International Journal of Pharmaceutics 491 (2015) 49–57

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International Journal of Pharmaceutics journal homepage: www.elsevier.com/locate/ijpharm

Pharmaceutical nanotechnology

Enzymatic action of phospholipase A2 on liposomal drug delivery systems Anders H. Hansena,b , Ole G. Mouritsena,b , Ahmad Arouria,b,* a

MEMPHYS-Center for Biomembrane Physics, Department of Physics, Chemistry, and Pharmacy, University of Southern Denmark, Odense, Denmark The Lundbeck Foundation Nanomedicine Research Center for Cancer Stem Cell Targeting Therapeutics (NanoCAN), University of Southern Denmark, Odense, Denmark b

A R T I C L E I N F O

A B S T R A C T

Article history: Received 9 April 2015 Received in revised form 3 June 2015 Accepted 4 June 2015 Available online 6 June 2015

The overexpression of secretory phospholipase A2 (sPLA2) in tumors has opened new avenues for enzyme-triggered active unloading of liposomal antitumor drug carriers selectively at the target tumor. However, the effects of the liposome composition, drug encapsulation, and tumor microenvironment on the activity of sPLA2 are still not well understood. We carried out a physico-chemical study to characterize the sPLA2-assisted breakdown of liposomes using dye-release assays in the context of drug delivery and under physiologically relevant conditions. The influence of temperature, lipid concentration, enzyme concentration, and drug loading on the hydrolysis of 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC, Tm = 42
C) liposomes with snake venom sPLA2 was investigated. The sensitivity of human sPLA2 to the liposome composition was checked using binary lipid mixtures of phosphatidylcholine (PC) and phosphatidylglycerol (PG) phospholipids with C14 and C16 acyl chains. Increasing temperature (36– 41
C) was found to mainly shorten the enzyme lag-time, whereas the effect on lipid hydrolysis rate was modest. The enzyme lag-time was also found to be inversely dependent on the lipid-to-enzyme ratio. Drug encapsulation can alter the hydrolysis profile of the carrier liposomes. The activity of human sPLA2 was highly sensitive to the phospholipid acyl-chain length and negative surface charge density of the liposomes. We believe our work will prove useful for the optimization of sPLA2-susceptible liposomal formulations as well as will provide a solid ground for predicting the hydrolysis profile of the liposomes in vivo at the target site. ã2015 Elsevier B.V. All rights reserved.

Keywords: Bio-responsive liposomes Drug delivery system Anticancer Phospholipase A2 enzyme Triggered drug release Dye-release assay

1. Introduction The concept of using liposomes for drug delivery in cancer chemotherapy was introduced by Gregoriadis et al. (1974) back in the early seventies). Twenty years later, the first liposomal formulation loaded with the anticancer drug doxorubicin (Doxil1) was authorized for cancer treatment, which was followed by few other medicinal products (Barenholz, 2012; Chang and Yeh, 2012). Although this approach can improve the drug pharmacokinetics and reduce off-target toxicities, relying on passive drug release does not secure and improve the drug efficacy (Barenholz, 2012). With an aim to enhance drug efficacy, there has been extensive research in the past decade to develop tools to actively deliver the

* Corresponding author at: MEMPHYS-Center for Biomembrane Physics, Department of Physics, Chemistry, and Pharmacy, University of Southern Denmark, Campusvej 55, DK-5230 Odense, Denmark. Fax: +45 6550 4048. E-mail addresses: [email protected] (A.H. Hansen), [email protected] (O.G. Mouritsen), [email protected], [email protected] (A. Arouri). http://dx.doi.org/10.1016/j.ijpharm.2015.06.005 0378-5173/ ã 2015 Elsevier B.V. All rights reserved.

drugs to the target cell in sufficiently high concentrations (Park, 2014). One example is bio-responsive nanoparticles, including liposomes, that change their properties in response to the surrounding microenvironment or to an external stimulus (Arouri et al., 2013). Phospholipase A2 (PLA2) is a large family of lipolytic enzymes that can hydrolyze the sn-2 ester bond of phospholipids producing an equimolar amount of the corresponding fatty acid and lysolipid (Burke and Dennis, 2009; Kudo, 2004; Murakami et al., 2011; Six and Dennis, 2000). The family is constituted of three main groups; cytosolic PLA2 (cPLA2), intracellular calcium-independent PLA2 (iPLA2), and secretory PLA2 (sPLA2) (Murakami and Taketomi, 2015). The latter is present extensively in many mammalian tissues like pancreas and liver as well as in insect and snake venoms (Six and Dennis, 2000). Unlike classical Michaelis–Menten enzymes, sPLA2 has preference for aggregated lipid superstructures (the enzyme is not active on single lipid molecules) and therefore several steps are involved in one catalytic cycle (Berg et al., 2001; Lambeau and Gelb, 2008).

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The enzyme requires calcium as a cofactor, and the enzymatic activity is determined by the composition, morphology, and physico-chemical properties of the lipid membrane (Halperin and Mouritsen, 2005; Hønger et al., 1996; Høyrup et al., 2004; Mouritsen et al., 2006). Furthermore, sPLA2 enzymes display a lag-burst behavior, implying an increase in catalytic activity by two to three orders of magnitude after an intrinsic lag-time with very low enzymatic activity (Mouritsen et al., 2006). The activation of the enzyme has been linked to the alterations in the composition and properties of the lipid membrane after the initial hydrolysis as well as to conformational changes in the enzyme structure and formation of sPLA2 aggregates (Code et al., 2008; Mouritsen et al., 2006). The extensive hydrolysis of the lipid membrane and the sPLA2 aggregation leading to the formation of macroscopic amyloid fibrils will eventually attenuate the activity of sPLA2 (Arouri and Mouritsen, 2012; Code et al., 2008). The intrinsic lag-time can range from seconds to hours (Hønger et al., 1996; Høyrup et al., 2001; Mouritsen et al., 2006) and is modulated by the heterogeneities and defects in the lipid bilayer, for instance, formed during membrane phase transitions, at lipid phase segregations, and in the presence of lipopolymers (Høyrup et al., 2004; Leidy et al., 2006; Mouritsen et al., 2006; Simonsen, 2008; Vermehren et al., 1998). In the human body, PLA2 enzymes are involved in the downregulation of cell signals via the cleavage “deactivation” of bioactive phospholipids. Pathologically, PLA2-assisted release of fatty acids and lysolipids can provoke a variety of proinflammatory and metabolic conditions (Hui, 2012; Murakami and Kudo, 2003; Tselepis and John Chapman, 2002). Due to their suspected role in tumorigenesis and metastasis, PLA2 enzymes, like sPLA2 subtype IIA, are overexpressed in several cancer types, like prostate, breast and colon cancer (Belinsky et al., 2007; Cai et al., 2012; Hu et al., 2011; Murata et al., 1993; Scott et al., 2010; Tosato et al., 2010; Tribler et al., 2007). The percentage of cancer patients with elevated levels of sPLA2 can range from 28% (e.g., lung or gastric cancer) to 100% (e.g., liver cancer) (Scott et al., 2010). The serum level of sPLA2 in healthy subjects is strictly low, with an average level of 2.2  0.1 ng mL1 (range 1.4–4.2 ng mL1) (Matsuda et al., 1991) or 0.17 nM using a sPLA2 molecular weight of 13,118 g mol1 (Martin-Moutot and Rochat, 1979). The mean concentration of sPLA2 found in effusions of eight different cancer types was 54.3  44.3 ng mL1 (range 9.9–188.3 ng mL1) (Abe et al., 1997) corresponding to 4.1 nM. The serum level of sPLA2 in cancer patients is mostly only slightly higher than in healthy subjects. For instance, the average serum level of sPLA2 in patients with different types of cancer was 9 ng mL1 (range 2–100 ng mL1) (Abe et al., 1997), and the average concentration found in the serum of colon cancer patients was 6.8 ng mL1 (range 1– 22 ng mL1) (Tribler et al., 2007). The serum level of sPLA2 is

strongly dependent of the stage of cancer and the presence of distant metastases, whereas factors like the tumor size, histological grade, and organ site are of less importance (Abe et al., 1997; Yamashita et al., 1994). The elevated levels of sPLA2 in tumors have been exploited to develop sPLA2-susceptible liposome-based drug delivery systems loaded with anticancer drugs. After their passage through the vascular fenestrae of cancer tissues with elevated levels of sPLA2 (Matsumura and Maeda, 1986; Moghimi and Farhangrazi, 2014), the enzyme will trigger the release of the drugs precisely at the target site. This strategy will aid to overcome some of the limitations of conventional liposomal formulations (Andresen et al., 2005). The feasibility of this platform has been demonstrated before for doxorubicin- and cisplatin-loaded liposomes (Andresen et al., 2005; de Jonge et al., 2010), liposome-forming lipid-like anticancer prodrugs (Arouri et al., 2013; Pedersen et al., 2009, 2010), as well as certain double lipid-prodrug systems (Arouri and Mouritsen, 2011, 2012). Several studies have postulated that the hydrolytic products, i.e., fatty acids and lysolipids, due to their membraneperturbing effects, can reduce the permeability barrier of the target cancer cell membrane and thereby enhance the drug cellular uptake (Andresen et al., 2005; Arouri and Mouritsen, 2013; Davidsen et al., 2002, 2003; Jespersen et al., 2012). Despite the appreciable number of studies already available on sPLA2 enzymes, there is an insufficient understanding of the effect of the cancer dynamic and complex microenvironment on the hydrolysis of liposomal formulations. The aim of the present study is to explore the enzymatic activity of sPLA2 on model liposomal formulations under physiologically relevant conditions with respect to temperature, enzyme concentration, and lipid concentration. The results of the study should be helpful for the optimization of a targeting strategy involving sPLA2-assisted drug delivery by liposomes. We investigated the effect of temperature (36–41
C), lipid concentration (0.5–5 mM), and enzyme concentration (5–200 nM) on the hydrolysis of 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) liposomes using snake venom sPLA2 (CsPLA2). The major difference between CsPLA2 and human sPLA2 (HsPLA2) is in their substrate specificity, namely HsPLA2 needs a certain threshold of anionic lipids for activity (Buckland and Wilton, 2000; Leidy et al., 2006; Pedersen et al., 2010). Therefore, in order to allow for testing single-component liposomes, i.e., DPPC, CsPLA2 was used in these experiments. Based on available literature, we believe that both snake venom and human sPLA2 will react to changes in the investigated conditions in a largely comparable way. DPPC was chosen because it has a gel-to-fluid phase transition temperature around 42
C (Arouri and Mouritsen, 2011; Garidel et al., 1997), slightly higher than the human body temperature. Also, the sPLA2-

Fig. 1. A cartoon illustrating the principle of the calcein release assay as well as the enzymatic hydrolysis of DPPC.

A.H. Hansen et al. / International Journal of Pharmaceutics 491 (2015) 49–57

assisted hydrolysis of DPPC is well characterized (Arouri and Mouritsen, 2012; Hønger et al., 1996; Høyrup et al., 2001; Mouritsen et al., 2006). The cleavage of DPPC produces 1palmitoyl-2-hydroxy-sn-glycero-3-phosphocholine (lyso-PPC) and palmitic acid (PA) (see Fig. 1). The membrane perturbing effect of free fatty acids and lysolipids on various types of liposomes/vesicles has been the subject of several earlier publications (Arouri and Mouritsen, 2013; Davidsen et al., 2002; Jespersen et al., 2012; Rosholm et al., 2012). For following the enzymatic hydrolysis of DPPC, calcein-loaded ether-linked 1,2-di-O-stearyl-sn-glycero-3-phosphocholine (Di-OSPC) liposomes were co-added to the reaction mixture to serve as a reporter system. The use of calcein-loaded DPPC was not practically possible due to low calcein retention at DPPC phase transition temperature, which would increase nonspecific calcein release. In addition, the calcein–DPPC interaction could substantially alter the hydrolysis profile of DPPC liposomes. The principle of the assay is depicted in Fig. 1. The assay is well established, robust and reproducible and has been proved efficient for following the sPLA2-assisted lipid hydrolysis in several earlier studies (Arouri and Mouritsen, 2012; Davidsen et al., 2002, 2003; Leidy et al., 2006). The Di-O-SPC liposomes (the reporter system) cannot be hydrolyzed by sPLA2 and are highly stable below 53
C (Di-O-SPC phase transition) showing no passive calcein release. In addition, the permeability-enhancing effect of free fatty acids and lysolipids on Di-O-SPC liposomes is almost instantaneous and concentration-dependent (Davidsen et al., 2002). In order to understand the influence of loading the liposome with a drug on the enzymatic hydrolysis profile, we followed the enzyme-assisted luciferin release from DPPC liposomes remoteloaded with luciferin. The release of luciferin was determined via the addition of luciferase enzyme to the reaction mixture and measuring the bioluminescence signal emitted from the ATPdriven luciferase-catalyzed oxidative decarboxylation of luciferin to oxyluciferin (Nakata and Kamidate, 2001). To investigate the substrate specificity of human sPLA2 (HsPLA2), HsPLA2 extracted from tear fluid was used. The liposomes were composed of binary lipid mixtures of phosphatidylcholine (PC) and phosphatidylglycerol (PG) lipids with C14 and C16 carbon acyl chains in order to check the effect of the acyl chain length and negative surface charge density. Since C14 and C16 phospholipids are completely miscible both in the gel and fluid phases (Garidel and Blume, 2000; Garidel et al., 1997), it is not expected that the acyl-chain difference will significantly affect the homogeneity and surface morphology of the liposomes. This study allowed us to quantify the enzymatic activity of sPLA2 under tumor-relevant conditions as well as understand the effect of the liposome composition and drug loading on the hydrolysis process. 2. Materials and methods 2.1. Materials 1,2-Dipalmitoyl-sn-glycero-3-phosphocholine (DPPC), 1,2dimyristoyl-sn-glycero-3-phosphocholine (DMPC), 1,2-dipalmitoyl-sn-glycero-3-phosphoglycerol (DPPG) and 1,2-dimyristoylsn-glycero-3-phosphoglycerol (DMPG) were purchased from Corden Pharma LLC (Switzerland). 1,2-Di-O-distearoyl-sn-glycero-3-phosphocholine (Di-O-SPC) was obtained from Avanti Polar Lipids, Inc. (USA). Luciferin (sodium salt) was purchased from Regis Technologies (USA). Luciferase (from P. pyralis), sPLA2 (from N. m. mossambica), Triton X-100 (TX-100), calcein, magnesium acetate, potassium sulfate, sodium carbonate, magnesium chloride, calcium chloride, HEPES, perchloric acid, ammonium molybdate, ascorbic acid, 0.65 mM phosphate standard, chloroform, methanol,

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ATP, bovine serum albumin (BSA), dithiothreitol (DTT), and EDTA were purchased from Sigma–Aldrich (Germany). 2.2. Preparation of liposomes DPPC was dissolved in chloroform and the lipid mixtures in chloroform/methanol 2:1 (v/v) after which the organic solvent was evaporated under vacuum. The liposomes were freshly prepared by hydration of the lipid film in HEPES buffer (25 mM HEPES, 2 mM EDTA, 2.4 mM CaCl2, pH 7.5), followed by freeze-thawing of the formed lipid dispersion and extrusion 15 times through two 100 nm diameter polycarbonate filters using the Avanti Mini Extruder (Avanti Polar Lipids, Inc., USA) at a temperature higher than the phase transition temperature of the lipid or the lipid mixture, respectively. 2.3. Remote loading of luciferin into preformed DPPC liposomes DPPC liposomes (30–40 mg/mL, 2 mL) were prepared in 120 mM magnesium acetate solution (pH 6.0) by extrusion. The produced liposomes were passed through a size-exclusion column (Bio-Rad) containing Sephacryl S-200 (GE Healthcare) to exchange the bulk solution with 120 mM potassium sulfate (pH 6.0). Luciferin was remote-loaded into DPPC liposomes using the established acetate/acetic acid gradient. Untrapped luciferin was subsequently removed by size-exclusion chromatography (Sephacryl S-200). Using a FLUOstar Omega Microplate Reader (BMG LAB-TECH) in a 96-well microplate mode, the amount of encapsulated luciferin was determined by measuring the absorbance of luciferin in 0.5 M carbonate buffer (pH 11.5) at 385 nm after lysing the liposomes with Triton X-100. The phospholipid concentration was determined using a procedure adapted from Bartlett’s phosphate assay (Bartlett, 1959). 2.4. Source and quantification of human sPLA2 Human sPLA2 (HsPLA2) was obtained from tear fluid collected in our lab from 10 healthy subjects. When not in use, all samples were kept at 80
C. Tear fluid was then diluted in Tris-glycine sodium dodecyl sulfate (SDS) Sample Buffer 2 stock (to obtain 1 stock) from Invitrogen and NuPAGE Sample Reducing Agent 10 stock (to obtain 1 stock). The samples were shaken (700 rpm, 60
C) for 15 min. To visualize HsPLA2 in the collected tear samples, gel electrophoresis was performed for all tear samples using a power supply (Buch and Holm Consort EV243) to apply a voltage (120 V, 22 mA, 3 W) over a 8–16% Tris-glycerine gel 1.0 mm well (Invitrogen). The reference was SeeBlue Plus2 Prestained Standard (Invitrogen), and the running buffer was 1 Tris SDS. After 1.5 h, the gel was stained with SimplyBlue SafeStain using the microwave procedure (Invitrogen). This latter step was repeated 4 times. Milli-Q water was added to de-stain the gel over night. This procedure was adapted from (Bjerrum and Prause, 1994). A 13–15 kDa band corresponding to the HsPLA2 enzyme was observed for all tear fluid samples used in this study. A NanoDrop UV/Vis spectrophotometer (ND-1000, Thermo Scientific) was used to quantify the approximate HsPLA2 concentration in the tear fluid samples (1.1–2.63 mg mL1). 2.5. Preparation of sPLA2 stocks A concentrated stock of 0.15 mg mL1 of cobra sPLA2 (CsPLA2) was prepared by dissolving 1 mg of lyophilized powder (90% protein, specific activity 1500 units mg1 protein, http://www. sigmaaldrich.com) in 6.9 mL of HEPES buffer (25 mM HEPES, 2 mM EDTA, 2.4 mM CaCl2, pH 7.5). This concentration corresponds to

A.H. Hansen et al. / International Journal of Pharmaceutics 491 (2015) 49–57

2.8. Luciferin release experiments

TX-100

A o

39

o

Co ntr ol (41

C)

o

36

C

o

37

C

o

0.5

38

C

C

C

1.0

C

Di-O-SPC liposomes (50 nm in diameter) loaded with 50 mM calcein (at self-quenching concentration) were prepared by sonication of the lipid in a calcein solution (50 mM) for 30 min at room temperature. HEPES buffer (25 mM HEPES, 2 mM EDTA, 2.4 mM CaCl2, pH 7.5) was used to dilute the lipid sample. Untrapped calcein was removed by dialysis (3.5 kDa), and the lipid concentration was determined using the phosphate assay (Bartlett, 1959; Pokorny et al., 2002). The experiments were performed in duplicates or more using FLUOstar Omega Microplate Reader (BMG LABTECH) in a 96-well microplate (Nunc). After equilibrating the sample at the required temperature, the enzymatic reaction was initiated by the addition of sPLA2 to a mixture containing the lipid substrate and 500 nM calcein-loaded Di-O-SPC (150 ml final volume). The emitted light was recorded at 520 nm using an excitation wavelength of 490 nm. To achieve full calcein release, Triton X-100 was added to the reaction mixture (1 vol% final concentration) at the end of the experiment. The percentage of calcein release was calculated using the following equation: calcein release% = (It – IB)/(Itot – IB), where It is calcein fluorescence at time t, IB is background fluorescence and Itot is calcein fluorescence after lysing the liposomes with TX-100.

o

2.7. Calcein release experiments

o

DSC measurements were carried out using a Nano-DSC (Calorimetry Science, Provo, Utah, USA). The liposomes were freshly prepared, and a total lipid concentration of 1–2 mM was used for each experiment. The samples were degassed for 10 min before being loaded into the DSC cell, and the reference cell was filled with buffer. A scanning rate of 1
C min1 was applied.

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2.6. Differential scanning calorimetry (DSC)

calcein from Di-O-SPC liposomes causing an increase in calcein fluorescence signal due to calcein de-quenching (i.e., dilution). From the calcein release profile, it is possible to determine the percentage of calcein release (using Triton X-100 surfactant to set the 100% release), the calcein release lag-time t (time between the mixing of the reactants and the observed increase in calcein fluorescence), and the initial slope of calcein release profile. These indicators can be used to characterize the enzymatic hydrolysis process. For instance, the initial slope of calcein release and the total calcein release are dependent on the initial lipid hydrolysis rate and the total amount of lipid hydrolyzed during the course of the experiment (i.e., the amount liberated of the fatty acid and the lysolipid), respectively. Calcein release lag-time can be linked to the intrinsic lag-time of sPLA2, which is an intrinsic property of the enzyme related to its lag-burst behavior (Mouritsen et al., 2006). The former is expected to be longer because in addition to the intrinsic lag-time of sPLA2, it is also controlled by the time needed by the fatty acid and the lysolipid to leave the liposomal bilayer, diffuse in solution, and partition into Di-O-SPC liposomes as well as the time needed for subsequent calcein efflux. The effect of temperature (36–41
C) on the enzymatic hydrolysis of 80 mM DPPC liposomes (main phase transition temperature Tm around 42
C) by CsPLA2 was indirectly assessed by following the calcein release from Di-O-SPC liposomes. As shown in Fig. 2A, the calcein release curves were shifted to the left with

41

10 mM enzyme, using an estimated enzyme molecular weight of 13,120 g mol1 (Martin-Moutot and Rochat, 1979). The molar catalytic activity of HsPLA2 was estimated to be 600 units per liter tear fluid, corresponding to a HsPLA2 concentration of 1.5 mg L1 (Nevalainen et al., 1994) and a molar concentration of 110 nM (using an estimated enzyme molecular weight of 13,120 g mol1 (Martin-Moutot and Rochat, 1979).

Normalized fluorescence [RLU]

52

sPLA 2

0.0 0

10

20

30

Time [min]

B

Luciferase stocks were prepared by dissolving 1 mg luciferase in 50 mL HEPES buffer (25 mM HEPES, 2 mM EDTA, 24 mM magnesium acetate, 3 mg BSA, 2 mg DTT, pH 7.5). The luciferase solution was filtered through a 0.2 mm sterile filter. ATP stocks were prepared by dissolving ATP in HEPES buffer (25 mM HEPES, 2 mM EDTA, 2.4 mM CaCl2, pH 7.5). The reactions were performed in a 96well microplate at 37
C in HEPES buffer (25 mM HEPES, 2 mM EDTA, 2.4 mM CaCl2, pH 7.5), containing 17 nM luciferase, 12.5 mM ATP, 0.24 mM magnesium acetate, 60 mg mL1 BSA, and 40 mg mL1 DTT at pH 7.5. In each reaction well, 10 mM DPPC liposomes (100 nM lipid) remote-loaded with luciferin were incubated for different times (5, 10, 20, 48, 66, and 90 min) with 7 nM CsPLA2. The enzymatic oxidation of luciferin by luciferase was followed in the fast kinetic luminescence mode using FLUOstar Omega Microplate Reader. 3. Results The sPLA2-assisted hydrolysis of the liposomes was indirectly monitored via Di-O-SPC liposomes loaded with calcein at a selfquenching concentration (50 mM) co-added to the reaction mixture as a reporter system. The membrane-perturbing effects of the liberated lysolipids and fatty acids will eventually release

Fig. 2. (A) Normalized calcein release profiles from Di-O-SPC liposomes upon the hydrolysis of DPPC by CsPLA2 at different temperatures (36–41
C). (B) Effect of temperature on the lag-time t and the percentage of calcein release, which represents the maximum calcein release right before the addition of TX-100. Lines are inserted only as a guide to the eye. A part of the DSC thermogram of DPPC is also shown. The measurements were carried out in quintuplicates in an aqueous medium (pH 7.5) containing 200 nM CsPLA2, 80 mM DPPC (100 nm in diameter), 500 nM Di-O-SPC liposomes (50 nm in diameter), 25 mM HEPES, 2 mM EDTA, and 2.4 mM CaCl2.

A.H. Hansen et al. / International Journal of Pharmaceutics 491 (2015) 49–57

Normalized fluorescence [RLU]

increasing temperature indicating a shorter calcein release lagtime. The curves in Fig. 2A also show that the initial slope of calcein release and the maximum calcein release increased with temperature. The percentage of calcein release and the calcein release lag-time extracted from the raw curves are displayed in Fig. 2B. The lag-time was determined from the maximum of the second derivative of the profiles. The percentage of calcein release increased from 82% at 36
C to around 100% at 38 to 41
C, whereas the lag-time decreased from 16 min at 36
C to 5 min at 41
C. Our results concur with earlier studies (Leidy et al., 2006; Mouritsen et al., 2006), and the enhanced activity of sPLA2 at the lipid Tm (coexistence region) was due to the membrane defects formed during the phase transition (Mouritsen et al., 2006) as well as to the temperature-dependent increase in the enzyme activity (Menashe et al., 1980; Nair et al., 1976). Although temperature may accelerate the effect of fatty acids and lysolipids, this will increase the slope of the calcein release curves, whereas the end-state calcein release is linked to the total amount liberated of fatty acids and lysolipids. Furthermore, the order and temperature-dependence of the lag-time values in our study, measured with the calcein-release assay, agree very well with the enzyme intrinsic lag-time values and tendency reported in literature (Mouritsen et al., 2006), all of which indicate that the temperature-induced changes in the calcein release curves are truly due to changes in the activity of sPLA2. To study the effect of DPPC concentration on the enzymatic activity of CsPLA2, low lipid concentrations were utilized (i.e., 0.5– 5.0 mM). The experiments were performed at 37
C using an appreciable amount of CsPLA2 (200 nM) to avoid the enzyme concentration being the limiting factor. However, this concentration is well above physiologically relevant conditions. The measured calcein release profiles are shown in Fig. 3A, and the

TX-100

A

1.0

5.0 M 4.0 M 3.0 M 2.5 M

0.5

1.5 M sPLA 2 0.5 M

0.0 0

10

20

30

53

initial slope of calcein release and maximum calcein release values extracted from the profiles are plotted in Fig. 3B. The percentage of calcein release and the initial slope were proportional to DPPC concentration. This is understandable giving that a higher DPPC concentration, in the presence of sufficient sPLA2, will generate more PA and lyso-PPC. A complete calcein release was attained at 4–5 mM DPPC. Due to the limited Di-O-SPC concentration (500 nM), DPPC concentrations higher than 5 mM will not have any additional effect despite the larger amount of the fatty acid and the lysolipid that will be released. It should however be noted that the maximum calcein release depends on the length of the experiment, i.e., the complete calcein release would have been observed at DPPC concentrations lower than 4–5 mM if the course of the experiment had been longer. Earlier studies also showed a proportional correlation between the lipid concentration and the enzymatic hydrolysis rate, and a plateau was observed when the capacity of the enzyme was reached (Petkovic et al., 2002). Interestingly, the calcein release lag-time was independent of DPPC concentration in the studied range 0.5–5 mM (around 6 min). However, a much higher DPPC concentration of 80 mM (see Fig. 2B) eventually extended the lag-time from 6 to 12 min. This was probably due to the low number of sPLA2 molecules per liposome (i.e., high lipid-to-enzyme ratio); therefore, a longer time was needed for the creation of membrane defects essential for the activation of the enzyme. In our attempts to mimic the enzymatic reaction in a physiologically relevant setting, we reproduced the calcein release experiments using low micromolar substrate concentrations and constrained low nanomolar sPLA2 concentrations. In Fig. 4, a comparison is made to show the effect of reducing the sPLA2 concentration. Using 200 nM sPLA2 and 5 mM DPPC, the calcein release lag-time was 6 min and 95% calcein release was attained at the end of the experiment. Reducing sPLA2 concentration to 7 nM prolonged the lag-time to 14 min and only 85% calcein release was reached. Again, and using 3 mM DPPC, reducing the sPLA2 concentration from 200 nM to 5 nM protracted calcein release lag-time from 6 to 18 min and decreased calcein release from 70% to 45%. These observations indicate that a high lipid-to-enzyme ratio will not only delay the activation of the enzyme but it will also slow down the rate of formation of PA and lyso-PPC, thereby diminishing their membrane perturbing effects. To understand the influence of drug loading on CsPLA2 activity on DPPC liposomes, we carried out dye-release experiments using liposomes remotely loaded with luciferin as a model drug. After

40

Time [min] Calcein release

3 2

50 1

Calcein release [%]

B

Initial slope [min-1]

Calcein release [%]

100

150

Initial slope

5 M DPPC 6 min

100

3 M DPPC 6 min

5 M DPPC 14 min

3 M DPPC 18 min

50

0

0 0

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6

DPPC concentration [µM]

0 200

Fig. 3. (A) Normalized calcein release profiles from Di-O-SPC liposomes upon the hydrolysis of various DPPC concentrations (0.5–5 mM) by CsPLA2 at 37
C. (B) Effect of DPPC concentration on the initial and maximum calcein release. Lines are inserted only as a guide to the eye. The measurements were carried out in an aqueous medium (pH 7.5) containing 200 nM CsPLA2, 0.5–5 mM DPPC (100 nm in diameter), 500 nM Di-O-SPC liposomes (50 nm in diameter), 25 mM HEPES, 2 mM EDTA, and 2.4 mM CaCl2.

200

7

5

sPLA2 concentration [nM] Fig. 4. The effect of CsPLA2 and DPPC concentrations on the percentage of calcein release and calcein release lag-time. The measurements were carried out in an aqueous medium (pH 7.5) containing 500 nM Di-O-SPC liposomes (50 nm in diameter), 25 mM HEPES, 2 mM EDTA, and 2.4 mM CaCl2.

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the incubation of the remote-loaded DPPC liposomes with CsPLA2 (7 nM) for varying times, luciferase was added to the reaction mixture and the luminescence resulting from luciferase–luciferin raction was measured in order to follow CsPLA2-triggered luciferin release. The luminescent signal was only measured for a short time (5 s) at 37
C to avoid possible proteolysis of luciferase enzyme over time (Ataei et al., 2009). Unlike unloaded DPPC liposomes, which were quickly degraded by sPLA2 (see Figs. 2 A and 3 A and Refs. (Arouri and Mouritsen, 2012; Petkovic et al., 2002)), Fig. 5 shows that the hydrolysis of luciferin-loaded DPPC liposomes was much slower and occurred in two steps. This observation is likely due to the perturbing effects of luciferin on DPPC bilayer and domains formation (unpublished DSC results). The phenomenon of a twostep hydrolysis was observed before with lipid mixtures (Høyrup et al., 2001) and lipid prodrugs (Arouri and Mouritsen, 2012), and was correlated with the continuously changing properties of the membrane during the hydrolysis, rendering the membrane, at some point, less appealing to sPLA2. The reaction to which no CsPLA2 was added showed a background signal from free luciferin, possibly because of nonspecific luciferin release. In all experiments discussed above, sPLA2 from snake venom (CsPLA2) was used to characterize the enzymatic action on lipid membranes. In contrast to CsPLA2, human sPLA2 (HsPLA2) is more selective and necessitates the presence of considerable amounts of anionic lipids like phosphatidylglycerol (PG) and phosphatidylserine (PS) (Buckland and Wilton, 2000; Leidy et al., 2006). Therefore, in order to fine-tune the lipid formulation, we investigated the

Fig. 5. (A) Luminescence and (B) maximum luminescence generated from luciferase–luciferin reaction after the incubation of luciferin-loaded DPPC liposomes with CsPLA2 for different times (5–90 min). Lines are inserted only as a guide to the eye. The reactions were carried out at 37
C in an aqueous medium (pH 7.5) containing 10 mM DPPC liposomes (100 nm in diameter), 7 nM CsPLA2, 17 nM luciferase, 25 mM HEPES, 2 mM EDTA, 12.5 mM ATP, 0.24 mM magnesium acetate (co-factor), 60 mg mL1 BSA, 40 mg mL1, and DTT.

action of HsPLA2 from tear fluid on PC/PG mixed membranes with C14 and C16 acyl chains. In general, as shown in Fig. 6, the percentage of calcein release increased with increasing the PG molar fraction in the mixed systems (from 0.26 to 0.34 PG). This effect is probably due to a higher accumulation of the cationic HsPLA2 on increasingly more anionic surfaces (Lambeau and Gelb, 2008). Interestingly, the DPPG-containing mixed systems – DPPC/DPPG (Tm is around 40
C) and DMPC/DPPG (Tm is around 23
C) (Garidel et al., 1997) – were associated with the highest calcein release (Fig. 6) as well as with the highest initial slope and the shortest calcein release lag-time (data not shown). DMPC/DMPG (Tm is around 23
C (Garidel et al., 1997)) mixed systems were not efficient substrates for HsPLA2, and DPPC/DMPG (Tm is around 34
C (Garidel et al., 1997)) mixed systems showed an intermediate effect. At 37
C, DPPC/DPPG mixed systems are in the gel–fluid coexistence region, whereas the other mixed systems are in the fluid phase. This might explain the enhanced hydrolysis of the DPPC/DPPG liposomes, namely due to the membrane defects formed during the phase transition (Mouritsen et al., 2006). The higher calcein release associated with the C16 lipids DPPC and DPPG could be either because they were more favorable enzyme substrates or because their hydrolytic products were more efficient permeability enhancers; yet, further studies are needed to explain this phenomenon. 4. Discussion The use of sPLA2-susceptible liposomes for the selective delivery of anticancer drugs is a promising approach furnished by its applicability in vivo (Andresen et al., 2005). However, despite the large body of data available, not much is known about the hydrolysis process in the cancer vicinity which calls for additional studies pertinent to physiological conditions. The intrinsic lag-time of sPLA2 is shortest at the main phase transition temperature of lipid bilayers, and in mixed lipid membranes the minimum lag-time occurs over a rather broad temperature range (Mouritsen et al., 2006). It appears from our results as well as from earlier reports (Davidsen et al., 2003; Leidy et al., 2006; Mouritsen et al., 2006) that going away in temperature from the liposome main phase transition temperature (Tm) tends to delay the activation of sPLA2 (i.e., prolong the lag-time), without significantly affecting the sigmoid-like increase in lipid hydrolysis rate. Therefore, it might actually be beneficial to design a liposomal drug delivery system that has a Tm somewhat higher than the body temperature in order to delay the hydrolysis until after the liposomes have accumulated in the target cancer. Since the permeability and instability of liposomes are largest at Tm (Landon et al., 2011), using liposomes with Tm higher than 37
C will also help to improve the stability and drug retention of the liposomal formulation in the body during the biodistribution process. In addition to the reaction conditions and membrane properties, we also found the lag-time to be dependent on the lipid-toenzyme ratio. As illustrated in Fig. 7, there appears to be a “threshold” lipid-to-enzyme molar ratio of 40–50 (DPPC/CsPLA2), above which the lag-time starts to increase. A low number of enzyme molecules per liposome means more time is required for the creation of membrane defects and the aggregation of sPLA2 molecules, both of which are needed for the activation of the enzyme (Code et al., 2008; Mouritsen et al., 2006). Increasing the lipid-to-enzyme ratio above the threshold will not only extend the lag-time, but it will also reduce the overall hydrolysis rate of the lipid (compare Figs. 2 A and 3 A). It is anticipated that the initial serum lipid concentration after the parenteral administration of a liposomal medicinal product will be on the low end of the micromolar range, and therefore the lipid-to-enzyme ratio will be relatively high. This will ensure that

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Fig. 6. Percentage of calcein release from Di-O-SPC liposomes upon the enzymatic hydrolysis of different liposomal formulations by human sPLA2 (HsPLA2) from tear fluid. The measurements were carried out at 37
C in an aqueous medium (pH 7.5) containing 2 nM HsPLA2, 20 mM lipid (100 nm in diameter), 500 nM Di-O-SPC liposomes (50 nm in diameter), 25 mM HEPES, 2 mM EDTA, and 2.4 mM CaCl2.

Lag-time [min]

20 15 10 5 0 1

10

10 0

100 0

DPPC to sPLA2 molar ratio Fig. 7. Correlation between DPPC to CsPLA2 molar ratio and lag-time of calcein release at 37
C. The line is inserted only as a guide to the eye.

there will be sufficient time for the biodistribution of the liposomes and their accumulation in the target cancer before any significant liposome hydrolysis. Because of the higher sPLA2 level in the cancer tissue compared with serum (on average 6–8 times) together with the initially low lipid concentration during the accumulation phase, the lipid-to-enzyme ratio at the cancer site will be substantially low enough to guarantee a faster lipid hydrolysis and consequently a more efficient drug release. Notwithstanding our comment, the mode of hydrolysis of the liposomes will be greatly affected by the proteins and other components present in the blood and interstitial fluid.

So far, almost all studies on the action of sPLA2 have been conducted using simple and “empty” model lipid bilayers. It is clear from our results that the payload-induced alterations in the substrate lipid bilayer can affect the activity of sPLA2. However, we believe that the drug release profile will strongly depend on the mode of drug-membrane interaction and therefore our results cannot be generalized. Furthermore, our results show that the drug release takes place gradually during the hydrolysis process, where the membrane defects or openings produced during the hydrolysis tend to re-seal quickly due to the fast diffusional rearrangement of the membrane components (Davidsen et al., 2002; Rosholm et al., 2012). Somewhat surprisingly, we found that the activity of HsPLA2 is not only dependent on the surface charge of the lipid bilayer but also on the acyl-chain length of the phospholipid, in such a way that the overall calcein release was much reduced when a C16 acyl chain phospholipid was replaced with a C14 acyl chain phospholipid. The reason for this effect is unknown and requires further investigation. However, in the context of formulating a drug delivery system, DPPC/DPPG liposomes showed superior properties over the other tested formulations. The differences between venom and human sPLA2 in terms of activity and substrate specificity have been the subject of several earlier studies (Buckland and Wilton, 2000; Leidy et al., 2006; Pedersen et al., 2010). It appears that the major difference between venom and human sPLA2 refers in the substrate specificity, whereas their global behavior and activity are essentially comparable. Venom sPLA2 is highly active on uncharged PC lipids, whereas the activity of human sPLA2 on PC lipids is very modest

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(Leidy et al., 2006). The incorporation of anionic lipids like PG was found to enhance the activity of both types of sPLA2, but the effect was more pronounced for the human enzyme (Leidy et al., 2006). A PG threshold of 30 mol% for the activation of human sPLA2 was proposed, above which the activity of the enzyme was found to be proportional to the membrane negative surface charge density (Leidy et al., 2006). The presence of anionic lipids facilitates the accumulation of the highly cationic human sPLA2, which has several arginine and lysine amino acids scattered around the surface of the enzyme (Berg et al., 2001). It has been speculated that human sPLA2, which is able to make contact to 20–40 phospholipid molecules at the interface (Ramirez and Jain, 1991), may actually induce phase segregation separating anionic lipids from zwitterionic ones (Gelb et al., 1994; Lambeau and Gelb, 2008). The local heterogeneities created at the membrane interface were found to be essential for the activity of human sPLA2 (Mouritsen et al., 2006). Even though the affinity of human sPLA2 is higher for anionic lipids, the catalytic efficiency of the enzyme was found to be independent of the net charge of the lipid molecule (Singer et al., 2002). Unlike for human sPLA2, the electrostatics was found to be of minor importance for venom sPLA2, where the enhanced activity of venom sPLA2 in the presence of anionic lipids has been linked to the anionic-lipid-induced membrane perturbations and lower membrane packing density, which would facilitate non-polar interactions with the membrane (Buckland and Wilton, 2000). Our results concur well with the published reports, which strongly support our assumption that the knowledge obtained in this study with snake venom sPLA2 can be translated with a high degree of confidence to human sPLA2, qualitatively and maybe also semiquantitatively. The current paper will aid filling the gap between our fundamental understanding of the molecular mechanism of sPLA2 and the lipidology-based bio-relevant knowledge needed for the rational design of sPLA2-susceptible liposomes as well as for the elucidation of their fate in vivo. In order to further support the applicability of this platform, the focus of future studies should be on establishing a closer relation between the activity of sPLA2 on simple model systems and the real activity of the enzyme under in vivo biological conditions (Arouri et al., 2015). Acknowledgments This work was supported by The Lundbeck Foundation Center of Excellence NanoCAN (Nanomedicine Research Center for Cancer Stem Cell Targeting Therapeutics), the Danish Council for Independent Research-Technology and Production Sciences (DFF-FTP), and a scholarship from Novo Nordisk Foundation to AHH. References Abe, T., Sakamoto, K., Kamohara, H., Hirano, Y., Kuwahara, N., Ogawa, M., 1997. Group II phospholipase A2 is increased in peritoneal and pleural effusions in patients with various types of cancer. Int. J. Cancer 74, 245–250. Andresen, T.L., Jensen, S.S., Jørgensen, K., 2005. Advanced strategies in liposomal cancer therapy: problems and prospects of active and tumor specific drug release. Prog. Lipid Res. 44, 68–97. Arouri, A., Hansen, A.H., Rasmussen, T.E., Mouritsen, O.G., 2013. Lipases, liposomes and lipid-prodrugs. Curr. Opin. Colloid Interface Sci. 18, 419–431. Arouri, A., Mouritsen, O.G., 2011. Anticancer double lipid prodrugs: liposomal preparation and characterization. J. Liposome Res. 21, 296–305. Arouri, A., Mouritsen, O.G., 2012. Phospholipase A2-susceptible liposomes of anticancer double lipid-prodrugs. Eur. J. Pharm. Sci. 45, 408–420. Arouri, A., Mouritsen, O.G., 2013. Membrane-perturbing effect of fatty acids and lysolipids. Prog. Lipid Res. 52, 130–140. Arouri, A., Trojnar, J., Schmidt, S., Hansen, A.H., Mollenhauer, J., Mouritsen, O.G., 2015. Development of a cell-based bioassay for phospholipase A2-triggered liposomal drug release. PLoS One 10, e0125508.

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