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DOI: 10.1177/0883911512440536jbc.sagepub.com
JOURNAL OF
Bioactiveand
CompatiblePolymers
Poly(butylene/diethylene glycol succinate) multiblock copolyester as a candidate biomaterial for soft tissue engineering: Solid-state properties, degradability, and biocompatibility
Chiara Gualandi1,2, Michelina Soccio3, Marco Govoni4, Sabrina Valente5, Nadia Lotti3, Andrea Munari3, Emanuele Giordano4, Gianandrea Pasquinelli6 and Maria Letizia Focarete1
AbstractA multiblock bioresorbable copolyester, poly(butylene/diethylene glycol succinate), was synthesized by reactive blending, and it was used, together with the corresponding poly(butylene succinate) homopolymer, to form films and to fabricate biomimetic electrospun scaffolds. The poly(butylene/diethylene glycol succinate) scaffold had a more pronounced elastomeric behavior than poly(butylene succinate). It also underwent hydrolytic degradation faster than poly(butylene succinate) since the incorporated diethylene glycol succinate units rendered the copolymer more
1 Department of Chemistry “G Ciamician” and National Consortium of Materials Science and Technology (INSTM, RU Bologna), University of Bologna, Bologna, Italy
2 Advanced Applications in Mechanical Engineering and Materials Technology Interdepartmental Center for Industrial Research (CIRI MAM), University of Bologna, Bologna, Italy
3Department of Civil, Environmental, and Materials Engineering (DICAM), University of Bologna, Bologna, Italy4 Department of Biochemistry ‘G. Moruzzi’ Laboratory of Cellular and Molecular Engineering and National Institute for Cardiovascular Research, University of Bologna, Cesena, Italy
5Anaestesiological and Surgical Sciences, University of Bologna, Bologna, Italy6Clinical Department of Radiological and Histocytomorphological Sciences, University of Bologna, Bologna, Italy
Corresponding author:Maria Letizia Focarete, Department of Chemistry “G Ciamician” and National Consortium of Materials Science and Technology (INSTM, RU Bologna), University of Bologna, via Selmi 2, 40126 Bologna, Italy Email: [email protected]
440536 JBC27310.1177/0883911512440536Gualandi et al.Journal of Bioactive and Compatible Polymers2012
Article
Gualandi et al. 245
hydrophilic than poly(butylene succinate). The films degraded faster than electrospun samples due to the autocatalytic effect of carboxylic end-groups. The biodegradable poly(butylene/diethylene glycol succinate) scaffold supported the growth and preserved the cardiac phenotype markers of H9c2 cells, demonstrating its potential utility in soft tissue engineering applications.
Tissue engineering for regenerative medicine uses natural or synthetic polymers that, when formed into appropriate scaffolds, are able to accommodate the functional requirements of the seeded cells. Soft tissue engineering scaffolds must possess suitable mechanical properties, since in vivo cell functions and extracellular matrix synthesis are largely regulated by mechanical signals.1
Commercial biodegradable thermoplastic materials typically employed in tissue engineering, such as poly(lactic acid), poly(glycolic acid), and their copolymers, are often too rigid to be replacements for soft tissue, with elastic moduli, E, in the range 1–10 GPa.2 Materials with lower elastic moduli are required for soft tissue engineering applications, with E in the range 0.1–100 MPa, depending on the tissue.2–4 Progress in this field has been limited by the scarcity of biode-gradable polymeric material possessing elastomeric mechanical properties, which would enable them to withstand cyclic deformation and to transfer mechanical stress to cultured cells, prerequi-sites for candidate scaffolds. Development of new elastomeric materials with controllable degrada-tion rates has indeed increased significantly in the last decade.5 Biodegradable polyurethane,6,7 trimethylene carbonate–based polymers,8 caprolactone copolymers,9,10 and sebacate-based poly-mers11 have been proposed for these applications.
It is not only the design of new elastomeric polymer systems but also the selection of appropri-ate polymer synthetic routes that is a key issue. Reactive blending, which occurs when two poly-mers are melt mixed together,12 is a simple and versatile solvent-free method to synthesize new copolymers displaying final properties that can be favorably modulated, depending on the type, relative amount, and distribution of the comonomeric units along the polymer chain.
We have previously demonstrated the possibility of obtaining a wide range of new copolymers by mixing, in the molten state, different types of polyesters.13,14 In particular, new poly(butylene/diethylene glycol succinate) (P(BS-b-DGS)) multiblock copolymers were synthesized starting from the parent homopolymers—poly(butylene succinate) (PBS) and poly(diethylene glycol suc-cinate) (PDGS)—in the presence of Ti(OBu)4 as a catalyst.14 PBS is a biodegradable aliphatic polyester with mechanical properties that make it a suitable alternative to nondegradable polyole-fins for packaging applications.15,16 Recently, PBS has also been tested as a biomaterial, and its biocompatibility has been demonstrated.17,18 However, while PBS is suited to applications requir-ing high load resistance, it is too rigid to be exploited for soft tissue engineering applications. PDGS displays a molecular structure very similar to that of PBS, differing only in the presence of an ether oxygen atom in the glycol subunit. As previously reported,13 the introduction of diethylene glycol succinate (DGS) into the PBS chain significantly changes the phase structure of the material and, consequently, its thermal properties, in a way that depends on the arrangement of DGS units along the polymer chain. It was found that, in the obtained multiblock copolymers, the block length remarkably affects the capability of the material to crystallize. It would therefore be anticipated
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that the introduction of DGS units into the PBS chain could also change its mechanical properties, the latter being strictly dependent on polymer morphology.
Taking into account previous thermal characterization results, in this work we prepared a multi-block copolymer (P(BS-b-DGS)), by melt mixing PBS and PDGS, with a block length specifically selected to allow the development of a small amount of rigid crystal phase coexisting with a soft amorphous rubbery phase. This is an important factor in the realization of elastomeric scaffolds with structural stability. The aim of this work was to evaluate how the introduction of DGS units modifies thermomechanical properties, wetting behavior, and hydrolytic degradation profile of PBS, and to check the biocompatibility of cell culture frameworks fabricated with these new mate-rials. The latter experiments were performed by using the H9c2 cell line, derived from embryonic rat heart, employed as a cardiac muscle model. In view of producing scaffolds for soft tissue engi-neering, PBS and P(BS-b-DGS) scaffolds resembling the morphology of the fibrous extracellular matrix component were prepared by electrospinning. Solid-state properties, degradability, and bio-compatibility of the electrospun scaffolds were evaluated and compared to those of samples in form of film.
Materials and methods
Materials
Dimethylsuccinate (DMS), 1,4-butanediol (BD), diethylene glycol (DG), and titanium tetrabutox-ide (TBT) (Sigma–Aldrich Co., Milan, Italy) were reagent grade products; DMS and BD were used as supplied, whereas DG and TBT were distilled before use. Chloroform, dichloromethane (DCM), 2-chloroethanol (CE), and ethanol (EtOH) were purchased from Sigma–Aldrich Co. and were used without any further purification.
Synthesis of PBS and PDGS
The syntheses of PBS and PDGS were performed starting from the monomers according to the usual two step procedure for polyester synthesis as follows. DMS (100.0 g, 0.684 mol) and BD (73.99 g, 0.821 mol) or DG (87.12 g, 0.821 mol), according to the polymer to be synthesized, were introduced in a 250-mL three-neck round-bottom flask, provided with a mechanical stirring, a torque meter, and a condenser. TBT (about 0.2 g of TBT/kg of polymer) was added to the reagents, and the flask was placed in an oil bath at 150°C. In the first stage, under pure nitrogen flow, the temperature was increased to 180°C and maintained until no significant increment in distillate volume was observed (120 min from the catalyst addition). The condenser was removed, and the reactor was connected to a liquid nitrogen cooling trap. In the second stage, the reaction tempera-ture was increased to 210°C, and a dynamic vacuum was slowly applied. The reaction was con-ducted at 0.1 mbar until no increase in torque signal was observed. Prior to characterization, the obtained polymers were carefully purified by dissolution in chloroform and reprecipitation in methanol. The purification procedure was repeated three times. Finally, the samples were kept under vacuum at room temperature (RT) for several days in order to remove the residual solvent.
Synthesis of P(BS-b-DGS) block copolyester
Block P(BS-b-DGS) copolyester was obtained by melt-mixing PBS and PDGS without purifica-tions. The two homopolymers were mixed in a 1:1 molar ratio in a 200-mL glass reactor at 225°C,
Gualandi et al. 247
under a nitrogen atmosphere to prevent hydrolytic degradation. During the process, the sample was taken out of the reactor after 30-min reaction time and cooled in air. Copolymer synthesis was cata-lyzed by the residual TBT, introduced during the homopolymerization of PBS and PDGS. The copolymer was purified according to the procedure described above for the homopolymers.
Film and scaffold preparation
Films of PBS homopolymer and of P(BS-b-DGS) copolymer were obtained by compression molding the polymers between Teflon® plates, with an appropriate spacer, at 150°C and 140°C, for PBS and P(BS-b-DGS), respectively, for 1 min under a pressure of 2 t/m2 (laboratory press, Carver C12, Carver Inc., Indiana, USA). The obtained films (0.2 mm thick) were labeled PBS-f and P(BS-b-DGS)-f.
Electrospun mats were fabricated by using an electrospinning apparatus constructed in-house, comprised of a high-voltage power supply (SL 50 P 10/CE/230; Spellman, New York, USA), a syringe pump (KD Scientific 200 series, Massachusetts, USA), a glass syringe, and a stainless-steel blunt-ended needle (inner diameter = 0.84 mm) connected with the power supply electrode and a grounded aluminum plate-type collector (7 cm × 7 cm). Polymer solution was dispensed through a Teflon tube to the needle, which was placed vertically on the collecting plate. PBS and P(BS-b-DGS) were individually dissolved in a mixed solution of DCM and CE (80:20, vol.%) at a concentration of 17% (w/v) and 22% (w/v), respectively. Electrospun scaffolds were produced at RT and a relative humidity of 40% ± 5%. For both polymers, the solution was dispensed at a con-stant rate of 0.01 mL/min, and the distance between the needle and the collector was 15 cm. Voltages of +14 kV and of +20 kV were used for PBS and P(BS-b-DGS), respectively. Electrospun mats, labeled PBS-es and P(BS-b-DGS)-es, were kept under vacuum over P2O5 at RT overnight in order to remove residual solvents.
Instrumental methods
Polymer structure and composition were determined by means of proton nuclear magnetic reso-nance (1H-NMR) spectroscopy. The samples were dissolved (15 mg/mL) in deuterated chloroform solvent with 0.03 vol.% tetramethylsilane added as an internal standard. The measurements were carried out at RT, employing a Varian Inova 400-MHz instrument (Agilent Technologies, USA). The molecular weights were obtained by gel-permeation chromatography (GPC) at 30°C using a 1100 HPLC system (Agilent Technologies, USA) equipped with PLgel 5-µm MiniMIX-C column (Agilent Technologies, USA) (length = 250 mm, internal diameter = 4.6 mm). A refractive index was employed as detector. In all cases, chloroform was used as eluent with a 0.3 mL/min flow and sample concentrations of about 2 mg/mL. A molecular weight calibration curve was obtained with polystyrene standards in the range of molecular weight 2000–100,000 g/mol.
Differential scanning calorimetry (DSC) measurements were carried out using a TA Instruments’ Q100 DSC (Delaware, USA) equipped with a liquid nitrogen cooling system (LNCS) accessory. DSC scans were performed from −100°C to 180°C in helium atmosphere. A rate of 20°C/min was used during heating scans, whereas the cooling scans were performed at a rate of 10°C/min. The glass transition temperature (Tg) was taken at half-height of the glass transition heat capacity step, and the melting temperature (Tm) was taken at the peak maximum of the melting endotherm.
Scanning electron microscopic (SEM) observations were carried out using a Philips 515 SEM (FEI, Eindhoven, The Netherlands) at an accelerating voltage of 15 kV, on samples sputter coated with gold. The distribution of fiber diameters was determined through the measurement of approxi-mately 250 fibers by means of an acquisition and image analysis software (EDAX Genesis, New York, USA), and the results were given as the average diameter ± standard deviation.
248 Journal of Bioactive and Compatible Polymers 27(3)
Wide angle X-ray scattering (WAXS) measurements were carried out at RT with a PANalytical X’Pert PRO diffractometer (Almelo, The Netherlands) equipped with an XCelerator detector (Almelo, The Netherlands). Cu anode was used as the X-ray source (λ1 = 0.15406 nm, λ2 = 0.15443 nm). The amorphous and crystalline contributions were calculated by fitting method, using the Fityk program. The degree of crystallinity (χc) was evaluated as the ratio of the crystalline peak areas to the total area under the scattering curve.
Stress–strain measurements were performed at RT with an Instron 4465 (Instron, Massachusetts, USA) tensile testing machine on rectangular films (5 mm wide and 0.2 mm thick) and on rectan-gular sheets cut from electrospun mats (5 mm wide and 20 ÷ 50 µm thick, measured by a microcali-per). The gauge length was 20 mm, and the crosshead speed was 0.5 mm/min. In order to improve electrospun sample handling during mechanical test, the procedure described by Huang et al. was applied.19 Load-displacement curves were obtained and converted to stress–strain curves. Tensile elastic modulus was determined from the initial linear slope of the stress–strain curve. At least six replicate specimens were run for each sample, and the results were provided as the average value ± standard deviation.
Static contact angle measurements were performed on polymer films using a KSV CAM101 (KSV, Espoo, Finland) instrument at ambient conditions, recording the side profiles of deionized water drops for image analysis. Five drops were observed on different areas for each film, and contact angles were reported as the average value ± standard deviation.
Hydrolytic degradation
Hydrolytic degradation studies were carried out on PBS and P(BS-b-DGS), using both films and electrospun samples. Prior to degradation experiments, each specimen was dried over P2O5 under vacuum at RT for 2 days, then weighed to determine initial sample weight. In order to ensure com-plete wetting, electrospun mats were prewetted in ethanol for 15 min; the ethanol was then replaced by deionized water by means of repeated rinses. Films and wet electrospun samples were immersed in phosphate buffered solution (0.1 M, pH = 7.4) and incubated in a SBS30 Stuart Scientific shak-ing bath (Redhill, UK) at 37°C and 50 revs/min. The buffer solution was periodically changed to keep the pH constant for the entire duration of the degradation experiments. At various intervals, triplicate specimens of each sample were recovered from the bath, repeatedly washed with deion-ized water, and dried over P2O5 under vacuum for 2 days to constant weight. The mass loss was determined gravimetrically by comparing the dry weight remaining at a specific time with the initial sample weight. The molecular weight of each recovered sample was determined by GPC, and the changes in thermal properties were evaluated by comparing DSC data of the initial samples with those of the degraded ones. 1H-NMR analysis was performed to detect changes in the molecu-lar composition of P(BS-b-DGS) samples during the hydrolysis experiments. Samples were labeled as PBS-f-x, PBS-es-x, P(BS-b-DGS)-f-x, and P(BS-b-DGS)-es-x, where x indicates the days of buffer exposure.
Indirect cytotoxicity
Indirect cytotoxicity evaluations of PBS and P(BS-b-DGS) were performed in accordance with the ISO10993-5 international standard for biological evaluation of medical devices. Both polymers (in film form) were sterilized in a laminar flow culture hood by immersion in 70% ethanol for 15 min, followed by repeated washes in phosphate buffered saline containing 100 U/mL penicillin/strepto-mycin and 0.2% fungizone (Sigma–Aldrich Co., Milan, Italy). To obtain culture medium contain-ing potential polymer extracts, samples (5 mg polymer/1 mL medium) were kept for 24 h in
Gualandi et al. 249
Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS), 2 mM of l-glutamine, and 100 U/mL of penicillin/streptomycin, at 37°C in a humidified atmosphere containing 5% CO2. The resulting media were filtered through a 0.2 µm porosity nitrocellulose filter before administration to embryonic rat cardiac H9c2 cells, obtained from the European Collection of Cell Cultures (ECACC). Cells were seeded in a 12-well culture plate (2.5 × 104 cells/well) in standard DMEM to allow their attachment and were quantified by resazurin (i.e. alamarBlue®) fluorescence assay (Invitrogen, Monza, Italy). After 48 h, the culture medium was discarded, the polymer-extract media were added to the wells, and the cells were further incubated for 24 h. At the end of this incubation time, alamarBlue assay was performed again for cytotoxicity screening. The alamarBlue fluorescence (Ex/Em = 540/590 nm) was read in a Wallac Victor2 multilabel multiplate reader (PerkinElmer, Massachusetts, USA). Three separate experiments (n = 3), two replicates each, were performed. The signal obtained from cells cultured in DMEM was used as the negative control, while a cytotoxic response (positive control) was obtained by addition of 1 mM H2O2 for 120 min to cells (48 h after cell seeding). Fluorescence values (mean ± standard error of the mean) were calculated, and the unpaired t-test was used to evaluate statistical differences between samples.
Cell adhesion and proliferation
Evaluation of both cell adhesion and cell proliferation was performed in accordance with ISO10993-5. Polymeric substrates (in both film and scaffold forms) were fixed at a plastic ring (Tecaflon®, internal diameter = 17 mm and external diameter = 20 mm) by using silicone rubber (RTV 108Q; GE Silicones, North Carolina, USA), in order to obtain a cell leakage–proof well with the substrate fixed at the bot-tom. Sterilization (ring plus polymeric substrate) was performed in a laminar flow culture hood by immersion in 70% ethanol for 15 min, followed by washings in phosphate buffered saline containing 200 U/mL of penicillin/streptomycin and 0.2% fungizone. In order to evaluate cell adhesion and pro-liferation on different polymer substrates, 2.5 × 104 H9c2 cells in 1 mL of DMEM were seeded onto PBS and P(BS-b-DGS) films and electrospun scaffolds, assembled with the Tecaflon rings as described above and placed in standard culture multiplates. The amount of viable cells was quantified at day 1, 7, and 14 with the alamarBlue fluorescence assay. Control signal was acquired from H9c2 cells cul-tured in standard polystyrene wells (Sarstedt, Numbrecht, Germany). Three separate experiments (n = 3), two replicates each, were performed. Fluorescence values (mean ± standard error of the mean) were calculated, and the unpaired t-test was used to evaluate statistical differences between samples. In selected samples intended for further microscopy analysis, cells were grown up to 35 days.
SEM
After 7, 14, and 35 days in culture, the cell-seeded substrates were washed with 0.15 M phosphate buffer to remove culture medium and then fixed in 2.5% phosphate buffered glutaraldehyde (TAAB Laboratories, Aldermaston, UK) at pH 7.4 overnight at 4°C. Afterward, the samples, carefully removed from the Tecaflon ring with a scalpel, were washed, postfixed in 1% buffered osmium tetroxide, rinsed in distilled water, and dehydrated in a graded series of ethanol for 15 min each step at RT. Drying was performed in a solution of 50% absolute ethanol/50% hexamethyldisilazane (HMDS; Fluka Analytical, Sigma–Aldrich Co., Milan, Italy), followed by pure HMDS for 30 min at RT and finally air-dried. Cell-seeded substrates were mounted on aluminum stubs with double-sided adhesive tape, coated with a 10-nm-thick gold film using a sputtering device and observed with a Philips 505 SEM at 15 kV.
250 Journal of Bioactive and Compatible Polymers 27(3)
Immunohistochemistry
After 35 days of culture with H9c2 cells, PBS-es and P(BS-b-DGS)-es scaffolds were fixed in 10% buffered formalin and embedded in paraffin20; 4-µm-thick sections were used for histological and immunohistochemical analysis. For a histological analysis, the sections were stained with hema-toxylin and eosin (H&E). Paraffin-embedded sections were used to perform immunohistochemical analysis on H9c2 cells that were not made to adhere to any substrate and on H9c2 cells after adhe-sion on PBS-es and P(BS-b-DGS)-es scaffolds. Briefly, the samples were dewaxed, rehydrated through ethanol (from 100% to 70%), rinsed in distilled water, and subjected to antigen retrieval treatment using citrate buffer (pH 6), at 120°C and 1 atm for 21 min. After cooling and washing, the endogenous peroxidase activity was quenched with 3% H2O2 in absolute methanol in the dark for 5 min at RT. Then the sections were processed according to the nonbiotin-amplified method (NovoLink™ Polymer Detection System; Novocastra Laboratories Ltd, Newcastle Upon Tyne, UK) using the manufacturer instructions. The following mouse antibodies (mAbs) were used: anti α-smooth muscle actin (α-SMA) (1:9000, clone 1A4; Sigma–Aldrich Co., Milan, Italy), anti desmin (1:300, clone D33; DakoCytomation, Milan, Italy), and anticardiac troponin I (Tn-I) (1:250, clone 284 (19C7); Abcam, Cambridge, UK); mAbs were diluted in 1% buffered bovine serum albumin (BSA) overnight at 4°C. After counterstaining with Gill’s hematoxylin, samples were viewed in a Leitz Diaplan light microscope connected with a video camera (JVC 3CCD video camera, KY-F55B, Milan, Italy). Digital images were processed using the Image-Pro Plus® 6 pro-gram. Negative control consisted of omission of primary antibody from sections.
Results
The two homopolymers PBS and PDGS were synthesized by polycondensation reaction, while the copolymer P(BS-b-DGS) was obtained by reactive blending of the two homopolymers. At RT, the PBS and P(BS-b-DGS), as prepared, are opaque and light yellow colored solid powders, whereas PDGS is a sticky rubber. Since it is impossible to process the PDGS homopolymer to obtain films and electrospun mats, in this work, we investigated only PBS and P(BS-b-DGS), whose chemical structures are shown in Figure 1. Butylene succinate (BS) and DGS units are very similar, both containing two ester groups along a saturated aliphatic chain. The difference between the two units is the presence in the DGS unit of an ether oxygen atom (in the glycol subunit), which is absent along the BS unit.
Data concerning molecular characterization of PBS and P(BS-b-DGS)—namely, weight aver-age molecular weight (Mw) and molecular weight polydispersity index (PDI), block length, and degree of randomness (b)—are reported in Table 1 together with those of PDGS for comparison.
Figure 1. Chemical structure of (a) PBS and (b) P(BS-b-DGS).PBS: poly(butylene succinate); P(BS-b-DGS): poly(butylene/diethylene glycol succinate).
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All the polymers prepared have similar, relatively high molecular weights. In all cases, the 1H-NMR spectra are consistent with the expected structures (see as an example the 1H-NMR spectrum of P(BS-b-DGS) shown in Figure 2). The copolymer composition, calculated from the relative areas of the 1H-NMR resonance peak of the b aliphatic proton of the butanediol subunit located at 4.11 ppm and of the d protons of the methylene groups of the diethylene diol subunit at 4.25 ppm was close to that of the feed, that is, 50 mol% BS and 50 mol% DGS. It is well known that informa-tion on the arrangement of the comonomeric units in the macromolecular chain may be deduced by the degree of randomness b, which can in turn be determined by 1H-NMR spectroscopy: b is equal to 1 for random copolymers, 2 for alternate copolymers, and around zero for a mixture of two
Table 1. Molecular characterization data obtained by GPC and 1H-NMR analysis
GPC: gel-permeation chromatography; 1H-NMR: proton nuclear magnetic resonance; Mw: weight average molecular weight; PDI: polydispersity index; DGS: diethylene glycol succinate; L: block length; b = degree of randomness; PBS: poly(butylene succinate); P(BS-b-DGS): poly(butylene/diethylene glycol succinate); PDGS: poly(diethylene glycol succinate).aCalculated by GPC analysis.bCalculated by 1H-NMR.
Figure 2. 1H-NMR spectrum of P(BS-b-DGS) and resonance assignments with expansion of interesting aliphatic region between 2.55 and 2.75 ppm.1H-NMR: proton nuclear magnetic resonance; P(BS-b-DGS): poly(butylene/diethylene glycol succinate).
252 Journal of Bioactive and Compatible Polymers 27(3)
homopolymers, while 0 < b < 1 for block copolymers. The calculation of b was carried out accord-ing to the procedure described elsewhere,21 and the value reported in Table 1 indicates that the copolymer under investigation has a multiblock architecture, with an average L of 9 units for both BS and DGS blocks.
The different chemical structure of the polymers under investigation (Figure 1) affects their behavior upon water contact. Measurements performed on flat films provided the following values of water contact angle (WCA)—WCAPBS = 96 ± 3 and WCAP(BS-b-DGS) = 86 ± 4—revealing a slightly higher hydrophilicity of P(BS-b-DGS) with respect to PBS.
Scaffolds of PBS and P(BS-b-DGS) were fabricated by electrospinning, using the instrumental parameters described in the experimental section. SEM micrographs of the electrospun scaffolds at different magnifications are shown in Figure 3. The scaffolds appear to be highly porous mats with microscale interstitial pores. Both polymers yielded fibers free of bead defects, randomly oriented, and possessing submicrometric diameters (PBS fiber diameter distribution = 340 ± 90 nm; P(BS-b-DGS) fiber diameter distribution = 700 ± 410 nm).
DSC was used to investigate the thermal transitions occurring in the polymers, in both films and electrospun mats (see data in Table 2). Both PBS and P(BS-b-DGS) films are semicrystalline, with Tg values below RT. It is also notable that P(BS-b-DGS), both in film and electrospun forms, dis-plays a single Tg indicating the presence of a homogeneous amorphous phase. Above Tg, the DSC heating scans of PBS-f show slight premelting followed by a cold crystallization peak (data not shown), which can be attributed to crystal phase rearrangements, and finally a melting peak (Tm = 113°C). The DSC curve of P(BS-b-DGS)-f displays a broad melting endotherm with a peak (Tm) at 99°C corresponding to the fusion of the crystal phase developed by BS units.13 The significantly
Figure 3. SEM micrographs of (a and b) PBS electrospun scaffold and of (c and d) P(BS-b-DGS) electrospun scaffold. For each scaffold two different magnifications are reported.SEM: scanning electron microscopy; PBS: poly(butylene succinate); P(BS-b-DGS): poly(butylene/diethylene glycol succinate).
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lower value of both Tm and melting enthalpy (ΔHm) in the case of the copolymer indicates the reduced level of crystallinity in P(BS-b-DGS)-f compared to PBS-f. This was also confirmed by WAXS analysis that revealed 59% and 49% of crystallinity degree (χc) for PBS-f and P(BS-b-DGS)-f, respectively.
Data in Table 2 show that thermal properties of films and of the corresponding electrospun mats differ only in the melting enthalpy of the first heating scan, given the lower amount of crystal phase displayed by electrospun samples (χc-PBS-es = 55% and χc-P(BS-b-DGS)-es = 44%, by WAXS) compared to the corresponding films.
The tensile behavior of the polymers investigated is shown in Figure 4, in which load is plotted as a function of displacement. In Table 3, the corresponding mechanical data (elastic modulus, E; stress at break, σb; and deformation at break, εb) of both films and electrospun samples are recorded. As expected, the highly porous electrospun samples were characterized by a lower elastic modulus, were less strong, and displayed higher deformation at break compared with the corresponding samples in form of films. In addition, PBS and P(BS-b-DGS) show different tensile behaviors, in the form of both films and electrospun mat; the P(BS-b-DGS) had lower elastic modulus than PBS, a similar elongation at break, and lower stress at break, indicating that PBS was stronger than P(BS-b-DGS).
PBS and P(BS-b-DGS) samples subjected to hydrolytic degradation in buffered solution at 37°C were recovered at different time intervals, up to approximately 250 days, in order to investi-gate the degradation behavior of the two polymers, both as films and electrospun mats. A macro-scopic inspection of the retrieved samples revealed that, in the case of PBS, neither films nor scaffolds showed evident changes in shape or color within the degradation time range investigated.
Table 2. Calorimetric data of films and electrospun mats of PBS and P(BS-b-DGS) before and after selected hydrolytic degradation intervals
PBS: poly(butylene succinate); P(BS-b-DGS): poly(butylene/diethylene glycol succinate); f: films; es: electrospun mats.aHeating scan performed after the cooling scan.
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On the contrary, the P(BS-b-DGS) samples, especially those in form of films, became more fragile as the degradation experiment progressed. SEM analysis did not reveal any microscopic change in the morphology of any of the investigated samples. In Figure 5(a), the percentage of residual weight (mres%) is reported as a function of time, while in Figure 5(b), the percentage of residual weight average molecular weight (Mw-res%) is plotted as a function of time. It is evident that PBS, both in form of film and fibrous scaffold, did not lose weight in the time interval investigated, whereas the weight of P(BS-b-DGS) samples significantly decreased in the course of degradation. The weight loss was more evident in the case of P(BS-b-DGS)-f: after 235 days of buffer exposure, it lost almost 20% of its initial weight.
All samples under investigation underwent a significant decrease of Mw, which commenced at the very beginning of the experiment (Figure 5(b)). At the end of the experiment, the molecular weight of each sample was less than 40% of its initial value. It is evident that, for both polymers, the film underwent a faster decrease of polymer molecular weight than the corresponding electrospun scaffold. In addition, the change of molecular weight seems not to be affected by the type of polymer investigated. It was observed that the residual number average molecular weight (Mn-res%) dis-played the same trend of Mw-res%.
In order to get information about hydrolysis mechanism of P(BS-b-DGS), 1H-NMR measure-ments were performed on the degraded retrieved samples of P(BS-b-DGS)-f and P(BS-b-DGS)-es. DGS content (mol%) as a function of degradation time is shown in Figure 6. A decrease in DGS content was observed as degradation proceeded; this was more consistent for the film than for the electrospun sample. After 235 days of exposure to water, DGS content decreased from the initial
Table 3. Mechanical data of PBS and P(BS-b-DGS) in the form of both films and electrospun mats
Figure 4. Stress–strain curves of PBS and P(BS-b-DGS) in the form of (a) films and (b) electrospun mats.PBS: poly(butylene succinate); P(BS-b-DGS): poly(butylene/diethylene glycol succinate); f: film; es: electrospun mats.
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50 mol% of the nondegraded samples down to 41% and 47 mol% (film and electrospun mats, respectively).
DSC analyses of degraded samples at some selected times of buffer exposure are reported in Table 2. The most significant difference between the thermal properties of degraded samples and the corresponding nondegraded ones was found in the values of ΔHm. For every sample, the ΔHm, and thus, the amount of crystal phase present, increased during degradation. The increase of ΔHm was more evident for P(BS-b-DGS)-f compared to the other samples.
The alamarBlue fluorescence assay was used to assess potential indirect cytotoxicity of PBS-f and P(BS-b-DGS)-f on H9c2 cells. Data shown in Figure 7 indicate that fluorescence output was comparable for cells grown during 24 h in PBS-f or P(BS-b-DGS)-f extraction medium (128500 ± 3023 au and 122400 ± 2199 au, respectively) and in standard DMEM control (125300 ± 3174 au).
Figure 5. (a) Percentage of residual weight (mres%) and (b) percentage of residual weight average molecular weight (Mw-res%) of PBS and P(BS-b-DGS), both in the form of films and electrospun mats, as a function of days of buffer exposure.PBS: poly(butylene succinate); P(BS-b-DGS): poly(butylene/diethylene glycol succinate); f: film; es: electrospun mats.
Figure 6. DGS mol% content in P(BS-b-DGS) as a function of degradation time calculated by 1H-NMR measurements carried out on the retrieved samples.1H-NMR: proton nuclear magnetic resonance; DGS: diethylene glycol succinate; P(BS-b-DGS): poly(butylene/diethylene glycol succinate); f: film; es: electrospun mats.
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When exposed to 1 mM H2O2 for 120 min, as a positive cytotoxicity control, all the cells were killed (data not shown).
The alamarBlue fluorescence assay was also used to evaluate cell adhesion and prolifera-tion: quantification was performed by fluorescence measurements (expressed in arbitrary units) on aliquots of medium withdrawn at days 1, 7, and 14 (Figure 8). Our results show that after 24 h from cell seeding, films and electrospun scaffolds of homo- and copolymer hosted about the same number of H9c2 cells (PBS-f = 31350 ± 1572 au, PBS-es = 28000 ± 3769 au, P(BS-b-DGS)-f = 31610 ± 6997 au, and P(BS-b-DGS)-es = 23480 ± 2868 au) (Figure 8). For all the tested conditions, this is about 50% of the number of H9c2 cells, which adhered to the
Figure 7. Evaluation of indirect cytotoxicity of PBS-f and P(BS-b-DGS)-f.DMEM: Dulbecco’s modified Eagle’s medium; PBS: poly(butylene succinate); P(BS-b-DGS): poly(butylene/diethylene glycol succinate); f: films.
Figure 8. Evaluation of cell adhesion and proliferation on PBS and P(BS-b-DGS) films and electrospun scaffolds at days 1, 7, and 14. At day 7, * = significantly different (P < 0.05) versus PBS-f; at day 14, ° = significantly different (P < 0.001) versus PBS-f, PBS-es, and P(BS-b-DGS)-es.PBS: poly(butylene succinate); P(BS-b-DGS): poly(butylene/diethylene glycol succinate); f: film; es: electrospun mats.
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control polystyrene surface (61432 ± 6997 au). At day 7, the number of cells growing onto PBS-f (67250 ± 7102 au) increased twofold, showing a significant difference (unpaired t-test, P < 0.05) compared to the number of the cells proliferating onto PBS-es (29950 ± 3546 au), P(BS-b-DGS)-f (25580 ± 2061 au), and P(BS-b-DGS)-es (33070 ± 6611 au). At the end of the experiments (day 14), the number of cells on PBS-es (90950 ± 6960 au) and P(BS-b-DGS)-es (83500 ± 4785 au) increased considerably, approaching the fluorescence values obtained by cells seeded on PBS-f (112000 ± 14990 au). In contrast, the number of cells on P(BS-b-DGS)-f (23400 ± 10760 au) remained almost the same evaluated at the beginning of the experiment, significantly different (unpaired t-test, P < 0.001) from the results obtained in all the other tested conditions.
In agreement with data on cell proliferation, SEM analysis showed that H9c2 cells adhered effectively on PBS-f, PBS-es, and P(BS-b-DGS)-es substrates. By comparing the SEM observa-tions taken up to 14 days, the substrate surfaces were covered with an increased number of elon-gated, sheet-like H9c2 cells. In contrast, H9c2 cells seeded on P(BS-b-DGS)-f were seen to adhere randomly onto the substrate surface, thus suggesting that cell growth remained almost unchanged during the time of culture. The H9c2 cells remained adherent to the scaffold surface even after 35 days of culture. Fully spread H9c2 cells covered, as a confluent monolayer, the entire surface of P(BS-b-DGS)-es. The cell adhesion was lower on the other substrates, and P(BS-b-DGS)-f showed the lowest surface cell coverage (Figure 9).
Immunohistochemistry was used to establish whether culture on polymeric electrospun fibrous scaffolds could modify the lineage phenotype of H9c2 cells. Before culturing, H9c2 cells showed a marked and diffuse α-SMA expression and a focal dot-like positivity for car-diac troponin I; desmin was negative. After 35 days of culture, a diffuse cardiac troponin I expression was seen in the H9c2 cells adhering to PBS-es and P(BS-b-DGS)-es. In these scaf-folds, cardiac troponin I also increased in intensity; α-SMA remained unchanged, and desmin remained negative (Figure 10).
Figure 9. SEM analysis of H9c2 cells seeded on PBS and P(BS-b-DGS) films and electrospun scaffolds after 35 days of culture. Scale bars = 100 µm.SEM: scanning electron microscopy; PBS: poly(butylene succinate); P(BS-b-DGS): poly(butylene/diethylene glycol succinate).
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Discussion
The copolyester P(BS-b-DGS) studied in the present work is a multiblock copolymer of BS and DGS sequences, obtained by reactive blending of the two parent homopolymers. In general, when two polymeric chains with reactive functional groups—in the present case ester groups and carboxylic and hydroxyl end-groups—are mixed in the molten state for an appropriate period, exchange reactions occur leading to the formation of copolymers with different archi-tectures, according to the mixing time. Reactive blending is a well-known synthetic method, generally employed to improve mutual miscibility between two otherwise immiscible polymers by in situ reactive compatibilization22: it is also effective for obtaining block, random, or grafted copolymers, according to the blending conditions.12,23,24 In this work, we used reactive blending as a synthetic approach to produce new biomaterials through copolymerization of selected homopolymers. PBS and PDGS homopolymers were initially synthesized by a polycondensa-tion reaction, and the P(BS-b-DGS) copolymer was then obtained by melt reactive blending. The relatively high molecular weight of the synthesized samples (see Table 1) indicated that appropriate synthesis conditions and a good polymerization control were obtained. The consist-ency between the experimental and theoretical NMR spectra demonstrates the absence of side reactions.
Figure 10. Histological and immunohistochemical analysis for lineage markers of H9c2 before culturing and of H9c2 after 35 days of culture on PBS-es and P(BS-b-DGS)-es. Scale bars = 50 µm.PBS: poly(butylene succinate); P(BS-b-DGS): poly(butylene/diethylene glycol succinate); es: electrospun mats; H&E: hematoxylin and eosin SMA: smooth muscle actin; Tn-I: troponin I.
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In order to highlight how the introduction of DGS units into PBS chain affects solid-state prop-erties and the degradation behavior of the resulting copolymer, films and electrospun scaffolds of PBS and P(BS-b-DGS) were prepared, and thermomechanical properties, degradation, and bio-compatibility were assessed. Electrospinning was chosen as the processing technique to produce biomimetic nonwoven flexible mats expected to be suitable scaffolds in engineering soft tis-sues,25,26 such as skin,27,28 nervous,29,30 vascular,31,32 and cardiac33–35 tissues.
Thermal characterization performed on PBS and P(BS-b-DGS) (Table 2) revealed that, at RT, both polymers possess a rubbery amorphous phase (Tg < RT) and a rigid crystal phase. However, the introduction of DGS units into the PBS chain decreases the degree of crystallinity and the per-fection of the crystals developed during the cooling from the melt (in the case of film production) and during fiber solidification (in the case of electrospun mat production). For both PBS and P(BS-b-DGS), electrospun mats display a lower degree of crystal phase compared to the corresponding films. Suppression of crystal phase formation is typically observed in electrospun materials, and it is ascribed to the extremely fast evaporation of the solvent that prevents macromolecular organiza-tion in a crystal structure before the occurrence of fiber solidification.36–38
Characterization of mechanical properties (Figure 4 and Table 3) indicated that P(BS-b-DGS) possesses a lower elastic modulus and a lower tensile stress at break when compared with PBS (P(BS-b-DGS)-f: E = 96 ± 12 MPa, σb = 6.4 ± 0.5 MPa; PBS-f: E = 488 ± 69 MPa, σb = 34 ± 1 MPa). Since the investigated polymers display a soft amorphous phase with comparable mobil-ity—that is, similar Tg values—the observed decrease in stiffness when DGS units are incorporated into PBS chain might be ascribed to the decrease in crystallinity degree (χc-PBS-f = 59% and χc-P(BS-b-DGS)-f = 49%). An additional point to be taken into consideration is that when polymers are processed into electrospun nonwoven scaffolds, the stiffness of the mat decreases and the elongation at break increases with respect to corresponding films. This is related to the porous structure of the electrospun mat and to the lower amount of crystal phase developed during electrospinning.
Because of the intended use of these new P(BS-b-DGS) electrospun scaffolds for soft tissue engineering applications, knowledge of their hydrolytic degradation behavior is needed. To that end, in this study hydrolysis of both P(BS-b-DGS) and PBS—as a film and as an electro-spun scaffold—was studied, with the intention of determining how the chemical structure and the fibrous nanostructure architecture affects the degradation profile. Several hydrolytic deg-radation studies have been performed on polyesters—most of them examining poly(lactic acid) and poly(lactide-co-glycolide) copolymers—to clarify hydrolytic mechanisms within this polymer family.39–41 In general, the main factors affecting hydrolysis rate are the materi-al’s hydrophilicity and morphology—in essence, the amorphous-to-crystalline phase ratio—and specimen’s dimensions. When the sample is in contact with aqueous medium, water absorption occurs, at a rate depending on material’s hydrophilicity and on inter-/intrachains free volume, and hydrolysis of the ester bonds begins, leading to a random chain cleavage. However, at neutral pH, diffusion of water into the polymer is faster than the hydrolysis of ester bonds, and aliphatic polyesters therefore degrade by a bulk degradation process.41 In the first stage, macromolecules in the amorphous phase are hydrolyzed, while the closely packed crystal phase is eroded by water only at a later stage. As the hydrolysis of the amorphous phase proceeds, chain fragments of low molecular weight diffuse and dissolve into the aqueous medium, leading to sample weight loss. It has been extensively reported that the carboxylic end-groups produced during hydrolysis may display an autocatalytic effect on the kinetics of degradative process,42 especially if the diffusion of acidic polymeric fragments out of the sam-ple is limited and they accumulate within the specimen core. In these cases, degradation rate
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is faster in the bulk than at the sample surface.39,40 Evidence in the literature demonstrates that larger specimens degrade faster compared to smaller ones,43–45 owing to the effect of autoca-talysis in the former case. However, the dimensional lower limit of specimen below which the autocatalytic effect becomes insignificant is not yet clear, especially in the case of the submi-crometric fibers such as those that may be obtained by electrospinning. Only a limited number of studies concerning the hydrolytic degradation of electrospun fibers are available to date,35,46–48 and their results are not at all exhaustive and often are conflicting.
Results of degradation experiments obtained in the present work show that all samples (i.e. PBS-f, PBS-es, P(BS-b-DGS)-f, and P(BS-b-DGS)-es) underwent a decrease of polymer molecu-lar weight during water exposure (Figure 5(b)). By separately comparing films and scaffolds made out of the two different materials, it emerged that the decrease of Mw, which is correlated to the rate of ester cleavage, was the same for PBS and P(BS-b-DGS). This result indicated that the slight increase of hydrophilicity, provided by the incorporation of DGS units into PBS, and evidenced by contact angle measurements, did not affect the rate of ester cleavage along the chain. Unlike the P(BS-b-DGS) samples (Figure 5(a)), the PBS samples (both as films and scaffolds) did not lose weight during the degradation experiment, despite the similar rate of ester cleavage displayed by PBS and P(BS-b-DGS) (Figure 5(b)). In both materials, macromolecules in the amorphous phase are highly mobile, Tg being below RT. Therefore, once the degrading chains are short enough to dissolve in water, they should be able to diffuse out of the sample, leading to sample weight loss. However, if short fragments are anchored to the rigid crystal phase, their diffusion is unlikely. The higher amount of crystal phase in the PBS, plus the greater hydrophobicity of PBS fragment, com-pared to that of a P(BS-b-DGS) fragment of the same length, may explain the constant weight observed for PBS samples during the degradation experiment.
For the same material, there was a difference in the rate of Mw decrease observed in the film and scaffold forms: the reduction of Mw proceeded faster for films than for electrospun samples (Figure 5(b)). This finding could be ascribed to the autocatalytic effect of carboxylic end-groups exerting greater influence on the degradation kinetics in 200-µm-thick films than in submicrometric electro-spun fibers. In the latter, the small fiber diameter allows a more effective diffusion of the acidic degradation fragments into the surrounding buffer in comparison with thick films. The faster degra-dation rate of films compared to fibrous scaffolds is also indicated by the results presented in Figure 5(a), showing that weight loss in the P(BS-b-DGS)-f sample was greater than that in the correspond-ing P(BS-b-DGS)-es sample. DSC analysis performed on degraded samples (Table 2) showed an increase of ΔHm, especially for P(BS-b-DGS)-f. Since, typically, degradation occurs mostly in the amorphous phase, it is reasonable to assume that the increase of crystal phase is the consequence of macromolecules released from the amorphous phase, particularly for P(BS-b-DGS)-f and P(BS-b-DGS)-es, where a significant weight loss was observed. However, for all samples investigated, an annealing process during buffer exposure cannot be excluded.
A change in P(BS-b-DGS) composition was observed during degradation (Figure 6). The decrease of DGS content in degraded P(BS-b-DGS) samples demonstrates that the soluble frag-ments, lost from the retrieved samples, were richer in DGS units. This would be anticipated, given the greater hydrophilicity of the DGS unit compared to the BS unit. Composition change of the copolymer during degradation was more apparent in the film than in the scaffold, again confirming the higher rate of degradation of the film compared with the electrospun scaffold.
A practical conclusion to be drawn from the above is that it could be possible to design and fabricate an electrospun scaffold made out of a P(BS-b-DGS), which would have physical proper-ties close to those typical of the extracellular matrix in soft tissues. For a preliminary assessment of our new P(BS-b-DGS) electrospun scaffold for tissue engineering applications, we used the
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H9c2 cell line, derived from embryonic rat heart and largely employed as a cellular muscle model.49 Biocompatibility assays were performed in accordance with ISO10993-5. In particular, the alamar-Blue fluorescence assay was used to assess potential indirect cytotoxicity and degree of cell adhe-sion and proliferation when using PBS and P(BS-b-DGS) substrates. As expected, absence of potentially cytotoxic products released into the culture medium by the investigated polymer sam-ples was recorded (Figure 7), indicated by an essentially equivalent metabolic activity in cells cultured either in standard DMEM or in PBS and P(BS-b-DGS) extraction culture medium.
Cell adhesion on all films and electrospun scaffolds, whether made from PBS homopoly-mer or P(BS-b-DGS), was broadly similar, and we assessed that about the same number of cells was resident on all the substrates 24 h after cell seeding (Figure 8, day 1). Although the number of adhering cells on these substrates was lower than the number of cells attached to the control polystyrene surface after the same interval, our data show that all our fabricated substrates support a physical environment where cells can establish colonies. After 1 week in culture, only the cells seeded onto PBS-f were able to grow at a significant rate (Figure 8, day 7). The molecular mechanism underlying this observation is not clear, in particular in light of data collected on the same substrates 1 week later. In fact, at day 14, we could consistently score in the three separate experiments, two replicates each, that the cells adhering on PBS-es and P(BS-b-DGS)-es proliferated and became about three times more than 1 week before (Figure 8, day 7). At this time point of the experiment, the population cultured on PBS-es and P(BS-b-DGS)-es resulted in the same order of magnitude of the one growing on PBS-f. No changes were observed throughout the duration of the experiment in the number of cells cul-tured onto P(BS-b-DGS)-f (Figure 8). These results, based on the evaluation of the metabolic function of the cells, which actively reduce resazurin proportionally to their number, were confirmed by SEM. SEM evaluation showed that the PBS-f, PBS-es, and P(BS-b-DGS)-es substrates were covered with elongated, sheet-like cells at day 14. Randomly adherent cells were otherwise scored on P(BS-b-DGS)-f substrate surface, confirming that cells did not grow during the time course of culture. Although not proliferating, these cells were viable and meta-bolically active, as indicated by their ability to reduce resazurin partially. Additional SEM analysis was performed on cells grown up to 35 days. After 1 month in culture, fully spread cells covered, as a confluent monolayer, the entire surface of P(BS-b-DGS)-es (Figure 9). Comparable results were collected processing the PBS-f and PBS-es substrates. However, even after such a prolonged time of culture, P(BS-b-DGS)-f showed a lower percentage of surface cell coverage (Figure 9).
It is known, and confirmed in this work by immunohistochemistry, that H9c2 cells harbor a par-tially differentiated phenotype. Even before seeding on our substrates, the H9c2 cells showed dif-fuse α-SMA expression and a focal dot-like positivity for cardiac troponin I (Figure 10). After 35 days in culture, an increased and diffuse cardiac troponin I expression was seen in H9c2 cells grow-ing in PBS-es and P(BS-b-DGS)-es, with a virtually unchanged signal for α-SMA (Figure 10). The increase in this cardiac marker indicated that, growing in a morphologically biomimetic environ-ment, muscle-committed cells might proliferate while maintaining desired markers of the cardiac phenotype.
Conclusions
Electrospun scaffolds made from a biodegradable polyester, a multiblock copolymer of BS and DGS (P(BS-b-DGS)), were investigated for soft tissue engineering applications. A reac-tive blending process was used as a simple, versatile, and cost-effective synthetic strategy to
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obtain the copolymer. The introduction of DGS units into PBS chain had the effect of changing polymer mechanical properties, as a consequence of reducing the degree of polymer crystal-linity, shifting them closer to elastomeric behavior, which is particularly important in the context of soft tissue engineering applications. The specimen dimensions affected both the mechanical properties and the degradation profile of the material. The electrospun scaffolds degraded slower than the corresponding thick, nonporous films; thus, the thinner electrospun fibers were hydrolyzed to a lesser extent than the thick films. As a consequence, more limited sudden release of acidic by-products is expected during in vivo degradation of electrospun fibers, thus reducing inflammatory reactions. The physical properties of the P(BS-b-DGS) electrospun materials, together with the encouraging cell culture results, suggest a high poten-tial for its application in regenerative medicine of soft tissues, such as injured cardiac tissue, and its suitability for further studies in the field of tissue engineering for regenerative medi-cine of mesenchyme-derived tissues. This study represents an initial step in the long path to providing more experimental evidence to justify such an application. It does, however, pro-vide a reasonable approach, which starts from thorough investigation of chemical and physical properties of the polymer and the corresponding biocompatible scaffold.
Funding
This research was carried out with the financial support of Regione Emilia Romagna (Programma di Ricerca Regione Università 2007–2009, Area 1b “Medicina rigenerativa”). C.G. is the recipient of a fellowship awarded from the Spinner Consortium of Regione Emilia Romagna. The postdoctoral position filled by M.G. was open with the financial support of the Istituto Nazionale per le Ricerche Cardiovascolari (INRC) to E.G.
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