Article pubs.acs.org/journal/abseba
Highly Bioactive SDF-1α Delivery from Low-Melting-Point, Biodegradable Polymer Microspheres Dimitra A. Louka, Nathan Holwell, Brandon H. Thomas, Fei Chen, and Brian G. Amsden* Department of Chemical Engineering, Queen’s University, Kingston, Ontario K7L 3N6, Canada ABSTRACT: Aliphatic polyester biodegradable microspheres have been extensively studied for controlled and minimally invasive in situ protein delivery. However, they are commonly characterized by protein denaturation via acidic polyester degradation products, whereas their supraphysiologic modulus contributes to the inflammatory response upon implantation. To address these limitations, low-melting-point poly(ε-caprolactone-co-glycolide)-b-poly(ethylene glycol)-b-poly(ε-caprolactone-co-glycolide) (PEG-(PCG)2) copolymers were prepared and characterized for their ability to release bioactive stromalderived factor-1α (SDF-1α) as a representative therapeutic protein. The PEG molecular weight was chosen such that it would be crystalline at room temperature to promote easy handling of the microspheres, whereas the molecular weight and composition of the hydrophobic PCG blocks were adjusted to ensure the polymer was a viscous amorphous liquid at 37 °C. Microspheres prepared from the triblock copolymers completely degraded within 8 weeks in vitro with a minor decrease in microenvironmental pH. A prolonged release of SDF-1α was observed with its bioactivity highly retained after encapsulation and release. KEYWORDS: triblock copolymers, SDF-1α, caprolactone, glycolide, microspheres, controlled release
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INTRODUCTION Protein therapeutics have the potential to treat a wide range of conditions, are highly specific in their action, are expected to be less toxic than synthetically derived molecules, and behave more predictably in vivo.1 For many therapeutic proteins, because of their low effective concentrations and paracrine action, localized and sustained delivery would be advantageous.2−4 Biodegradable microsphere formulations in which the protein is distributed as solid particles have been extensively explored for this purpose. Their small size and thus ready injectability as well as the ability to tune polymer degradation and thus release kinetics make this formulation approach attractive for protein delivery. The most widely investigated biodegradable polymers for preparing microspheres have been the copolymers of poly(lactide-co-glycolide) (PLG). This popularity stems from their commercial availability in a variety of molecular weights and compositions, their readily adjustable degradation rates, and their long history of in vivo safety as components of biodegradable devices such as sutures and screws.5,6 However, the internal accumulation of hydrolytic degradation products of PLGs has been implicated in the denaturation of multiple protein therapeutics.7−13 Moreover, PLGs have high moduli of elasticity, ranging from 0.2 to 7 GPa, depending on glycolide to lactide monomer ratio and molecular weight,14,15 whereas many soft tissues into which these microspheres will be implanted have moduli ranging from a tens of Pa to hundreds of kPa.16,17 The mismatch in stiffness between the PLG and the surrounding tissue contributes to an enhanced inflammatory © XXXX American Chemical Society
response, above that contributed by the release of acidic degradation products as a result of the hydrolysis of PLG,18 because of shear resulting in the generation of a thick fibrotic capsule surrounding the microspheres19 that impedes drug movement into the surrounding tissue as well as induces patient discomfort. In an effort to reduce tissue irritation and the influence of acidic degradation products on protein denaturation, we have explored the concept of preparing microspheres using a biodegradable polymer with a melting point between room temperature and body temperature.20 At room temperature, the semicrystalline nature of the polymer allows for good flow and handling properties of the microspheres while the low melting point ensures that the polymer microspheres are similar to viscous liquid droplets once warmed to body temperature, and thus are not mechanically irritating. These polymers were based on ε-caprolactone and trimethylene carbonate, as highmolecular-weight poly(ε-caprolactone) has a glass transition temperature of −60 °C and a melting point of 63 °C, which can be reduced by decreasing its molecular weight as well as by copolymerization,21,22 whereas poly(trimethylene carbonate) is amorphous and has a glass transition temperature of −26 to −15 °C, depending on molecular weight.23 A low glass transition temperature is desirable to produce a low viscosity Special Issue: Biomaterials in Canada Received: June 23, 2017 Accepted: September 25, 2017 Published: September 25, 2017 A
DOI: 10.1021/acsbiomaterials.7b00403 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX
Article
ACS Biomaterials Science & Engineering liquid copolymer upon melting.24 To provide for effective dispersion in aqueous media, we used a low-molecular-weight (1000 Da) poly(ethylene glycol) (PEG) diol as an initiator, resulting in a triblock copolymer. In this previous design, the hydrophobic poly(ε-caprolactone-co-trimethylene carbonate) blocks provided the desired melting properties. However, although the resulting hydrophobic blocks of the copolymer had a melting range of 19 to 34 °C and could be formulated into microspheres for prolonged and highly bioactive growth factor delivery, the microsphere degradation rate was very slow; after 28 weeks, only approximately 5% mass loss was observed in vitro.20 It was the objective of this work to create a faster degrading copolymer for the same purpose. To achieve this objective, the trimethylene carbonate in the hydrophobic block was replaced with glycolide, which is more readily hydrolyzed,5 despite its hompolymer possessing a higher glass transition temperature (35−40 °C25) and thus leading to an increase in the viscosity of the hydrophobic block region when above its melting point. Furthermore, instead of relying on the hydrophobic blocks to provide the desired crystallinity at room temperature, the thermal properties of PEG were exploited. The 1000 Da PEG used previously was replaced with 4000 Da PEG, which has a higher melting point (53−60 °C) than does 1000 Da PEG (35−40 °C).26 To assess the potential of this formulation approach, stromal-derived factor-1 alpha (SDF-1α) was used as a representative therapeutic protein. SDF-1α is of potential therapeutic interest because of its ability to recruit monocytes and endogenous stem cells to facilitate wound repair and angiogenesis.27−31
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The supernatant was removed by centrifuging at 3200 rpm for 5 min. The obtained purified copolymer was dried at 50 °C under vacuum overnight. Copolymer Characterization. The composition analysis was performed in chloroform-d using 1H nuclear magnetic resonance (NMR) spectroscopy on a Bruker Avance spectrometer 400 MHz. The number-average molecular weight of the copolymer (Mn) was determined from the 1H NMR spectra integrations of PEG methoxy protons (IPEG) compared to the integrations of the methylene protons of glycolyl (−OCH2CO−) (G) (IG) and caproyl (−OCH2CH2CH2CH2CH2CO−) (C) (IC) monomer units that are adjacent to the carbonyl group. The degree of randomness (R) was calculated using the integrations of the peaks corresponding to the glycolyl and caproyl sequences of the 1H spectra as reported by Dobrzynski et al.32 Briefly, the experimental lengths of the two sequences were calculated from eqs 1 and 2:
LC = (ICC + ICG)/ICG LG =
(1)
LC k
(2)
where LC and LG are the experimental average lengths of caproyl and glycolyl sequences, respectively. ICC is the integration of the caproyl methylene peak corresponding to caproyl−caproyl (CC) adjacent sequences (δ = 4.1 ppm) and ICG represents the integration of the caproyl peak corresoponding to caproyl-glycolyl (CG) adjacent sequences (δ = 4.2 ppm). k is the molar ratio of caproyl to glycolyl unit, given by k = IC/IG
(3)
The lengths of the caproyl and glycolyl units in randomly structured copolymers were calculated from eqs 4 and 5:
k+1 k
(4)
LCR = k + 1
(5)
LGR =
EXPERIMENTAL SECTION
Materials and Methods. Poly(ethylene glycol) diol (4000 Da) (PEG 4000), ε-caprolactone (C), and anhydrous toluene were purchased from Acros Organics, USA. Glycolide monomer (G) was purchased from Altasorb, USA and anhydrous tin(II) 2-ethylhexanoate (Sn(Oct)2) was from Alfa Aesar (USA). Chloroform-d, lysozyme from chicken egg white, trehalose, bovine serum albumin (BSA), fetal bovine serum (FBS), and HyClone penicillin−streptomycin (P/S) solution were obtained from Sigma-Aldrich, Canada. Methanol, dichloromethane (DCM), anhydrous ethyl alcohol, Pierce bicinchoninic acid (BCA) protein assay reagents, phosphate buffered saline 10× (PBS), SNARF-1, Lysosensor yellow/blue fluorescence dyes, and RPMI-1640 medium (ATCC modified) were purchased from Fisher Scientific, Canada. Human stromal derived factor-1-alpha (SDF-1α) and the appropriate ELISA kit were purchased from PeproTech Inc., USA. The NCI-H69 male human small cell lung carcinoma cell line was purchased from ATCC (USA), whereas MTT assay reagent was obtained from Invitrogen, and a QuantiFluor dsDNA assay kit was purchased from Promega. Copolymer Synthesis. Poly(ε-caprolactone-co-glycolide)-b-poly(ethylene glycol)-b-poly(ε-caprolactone-co-glycolide) (PEG-(PCG)2) triblock copolymers were prepared in oven-dried glass ampules under Ar atmosphere. Appropriate amounts of monomers and PEG were weighed directly into the ampule and melted at 110 °C. The ampule was taken out of the 110 °C oven and an appropriate volume of the catalyst solution in dry toluene (dilution factor = 1:3.5) was added to the mixture; 1 mmol of stannous 2-ethylhexanoate catalyst for each mol of monomer was used. Ar gas was gently blown into the ampule and the ampule was then vortexed and flame-sealed under vacuum (ca. 4 kPa). Polymerization took place in an oven at 150 °C for 2.5 h. Brief (5 s) vortex was applied every 20 min during polymerization. After polymerization, the glass ampule was snapped open and the polymerization reaction was stopped by dissolving all contents in cold DCM. The resulting copolymers were purified twice by dissolution in DCM and precipitation in −20 °C methanol for 12 h.
Finally, the degree of randomness was then obtained from
R=
LGR LR = C LG LC
(6)
The degree of randomness for completely random chains is 1 and is 0 for diblock copolymers. A high concentration of alternating monomer sequences is implied when R is greater than 1. Thermal analysis was performed using a Mettler Toledo DSC1 differential calorimeter calibrated with an indium standard. Approximately 10 mg samples were subjected to a heating−cooling−heating sequence of −80 to 270 °C to −80 °C at a heating rate of 10 °C/min for all cycles. The enthalpy of fusion (ΔHf) was calculated from the area of the endothermic melting peak integrated using the instrument’s software and used to calculate the percent crystallinity of the copolymers. The percent crystallinity (% X) at 37 °C was calculated as the enthalpy of melting of the sample divided by the enthalpy of melting of pure 100% crystalline poly(ε-caprolactone) (ΔHm,PCL = 139.6 J/g).33 The dispersity (Đ), number and weight-average molecular weights of the copolymers were measured using a Waters GPC 2690 equipped with four Waters Styragel HR columns connected in series and a multiangle laser light scattering Wyatt Technology DAWN EOS detector. HPLC grade tetrahydrofuran (THF) was used as the mobile phase with the flow rate set at 1 mL/min and temperature at 25 °C. Samples consisted of 5 mg/mL polymer solution in THF. The refractive index increment (dn/dc) was determined using a Wyatt Optilab rEX refractometer at a wavelength of 690 nm. To measure the viscosity of the hydrophobic blocks at 37 °C, we prepared copolymers of the same number-average molecular weight and comonomer composition as the triblock copolymers under the same conditions, but with 1-octanol (Oct) as the initiator instead of PEG 4000. The viscosities of Oct-PCG copolymers were measured at 37 °C using a TA Instruments AR2000 controlled stress rheometer. B
DOI: 10.1021/acsbiomaterials.7b00403 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX
Article
ACS Biomaterials Science & Engineering To measure the ability of the copolymers to absorb water, the copolymers were heat molded into 3 mm thick × 1.5 cm square thin, nonporous films (n = 3), immersed in an aqueous environment at 37 °C. To create the films, 100 mg (mdry) of the copolymer was placed in a square mold and into a 100 °C oven for 10 min to melt flat at the bottom of the mold. The molds were quickly transferred to a desiccator and remained there to cool at room temperature and crystallize before being removed from the mold. The weight of the dry films was recorded at room temperature and then the films were placed dry in a 37 °C incubator for 2 h. Phosphate-buffered saline pH 7.4 at 37 °C was added to the films and the samples were incubated for 24 h. After 24 h, the buffer was removed and the excess moisture was blotted off the surface of the film. The weight was recorded again at room temperature (mwet) and the samples were dried overnight at 110 °C in an oven. After drying, the films were transferred to a desiccator to cool to room temperature, after which the weight of the dry film was recorded. The equilibrium water content (EWC) was calculated as EWC =
m wet − mdry mdry
piece of aluminum plaque connected to the second electrode of the circuit that was immersed into the collector bath (cooled at −70 °C in dry ice bath). The voltage applied was 7 kV and the current was set at 0.1 mA. The distance from the tip of the needle to the surface of the collector solution was 7 cm. The polymer solution or polymer/protein suspension was ejected through a flat tip 20 gauge needle syringe using the syringe pump set at 0.5 mL/min. Solid SDF-1α or lysozyme (model protein) containing particles were added to a 0.7 mg/mL polymer solution and briefly sonicated prior to electrospraying, at a concentration of 1.5 or 3% wt. of dry polymer. Blank, nonprotein loaded microspheres were fabricated for an in vitro degradation study. The droplets produced quickly solidified upon contact with the cold ethanol. The bath was constantly stirred at 400 rpm and small dry ice pieces were added several times to the ethanol bath throughout the process. The resulting microspheres were held for 48 h in ethanol at -80 °C to allow for the DCM to be extracted to the ethanol. The suspended microspheres were then transferred to a−20 °C freezer to allow for polymer crystallization and were retained at this temperature for 48 h. After crystallization, microspheres were subjected to size separation. Sieving was applied using two sieves with opening diameters of 45 and 100 μm. This size fraction was dried of solvent and used for degradation and release experiments. The average diameter of the microspheres was determined from scanning electron microscopy (SEM) images using Adobe Photoshop CS6 image software. The scale on the SEM images was used to calibrate the integrated scale of the software to the corresponding number of pixels. An average diameter was calculated from the measurements of 100 microspheres (n = 3). The encapsulation efficiency of the loaded protein was determined by dissolving 10 mg of protein-loaded microspheres in 1 mL DCM. The solution was vortexed for 2 min at maximum speed and centrifuged at 500 000 rpm for 5 min. The supernatant was removed and the precipitated protein particles were air-dried. The dry particles were dissolved in a PBS pH 7.4 buffer solution and the protein content was measured using Pierce BCA assay at 562 nm with an EnSpire 2300 microplate reader (PerkinElmer). The analysis was carried out in triplicate. The encapsulation efficiency (E) was calculated from m E= 100% mo (8)
100% (7)
Protein Particle Preparation. The protein particles and loaded microsphere formulation were optimized using lysozyme as a model protein. Three different compositions, 10, 40, and 60% of trehalose in lysozyme, were codissolved in pH 7.4 PBS at a concentration of 5% w/ v, frozen in liquid nitrogen, and lyophilized. The lyophilized particles were reduced in size by grinding and sieving through U.S Standard No.325 (45 μm) and No. 500 (25 μm) sieves. The
