Similar results were observed when ethanol was used as the solvent for phospholipids and liposome formation was carried out by MHF-mediated core-annular flows in channels with circular cross sections (Phapal and Sunthar, 2013). Though the reasons for liposome size reduction with FRR remain to be completely understood, it seems that diffusive mixing of the alcohol and aqueous phases favours larger liposomes while advective mixing leads to smaller liposomes (Phapal and Sunthar, 2013). Recently, it was demonstrated that the smaller plasmid DNA-cationic liposome (pDNA-CL) complexes were obtained in channels having periodic bends compared to straight channels (Balbino et al., 2013) further implicating advective mixing in the formation of smaller liposomes. In a recent work, Jahn and coworkers observed bilayer disc intermediates inside the micro-channel by cryo-SEM on rapid freezing after stable flows were generated by MHF (Jahn et al., 2013). Increase in FRR was recently reported to result in reduced yield of liposomes (Wi et al., 2012). Independent studies have shown that liposome size increased with an increase in concentration of the phospholipid in alcohol (Mijajlovic et al., 2013; Phapal and Sunthar, 2013). Moreover, another study showed that liposome formation by MHF method in presence of sonication resulted in smaller liposomes (Huang et al., 2010). It was also observed reported that an increase in FRR adversely affected PEG-lipid and folate incorporation into liposomes formed by MHF method (Hood et al., 2013). Furthermore, it was also shown that MHF can be employed for synthesis of liposome encapsulated hydrogels (Hong et al., 2010) with the FRR dependence of size similar to conventional liposomes prepared by MHF (Jahn et al., 2010). Taken together, these studies underscore the fact that yield, size, polydispersity and encapsulation efficiency of liposomes may be modulated by varying FRR, and not by independently varying flow rates of aqueous and alcohol streams. B. Microfluidic droplets for formation of giant vesicles. It has been shown that when two immiscible phases such as oil and water are allowed to flow in a microchannel, small droplets of one phase having uniform size can be generated under certain conditions (Anna et al., 2003; Thorsen et al., 2001). Sugiura and co-workers prepared

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Fig. 3. Microfluidic droplets for preparing giant vesicles. Using microfluidics W/O/W emulsions are prepared by first generating water droplets in oil and then encapsulating these again in water. The phospholipid monolayers are arrayed at the oil water interfaces. When the oil phase evaporates the monolayers merge to form bilayer of a giant vesicle. Schematic diagram not to scale (Shum et al., 2008).

a water-in-hexane emulsion by the microfluidic method using sorbitan monooleate and stearylamine as emulsifiers (Kuroiwa et al., 2009; Sugiura et al., 2008). The water droplets were then frozen to prevent coalescence and removed from the dispersed hexane phase by precipitation (Sugiura et al., 2008). The frozen ice particles were reintroduced in a hexane solution containing a phospholipid, which replaced the sorbitan monooleate at the droplet surface (water–hexane interface). The hexane was evaporated and water added to the system to obtain giant phospholipid vesicles having mean diameters in the range 4–20 ␮m which were similar to that of the water droplets (Sugiura et al., 2008). Entrapment of the fluorophore, calcein by the giant vesicles was poor with the encapsulation efficiency below 20%. Recently, a novel microfluidic device was developed in which the generation water in oil-emulsion, subsequent removal of the oil phase and transfer of giant vesicles into an aqueous phase takes place on the same device (Matosevic and Paegel, 2011). Shum and co-workers prepared water-in-oil-in-water (w/o/w) double emulsions by microfluidic method with the phospholipid dissolved in the middle oil phase which was a volatile mixture of toluene and chloroform (Fig. 3) (Shum et al., 2008). On removal of the organic layer by evaporations, giant vesicles were obtained

(Shum et al., 2008). The w/o/w double emulsion method may be implemented in a relatively straightforward manner compared to the w/o emulsions used by Suguira et al. (Sugiura et al., 2008). Davies and coworkers varied channel wettability and 3D flow focussing to obtain stable double emulsions that do not require amphiphiles to stabilize the droplets (Davies et al., 2012). It was also demonstrated that w/o/w double emulsions enable easy incorporation of both hydrophilic and hydrophobic drugs into the vesicles (Davies et al., 2012; Shum et al., 2008). Droplet emulsion methods have also been used to carry out encapsulation of drugs, proteins, cells and microparticles into giant vesicles (Davies et al., 2012; Shum et al., 2008; Tan et al., 2006). A possible drawback of double emulsion technique is the presence of residual organic solvent in the final liposome product which may be highly toxic for biologically active ingredients and may also adversely influence the stability of liposomes (Dwivedi, 1987; Vemuri and Rhodes, 1995). C. Pulsed jet flow microfluidics for giant vesicles. Another method for generating giant vesicles involves a pulsed jet flow of an aqueous solution from a micro-capillary at a phospholipid bilayer (Funakoshi et al., 2007). The bilayer is generated by bringing two macro-sized aqueous drops together in a phospholipid containing oil phase (Fig. 4). As the oil film between the drops drains, a planar bilayer is formed into which a pulsed jet of aqueous solution is injected using a microdispensor (Funakoshi et al., 2007). Stachiowic and coworkers demonstrated that giant vesicles of uniform size may be prepared in a reproducible manner (Stachowiak et al., 2008). They reported that the size of the vesicle was a function of amplitude and number of pulses during periodic jetting (Stachowiak et al., 2009). The major advantage of the method is the high encapsulation efficiency. However, automation is a challenge as the microcapillary positioning near the bilayer has to be manually controlled. D. Thin film hydration in microtubes. In this method, phospholipids dissolved in chloroform were dried inside microtubes of diameters ranging from 200 to 530 ␮m, and the resulting thin film was hydrated by perfusing an aqueous buffer through the tube (Suzuki et al., 2008). Though the mean liposomes size was much smaller than that of the tubes, smaller LUVs were obtained in tubes having smaller diameters. Moreover, the liposome size was found to be inversely related to the average fluid velocity in the tube (Suzuki et al., 2008). In another study, liposomes prepared by conventional thin film hydration were perfused though a 330 ␮m

Fig. 4. Pulsed-jet flow method for preparing giant vesicles. The phospholipids, dissolved in the oil phase, assemble as monolayers at the oil–water interface of two drops placed in oil (A). Periodic pulses of a fluid jet are directed at the interface of the two water drop brought in close proximity, resulting in giant vesicles in one of the water drops (B). Schematic diagram not to scale (Funakoshi et al., 2007).

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diameter microtube that was placed in a bath sonicator (Yamashita et al., 2010). It was observed that, in contrast to bulk sonication of liposome suspensions, sonication in microtubes resulted in a highly monodisperse population of liposomes (Yamashita et al., 2010). 8. Supercritical fluids for liposome formation Supercritical fluids (SCFs) possess some desirable properties of both liquids and gases. For instance, a small change in pressure or temperature leads to large changes in density of the SCF and solubility of various species in the SCF. SCFs are increasingly replacing organic solvents as they enable more efficient separation and purification. The use of SCFs in liposome preparation has been reviewed by Meur and coworkers (Meure et al., 2008). The SCF used in most studies is carbon dioxide. Castor and Chu proposed two methods where the phospholipid, an organic co-solvent and SCF are brought in contact with an aqueous phase (Castor, 1996; Castor and Chu, 1998). In the first method, the compressed mixture of the phospholipid, SCF and organic cosolvent is injected into the aqueous phase through a nozzle. In the second method, the aqueous phase is first mixed with the phospholipid, SCF and co-solvent mixture and later decompressed by spraying through a nozzle. The size of liposomes obtained was in the range of 0.2–4 ␮m (Castor, 1996; Castor and Chu, 1998). Similar to the decompression method of Castor and Chu (Castor and Chu, 1998), Otake and coworkers suggested the supercritical reverse phase evaporation method (Otake et al., 2001). The two methods differed in the rate at which decompression of the mixture of phospholipids, water, SCF and organic co-solvent occurs. While Castor and Chu proposed rapid decompression of the mixture by spraying through a nozzle (Castor and Chu, 1998), Otake and coworkers used a variable volume chamber to gradually reduce system pressure (Castor and Chu, 1998). The drug encapsulation efficiency of liposomes, prepared using SCFs, was superior to that in conventional methods of liposome preparation. In further works, Otake and coworkers showed that liposome size and trapping efficiency may be modulated by varying the pressure and organic co-solvent concentration of the system (Imura et al., 2003a, 2003b). Karn and co-workers demonstrated that, compared to a modified Bangham film hydration method, SCF reverse phase evaporation provided superior entrapment efficiency and drug loading characteristics for cyclosporin A (Karn et al., 2013). A supercritical anti-solvent (SAS) method has been reported, in which the organic co-solvent containing the phospholipid is sprayed into the supercritical fluid, which acts as the antisolvent, leading to precipitation of micronized particles of the lipids (Magnan et al., 2000). The size of the particles was correlated with the droplet size of the spray as well as the concentration of the phospholipid in the co-solvent (Magnan et al., 2000). Next, the particles are hydrated in an aqueous buffer to obtain liposomes. Lesoin and coworkers studied the size of lipid particles formed by the SAS method and that of liposomes resulting from hydration of the particles (Lesoin et al., 2011). They reported that increasing the pressure of the system or the SCF/co-solvent ratio led to a reduction in the fraction of small liposomes in the system (Lesoin et al., 2011). It has been argued that the SCF methods can be scaled-up without much difficulty. 9. Modified electroformation methods for preparation of giant vesicles The conventional electroformation protocol yields GUVs only under certain constraints. For instance, in the presence of physiological ionic solutions, GUVs are obtained only if charged phospholipids are used (Akashi et al., 1996). It was shown that,

13

by incorporation of 10 mol% of negatively charged phospholipids, GUVs could be obtained in 100 mM potassium chloride solutions (Akashi et al., 1996). It was also shown that 2–5 times more GUVs were electroformed, when uniform thin films of phospholipids obtained by spin coating of lipid solution on ITO slides were used instead of phospholipid films obtained by conventional droplet spreading method (Estes and Mayer, 2005). GUVs of physiological ionic strength were obtained by carrying out electroformation in a flow chamber to allow switching of the initial aqueous solution of glycerol, used to rehydrate the phospholipid film, to an ionic solution while GUVs were still attached to the electrode (Estes and Mayer, 2005). Pott and coworkers were able to electroform GUVs in solution of high ionic strength by applying higher voltages across the electrodes and by using phospholipid films obtained by drying of aqueous phospholipid suspensions in place of organic solutions (Pott et al., 2008). In another study, it was reported that GUV electroformation was superior with damp lipid films compared to that from hydration of dry lipid films deposited by removal of the organic solvent (Baykal-Caglar et al., 2012). It was also shown that damp lipid films formed by the drying of preformed liposomes on ITO slides in a chamber under controlled humid conditions exhibited lower lipid compositional heterogeneity and more uniform cholesterol distribution in the bilayer (Baykal-Caglar et al., 2012). Berre and coworkers demonstrated electroformation of GUVs on functionalized and micropatterned silicon surfaces (Le Berre et al., 2008). They observed larger and more polydisperse giant vesicles by carrying out electroformation on silicon surface compared to ITO surfaces (Le Berre et al., 2008). Moreover, they observed that grafting silicon surface with phenyl trimethoxysilane resulted in larger GUVs compared to native silicon surface, while electroformation carried out on OH- and HF-terminated silicon produced very few GUVs (Le Berre et al., 2008). These data suggest that highly hydrophilic or hydrophobic substrates are not conducive to GUV formation. Furthermore, etching microstructures on silicon dioxide surface prior to phospholipid film deposition and subsequent electroformation significantly influenced GUV formation (Le Berre et al., 2008). The mean size and polydispersity of electroformed GUVs was found to be significantly smaller when phospholipid films were deposited on the silicon dioxide surface having finer microstructures, compared to coarser ones (Le Berre et al., 2008). Herold and coworkers observed that yield of GUVs of cationic phospholipids deteriorated significantly with the age of the ITOcoated glass electrodes (Herold et al., 2012). They implicated a carbon-rich contamination layer on aged ITO electrodes for the reduced GUV yield and demonstrated that GUV electroformation can be restored on annealing of the ITO electrodes (Herold et al., 2012). Okumura and coworkers showed that GUV electroformation does not require deposition of the phospholipid film on the electrode, but presence of electric field is necessary and sufficient for GUV electroformation (Okumura and Sugiyama, 2011; Okumura et al., 2007). They were able to carry out GUV electroformation on a glass surface and a poly(ethylene terephthalate) mesh thereby eliminating the need for an electroconductive substrate (Okumura et al., 2007). Electroformation was also employed for the formation of GUVs from erythrocyte ghosts under physiological ionic conditions (Montes et al., 2007) as well as under low salt conditions (Mikelj et al., 2013). These new studies have removed several of the constraints associated with conventional electroformation methods for preparing GUVs.

10. Size reduction of MLVs and GUVs To obtain highly mono-disperse liposomes having size in the range 40–200 nm for use in drug delivery applications, MLV

14

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suspensions are typically subjected to extrusion or sonication (Hope et al., 1985; Huang, 1969). Richardson and coworkers suggested that the primary mechanism size reduction of liposomes by ultrasound waves was due to a shear-mediated elongation and subsequent rupture of cylindrical liposomes, caused by microstreaming around the bubbles, and not a result of the collapse of cavitated bubbles (Richardson et al., 2007). They also reported that increasing sonication power resulted in smaller liposomes whereas larger liposomes were obtained when the system pressure was increased (Richardson et al., 2007). Yamaguchi and coworkers have reported that the size-reduction of liposomes by sonication is a function of ultrasound frequency (Yamaguchi et al., 2009). They suggested that low frequency waves lead to larger amplitude oscillations and higher microjet streaming which is responsible for formation of smaller liposomes (Yamaguchi et al., 2009). Silva and coworkers quantified the effect of distance of ultrasound source on liposome size (Silva et al., 2010). They reported that liposomes of smaller size and lower polydispersity index were obtained when the distance of probe corresponded to the antinodal point, where the amplitude of the wave is maximal (Silva et al., 2010). Tejera-Garcia and coworkers used adaptive focused ultrasound to maximize the mechanical energy from ultrasound at a focal point and varied the number of pressure wave (cycles) per burst and duty cycle (fraction of time for which the pulse is switched on in the total pulse period) (Tejera-Garcia et al., 2011). The optimal number of cycles per pulse for obtaining the smallest liposomes with lowest value of PDI was in the range of 100–500, depending upon the type of phospholipid and the intensity of the ultrasound wave (Tejera-Garcia et al., 2011). Frisken and coworkers have shown that the liposome size obtained by extrusion of MLVs through track-etched polycarbonate membranes is a function of the membrane pore size, extrusion pressure and number of extrusion cycles (Frisken et al., 2000; Hunter and Frisken, 1998; Patty and Frisken, 2003). Recently, we have demonstrated that, in contrast to the multiple high pressure extrusion cycles required for MLV size reduction, only a single extrusion of electroformed GUVs at moderate extrusion pressures is sufficient to obtain monodisperse liposome populations (Patil et al., 2012, 2013). We also showed that the mean size is a function of the membrane pore size and the suspension velocity inside the pore (Patil et al., 2012, 2013). Furthermore, we showed that the entrapment efficiency of GUV-derived liposomes was significantly higher, compared to that of MLV-derived liposomes (Patil et al., 2012).

11. Other novel methods for preparing liposomes A. Freeze drying of double emulsions. Freeze drying of liposomeforming lipids and water-soluble carrier materials dissolved in tert-butyl alcohol/water co-solvent systems results in cakes of an isotropic monophasic solution. On addition of water, the freezedried product spontaneously forms a homogenous dispersion of MLVs which may then be downsized by extrusion. However, one of the limitations of freeze drying was the relatively low encapsulation efficiency of freeze dried liposomes. Wang and coworkers proposed a novel scheme comprising of freeze drying a double emulsion W1/O/W2, where W1 and W2 are aqueous phases and O is the organic phase with a dissolved phospholipid (Wang et al., 2006). The liposomes obtained were below 200 nm in size while encapsulation efficiency varied with type of species that was encapsulated. For instance, the efficiency of calcein entrapment was 87% while that of 5-fluorouracil was only 19% (Wang et al., 2006). Freeze drying of water-in-oil (W/O) emulsions that contained phospholipids, resulted in lyophilates, which on rehydration yielded liposomes having mean size less than 200 nm with encapsulation efficiency greater than 60% for three different drugs (Wang et al., 2011). Freeze

Fig. 5. Liposome formation in a membrane contactor. An alcohol solution of phospholipids is extruded into an aqueous phase through a membrane. Dispersion of alcohol droplets into the aqueous phase and their dissolution results in selfassembly of phospholipid molecules into liposomes. Schematic diagram not to scale (Laouini et al., 2011).

drying technique is also used to solve the long term liposome stability problems of thermolabile products which are sensitive to heat drying. This method involves the removal of water from liposomal products in the frozen state at extremely low pressures in presence of certain sugars (sucrose, trehalose) to prevent leakage of encapsulated materials and increase in liposome size during rehydration (Crowe et al., 1985, 1987; Stark et al., 2010). B. Membrane contactor for preparation of liposomes. Charcosset and coworkers have developed a modified ethanol injection method where phospholipid solution in ethanol was extruded into an aqueous phase using a membrane contactor (Fig. 5) (JaafarMaalej et al., 2011). In the first of these studies, a tubular Shirazu porous glass (SPG) membrane with mean pore size of 900 nm was used, in which the mean liposome size was observed to decrease from 203 to 61 nm when the aqueous to organic phase flow ratio was increased from 1.6 to 2 (Jaafar-Maalej et al., 2011). Similar trends were observed in another study where polypropylene hollow fibres were preferred due to access to larger membrane areas and uniform flows (Laouini et al., 2011). Drug encapsulation efficiency observed in these studies was over 90% (Jaafar-Maalej et al., 2011; Laouini et al., 2011). The group also used micro-engineered membranes with well-defined array of uniform pores in the range 5–40 ␮m, where the mean liposome size was found to increase with increasing membrane pore size (Laouini et al., 2013). Mean liposome size was also found to decrease with an increase in the distance between pores, which was explained by a reduced probability of drop coalescence (Laouini et al., 2013). Advantages of the membrane contactor, apart from greater control on liposome size and high encapsulation efficiency, include the option for continuous operation and scale-up of the process. C. Hydration of phospholipids deposited on nanostructured materials. Yu and coworkers prepared monodisperse liposomes by hydration of phospholipids deposited on electrospun amphiphilic nanofibres, composed of the hydrophilic polymer polyvinyl pyrrolidone and soybean lecithin (Yu et al., 2011). The templating and confinement properties of the nanofibers enabled spontaneous self-assembly of phosphatidyl choline. Liposomes were spontaneously formed when the fibres were added to water. The hydrophilicity of the nano-fibres, which was modulated by varying the phospholipid to polymer ratio, was found to dictate the size of the liposomes formed (Yu et al., 2011). D. Liposome preparation by curvature-tuning. The contributions of bilayer spontaneous curvature and curvature-stress modulations (brought about by different lipids, sterols, and proteins) to liposome self-assembly, have been recently reviewed (Mouritsen, 2011). Spontaneous vesiculation by rapid pH change was first reported by Hauser and coworkers where MLV suspensions were subjected to pH jump resulting in SUVs (Hauser, 1989; Hauser and Gains, 1982). Genc and coworkers demonstrated a solvent-free method for the preparation of monodisperse SUVs by spontaneous vesiculation, where charged or zwitterionic lipids, mixed with

Y.P. Patil, S. Jadhav / Chemistry and Physics of Lipids 177 (2014) 8–18

15

Table 1 Novel methods of liposome preparation discussed in this review. Preparation methods

Properties Type of liposome

Encapsulation efficiency

Scale up/scale down

Operator scale/ automation

References

Monodisperse SUVs and LUVs

Low

−/+

+/−

High

−/+

+/−

C. Pulsed jet flow microfluidics D. Thin film hydration in micro-tubes

Monodisperse GUVs Monodisperse GUVs Monodisperse MLVs and LUVs

High

−/+

+/−

−

−/+

+/−

Hong et al. (2010), Hood et al. (2013), Huang et al. (2010), Jahn et al. (2004, 2010, 2013), Phapal and Sunthar (2013) and Wi et al. (2012) Davies et al. (2012), Shum et al. (2008) and Sugiura et al. (2008) Funakoshi et al. (2007) and Stachowiak et al. (2008, 2009) Suzuki et al. (2008) and Yamashita et al. (2010)

2. Supercritical fluids (SCFs) method

Monodisperse LUVs

High

+/−

−/+

Castor (1996), Castor and Chu (1998), Imura et al. (2003a, 2003b), Karn et al. (2013) and Otake et al. (2001)

3. Modified electroformation method

Polydisperse GUVs

High

−/+

+/−

Baykal-Caglar et al. (2012), Herold et al. (2012), Le Berre et al. (2008), Mikelj et al. (2013), Okumura and Sugiyama (2011), Okumura et al. (2007) and Pott et al. (2008)

4. Size reduction of MLVs and GUVs

Monodisperse SUVs and LUVs

High

+/−

−/+

Patil et al. (2012, 2013), Richardson et al. (2007), Silva et al. (2010), Tejera-Garcia et al. (2011) and Yamaguchi et al. (2009)

5. Freeze drying of double emulsions

Monodisperse SUVs

High

+/−

+/−

Wang et al. (2006, 2011)

6. Membrane contactor method

Monodisperse LUVs

High

+/−

−/+

Jaafar-Maalej et al. (2011) and Laouini et al. (2013)

7. Hydration of phospholipids deposited on nanostructured material

Monodisperse SUVs and LUVs

−

8. Liposome formation by curvature tuning

Monodisperse SUVs

−

−/+

+/−

Genc et al. (2009) and Mouritsen (2011)

9. Biomimetic reaction for vesicular self-assembly

Polydisperse GUVs

−

−/+

+/−

Budin and Devaraj (2012)

1. Microfluidic methods A. Micro hydrodynamic focusing (MHF) B. Microfluidic droplets

Yu et al. (2011)

lyso-palmitoylphosphatidylcholine were subjected to rapid pH change (Genc et al., 2009). By directly adding the phospholipids to an aqueous buffer, the need to first prepare an MLV suspension was eliminated. The time interval of pH jump, equilibration time, temperature, and lipid type were critical parameters that influenced the size, shape, and polydispersity of the liposomes (Genc et al., 2009). 12. Conclusions and perspectives Several liposomal formulations are already on the market, while quite a few are still in the pipeline. Conventional techniques for liposome preparation and size reduction remain popular as these are simple to implement and do not require sophisticated equipment. However, not all laboratory scale techniques are easy to scale-up for industrial liposome production. Many conventional methods, for preparing small and large unilamellar vesicles, involve use of either water miscible/immiscible organic solvents or detergent molecules. The double emulsion method (Shum et al., 2008) and the reverse-phase evaporation technique (Szoka Jr and Papahadjopoulos, 1978) are based on the replacement of a waterimmiscible solvent by an aqueous phase. Residual organic solvents may become toxic and degrade the active ingredient representing a possible risk for human health (Birnbaum et al., 2000). In the case of protein drugs, detergents or organic solvents can cause denaturation of proteins and affect the membrane properties (Seddon et al., 2004). This influences the stability of vesicles and turns out

to be the limiting factor in scaling up of the liposome preparation process (Wagner and Vorauer-Uhl, 2011; Wagner et al., 2002). Scaling down of conventional protocols for use in point-ofcare devices also remains a challenge. Moreover, fresh obstacles are encountered during incorporation of new bioactive molecules into liposomes. To address the aforementioned limitations, several novel techniques of liposome production as well as modifications to conventional methods have been proposed over the last decade (Table 1). However, steps in this direction remain in a nascent stage and much work needs to be done towards scaling, design, control and optimization of processes for liposome formation. Technologies for point-of-care use of liposomes in diagnostic and therapeutic applications remain to be developed and tested. A major impetus to research in this area is due to rapid advances in micro-scale fabrication and nanoscale self-assembly techniques. In the next decade, one can foresee progress in automation and control of processes for production of conventional liposomes as well as innovations in the development of multifunctional liposomes. The ability to precisely manipulate the position and orientation of liposomes or form nano-arrays on various substrates has also expanded their potential use to biosensor applications (Bally et al., 2010; Prakash and Kaler, 2013). Liposomes have also been used as templates for the production of nano-particles (An et al., 2009) and as femto-litre compartments for proteins synthesis (Sunami et al., 2006). In vitro processes mimicking physiological liposome synthesis have also been developed (Nawroth et al., 2011). The need for improvements in the design and stability of liposomal diagnostic

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