Lipid-Based Drug Delivery Systems in Cancer Therapy: What Is Available and What Is Yet to Come

1. Multilamellar Vesicles.

MLVs usually range in size between 0.05 and 10 μm and consist of multiple phospholipid bilayers (Fig. 2) (Sharma and Sharma, 1997). They can easily be prepared and are widely studied along with ULVs. The simplest and the most popular method for the preparation of MLVs is thin film hydration (Sharma and Sharma, 1997). In this method, MLVs can be formed spontaneously by adding an excess volume of aqueous buffer to a thin film of dry lipids at a temperature above the phase transition temperature (PTT) of lipids (Vemuri and Rhodes, 1995; Sharma and Sharma, 1997). For thin film hydration, the desired drug to be encapsulated within MLVs can either be included in the aqueous hydration buffer for hydrophilic drugs or in the lipid film for lipophilic drugs (Sharma and Sharma, 1997). Although it is easy to prepare MLVs using thin film hydration, such a method provides relatively poor drug encapsulation efficiency (5–15%). Therefore, the preparation procedure must be optimized and vesicles must be characterized carefully, as minor changes in the preparation can result in alternations to the liposomal characteristics and behavior (Vemuri and Rhodes, 1995).

In the preparation of MLVs using thin film hydration, a thin film of lipids is preferred because this enhances the encapsulation efficiency of drug (Bangham, 1982). This thin film of lipid can be prepared by removing an organic solvent from the lipid mixture under vacuum using a rotary evaporator (Bangham, 1982). The time allowed for the hydration of lipid film with aqueous or drug solution also influences the amount of drug trapped within liposomal vesicles (Olson et al., 1979). It was demonstrated by Olson et al. (1979) that lipid films with similar compositions could achieve greater drug entrapment by using a slower rate of hydration and gentle mixing. In addition, a small proportion (10–20 mol%) of negatively charged lipids, such as PS, PI, and PG, or positively charged lipids, such as stearylamine, can be added to a liposome formulation to increase the interlamellar distance between successive bilayers, and thus, increase the encapsulation capacity of MLV liposomes (Vemuri and Rhodes, 1995). These lipids with charged moieties can also reduce vesicular aggregation after MLV synthesis (Vemuri and Rhodes, 1995).

As thin film hydration provides low encapsulation efficiency, other methods of MLVs preparation with higher encapsulation efficiency have been proposed. One method involves the hydration of the lipid in the presence of an organic solvent using sonication or vortex mixing (Papahadjopoulos and Watkins, 1967; Gruner et al., 1985). In this method, MLVs are formed as the organic phase is removed by passing a stream of nitrogen through the system after sonication or vortex mixing (Papahadjopoulos and Watkins, 1967; Gruner et al., 1985). This technique has been reported to allow up to 40% encapsulation efficiency to be achieved (Gruner et al., 1985). However, large amounts of organic solvent are often left within the liposome formulation (Papahadjopoulos and Watkins, 1967; Gruner et al., 1985). Another method of preparation previously described involves the mixing of a preformed SUVs dispersion with an aqueous solution of the drug to be encapsulated, followed by lyophilization and rehydration of the mixture (Kirby and Gregoriadis, 1984; Ohsawa et al., 1984). This technique also allows for the formation of MLVs that achieve an encapsulation efficiency of up to 40% (Kirby and Gregoriadis, 1984; Ohsawa et al., 1984).

2. Large Unilamellar Vesicles.

LUVs are liposomal vesicles consisting of a single phospholipid bilayer and are considered to be of a size greater than 100 nm (Fig. 2) (Sharma and Sharma, 1997). However, the size range of LUVs is debatable, because some investigators previously described unilamellar vesicles in a size range of 50–100 nm as LUVs (Vemuri and Rhodes, 1995). Unlike MLVs, LUVs have the ability to hold a larger volume of solution within their cavity. Therefore, LUVs have higher encapsulation efficiency than MLVs (Hauser, 1982; Vemuri and Rhodes, 1995). As LUVs exhibit a high encapsulation efficiency, they are more economical in terms of manufacturing as liposomal drug formulations, since a larger amount of drug can be encapsulated within a smaller quantity of lipids (mg of drug per mg of lipid) (Tyrrell et al., 1976; Vemuri and Rhodes, 1995). Other than high encapsulation efficiency, LUVs also provide a reproducible drug release rate, which is a crucial factor in a drug delivery system (Tyrrell et al., 1976). LUVs can be prepared by reverse-phase evaporation or detergent removal techniques, which are discussed below.

In the reverse-phase evaporation technique, a water-in-oil (w/o) emulsion is formed between water and phospholipids in an excess of organic solvent by mechanical means or by sonication (Szoka and Papahadjopoulos, 1978). When the organic solvent is removed from the mixture under vacuum, phospholipid droplets containing water are formed (Szoka and Papahadjopoulos, 1978). These droplets come together to form a gel-like matrix, which transforms into a smooth paste of LUV suspension once the organic solvent is completely removed (Szoka and Papahadjopoulos, 1978). The reverse-phase evaporation technique has been reported to allow a drug encapsulation efficiency of up to 60–65% to be achieved (Szoka and Papahadjopoulos, 1978). Although high encapsulation efficiency can be achieved using the reverse-phase evaporation technique, this method exposes drugs and biologically active molecules to organic solvents and mechanical forces (Vemuri and Rhodes, 1995). Thus, these molecules may undergo conformational changes, protein denaturation, or DNA strand breakage due to the harsh conditions used (Vemuri and Rhodes, 1995).

Alternatively, LUVs can also be prepared using a detergent removal technique. In this method, phospholipids and a detergent are allowed to mix together forming micelles. The detergent is then removed from the preparation, which results in micelles progressively becoming richer in phospholipids. Subsequently, these phospholipid-rich micelles come together to form single bilayer vesicles (Kagawa and Racker, 1971; Milsmann et al., 1978). It is important that detergents used for this technique are those with high critical micelle concentration (CMC) in the range of 10–20 mM, such as sodium cholate, sodium deoxycholate, and octylglucoside (Vemuri and Rhodes, 1995).

In the detergent removal technique, the detergent can be removed by different methods, such as dialysis, column chromatography, or adsorption using Bio-Beads (Bio-Rad Laboratories, Hercules, CA). The removal of the detergent by dialysis was first reported by Milsmann et al. (1978). In this technique, the detergent is allowed to flow through a dialysis cell from a phospholipid-detergent mixture, resulting in a homogeneous population of single bilayer liposome vesicles ranging in size between 50 and 100 nm in diameter (Milsmann et al., 1978). The use of column chromatography to remove detergents was reported by Enoch and Strittmatter (1979). Using column chromatography, Enoch and Strittmatter separated deoxycholate from the phospholipid-deoxycholate mixture by passing the mixture through a Sephadex (GE Healthcare, Chicago, IL) G-25 column, which yielded single layer phospholipid vesicles with a diameter of 100 nm. Alternatively, adsorption using Bio-Beads can be implemented to remove detergents (Gerritsen et al., 1978). It was reported that Bio-Beads SM-2 were added directly into the phospholipid-detergent mixture (Gerritsen et al., 1978). This allowed the detergent to bind to Bio-Beads selectively and rapidly, separating phospholipid vesicles from the detergent (Gerritsen et al., 1978). However, a nonionic detergent must be used in this method (Gerritsen et al., 1978).

3. Small Unilamellar Vesicles.

SUVs are usually found with diameters ranging between 25 and 50 nm (Fig. 2) (Huang, 1969). They can be prepared from MLVs or LUVs using sonication (Saunders et al., 1962; Huang, 1969) or extrusion under high pressure (Hamilton et al., 1980). In the preparation of SUVs using sonication, the MLVs or LUVs suspension is sonicated under nitrogen or argon gas to reduce the size of the vesicles to the SUV size range (Saunders et al., 1962; Huang, 1969). Both types of sonication, namely a bath or a probe sonicator, can be used to generate SUVs (Vemuri and Rhodes, 1995). However, bath sonication offers advantages over probe sonication, because the preparation of SUVs can be performed aseptically with sealed containers and the temperature can be controlled throughout the preparation (Vemuri and Rhodes, 1995). The characterization of SUVs generated by sonication has previously demonstrated that this method is able to synthesize liposomes with sizes ranging between 25 and 50 nm (Huang, 1969). Alternatively, MLVs or LUVs can be converted to SUVs by extrusion (Hamilton et al., 1980). In this method, MLVs or LUVs are forced multiple times through a narrow orifice under high pressure (20,000 psi) at 4°C, resulting in SUVs with sizes ranging between 15 and 30 nm (Hamilton et al., 1980). This technique allows for the reproducible production of SUVs (Hamilton et al., 1980). However, the temperature of the preparation cannot be accurately controlled using this technique, resulting in temperature fluxes during preparation that can affect phospholipid packing within the liposome (Hamilton et al., 1980).

In addition, SUVs can also be prepared directly by the solvent injection method using diethyl ether (Deamer and Bangham, 1976) or ethanol (Batzri and Korn, 1973). In general, using this method, lipids dissolved in an organic solvent are injected into an excess amount of aqueous solution or water by a syringe-type infusion pump, forming SUVs spontaneously (Batzri and Korn, 1973; Deamer and Bangham, 1976). The organic solvent is then removed from the preparation completely. This method has been reported to produce SUVs with sizes between 50 and 200 nm (Deamer and Bangham, 1976). Similar to other liposome preparations, it is almost impossible to completely remove organic solvents from the preparations when using this technique (Vemuri and Rhodes, 1995).

1. Cancer Cell Targeting.

The aim of cancer cell targeting is to improve the cellular uptake of nanoparticles into these cells (Fig. 6). In this strategy, it is the enhanced cellular internalization rather than the enhanced tumor accumulation that improves the efficacy of nanoparticles (Kirpotin et al., 2006). This cellular targeting will also cause a direct cell kill from the release of an encapsulated drug rather than starving cancer cells of nutrients and oxygen by blocking tumor vasculature (Pastorino et al., 2006). The following list of receptors and specific cell surface moieties (Table 1) are capable of being internalized and have been studied extensively with nanoparticles for active cancer cell targeting (Danhier et al., 2010), namely, transferrin receptor, folate receptor, cell surface glycoproteins, epidermal growth factor receptor (EGFR), and ssDNA and RNA aptamers.

2. Tumoral Endothelium Targeting.

In 1971, Judah Folkman (1971) suggested that the growth of tumors could be suppressed by preventing tumors from forming new blood vessels. This suggestion forms the basis of the design of nanomedicines that actively target tumor endothelial cells (Lammers et al., 2008). In contrast to directly targeting cancer cells, the targeting of tumoral endothelium aims to kill cancer cells by depriving cancer cells of oxygen and nutrients. In this strategy, ligand-targeted nanoparticles bind to and kill angiogenic blood vessels (Fig. 6), which in turn kills cancer cells that these blood vessels support in terms of providing nutrients. By inhibiting blood supply to tumors, the size and metastatic capabilities of tumors can be reduced (Danhier et al., 2010). The active targeting of the tumoral endothelium is also advantageous in that it can solve the issue of insufficient drug delivery to hypovascular tumors to some extent as: 1) the tumor vascular networks are more accessible to circulating nanoparticles than cancer cells localized in the tumoral interstitial matrix; 2) the extravasation of nanoparticles is not required to reach the targeted site; 3) the binding of ligands to their receptors is possible immediately after intravenous injection; 4) endothelial cells are genetically more stable than cancer cells and the risk of endothelial cells developing drug resistance is lower; 5) it has been estimated that the destruction of one endothelial cell could lead to the death of around 100 neoplastic cells, whereas the targeting of cancer cells requires the direct eradication of both accessible and inaccessible tumor cells to be considered effective; 6) the targeting of tumor vascular endothelium can achieve therapeutic efficacy without having to penetrate deep into the tumor, therefore, it is not heavily affected by tumor interstitial hypertension and does not require highly perfused or vascularized blood vessels; and 7) most endothelial cell markers are expressed across a wide range of tumor-types, and thus, broad application is possible (Denekamp and Hobson, 1982; Denekamp, 1984; Danhier et al., 2010).

The following proteins are the main candidates for tumoral endothelium targeting, namely, vascular endothelial growth factor (VEGF) and its receptors, VEGFR-1 and VEGFR-2; α_vβ₃ integrin; vascular cell adhesion molecule-1 (VCAM-1); and matrix metalloproteinases (MMPs) (Table 1). Alternatively, cationic liposomes can be used to target the negatively charged surface of tumor endothelial cells via an electrostatic interaction (Campbell et al., 2009; Weijer et al., 2015).

1. pH-sensitive Liposomes.

Due to defects in their mitochondrial respiratory chain and hypoxic nature, tumors generally exhibit a lower extracellular pH than normal tissue (Feron, 2009). This is due to their metabolism heavily relying on glycolysis to produce energy and is known as the Warburg effect (Warburg, 1956a,b). In the tumor microenvironment where glycolysis is highly active, lactate is constantly being produced as a byproduct of pyruvate to generate nicotinamide adenine (NAD⁺) required by various glycolytic enzymes. Consequently, lactate is transported out of tumor cells via monocarboxylate transporters to avoid lactate cytotoxicity and maintain a high metabolic activity (Dimmer et al., 2000; Feron, 2009). The process of lactate elimination progressively increases the acidity of the tumor extracellular space as the monocarboxylate transporter also simultaneously exports one proton out of tumor cells with each lactate molecule (Dimmer et al., 2000; Cardone et al., 2005). It has been shown that the extracellular pH of tumor tissues is between 6.0 and 7.0, whereas the extracellular pH of normal tissues and the blood is approximately 7.4 (van Sluis et al., 1999; Cardone et al., 2005).

By using pH-sensitive lipids and polymers, nanoparticles that release encapsulated drugs in the presence of low pH within tumors, as well as in endosomes and lysosomes, can be prepared (Shenoy et al., 2005a,b). As liposomes are versatile drug delivery systems, pH-sensitive liposomes have been investigated.

Originally, PE and its derivatives were often used in the preparation of pH-sensitive liposomes together with amphiphilic compounds containing an acidic group that acts as a stabilizer at neutral pH (Ellens et al., 1985; Liu and Huang, 1989; Duzgunes et al., 2001; Torchilin, 2005; Karanth and Murthy, 2007). Unlike other phospholipids, PE consists of a small head group that is weakly hydrated and which occupies a low molecular volume relative to its acyl chain (Cullis and de Kruijff, 1979; Seddon et al., 1983). This characteristic gives PE a cone shape instead of a cylinder shape observed in bilayer stabilizing phospholipids and impedes the formation of a lamellar phase (Cullis and de Kruijff, 1979; Seddon et al., 1983). In contrast, the cone shape of PE molecules favor the formation of a strong intermolecular interaction between the amine and phosphate groups of the phospholipid polar heads (Karanth and Murthy, 2007). As a result, PE molecules have a strong tendency to possess an inverted hexagonal phase above its PTT (Karanth and Murthy, 2007).

The incorporation of amphiphilic molecules containing an acidic group, which can be protonated, in between PE molecules can promote electrostatic repulsion allowing a bilayer structure to be formed, and ultimately, the generation of liposomes under physiologic pH and temperature (Duzgunes et al., 1985; Lai et al., 1985). Although stable liposomes containing PE can be prepared under physiologic pH, destabilization of liposomes can occur when the negatively charged acidic groups of amphiphilic molecules are protonated at low pH (Duzgunes et al., 1985; Lai et al., 1985; Torchilin et al., 1993; Karanth and Murthy, 2007). This is because under this latter condition, amphiphiles lose their bilayer stabilizing capacity, allowing PE molecules to acquire their inverted hexagonal phase (Duzgunes et al., 1985; Lai et al., 1985; Torchilin et al., 1993; Karanth and Murthy, 2007). It is the tendency of PE to form an inverted hexagonal phase in the absence of sufficient bilayer stabilizing agents at low pH that gives liposomes containing PE their pH-sensitive property (Torchilin et al., 1993).

Various other pH-sensitive polymers and strategies have been used to exploit the acidic microenvironment of tumors. For example, succinylated poly(glycidol), PEG derivatives containing carboxyl groups, were reported previously to promote the fusion of liposomes under acidic conditions (Kono et al., 1994; Kono et al., 1997). The incorporation of these PEG derivatives into liposomes composed of egg yolk-PC demonstrated enhanced calcein leakage from liposomes, liposomal aggregation and fusion as the pH decreased (Kono et al., 1994). Moreover, these liposomes containing succinylated poly(glycidol) were relatively stable under normal physiologic conditions at pH 7.4 (Kono et al., 1994). A lipid mixing assay further revealed that egg yolk-PC liposomes bearing succinylated poly(glycidol) fused with endosomal and/or lysosomal membranes (Kono et al., 1997). These studies suggested that these liposomes delivered their contents into the cytoplasm by fusing with the endosomal membrane once they were internalized by cells via endocytosis (Kono et al., 1997). Additionally, an examination of egg yolk-PC liposomes bearing succinylated poly(glycidol) in normal CV-1 renal cells also demonstrated that once internalized, these liposomes were able to deliver calcein into the cytoplasm more efficiently than liposomes made up of egg yolk-PC only (Kono et al., 1997).

The use of pH-sensitive liposomes has shown some promising results for their future application in cancer therapy. For instance, pH-sensitive octylamine grafted polyaspartic acid was incorporated into cytarabine encapsulated liposomes (Wang et al., 2014). This new liposomal formulation was compared with conventional non-pH-sensitive cytarabine encapsulated liposomes and standard cytarabine (Fig. 1) for its antitumor effect and cytotoxicity in both human HepG2 hepatoma cells and normal human liver L02 cells (Wang et al., 2014). The new formulation was found to be more effective at killing cancer cells than both the non-pH-sensitive liposomal formulation and standard cytarabine at all concentrations tested after 48 hours (Wang et al., 2014). In contrast, the non-pH-sensitive liposomal formulation of cytarabine and standard cytarabine were more cytotoxic to normal liver cells than the pH-sensitive liposomal formulation of cytarabine, with the latter formulation showing 100% cell viability at all concentration tested after 48 hours (Wang et al., 2014). Similar results were observed when PEG-poly(monomethyl itaconate)-CholC6 copolymer was incorporated into rapamycin encapsulated liposomes (Ghanbarzadeh et al., 2014). This formulation was able to deliver rapamycin (Fig. 1) to HT-29 colon cancer cells more efficiently than conventional liposomes, while showing less cytotoxicity to human umbilical vein endothelial cells than the latter formulation (Ghanbarzadeh et al., 2014). In addition, the incorporation of either oleyl alcohol or dimethyldioctadecylammonium bromide into liposomes composed of egg-PC, 3β-hydroxy-5-cholestene 3-hemisuccinate (CHEMS) and Tween-80 was also reported to demonstrate pH sensitivity under an acidic environment (Shi et al., 2002; Sudimack et al., 2002). Both formulations were able to efficiently retain encapsulated calcein within liposomes at pH = 7.4, while undergoing rapid calcein release and liposome destabilization upon acidification (Shi et al., 2002; Sudimack et al., 2002). Furthermore, both types of liposomes (i.e., those containing either oleyl alcohol or dimethyldioctadecylammonium bromide) showed better retention of their pH sensitivity in 10% serum compared with pH-sensitive liposomes composed of 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE) (Shi et al., 2002; Sudimack et al., 2002). The conjugation of folate to these two formulations for the targeting of the folate receptor in human oral cancer-derived KB cells also demonstrated improved cytoplasmic delivery of cytosine-β-d-arabinofuranoside in comparison with folate receptor-targeted non-pH-sensitive liposomes (Shi et al., 2002; Sudimack et al., 2002). This was reflected in a 17-and 11-fold increase in cytotoxicity of cytosine-β-d-arabinofuranoside when encapsulated in folate receptor-targeted liposomes containing either oleyl alcohol or dimethyldioctadecylammonium bromide, respectively, compared with folate receptor-targeted non-pH-sensitive liposomes (Shi et al., 2002; Sudimack et al., 2002).

Other strategies reported previously that can be employed to target the acidic tumor environment include the incorporation of fusogenic peptide together with encapsulated substances and zinc oxide-liposome nanocomplexes. It was reported that the coencapsulation of both pH-dependent fusogenic peptide (diNF-7) and diphtheria toxin A chain into non-pH-sensitive immunoliposomes targeting the EGFR could induce fusion and leakage of encapsulated contents at low pH (Mastrobattista et al., 2002). Consequently, the destabilization of liposomes observed led to an increase in cytosolic delivery of liposomal contents (Mastrobattista et al., 2002). Furthermore, the coencapsulation of diNF-7 and diphtheria toxin A chain resulted in increased cytotoxicity of EGFR-targeted immunoliposomes against ovarian carcinoma OVCAR-3 cells compared with immunoliposomes without the diNF-7 peptide (Mastrobattista et al., 2002). More recently, zinc oxide nanoparticles were incorporated into the liposomal bilayer, resulting in the formation of zinc oxide-liposome nanocomplexes (Tripathy et al., 2015). This complexation gave liposomes a pH-sensitive property, as zinc oxide nanoparticles are dissociated under the acidic condition of lysosomes, and thus, releasing the encapsulated content from the liposomes (Tripathy et al., 2015). These nanocomplexes encapsulating daunorubicin were found to be significantly more cytotoxic to alveolar adenocarcinoma A549 cells than conventional daunorubicin (Fig. 1) encapsulated liposomes and also than the unencapsulated drug, because they rapidly released their contents upon lysosomal localization (Tripathy et al., 2015).

2. Temperature-sensitive Liposomes.

The concept of temperature-sensitive liposomes was first introduced in 1978 by Yatvin et al. (1978) who reported that the release of the encapsulated content from liposomes could be achieved by applying heat to the target site to create a mild local hyperthermia. These liposomes were observed to be stable under normal physiologic temperature, while becoming leaky at higher temperature (Yatvin et al., 1978). In cancer treatment, the use of temperature-sensitive liposomes would involve local heating of the tumor site to trigger liposomal drug release (Deshpande et al., 2013). Furthermore, the hyperthermic condition would also enhance the liposomal drug delivery to tumors by increasing vascular perfusion and extravasation of liposomes across tumoral blood vessels (Ponce et al., 2006). These enhancements were observed in tumor-bearing mice, where the uptake of non-temperature-sensitive liposomes increased by ∼2- to 4-fold in heated tumors in comparison with nonheated tumors (Kong et al., 2000; 2001). A 2- to 16-fold increase in liposome delivery to heated tumors was also observed in cats with soft tissue sarcomas relative to nonheated tumors (Matteucci et al., 2000).

As phospholipid bilayers can undergo phase transition, such as gel-to-liquid crystalline and lamellar-to-hexagonal transition, it is possible for them to become highly permeable to water soluble molecules (Ganta et al., 2008). This allows certain phospholipids to be used in the generation of temperature-sensitive liposomes. Often, 1,2-dipalmitoyl phosphatidylcholine (DPPC) is used as the main component of temperature-sensitive liposomes, as this phospholipid undergoes gel-to-liquid crystalline phase transition at 41°C and causes the leakage of encapsulated substances (Yatvin et al., 1978; Jeong et al., 2009).

Initially, temperature-sensitive liposomes prepared by Yatvin et al. (1978) included DPPC together with 1,2-distearoyl-sn-glycerol-3-phosphocholine (DSPC) at a ratio of 7:1 (Yatvin et al., 1978). Consequently, the inclusion of DSPC increased the PTT of the liposomal membrane from 41°C to 43–45°C, where the maximum content release from liposomes was observed (Yatvin et al., 1978; Yatvin et al., 1981; Mills and Needham, 2005; Marsh, 2013). The content release of this formulation was reported to be slightly improved over non-temperature-sensitive liposomes (Mills and Needham, 2005). However, the release was still too slow to be used for therapeutic purposes (Mills and Needham, 2005). Moreover, the temperature required to trigger release was marginally higher than that achievable clinically (Kong and Dewhirst, 1999). As a result, micelle-forming lysolipids, such as 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphocholine (MPPC) and 1-steroyl-2-hydroxy-sn-glycero-3-phosphocholine (MSPC), have been incorporated into temperature-sensitive liposome formulations to increase the permeability of the liposomal membrane to their encapsulated contents (Needham et al., 2000; Needham et al., 2013).

Liposomal DOX composed of DPPC, MPPC, and DSPE-PEG_2000Da, at a molar ratio of 90:10:4, was found to release ∼45% of the encapsulated drug within 20 seconds of being exposed to a temperature of 42°C (Needham et al., 2000). In contrast, liposomes composed of pure DPPC released 20% of their encapsulated DOX over 1 hour at 42°C, whereas non-temperature-sensitive liposomes demonstrated no release at the same temperature (Needham et al., 2000). Liposomes containing MPPC were also more superior than the traditional temperature-sensitive liposomes composed of DPPC, hydrogenated soy-PC, cholesterol, and DSPE-PEG_2000Da (molar ratio 100:50:30:6), which released 40% of their contents after 30 minutes at 42°C (Needham et al., 2000). Furthermore, the trigger temperature for the release of DOX from temperature-sensitive liposomes containing MPPC was found to be as low as 39–40°C (Needham et al., 2000).

In mice bearing human squamous cell carcinoma (FaDu) xenografts, temperature-sensitive MPPC liposomal DOX demonstrated essentially no antitumor activity at 34°C (Needham et al., 2000). However, at a tumor temperature of 42°C, this liposomal DOX formulation showed dramatic anticancer activity, with all 11 mice in this group achieving a complete response and none showed any cancer regrowth. Animals in this group also remained tumor-free for up to 60 days (Needham et al., 2000). In comparison, non-temperature-sensitive liposomes also showed some antitumor activity at 42°C (Needham et al., 2000). However, the efficacy of this formulation was not as great in extending the tumor growth inhibition time of these animals relative to the temperature-sensitive liposomes (Needham et al., 2000). Similarly, the inclusion of MSPC into the liposomal formulation led to a rapid release of liposomal contents under mild hyperthermic conditions (Needham et al., 2013). In fact, incorporation of MSPC into the liposomal formulation at 5.0, 7.4, 8.5, and 9.3 mol% accelerated the initial DOX release rate, with the 8.5 and 9.3 mol% formulations releasing 80% of their content within 4 and 3 minutes, respectively (Needham et al., 2013).

The rapid release of encapsulated content from temperature-sensitive liposomes containing lysolipids can be explained by the fact that these single chain lipids can increase the fluidity of the liposomal membrane and reduce the PTT slightly by ∼1°C (Needham et al., 2013). More importantly, lysolipids can form porous defects that are stabilized by DSPE-PEG_2000Da within the liposomal bilayer along the grain boundary between solid and liquid lipid at the PTT (Needham et al., 1997; Zhelev, 1998; Mills and Needham, 2005; Needham et al., 2013).

The use of MSPC in the preparation of temperature-sensitive liposomes has demonstrated to be quite effective at inducing rapid liposomal drug release under mild hyperthermic conditions. This led to the development of ThermoDox, a temperature-sensitive liposomal DOX formulation. ThermoDox (Celsion Corporation, Lawrenceville, NJ) is composed of DPPC:MSPC:DSPE-PEG_2000Da at 86.5:9.7:3.8 mol% (Needham et al., 2013). In rabbits bearing the Vx2 tumor, the use of ThermoDox resulted in a 3- to 4-fold increase in DOX levels within heated tumors compared to nonheated tumors and an 8-fold increase in comparison to free DOX (Ranjan et al., 2012). Similarly, mice bearing fibrosarcoma FSA-1 tumor xenografts were found to have a 10-fold higher DOX level when administered as ThermoDox to hyperthermic tumors compared with control tumors at 37°C (Ponce et al., 2007). The higher DOX concentration also resulted in enhanced antitumor activity in mice treated with ThermoDox and hyperthermia (Ponce et al., 2007). Currently, ThermoDox is being investigated in phase II clinical trials for breast cancer and colorectal liver metastases and also in phase III trials for the treatment of hepatocellular carcinoma (May and Li, 2013). For more information on ThermoDox, please refer to Grull and Langereis (2012) and May and Li (2013).

There are many ways in which mild tumor hyperthermia (∼39–42°C) can be generated to trigger drug release from temperature-sensitive liposomes. This includes implementing: 1) radio frequency heating via an array of antennas (van der Zee et al., 2000) or a catheter (Wood et al., 2012); 2) focused microwaves (Hauck et al., 2006); 3) infrared laser (Salomir et al., 2005); and 4) ultrasound (Deckers et al., 2008; Frenkel, 2008). More recently, high-intensity focused ultrasound (HIFU) has emerged as a promising technology for noninvasively applying heat to deep-seated tumors by focusing multiple ultrasound waves at the focal point (Grull and Langereis, 2012). This results in the deposition of high acoustic intensity and subsequent local heating of the tumor (Grull and Langereis, 2012). The induction of hyperthermia by ultrasound is associated with the absorption of acoustic energy by the fluids and tissues and occurs as a result of a rise in power density when HIFU is focused on the target tissue (Ahmed et al., 2015). For more information, please refer to Lu et al. (1996), Huber et al. (2001), and Leighton (2007).

HIFU has been used in combination with magnetic resonance imaging for magnetic resonance guided HIFU (MR-HIFU) treatment (Grull and Langereis, 2012). For temperature-sensitive liposomal drug delivery, this strategy would allow the heating of a predefined volume of tumor to be more controlled and precise (Grull and Langereis, 2012). This is because magnetic resonance imaging can map in detail the treatment field and monitor near-real time temperature changes (Grull and Langereis, 2012). The benefits and problems of this technology were apparent in animal studies using ThermoDox, where DOX levels within the heated tissue were significantly higher compared with nonheated tissue and free DOX (Ponce et al., 2007; Ranjan et al., 2012; Staruch et al., 2012). However, partial thermal ablation was also observed in the heated tissue, which is believed to be due to imperfect temperature control (Ranjan et al., 2012). It is worth noting that MR-HIFU will be used in phase II clinical trials investigating the use of ThermoDox for the treatment of prostate cancer metastases to bone (May and Li, 2013). For more information on HIFU, please refer to Grull and Langereis (2012) and May and Li (2013).

Apart from the incorporation of lysolipids to generate temperature-sensitive liposomes, polymers with a low critical solution temperature (LCST) have also been investigated as an alternative composition for preparing this type of stimuli-sensitive liposomes (Ganta et al., 2008). When polymers with a LCST are incorporated into liposomes, they remain hydrated below the LCST and stabilize the liposomes (Ganta et al., 2008). However, above the LCST, these polymers become dehydrated and destabilize the liposomal membrane, causing the release of their encapsulated contents (Ganta et al., 2008).

By using polymers with a LCST that is slightly above normal physiologic temperature, liposomes with thermosensitive properties can be prepared (Sawant and Torchilin, 2010). Polymers that have been investigated for their use in temperature-sensitive liposome preparations include poly(N-isopropylacrylamide) (Kono et al., 2002), poly(N-iso isopropylacrylamide-co-acrylamide) (Han et al., 2006) and Pluronic F127 (Chandaroy et al., 2001). However, poly(N-isopropylacrylamide) is not biodegradable, and therefore, can result in unwanted side effects associated with polymer accumulation (Sawant and Torchilin, 2010). Please see a review by Kono (2001) for detailed information on temperature-sensitive polymer modified liposomes.

The use of pH-sensitive and temperature-sensitive drug release is considered to be an effective stimuli-responsive strategy, which shows promising results for clinical use in cancer therapy. However, other stimuli-sensitive liposomes also exist and are further discussed in section VIII.

1. Doxorubicin.

DOX (Fig. 1) is an anticancer drug of the anthracycline family (Blum and Carter, 1974). Like other members within this family, it is produced by Streptomyces bacteria (Streptomyces peucetius var. caesius) (Blum and Carter, 1974). DOX interacts with nucleic acids of dividing cells by two mechanisms (Yang et al., 2014a). First, it inhibits the synthesis of DNA and RNA by intercalating between base pairs of the nucleic acid strand with high affinity, and thus, preventing rapidly dividing cancer cells from undergoing replication and transcription (Yang et al., 2014a). Second, it inhibits the topoisomerase IIα enzyme, impeding the ability of supercoiled DNA from unfolding, and thus, further blocking DNA replication and transcription (Gewirtz, 1999). Apart from interacting with nucleic acids, DOX can also form iron complexes to generate redox-active free radicals that are detrimental to nucleic acids, proteins, and membrane lipids (Keizer et al., 1990).

The redox activity of DOX may contribute partially to the drug’s high cumulative dose-dependent cardiotoxicity, because cardiac muscle is enriched with mitochondria, which contain a high level of anionic diphosphatidylglycerol (cardiolipin) that interacts strongly with DOX (Goormaghtigh et al., 1980). This can lead to lipid peroxidation within cardiac tissue (Keizer et al., 1990). Similarly, as hepatocytes also contain high levels of mitochondria, it is possible that the redox activity of DOX may cause lipid peroxidation within the liver, leading to hepatotoxicity (Keizer et al., 1990; Bagchi et al., 1995; Damodar et al., 2014). The redox activity of DOX within the liver was supported by an increase in malondialdehyde, as well as hepatic enzymes involved in free radical metabolism, such as superoxide dismutase, glutathione peroxidase, and catalase (Kalender et al., 2005). However, the severity of DOX-induced hepatotoxicity may not be as severe as the cardiotoxicity. This may be because of the fact that semiquinone radicals of DOX tend to react with oxygen to form less harmful superoxide radicals in the liver, whereas they react preferentially with hydrogen peroxide to form highly reactive hydroxyl radicals in cardiac tissue (Nohl and Jordan, 1983). In addition, the lower level of NADPH cytochrome P-450 reductase in sarcosomes of the cardiac tissue compared to liver microsomes can lead to an inefficient reduction of DOX to its semiquinone form, which may contribute to the preferential reaction of DOX with hydrogen peroxide in the heart (Nohl and Jordan, 1983).

Typically, the standard treatment of cancers using conventional DOX is performed by an intravenous infusion at a dose between 10 and 60 mg/m², with an upper accumulative dose of 550 mg/m² to reduce its side effects and toxicities (Skeel and Khleif, 2011; Easson and Pointon, 1985). Other side effects of DOX, apart from cardiotoxicity, include myelosuppression, nausea and vomiting, stomatitis, alopecia, local tissue damage, and hyperpigmentation of skin (Peng et al., 2005; Takimoto and Calvo, 2008; Skeel and Khleif, 2011).

Although DOX exhibits a high level of toxicity, it was still selected as a candidate for liposomal drug delivery systems because of its effectiveness against cancers (Barenholz, 2012b). DOX is considered to be one of the most effective anticancer drugs ever discovered with activity against a broad range of cancer types, including both hematologic cancers (leukemia and lymphoma) and solid cancers (breast, uterine, ovarian, and lung cancers) (Weiss, 1992). Until today, it remains one of the most effective first line treatments for cancer (Weiss, 1992). Other practical reasons that lead to the selection of DOX for a liposomal drug delivery system are its distinct absorbance and fluorescence properties (Barenholz, 2012b). With a fluorescence excitation wavelength of 480 nm and a fluorescence emission wavelength of 550 nm, the easy and accurate quantification of DOX concentration, its chemical degradation, its state of aggregation, and changes in its local environment due to the difference in its fluorescence excitation and emission spectra, can be examined (Karukstis et al., 1998). The fluorescence properties of DOX also enable the drug’s DNA-binding activity to be detected through the fast quenching of the fluorescent DOX (Barenholz, 2012b). Altogether, the molecular characteristics of DOX allow its pharmacokinetic and biodistribution to be studied over a long period of time (Gabizon et al., 1991; Amselem et al., 1993a).

2. The Early Days of Liposomal Doxorubicin Formulation.

The first generation of liposomal DOX that led to the later development of Doxil was initially prepared in the 1980s by Alberto Gabizon and Yechezkel Barenholz (Barenholz, 2012a). This negatively charged, medium-size multilamellar liposomal DOX formulation prepared from low T_m egg-derived PC, negatively charged egg-derived PG, and cholesterol (OLV-DOX) was able to enter into the “first in man” clinical trial in 1989 (Gabizon et al., 1989). Notably, OLV-DOX reduced DOX toxicity by lowering the peak level of the free drug and changing the drug’s biodistribution, leading to a higher maximum tolerated dose (MTD) compared with conventional DOX (Gabizon et al., 1989). Despite these pharmacological improvements, OLV-DOX was not granted a further clinical trial as a result of its inferior therapeutic efficacy (Gabizon et al., 1989).

Through studies focusing on the pharmacokinetics and biodistribution of OLV-DOX, it was concluded that the ineffectiveness of OLV-DOX was a result of two separate issues with the formulation: 1) the rapid clearance of OLV-DOX by hepatic and splenic RES, while the accumulation of OLV-DOX in the tumor was avoided; and 2) the plasma clearance of fast released free DOX from liposomes (Gabizon et al., 1991; Amselem et al., 1993a). It was believed that the high clearance level of OLV-DOX was contributed to by its inferior physicochemical characteristics. These characteristics included: 1) the drug localization within the liposome membrane bilayer instead of being encapsulated inside the liposome interior, leading to rapid release upon a large dilution, such as during an intravenous infusion to humans (Amselem et al., 1993a,b; Barenholz and Cohen, 1995; Barenholz, 2003); 2) the high level of negative charge on the liposome due to the incorporation of a high concentration of PG into liposome bilayer, which may increase uptake by RES and induce complement activation (Gabizon and Papahadjopoulos, 1988; Szebeni et al., 2007; Szebeni et al., 2011); and 3) the large size of OLV-DOX (200–500 nm) inhibited the extravasation of liposomes into tumoral tissue, and thus, decreased the EPR effect (Hwang, 1987).

Although the development of OLV-DOX for human use was a failure, the inferior characteristics of OLV-DOX did shed some light on the improvements that could be made for the development of a new generation of liposomal DOX, namely Doxil. Doxil was a product developed by Liposome Technology Inc., which changed its name to Sequus and was eventually owned by Johnson & Johnson (Barenholz, 2012b). For a more effective delivery system, Doxil was designed to be in the nano-scale (∼100 nm) range to take advantage of the EPR effect and the permeable tumoral blood vessles. However, this approach posed some challenges, because nanosized liposomes exhibit an extremely small volume, whereas DOX required a relatively high dosage to achieve therapeutic efficacy (10–50 mg/m²). Moreover, the low solubility of DOX also limited the use of conventional drug loading methods. It was rationalized that an appropriate nanosized liposome must be able to extend the plasma circulation time compared with OLV-DOX to improve its efficacy. As a result, a remote loading technique and sterically stabilized liposomes were developed by Liposome Technology Inc. for the manufacture of Doxil (Barenholz, 2012b).

3. The Development of Doxil.

a. Remote loading.

In remote loading (also known as active loading), a transmembrane gradient of ammonium sulfate, [(NH₄)₂SO₄]_liposome>> [(NH₄)₂SO₄]_medium, is used as a driving force for the efficient and stable loading of amphipathic weak base drugs, such as doxorubicin-NH₂ (DOX-NH₂), into preformed nanoliposomes (Fig. 8) (Haran et al., 1993; Bolotin et al., 1994; Lasic et al., 1995). This strategy involves preparing nanosized liposomes that exhibit a transmembrane concentration gradient consisting of a high intraliposomal concentration of (NH₄)₂SO₄ and a low extraliposomal concentration of (NH₄)₂SO₄ (Haran et al., 1993). Depending on the pH, intraliposomal (NH₄)₂SO₄ can exist in the ionized form and the ionized species (NH₄⁺ and SO₄²⁻) usually exhibit very low permeability coefficients. Therefore, these ions either do not, or very slowly, permeate through the liposomal lipid bilayer (Haran et al., 1993). Only upon base exchange between NH₄⁺ and DOX-NH₂ during drug loading will the unionized ammonia molecules (NH₃) be formed, which can then traverse across the liposomal lipid bilayer. The counterion, SO₄^2-, also plays an important role in the drug loading process. This anion regulates the state of aggregation and precipitation/crystallization of DOX inside the liposome by forming a (DOX-NH₃)₂SO₄ salt, which can be observed as long and fiber-like crystals within liposomes (Fig. 9). In doing so, SO₄²⁻ controls the efficiency and stability of both remote loading and drug release (for more information on remote loading see: Barenholz, 2007; Barenholz and Haran, 1994; Bolotin et al., 1994; Haran et al., 1993; Lasic et al., 1995).

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Fig. 8.

Remote loading of DOX into Doxil liposomes using (NH₄)₂SO₄ gradient. The remote loading of DOX into a Doxil liposome uses a transmembrane concentration gradient consisting of a high intraliposomal concentration of (NH₄)₂SO₄ and a low extraliposomal concentration of (NH₄)₂SO₄ as a driving force for DOX loading. In this process, the ammonium salt of DOX (DOX-NH₃⁺) donates H⁺ to form an amphipathic weak base DOX-NH₂, which can diffuse across the phospholipid bilayer of the liposome. Within the liposome, (NH₄)₂SO₄ dissociates to form NH₄⁺ and SO₄^2-. Then NH₄⁺ undergoes a base exchange with DOX-NH₂ to form NH₃ and DOX-NH₃⁺ within the liposome. Two molecules of DOX-NH₃⁺ quickly precipitate or crystallize with SO₄^2- to form the (DOX-NH₃)₂SO₄ salt inside the liposome cavity, whereas NH₃ diffuses across liposome membrane into the external medium.

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Fig. 9.

Cryotransmission electron microscopy (cryo-TEM) of commercial Doxil liposomes. In a commercial Doxil formulation, remote loading produces doxorubicin sulfate [(DOX-NH₃)₂SO₄] crystals within PEGylated nanoliposomes. These crystals are clearly shown by cryo-TEM as long and fiber-like structures within liposomes. In addition, it can also be seen from cryo-TEM that (DOX-NH₃)₂SO₄ rods come into contact with the liposome membrane, slightly changing the shape of spherical liposomes. Reprinted with permission from Elsevier from Barenholz (2012b).

By using remote loading, it is possible to encapsulate more than 90% of DOX within liposomes. However, the efficiency of this loading technique and the stability of liposomal DOX, such as Doxil, depend on the following factors (Barenholz, 2012b):

• The large difference in permeability coefficient of the uncharged NH₃ (10⁻¹ cm/s) and the anionic SO₄^2- (>10⁻¹² cm/s).

• The initial pH gradient having a higher concentration of intraliposomal H⁺ than extraliposomal H⁺.

• The low solubility of the (DOX-NH₃)₂SO₄ salt, which dramatically reduces the intraliposomal osmotic pressure and, therefore, helps prevent liposomes from collapsing (Fig. 9).

• The asymmetry of the DOX partition coefficient (K_p) with higher extraliposomal K_p than intraliposomal K_p.

Not only was Doxil designed as a nanoscale particle, it was also engineered to be sterically stabilized in order to extend its circulation time within human plasma. To improve the short plasma half-life of OLV-DOX, DSPE-PEG_2000Da was incorporated into the lipid bilayer of Doxil. According to Barenholz (2012b), who played a key role in the design and development of Doxil, this idea came from the work of Frank Davis, Abraham Abuchowski, and colleagues (Abuchowski et al., 1977). These investigators conjugated PEG to proteins (Abuchowski et al., 1977) and later demonstrated that PEGylation helped reduce the immunogenicity of therapeutics as well as improve their safety and efficacy (Veronese, 2001; Veronese and Pasut, 2005). Through experimentation with PEG polymers of different sizes (350–15000 Da) and some consideration regarding the metabolism of PEGylated lipids and their kidney excretion rate, a 2000 Da PEG residue was the polymer of choice for the development of Doxil (Barenholz, 2012b). The addition of PEG polymers to Doxil was believed to reduce nonspecific protein-binding and cell interaction, allowing liposomes to stay in the circulation for an extended period of time (Barenholz, 2012b). The current formulation of Doxil is composed of hydrogenated soy-PC:cholesterol:DSPE-PEG_2000Da at a ratio of 56.4: 38.3:5.3 (Mamidi et al., 2010).

4. Preclinical Studies of Doxil.

Through a number of animal studies, it is believed that Doxil liposomes can escape highly permeable tumor blood vessels and subsequently accumulate within the tumor as intact liposomes (Huang et al., 1992; Working et al., 1994; Yuan et al., 1994; Vaage et al., 1997). Within the tumor tissue, Doxil liposomes travel and distribute themselves across the tumor mass by diffusion in a similar manner as macromolecules (Gabizon et al., 2003b). In contrast, free DOX is distributed throughout the body, but its movement toward the tumor is restricted by the tumor interstitial hypertension, which decreases the free drug’s ability to enter tumor tissue (Jain, 1987; Jain et al., 2007). Unfortunately, Doxil’s mechanism of drug release and its internalization are still unclear. However, two mechanisms have been proposed to be responsible for DOX release and the internalization of Doxil (Barenholz, 2012b): 1) intact Doxil liposomes are taken up by cells followed by intracellular drug release; and/or 2) DOX is released extracellularly into the tumor interstitial fluid, which is then taken up by cells as the free drug. Factors that may cause the release of DOX from Doxil include the collapse of the (NH₄)₂SO₄ gradient and the liposomal destabilization of Doxil by phospholipases that hydrolyze phospholipids (Huang et al., 1992; Working et al., 1994; Mouritsen and Jorgensen, 1998). However, the latter factor may only be partially involved, because the presence of cholesterol in the liposome bilayer can dramatically decrease phospholipase activity (Mouritsen and Jorgensen, 1998). Interestingly, it was recently suggested that ammonia produced at the tumor site by glutaminolysis may enhance the release of DOX from Doxil (Silverman and Barenholz, 2015). This may also represent a new strategy for stimuli-responsive drug release, as the level of ammonia observed in the tumor was much higher than in the plasma. For more details, please see Silverman and Barenholz (2015).

Doxil treatment has been shown to be more superior than free DOX in terms of its pharmacokinetic and pharmacodynamic properties in vivo (Lu et al., 2004; Laginha et al., 2005b; Anders et al., 2013). In mice bearing murine hepatocarcinoma (H₂₂) solid tumor xenografts, a single dose of 5 mg/kg Doxil was able to prolong the elimination half-life of DOX, extending the duration from ∼24 hours (observed with a single dose of 5 mg/kg free DOX) to ∼46 hours (Lu et al., 2004). Furthermore, the AUC upon Doxil treatment was shown to be 6.8-fold higher than free DOX after a single 5 mg/kg dose of each treatment (Lu et al., 2004). At either a 5 or 10 mg/kg dose administered intravenously, Doxil was demonstrated to be more effective than free DOX, showing a significantly higher tumor inhibition rate in H₂₂ hepatocarcinoma-bearing mice after 2 and 7 days (Lu et al., 2004). Another pharmacokinetic study comparing Doxil to free DOX has also been reported using murine mammary carcinoma (4T1)-bearing mice (Laginha et al., 2005b). This study measured the amount of DOX released from Doxil (at an equivalent DOX dose of 9 mg/kg) in comparison with free DOX (at a single dose of 9 mg/kg) (Laginha et al., 2005b). It was reported that although ∼49% of DOX was bioavailable from Doxil compared with ∼95% of free DOX, the AUC of DOX in the tumor available from a single dose of Doxil (measured 7 days after administration) was 87-fold higher than that of a single dose of free DOX (measured 24 hours after administration) (Laginha et al., 2005b). Hence, these studies demonstrated the far greater half-life and distribution of Doxil compared to free DOX within the animal. In fact, Doxil was demonstrated to increase the half-life of DOX from 25 hours observed for “free DOX” to 151 hours and reduce the clearance rate from 89 g/h/kg for free DOX to 1.5 g/h/kg for Doxil (Laginha et al., 2005b).

For the purpose of studying breast cancer brain metastases, athymic mice were inoculated intracerebrally with invasive human breast adenocarcinoma MDA-MB-231-BR-luciferase-expressing cells and then treated with either Doxil or free DOX (Anders et al., 2013). It was reported that Doxil improved the serum and intracranial tumor AUC of DOX in these mice in comparison to free DOX by 1500- and 20-fold, respectively (Anders et al., 2013). Doxil also improved the median survival time of these intracranial tumor-bearing mice compared with free DOX, in this case, by 32 versus 23.5 days (Anders et al., 2013). Additionally, the toxicity of Doxil in vivo was shown to be less than free DOX. In fact, intravenous administration of a single dose of Doxil (10 mg/kg) to H₂₂ hepatocarcinoma-bearing mice led to no significant toxicity to the gastric mucosa, intestinal mucosa, and heart muscle (Lu et al., 2004). However, severe damage was observed when free DOX of an equivalent dose was used, most likely because of the higher level of this drug in these tissues (Lu et al., 2004).

5. Clinical Studies of Doxil.

The “first in human” clinical trial for Doxil aimed to study the plasma pharmacokinetics and the accumulation of DOX administered as Doxil compared with free DOX (Gabizon et al., 1994). This study clearly demonstrated that a significantly higher level of DOX was present in both tumor cells and tumor interstitial fluid after administration of Doxil compared with free DOX (Gabizon et al., 1994). Moreover, using cationic ion exchangers, it was shown that more than 98% of the plasma DOX remained associated with liposomes after the intravenous administration of Doxil (Gabizon et al., 1994), indicating the improved stability of Doxil over the former OLV-DOX formulation. It was also demonstrated that the elimination time of Doxil followed a biexponential curve having a half-life of 2 and 45 hours, with most of the drug being eliminated after the second half-life (Gabizon et al., 1994). In addition, the clearance rate of DOX derived from Doxil was shown to be much slower than that of free DOX (0.1 l/h for Doxil compared with 45 l/h for free DOX) as well as its volume of distribution, which was lower for Doxil (4 liters) compared with free DOX (254 liters) (Gabizon et al., 1994). As expected, DOX metabolites derived from the urine of patients treated with Doxil were identical to those from patients treated with free DOX. However, the total amount of metabolites from daily urinary excretion was significantly lower in patients treated with Doxil (Gabizon et al., 1994).

The improvement of Doxil over conventional DOX was also indicated by the levels of the drug in malignant effusions, which was 4–16 times higher than that of free DOX (Gabizon et al., 1994). Furthermore, the tumoral DOX levels peaked between 3 and 7 days after the administration of Doxil, suggesting the exposure of tumor cells to the drug was much higher and for a longer period (Gabizon et al., 1994, 2003b; Solomon and Gabizon, 2008). Overall, Doxil with PEG modification and remote loading exhibited more superior characteristics than free DOX in a number of clinical trials (see section VII.A.7 below). For more information on Doxil’s performance, see the review by Gabizon et al. (2003b).

6. Side Effects and Safety of Doxil.

Although the development of Doxil significantly reduced the cardiotoxicity of DOX, allowing a higher accumulated dose and extended treatment duration to be achieved, two side effects typically not observed with free DOX are apparent after Doxil treatment. The first and more dominant side effect is grade 2 or 3 desquamating dermatitis, also known as palmar plantar erythrodysthesia (PPE) or “foot and hand syndrome,” which appears as redness, tenderness, and peeling of the skin (Solomon and Gabizon, 2008). The severity of PPE increases with dosage and is more pronounced after treatment intervals of 3 weeks rather than 4 weeks. Thus, the latter interval time is used to lessen PPE symptoms, because there is no solution to counteract this side effect (Solomon and Gabizon, 2008). The second side effect is an infusion-related reaction, which is a common side effect of nano-scale drug delivery systems and appears as flushing and shortness of breath. This Doxil-induced infusion reaction is a complement activation-related pseudoallergy, an acute hypersensitivity, which can be reduced by slowing the infusion rate and by appropriate premedication (Solomon and Gabizon, 2008; Barenholz, 2012b). Although Doxil shows minimal hepatotoxicity during clinical trials (Northfelt et al., 1998; Gordon et al., 2001; O'Brien et al., 2004; Orlowski et al., 2007), interestingly, changes in size (from 102 to 196 and 336 nm) and composition (from hydrogenated soy-PC:cholesterol: DSPE-PEG_2000Da 56.4:38.3:5.3 to 52.7:38.3:9) of the Doxil formulation have resulted in significantly elevated liver enzymes in the blood of mice, indicating liver damage (Mamidi et al., 2010).

7. Indications of Doxil.

Due to the improved therapeutic index profiles reported during phase III clinical trials, Doxil was approved by the FDA and/or EMA for the following cancer types:

8. The Shortage of Doxil and the Lack of Generic Doxil.

In 2011, Ben Venue Laboratories, the sole supplier of Doxil, made the decision to shut down its Bedford site in Ohio temporarily because of the lack of Good Manufacturing Practices cited by the FDA (Barenholz, 2012b). However, the shutdown was made permanent in 2013 because of legal and financial reasons. As a result, the supply of Doxil has been scarce since 2011, with thousands of people still waiting for Doxil treatment. Fortunately, in 2013 the FDA took action to temporarily approve a generic version of Doxil called Lipodox, supplied by Sun Pharmaceutical Industries Ltd (Mumbai, India) (Gaspani and Milani, 2013).

Recently, one study demonstrated that Lipodox did not show equivalent efficacy as Doxil in patients for the treatment of ovarian cancer (Smith et al., 2015). In this study, data from ovarian cancer patients receiving Lipodox (from 21st February 2012–1st March 2013) was compared with data from ovarian cancer patients receiving Doxil (from 1st January 1996–30th June 2006). For every three patients receiving Doxil (n = 120), their data were compared with the results from one patient treated with Lipodox (n = 40) by matching their age, stage of cancer, dose of liposomal DOX, platinum sensitivity, and number of prior treatments. It was found that the overall response rate of ovarian cancer patients treated with Lipodox was 4.3% compared with 18% for those treated with Doxil, whereas the mean time to progression was 4.1 ± 2.8 months for the Lipodox-treated group and 6.2 ± 7.2 months for the Doxil-treated group (Smith et al., 2015).

An indirect comparison between two separate clinical trials examining the efficacy of Lipodox and Doxil also demonstrated the lower anticancer activity of Lipodox (Gordon et al., 2001; Chou et al., 2006). In a study by Chou et al. (2006), 29 recurrent platinum-resistant/refractory ovarian cancer patients were treated with Lipodox at a dose of 45 mg/m² for an average of 4.6 cycles. This study reported a median progression free survival of 5.4 months. On the other hand, a study by Gordon et al. (2001) reported a progression free survival of 7.2 months for recurrent epithelial ovarian carcinoma patients treated with Doxil. In this investigation, 239 patients were given Doxil at a dose of 50 mg/m² for six cycles (Gordon et al., 2001).

The inferior efficacy of Lipodox relative to Doxil may be due to the lower uptake of Lipodox in tumor tissue compared with Doxil (Smith et al., 2016). This explanation was suggested by a preclinical study of human ovarian cancer (SKOV₃-GFP-LUC) tumor-bearing mice treated with either Lipodox or Doxil at a dose of 5 and 10 mg/kg for three cycles (Smith et al., 2016). It was found that the intratumoral concentration of Lipodox was lower than that of Doxil at both the 5 mg/kg (1.0–25.5 ng/ml versus 2.7–42.2 ng/ml) and 10 mg/kg (2.9–35.6 ng/ml versus 2.0–76 ng/ml) doses (Smith et al., 2016). Furthermore, compared with Doxil, there was a 15.7% and 21.3% decrease in the efficacy of Lipodox at both 5 and 10 mg/kg, respectively (Smith et al., 2016). This suggested that the lower efficacy of Lipodox observed might have been the result of lower tumor accumulation (Smith et al., 2016). It is worth noting that although both the preclinical and data comparison studies have suggested the lower anticancer activity of Lipodox compared to Doxil, a direct comparative clinical study between Lipodox and Doxil must be performed to ensure a more accurate assessment.

Despite the fact that the patent protection of Doxil in the United States ended in early March 2010 (Barenholz, 2012b), there is still a surprising lack of generic PEGylated liposomal DOX formulations approved by the FDA and EMA. The reason for the lack of generic Doxil may be because of the complexity of the approvals required by the authorities. In fact, generic Doxil does not only need to meet the requirements for the approval of generic low molecular weight drugs and biologicals, but it must also meet the physical and physiochemical requirements for liposomal products (Barenholz, 2012b).

1. The Development of Myocet.

The current formulation of Myocet is composed of egg PC and cholesterol in the ratio of 55:45 mol% (Swenson et al., 2001). Similar to Doxil, the encapsulation of DOX in Myocet is carried out by remote loading. However, the preparation of Myocet uses a pH gradient rather than the (NH₄)₂SO₄ gradient as a driving force for DOX remote loading (Fig. 10) (Swenson et al., 2001). In this approach, sodium carbonate is added to an aqueous suspension of proton-rich liposomes containing citrate buffer, and thus, an acidic environment inside the liposomes (pH = 4) is created with a neutral pH environment outside the liposomes (pH = 7.8) (Swenson et al., 2001). Once added to the suspension, DOX is forced to accumulate within the liposomes by the lower intraliposomal pH. This condition allows more DOX to be loaded into the liposomes that is well beyond its solubility limit (Swenson et al., 2001). The accumulation of DOX within the liposomes is further facilitated by the unique complexation of DOX with citrate anions, which forms bundle-like structures within the liposomes (Fig. 11). This complexation also reduces the rate at which DOX diffuses from the acidic liposomes (Li et al., 2000).

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Fig. 10.

Remote loading of DOX into Myocet liposomes using a pH gradient. In contrast to Doxil, Myocet uses a pH gradient as a driving force to actively load DOX into liposomes. In this approach, sodium carbonate buffer creates a neutral pH environment (pH = 7.8) outside the liposome, whereas citrate buffer creates an acidic environment (pH = 4) inside the liposome. Outside the liposome, DOX-H⁺ is deprotonated to DOX, which can diffuse through liposomal membrane into the liposome cavity. Within the liposome, the acidic pH protonates DOX back to DOX-H⁺, which interacts with citrate anions to form the DOX-citrate salt that precipitates into a crystal.

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Fig. 11.

Cryotransmission electron microscopy (cryo-TEM) of Myocet liposomes. The unique complexation of DOX with citrate anions during the remote loading of DOX into Myocet liposomes results in the formation of bundles of DOX fibers within the liposome. (A) The arrow shows the end of a DOX fibrous bundle with each fiber within the bundle being ∼3–3.5 nm apart. (B) The flexibility of these DOX fibers is demonstrated by their ability to exist as fibrous bundles in different geometries, including U-shape, circle, and straight. (C) The arrow shows striated regions in DOX fibrous bundles that repeat approximately every 50 nm. Reprinted with permission from Elsevier from Li et al. (1998).

The exact interaction between citrate anions and DOX inside Myocet remains unclear. However, it is certain that the multivalency of the citrate anion is crucial for DOX complexation, because similar DOX bundle formations were observed in di-anionic sulfate containing liposomes (Lasic et al., 1992) but not in liposomes containing mono-anionic species (Li et al., 1998). By using pH gradient remote loading, it is possible to encapsulate more than 95% of DOX inside liposomes of size ∼150 nm with a final DOX/lipid molar ratio of 0.27 (Swenson et al., 2001). Upon physical characterization of Myocet, DOX appears as bundles of fibers inside the liposomes, where DOX monomers stack to form each fiber (Fig. 11A). These fibers are believed to be flexible, because cryoelectron microscopy revealed them to be in different geometries, such as straight, curved, and circulars (Fig. 11, B and C) (Li et al., 1998).

2. Preclinical Studies of Myocet.

The higher efficacy of Myocet over conventional DOX is thought to be due to an increase in its circulation time and tumor accumulation, as well as a reduction in DOX toxicity. In fact, an injection of ¹⁴C-labeled DOX into male beagle dogs as either Myocet or the free drug demonstrated that Myocet localized less readily to normal tissue and was eliminated in the bile much slower than free DOX (Potchoiba et al., 1996). The radioactivity of ¹⁴C-labeled DOX also revealed that the AUC of Myocet in myocardial tissues of beagle dogs to be ∼67% of that found for conventional DOX-treated animals, suggesting that Myocet would potentially cause less cardiotoxicity (Potchoiba et al., 1996).

The reduction in cardiotoxicity after Myocet treatment was demonstrated in another study in which beagle dogs were administered intravenously with either Myocet or free DOX at a dose of 1.5 mg/kg every 3 weeks for a total of 8 cycles (Kanter et al., 1993). It was shown that all dogs treated with free DOX exhibited histologic evidence of anthracycline-induced cardiotoxicity by the end of the treatment (157–164 days) (Kanter et al., 1993). In contrast, none of the dogs in the Myocet-treated group showed any signs of microscopic or macroscopic damage to the heart by the end of the treatment (Kanter et al., 1993). Furthermore, dogs treated with free DOX were found to be five times more likely to experience vomiting and diarrhea compared with those treated with Myocet (Kanter et al., 1993). Alopecia and gastrointestinal bleeding were also present in the free drug-treated group, whereas none was observed in the Myocet-treated group (Kanter et al., 1993). This improved safety may be due to the lower accumulation of DOX in normal tissue when administered as Myocet, because normal tissues do not possess permeable blood vessels from which the liposomes can escape (Kanter et al., 1993).

The improved anticancer activity of Myocet in comparison with free DOX was observed in mice bearing the SC115 Shionogi mouse mammary tumor (Mayer et al., 1990a). In this comparative study, tumor-bearing mice were administered with either Myocet or free DOX at 6.5 mg/kg (the MTD for free DOX) weekly for 3 weeks. At this dose, DOX levels within the tumor after Myocet treatment increased from 2.6 µg/g of tissue after 1 hour to 5.5 µg/g of tissue after 24 hours (Mayer et al., 1990a). In contrast, the DOX levels within the tumor for the conventional free drug was 2.0 µg/g of tissue after 1 hour and remained constant over 24 hours (Mayer et al., 1990a). In addition, there was an increase in tumor growth inhibition observed in mice treated with Myocet compared with free DOX (Mayer et al., 1990a).

A further increase in the administration dose of Myocet to 13 mg/kg (the MTD for Myocet) weekly for 3 weeks resulted in greater tumor accumulation of DOX and tumor growth inhibition. It was found that at this dose, the DOX tumor concentration was 5.7 µg/g of tissue after 1 hour and 10.2 µg/g of tissue after 24 hours (Mayer et al., 1990a). Moreover, this treatment led to a significant reduction in tumor weight, from 5 to 0.5 g (Mayer et al., 1990a). Complete tumor regression was also observed in 25% of mice treated with 13 mg/kg of Myocet, which lasted over the 50-day study period (Mayer et al., 1990a).

3. Clinical Studies of Myocet.

The advantage of Myocet over conventional DOX has been demonstrated previously for the treatment of metastatic breast cancer. The plasma level of total DOX was shown to be substantially higher for Myocet than for conventional DOX, whereas peak plasma levels of free DOX were shown to be lower for Myocet than for conventional DOX (Swenson et al., 2001). Three pivotal phase III clinical trials evaluating Myocet for the treatment of metastatic breast cancer reported the improved safety of DOX when administered as Myocet with patients showing a lower incidence of cardiotoxicity, whereas the therapeutic efficacy of Myocet remained comparable to that of conventional DOX (Batist et al., 2001; Harris et al., 2002; Chan et al., 2004).

In a study by Batist et al. (2001), metastatic breast cancer patients were given either a combination of cyclophosphamide (600 mg/m²) and Myocet (60 mg/m²) or conventional DOX (60 mg/m²) for a median of 6 cycles/patient. Both groups showed comparable antitumor activity with a response rate of 43%. Additionally, the combination of cyclophosphamide with Myocet or conventional DOX resulted in a median time to progression of 5.1 versus 5.5 months, a median time to treatment failure of 4.6 versus 4.4 months, and a median survival time of 19 versus 16 months, respectively (Batist et al., 2001). Although the therapeutic efficacy of Myocet and conventional DOX was similar, there were fewer patients in the Myocet-treated group that developed cardiotoxicity compared with the group receiving conventional DOX treatment (6 versus 21%, respectively) (Batist et al., 2001). None of the patients treated with Myocet suffered congestive heart failure, although five cases were found in the conventional DOX-treated patients (Batist et al., 2001). The estimated median cumulative lifetime dose of DOX at which cardiotoxicity first appears was reported to be >2220 mg/m² for Myocet compared with 480 mg/m² for conventional DOX (Batist et al., 2001). Finally, patients receiving Myocet also showed fewer incidence of grade 4 neutropenia compared with conventional DOX-treated patients (61 versus 75%, respectively) (Batist et al., 2001).

A study on Myocet as a monotherapy for the treatment of metastatic breast cancer was also investigated (Harris et al., 2002). In this study, 75 mg/m² of Myocet (n = 108) was compared with a dose of 75 mg/m² of conventional DOX (n = 116) for a median of four cycles. Both treatment groups demonstrated an overall response rate of 26% and a similar time to disease progression of 3.8 months for the Myocet-treated group versus 4.3 months for the conventional DOX-treated group (Harris et al., 2002). The survival time was reported to be 16 months for patients receiving Myocet and 20 months for patients receiving conventional DOX (Harris et al., 2002). Furthermore, it was shown in the study that cardiac events sufficient to remove patients from the trial were more than twice as likely in patients receiving conventional DOX than in those receiving Myocet (29 versus 13%, respectively). Nine cases of congestive heart failure were found in the conventional DOX-treated group, whereas only two were found in the Myocet-treated group (Harris et al., 2002). In addition to lower cardiotoxicity, Myocet-treated patients also showed fewer incidences of grade 3 or 4 infections and grade 3 or 4 nausea/vomiting (Harris et al., 2002). Similar to the previous mentioned clinical trial by Batist et al. (2001), the estimated median cumulative lifetime dose of DOX at which cardiotoxicity first appeared was higher for Myocet than for conventional DOX (785 versus 533 mg/m², respectively) (Harris et al., 2002).

As patients with high risk factors for cardiotoxicity have a greater possibility of suffering cardiac damage when treated with conventional DOX, it is important that they are considered for Myocet treatment. In light of this, a subset analysis of high risk patients (namely those with previous DOX exposure or history of cardiac disease) was performed in individuals treated with Myocet (Batist et al., 2006). This analysis included 68 patients who had been previously treated with adjuvant conventional DOX. Although these patients received a higher cumulative dose of DOX in the Myocet-treated group (308 mg/m²) compared to the conventional DOX-treated group (225 mg/m²), the incidence of cardiac events was still significantly lower in the Myocet-treated group relative to conventional DOX (22 versus 39%, respectively). The median cumulative lifetime dose at which cardiotoxicity first appeared was 780 mg/m² for Myocet versus 580 mg/m² for convention DOX (Batist et al., 2006). Furthermore, antitumor efficacy was shown to be significantly higher in the Myocet-treated group with a response rate of 31 versus 11% in conventional DOX-treated group (Batist et al., 2006). The median time to treatment failure was also higher in Myocet-treated group than the conventional DOX-treated group (4.2 versus 2.1 months, respectively). However, there was no difference in the survival time between both treatment groups (Batist et al., 2006).

In another phase III clinical trial evaluating Myocet for the treatment of metastatic breast cancer, patients were randomized to receive either Myocet (75 mg/m²) or epirubicin (75 mg/m²), a stereoisomer of DOX known to be less cardiotoxic (Chan et al., 2004). Both groups were treated in combination with cyclophosphamide (600 mg/m²) for a maximum of 8 cycles. Myocet was reported to be more effective than epirubicin in this study, showing a significantly longer time to progression (7.7 versus 4.4 months, respectively) and time to treatment failure (5.7 versus 4.4 months, respectively). There were no significant differences observed between the Myocet or epirubicin groups regarding the overall response rate (46 versus 39%, respectively) or overall survival time (18.3 versus 16 months, respectively) (Chan et al., 2004). Both the Myocet and epirubicin treatment groups also showed a lower incidence of cardiotoxicity with only 11.8 versus 10.2%, respectively, showing asymptomatic impairment in the left ventricular ejection fraction (Chan et al., 2004). No clinical congestive heart failure was observed in both groups. However, Myocet in combination with cyclophosphamide did exhibit increased hematologic toxicity compared with epirubicin showing a significantly higher incidence of grade 4 neutropenia (87 versus 67%, respectively), although the frequency of prolonged or febrile grade 4 neutropenia was similar in both groups (Chan et al., 2004). This may be because of the difficulty in finding a comparable equivalent dose between the two drugs.

4. Myocet versus Doxil.

No direct comparative studies between Myocet, Doxil, and conventional DOX have been conducted in humans to date. This may be because each of these formulations exhibits strikingly different DOX pharmacokinetics (Swenson et al., 2001; Leonard et al., 2009), and thus, making it difficult to determine a comparable effective dose for each formulation. However, Myocet (∼46.7 l/h) is believed to have a slower clearance rate than conventional DOX (∼5.1 l/h), but is not as slow as Doxil (Leonard et al., 2009). Due to its long circulation time and low clearance rate, Doxil is able to penetrate into skin tissues readily, making it more effective for the treatment of AIDS-related Kaposi sarcoma (Gordon et al., 1995; Swenson et al., 2001). However, as previously mentioned, Doxil has been observed to result in PPE as a side effect (Gordon et al., 1995). This problem is rarely observed in Myocet-treated metastatic breast cancer patients (<0.5%), which may be due to the fact that non-PEGylated liposomes are phagocytosed by mononuclear phagocytes more frequently than PEGylated liposomes (Sparano and Winer, 2001).

1. The Development of DaunoXome.

a. Identification of lipid compositions for liposomal formulation and the selection of daunorubicin.

DaunoXome is a liposomal formulation of daunorubicin (Fig. 1) (Forssen, 1997), a member of anthracycline family that is similar to DOX (Forssen, 1997). Like DOX, the anticancer activity of daunorubicin is due to: 1) its ability to inhibit DNA synthesis by DNA intercalation and/or inhibition of DNA polymerase activity; 2) interference with topoisomerase to induce DNA damage; and 3) generation of free radicals by redox reactions that can cause molecular damage, such as lipid peroxidation. Interestingly, it was reported that daunorubicin is more effective at inhibiting DNA than RNA synthesis compared with DOX (Bremerskov and Linnemann, 1969). Apart from nucleic acid and oxidative damage that contributes to daunorubicin’s anticancer activity, it was demonstrated that daunorubicin can also induce cancer cell apoptosis by initiating sphingomyelin hydrolysis, which subsequently produces ceramide (Jaffrezou et al., 1996). Currently, conventional daunorubicin is primarily used for the treatment of acute myeloid leukemia in contrast to DOX, which is implemented for the treatment of various solid tumors (Gewirtz, 1999).

The reason that daunorubicin was selected over other anthracyclines for the DaunoXome liposomal formulation was due to its enhanced stability in aqueous solution (Bosanquet, 1986) and its significant activity against various solid tumors (Hortobagyi, 1997) with therapeutic efficacy comparable to that of DOX (Michieli et al., 1993; Iwasaki et al., 1995; Nagasawa et al., 1996). Furthermore, daunorubicin is less cardiotoxic than DOX on a cumulative basis after prolonged administration (Forssen, 1997). DaunoXome was approved by the FDA and EMA in 1996 for the treatment of AIDS-related Kaposi sarcoma (Gill et al., 1996). It was also approved by the FDA for the treatment of acute myeloid leukemia in 2008 (Latagliata et al., 2008).

The development of DaunoXome began with the identification of suitable liposome compositions and physical characteristics that would allow for the maximum uptake of the encapsulated therapeutic by tumors (Forssen, 1997). This identification step was conducted by an in vivo screening approach developed by Mauk and colleagues (Hwang and Mauk, 1977; Mauk and Gamble, 1979b; Mauk et al., 1980). In this approach, a radioactive gamma emitter, indium-111 (¹¹¹In), was actively loaded into liposomes (Fig. 12), enabling the investigators to assess the stability and release of the encapsulated content from liposomes as a function of the formulation (Mauk and Gamble, 1979a). The radioactive indium was actively loaded into a preformed liposome by incorporating ionophores (A23187) into the liposomal membrane bilayer, allowing ¹¹¹In³⁺ ions to pass through the membrane (Fig. 12).

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Fig. 12.

Remote loading of ¹¹¹In into liposomes. The ionophore A23187 is inserted into the lipid bilayer of liposomes. The incorporation of ionophores into liposomal membrane allows ¹¹¹In³⁺ ions to cross the membrane into the liposome cavity. The weak chelator, nitrilotriacetic acid, inside the liposome then traps ¹¹¹In by forming a weak complex with ¹¹¹In³⁺ ions.

Once inside the liposome, ¹¹¹In³⁺ ions were trapped within the liposomal cavity upon complexation with the weak chelating agent, nitrilotriacetic acid (NTA) (Mauk and Gamble, 1979a). Although NTA could trap ¹¹¹In³⁺ within the liposome, only weak binding between ¹¹¹In³⁺ and NTA occurred, and thus, allowing the high affinity association of ¹¹¹In³⁺ with biologic macromolecules to occur (Mauk and Gamble, 1979a). By using active loading, >90% of ¹¹¹In³⁺ could be encapsulated within liposomes. As ¹¹¹In³⁺ binds strongly to tissues and is known to have minimal redistribution (Hwang et al., 1982), the total ¹¹¹In³⁺ released from liposomes into the targeted tissue can be estimated with accuracy.

From extensive in vivo screening of different liposomal formulations, liposomes composed of DSPC and cholesterol in a 2:1 molar ratio and with a diameter between 40 and 80 nm were the most effective formulation for the delivery of encapsulated ¹¹¹In to tumor tissues in vivo (Proffitt et al., 1983b; Turner et al., 1988; Williams et al., 1988). These DSPC:cholesterol liposomes were able to deliver encapsulated ¹¹¹In to tumor cells selectively in a number of murine tumor models, including the mammary adenocarcinomas, EMT-6 and 16C, B16 melanoma, Lewis lung carcinoma, P1798 lymphosarcoma, sarcoma 180, and colon carcinoma 51 (Proffitt et al., 1983a,b; Patel et al., 1985). This liposomal formulation was subsequently developed into an ¹¹¹In-based tumor imaging agent (VesCan), which could image a wide range of solid neoplasms (Presant et al., 1988, 1990). In particular, Kaposi sarcoma demonstrated the highest level of ¹¹¹In uptake of all tumors investigated (Presant et al., 1990). It was suggested that the high uptake of ¹¹¹In in Kaposi sarcoma lesions might be due to their highly vascularized nature, which allowed small particles, such as liposomes, to extravasate readily (Wu et al., 1993; Brown et al., 1996). As a result of the high accumulation of liposomal ¹¹¹In within Kaposi sarcoma lesions, the development of DaunoXome using DSPC:cholesterol (2:1 molar ratio) began.

Apart from delivering liposomal contents efficiently, liposomes prepared from a DSPC:cholesterol (2:1 molar ratio) formulation also posed other advantages (Forssen, 1997). First, liposomal membranes composed of these lipid compositions exhibit a high PTT (Forssen, 1997). This means that, under physiologic or ambient conditions, DSPC:cholesterol liposomes remain stable and are more resistant to the leakage of their entrapped contents. Second, having only two key components in a liposomal formulation simplifies the manufacturing conditions and formulation parameters suitable for both the active ingredients and the carrier (Forssen, 1997). Third, as the in vivo delivery mechanism of these liposomes depends mostly upon the physiologic properties of tumors and their highly vascularized nature, these liposomes can target a wide range of solid tumors and are less affected by targeting problems, such as antigenic drift (Forssen, 1997). Although antigenic drift usually occurs in viruses where the virus undergoes mutations in the antigenic epitopes (antibody binding site) to escape recognition by antibodies and cytotoxic T-lymphocytes, tumors have also been reported to use a similar process as one of the mechanisms to avoid the immune system (Bai et al., 2003). For more detail, please see Bai et al. (2003).

2. Preclinical Studies of DaunoXome.

Once it was established that the formulation in which liposome encapsulated daunorubicin within its interior was the most effective delivery approach, preclinical studies were investigated to evaluate the efficacy, toxicity, and pharmacokinetic profile of this formulation.

The examination of the in vitro cytotoxicity of DaunoXome and conventional daunorubicin against P1798 tumor cells demonstrated that the relative potency of these two forms of daunorubicin varied widely depending on the incubation time (Forssen et al., 1996). For incubation periods under 8 hours, DaunoXome was observed to be less cytotoxic than free daunorubicin. This could be explained by the high membrane permeability of the conventional drug, allowing it to become fully available immediately after it was added to the culture medium. However, after a longer incubation period, DaunoXome was more cytotoxic than conventional daunorubicin (Forssen et al., 1996). A similar result was observed by confocal microscopy imaging of P1798 cells incubated with either free daunorubicin or DaunoXome (Forssen et al., 1996).

It was suggested that DaunoXome is internalized by cells via endocytosis (Forssen, 1997) and subsequently enters the lysosomal compartment, like other nanoparticles (Harding et al., 1991; Turek et al., 1993; Naito et al., 1996; Nam et al., 2009). The study of DaunoXome’s mechanism of uptake was examined by Forssen (1997). In these studies, the indigo dye precursors, 5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside or 5-bromo-4-chloro-3-indolyl-β-d-phosphate, were encapsulated within DSPC:cholesterol liposomes and used to investigate the mechanism of uptake of these liposomes into cells. These precursors, when not encapsulated within liposomes, were poorly taken up by cells both in vitro and in vivo. Furthermore, when activated by the lysosomal enzymes β-galactosidase (for 5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside) or alkaline phosphatase (for 5-bromo-4-chloro-3-indolyl-β-d-phosphate), these agents form an insoluble blue-violet dye (Tsou et al., 1972). When liposomes containing these precursors were examined in tissue culture, a compartmentalized blue-violet dye was observed within the cytoplasm (Forssen, 1997). In contrast, this dye formation was absent when the same compounds in their free forms were used to treat these cells (Forssen, 1997). Thus, these observations were consistent with liposome endocytosis followed by lysosomal fusion.

Two murine models bearing P1798 lymphosarcoma and mammary adenocarcinoma MA16C xenografts were used in the preclinical animal studies of DaunoXome (Forssen et al., 1992). In the fast growing P1798 lymphosarcoma animal model, the single-dose efficacy of DaunoXome and conventional daunorubicin were compared. At equivalent daunorubicin doses, DaunoXome generated a significantly higher median survival time than the conventional drug at all doses investigated (Forssen et al., 1992). The maximum median survival time of tumor-bearing mice achieved for the conventional drug-treated group was 18.5 days at 30 mg/kg, whereas a dose of 40 mg/kg became too toxic, reducing the median survival time to 12.5 days (Forssen et al., 1992). There were no long-term survivors observed in the conventional drug-treated group. In comparison, the maximum median survival time for the DaunoXome-treated mice at 30 mg/kg was 21.5 days. Moreover, there were three long-term survivors observed (>60 days) in this group (Forssen et al., 1992).

In the slow growing mammary adenocarcinoma MA16C model, tumor-free long-term survivors were observed in mice treated with conventional daunorubicin at a dose of 10 and 20 mg/kg, with one and four survivors observed, respectively (Forssen et al., 1992). The remaining mice in this group either developed detectable tumors or died of drug toxicity at higher doses (Forssen et al., 1992). In the DaunoXome-treated group, the occurrence of tumor-free long-term survivors was much more frequent. Forty-nine mice in this group were treated with DaunoXome at doses ranging from 2 to 35 mg/kg, with only three mice having small residual tumors after treatment (two mice at 2 mg/kg and one at 20 mg/kg). In contrast, the other remaining 46 mice were observed to be tumor free (Forssen et al., 1992). Additionally, an apparent complete cure was observed in DaunoXome-treated mice at 25 mg/kg, with all mice becoming tumor free at this dose (Forssen et al., 1992).

The direct cytotoxic effect of DaunoXome against tumors in vivo was also demonstrated by a significant reduction in tumor volume of DaunoXome-treated mice in comparison with conventional drug-treated mice (Forssen et al., 1992). In the conventional drug-treated group, regardless of the dose level, each established tumor eventually grew to a size of >500 mg (Forssen et al., 1992). In contrast, of the 40 mice treated with DaunoXome at doses between 15 and 30 mg/kg, 14 of them were observed to have tumors that regressed and disappeared completely. Based on these data, it was suggested that the tumor growth suppression of DaunoXome at 2 mg/kg was approximately equivalent to the effect observed with conventional daunorubicin at 15 mg/kg (Forssen et al., 1992).

The effectiveness of DaunoXome over conventional daunorubicin can be explained in terms of altered pharmacokinetics. Significant differences in the pharmacokinetics of these two formulations were observed in plasma and also using P1798 solid tumor xenografts (Forssen et al., 1992). The plasma level of daunorubicin-eq (daunorubicin fluorescent equivalent; parent drug plus fluorescent metabolites) produced by DaunoXome after 1 hour was 268 µg/ml, a 185-fold increase over the level produced by the conventional drug (i.e., 1.4 µg/ml). Additionally, the plasma AUC value of DaunoXome was 227-fold higher than the conventional drug (Forssen et al., 1992). The daunorubicin-eq levels in tumors 1 hour postinjection of the conventional drug was 9.6 µg/g. On the other hand, DaunoXome produced an accumulation of daunorubicin-eq within the tumor that continued to increase 8 hour postinjection, peaking at 100 µg/g (Forssen et al., 1992).

The tissue daunorubicin AUC ratios for DaunoXome and the conventional drug were also compared (Forssen et al., 1992). The comparison demonstrated a 10-fold increase in the AUC ratio produced by DaunoXome in tumor tissue relative to conventional daunorubicin (Forssen et al., 1992). In contrast, the AUC ratios of DaunoXome for hepatic, splenic, and brain tissue increased slightly, with no significance difference being found relative to conventional daunorubicin (Forssen et al., 1992). In addition, a slight decrease in the AUC ratio was observed in both the heart and lungs relative to daunorubicin. Hence, as shown by these results, DaunoXome was able to accumulate daunorubicin within tumor tissues at a higher level than other tissue types compared with conventional daunorubicin (Forssen et al., 1992). From preclinical data, it was concluded that DaunoXome operated through targeted delivery, as shown by a 10-fold increase in the tumor AUC value in comparison with a 0.4- to 1.6-fold change observed in normal tissues (Forssen et al., 1992). This finding demonstrated the improved tumor selectivity of DaunoXome over free daunorubicin.

3. Clinical Studies of DaunoXome.

3. Side Effects and Safety of DaunoXome.

Overall, DaunoXome provides an effective and safe alternative to conventional drugs for the treatment of AIDS-related Kaposi sarcoma and acute myeloid leukemia. It has demonstrated a significant rate of tumor regression even in patients who have failed prior chemotherapy (Gill et al., 1996). Side effects of DaunoXome have been shown to be milder than free daunorubicin, with no evidence of anthracycline-related cardiotoxicity. Furthermore, because of the minimal myelosuppression observed with DaunoXome, anti-HIV therapies, such as azidothymidine, can be used together with DaunoXome, further improving patient treatment (Forssen and Ross, 1994; Forssen, 1997).

1. Vincristine.

Vincristine (Fig. 1) is a cell-cycle dependent anticancer drug that has been commercially available on the market since 1963 (Silverman and Deitcher, 2013). It directly binds to tubulin, causing microtubule depolymerization, M-phase arrest, and apoptosis in cells undergoing mitosis (Silverman and Deitcher, 2013). The most effective treatment strategy for vincristine occurs when cancer cells are exposed to a high concentration of vincristine over a long period of time. Unfortunately, this is unachievable because vincristine also induces autonomic and peripheral sensory-motor polyneuropathy, which is due to a blockage in axon transport and axon degradation caused by nonfunctional microtubules (Moore and Pinkerton, 2009). As a result of its dose-limiting toxic side effects, vincristine dosage is usually restricted to 2 mg/m² (Moore and Pinkerton, 2009). Hence, vincristine sulfate liposomal injection (i.e., Marqibo) was developed to improve the pharmacokinetic and pharmacodynamic profiles of vincristine.

2. The Development of Marqibo.

Early studies investigating the liposomal formulation of vincristine examined the encapsulation of the drug within egg PC/cholesterol or DSPC/cholesterol liposomes, both at a lipid composition ratio of 55:45 (Mayer et al., 1990b; Boman et al., 1995; Waterhouse et al., 2005). The loading of vincristine into these liposomes was conducted by remote loading using a pH gradient, where the intra- and extraliposomal pH were 4.0 and 7.5, respectively (Mayer et al., 1990b). Although remote loading was able to encapsulate a high concentration of vincristine with an encapsulation efficiency of 98% in both liposome formulations, the release of vincristine from these liposomes was not as prolonged and sustained as expected (Mayer et al., 1990b). In fact, the egg PC/cholesterol liposomal formulation of vincristine showed a release of 96% of the entrapped content into whole blood after just 24 hours at 37°C (Mayer et al., 1993). Only a small improvement was observed when egg PC was replaced by DSPC, with ∼80% of vincristine being released after 24 hours at 37°C (Mayer et al., 1993). A similar result was observed in vivo, where encapsulated vincristine escaped the liposomes rapidly into the plasma after an intravenous injection into mice inoculated with L1210 leukemia (Mayer et al., 1993).

However, a significant improvement in terms of prolonged vincristine release was observed when DSPC was substituted with sphingomyelin. Such substitution led to a reduction in both the liposomal vincristine leakage rate and hydrolysis rate by 1.42-fold and 100-fold in vitro, respectively (Webb et al., 1995) and a ∼75% retention of encapsulated vincristine within liposomes 24 hours after an intravenous injection into mice (Boman et al., 1994, 1995; Webb et al., 1995; Johnstone et al., 2001; Krishna et al., 2001). Furthermore, the mean particle size of ∼100 nm helped prolong the circulation time of liposomal vincristine (Peixoto Júnior et al., 2009; Thomas et al., 2009; Silverman and Deitcher, 2013). A negligible level of bovine and human plasma protein binding was also observed on the surface of liposomal vincristine during in vitro studies, which was probably due to the neutral charge of the sphingomyelin/cholesterol composition and a tight lipid arrangement (Webb et al., 1995; Oja et al., 1996). This minimal protein binding may have further extended the circulation time of sphingomyelin/cholesterol liposomes (Webb et al., 1995). As such, the sphingomyelin/cholesterol (55:45) liposomal formulation was used to encapsulate vincristine in the production of Marqibo.

3. Preclinical Studies of Marqibo.

Preclinical studies demonstrated Marqibo to be more effective than free vincristine without imposing additional toxicities by means of prolonged plasma circulation time, increased drug penetration, and increased plasma drug concentration (Kanter et al., 1994; Webb et al., 1998). A significantly higher AUC and a lower total vincristine clearance and volume of distribution compared with standard vincristine were observed across mice, rats, and dog models treated with Marqibo, suggesting that the encapsulated drug remained within the body for a longer period of time (Kanter et al., 1994). This was supported by the delay in vincristine excretion in rats injected with Marqibo. In fact, a radiolabeled mass-balance study in rats demonstrated that 90% of Marqibo administered was detected in the urine and feces more than 72 hours after the initial administration (Kanter et al., 1994), a delay of 12–48 hours compared with standard vincristine (Castle et al., 1976).

In addition, Webb et al. (1998) demonstrated that vincristine concentration in the blood and plasma were at least 2-fold higher in mice treated with Marqibo (2 mg/kg of encapsulated vincristine) than in those given unencapsulated vincristine at an identical dose. There was also at least a 10-fold higher drug concentration accumulated within lymph nodes, heart, kidney, liver, skin, small intestine, and spleen in mice treated with liposomal vincristine than in those treated with free drug 1 day after administration (Webb et al., 1998). The study further showed that the increased drug exposure as a result of liposomal encapsulation was not associated with increased drug toxicity. It was reported that mice bearing a P388 ascitic tumor given Marqibo (2, 3, or 4 mg/kg) had a survival rate of at least 50% in all treatment groups, with a 4 mg/kg dose being the most effective (8–9 of 10 mice survived past 60 days depending on drug:lipid ratio). In contrast, mice treated with free vincristine at an identical dose displayed only a 33–38% survival rate (Webb et al., 1998).

By using the liposomal formulation, vincristine was proven to be more efficacious and tolerable than its standard formulation (Shan et al., 2006; Silverman and Deitcher, 2013). In these studies, mice bearing Namwala tumor xenografts were injected intravenously with 1.0, 1.5 and 2.5 mg/kg of Marqibo and compared with those injected with 0.5, 1.0, and 1.5 mg/kg of standard vincristine (Silverman and Deitcher, 2013). The mice injected with Marqibo demonstrated greater tumor suppression than those given standard vincristine, with the Marqibo dose of 2.5 mg/kg showing the highest activity. The MTD for Marqibo was 2.5 mg/kg, a dose unachievable by standard vincristine treatment, where a MTD of 1.5 mg/kg was observed (Silverman and Deitcher, 2013). A similar result was reported using the LX-1 human small-cell lung carcinoma xenograft mouse model, in which Marqibo was shown to have greater antitumor activity than its free drug formulation at the equivalent dose of 1.0 mg/kg (Shan et al., 2006). This study also investigated the extravasation kinetics and accumulation of liposomal encapsulated vincristine using intravital microscopy imaging. Notably, significantly faster extravasation of liposomes was observed in tumor blood vessels than in normal tissues after a single dose of fluorescently labeled liposomal encapsulated vincristine (Shan et al., 2006). Furthermore, the accumulation of liposomal vincristine within the interstitium was ∼70-fold higher in tumor tissue than in normal tissues at 1 hour and remained greater even after 48 hours (Shan et al., 2006).

Silverman and Deitcher (2013) summarized the effect of Marqibo over standard vincristine in 18 animal tumor models representing 11 different cancer types to show the superior anticancer activity of Marqibo. In all cases, Marqibo showed equivalent or higher anticancer activity than standard vincristine for the same dose level. The dose-dependent antitumor activity of Marqibo was observed in 13 of 18 models. Three models were not sensitive to either Marqibo or standard free drug, whereas one was highly sensitive to both treatments (Silverman and Deitcher, 2013).

4. Clinical Studies of Marqibo.

As observed in animal models, Marqibo administered to humans demonstrated a similar improvement over standard vincristine (Thomas et al., 2009; Yan et al., 2012; O'Brien et al., 2013). Indeed, higher doses of Marqibo were able to be administered to patients compared with standard vincristine (Hagemeister et al., 2013; O'Brien et al., 2013). As vincristine in excess of 2 mg has been associated with severe neuropathy, a standard vincristine dose of 1.5 mg/m² has a dose cap of 2.0 mg per single administration, regardless of the total body surface area (Haim et al., 1994). This helps limit the amount of vincristine exposed to patients with a total body surface area >1.33 m² (Raj et al., 2013). In contrast, this dose cap was not assigned to the liposomal encapsulated vincristine formulation Marqibo during two clinical studies (Hagemeister et al., 2013; O'Brien et al., 2013). It was also identified in two phase I clinical studies that the dose-limiting toxicities and MTD of Marqibo monotherapy were 2.8 and 2.4 mg/m², respectively, whereas the dose-limiting toxicities and MTD of Marqibo in combination with dexamethasone were 2.4 and 2.25 mg/m², respectively (Gelmon et al., 1999; Thomas et al., 2009).

Liposomal encapsulation of vincristine was also recently reported to improve the pharmacokinetics of this drug. A single dose injection of 2.0 mg/m² Marqibo into patients with solid tumors demonstrated an increase in the plasma AUC and a reduction in the clearance rate of encapsulated vincristine compared with patients treated with a single 2.0 mg/m² dose of standard vincristine (Yan et al., 2012). However, the concentration-dependent efficacy of Marqibo was not significantly different in this study when the Marqibo dose was changed to 1.5 or 2.3 mg/m².

Further positive results with Marqibo treatment in humans were reported in clinical trials conducted for its approval (Thomas et al., 2006; O'Brien et al., 2013). In a phase II clinical trial for Marqibo monotherapy (Thomas et al., 2006), 16 adult patients with recurrent or refractory acute lymphocytic leukemia were given liposomal encapsulated vincristine/sodium phosphate mixture (concentration of 0.16 mg/ml) via an intravenous infusion over 60 minutes at a dose of 2 mg/m². The administration was repeated every 2 weeks provided that there was no sign of rapid disease progression or dose-limiting toxicities observed in each patient. Of 14 patients eligible for evaluation, 2 patients (14%) met the overall objective response criteria. This result consisted of one complete response after three doses of liposomal encapsulated vincristine and one partial response after two doses (Thomas et al., 2006).

In the landmark phase I trial of Marqibo that led to its approval by the FDA, some positive responses were observed in adult patients with advanced relapsed or refractory Philadelphia chromosome-negative–acute lymphoblastic leukemia (O'Brien et al., 2013). Sixty-five patients were treated with Marqibo on a weekly basis at a dose of 2.25 mg/m² with no dose capping via intravenous infusion over 60 minutes (O'Brien et al., 2013). The primary efficacy end point was defined as the proportion of patients who achieved a complete response or a complete response with incomplete recovery of peripheral blood neutrophil counts or platelet counts. The overall response rate obtained was 35%, with 20% achieving either a complete response or a complete response with incomplete recovery (O'Brien et al., 2013). Complete response/complete response with incomplete recovery was observed in 25% of patients with an untreated relapse and 14% of patients with a relapse previously refractory to single- or multi-agent antileukemic therapy (O'Brien et al., 2013). Median complete response/complete response with incomplete recovery duration was 23 weeks, with 12 patients proceeding to hematopoietic cell transplantation after Marqibo treatment, whereas 5 patients were long-term survivors (O'Brien et al., 2013).

Apart from being studied as a monotherapy, Marqibo had been investigated in combination with various anticancer drugs. For example, a phase I multi-center study assessed Marqibo in combination with dexamethasone for the treatment of relapsed or refractory acute lymphoblastic leukemia (Thomas et al., 2009). In this investigation, 36 patients were administered with Marqibo (i.v.) weekly at 1.5, 1.825, 2.0, 2.25, or 2.4 mg/m², whereas dexamethasone (40 mg) was given on days 1 through 4 and on days 11 through 14 of each 4-week cycle (Thomas et al., 2009). This study reported the complete response rate of 29% for the 14 patients who underwent therapy as their first salvage attempt (Thomas et al., 2009).

Furthermore, a phase II clinical trial examining the long-term effects of Marqibo in multidrug therapy where conventional vincristine was replaced with Marqibo also demonstrated positive results in patients with hematologic cancer (Hagemeister et al., 2013). This study included 72 patients with untreated aggressive non-Hodgkin lymphoma, 60 of whom had diffuse large B-cell lymphoma. Patients received a combination of cyclophosphamide, DOX, Marqibo (2.0 mg/m² without dose cap), and prednisone with or without rituximab via intravenous infusion for six cycles. Furthermore, patients with disease regression were prescribed up to eight cycles, whereas patients with no enlargement of lymph node larger than 5 cm were subjected to three cycles followed by local radiotherapy (Hagemeister et al., 2013). The overall response rate was 96% (69/72), including 65 (90%) complete responses, 2 (3%) with unconfirmed complete responses, and 2 (3%) with partial responses. The median progression free survival and overall survival were not reached at median follow up of 8 and 10.2 years, respectively. In addition, the 5 and 10 year progression free survival and overall survival were 75 and 63, 87 and 77%, respectively (Hagemeister et al., 2013). Apart from this study, Marqibo is also currently being investigated as a multidrug therapy (prednisone with or without rituximab) in a phase III clinical trial for elderly patients with untreated diffuse large B-cell lymphoma (Hagemeister et al., 2013).

5. Side Effects and Safety of Marqibo.

Despite higher vincristine exposure in Marqibo compared with standard vincristine, Marqibo treatment was well tolerated in most patients (Thomas et al., 2006, 2009; Hagemeister et al., 2013; O'Brien et al., 2013). Thomas et al. (2006) reported two patients with grade 1 peripheral neuropathy after 2 and 4 doses of Marqibo. However, both patients had a history of vincristine-related peripheral neuropathy. They also reported one patient with grade 2 orthostasis and intermittent headaches (Thomas et al., 2006). O’Brien et al. (2013) reported that of 65 patients, 86% had neuropathy-associated adverse events, with 23% reported to be grade 3 peripheral neuropathy-related events. In addition, one grade 4 peripheral neuropathy-related adverse event was also observed in this study. The high incidence of adverse side effects could be explained by the fact that 77% of these patients had reported neuropathy-related symptoms before the study (O'Brien et al., 2013). Apart from neuropathy-related side effects, grade 1–2 constipation, nausea, and vomiting were observed in 34, 22, and 11% of patients, respectively (O'Brien et al., 2013).

In combination therapy studies, patients treated with Marqibo and dexamethasone were reported to exhibit constipation (67%), fatigue (61%), peripheral neuropathy (55%), anemia (50%), and pyrexia (50%), most of which were of grade 1–2 (Thomas et al., 2009). Furthermore, clinically relevant toxicities of grade 3–4 were only observed when the MTD of 2.25 mg/m² was reached (Thomas et al., 2009). Similarly, combination therapy of Marqibo with multiple anticancer agents was also reported to be well tolerated, despite the exposure of up to 35 mg of liposomal encapsulated vincristine (Hagemeister et al., 2013). All nervous system-related adverse events in this latter study were of grade 1–2 magnitude, except for grade 3 peripheral neuropathy that was reported in 2 of 72 patients. No patients reported grade 4 neuropathy-related adverse events (Hagemeister et al., 2013). Constipation of any grade was reported in 15% of patients with no patient reporting grade 3–4 constipation (Hagemeister et al., 2013).

1. Conventional Cytarabine Treatment and Neoplastic Meningitis.

NM, also known as leptomeningeal metastases, is characterized by the infiltration of leptomeninges by cancerous cells (Little et al., 1974; Olson et al., 1974; Theodore and Gendelman, 1981). Once inside the leptomeninges, cancerous cells use cerebrospinal fluid (CSF) as a mean to metastasize throughout the neuraxis (Chamberlain and Corey-Bloom, 1991). NM most often occurs in patients with primary hematologic malignancies, solid tumors of the breast and lungs, and melanoma (Kaplan et al., 1990). The main obstacle for the treatment of NM has always been the poor penetration of chemotherapeutic agents administered systemically across the blood-brain barrier into the CSF (Angst and Drover, 2006). As a result, the treatment of NM may involve radiation therapy and intrathecal chemotherapy (Shapiro et al., 1977). Although radiation therapy is effective in the treatment of leukemic meningitis, its long-term efficacy in other types of NM is uncertain (Murry and Blaney, 2000). Furthermore, craniospinal radiation has been shown to cause acute toxicity, such as myelosuppression, as well as long-term neurologic and neuroendocrine complications (Murry and Blaney, 2000). Consequently, intrathecal administration of the antimetabolites cytarabine and methotrexate has been chosen as a preferred standard treatment of NM (Murry and Blaney, 2000; Angst and Drover, 2006).

The cytotoxic effects of both cytarabine and methotrexate are cell-cycle specific and only occur during the synthesis of DNA (Graham and Whitmore, 1970). The optimal therapeutic efficacy of these cell-cycle-specific agents requires cancer cells to be exposed to low or moderate concentrations of these compounds for a prolonged period of time (Graham and Whitmore, 1970). However, such treatment is limited by the short half-life of these chemotherapeutic agents in the CSF (Bleyer et al., 1978; Zimm et al., 1984). Typically, to achieve the effective drug concentration and to solve the problem of their short half-life, cytarabine and methotrexate are administered by frequent intrathecal injections or a continuous infusion, both of which pose the risk of infections and are inconvenient as well as time consuming for patients (Angst and Drover, 2006). Thus, a sustained release delivery system generating a minimum cytotoxic drug concentration over an extended period of time after an initial intrathecal injection may solve the problems presented in the current standard treatment of NM (Angst and Drover, 2006).

2. The Development of DepoCyt.

DepoCyt is essentially cytarabine encapsulated in multivesicular liposomes (MVLs) generated by DepoFoam technology derived from a water-in-oil-in-water (w/o/w) double emulsification process (Mantripragada, 2002). DepoFoam MVLs containing cytarabine are microscopic spherical particles usually ranging between 3 and 30 μm in size (Murry and Blaney, 2000; Angst and Drover, 2006). Each particle is composed of numerous nonconcentric polyhedral aqueous compartments separated by a lipid bilayer membrane made of lipids, such as dioleoyl PC, dipalmitoyl PG, cholesterol, and triolein (Fig. 13) (Murry and Blaney, 2000; Angst and Drover, 2006). It is possible to imagine MVLs as architecturally analogous to aggregated soap bubbles (Angst and Drover, 2006). As MVLs are much larger than typical unilamellar or multilamellar liposomes, they provide a much greater drug loading capacity. Typical DepoFoam MVLs are composed of ∼4% lipid and 96% water, which are suitable for encapsulating hydrophilic drugs, such as cytarabine (Murry and Blaney, 2000). Although the cytarabine encapsulated MVLs (i.e., DepoCyt) are not considered nanoparticles, they are worth discussing, because they represent a different type of liposome that is capable of encapsulating hydrophilic drugs as well as sensitive molecules, such as proteins, and provide an alternative mean for sustained release.

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Fig. 13.

Morphology of DepoCyt liposomes prepared using DepoFoam Technology. The production of DepoCyt uses DepoFoam technology, which generates multivesicular liposomes (MVLs). Each DepoCyt MLV is composed of numerous non-concentric polyhedral aqueous compartments separated by a lipid bilayer membrane. The arrangement of the MVL gives the whole DepoCyt particle a resemblance of aggregated soap bubbles. DepoCyt MLVs are microscopic spherical particles that usually range in size between 3 and 30 μm.

3. Preclinical Studies of DepoCyt.

The evidence for DepoCyt providing sustained drug release was first demonstrated in animal studies (Kim and Howell, 1987a,b; Kim et al., 1987b). Studies in mice demonstrated that when DepoCyt was injected subcutaneously or intraperitoneally, the serum half-life of cytarabine was significantly longer than that found with conventional cytarabine (Kim and Howell, 1987a,b; Kim et al., 1987b). For example, after a single subcutaneous injection of DepoCyt into BDF1 mice inoculated with L1210 leukemia cells, the half-life of cytarabine was 4 days (96 hours) compared with 10 minutes when unencapsulated cytarabine was administered (Kim and Howell, 1987b). Furthermore, a single dose of DepoCyt was demonstrated to be curative to systemic leukemia in these mice (Kim and Howell, 1987b). A similar result was observed in mice administered with DepoCyt via the intraperitoneal route, showing a terminal half-life of 21 hours versus 16 minutes for the unencapsulated cytarabine (Kim and Howell, 1987a).

The lateral ventricular intrathecal administration of DepoCyt into Sprague-Dawley rats demonstrated an increase in the CSF terminal half-life of cytarabine by a factor of 55 (from 2.7 to 148 hours) and resulted in the maintenance of cytarabine levels above the minimal cytotoxic CSF level of 0.1 μg/ml for at least 14 days (Kim et al., 1987a). Rhesus monkeys (Macaca mulatta) have also been used to study the pharmacokinetics of DepoCyt (Kim et al., 1993b). In this study, six rhesus monkeys of similar weight (9–12 kg) were injected with a single dose of DepoCyt (2 mg/monkey) via an intralumbar route, and one, also of similar weight, was injected with a single dose of conventional unencapsulated cytarabine (2 mg/monkey). Both DepoCyt and unencapsulated cytarabine displayed a biexponential elimination pattern (Kim et al., 1993b). The cytarabine terminal half-life of 156 hours was observed after the intrathecal administration of DepoCyt, whereas a terminal half-life of only 0.74 hour was observed for unencapsulated cytarabine (Kim et al., 1993b). In addition, the cytarabine concentration was maintained above the minimal cytotoxic concentration of 0.1 μg/ml for a longer period of time after DepoCyt administration compared with unencapsulated cytarabine administration (Kim et al., 1993b).

4. Clinical Studies of DepoCyt.

Due to the positive results observed in animal studies, clinical trials focusing on the antitumor activity, pharmacokinetics, and safety of these multivesicular liposomes encapsulating cytarabine were investigated further in patients with NM secondary to various hematologic and nonhematologic malignancies. In a phase I clinical trial, nine patients with leptomeningeal metastasis were given 1–7 cycles of DepoCyt (cytarabine 10 mg/ml) into the lateral ventricle using an Ommaya reservoir (Chamberlain et al., 1993). DepoCyt was administered such that the final quantity of cytarabine ranged from 25 to 125 mg/dose. It was reported in this study that the ventricular concentration of free cytarabine released from DepoCyt into CSF exhibited a biexponential half-life with an initial half-life of 7.2 ± 1.7 hours and a terminal half-life of 140 ± 49 hours. Five of six patients who were eligible for CSF cytologic evaluation responded well to the treatment, with the CSF being cleared of malignant cells within 3 weeks after initial treatment (Chamberlain et al., 1993). The duration of the response was found to be between 2 to greater than 14 weeks (median of 11 weeks), whereas the dose limiting toxicities and MTD were identified to be at a DepoCyt dose of 125 and 75 mg, respectively (Chamberlain et al., 1993).

The second clinical study conducted by the same group of investigators further demonstrated the improved pharmacokinetics of cytarabine in the form of a DepoCyt injection (Kim et al., 1993a). In this second investigation, 12 patients received an increasing dose of DepoCyt by mean of a single intraventricular injection or lumbar intrathecal injection every 2–3 weeks for 15–27 cycles. Therapeutic ventricular CSF concentration of cytarabine was maintained for 9 ± 2 days after the intraventricular injection (Kim et al., 1993a). On the other hand, therapeutic intralumbar concentration of cytarabine was maintained for up to 14 days after the intralumbar injection, with 7 of 9 patients receiving DepoCyt via this route exhibiting some cytologic responses (Kim et al., 1993a). This response was defined as two consecutive negative CSF cytology examinations at least 1 week apart. Although both routes were able to maintain cytarabine concentrations above the minimal cytotoxic level for about 14 days, intralumbar administration was shown to result in higher cytarabine CSF concentrations (Kim et al., 1993a).

A pivotal randomized trial of 28 patients with lymphomatous meningitis was conducted to examine the efficacy and safety of DepoCyt in comparison with conventional cytarabine (Glantz et al., 1999b). From this trial, it was demonstrated that DepoCyt injection (50 mg) every 2 weeks was more effective than a conventional 50 mg injection of cytarabine every 2 weeks. The elimination of malignant cells from CSF was significantly better in DepoCyt-treated patients relative to conventional cytarabine (71 versus 15%, respectively) and the time to neurologic progression of disease (79 versus 42 days, respectively) as well as survival time (100 days versus 63 days, respectively) were observed to be longer (Glantz et al., 1999b). Additionally, patients receiving DepoCyt were observed to be more functional during their daily activities with no sign of cumulative toxicity present. However, DepoCyt treatment was frequently associated with transient symptoms of arachnoiditis, such as headache (27 versus 2%) and nausea/vomiting (9 versus 2%) relative to cytarabine.

Two other studies involving larger numbers of participants were also investigated previously (Jaeckle et al., 2001, 2002). In the first study examining 51 patients diagnosed with NM due to breast cancer, patients were given a 50 mg intrathecal injection of DepoCyt for 1 month followed by an additional 3 months of therapy if patients responded to the initial treatment (Jaeckle et al., 2001). Of 43 patients cytologically evaluated, 28% showed no sign of malignant cells in the CSF, whereas the median time to neurologic progression was 49 days and median survival time was 88 days. In the second study, 110 patients diagnosed with NM were given the same treatment as the first study above (Jaeckle et al., 2002). Similarly, 19 of 70 patients cytologically evaluated (27%) showed some responses, whereas the median time to neurologic progression and median survival time were 55 days and 95 days, respectively (Jaeckle et al., 2002). Only mild headache (4–11%) and arachnoiditis (6–19%) were observed throughout both studies (Jaeckle et al., 2001, 2002).

Apart from being investigated in comparison with conventional cytarabine, DepoCyt was also previously compared with an alternative drug, methotrexate, in patients with NM secondary to solid tumors (Glantz et al., 1999a). On an intent-to-treat basis, 8 of 31 patients (26%) receiving DepoCyt compared with 6 of 30 (20%) patients receiving methotrexate, demonstrated some responses. The median time to disease progression was 58 days for DepoCyt-treated patients compared with 30 days for those treated with methotrexate. Median survival time was also found to be higher for patients receiving DepoCyt compared with those receiving methotrexate (105 versus 78 days), although the improvement was not significant (Glantz et al., 1999a).

5. Side Effects and Safety of DepoCyt.

The toxic side effects associated with DepoCyt are similar to those of conventional cytarabine. In the phase I clinical study, the predominant toxic side effects of DepoCyt observed at the MTD of 75 mg were symptoms associated with drug-induced chemical arachnoiditis (Chamberlain et al., 1993). These symptoms included grade 1–2 fever (50%), grade 1–2 headache (38%), grade 1–2 back and/or neck pain (38%), and grade 1–2 nausea or vomiting (25%). Less than 15% of patients had grade 3–4 nausea or vomiting. The observed side effects of DepoCyt were transient, occurring typically on day 2 to day 4 and were not apparent on day 1 and days 5 to 7. These toxicities were confirmed in two larger studies (Jaeckle et al., 2001, 2002). Both studies (51 and 110 patients) reported symptoms of headache, nausea, vomiting, and arachnoiditis, predominately at a grade 1–2 magnitude (Jaeckle et al., 2001, 2002). All side effects were also generally transient, resolving after the end of the treatment cycle they occurred in. In all studies, the toxic side effects could be ameliorated by a concomitant systemic administration of dexamethasone 2–4 mg twice daily for 5 days, beginning on the day of initial DepoCyt administration (Chamberlain et al., 1993; Jaeckle et al., 2001, 2002).

DepoCyt is currently formulated as a sterile suspension in NaCl 0.9% weight/volume solution for intrathecal injection. It is available in a ready-to-use, single-use vial, with each vial containing 50 mg cytarabine in 5 mL of NaCl solution (Murry and Blaney, 2000). A single dose of DepoCyt administered intrathecally at 50 and 75 mg was shown to elevate the levels of cytarabine above its minimum cytotoxic concentration (>0.1 μg/mL) for 2 weeks (Murry and Blaney, 2000). Thus, the recommended DepoCyt dosage for adults is 50 mg every fortnight. In addition, as DepoFoam particles are generally denser than the suspending medium, DepoCyt must be resuspended by gently shaking the vial immediately before injection (Murry and Blaney, 2000).

A. Redox-sensitive Liposomes.

A high redox potential difference exists between the intracellular compartment and extracellular space due to the abundant amount of reducing agents, such as glutathione, within cells compared with extracellular fluid (Schafer and Buettner, 2001). The distinction in redox potential between the two compartments may be exploited by redox-sensitive liposomes. Disulfide bonds can be used as a linker to conjugate specific ligands onto the surface of liposomes or it can be used as a disulfide bridge to prepare thiolated lipid, which can be incorporated into liposomes (Ganta et al., 2008). Upon arrival into intracellular compartment, ligands conjugated by disulfide bond or thiolated liposomes can be destabilized by the reduction of the disulfide bond by glutathione, resulting in the release of the encapsulated contents (Ganta et al., 2008).

Disulfide-mediated redox-sensitive liposomes are generally prepared by using a mixture of standard phospholipids and a small amount of lipids containing disulfide bonds linking their hydrophobic and hydrophilic regions together (Ganta et al., 2008). The coupling of PEG_2000Da to DSPE via 3,3′dithiopropionate resulted in a DSPE-PEG_2000Da derivative containing a disulfide bond, known as PEG-α-aminocarbonylethyl-dithiopropionyl-DSPE, or PEG-DTP-DSPE (Kirpotin et al., 1996). When PEG-DTP-DSPE was incorporated into DOPE liposomes, liposomal fusion and the rapid and complete release of encapsulated contents was observed after thiolytic cleavage of PEG from liposomes (Kirpotin et al., 1996). Similarly, anti-CD19-targeted liposomes composed of DOPE, CHEMS and PEG-DTP-DSPE exhibiting both pH and redox sensitivity also demonstrated improved DOX delivery and anticancer activity against B-lymphoma cells compared with non-pH-sensitive liposomes (Ishida et al., 2001b). The in vivo study of these liposomes in mice bearing B-cell lymphoma also showed their improved antitumor activity over non-pH-sensitive anti-CD19-targeted liposomes, despite having a faster rate of drug release and higher clearance (Ishida et al., 2001b). Another DSPE-PEG_2000Da derivative containing the disulfide bond linkage was prepared using dithiobenzylurethane (Zalipsky et al., 1999). This derivative is believed to be more effective than PEG-DTP-DSPE because it does not require potent thiolytic agents for its cleavage (Zalipsky et al., 1999). The cleavage of the dithiobenzylurethane linkage by cysteine, a mild thiolytic agent, was reported to be relatively efficient, resulting in a release of dye from DOPE liposomes within 60 minutes (Zalipsky et al., 1999). For more information, please refer to Sawant and Torchilin (2010), Ganta et al. (2008), and Saito et al.(2003).

Although enhanced drug delivery by redox-sensitive liposomes may be promising, the cleavage of these liposomes requires high concentration of reducing agents, which may not be present in some pathologic tissues (Sawant and Torchilin, 2010). In addition, the reduction of redox-sensitive liposomes may not be optimal, because many liposomes enter cells via endosomes, whereas the reduction process takes place within the cytosol (Sawant and Torchilin, 2010). Therefore, redox-sensitive liposomes must escape from the endosome first before destabilization (Sawant and Torchilin, 2010).

B. Ultrasound-responsive Liposomes.

Ultrasound can be classified into low frequency and high frequency ultrasound (Ahmed et al., 2015). High frequency ultrasound is often associated with thermal effects and can be used to trigger drug release from temperature-sensitive liposomes as described above in section IV.C.2 (Ahmed et al., 2015). In contrast, low frequency ultrasound (LFUS) is associated with mechanical effects, such as oscillation and cavitation (Nyborg, 2001). Acoustic cavitation occurs as a result of the interaction between acoustic waves and gas bubbles (Leighton, 2007). When ultrasound excites gas bubbles, they oscillate in response to acoustic pressure (Ahmed et al., 2015). When acoustic pressure is increased further, the oscillation becomes nonlinear (Ahmed et al., 2015). This can lead to the increase in size of the gas bubble and their eventual collapse (Ahmed et al., 2015). It was shown that LFUS can be used to trigger the release of liposomal contents mechanically.

A study on the release of DOX from Doxil after exposure to LFUS (20 kHz) for 30 minutes reported that 85% of DOX was released from Doxil in saline solution, and 61% was released in human plasma (Schroeder et al., 2009b; Ahmed et al., 2015). In contrast, exposure to 30 minutes of higher frequency ultrasound (1 MHz) resulted in the release of 58% of DOX from Doxil in saline and 5% release in human plasma. A further reduction in DOX release was observed when 3 MHz ultrasound was used (Schroeder et al., 2009b; Ahmed et al., 2015). In mice bearing J6456 murine lymphoma tumors, exposure to LFUS for 2 minutes resulted in the release of nearly 70% of cisplatin from PEGylated liposomes (hydrogenated soybean PC:cholesterol:DSPE-PEG_2000Da 51:44:5 mol%) compared with the release of <3% of cisplatin from PEGylated liposomes without LFUS (Schroeder et al., 2009a). Furthermore, when these liposomes were injected intravenously into mice bearing C26 colon adenocarcinoma tumors followed by LFUS 24 hours postadministration, they demonstrated more efficacious antitumor activity than free cisplatin and PEGylated liposomal cisplatin without LFUS (Schroeder et al., 2009a). For more information on ultrasound-responsive liposomes, please refer to reviews by Ahmed et al. (2015) and Schroeder et al. (2009b).

As liposomes contain no gas, they are only partially sensitive to ultrasound (Ahmed et al., 2015). Recently, liposomes with echogenic properties were developed that contain a gas phase or an emulsion that is vaporizable, making them more responsive to ultrasound (Ahmed et al., 2015). For instance, liposomes containing a liquid emulsion of perfluorocarbons, called eLiposomes, have been investigated (Javadi et al., 2012; Lattin et al., 2012; Javadi et al., 2013). These liposomes can be ruptured by reducing the local pressure below the vapor pressure of the perfluorocarbon emulsion, allowing it to vaporize and making the liposome sensitive to ultrasound (Ahmed et al., 2015). However, more studies on these eLiposomes are needed because their design still requires optimization. For more information on these liposomes, please refer to Javadi et al. (2012, 2013) and Lattin et al. (2012).

Alternatively, liposomal DOX containing perfluorobutane microbubbles has been investigated in comparison with normal liposomal DOX formulations (Lentacker et al., 2010). This microbubble-liposome system was demonstrated to increase BLM melanoma cell death in vitro by 2-fold when exposed to ultrasound (1 MHz) compared with standard liposomal DOX (Lentacker et al., 2010). This was suggested to be due to the sonoporation of the cell membrane by the implosion of microbubbles upon ultrasound exposure, causing an increase in the cellular uptake of DOX (Lentacker et al., 2010).

C. Magnetic Liposomes.

The use of magnetic liposomes is a relatively recent approach in targeted drug delivery using a magnet as a means of targeting (Nobuto et al., 2004). In this strategy, liposomes are loaded with a drug and a ferromagnetic material and are infused into the subject intravenously. The subject’s tissue affected by cancer is subsequently placed between two poles of a magnet to direct nanoparticles to the targeted site. Generally, iron oxide nanoparticles of size less than 10 nm, namely, magnemite and magnetite, are incorporated into liposomes to achieve the magnetic property (Sawant and Torchilin, 2010; Deshpande et al., 2013). In one study, hamsters bearing a limb osteosarcoma were intravenously administered with magnetic liposomes encapsulating DOX, and the limb was subsequently placed between two poles of a 0.4 Tesla magnet (Nobuto et al., 2004). After 60 minutes, a 4-fold increase in the tumor drug concentration was observed (Nobuto et al., 2004). Alternatively, the magnet can be implanted within the tumor. Using the same osteosarcoma-bearing hamster model, this strategy demonstrated enhanced accumulation of DOX encapsulated magnetic liposomes in the tumor vasculature and improved inhibition of tumor growth (Kubo et al., 2001). This strategy also demonstrated a 25-fold increase in the accumulation of ^99mTc-albumin loaded magnetite-containing liposomes within the kidney implanted with a magnet (Babincova et al., 2000). However, because iron oxide nanoparticles must be incorporated into magnetic liposomes together with the encapsulated drug, it is important that the incorporation of these particles does not affect the efficacy of the entrapped drug (Sawant and Torchilin, 2010). It was reported that the drug loading efficiency of magnetic liposomes was affected at high magnetite concentrations, and thus, the incorporation of these iron oxide nanoparticles decreased drug encapsulation (Dandamudi and Campbell, 2007).

It is also possible to modify magnetic liposomes to give them temperature-sensitive and active targeting properties. For example, magnetic nanoparticles and DOX have been incorporated into temperature-sensitive liposomes composed of DPPC, cholesterol, DSPE-PEG_2000Da, and DSPE-PEG_2000Da-folate at a molar ratio of 80:20:4.5:0.5 (Pradhan et al., 2010). When these liposomes were targeted to KB and HeLa cell cultures by a magnetic field, the cellular uptake of DOX was increased substantially compared with Doxil, nonmagnetic folate receptor-targeted liposomal DOX, and free DOX (Pradhan et al., 2010). Furthermore, the application of hyperthermia (at 42.5 and 43.5°C) and a magnetic field synergistically increased the antitumor activity of these folate receptor-targeted temperature-sensitive magnetic liposomes containing DOX (Pradhan et al., 2010).

D. Enzyme-sensitive Liposomes.

Stimuli-sensitive liposomes can be generated to exploit the expression of various enzymes within the tumor (Deshpande et al., 2013). Enzymes that have been found to be overexpressed within tumor tissue include MMPs, phospholipase A₂, alkaline phosphatase, transglutaminase, and PI-specific phospholipase (Arias, 2011). In the presence of these catalytic proteins, enzyme-sensitive liposomes can be engineered to destabilize and release their encapsulated contents at the tumor site (Arias, 2011). A linker that is cleavable by enzymes that are overexpressed in tumors can be used to modify the liposomes (Danhier et al., 2010; Zhu et al., 2012).

For instance, a matrix metalloproteinase 2 (MMP-2)-cleavable octapeptide linker composed of Gly-Pro-Leu-Gly-Ile-Ala-Gly-Gln has been used to conjugate PEG_2000Da with DSPE and incorporated into liposomal formulations (Zhu et al., 2012). It was reported that these MMP-2-sensitive DSPE-PEG_2000Da conjugates were incorporated into liposomes modified with cell-penetrating peptide, namely transactivator of transcription peptide (TATp), and antinucleosome antibody 2C5 (Zhu et al., 2012). Consequently, MMP-2-sensitive DSPE-PEG_2000Da acts as a steric shield for these liposomes, which become detached in the presence of high concentrations of MMP-2 enzymes, leading to the exposure of other surface functionalities (Zhu et al., 2012). During in vitro studies, Gly-Pro-Leu-Gly-Ile-Ala-Gly-Gln octapeptides were cleaved in the presence of MMP-2, removing the PEG shield from liposomes and exposing other active-targeting moieties (Zhu et al., 2012). Subsequently, the enhanced cellular internalization of these liposomes by mouse breast cancer cells (4T1) was observed (Zhu et al., 2012).

Additionally, a lipopeptide generated by the conjugation of matrix metalloproteinase-9 (MMP-9) substrate peptide to a fatty acid chain, has been incorporated into liposomes (Banerjee et al., 2009; Deshpande et al., 2013). These liposomes were reported to rapidly release a significant amount of encapsulated contents in breast cancer cells (MCF-7) expressing high levels of MMP-9, while slowly releasing their contents in colorectal adenocarcinoma cells (HT-29) with low levels of MMP-9 (Banerjee et al., 2009).

1. Preparation of Solid Lipid Nanoparticles.

2. Drug Incorporation and Drug Loading Capacity.

The drug loading capacity of SLNs is determined by the following parameters: 1) the solubility of the drug in the molten lipid; 2) the miscibility of the melted drug and molten lipid; 3) the chemical and physical structures of solid lipid matrix; and 4) the polymorphic state of the lipid material (Muller et al., 2000b).

To achieve adequate drug loading in SLNs, the encapsulated drug must be sufficiently soluble in the molten lipid. As solubility decreases with decreasing temperature, it is also necessary for the drug to have sufficiently high solubility to avoid separation from the molten or solid lipid (Mehnert and Mader, 2001). Solubilizers can be added to improve drug solubility in the molten lipid (Muller et al., 2000b). Additionally, the presence of mono- and diglycerides in the lipid mixture used to prepare the SLN matrix can further improve drug solubilization (Muller et al., 2000b).

The nature of lipid, including its chemical and physical structure, also plays an important role in drug loading capacity. Lipids that form particles with a perfect crystalline lattice matrix will cause the expulsion of the drug (Westesen et al., 1997). In contrast, a complex lipid mixture consisting of different types of lipids and fatty acid chain lengths produces imperfect crystalline lattices within the SLN matrix, and thus, will accommodate more drugs within the matrix as a result of lattice imperfections (Muller et al., 2000b).

As the degree of crystallization of the lipid matrix determines the drug loading capacity of SLNs, lipid crystal polymorphism may also play a crucial role in the drug loading efficiency of SLNs (Muller et al., 2000b). The crystallization of the lipids in nanoparticles is different from that of the bulk material (Westesen et al., 1993). Although the lipids in nanoparticles recrystallize partially into a less stable α-crystalline form, the bulk lipid usually recrystallizes preferentially and rapidly into the more stable β-crystalline form via β-modification (Westesen et al., 1993). With increasing levels of the stable β-crystalline form within SLNs, the crystalline lipid lattice becomes more perfect and the number of imperfections decreases (Westesen et al., 1993, 1997). Thus, fewer drug molecules can be accommodated within SLN matrices with higher levels of the β-crystalline form, which promotes drug expulsion and the lowering of the drug loading capacity (Bunjes et al., 1996; Westesen et al., 1997). On the other hand, an increase in the α-crystalline form within SLNs improves the drug loading capacity of these particles (Westesen et al., 1997).

3. Studies of Solid Lipid Nanoparticles in Cancer Therapy.

A number of anticancer drug classes have been encapsulated within SLNs and examined both in vitro and in vivo since the early 1990s, including anthracyclines, taxanes, camptothecins, etoposide, flurodeoxyuridine, and retinoic acid (Cavalli et al., 1993; Yang et al., 1999; Wang et al., 2002; Lim et al., 2004; Serpe et al., 2004; Harivardhan Reddy et al., 2005).

Animal studies have shown that SLNs can increase the AUC of encapsulated drugs by 3- to 20-fold, as well as significantly extending the half-life of the encapsulated agent compared with the corresponding free drug (Yang et al., 1999; Zara et al., 1999; Fundaro et al., 2000; Wang et al., 2002; Zara et al., 2002). Moreover, the stealth form of SLNs that are generated by PEGylation further improves the AUC and the half-life of encapsulated drugs to a greater extent than non-stealth SLNs (Fundaro et al., 2000; Zara et al., 2002). The cytotoxicity of cholesteryl butyrate, DOX, and PTX encapsulated SLNs were previously investigated in the human colorectal HT-28 cancer cell line (Serpe et al., 2004). SLNs containing cholesteryl butyrate and DOX demonstrated higher cytotoxicity than their conventional drug formulations, showing lower IC₅₀ values (Serpe et al., 2004). Similarly, PTX encapsulated SLNs exhibited higher cytotoxicity than an equivalent amount of the conventional PTX formulation (Serpe et al., 2004).

To overcome resistance to anticancer agents, P-glycoprotein inhibiting chemosensitizers have been incorporated into SLNs (Fischer et al., 1998; Krishna and Mayer, 2000; Ugazio et al., 2002; Planting et al., 2005). The incorporation of chemosensitizers into SLNs could potentially improve the potency, safety, and specificity of these compounds (Krishna and Mayer, 2000). Additionally, their incorporation may reduce the interaction between chemosensitizers and cytotoxic drugs observed in conventional formulations when administered simultaneously (Fischer et al., 1998; Planting et al., 2005). Previously, the chemosensitizer cyclosporin-A was successfully encapsulated within SLNs (Ugazio et al., 2002). Such a delivery system was shown to release cyclosporin-A slowly over a long period of time with less than 4% of the chemosensitizer released after 120 minutes (Ugazio et al., 2002).

The enhanced specificity by active targeting has been investigated in a number of nanoparticles for many years (Chang et al., 2009; Low and Kularatne, 2009; Ulbrich et al., 2009). Recently, this strategy was assessed on SLNs encapsulating anticancer drugs (Stevens et al., 2004). Folate receptor-targeting SLNs containing the PTX-prodrug were shown to improve both the uptake and cytotoxicity of this agent in cells expressing the folate receptor compared with nontargeted SLNs (Stevens et al., 2004). This improvement was also observed in vivo, where folate receptor-targeted SLNs containing the PTX-prodrug significantly increased tumor growth inhibition and animal survival time relative to nontargeted SLNs and PTX in Cremophor EL (BASF SE, Lugwigshafen, Germany) (Stevens et al., 2004).

Similar to liposomes, SLNs have been investigated for their potential use in gene delivery (Tabatt et al., 2004a,b), because it is believed that these particles should be at least as effective as liposomes in delivering genetic material (Wong et al., 2007). Both SLNs and liposomes incorporating the same cationic lipid were demonstrated to have similar transfection efficiency in vitro in Cos-1 monkey kidney fibroblast-like cells (Tabatt et al., 2004a). Furthermore, alterations in the lipid composition between the two delivery systems were able to influence the transfection efficiency (Tabatt et al., 2004a). In addition, low toxicity and high transfection efficiency could be achieved by selecting combinations of two-tailed cationic lipids and matrix lipids (Tabatt et al., 2004b). Hence, there is a great possibility that SLN delivery system may be used in gene therapy for cancer management (Wong et al., 2007).

4. Disadvantages of Solid Lipid Nanoparticles.

Although SLNs provide many advantages over existing nanoparticle drug delivery systems, they do possess some drawbacks, particularly low encapsulation of hydrophilic drugs and nonuniform drug release (Wong et al., 2007). SLNs exhibit poor encapsulation of hydrophilic drugs as the drug to be encapsulated must be adequately dissolved within the melted lipid droplets to achieve high drug loading during SLN preparation (Mehnert and Mader, 2001). The addition of organic counterions during SLN preparation to form ionic pairs with charged drug molecules has been proposed to solve this problem (Cavalli et al., 1993, 2000). Another strategy involves the formation of polymer-lipid hybrid nanoparticles (PLNs), in which ionic drugs are electrostatically neutralized by polymer counterions and the drug-polymer complexes are subsequently incorporated into lipids for nanoparticle preparation (Fig. 15) (Wong et al., 2004, 2006; Li et al., 2006; Wong et al., 2006). Such a strategy has been shown to improve the encapsulation efficiency of ionic drugs, such as DOX-HCl and verapamil HCl from 20 to >80% (Wong et al., 2004, 2006). PLNs are not to be confused with lipid-polymer hybrid nanoparticles discussed below (see section IX.C).

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Fig. 15.

Incorporation of hydrophilic drugs into SLNs using polymers. One of the strategies for incorporating hydrophilic drugs into SLNs is to use charged polymers. In this procedure, the ionic form of the hydrophilic drug is electrostatically neutralized by counterions on the polymer. The drug-polymer complexes are subsequently incorporated into lipids for SLN preparation. This strategy gives rise to polymer-lipid hybrid nanoparticles (PLNs), which are not to be confused with lipid-polymer hybrid nanoparticles (LPNs).

The issue of nonuniform drug release by SLNs is related to the burst release or burst effect, which is the fast initial release of a large dose of the encapsulated drug followed by slow incomplete release (zur Muhlen et al., 1998). Burst release can create serious complications because of the cytotoxicity of anticancer drugs (Wong et al., 2007). Such behavior is attributed to the uneven distribution of the drug within SLNs due to the perfect lipid crystal structure that forms the SLN matrix (Westesen et al., 1993; Siekmann and Westesen, 1994; Bunjes et al., 1996), which lacks free space to accommodate large quantities of drug molecules. Thus, the majority of the drug is concentrated at or near the surface of the particles, which can then diffuse away rapidly into the surrounding medium, resulting in the burst release effect (zur Muhlen et al., 1998). This problem can be avoided by: 1) reducing the surfactant concentration; 2) cooling the lipid emulsion quickly to allow nanoparticles to be made up mainly of solid drug; and/or 3) by selecting lipid compositions that do not form perfect crystal lattices, such as mono-, di-, or triglycerides of different chain lengths (Wissing et al., 2004).

1. Preparation of Nanostructured Lipid Carriers.

NLCs can be produced by several methods, most of which are adopted from polymeric nanoparticle preparations. These methods include high-pressure homogenization (Stecova et al., 2007; Huang et al., 2008; Ruktanonchai et al., 2009); microemulsion (Doktorovova et al., 2010; Souza et al., 2011); phase inversion (Souto et al., 2007); emulsification by sonication (Das and Chaudhury, 2011); emulsification-solvent-evaporation (ESE) (Lin et al., 2010); solvent diffusion and solvent injection/solvent displacement (Schubert and Muller-Goymann, 2003); and the membrane contactor method (Charcosset et al., 2005). However, the most preferred method of NLC preparation is high-pressure homogenization (Iqbal et al., 2012).

2. Studies of Nanostructured Lipid Carriers in Cancer Therapy.

As NLCs are a relatively new development, not many studies have been focused on their use with anticancer drugs. Docetaxel encapsulated NLCs (DTX-NLCs) were developed for in vitro assessment against three human cancer cell lines (HepG2, SKOV3, and A549) and one murine B16 malignant melanoma cell line (Liu et al., 2011). In comparison with Duopafei (Qilu Pharmaceutical Co. Ltd, Jinan, China; the standard formulation of Docetaxel), DTX-NLCs were demonstrated to have a significantly lower IC₅₀ value than Duopafei in all cell lines and were more cytotoxic against A549 cells by inducing a higher level of apoptosis and G₂/M arrest (Liu et al., 2011). This study further investigated the in vivo anticancer activity of DTX-NLCs against Duopafei in Kunming mice bearing murine B16 malignant melanoma xenografts (Liu et al., 2011). Consequently, DTX-NLCs were reported to be more effective than Duopafei in vivo, and showed a higher inhibition rate at doses of 10 and 20 mg/kg (62.69 and 90.36% compared to the control, respectively) relative to Duopafei at 10 mg/kg, which exhibited an inhibition rate of 42.7% of the control (Liu et al., 2011).

PTX- and DOX-loaded NLCs, as well as NLCs that have been surface modified with folic acid and stearic acid have also been investigated against cancer cells, including multi-drug resistant (MDR) cells (Zhang et al., 2008b). Indeed, PTX-NLCs demonstrated a high level of cytotoxicity in MCF-7 and MCF-7/ADR cells, whereas DOX-NLCs only showed a high level of cytotoxicity against MCF-7/ADR cells (Zhang et al., 2008b). The ability of PTX-NLCs and DOX-NLCs to reverse MDR was 34.3- and 6.4-fold greater than the free drugs, respectively (Zhang et al., 2008b). A similar trend in cytotoxicity was also observed using SKOV3 and SKOV3-TR30 cells, where DOX-NLCs and PTX-NLCs were 2.2- and 31.0-fold more effective at reversing drug resistance, respectively, than the free drugs (Zhang et al., 2008b). In addition, the surface modification of NLCs with folic and stearic acid further enhanced the cytotoxicity of PTX and DOX against cancer cells, including MDR cells (Zhang et al., 2008b).

Additionally, PTX has been incorporated into hyaluronic acid-coated NLCs (HA-NLCs) to improve nanoparticle specificity (Yang et al., 2013). It was demonstrated that PTX was released slowly from PTX-loaded HA-NLCs and these nanoparticles were more cytotoxic than standard PTX in three cancer cell lines (i.e., the B16, CT26, and HCT116 cell lines) (Yang et al., 2013). In vivo studies further showed that PTX-loaded HA-NLCs exhibited a higher anticancer efficacy and were better tolerated by B16-bearing Kumming mice compared to conventional PTX. This could be due to the prolonged circulation time of PTX-loaded HA-NLCs and the increased accumulation of these nanoparticles in the tumor (Yang et al., 2013).

PEG-NLCs have also been prepared to encapsulate 10-hydroxycamptothecin (HCPT) and compared with free HCPT and HCPT encapsulated NLCs (HCPT-NLCs) (Zhang et al., 2008c). During in vitro studies, such PEGylated particles were able to accumulate within human lung adenocarcinoma epithelial A549 cells and, during in vivo studies, had longer circulation times and decreased uptake by the RES compared with free HCPT and unmodified HCPT-NLCs (Zhang et al., 2008c). Moreover, PEGylated HCPT-NLCs demonstrated superior antitumor efficacy against A549 lung cancer xenografts in vivo compared to free HCPT and unmodified HCPT-NLCs (Zhang et al., 2008c).