Influence of Additives on the Properties of Nanodroplets Formed in

Jul 5, 2013 - and Lynne S. Taylor*. ,†. †. Department of Industrial and Physical Pharmacy, College of Pharmacy, Purdue University, West Lafayette,...
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Influence of Additives on the Properties of Nanodroplets Formed in Highly Supersaturated Aqueous Solutions of Ritonavir Grace A. Ilevbare,† Haoyu Liu,‡ Junia Pereira,‡ Kevin J. Edgar,‡ and Lynne S. Taylor*,† †

Department of Industrial and Physical Pharmacy, College of Pharmacy, Purdue University, West Lafayette, Indiana Department of Sustainable Biomaterials, College of Natural Resources and Environment, Virginia Tech, Blacksburg, Virginia

‡

S Supporting Information *

ABSTRACT: The formation of colloidal drug aggregates of lipophilic drugs is thought to be of relevance for the oral delivery of poorly water-soluble drugs. In this study, the underlying basis for colloid formation from amorphous solid dispersions and the impact of additives on colloidal stability were evaluated. A relationship was found between the concentration at which colloidal droplets formed upon dissolution of an amorphous solid dispersion and the liquid− liquid phase separation (LLPS) transition concentration, whereby the latter is related to the theoretical amorphous “solubility” value. The composition of the dispersed phase in ritonavir−polymer−water solutions was confirmed to be a noncrystalline, ritonavir-rich droplet phase. Additives were found to impact the size, stability, and crystallization behavior of the colloidal phase. In general, charged additives reduced the kinetics of droplet coalescence, but had a variable effect on crystallization kinetics, either promoting or inhibiting crystallization. Through proper selection of formulation components, it thus appears possible to promote the formation of ∼250−350 nm colloidal droplets of ritonavir following dissolution of an amorphous solid dispersion, and to inhibit coalescence and crystallization from these two-phase supersaturated solutions. KEYWORDS: ritonavir, amorphous solid dispersions, nanodroplet formation, colloid formation, liquid−liquid phase separation, supersaturation, cellulose ω-carboxyalkanoates, 6-carboxypullulan ethers

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INTRODUCTION Formulation of poorly water-soluble drugs is a challenging problem encountered during the drug development process.1,2 It is estimated that approximately 40% of active substances currently in development fail because of low efficacy, often as a result of poor bioavailability, which can arise from ineffective intestinal absorption and/or undesirable metabolic activity.3−6 Colloidal dispersions are increasingly being used in drug delivery. Nanosuspensions of drugssubmicrometer colloidal dispersions of nearly pure drug particleshave been used to enhance the delivery of poorly water-soluble compounds for both oral and parenteral delivery.7−11 Nanodispersions can be either crystalline or noncrystalline and can be produced using several techniques.11−15 The top down wet milling approach tends to generate crystalline particles. In contrast, the bottom up solvent-shifting technique (also known as the cosolvent or antisolvent technique), a method which involves adding a nonsolvent such as water to an organic solution of the drug compound, can result in either crystalline or noncrystalline colloidal dispersions.13,14 It has also been demonstrated that supersaturating bioactive delivery systems, consisting of amorphous solid dispersions (ASD) of the poorly water-soluble compound with a water-soluble polymer, are capable of generating colloidal dispersions when dissolved in aqueous solution.16−22 Little is known about the mechanism by which colloidal species are produced from amorphous solid © 2013 American Chemical Society

dispersions although it has been suggested that they are important to the bioavailability enhancement often seen with amorphous solid dispersions.18 Furthermore, Frenkel et al. proposed that the formation of colloidal aggregates (especially aggregates with hydrodynamic radii less than 100 nm) may be responsible for the enhanced bioavailability of non-nucleoside reverse transcriptase inhibitors (NNRTIs) by promoting their absorption by particle-recognizing microvilli cells in Peyer’s patches of mucosa-associated lymphoid tissue.23 In order to take advantage of the properties of noncrystalline colloidal dispersions, the submicrometer droplets must be stabilized. The main problems to be overcome are coalescence of the aggregates to form larger droplets and crystallization; colloidal droplets are generated through the formation of highly supersaturated solutions, hence crystallization is likely to occur. Additives, such as surfactants, may offer a means to prevent coalescence of aggregates; however, their effect on the rate of drug crystallization (nucleation and crystal growth) has not been widely explored.14,15,24 Polymers have also been used to stabilize colloidal dispersions14−17 and can also serve as antinucleating agents and inhibit crystal growth.25,26 Received: Revised: Accepted: Published: 3392

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vinyl acetate) (K 28), poly(acrylic acid) (PAA), pyrene, sodium dodecyl sulfate (SDS), and Tween 80 were obtained from Sigma-Aldrich (St. Louis, MO). Hydroxypropyl methyl cellulose (HPMC) 606 grade and hydroxypropyl methyl cellulose acetate succinate (HPMCAS) AS-MF grade were obtained from Shin-Etsu Chemical Co., Ltd. (Tokyo, Japan). Cationically modified hydroxyethyl cellulose, Polyquaterium-10 (PQ-10, Figure 2a), was obtained from Making-Cosmetics, Inc. (Renton, WA), hydrophobically modified hydroxylethyl cellulose (HM-HEC) was obtained from Ashland Inc. (Covington, KY), and hydrophobically modified hydroxypropyl methylcellulose, HM-HPMC (hydroxylpropyl methyl cellulose stearoxy ether 90L), was obtained from Daido Chemical Co. (Osaka, Japan); chemical structures are shown in Figure 2. The syntheses of the cellulose ω-carboxyalkanoates (Figure 3) and 6-carboxypullulan ethers are described in refs 27 and 28, respectively. Table 1 comprises a list of additives used in this study and their abbreviations, while some of the properties of the synthesized polymers are presented in Table 2. The additives were used as received. Methods. Preparation of Amorphous Solid Dispersions (ASDs) of Ritonavir. Amorphous solid dispersions of ritonavir and additive(s) were prepared by the solvent evaporation method. For the ritonavir/PVP solid dispersion, a 10:90 (wt %) ratio of ritonavir to PVP was used. Amorphous solid dispersions containing the other additivesHPMC, HPMCAS, CAAdP 0.33, CAAdP 0.85, and Tween 80were prepared using a 10:80:10 wt % ratio of ritonavir to PVP to one of the aforementioned additives. These additives (which mostly comprise moderately hydrophobic polymers) were combined with PVP, a hydrophilic polymer, to ensure the rapid release of ritonavir and other components from the solid matrix. Mixtures containing combinations of PVP, CAAdP 0.33, CAAdP 0.85,

In this study, we assessed the properties of colloidal aggregates, generated in the presence of several polymers and surfactants, including a number of recently synthesized cellulose derivatives with greater chemical diversity than those that are commercially available.25,26 Ritonavir, a poorly water-soluble compound which has been shown to form colloidal dispersions,29 was selected as the model compound. The specific goals of this study were to (1) investigate the mechanism of aggregate formation as well as aggregate structure; (2) examine the impact of additives on the initial size and size stability of the aggregates and; (3) probe the crystallization behavior of solutions containing aggregates.

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EXPERIMENTAL SECTION Materials. Ritonavir (Figure 1) was purchased from Attix Corporation, Toronto, Ontario, Canada. Methanol was

Figure 1. Molecular structure of ritonavir.

purchased from Macron Chemicals (Phillipsburg, NJ). Poly(vinyl pyrrolidone) K29/32 (PVP), poly(vinyl pyrrolidone

Figure 2. Molecular structure of (a) polyquaterium 10 (PQ-10), (b) the recently synthesized polysaccharide derivative, butyl pullulan-6-carboxylate (butyl-6-CO2HPull), (c) hydrophobically modified hydroxylethyl cellulose (HM-HEC) and (d) hydrophobically modified hydroxypropyl methylcellulose (HM-HPMC). PQ10, HM-HEC, and HM-HPMC are not regioselectively substituted; oxidation of pullulan at C-6 is regioselective, but the position of the butyl ethers (between O-2, -3, -4 positions) is not. 3393

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constant at 50 mg. Solvent was removed in a rotary evaporator (Brinkman Instruments, Westbury, NY), with the water bath at 65 °C. The sample was then placed under vacuum for 24 h to remove residual solvent. The method used in determining the equilibrium crystal solubility, as well as the experimental and theoretical amorphous “solubilities” of ritonavir, was previously reported in ref 29. Determination of the Liquid−Liquid Phase Separation Concentration in the Presence of Additives. The liquid− liquid phase separation (LLPS) concentration of ritonavir in buffer has been measured previously.29 The LLPS transition concentration of ritonavir in the presence of 180 μg/mL of polymer solution was determined and compared to the value in the absence of the polymer. Solubilized ritonavir (4 mg/mL) in methanol was titrated into a solution maintained at 37 °C, and buffered to pH 6.8, which contained the additive of interest, using a syringe pump (Harvard Apparatus, Holliston, MA). The solution was stirred at a speed of 300 rpm. The liquid−liquid phase (LLPS) transition concentration (also referred to as the CL−L boundary)29 was determined as the concentration at which an increase in intensity of light scattered from the drug solutions was observed. Light scattering was detected by monitoring the extinction at nonabsorbing wavelengths ranging from 280 to 450 nm using an SI Photonics UV/vis spectrometer (Tuscon, Arizona), fiber optically coupled with a dip probe (path length 10 mm), while solubilized drug was continuously added to the solution using the syringe pump. Evolution of turbidity was characterized by an increase in the extinction between 280 and 450 nm. Characterization of Ritonavir Colloidal Phases. Simultaneous UV/Vis Spectroscopy and Dynamic Light Scattering (DLS). The properties of the liquid-like dispersed phase (colloidal phase) and the occurrence of various phase transformations (formation of colloidal phase, dissolution of colloidal phase, and crystallization) were studied by monitoring mean count rate and dispersed phase size using DLS, combined with simultaneous extinction (scattered light intensity) and absorption measurements using the above-mentioned UV/vis spectrometer. DLS experiments were performed using a NanoZetasizer (Nano-ZS) from Malvern Instruments (Westborough, MA) and its software, dispersion technology software (DTS). A backscatter detector was used, and the scattered light was detected at an angle of 173°. This optical configuration maximizes the detection of scattered light while maintaining signal quality. A quartz flow-through cuvette (Malvern Instruments (Westborough, MA)), coupled with a MasterFlex Easy-Load peristaltic pump (Cole Parmer, Vernon Hills, IL), was used for continuous sampling. The flow rate used was 1 mL/min. Measurements were made approximately every 5 min. The Z-average size of the particles was reported. The following experiments, described below, were performed using this set up. Formation of Dispersed Phase from Dissolution of Amorphous Solid Dispersions. An amorphous solid dispersion of ritonavir/PVP (10:90 wt %) dissolved in 100 mM sodium phosphate buffer solution (pH 6.8 and 37 °C) was used to generate a supersaturated solution of ritonavir. In order to acquire baseline turbidity measurements, data collection was started before the solid dispersion was added to the buffer solution (i.e., there was 0 μg/mL of ritonavir in solution for the first 20 min of the experiment). The solid dispersion was added to 25 mL buffer solution in small increments of ∼3.0 mg up to a total ritonavir concentration of approximately 20 μg/mL.

Figure 3. Molecular structure of the novel synthesized cellulose derivatives and the substituent groups. These cellulose derivatives are not regioselectively substituted. The abbreviations are presented in Table 1.

Table 1. Additives and Abbreviations Used in This Study polymer

abbreviation

poly(vinylpyrrolidone) (K 29/32) poly(vinylpyrrolidone vinyl acetate) (K 28) poly(acrylic acid) polyquaterium 10 sodium dodecyl sulfate Tween 80 hydroxypropyl methyl cellulose (606 grade) hydroxypropyl methyl cellulose acetate succinate (ASMF) hydrophobically modified hydroxylethyl cellulose hydrophobically modified hydroxypropyl methyl cellulose butyl pullulan-6-carboxylate cellulose acetate propionate 504-0.2 adipate 0.33 cellulose acetate butyrate 553-0.4 adipate 0.81 cellulose acetate propionate 504-0.2 adipate 0.85 cellulose acetate 320S suberate 0.90

PVP PVPVA PAA PQ-10 SDS Tween 80 HPMC HPMCAS HM-HEC HM-HPMC butyl-6-CO2HPull CAAdP 0.33 CAAdB 0.81 CAAdP 0.85 CA Sub 0.90

Table 2. Degree of Substitution (DS) of the Cellulose-Based Synthesized Polymers polymer

DS (CO2H)a

CAAdP 0.33 CAAdP 0.85 CAAdB 0.81 CA Sub 0.90

0.33 0.85 0.81 0.90

DS (other)b Ac, Ac, Ac, Ac,

0.04; Pr, 2.09 0.04; Pr, 2.09 0.14; Bu, 1.99 1.82

DS (total) 2.46 2.98 2.84 2.72

a

DS (CO2H): DS of the carboxylic-bearing substituent. bAbbreviations: acetate (Ac), propionate (Pr), butyrate (Bu).

and Tween 80 were dissolved in 100% methanol, while mixtures containing HPMC and HPMCAS were dissolved in a solution of dichloromethane/methanol (1:1 wt % ratio). A total of 500 mg of ritonavir and polymer(s) was dissolved in the appropriate solvent(s). For example, the ritonavir/PVP amorphous solid dispersion was prepared by dissolving 50 mg of ritonavir and 450 mg of PVP in 100 mL of 100% methanol, while ritonavir/PVP/HPMC amorphous solid dispersion was prepared by dissolving 50 mg of ritonavir, 400 mg of PVP, and 50 mg of HPMC in 100 mL of dicholoromethane/methanol (1:1 wt % ratio); for all the solid dispersions prepared, the amount of ritonavir was held 3394

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After ∼60 min, the test solution was diluted to make a final ritonavir solution concentration of 10 μg/mL. Additional amorphous solid dispersions of ritonavir were also evaluated by dissolving them in aqueous solution (100 mM sodium phosphate buffer (pH 6.8), 100 mM NaCl or distilled water) at 37 °C. An excess amount of solid dispersion, ∼20 mg, was added to 50 mL aqueous solution. The solution was stirred at a speed of 300 rpm, using a stir bar and digital stir plate (Corning, PC 420D, Corning Inc., NY). Data collection (UV and DLS) commenced after the dissolution of solid dispersion. UV absorption and extinction data were used to further monitor solution properties and phase transformations as described previously.29 Formation of Dispersed Phase from Solvent-Shifting Method and Induction time Measurements. Supersaturated solutions were generated by adding a small volume of predissolved drug in methanol to buffer. Solubilized drug (4 mg/mL) in methanol was titrated into 100 mM sodium phosphate buffer solution (50 mL) equilibrated at 37 °C, at pH 6.8, using a syringe pump (Harvard Apparatus, Holliston, MA). Agitation of the solution at 300 rpm was achieved by use of a magnetic stirrer (Corning PC-420D, Fisher Scientific, Pittsburgh, PA). DLS and induction time experiments were performed in the absence and presence of predissolved additives. With the exception of butyl-6-CO2HPull (2 μg/ mL), an additive concentration of 5 μg/mL was used. All experiments were performed in triplicate. An initial ritonavir solution concentration of 20 μg/mL was used, and this concentration is just above the theoretical amorphous “solubility” (CL−L boundary) of ritonavir. Data collection (UV and DLS) commenced immediately after the addition of the drug solution to the test medium. The onset of crystal formation was measured using the method previously described in ref 26. Crystal formation was characterized by measuring the induction time. The onset of crystal formation at an equilibrium temperature (37 °C) was determined from the increase in intensity of light scattered (extinction) from the drug solutions. Light scattering was detected by monitoring the extinction at 280 nm using an SI Photonics UV/vis spectrometer (Tuscon, Arizona), fiber optically coupled with a dip probe (path length 10 mm); ritonavir has no absorbance at this wavelength. Wavelength scans (200−450 nm) were performed at 30 s time intervals. Fluorescence Spectroscopy. A fluorescence probe technique was used to further evaluate the nature of the colloidal phase generated during the dissolution of an amorphous solid dispersion of ritonavir/PVP (10:90 wt %). Fluorescence spectra were obtained using a Shimadzu spectrofluorophotometer RF5301PC (Shimadzu Scientific Instruments, Inc., Columbia, MD). Pyrene, which exhibits different fluorescence characteristics depending upon the properties of the solubilizing medium,30,31 was used as a fluorescence probe. A known amount of pyrene solubilized in DMSO was added to distilled water to generate a pyrene concentration of 0.5 μg/mL. Thirty milligrams of ritonavir/PVP solid dispersion was dissolved in 50 mL of distilled water containing 0.5 μg/mL of pyrene. Emission spectra of pyrene were obtained by exciting the samples at 334 nm. Spectra were analyzed using GRAMS/AI V.7.02 software (Thermal Fisher Scientific, Inc., Waltham, MA). The time point of crystallization was characterized by an increase in the ratio of intensity of the first (I1 at 373 nm) and the third peaks (I3 at 383 nm) of the pyrene emission spectra;

I1/I3 is a sensitive parameter characterizing the polarity of the probe’s environment.30 Scanning Electron Microscopy. Scanning electron microscopy was used to characterize the colloidal dispersion generated after the dissolution of amorphous solid dispersions of ritonavir/PVP (10:90 wt %) and ritonavir/PVP/Tween 80 (10:80:10 wt %) in distilled water at 37 °C. Approximately 20 μL samples of the solution were withdrawn at 30 min intervals for 2 h, and each sample was placed on a glass slide. The glass slides were allowed to dry overnight in a vacuum oven at room temperature for ∼12 h. The glass slides were mounted using double sticky copper tape and sputter-coated with Pt for 60 s prior to imaging. Subsequently, samples were imaged with a FEI NOVA nanoSEM field emission SEM using the EverhartThornley (ET) detector and through-the-lens detector (TLD). Parameters were 5 kV accelerating voltage, ∼4−5 mm working distance, beam spot size of 3, and 30 μm aperture. Magnifications were 100−10000×. The SEM images were analyzed using ImageJ, processing and analysis in Java (National Institutes of Health). Differential Scanning Calorimetry. The precipitated (colloidal) phase generated during the dissolution of an amorphous solid dispersion of ritonavir/PVP (10:90 wt %) in distilled water at 37 °C was also characterized using a differential scanning calorimeter (DSC). The supernatant was separated from the precipitated phase by ultracentrifugation for 15 min at 40,000 rpm (equivalent of 274356g) in an Optima L100 XP ultracentrifuge equipped with Swinging-Bucket Rotor SW 41 Ti (Beckman Coulter, Inc., Brea, CA). Subsequently, the precipitate was analyzed using DSC. Thermal transitions were measured using a TA Q2000 DSC (TA Instruments, New Castle, DE) attached to a refrigerated cooling accessory (RCS) (TA Instruments, New Castle, DE). Both the DSC and RCS were purged with nitrogen gas. Tin was used for temperature calibration, while cell constant and enthalpy calibrations were performed using indium. Baseline calibration was performed by heating the empty cell from −50 to 300 °C at 20 °C/min. The reference and sample pans were identical. The precipitate was sealed in an aluminum pan with a pinhole in the lid. The thermogram was obtained by first cooling the sample, and then heating the sample at a rate of 20 °C/min in order to determine the glass transition temperature. The temperature range used was −40 to 60 °C. Thermal transitions were viewed and analyzed using the analysis software Universal Analysis 2000 for Windows 2000/XP provided with the instrument.

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RESULTS Characterization of Ritonavir Colloidal Phase. To characterize the properties of the dispersed phase generated during dissolution of an amorphous solid dispersion of ritonavir, a combination of analytical techniques were used: UV/vis spectroscopy, dynamic light scattering (DLS), fluorescence spectroscopy, differential scanning calorimetry (DSC), and scanning electron microscopy (SEM). Figure 4 shows a plot of mean count rate [in kilocounts per seconds (kcps)] and extinction (at a wavelength of 280 nm, UV measurement) as a function of time (in minutes). For a homogeneous solution with no scattering centers, the UV extinction at a nonabsorbing wavelength should be minimal and the count rate measured by DLS should be low. When small portions of a 10:90 wt % ritonavir/PVP ASD were added sequentially to a solution, extinction was minimal until 3395

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phase that contains a few percent of water (water is known to depress the glass transition event34). To further investigate the structure of the dispersed phase formed upon dissolution of the amorphous solid dispersion, fluorescence and SEM experiments were performed. A fluorescence probe was used to investigate the local environments in the solution. If the colloidal phase formed is noncrystalline in nature as suggested by the results discussed above (Figure 4), then the hydrophobic probe molecule would be expected to partition into this phase and consequently undergo a change in emission characteristics relative to the emission spectrum in water. For pyrene, the hydrophobic probe molecule used in this study, it has been shown that the I1/I3 emission peak ratio is sensitive to the polarity of the probe environment.30 Figure 5 shows the dependence of the pyrene Figure 4. Plot of mean count rate and extinction with time during dilution experiment. An amorphous solid dispersion of ritonavir/PVP 10:90 wt % ratio was used. At concentrations below the LLPS concentration (18.3 ± 0.46 μg/mL), meaningful particle size data (DLS) could not be obtained since formation of the dispersed phase starts at ∼18.3 μg/mL; initial droplet size at 18.3 μg/mL was ∼350 nm.

sufficient ASD had been added to yield a ritonavir concentration of ∼18 μg/mL; the resultant solution was also then observed to be cloudy. At this concentration, the mean count rate also became significant and the median diameter of the dispersed phase was around 350 nm. At concentrations lower than ∼18 μg/mL, meaningful size data (using DLS) could not be obtained since the count rate was very low. Both the extinction and DLS data are consistent with the formation of a second phase which scatters light. Interestingly, a threshold concentration of ritonavir needs to be generated by dissolving a sufficient amount of the ASD before the second phase is produced; this concentration is approximately the same as the liquid−liquid phase separation (LLPS) concentration (18.3 μg/ mL) and the experimental amorphous “solubility” (19.8 μg/ mL) of ritonavir in the same medium.29 The colloidal phase subsequently disappeared when the mixture was diluted below the LLPS concentration (Figure 4). At half the LLPS concentration (10 μg/mL), the mean count and extinction returned to the background signal, comparable to when the ritonavir concentration was 0 μg/mL, showing that the second phase will not persist if diluted below the LLPS concentration. This type of behavior has been observed previously for colloidal aggregates.32,33 These results strongly support the formation of drug-rich liquid-like droplets that form when sufficient ASD has been dissolved so that the LLPS concentration is exceeded. If the second phase consisted of crystalline particles, they would not dissolve at a solution concentration (10 μg/mL) well above the equilibrium solubility of ritonavir (1.3 μg/mL at pH 6.8, 37 °C). The DSC thermogram of the isolated dispersed phase (Figure S1 in the Supporting Information) showed a thermal event characteristic of a glass transition (Tg) at around 1.05 °C, indicating that it is a supercooled liquid at the experimental temperature. The melting point of the form II polymorph of ritonavir is 122.7 °C,26 therefore at this much lower temperature, the liquid form of ritonavir is metastable with respect to this crystalline form. The Tg of pure amorphous ritonavir is 50 °C;21,25 the lower Tg associated with the dispersed phase generated by dissolving the amorphous solid dispersion is consistent with the formation of a ritonavir-rich

Figure 5. Ratio of intensity of the first (I1 at 373 nm) and the third peaks (I3 at 383 nm) of pyrene emission spectra as a function of concentration for ritonavir at an equilibrium temperature of 37 °C. A significant decrease in I1/I3 occurred at the concentration corresponding to the LLPS concentration. The increase in I1/I3 at ∼120 min corresponds to crystallization. An amorphous solid dispersion of ritonavir/PVP 10:90 wt % ratio was used.

I1/I3 emission peak ratio on time following the addition of sufficient ritonavir/PVP (10:90 wt %) ASD to 100 mM sodium phosphate buffer to generate an initial concentration of ∼20 μg/mL ritonavir. I1/I3 decreased sharply after the amorphous solid dispersion dissolved completely and remained constant for up to ∼100 min. The decrease of the I1/I3 values indicates that pyrene is in a more hydrophobic environment than water (ritonavir has a Log P of 5.9835 and thus is much more hydrophobic than water), which is consistent with the formation of a noncrystalline ritonavir colloidal phase into which the probe partitions. If the colloidal phase were crystalline, minimal change in probe emission spectrum would be expected since pyrene would not be able to penetrate the crystal lattice. After ∼100 min, the I1/I3 emission peak ratio increased significantly, consistent with the formation of crystalline particles from which pyrene is excluded, hence the pyrene encounters a more hydrophilic environment again. This supposition is supported by our previous observations that, under similar experimental conditions (initial concentration of ∼20 μg/mL in buffer at 37 °C), ritonavir was found to have a crystal-nucleation induction time of ∼120 min.26 SEM experiments also supported the noncrystalline nature of the colloidal phase. The test solution (dissolved amorphous 3396

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dispersion, a number of amorphous solid dispersions containing cellulose-based polymers (both commercially available and those designed and synthesized for the purpose) and surfactant (Tween 80) were evaluated. The additives (which are composed mostly of moderately hydrophobic polymers) investigated herein were combined with PVP, a hydrophilic polymer; PVP was included in the matrix due to its high aqueous solubility, facilitating rapid dissolution of the solid dispersion. Figures 7 and 8 show the size variation of the

solid dispersion of ritonavir/PVP 10:90 wt % in water) was analyzed at 30 min intervals for 2 h. Examination of this system by SEM revealed that the dispersed phase consisted of smooth, spherical liquid-like droplets with an initial size ∼500 nm (Figure 6a−c). This observation is consistent with those made

Figure 7. Plot of mean aggregate size (diameter, nm) versus time, showing the size of the colloidal aggregates produced during the dissolution of various amorphous solid dispersions of ritonavir (RTV). The error bars represent one standard deviation (n = 3).

colloidal aggregates with time for the different amorphous solid dispersions investigated herein. It is clear from the results that the additives have different effects on the initial size and stability of the colloidal aggregates. The ritonavir/PVP solid dispersion produced a colloidal phase with an initial average size of approximately ∼500 nm, which gradually increased with

Figure 6. Scanning electron micrograph of the ritonavir dispersed phase formed during the dissolution of ritonavir/PVP 10:90 wt % amorphous solid dispersion. Images were taken at different time points: (a) 5 min, (b) 30 min, (c) 60 min, (d) 90 min, (e) 120 min, and (f) 480 min.

in previous studies of systems undergoing LLPS.13,17,29 The structure of these colloidal aggregates is a sharp contrast to the needle-shaped particles characteristic of crystalline ritonavir. With time, coalescence of the aggregates and an increase in aggregate size was observed, a known characteristic of a system undergoing LLPS29,36 (Figure 6d−f). After ∼90 min, the morphology of the aggregates began to change, with the appearance of needle-like particles, suggesting that crystals had formed, in line with the results from the fluorescence experiment and previous induction time studies. In combination, these results indicate that the colloidal droplets created upon dissolution of an amorphous solid dispersion of ritonavir and PVP form at a concentration similar to that of the LLPS, and have similar properties to a ritonavir dispersed phase formed in highly supersaturated solutions generated by antisolvent addition.29 Influence of Amorphous Solid Dispersion Composition on Size and Stability of the Colloidal Aggregates. It was of interest to investigate the effect of various additives on the stability and size of the colloidal aggregates produced during dissolution of the various amorphous solid dispersions, since these factors have been proposed to have an impact on absorption and bioavailability of aggregate forming drug compounds.23,36 In addition to the ritonavir/PVP solid

Figure 8. Plot of mean aggregate size (diameter, nm) versus time, showing the size of the colloidal aggregates produced during the dissolution of various amorphous solid dispersions of ritonavir (RTV) containing Tween 80. The RTV/PVP 10:90 wt % amorphous solid dispersion was included for comparison. The arrows indicate the point of crystallization. The error bars represent one standard deviation (n = 3). 3397

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time, consistent with coalescence of the droplets. The significant increase in particle size at ∼90 min is consistent with crystal formation which was observed during fluorescence (Figure 5) and SEM (Figure 6) experiments. The addition of HPMC to the solid dispersion (ritonavir/HPMC/PVP ASD) resulted in a colloidal phase with an initial average size >1000 nm, which increased gradually with time to ∼2200 nm; in other words, HPMC was unable to stabilize the colloidal aggregates against coalescence. However, unlike the ritonavir/PVP ASD, crystallization of ritonavir was not detected during the time course of the experiment (2 h) when HPMC was present. ASDs containing the more hydrophobic negatively charged cellulose-based polymers, HPMCAS, CAAdP 0.33, and CAAdP 0.85, were more effective in stabilizing the colloidal aggregates; colloidal aggregates with an average size of ∼350 nm were initially generated during the dissolution of ASDs containing these polymers and showed little growth over the time frame of the experiment. The newly synthesized polymers, CAAdP 0.33/ 0.85, were more effective in stabilizing the colloidal dispersions, maintaining the size of the colloidal aggregates at ∼350 nm over the entire duration of the experiment (Figure 7). Figure 8 compares the effect of polymer/surfactant combinations, PVP/Tween 80 and PVP/Tween 80/CAAdP 0.33, on the size and stability of the colloidal dispersions. In the presence of the polymer/surfactant combinations, colloidal aggregates with an initial size of ∼250 nm were produced; Tween 80 was quite effective in preventing the coalescence of the colloidal aggregates. Several nonionic surfactants including Triton X-100, tylozapol, and β-octylglucoside have been found to be very effective stabilizers for NNRTI colloidal aggregates,23 preventing droplet coalescence. However, an abrupt change in mean particle size was observed at ∼30 and ∼90 min post dissolution for ASDs containing PVP/Tween 80 and PVP/ Tween 80/CAAdP 0.33, respectively. For the ritonavir/PVP/ Tween 80 ASD, SEM results indicate that the onset of crystal formation was ∼30 min (Figure S2 in the Supporting Information). Thus, although the nonionic surfactant Tween 80 prevented droplet growth via coalescence, it promoted crystallization. Furthermore, in comparison to the aspect ratio (length/width of crystal, based on SEM images) of the crystalline particles in the presence of PVP alone (∼3.9 after ∼90 min), the aspect ratio of the crystalline particles in the presence of Tween 80 was significantly higher (∼22.4). In other words, the crystalline particles formed in the presence of PVP/Tween 80 are more needle-like (elongated) compared to the crystalline particles formed in the presence of PVP alone. Because DLS results are less reliable for particles with irregular shapes such as needle-like particles (the particle size of irregular particles is expressed in terms of a spherical equivalent diameter, that is, the diameter of the sphere that would produce an equivalent light scattering pattern to the measured particle), the sizes reported for the Tween containing dispersions are likely to be unreliable once crystallization has occurred. Influence of Ionic Strength on Size of the Colloidal Aggregates. The effect of ionic strength on the size and stability of the colloidal aggregates was investigated by comparing the mean droplet size of the colloidal aggregates produced when ritonavir/PVP 10:90 wt % ASD was added to different aqueous media: distilled water or 100 mM sodium phosphate, with results shown in Figure 9. At the higher ionic strength (100 mM buffer), aggregate size was approximately 50% larger than that of the aggregates formed in distilled water.

Figure 9. Plot of mean aggregate size (diameter, nm) versus time, showing the effect of ionic strength on the colloidal aggregate size produced during the dissolution of RTV/PVP 10:90 wt % amorphous solid dispersion. The error bars represent one standard deviation (n = 3).

The colloidal aggregates had an initial average size of approximately 237 and 500 nm in water and 100 mM buffer, respectively. The size of the colloidal aggregates was comparable in two different types of aqueous media of the same ionic strength, namely, sodium phosphate buffer and NaCl solution (data not shown). Influence of Polymers on the Liquid−Liquid Phase Separation (LLPS) Concentration of Ritonavir. It has been demonstrated that an additive can change the LLPS concentration if it increases the equilibrium solubility of the crystalline form;29 therefore, it was of interest to investigate the effect of selected polymers on the LLPS concentration of ritonavir. The concentration at which the liquid−liquid phase transition was observed for ritonavir in the presence of 180 μg/ mL polymer solution (this is the concentration of polymer that would be present in solution following complete dissolution of the ASD) is summarized in Table 3. The LLPS concentration Table 3. Impact of Polymers on LLPS Concentration of Ritonavir (RTV) polymera RTV (no polymer) RTV-PVP RTV-PAA RTV-HPMC RTV-HPMCAS RTV-CAAdP 0.85 a

LLPS concn (μg/mL) 18.2 18.3 18.3 18.7 18.2 21.3

± ± ± ± ± ±

0.35 0.46 0.32 0.58 0.37 1.22

Concentration of polymer in solution: 180 μg/mL.

of ritonavir in the presence of the polymers was found to be similar to the LLPS concentration in the absence of an additive (Table 3), suggesting that the polymers are not substantially incorporated into the ritonavir-rich colloidal (second) phase or at least do not change the thermodynamic activity of this phase.29 Influence of Additives on Stability of the Colloidal Phase and Nucleation: Solvent Shifting Method. We and others have demonstrated that colloidal species can be readily generated by dissolving amorphous solid dispersions,17,18,20,22,29 but these experiments are somewhat compli3398

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cated by the dissolution step. In order to gain a better understanding of the properties of additives that are important for inhibiting droplet coalescence (i.e., size stability) and preventing the crystallization (i.e., phase stability) of the ritonavir-rich dispersed phase, the effect of a larger number of additives on the size of the colloidal aggregates and crystallization-induction time of ritonavir was evaluated, using the solvent shifting method to generate the colloidal dispersions. When a high concentration of ritonavir solubilized in a small volume of organic solvent was added to buffer solution, the resultant solution (20 μg/mL ritonavir concentration) was observed to be cloudy with a slight bluish color, characteristic of the presence of a second scattering phase. A solution concentration of 20 μg/mL corresponds to just higher than the LLPS concentration of ritonavir in 100 mM sodium phosphate buffer (Table 3). A comparison of the effect of selected predissolved additives (present at a concentration of 5 μg/mL) on the size and stability of the colloidal aggregates with time is shown in Figures 10−12. Fourteen additives with

Figure 11. Plot of mean aggregate size (diameter, nm) versus time, comparing the effect of predissolved additives (group 2 additives, 2 and 5 μg/mL polymer concentration for butyl-6-CO2HPull and PQ10, respectively) on the ritonavir (RTV) colloidal aggregates produced using the solvent-shifting method. The initial concentration of ritonavir in solution was 20 μg/mL. The error bars represent one standard deviation (n = 3).

Figure 10. Plot of mean aggregate size (diameter, nm) versus time, comparing the effect of predissolved additives (group 1 additives, 5 μg/mL additive concentration) on the ritonavir (RTV) colloidal aggregates produced using the solvent-shifting method. The initial concentration of ritonavir in solution was 20 μg/mL. The error bars represent one standard deviation (n = 3).

Figure 12. Plot of mean aggregate size (diameter, nm) versus time, comparing the effect of the predissolved cellulose-based polymers (group 3 and 4 additives, 5 μg/mL polymer concentration) on the ritonavir (RTV) colloidal aggregates produced using the solventshifting method. The initial concentration of ritonavir in solution was 20 μg/mL. The error bars represent one standard deviation (n = 3).

different structures and properties were employed, of which 3 are synthetic polymers (PVP, PVPVA, and PAA), one is a surfactant (SDS), one is a recently synthesized pullulan derivative, and the other 9 are cellulose-based polymers. Five of the 9 cellulose-based polymers are commercially available polymers, while the other 4 are recently synthesized cellulose esters, designed to span a wider range of chemical functionality than commercially available cellulose derivatives (the pullulan derivative is also novel and was recently synthesized for use in drug delivery). Although the initial sizes of the aggregates formed in the presence of the additives are comparable (∼180−250 nm), SDS, PVP, PAA, and HPMC were ineffective in preventing the growth of droplets of the colloidal phase. The size variation of the aggregates with time was similar to that in the absence of additive (Figures 10 and 12); the size of the aggregates increased gradually with time, indicative of droplet coalescence. In the presence of PAA and PVP, dispersed phase size evolved abruptly at approximately 100 and 130 min, respectively, while in the presence of SDS, a break in the curve was seen at around

40 min. The time points at which these changes to the size of the dispersed phase were observed are comparable to the crystallization-induction time of ritonavir in the presence of these additives. The crystallization-induction times were assessed by monitoring the UV absorbance and/or extinction as a function of time, as described previously.26 In other words, PAA, PVP, and SDS are ineffective crystallization inhibitors; in actuality, PAA and particularly SDS promoted crystallization from the colloidal dispersions (nucleation induction time of ritonavir in the absence of additive under the same experimental conditions is ∼120 min). The hydrophilic positively and negatively charged polymers, PQ-10 and butyl6-CO2HPull, were effective size stabilizers but induced crystallization of the droplets (Figure 11 and Table 5). In contrast, the more hydrophobic negatively charged cellulosebased polymers, HPMCAS, CAAdP 0.33, CAAdP 0.85, and CA 3399

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crystallization in ways similar to those of PVP and CAAdP 0.33, respectively, while the hydrophobically modified cellulose ethers HM-HEC and HM-HPMC were found to have similar influence on colloidal stability and crystallization to that displayed by PQ-10 and butyl-6-CO2HPull. In summary, the more hydrophilic uncharged additives were ineffective or less effective inhibitors of crystal formation and/or coalescence of the colloidal aggregates, while the more hydrophobic negatively charged cellulose-based polymers were effective in inhibiting crystal formation and stabilizing the droplets against coalescence (Table 5). Interestingly, certain additives promoted nucleation, leading to more rapid crystallization of ritonavir colloidal aggregates.

Table 4. Summary of Effect of Additives on Size, Stability, and Crystallization of Ritonavir (RTV) Colloidal Aggregates: Solvent Shifting Method additive RTV (without additive) PAA PVP PVPVA PQ-10 SDS Tween 80 HPMC HPMCAS HM-HEC HM-HPMC Butyl-6-CO2HPull CAAdP 0.85 CAAdP 0.33 CAAdB 0.81 CA Sub 0.90 a

size,a diam (nm) 188−830 213−922 188−631 215−913 183−303 268−511 178−182 203−714 235−352 292−385 331−395 189−208 203−248 210−214 204−210 220−276

induction time (min) 119.3 93.0 120.0 119.2 34.5 34.7 23.6 134.0 146.0 53.3 59.5 28.0 171.4 238.0 242.8 181.5

± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

8.14 1.50 3.00 1.00 4.95 1.04 1.59 6.00 7.90 1.06 2.56 2.65 5.83 4.82 7.66 4.95

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DISCUSSION Although the formation of drug colloidal aggregates has been widely reported, the mechanism of formation and the factors influencing the size and stability of the colloidal aggregates are poorly understood. In particular, the formation of colloidal aggregates is of interest in the context of better understanding how amorphous solid dispersions lead to enhanced bioavailability. It was first demonstrated that amorphous solid dispersions can lead to the formation of colloidal particles as far back as 1965, when Tachibana and Nakamura investigated the properties of solutions formed from the dissolution of amorphous solid dispersions of PVP and beta carotene.17 They found that these solutions contained colloidal sized aggregates. Since then, there have been numerous articles demonstrating that supersaturating dosage forms, consisting of an amorphous blend of the poorly water-soluble compound with a watersoluble polymer, are capable of generating colloidal sized species when dissolved in aqueous solution.16−18,20,38 Intriguingly, the formation of colloidal aggregates has been suggested to promote absorption18,23 and be important for preventing crystallization,38 although there is little direct evidence for either of these suppositions. In recent studies from our group, it has been demonstrated that certain poorly water-soluble compounds can be induced to undergo liquid−liquid phase separation (LLPS) when sufficiently high supersaturations are generated in aqueous media, for example by changing the pH from a low to a high value for a weak base,39 or by adding a concentrated organic solution of the drug to water.29 The resultant dispersed phase consists of submicrometer droplets that form when a certain concentration has been exceeded; this concentration is consistent for a given medium and temperature, is predictable, and can be related to the “amorphous solubility”. It was also demonstrated that the method of generating supersaturation does not impact the concentration at which LLPS occurs. It is thus highly likely that the colloidal species observed upon dissolution of the rapidly dissolving ASDs investigated herein originate from LLPS. It is well-known that ASDs dissolve to generate supersaturated drug solutions. If this concentration exceeds the LLPS concentration, then a drug-rich dispersed phase will be produced (as demonstrated in Figure 4) if crystallization does not occur first. Thus for the ritonavir ASDs, upon addition to the aqueous medium of a sufficient amount of ASD, a concentration higher than the LLPS concentration is generated following dissolution, and the excess drug phase separates as fine colloidal noncrystalline droplets of submicrometer size. The fluorescence experiment shown in Figure 5, as well as the SEM data, confirms the initially noncrystalline character of the dispersed phase. The characteristics of the

Aggregate size before crystallization.

Table 5. General Classification of Effect of Additive on Stability of Colloidal Phase

Sub 0.90, were effective size stabilizers, maintaining the aggregates at a size 280 min versus 120 min. Assuming that there is equilibrium between the dispersed phase and continuous phase, then the supersaturation is the same in each phase. Therefore, according to classical nucleation theory,44 the relative nucleation rate will be dependent upon the absolute concentration in each phase 3401

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dispersions and the effect of additives on the size stability and crystallization tendency of these droplets were examined. The concentration at which colloid formation was observed was comparable to the liquid−liquid phase separation concentration and thus to the theoretical amorphous “solubility” of ritonavir. The composition of the dispersed phase formed by dissolution of the amorphous solid dispersion was confirmed to be a noncrystalline, ritonavir-rich phase. Certain charged additives were found to both prevent droplet coalescence and inhibit crystallization, while other additives were found to promote crystallization. Therefore, the composition of the amorphous formulation will be critical in determining the formation and longevity of this colloidal phase. The insights into the mechanism of colloidal phase formation and factors that influence its stability have important implications for understanding the phase behavior of amorphous solid dispersions upon dissolution as well as the delivery of compounds with low aqueous solubility.

is more randomly substituted and has more polar (poly(ethylene oxide)) side chains, so might interact with colloidal ritonavir in quite different fashion from the other ethers. The cellulose ω-carboxyalkanoate polymers that are such effective inhibitors of size growth as well as crystallization of the ritonavir colloidal particles have pendant anionic substituents and are relatively randomly substituted (so do not have faces that differ in polarity as does butyl-6-CO2HPull). Their hydrophobic substituents are attached by ester linkages, and the longest such substituents are terminated by anionic carboxyl groups. It is not difficult to imagine that the cellulose ωcarboxyalkanoates are more localized on the exterior of the colloidal particles, and certainly this could help to explain their effectiveness in preventing size growth of these particles. Further studies are required to confirm whether and how these structural differences are responsible for the observed differences in crystallization behavior. In summary, with the exception of HPMC, the additives that are effective crystallization inhibitors are also effective colloidal dispersion stabilizers; however, the reverse does not hold true. The spontaneous formation of drug-rich nanodroplets upon dissolution of amorphous solid dispersions is also interesting in that it provides an alternative method of generating nanospecies and might be important not only for oral drug delivery but for other routes of delivery where it is desirable to form drug-rich nanodroplets. It is therefore important to consider factors impacting the size and size stability of the species formed. It has been proposed that the size and stability of colloidal aggregates depends on solution conditions (such as pH, buffer, and ionic strength) and the structure and properties of the molecules including the number of polar atoms, number of hydrogen bond donors and acceptors, conformational flexibility, and presence of ionized groups.23 The size of the colloidal aggregates reported in this study (the smallest aggregate size was ∼200 nm) is relatively larger than the sizes reported for a number of other aggregate forming compounds.23,32,33,42 The relatively large size of ritonavir colloidal aggregates may be attributed to the highly hydrophobic nature of ritonavir, its large molecular size, and/or the un-ionized nature of the drug compound at pH 6.8 (ritonavir pKa’s are 1.8 and 2.6). Analysis of aggregate size distributions for NNRTIs under various physiological conditions demonstrated dependence of aggregation size on solution pH;23 at constant ionic strength, aggregate size was found to increase with increasing pH (decreasing extent of ionization; these compounds are weak bases). In the current study, we found that it was possible to make much smaller and more stable aggregates when charged additives were present; ritonavir aggregates coalesced rapidly in the absence of a charged additive. This dependence of aggregate size on drug and/or additive ionization, and thus on pH, may have important implications for drug absorption since pH varies along the gastrointestinal (GI) tract. The size of the colloidal aggregates also increases with increasing ionic strength (Figure 9). This observation is in line with expectations, since it is well-known that interactions between hydrophobic groups in water are promoted by the presence of water structuring salts, which enhance hydrophobic interactions by making the solvent more polar.29,43

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ASSOCIATED CONTENT

S Supporting Information *

DSC thermogram of ritonavir dispersed phase, SEM micrographs of ritonavir aggregates (ritonavir/PVP/Tween 80 amorphous solid dispersion), and a comparison of SEM micrographs of ritonavir aggregates produced using the solvent-shifting method and from dissolution of an amorphous solid dispersion. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*Department of Industrial and Physical Pharmacy, College of Pharmacy, Purdue University, 575 Stadium Mall Drive, West Lafayette, Indiana 47907, United States. Tel: +1-765-496-6614. Fax: +1-765-494-6545. E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors thank Ashland Inc. and Daido Chemical Corporation for their kind donations of HM-HEC and HMHPMC, respectively. Support of the National Science Foundation through Grant DMR-0804609 is gratefully acknowledged. We also thank Chia-Ping Huang of the Life Sciences Microscopy Facility of Purdue University for performing SEM analyses.

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CONCLUSION In this study, the formation of a colloidal phase consisting of ritonavir nanodroplets upon dissolution of amorphous solid 3402

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