Adjustable Degradation Properties and Biocompatibility of Amorphous

Jun 10, 2014 - Radical Ring-Opening Polymerization: Scope, Limitations, and Application to (Bio)Degradable Materials. Antoine Tardy , Julien Nicolas ,...
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Adjustable Degradation Properties and Biocompatibility of Amorphous and Functional Poly(ester-acrylate)-Based Materials Jenny Undin, Anna Finne-Wistrand, and Ann-Christine Albertsson* Department of Fibre and Polymer Technology, School of Chemical Science and Engineering, KTH Royal Institute of Technology, SE-100 44, Stockholm, Sweden ABSTRACT: Tuning the properties of materials toward a special application is crucial in the area of tissue engineering. The design of materials with predetermined degradation rates and controlled release of degradation products is therefore vital. Providing a material with various functional groups is one of the best ways to address this issue because alterations and modifications of the polymer backbone can be performed easily. Two different 2-methylene-1,3-dioxepane/glycidyl methacrylatebased (MDO/GMA) copolymers were synthesized with different feed ratios and immersed into a phosphate buffer solution at pH 7.4 and in deionized water at 37 °C for up to 133 days. After different time intervals, the molecular weight changes, mass loss, pH, and degradation products were determined. By increasing the amount of GMA functional groups in the material, the degradation rate and the amount of acidic degradation products released from the material were decreased. As a result, the composition of the copolymers greatly affected the degradation rate. A rapid release of acidic degradation products during the degradation process could be an important issue for biomedical applications because it might affect the biocompatibility of the material. The cytotoxicity of the materials was evaluated using a MTT assay. These tests indicated that none of the materials demonstrated any obvious cytotoxicity, and the materials could therefore be considered biocompatible.

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INTRODUCTION A wide variety of biodegradable polymers have been developed in recent years and used primarily in the area of tissue engineering.1 The most commonly used aliphatic polyesters, poly(εcaprolactone) (PCL) and poly(L-lactide) (PLLA)2,3 and copolymers thereof, are most often synthesized through ringopening polymerization (ROP) of cyclic esters using metallic, organic, anionic, or cationic catalysts.4−7 However, a less frequently exploited route for preparing polyesters is through the radical ring-opening polymerization (RROP) of cyclic ketene acetals.8−10 RROP of the cyclic ketene acetal 2methylene-1,3-dioxepane (MDO) yields PCL with some branching, the extent of which is dependent on the reaction temperature.11,12 The resulting PCL will be totally amorphous instead of semicrystalline, as is the conventionally availably PCL. A major benefit of RROP is that it allows the copolymerization of cyclic ketene acetals with different functional vinyl monomers and, as a result, introduces functionality into degradable materials in a relatively simple manner. We have previously synthesized amorphous and functional polyesters through the copolymerization of 2-methylene-1,3dioxepane (MDO) and glycidyl methacrylate (GMA).13 GMA has a carbon−carbon backbone and is therefore not susceptible to main-chain hydrolysis. However, GMA has an ester group that is located proximal to the polymer backbone and can potentially degrade, but with a relatively slow degradation rate © 2014 American Chemical Society

and without affecting the main chain. One method of introducing degradability into the polymer backbone is to copolymerize vinyl monomers with cyclic ketene acetals. The applicability of copolymerization of different cyclic ketene acetals is an interesting platform for the creation of materials for future use in, e.g., biomedical applications.13−15 However, the commonly used polyesters have some inherent drawbacks that need to be addressed because their crystalline nature results in slow degradation and can be a concern when, for example, pieces of the material are detached during degradation in the body.16−18 Although these materials may be considered biocompatible, they can release acidic degradation products upon degradation, resulting in a decrease in pH and producing a very acidic local environment that may cause inflammatory responses. The degradation products are generated at the surface and within the bulk of the material; in some thick specimens, the carboxylic acid groups may be trapped, resulting in an inhomogeneous degradation and rapid release of degradation products.19−23 It is therefore important to consider acidity issues when designing polyester-based biomaterials, and more information is needed about the cytotoxic effects of the degradation products at the implantation site.24 Received: May 13, 2014 Revised: June 5, 2014 Published: June 10, 2014 2800

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H NMR (400.13 MHz, CDCl3): δ = 3.47 ppm (s, 2H, CCH2), δ = 3.76−3.91 ppm (m, 4H, 2 −OCH2-), δ = 1.72−1.78 ppm (m, 4H, −OCH2 CH2CH2CH2O−). 13C NMR (100.61 MHz, CDCl3): δ = 66.11 ppm (CCH2), δ = 70.14 ppm (−OCH2 CH2CH2CH2O−), δ = 28.98 ppm (−OCH2 CH2CH2CH2O−): δ = 162.88 (CCH2−). Polymerizations. Free Radical Ring-Opening Polymerization of MDO. The polymerization of MDO was performed in bulk using 0.2 mol % of the initiator 2,2-azobis(isobutyronitrile) (AIBN) under an inert atmosphere. The monomer and the initiator were weighed into a 25 mL round-bottom flask equipped with a three-way valve and a magnetic stirrer. The system was cooled, and air was removed by three freeze-vacuum-argon/nitrogen-thaw cycles. The flask was immersed in a thermostated oil bath at 60 °C for 48 h. The resulting polymer was dissolved in chloroform and precipitated in heptane; it was then dried at room temperature until all of the solvent had evaporated. Copolymerization of MDO and GMA. The copolymerization of MDO and GMA was performed in bulk using 2 mol % of the initiator 2,2-azobis(isobutyronitrile) (AIBN) under an inert atmosphere. The monomers and the initiator were weighed into a 25 mL roundbottomed flask equipped with a three-way valve and a magnetic stirrer. The system was cooled in liquid nitrogen, and air was removed by three freeze-vacuum-argon/nitrogen-thaw cycles. The flask was immersed in a thermostated oil bath at 60 °C for 3 h. The resulting polymer was dissolved in chloroform and precipitated in heptane; it was then dried at room temperature until all solvent had evaporated. The precipitation was repeated until all of the monomer, as detected with proton nuclear magnetic resonance spectroscopy (1H NMR), had been removed. Ring-Opening Polymerization of ε-Caprolactone (ε-CL). The homopolymer of ε-caprolactone (εCL) was synthesized in bulk with ethylene glycol as initiator and stannous octoate (Sn(Oct)2) as catalyst. The initiator, the catalyst, and the monomer were added into a silanized round-bottom flask under a nitrogen atmosphere inside a glovebox. The reaction was started by immersing the flask in a thermostated oil bath at 110 °C for 48 h. The monomer-to-catalyst ratio, [monomer]/[Sn(Oct)2] were set to 10 000, and the degree of polymerization (DP) was set to 400. The resulting polymer was dissolved in chloroform and precipitated in a mixture of cold heptane and methanol (95:5). Film Preparation. Films of the copolymers and the reference polymers were prepared by dissolving the polymers (1.5 g) in chloroform (CHCl3), followed by solution-casting in glass Petri dishes. The films were dried at room temperature by solvent evaporation and had an average thickness of approximately 0.2 mm. Hydrolysis. The different materials were subjected to hydrolytic degradation in PBS, pH 7.4, and in deionized water at 37 °C. The PBS was diluted to 5000 mL, and the pH was adjusted to 7.4 with 1 M NaOH; 0.04 wt % NaN3 was added to prevent microbial growth. Each sample, approximately 10 mg of polymer, was placed in a 20 mL vial containing 10 mL of deionized water or PBS. The sample vials were sealed with septa and placed in a thermostatically controlled roller incubator at 37 °C and 60 rpm. After different time periods between 1 and 133 days, triplicate samples of each material were withdrawn from each test environment, dried under reduced pressure, and analyzed by various techniques. Cell Proliferation/Viability. Extract Preparation. Films were incubated in culture medium for 24 and 72 h at 37 °C with constant shaking (60 rpm) to simulate closely the effect of the products in a dynamic environment. The extracts were then filtered (0.2 mm pore size) and maintained at −20 °C for further use. For the extraction tests, complete culture medium in tissue culture polystyrene (TCPS) served as the negative control. Methylthiazol Tetrazolium (MTT) Assay. Cell viability and proliferation were analyzed using the methylthiazol tetrazolium (MTT) mitochondrial reaction. Mouse L929 cells were seeded in 96-well cell culture plates (1 × 104 cells/cm2) and reached 80% confluence in 24 h. The 24- and 72-h extracts were placed in contact with the monolayers for 24 h, and a negative control was included. Viability tests were performed using the MTT assay on quadruplicate samples. The culture medium was replaced by a solution of MTT 1

Many of these monomers lack functional sites that allow alterations to and modifications of the polymer backbone.25 Functional polymers offer the ability to be tailored to suit special application areas, for example, postfunctionalization through the immobilization of heparin on polyester-based materials.26 A central challenge is thus to design materials containing functional groups that will allow the control of the degradation rate and release of degradation products.27−30 It is well-known that degradation starts in the amorphous regions and continues in the crystalline regions after almost all of the amorphous parts have been degraded.31 The crystallinity of materials with their heterogeneous degradation profiles has contributed to a variety of problems in the area of tissue engineering. The introduction of amorphous blocks, or fully amorphous polymers, with a controlled microstructure allows for better control of the degradation rate.32−34 An amorphous material that is very homogeneous is capable of a more uniform rate of hydrolysis. The aim was to use the previously described scientific concept and design a functional amorphous ester-based material with adjustable degradation properties. The hypothesis was that, by changing the amount of functional groups in the material, the degradation rate will change. As a result, there will be a change in the amount of acidic degradation products released from the material. In addition, this will prevent the rapid release of acidic degradation products that might adversely affect the biocompatibility of the material during the degradation process. The degradation pattern and biocompatibility of the synthesized and amorphous ester-based material, poly(MDOco-GMA), was defined. Poly(MDO-co-GMA) offers a predefined design pattern of active functional groups, resulting in a material that can be adjusted easily for use in the chosen area of application. Two different copolymers of poly(MDO-co-GMA) that were synthesized using different monomer feed ratios and the semicrystalline PCL were subjected to hydrolytic degradation in two different environments.

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EXPERIMENTAL SECTION

Materials. Chemicals. 2,2-Azobis(isobutyronitrile), AIBN, (Acros Organics, USA) was purified by recrystallization from methanol prior to use. ε-Caprolactone (ε-CL) (Aldrich) was dried over calcium hydride for at least 24 h and subsequently distilled at reduced pressure under an inert atmosphere prior to use. Stannous octoate (Sn(Oct)2) (Sigma-Aldrich, Sweden) was dried over molecular sieves (3 Å) before use. Glycidyl methacrylate, GMA, (Aldrich, Germany) was distilled under reduced pressure prior to use. Bromoacetaldehyde dimethyl acetal (Alfa Aesar; Germany), 1,4butanediol (Ridel deHaën, Germany), Dowex 50 (Acros Organics, USA), Aliquat 336 (Aldrich, Germany), potassium tert-butoxide (Aldrich, Germany), anhydrous tetrahydrofuran, THF, (LabScan, Ireland), chloroform (LabScan, Ireland), hexane (LabScan, Ireland), methanol (Fisher Scientific, Germany), chloroform-d (99.8%, with silver foil (Cambridge Isotope Laboratories), and LC-MS grade water (Merck) were used as received. Dulbecco’s phosphate-buffered saline (PBS, pH =7.4 Cat. No. H15−011) was purchased from PAA laboratories (Austria). Monomer Synthesis. 2-Methylene-1,3-Dioxepane (MDO). The monomer 2-methylene-1,3-dioxepane (MDO) was synthesized in a two-step process as previously described.9,15,35,36 Briefly, the first step was the ring-closure of bromoacetaldehyde dimethyl acetal with 1,4butanediol, followed by elimination to create the exomethylene functionality at position 2. 2801

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(Sigma, St Louis, MO, USA), and the cells were incubated for 4 h. Viable cells, i.e., those with functional mitochondrial dehydrogenase, were able to reduce the yellow MTT to a purple formazan product. The medium was discarded, and the precipitated formazan was dissolved in 0.5 mL DMSO containing 6.25% (v/v) 0.1 M NaOH. The end product was quantified using a microplate spectrophotometer (BMG LABTECH, GmbH, Germany) at a wavelength of 570 nm and expressed as optical density (OD) units. Characterization. Nuclear Magnetic Resonance Spectroscopy (NMR). Proton and carbon nuclear magnetic resonance spectroscopy, 1 H NMR (400.13 MHz) and 13C NMR (100.61 MHz), spectra were obtained using a Bruker Avance DPX-400 NMR instrument. For the 1 H NMR and 13C NMR measurements, 5−100 mg of sample was dissolved in 1 mL deuterochloroform (CDCl3) in a 5 mm diameter sample tube. The spectra were calibrated using residual solvent signals (δ = 7.26 ppm and δ = 77.0 ppm for 1H NMR and 13C NMR, respectively). Size Exclusion Chromatography (SEC). The number-average molecular weight (Mn) and dispersity (Đ) of the polymers were determined by SEC. The samples were prepared by adding 20−30 mg of the polymers and 10 mL of chloroform to SEC-vials (2−3 mg/mL). The polymers were analyzed using a Verotech PL-GPC 50 Plus system equipped with a PL-RI Detector and two PLgel 5 μm MIXED-D (300 × 7.5 mm) columns from Varian. The samples were injected using a PL-AS RT Autosampler for PL-GPC 50 Plus, and chloroform was used as mobile phase (1 mL/min, 30 °C). The system was calibrated, and a calibration curve was created using polystyrene standards with a narrow molecular weight distribution ranging from 160−371 000 g/ mol. Corrections for flow rate fluctuations were made using toluene as an internal standard. The Cirrus GPC Software was used to process the data. Differential Scanning Calorimetry (DSC). The thermal properties of the polymers were measured by DSC (Mettler Toledo DSC 820 module) under a nitrogen atmosphere (nitrogen flow rate 80 mL/ min) using 5−10 mg of polymer encapsulated in an aluminum pan. The measurements were performed from −70 to 125 °C at heating and cooling rates of 10 °C/min. The glass transition temperature (Tg) was noted in the DSC-thermogram as the midpoint temperature of the glass transition peak (endothermic peak) in the second heating cycle. pH-Measurement. The pH of the degradation solution was determined using a VWR symphony meter SB70P equipped with a Biotrode (Hamilton, USA). The electrode had a pH range of 0−14, and the reference was a PROTELYTE electrolyte. The instrument was calibrated using buffer solutions at pH 4 and pH 7. Electrospray Ionization-Mass Spectrometry (ESI-MS). A Finningan LCQ ion trap mass spectrometer (Finnigan, San Jose, CA) was used to analyze the oligomeric degradation products. The polyester specimens were dissolved in a water/methanol system (1:2 v/v) and continuously infused into the ESI source by the instrument’s syringe pump at a rate of 3 mL/min. The LCQ ESI source was operated at 5 kV, and the capillary heater was set to 250 °C. Nitrogen was used as a nebulizing gas. The ions of interest for ESI-MS experiments were isolated monoisotopically in the ion trap and collisionally activated. Helium, present as damping gas in the mass analyzer, acted as the collision gas. The RF amplitude, which had a significant voltage range, was set to a value that caused the peak height of the parent ion to decrease by at least 50%. A positive ion mode was used for analysis.

two different environments phosphate buffer solution (PBS) and deionized water, at 37 °C for up to 133 days. The reaction milieu and temperature were chosen to mimic the milieu in the body and because the temperature of 37 °C is above the glass transition temperature (Tg) for all of the polymers under evaluation. Water absorption and mass loss are facilitated as the degradation is carried out above the glass transition temperature of the copolymers. At temperatures below the Tg, the polymers lose their flexible nature. This leads to higher chain mobility, which allows penetration of more water and migration of the degraded fragments. The molecular weight, mass loss, pH, and hydrolysis-induced changes in the copolymer composition were all monitored, as was the release of acidic monomeric and oligomeric degradation products. To ensure that no unexpended and toxic degradation products were formed during the hydrolysis, the biocompatibility of the materials were tested in vitro. Material Properties Prior to Degradation. The structures of the copolymer and homopolymer are shown in Scheme 1. The copolymer compositions and the original Scheme 1. Structures of the Materials Subjected to Hydrolysis (I) PCL and (II) poly(MDO-co-GMA)

molecular weights are given in Table 1. The two different MDO-GMA-based copolymers consisted of MDO and GMA at feed ratios of 80/20 and 65/35, respectively. The different feed ratios were chosen to vary the two copolymers as much as possible. More MDO being present in the feed results in a copolymer with properties similar to the homopolymer of Table 1. Copolymer Compositions, Original Molecular Weights, and Dispersities Prior to Degradation

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RESULTS The design of materials with predetermined degradation rates and controlled release of degradation products is essential in the area of tissue engineering. A degradable material consisting of active functional groups enables for tuning the properties of the material toward a special application. Two copolymers of poly(MDO-co-GMA) that were synthesized using different monomer feed ratios and the semicrystalline PCL were subjected to hydrolytic degradation in

polymer

f MDOa

conversion [%]b,c

FMDOa,b,c

Mn [g/mol]d

Đd

Tg [°C]e

MDO100 MDO80 MDO65 PCL GMA

1.0 0.80 0.65 -

99.0 92.4 88.1 100 100

1.0 0.54 0.42 -

48 000 255 000 235 000 50 000 200 000

1.9 1.7 1.8 1.4 1.7

−59.0 −4.1 22.2 −60.0 59.2

a

f MDO is the mole fractions of MDO in the feed, and FMDO is the mole fraction of MDO in the polymer. b60 °C, bulk, 2 mol % AIBN, 3 h reaction time. cDetermined by 1H NMR in CDCl3. dDetermined by SEC using narrow polystyrene standards, chloroform as eluent, and toluene as internal standard. eDetermined by DSC as the midpoint temperature of the transition in the second heating cycle. 2802

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Figure 1. Composition of (left) MDO80 and (right) MDO65 after different hydrolysis times in PBS at 37 °C, as determined by 1H NMR by comparing the peak intensities of the comonomers, MDO (black) and GMA (gray).

Figure 2. Molecular weight changes of MDO80 (■), MDO65 (●) and PCL (▲) during 133 days of hydrolysis (left) in deionized water at 37 °C, and (right) in PBS at 37 °C.

copolymer produce more hydrophilic regions, which renders the copolymer more vulnerable to side-chain hydrolysis compared with more randomly distributed GMA blocks. Conversely, with more MDO distributed in the copolymer, more degradable units will be available in the backbone. As a result, the composition of the copolymer greatly affects its degradation rate. The changes in the composition of the copolymers during hydrolysis were determined by 1H NMR, as shown in Figure 1. The composition of both copolymers remained almost constant for the first 28 days. These results correlate well with the ESI-MS results, which did not exhibit any oligomeric or monomeric degradation products for the same period of time. As the time of hydrolysis increased, the composition shifted to a higher GMA content as the MDO degraded. A possible reason for this shift could be the existence of some steric hindrance by the bulky GMA chain, which hampers the hydrolysis of the ester side groups;38 the degradation that occurs will be primarily main-chain degradation. At the end of the 133-day hydrolysis period, the MDO content in the copolymers MDO80 and MDO65 was 23% and 28%, respectively. Molecular Weight Changes and Mass. The changes in molecular weight of the copolymers and reference materials were followed with SEC. A decrease in the molecular weight of all of the polymers was observed immediately after submersion in both deionized water and PBS, as shown in Figure 2. There was no substantial difference in the rate of decrease for the molecular weight between the copolymers in deionized water, although the molecular weight of the copolymer with highest degree of MDO, MDO80, decreased slightly faster. This was expected due to the higher amount of hydrolyzable ester groups in the backbone. However, a difference was observed between the hydrolysis in deionized water and that in

MDO, poly(2-methylene-1,3-dioxepane) (PMDO), which is a viscous liquid at room temperature. A copolymer with less than 65 mol % of MDO in the feed will have a Tg that is higher than the normal room temperature and, more importantly, higher than body temperature. This composition will render the copolymer stiff and brittle at body temperature with very little elasticity. This is also the reason why the homopolymer of GMA, poly(glycidyl methacrylate) (PGMA), is not suitable for film formation. Degradation. Polyesters have been shown to degrade primarily in the body via chemical hydrolysis of the hydrolytically unstable ester bonds, yielding oligomers with carboxyl acid end groups. The hydrolysis was performed in two parallel degradation media: deionized water and a phosphatebuffered saline solution. Deionized water was necessary during the electrospray ionization-mass spectrometry (ESI) analysis of the degradation products because salt is well-known to be destructive to the ion source of the ESI. The residual molecular weight, mass, and changes in pH were determined in both environments. Hydrolysis-Induced Changes in the Copolymer Composition. During degradation, the amorphous regions undergo hydrolysis faster than do the crystalline regions due to their higher rate of water uptake.37 The degradation process begins in the amorphous regions, which then provides the nondegraded chains with more space for mobility and reorganization. The degradation then continues in the crystalline regions, leading to an increasing loss of mass. The more accessible a polymer is to water, i.e., the more hydrophilic it is, the more susceptible it is to hydrolysis and, therefore, faster degradation. Our hypothesis is that the amorphous and hydrophilic PGMA will not be susceptible to main-chain hydrolysis than will the hydrophobic PMDO. Larger GMA blocks in the 2803

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Figure 3. Residual mass of MDO80 (■), MDO65 (●) and PCL (▲) during 133 days of hydrolysis (left) in deionized water at 37 °C, and (right) in PBS at 37 °C.

Figure 4. pH changes of MDO80 (■), MDO65 (●) and PCL (▲) during 133 days of hydrolysis (left) in deionized water at 37 °C, and (right) in PBS at 37 °C.

weight falls below a certain value, i.e., is small enough, that the oligomers become water-soluble and able to diffuse to the surrounding medium. For example, the residual molecular weight for MDO80 degraded in PBS at 37 °C for 28 days decreased only approximately 50%, whereas approximately 85% of the original mass was retained. This large difference between the residual molecular weight and mass clearly demonstrates the importance of determining the molecular weight and the mass loss when performing degradation studies.39 Changes in pH. A decrease in pH during hydrolysis is obtained because of the formation of carboxylic acid-terminated degradation products. A rapid decrease in pH can be expected because the hydrolytic degradation of aliphatic polyesters is autocatalyzed by the lower pH, which in turn is due to the carboxylic end-groups that are generated by chain-scission of the ester bonds.40 This phenomenon is also a major concern because the accumulation of acidic degradation products will result in a more rapid decrease in pH in the surrounding milieu. During the first 7 days of hydrolysis in deionized water, the pH decreased from 7.5 to 7.0 for both copolymers; thereafter, it declined more slowly (Figure 4). After 133 days, at the end of the study, a pH value of 6.5 was measured, which confirmed that there was no rapid release of large amounts of degradation products. The extremely slow change in pH during hydrolysis in PBS can be explained by the buffering capacity of PBS and because the oligomeric degradation products are larger than and will not influence the pH to the same extent as monomeric acids.41 These results demonstrate that all of the MDO-based materials tested degrade without a rapid release of acidic degradation products and without producing a substantial lowering of the pH that might diminish the biocompatibility of the material.

PBS. This difference can be explained by the buffering capacity of PBS, which neutralizes the acidic degradation products and thereby suppresses the autocatalytic degradation process. Deionized water lacks this buffering capacity, and the acid formed promotes further hydrolysis. The molecular weight of MDO80 remaining in deionized water after 91 days was reduced to approximately the same extent (50%) as that in PBS after 133 days. This fact should be considered when analyzing the migration of oligomers. After 133 days of hydrolysis in deionized water, approximately 40% of the initial molecular weight of both of the copolymers was retained. When hydrolyzed in deionized water, the degradation rates of MDO80 and MDO65 were almost the same compared with the rate in PBS, where there was an observable difference. By increasing the amount of functional groups in the copolymer, the release rate of degradation products in a body-like environment would subsequently decrease. The hydrolysis was also followed by observing the changes in mass after different degradation times, as shown in Figure 3. After an initial loss of mass, there was a steady loss, and approximately 80% of the copolymers remained after 133 days. For the homopolymer of PCL, there was a negligible mass loss at all during the hydrolysis time. The likely reason for this small loss is because water absorption is hindered in PCL due to its hydrophobic and semicrystalline character. An amorphous character of the copolymers will instead facilitate the degradation. The molecular weight is a good indicator of polymer degradation than is the mass loss because more significant changes are observed at an earlier stage. This can be explained by the bulk degradation mechanism. Cleavage of the chain will occur continuously during the degradation, beginning immediately upon contact with water, but it is not until the molecular 2804

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Figure 5. Positive ESI-MS spectra of water-soluble degradation products after 133 days of hydrolysis in deionized water at 37 °C. Left: MDO80. Right: MDO65.

Degradation Products. The migration of monomeric and oligomeric degradation products into the reaction milieu after each hydrolysis time was analyzed by ESI-MS. As a result of the mechanism of hydrolysis, the oligomers obtained from the copolymers and PCL were expected to be terminated primarily with hydroxyl and carboxyl end groups. No monomeric or oligomeric degradation products were detected from either MDO80 or MDO65 until after 49 days of degradation, although a drop in the pH, molecular weight, and mass were observed. Thereafter, the amount of degradation products increased relatively slowly. This observation can be explained by the fact that the released oligomers were not detected by ESI-MS until they had a very specific size, below a mass (m/z) of 2000. This has been observed previously during the degradation of PCL.42 One supplementary explanation could be attributed to the degradation medium, i.e., deionized water. Thus, an inaccurate result could be obtained if the chains are not completely dissolved in the medium. The continuous increase in released oligomeric degradation products in combination with the slow reduction in molecular weight suggests that the hydrolyzable ester groups of MDO might be situated in the terminal regions of the polymeric chain. As the formation of water-soluble degradation products increased, a shift of the degradation product patterns toward shorter oligomers was seen in the spectra with more degraded samples. Figure 5 shows the positive ESI-MS spectra in the mass range m/z 50−500 containing the water-soluble compounds that migrated from MDO80 and MDO65 to water after 133 days of hydrolysis. All of the oligomers detected form positively charged adducts with sodium. Several low-intensity peaks at m/z 76.80, 98.87, 211.07, and 360.87 and some higher intensity peaks at m/z 113.93 and 177.00 were observed in both copolymers. The peak at m/z 113.93 correspond to the cyclic monomeric product of MDO and the peaks at m/z 155 (Figure 5, left) and m/z 132.8 (Figure 5, right) might correspond to 6-hydroxyhexanoic acid (HHA). HHA is a common hydroxyl acid that is formed as a result of the random hydrolytic scission of ester bonds in aliphatic polyesters, such as PCL. Other peaks in the spectra are either of uncertain origin or are oligomeric products originating from the wide range of different products from backbiting reactions that might occur during polymerization. Indirect Cell Toxicity Test. Ester-based polymers are extensively used for biomedical purposes due to their biocompatibility and biodegradability.3,43 The homopolymer of GMA has no degradable units in the backbone, but if

copolymerized with MDO, it will acquire a degradable backbone. Hydrolysis of esters generates carboxylic acid groups, which can produce harmful interactions with cells. It has been shown that cell attachment and proliferation can be supported very well on polyester scaffolds, though the pH can reach a value of 6.2 during degradation.44 It has also been observed when using L-929 mouse fibroblast cells that there is no increase in the inhibition of cell growth until the pH reached a value below 6.5.24 Here, the pH reached a value of 6.5 after 133 days of degradation for all of the materials tested. Therefore, indirect cytotoxicity tests are required to confirm the innocuous nature of these materials. The potential toxicity of the different copolymers was evaluated using human bone marrow mesenchymal stem cells (BM-MSC). MSCs maintain an undifferentiated and stable phenotype over many generations in vitro and are progenitors for different types of somatic cells, such as osteocytes, chondrocytes and adipocytes. The MSCs were chosen because of their strong potential to be used as seed cells in tissue engineering and regenerative medicine due to their multiplelineage differentiation potential.45−47 To evaluate in vitro cytotoxicity, a methylthiazol tetrazolium (MTT) assay was performed on extracts of four different MDO/GMA-based pristine copolymers that were produced from a feed ratio of MDO ranging from 65 mol % to 80 mol %. The MTT assay reflects mitochondrial metabolism, and the MTT activity is measured by evaluating the mitochondrialdependent conversion of thiazolyl blue tetrazolium bromide to formazan. As shown in Figure 6, the MTT assay demonstrated good cell viability after exposure to the four different copolymers that were tested. The MTT activity was higher on all copolymers than it was on tissue culture polystyrene (TCPS), which served as a control. These results indicate that greater cell proliferation is obtained on the copolymers in comparison to TCPS. A clear trend was observed for all copolymers: MDO65 had a higher rate of cell proliferation than did the others. This higher rate can be due to the more hydrophilic nature of this copolymer; it has been shown that the hydrophobic/hydrophilic nature of the biomaterials affects the differentiation of mesenchymal stem cells.48,49 The cell proliferation decreased between 24 and 72 h, in part due to the cells’ inability to attach to the surface. This is a common problem that has challenged researchers; consequently, most polymers must be subjected to a surface treatment to render the material suitable for cell attachment.50,51 Different coupling techniques have been employed 2805

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greater influence on the hydrolysis rate than did the degradation environment. The molecular weight of the materials decreased significantly faster than did the mass loss, and this effect was observed regardless of the environment. The cytotoxicity of the MDO-GMA-based materials with different compositions was determined using a MTT assay. None of the materials exhibited any obvious cytotoxicity toward the cells and were therefore considered biocompatible. The incorporation of functional groups into the backbone of GMA through the radical copolymerization of MDO resulted in a functional, amorphous, and degradable polymer. This approach may prove to be a valuable tool for controlling the degradation rate of the polymers and may consequently have great potential for use in future biomedical applications.

Figure 6. Cell viability (MTT assay) after incubating cells on four different copolymers: MDO65, MDO70, MDO75 and MDO80. Each sample was assayed in triplicate. The results are expressed as the OD of supernatants from treated cells. Complete culture medium in TCPS served as the negative control.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]; Tel: +46-8-790 82 74; Fax: +468-20 84 77.

in the past to ensure the covalent binding of, for example, peptides to the surface of the materials. Here, our material is also very promising because of the epoxy functionality, which has been shown to allow the further modification and coupling of bioactive molecules.13 Several transplantation studies with MSCs have used green fluorescent protein (GFP)-positive cells so that they can be easily located in the receiving tissue. To image the GFP autofluorescence of MSCs and to quantify and monitor any potential evidence of toxicity to the cells, a spectrophotometer was used at a wavelength of 570 nm. Most cells were not able to attach to the MDO/GMA materials and were therefore rinsed away after 1 h. After increasing the incubation time to 24 h, some cells could be observed on the materials. Both MDO65 and MDO70 had very strong autofluorescence, which enhanced the images of the cells, as observed in Figure 7. On the other two copolymers, MDO75 and MDO80, cells could be observed, although no greatly enhanced cell activity was observed during this time period compared with the control. These results were in good agreement with the MTT results previously obtained and thus confirmed the innocuous nature of the four different copolymers synthesized in this study.

Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors gratefully acknowledge ERC Advanced Grant PARADIGM, Grant Agreement No. 246776, for financial support of this work. The authors would also like to acknowledge Zhe Xing at the Faculty of Medicine and Dentistry at the University of Bergen for the help with the cytotoxicity tests.

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REFERENCES

(1) Langer, R.; Vacanti, J. P. Science 1993, 260, 920−926. (2) Albertsson, A. C.; Varma, I. K. Biomacromolecules 2003, 4, 1466− 1486. (3) Danmark, S.; Finne-Wistrand, A.; Wendel, M.; Arvidson, K.; Albertsson, A.-C.; Mustafa, K. J. Bioact. Compat. Polym. 2010, 25, 207− 223. (4) Labet, M.; Thielemans, W. Chem. Soc. Rev. 2009, 38, 3484−3504. (5) Kricheldorf, H. R.; Kreisersaunders, I. Makromol. Chem. 1990, 191, 1057−1066. (6) Kobayashi, S.; Uyama, H.; Kimura, S. Chem. Rev. 2001, 101, 3793−3818. (7) Gross, R. A.; Kumar, A.; Kalra, B. Chem. Rev. 2001, 101, 2097− 2124. (8) Bailey, W. J. Polym. J. 1985, 17, 85−95. (9) Bailey, W. J.; Ni, Z.; Wu, S. R. J. Polym. Sci., Part A: Polym. Chem. 1982, 20, 3021−3030. (10) Agarwal, S. Polym. Chem. 2010, 1, 953−964. (11) Jin, S.; Gonsalves, K. E. Macromolecules 1997, 30, 3104−3106. (12) Jin, S.; Gonsalves, K. E. Macromolecules 1998, 31, 1010−1015. (13) Undin, J.; Finne-Wistrand, A.; Albertsson, A. C. Biomacromolecules 2013, 14, 2095−2102.

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CONCLUSIONS The functional MDO-GMA-based copolymers that we synthesized have been demonstrated successfully to have adjustable degradation properties when evaluated under physiological conditions in vitro. The pH of the degradation media decreased with increasing hydrolysis time but without any rapid release of degradation products that might cause inflammatory responses in the body at the site of implantation. The degradation rate was faster in water than it was in PBS, although the amount of incorporated ester units in the backbone and the amorphous nature of the polymers had a

Figure 7. Quantification of cell proliferation by imaging GFP autofluorescence of the MSCs (green) at a wavelength of 570 nm. The scale bars represent 100 μm. 2806

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Biomacromolecules

Article

(14) Undin, J.; Illanes, T.; Finne-Wistrand, A.; Albertsson, A. C. Polym. Chem. 2012, 3, 1260−1266. (15) Undin, J.; Plikk, P.; Finne-Wistrand, A.; Albertsson, A. C. J. Polym. Sci., Part A: Polym. Chem. 2010, 48, 4965−4973. (16) Mathisen, T.; Lewis, M.; Albertsson, A. C. J. Appl. Polym. Sci. 1991, 42, 2365−2370. (17) Chu, C. C.; Campbell, N. D. J. Biomed. Mater. Res. 1982, 16, 417−430. (18) Pistner, H.; Stallforth, H.; Gutwald, R.; Muhling, J.; Reuther, J.; Michel, C. Biomaterials 1994, 15, 439−450. (19) Li, S. M.; Garreau, H.; Vert, M. J. Mater. Sci.: Mater. Med. 1990, 1, 198−206. (20) Li, S. M.; Garreau, H.; Vert, M. J. Mater. Sci. - Mater. Med. 1990, 1, 123−130. (21) Li, S. M.; Garreau, H.; Vert, M. J. Mater. Sci.: Mater. Med. 1990, 1, 131−139. (22) Lofgren, A.; Albertsson, A. C. J. Appl. Polym. Sci. 1994, 52, 1327−1338. (23) Ignatius, A. A.; Claes, L. E. Biomaterials 1996, 17, 831−839. (24) Cordewener, F. W.; van Geffen, M. F.; Joziasse, C. A. P.; Schmitz, J. P.; Bos, R. R. M.; Rozema, F. R.; Pennings, A. J. Biomaterials 2000, 21, 2433−2442. (25) Olsen, P.; Undin, J.; Odelius, K.; Albertsson, A.-C. Polym. Chem. 2014, 5, 3847−3854. (26) Undin, J.; Finne-Wistrand, A.; Albertsson, A.-C. Biomacromolecules 2013, 14, 2095−2102. (27) Hoglund, A.; Hakkarainen, M.; Albertsson, A.-C. J. Macromol. Sci., Part A: Pure Appl. Chem. 2007, 44, 1041−1046. (28) Malberg, S.; Hoglund, A.; Albertsson, A.-C. Biomacromolecules 2011, 12, 2382−2388. (29) Wanamaker, C. L.; Tolman, W. B.; Hillmyer, M. A. Biomacromolecules 2009, 10, 443−448. (30) Agatemor, C.; Shaver, M. P. Biomacromolecules 2013, 14, 699− 708. (31) Hoglund, A.; Odelius, K.; Albertsson, A.-C. ACS Appl. Mater. Interfaces 2012, 4, 2788−2793. (32) Hakkarainen, M.; Hoglund, A.; Odelius, K.; Albertsson, A.-C. J. Am. Chem. Soc. 2007, 129, 6308−6312. (33) Hoglund, A.; Malberg, S.; Albertsson, A.-C. Macromol. Biosci. 2012, 12, 260−268. (34) Olsen, P.; Borke, T.; Odelius, K.; Albertsson, A.-C. Biomacromolecules 2013, 14, 2883−2890. (35) Plikk, P.; Tyson, T.; Finne-Wistrand, A.; Albertsson, A. C. J. Polym. Sci., Part A: Polym. Chem. 2009, 47, 4587−4601. (36) Bailey, W. J.; Zhou, L. L. Tetrahedron Lett. 1991, 32, 1539− 1540. (37) Hakkarainen, M., Aliphatic polyesters: Abiotic and biotic degradation and degradation products. In Degradable Aliphatic Polyesters, Albertsson, A. C., Ed.; Springer-Verlag Berlin: Berlin, 2002; Vol. 157, pp 113−138. (38) Ananthalakshmi, N. R.; Wadgaonkar, P. P.; Sivaram, S.; Varma, I. K. J. Therm. Anal. Calorim. 1999, 58, 533−539. (39) von Burkersroda, F.; Schedl, L.; Gopferich, A. Biomaterials 2002, 23, 4221−4231. (40) Pitt, C. G.; Gu, Z. W. J. Controlled Release 1987, 4, 283−292. (41) Odelius, K.; Hoglund, A.; Kumar, S.; Hakkarainen, M.; Ghosh, A. K.; Bhatnagar, N.; Albertsson, A. C. Biomacromolecules 2011, 12, 1250−1258. (42) Hakkarainen, M.; Adamus, G.; Hoglund, A.; Kowalczuk, M.; Albertsson, A.-C. Macromolecules 2008, 41, 3547−3554. (43) Idris, S. B.; Arvidson, K.; Plikk, P.; Ibrahim, L.; Finne-Wistrand, A.; Albertsson, A.-C.; Bolstad, A. I.; Mustafa, K. J. Biomed. Mater. Res., Part A 2010, 94A, 631−639. (44) Xie, S.; Zhu, Q.; Wang, B.; Gu, H.; Liu, W.; Cui, L.; Cen, L.; Cao, Y. Biomaterials 2010, 31, 5100−5109. (45) McKay, R. Nature 2000, 406, 361−364. (46) Pittenger, M. F.; Mackay, A. M.; Beck, S. C.; Jaiswal, R. K.; Douglas, R.; Mosca, J. D.; Moorman, M. A.; Simonetti, D. W.; Craig, S.; Marshak, D. R. Science 1999, 284, 143−147.

(47) Zuk, P. A.; Zhu, M.; Ashjian, P.; De Ugarte, D. A.; Huang, J. I.; Mizuno, H.; Alfonso, Z. C.; Fraser, J. K.; Benhaim, P.; Hedrick, M. H. Mol. Biol. Cell 2002, 13, 4279−4295. (48) Curran, J. M.; Chen, R.; Hunt, J. A. Biomaterials 2006, 27, 4783−4793. (49) Mattioli-Belmonte, M.; Biagini, G.; Lucarini, G.; Virgili, L.; Gabbanelli, F.; Amati, S.; Cecchet, F.; Albertsson, A. C.; Finne, A.; Andronova, N. J. Bioact. Compat. Polym. 2005, 20, 509−526. (50) Shin, H.; Jo, S.; Mikos, A. G. Biomaterials 2003, 24, 4353−4364. (51) Curtis, A. S. G.; Forrester, J. V.; McInnes, C.; Lawrie, F. J. Cell Biol. 1983, 97, 1500−1506.

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