ARTICLE pubs.acs.org/Biomac
Biobased Poly(propylene sebacate) as Shape Memory Polymer with Tunable Switching Temperature for Potential Biomedical Applications Baochun Guo,*,† Yongwen Chen,† Yanda Lei,† Liqun Zhang,*,‡ Wen You Zhou,§ A. Bakr M. Rabie,§ and Jianqing Zhao† †
Department of Polymer Materials and Engineering, South China University of Technology, Guangzhou 510640, China Key Laboratory of Beijing City for Preparation and Processing of Novel Polymer Materials, Beijing University of Chemical Technology, Beijing 100029, China § Discipline of Orthodontics, Faculty of Dentistry, The University of Hong Kong, 34 Hospital Road, Hong Kong ‡
bS Supporting Information ABSTRACT:
From the point of better biocompatibility and sustainability, biobased shape memory polymers (SMPs) are highly desired. We used 1,3-propanediol, sebacic acid, and itaconic acid, which have been industrially produced via fermentation or extraction with large quantities as the main raw materials for the synthesis of biobased poly(propylene sebacate). Diethylene glycol was used to tailor the flexibility of the polyester. The resulted polyesters were found to be promising SMPs with excellent shape recovery and fixity (near 100% and independent of thermomechanical cycles). The switching temperature and recovery speed of the SMPs are tunable by controlling the composition of the polyesters and their curing extent. The continuously changed switching temperature ranging from 12 to 54 °C was realized. Such temperature range is typical for biomedical applications in the human body. The molecular and crystalline structures were explored to correlate to the shape memory behavior. The combination of potential biocompatibility and biodegradability of the biobased SMPs makes them suitable for fabricating biomedical devices.
’ INTRODUCTION Shape memory polymers (SMPs) have a sensitive response to the external stimuli, such as temperature, pH, humidity, light, electricity, and so on. Until now, the majority of the reported SMPs are stimulated by temperature. All SMPs contain two networks, which are called the permanent networks and the temporary networks. The permanent networks are either covalent crosslinks or physical cross-links, and the “bonds” exist above transition temperature (Ttrans). The temporary networks typically rely upon vitrification,1,2 crystallization,3 or some other physical interaction such as hydrogen bonding and ionic bonding.46 SMPs have a permanent shape that is provided by a permanent network, but they can be deformed above Ttrans. Temporary network is fixed into a temporary shape when the SMP is cooled under stress to below Ttrans. When reheated above Ttrans without stress, the material assumes its permanent shape.7,8 Because of the sensitivity to the environmental conditions, the SMPs have found various applications, such as actuators, smart fabrics, heat shrinkable tubes for electronic packaging, self-deployable r 2011 American Chemical Society
sun sails in spacecraft, and self-disassembling mobile phones.9 The applications of SMPs in biomedical devices, such as cardiovascular applications, nerve regeneration, orthopedic and orthodontic applications, wound healing lesions, and so on have also been highlighted.10,11 Up to now, the reports about the SMPs in biomedical devices were concentrated on petroleum-dependent polymers such as trans-polyisoprene, poly(ε-caprolactone) (PCL), polyurethanes (PU), and their copolymers.1214 Because of the structural characteristics of biobased polymers, their biocompatibility and expected biodegradability make them specially suitable for biomedical applications. Meanwhile, the energy, resources, and serious environmental problems influence our life deeply today. The full use of renewable resources and reducing dependence on fossil fuels has been paid global Received: January 8, 2011 Revised: March 4, 2011 Published: March 07, 2011 1312
dx.doi.org/10.1021/bm2000378 | Biomacromolecules 2011, 12, 1312–1321
Biomacromolecules attention to by many scientists. Consequently, the SMPs derived from biomasses or their extracts are of great importance and highly desired. The newly developed SMPs with biocompatibility or biodegradability have been reported. Mather et al. first developed a series of biodegradable hybrids of polyurethane, PCL, and poly(lactide-co-glycolide) (PLGA), all incorporating polyhedral oligomeric silsesquioxanes (POSS) as SMP applications.1518 Cai et al. synthesized a biocompatible and biodegradable elastomer, poly(glycerol-sebacate) (PGS), which exhibited good shape recovery ratio above the Ttrans (Tm = 10.6 °C).19 Lu et al. reported SMPs based on poly(L-lactide-co-εcaprolactone) (PCLA) copolymers, but the biocompatibility and biodegradability research were not involved.20 The SMPs based on polyglycerol-dodecanoate (PGD) with the combination of good mechanical properties, proper Ttrans (Tg of 32 °C), biocompatibility, and biodegradability were reported to be attractive for biomedical applications.21 Although the SMPs derived from biomass have been reported,2224 the SMPs derived from the industrially produced biomass extracts or zymotic biomasses, especially those with large quantities, are limited. Several diacids and diols, such as 1,3propanediol (PDO) and itaconic acid (IA), have been industrially produced via fermentation. Sebacic acid could also be derived from castor oil. They are potential raw materials for biobased polyesters (BPEs) as SMPs. More importantly, the reported biocompatible SMPs showed limited mechanical properties, and their tunability has not been realized. For example, the SMPs from PGS exhibited quite low tensile strength below 0.5 MPa,25 which limited their application as biomedical materials or devices. For biomedical devices, it is desirable to thermally activate the SMP via physiological heat. In such applications, the deployment of the device occurs typically from a compressed state packaged at ambient temperature. Following deployment into the desired location, the device is exposed to higher physiological temperature and reverts to its initial, desired shape.26 Meanwhile, the delicate tunability is also important in the biomedical application of SMPs as different switching temperatures meet different applications.2629 In the present work, a series of BPEs based on PDO, sebacic acid (SA), and IA were synthesized. Diethylene glycol (DEG) was introduced to tailor the chain flexibility. At ambient temperature, BPEs cover from rigid plastics to flexible elastomers, which permits their applications as SMPs with tunable switching temperature. The effects of composition and curing extent on the SMP performance, such as microstructures, switching temperature, shape recovery, and shape fixity were investigated. The cell proliferation and cell attachments were examined to evaluate the biocompatibility of these BPE-based SMPs. The biodegradability was also evaluated in phosphate-buffered saline (PBS).
’ EXPERIMENTAL SECTION Raw Materials. PDO (purity of 98.0%), IA (purity of 99.9%), and SA (purity of 99.0%) were purchased from Hunan Rivers Bioengineering, Qingdao Kehai Biochemistry, and Tianjing Damao Chemicals, respectively. DEG with purity of 99.5% was provided by Sinopec Yangzi Petrochemical. These diacids and diols were used as received. Tetra-nbutyl titanate (TBT, 98%), p-hydroxyanisole, and analytic grade dicumyl peroxide (DCP) were obtained from SinoPharm Group and also purified by recrystallization. Synthesis of BPEs and Preparation of SMPs. The reaction mixture with diols/diacids molar ratio of 1.05 and SA/IA molar ratio of 9
ARTICLE
was charged into the 150 mL three-necked flask. p-Hydroxyanisole (0.5 wt % relative to total of reactants) was used as the free radical inhibitor. The mixture was precondensated under nitrogen atmosphere at 190 °C for 1 h. TBT (0.5 wt % relative to total of reactants) was used as the catalyst. After gradually heating to 220 °C under vacuum, the polycondensation lasted for 35 h until the Weisenberg effect was found. The products were purified by dissolving in chloroform and precipitating in cold methanol. The precipitate was filtered, washed with methanol, and vacuum-dried at 50 °C. The obtained polyesters were characterized by gel permeation chromatography (GPC). The numberaverage molecular weight and polydispersity index (Mw/Mn) of these polyesters were determined to be about 30 00040 000 g/mol and 2.004.89, respectively. For the preparation of the SMPs, the purified BPE was mixed with DCP in chloroform. The mixture was dried under vacuum and subjected to compression molding at 150 °C for 15 min. The n in sample code Dn denotes the sample with fed DEG molar percentage of n, similarly hereinafter. Measurements. Proton nuclear magnetic resonance (1H NMR) was used to determine the chain compositions of BPEs. 1H NMR spectra of BPEs with deuterated chloroform as the solvent were recorded with tetramethylsilane as the internal standard using a Varian UNITY INOVA-500 NMR at 22.5 °C. The molecular weight and its distribution were measured by GPC using tetrahydrofuran as the solvent (GPC, Agilent 1100). The X-ray diffraction (XRD) studies were conducted at ambient temperature on a Rigaku Dmax/III diffractometer using Cu KR ) with an accelerating voltage of 40 kV and a current radiation (λ = 1.54 Å of 30 mA. All the samples were scanned from 5 to 60° with a step length of 0.02° at 24 °C. Melting and recrystallization of the uncured and cured BPEs were determined with a Q20 differential scanning calorimeter (TA, New Castle, DE). The thermal history of the sample was eliminated by heating the sample to 150 °C. After cooling to 65 °C, the sample was reheated at 10 °C/min to measure the melting point (Tm). Tensile tests of the SMPs were performed at 28 °C by using a Zwick Roell Z010 instrument at a crosshead speed of 50 mm/min. The tensile tests at elevated temperature were performed by using a Q800 dynamic mechanical analyzer (TA). After equilibrating at Tm þ 10 °C for 10 min, the tensile was initiated at 0.05 N/min until fracture. The tensile tests at elevated temperature were repeated for at least three times. Dynamic mechanical analysis (DMA) was conducted with an EPLEXOR 500N DMTS instrument (GABO, Ahlden, Germany) under a tensile mode with a dynamic strain of 0.5%. The frequency and heating rate were set as 1 Hz and 3 °C/min, respectively. The thermomechanical cycles were also performed with the Q800 machine to characterize the shape memory behavior. Prior to the cycle, an equilibration at Tm þ 20 °C for 10 min was used. In step 1, the sample was deformed by ramping force from preload 0.005 N to a designed value at a rate of 0.05 N/min (deformation). In step 2, the sample was cooled at the rate of 3 °C/min to Tm 20 °C to fix the deformed sample under constant force (cooling). In step 3, the force to the sample was unloaded to preload value (0.005 N) at a rate of 0.05 N/min. After the unloading step, an additional 10 min of isothermal step was included to ensure shape fixing at Tm 20 °C (unloading and shape fixing). In the final step, the sample was reheated at the rate of 3 °C/min to Tm þ 20 °C and held at Tm þ 20 °C for 10 min to recover any residual strain (recovery). The thermomechanical cycle consisting of four steps was repeated four times on the same sample. To reveal the effect of cooling on the shape fixing, the thermomechanical cycles of D0 were also done with changed cooling step (down to 10 °C). The other steps were not changed. Another method with larger specimen (20.0 mm 5.0 mm 0.7 mm) was employed to determine the shape fixity (SF) and shape recovery (SR) of all samples. The sample was first heated to a loading temperature of Tm þ 20 °C and loaded to the strain (εm) of 100 or 50%, 1313
dx.doi.org/10.1021/bm2000378 |Biomacromolecules 2011, 12, 1312–1321
Biomacromolecules
ARTICLE
followed by cooling and unloading at Tm 30 °C. Upon unloading, part of the strain (εm εu) was instantaneously recovered, leaving an unload strain (εu). Then, the sample was reheated to the loading temperature of Tm þ 20 °C to recover the strain, leaving a permanent strain (εp). Reheating at the loading temperature is known to give the maximum recovery stress. These three steps complete one thermomechanical cycle. SF and SR are defined as below. SFð%Þ ¼
εu 100, εm
SRð%Þ ¼
εm εp 100 εm
In vitro fibroblasts response on BPEs was performed. NIH3T3 mouse fibroblast cell line (ATCC, Manassas, VA) used in the experiments was passages 5. Cells were cultured and passaged in regular culture media consisting of Dulbecco’s modified eagle medium (DMEM, GIBCO) supplemented with 10 vol % fetal bovine serum (FBS, Invitrogen) and antibiotics penicillin 100 units/mL and streptomycin 100 μL/mL (Invitrogen). Cells were incubated at 37 °C in a 5% CO2 incubator, and the medium was changed every 3 days. Prior to cell culture assays, all BPE films were sterilized by exposure to UV light for 15 min and immersed in PBS for 1 day. The film was attached with 2% agarose to avoid floatation. Cells were routinely removed from tissue culture polystyrene (TCPS) dish (75T, Costa) with 0.25% trypsin-EDTA and plated on different substrates. The seeding cell density is 2 105 cells/mL. SEM analyses were performed to study the morphology of NIH3T3 cells grown on the surface of the SMPs. At day 7, the medium was removed and the wells were washed twice with PBS. The cells were then fixed with 2.5% glutaraldehyde in cacodylate buffer (Karnovsky fixation solution) for 424 h at 4 °C. Then, the specimens were washed with cacodylate buffer several times and the dehydration of the cells attached to the films was done in ethanol at 30, 50, 70, 80, 90, 95, and 100%, immersed for 15 min at each step. The critical point drying of these samples was performed in a critical point dryer (BAL-TEC CPD 030) using liquid carbon dioxide as transitional fluid. A 10 nm thick gold palladium layer was deposited on the samples with the use of a sputter coater (BAL-TEC SCD 005). The morphologies of the cells attaching to the surfaces of BPE films were taken by a Hitachi S4800 FEG SEM. Cell viability on the SMPs films was evaluated by the Cell Counting Kit-8 (CCK-8) assay (Enzo, Life Sciences). CCK-8 is a kind of cell viability assay reagent with higher sensitivity and better reproducibility than MTT, MTS, or WST-1 by utilizing a highly water-soluble formazan dye. At designated time intervals (1, 4, and 7 days), 10 vol % of CCK-8 solution relative to medium was added to each well, and the plate was incubated for 3 h in the incubator. The absorbance was measured at 450 nm using a microplate reader. Cell viability was calculated by the following equation: Cell viability (%) = (As/Acontrol) 100, where As is the absorbance of the cells cultured on films and Acontrol is the absorbance of the cells incubated on TCPS only (positive control). The absorption value of phenol red in the culture medium was removed by subtracting the absorption value of a blank solution. Data were analyzed with a statistical analysis software (SPSS 15.0) and expressed as means ( standard deviation. The unpaired t tests with Welch correction were used to compare the cell viability on control and different films. Significance was set in advance at p < 0.05. In vitro degradation of BPEs was done in PBS at 37 °C. A proper solution-to-mass ratio, 50:1, was employed, that is, 10 mL of PBS solution for one sample, whose average mass was 0.2 g. Ten to fifteen slabs for each sample were immersed in PBS solution at pH of 7.4 at 37 °C in a shaking water bath with a rotation speed of 20 rpm.30 Over the course of study, the pH values of the PBS were closely monitored. Although no measurable changes in pH were observed, the PBS solution was changed fresh at least once a week. Samples were removed at the different time points, dried at 40 °C, and weighed again. Dry weight change was thus determined. Mass spectra were collected on an Esquire HCT PLUS
Figure 1. Preparation of cross-linked BPE and its structure. (Bruker Daltonics, Bremen, Germany) by electrospray-ionization mass spectrometry in scan mode (m/z 50500) with detection in the negative ion mode, and MS parameters were set to the following values: capillary potential 3.5 kV, nebulizer gas 20 psi, and drying gas flow and temperature were 10 L/min and 350 °C, respectively.
’ RESULTS AND DISCUSSION Tunability of Switching Temperature with Variable Composition and Curing Extent. The polycondensation of SA, IA,
PDO, and DEG results in a series of semicrystalline copolyesters with variable crystallinity. The pendant double bonds introduced by IA permit the free radical cross-linking of BPE by DCP into three-dimensional network. The cross-linked BPEs behave as elastomers above their Tm values although they are still crystallizable in the lowered temperature. BPEs exhibit quite low glasstransition temperature (Tg) (49.4 ∼ 39.7 °C), independent of composition and curing extent (Table S1 of the Supporting Information). Consequently, the cross-linked BPE consists of a physically bonded network (via crystallites) and a covalently bonded network structure, which meets the structural requirements of SMPs. The preparation of cross-linked BPE and its structure is schematically illustrated in Figure 1. For semicrystalline copolyester, the shape memory behavior is triggered by melting of the crystalline domain in the polymer. To alter Tm and accordingly tune the switching temperature, in this work, DEG is introduced to modify the chain sequences of BPE. Alternatively, the cross-linking of BPE would effectively restrain the crystallization and decrease Tm of BPE to some extent. By varying the composition and curing extent, BPE shifts from rigid plastics to flexible elastomers at ambient temperature. Figure 2 shows the Tm of BPE as function of DEG molar fraction and DCP content. The molar fraction of DEG in the glycols is set from 0 to 100%, and the content of DCP is set from 0 to 0.8 wt % related to BPE. The real DEG molar fraction in the chain was determined by 1H NMR (Table S2 of the Supporting Information). As shown in Figure 2, Tm of BPE is greatly changed when DEG/PDO molar ratio is changed. One can find that Tm of BPE containing both PDO and DEG is lower than that of BPE containing only PDO or 1314
dx.doi.org/10.1021/bm2000378 |Biomacromolecules 2011, 12, 1312–1321
Biomacromolecules
Figure 2. Tm of BPEs as a function of DEG molar fraction and DCP content.
DEG. This may be interpreted by the increase in sequence randomness in the copolyester, which was also observed in other copolyester systems.31,32 The increase in sequence randomness will be substantiated by the decrease in crystallinity, as indicated in the below section. When DEG fraction is 90%, BPE possesses a Tm >30 °C. The BPE with higher unbalance of glycol molar ratio has higher Tm. BPE with intermediate fraction of DEG (about 4070%) behaves as an elastomer under ambient temperature (30 °C). At different DEG fraction, the decreasing extent of Tm caused by cross-linking is different. The highest impact exerted by cross-linking is found in BPE without DEG. With increasing DCP content, Tm of BPE decreases continuously. When 0.8 phr of DCP is used, Tm is 12 °C lower than the uncured one. The cross-linking of BPE restrains the crystallization process. The cross-links suppress the chain mobility and increase sequence randomness, which therefore decreases the size of crystallite and lowers Tm. For the composition with very high fraction of DEG, the DCP content does not effectively influence Tm of BPE. This may be explained by the greater mobility caused by more flexible sequences containing DEG. As a consequence, the restriction of chain ordering into crystal lattices by cross-links would be insignificant. From Figure 2, it is clearly demonstrated that Tm of BPE could be finely tuned, from 12 to 54 °C, by changing the composition and the curing extent of BPE. In some previous reports, if the switching temperature is around or higher than body temperature to some extent, then the SMPs could potentially be used for various biomedical applications.11 Effects of Composition and Curing Extent on Crystal Structures. Figure 3 presents wide-angle X-ray diffraction (WAXD) patterns of the annealed BPE series as a function of composition. The diffraction peaks around 18.9, 20.3, 21.6, and 22.4° were found in all PDO-rich BPEs. Usually orthorhombic crystal is formed in the polyester of SA and PDO. The diffractions around 18.9, 20.3, 21.6, and 22.4° are assigned to (102), (016), (112), and (105) diffractions, respectively.33 For the DEG-rich samples, however, the diffractions are found to be around 18.7, 19.9, and 23.4°. For the samples containing from 0 to 40 mol % DEG, the SA-propanediol sequences crystallize whereas the SA-diethylene glycol sequences remain in the amorphous regions. On the contrary, in
ARTICLE
Figure 3. WAXD pattern evolution of BPEs with continuously changed DEG content. The inserted picture is the dependence of crystallinity of BPEs on DEG molar fraction by WAXD method.
Table 1. Characterization of D0 Samples with Different DCP Content samples (DCP, phr) Tm, °C Tc, °C ΔHc, J/g ΔHm, J/g crystallinity, %a
a
0
54.4
22.1
45.0
51.7
16.1
0.1
50.6
19.9
41.5
49.3
14.9
0.3 0.5
52.6 47.1
18.9 15.3
41.1 40.9
48.6 47.7
14.3 12.2
0.8
42.7
13.8
39.2
42.9
10.1
Calculated by WAXD method.
the samples containing 70100 mol % DEG, the SA-diethylene glycol sequences crystallize and the SA-propanediol sequences remain in the amorphous regions. Consequently BPEs with dominant content of SA-propanediol or SA-diethylene glycol segments show the features of their own diffraction peaks. Similar results were found in other copolyester systems.34 Only the samples containing 5060 mol % of DEG, in which the sequential randomness is sufficiently high, are amorphous at 25 °C, although they are still crystallizable at lowered temperatures. By deconvolution of diffraction patterns, the crystallinity of BPEs could be calculated. The dependence of the calculated crystallinities on DEG molar fraction is also depicted in Figure 3. Consistent with Tm changes, as shown in Figure 2, the crystallinity of BPE reaches a minimum in middle content of DEG (60 mol %). When the DEG content is lowered or increased, the crystallinity gradually increases. To evaluate the possibility of tuning Ttrans (i.e., Tm and Tc) by curing extent, DSC and WAXD for D0 samples cured with different DCP concentrations were performed. The analysis data are summarized in Table 1. Compared with the uncured one, although cross-linking of BPE does not qualitatively alter the melting and recrystallization behavior (Figures S1 and S2 in the Supporting Information), quantitative changes are found. Inspection of Table 1 reveals that the transition temperatures, Tm and Tc, along with associated enthalpies of melting and crystallization, modestly decrease with increasing DCP concentration. In 1315
dx.doi.org/10.1021/bm2000378 |Biomacromolecules 2011, 12, 1312–1321
Biomacromolecules
Figure 4. Typical stressstrain curves of the cured BPEs with changing DEG molar fraction. The samples were cured with 0.5 phr of DCP; tensile tests were performed at 28 °C.
particular, when 0.8 phr of DCP is used, melting enthalpy and crystallization enthalpy decrease by 17 and 13%, respectively. This may be attributed to a constraining effect of cross-link junctions on crystal growth. Specifically, the local chain mobility necessary for chain insertion into a growing crystal is suppressed in the neighborhood of a cross-link site.3 This hypothesis is supported by the reduction in crystallinity with increasing DCP content, which is also calculated by diffraction method (Table 1). Other researchers also observed the crystallization depression by cross-linking.3,3539 Mechanical Properties of BPEs. Mechanical strength and elongation are essential for the applications of SMPs.21,40 Figure 4 depicts the typical stressstrain curves of BPE with changing DEG molar fraction. It is revealed that BPEs with unbalanced glycol compositions behave like plastics at ambient temperature and apparent yielding phenomena is found. They also exhibit much higher tensile strength and break strain. Those with balanced glycol compositions, however, behave like elastomers at ambient temperature with much lower tensile strength and higher break strain. These observations are actually consistent with those of crystallinity, as described above. BPEs with intermediate glycol ratios possess much lower crystallinity and consequently much lower tensile strength and higher break strain. The dependences of tensile strength and break strain of BPEs on DEG molar fraction are summarized in Figure S3 in the Supporting Information. Curing extent also influences mechanical properties. With increasing DCP content, the cross-linking density increases; consequently, the tensile strength and elongation at break decrease slightly (Figure S4 in the Supporting Information). The decreased tensile strength may be attributed to the disrupted crystallization in the samples with higher DCP content, which has been substantiated by WAXD. Actually, the drawability of a simicrystalline polymer was reported to be influenced by the combined effects of decreased crystallinity and increased cross-link density. In the present systems, the slightly decreased elongation at break may be attributed to the increased cross-link density. The tensile tests of the BPEs were also performed above their Tm values. The typical stressstrain curves are depicted in Figure 5. As shown, all BPEs behave like soft elastomers above Tm, with tensile strength of 0.7 to 1.6 MPa and elongation at break of 350430% (an exception of D0 with break strain of only 120%). The exhibited tensile strength and drawability of BPEs are
ARTICLE
Figure 5. Tensile stressstrain curves of BPEs at elevated temperature. The samples were cured with 0.5 phr of DCP; tensile tests were performed at Tm þ 10 °C.
Figure 6. Storage modulus of BPEs with different molar fraction of DEG. The samples were cured with 0.5 phr of DCP.
compatible with many natural tissues, such as human inferior cava vein (1.17 MPa),9 ulnar cadaveric peripheral nerve (0.5 to 0.6 MPa),41 and porcine aortic heart valve (8.3 MPa).42 The dynamic analysis was conducted on all cross-linked BPEs. The storage modulus as the function of temperature is shown in Figure 6. In general, the modulus below the switching temperature (crystalline modulus) governs the strength of the SMP, while the modulus above the switching temperature (rubbery modulus) determines the recovery rate. The driving force of shape recovery is the elastic force generated during the deformation. Consequently, higher rubbery modulus generates higher elastic stress and hence faster recovery speed. Many works on SMPs had similar observation and interpretation.4346 The SMP with higher crystalline modulus would have higher strength. However, the SMP with higher rubbery modulus would possess faster recovery rate. As clearly shown in Figure 6 (all with 0.5 phr of DCP), when the content of DEG or PDO dominates (Group A), the crystalline modulus is higher than those with matchable glycol content 1316
dx.doi.org/10.1021/bm2000378 |Biomacromolecules 2011, 12, 1312–1321
Biomacromolecules
ARTICLE
Figure 7. Three-dimensional diagram of stress-controlled programming cycles for (a) D0, (b) D0 with colder cooling step, (c) D50, and (d) D100. All samples were cured with 0.5 phr of DCP.
(Group B). The higher crystalline modulus is due to the higher crystallinity, as revealed above. Group A is also mechanically stronger than Group B at lower temperature. Interestingly, the rubbery moduli of Group B are higher than those of Group A, indicating faster recovery rate for Group B. As the similar value of gel fraction of BPEs (Table S3 of the Supporting Information), the variation in rubbery modulus should be attributed to the variation in cohesive energy density of the chains rendered by changed chemical composition. For practical applications, both mechanical strength and recovery rate are essential. Actually, the recovery rate could be further tailored by nanoreinforcement of BPEs. The effects of reinforcement of BPEs on the shape memory behavior are still under investigation and will appear in another paper. In the present systems, the mechanical strength and recovery rate could be tuned with the composition of BPE. Shape Memory Behavior of BPEs. The shape memory behavior was activated by changing the switching temperature below and above the Tm of BPEs with different compositions. BPEs behave as soft rubber above their Tm in permanent shape and
could be deformed to a temporary shape under external force. When the deformed samples are cooled below switching temperature, their temporary shape is fixed without loading, and the deformed shape can be kept well. However, if the operation temperature is raised to above the switching temperature again, then BPEs will quickly return to their initial shapes. The stresscontrolled programming cycles (four times) of samples D0, D50, and D100 are depicted in Figure 7. It is demonstrated that all three samples exhibit excellent shape recovery. During the cooling step, the SMPs are generally further deformed because of the loading. However, compared with D0 and D100, D50 shows different deformation behavior during the cooling step. Under the same cooling condition (Tm þ 20 °C to Tm 20 °C), D0 (Figure 7a), D50 (Figure 7c), and D100 (Figure 7d) are cooled to 32.6, 5.6, and 16.4 °C, respectively. The crystallization peak temperatures revealed by DSC (Figure S5 of the Supporting Information) for D0, D50, and D100 are 15.3, 11.4, and 0.3 °C, respectively. Consequently, during such cooling, D50 is expected to crystallize more completely than D0 and D100 because the 1317
dx.doi.org/10.1021/bm2000378 |Biomacromolecules 2011, 12, 1312–1321
Biomacromolecules
ARTICLE
Figure 8. Transition from the temporary spiral shape to the permanent linear shape for D20 (upper photos) and D0 (lower photos). The samples were cured with 0.5 phr of DCP, and the recovery process was recorded after heating the samples to Tm þ 20 °C.
lower limit (5.6 °C) for D50 is closer to the crystallization temperature (11.4 °C). As expected, apparent crystallizationinduced deformation (reversed-S shape)47 is observed for D50 (Figure 7c). From the observation on the third step (unloading), one can find that D50 exhibits slightly better shape fixity than D0 and D100. The relatively higher shape fixity for D50 is also attributed to the more complete crystallization of D50. To confirm this effect on the shape fixity, the sample of D0 is cooled to lower temperature (10 °C) to facilitate the crystallization. The thermomechanical cycles are depicted in Figure 7b. Expectedly, the reversed-S-shaped deformation during cooling, which is induced by crystallization, is also observed. Also, the shape fixity is substantially improved. For multiple cycling, because of the possible orientation of chains, the cyclic curves of all samples could not be overlapped completely. The SR and SF values of all samples were determined by larger sample specimen, and the results are also summarized in Table S2 of the Supporting Information. SR and SF values are found near to 100% and independent of cycle times, indicating very good shape memory properties. The independency of SR and SF values on chemical composition and curing extent clearly indicates the fast crystallization process upon cooling and the good elasticity of the chemically cross-linked network. The independency of SR and SF values on cycle times could be attributed to the low capability of chain orientation during the cycling tests. According to the model proposed by Kim,48 a high elasticity ratio (glass-state modulus/rubbery modulus), preferably with a difference of two orders of magnitude, allows easy shaping at T > Ttrans and great resistance to deformation at T < Ttrans. In the present system, all samples possess very high elasticity ratio (a difference higher than two orders of magnitude), which is responsible for the observed good shape fixity ratio for all of the samples. Figure 8 demonstrates the comparison of the transition from the temporary spiral shape to the permanent linear shape for D0 (upper photos) and D20 (lower photos) (all with 0.5 phr of DCP). Very clearly, the former possesses much faster shape recovery. As described above, the rubbery modulus of the SMP determines the recovery speed. According to Figure 6, D20 possesses much higher rubbery modulus than that of D0, although the former is mechanically weaker at room temperature. In
Figure 9. CCK-8 viability assay of NIH3T3 cell after 1, 4, and 7 days of cell culture on D0, D40, and D70 films in comparison with tissue culture PS (TCPS) as control. The samples were cured with 0.5 phr of DCP.
conclusion, the mechanical strength, switching temperature, and recovery speed could be tuned by the composition and curing extent without detrimental effect on the shape recovery ratio. Cell Viability and in Vitro Degradation. The ability to support continued attachment and proliferation of cells is the key feature of a biomaterial. NIH3T3 cells were used as reference cells to evaluate the biocompatibility of synthesized BPEs. Figure 9 shows the viability of the seeded cells on the various substrates for 1, 4, and 7 days compared with those on TCPS. The viability of the cells seeded on TCPS at day 1 was used as the reference value to calculate the relative viability. The viability of the attached and proliferated cells on the surfaces of these substrates generally increased significantly with increasing culturing time, showing the good biocompatible nature of synthesized BPEs. The continually increased CCK-8 values showed that BPEs exhibited no cytostatic property (i.e., inhibition of cell proliferation) toward 1318
dx.doi.org/10.1021/bm2000378 |Biomacromolecules 2011, 12, 1312–1321
Biomacromolecules
ARTICLE
Figure 10. Representative SEM images of NIH3T3 cell that had been seeded or cultured on (a) D0, (b) D40, and (c) D70 film substrates for 7 days.
NIH3T3 fibroblast cells up to 1 week. The growth rate of cells on D0, D40 and D70 was slower compared with the control assay at the same time interval. This just showed the difference of biocompatibility degree of BPEs with TCPS. Similar results were found by Sangsanoh et al. on the viability of different cells on chitosan micro- and nanoscale fibers compared with TCPS.49 It was found that the viability of the cells on the surfaces of D0, D40, and D70 first increased (day 1) and then decreased (days 4, 7) with increasing DEG molar fraction. DEG seems to favor of NIH3T3 cell attachment but not proliferation. The toxicity of DEG (human fatal dose is ∼1 mL/kg after ingestion)50 is a great concern when it is used in biomedical materials, although some recent studies of biomedical materials involved the utilization of DEG.5153 A recent study showed that the mechanism for the toxicity results from metabolites of DEG (2-hydroxyethoxyacetic acid) and not DEG itself.54 To test the availability of trace DEG or its metabolites 2-hydroxyethoxyacetic acid in the degradants of BPE, the ESI/MS is performed on the degradants of D70, which was accumulated in PBS for 36 days. After careful inspection of the mass spectra of the degradants (Figure S6 of the Supporting Information), irrespective of the scanning direction, no traces of DEG and 2-hydroxyethoxyacetic acid were detected. Only diethylene glycol sebacate unit (m/z 288.5), 1,3-propanediol sebacate unit (m/z 258.5), and SA unit (m/z 200.5) were found. That is to say, only lower esters of DEG are formed during the degradation of BPE in PBS, and they could not be further degraded into DEG. Consequently, the biocompatibility of BPEs is not affected by the introduction of DEG, and BPEs are believed to be safe for biomedical applications. Cell viability assay (MTT or CCK-8) can demonstrate only one aspect of material’s biocompatibility by using mitochondria activity of viable cells. Phenotype and interaction of cells need using morphological techniques to evaluate. Figure 10ac represents the SEM micrographs of fibroblasts cultured for 7 days on D0, D40, and D70. Cells grown on both types of D0 and D40 substrates were well-expanded in typical spindle morphologies and forming intercellular tight junctions with adjacent cells. The proliferated cells also appeared to grow on top of one another, forming cellular clusters. Although the cell grown on the D70 substrates also formed cellular clusters, the typical spindle morphology was not evident. This may be due to the different surface chemistry and crystallinity of D70 with more DEG, which reduces the CCK-8 value from Figure 9. From the results above, the D0 and D40 substrates supported the attachment and proliferation of NIH3T3 cells very well. It can be concluded that BPEs are potentially biocompatible from the viability and SEM results. Relatively easy degradation in PBS at 37 °C of the SMPs was observed. Figure 11 depicts the weight loss evolution of the SMPs up to 50 days. It can be seen that all SMPs lose their weight in PBS gradually, and the degradation of D70 is much faster than that for D0 and D40. Although BPE was cross-linked
Figure 11. Weight loss of BPEs in PBS at 37 °C as a function of time.
by carboncarbon bonds, the backbone of the polyester is still susceptible to PBS. The gel fractions of BPEs before and after degradation were determined (Table S3 of the Supporting Information). The substantially decreased gel fraction after degradation evidently demonstrates the bulk degradation of BPEs. According to Lendlein’s results,13 the beginning cleavage of ester bonds led only to an increase in the swelling capability. During this period, the swollen and partially degraded chain segments were still connected to multifunctional netpoints. A considerable amount of the degradant in this study could not be dissolved in PBS. However, it could be dissolved in chloroform. Consequently, the determined degradation in PBS was underestimated. Lendlein et al. found that the polyurethanes based on polyester even had a longterm induction of degradation up to 2 months, in which the degradation took place in fact.13 It is well known that the hydrolysis of semicrystalline polyester is highly dependent on the crystallinity. The higher the crystallinity, the lower mass loss rate the polyester possesses. At 37 °C, D0 and D40 are crystallizable and D70 is amorphous from Figure 2. As a consequence, the mass loss of D70 is much faster than that of D0 and D40. The demonstrated biodegradability of BPEs, together with the potential biocompatibility, renders them as potential candidates for biomedical devices such as different stents.
’ CONCLUSIONS BPEs were successfully prepared with PDO, SA, and IA, which have been industrially produced via fermentation or extraction 1319
dx.doi.org/10.1021/bm2000378 |Biomacromolecules 2011, 12, 1312–1321
Biomacromolecules with large quantities as the main raw materials. The resulted BPEs were cross-linked to form SMPs with excellent shape recovery and fixity (near 100% and independent of thermomechanical cycles). The tunable switching temperature and shape recovery speed were realized by introducing DEG to BPE chains and varying the curing extent of BPEs. The switching temperature is typical for biomedical applications in human body. In vitro fibroblast response and degradation demonstrated that the SMPs from BPEs were potentially biocompatible and biodegradable, rendering them as potential candidates for biomedical devices such as different stents.
’ ASSOCIATED CONTENT
bS
Supporting Information. Tg values, SR/SF values, tensile properties, and gel fractions of BPEs, DSC, and WAXD traces of BPEs with variable DCP contents, and ESI/MS spectrum of the degradants. This material is available free of charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION Corresponding Author
*E-mail:
[email protected] (B.G.);
[email protected]. cn (L.Z.).
’ ACKNOWLEDGMENT This work was supported by National Natural Science Foundation of China (50933001), National Outstanding Youth Science Fund (50725310), Fundamental Research Project for the Central Universities (2009ZZ0007), and National Basic Research Program of China (2011CB606002). ’ REFERENCES (1) Zhou, J. W.; Schmidt, A. M.; Ritter, H. Macromolecules 2010, 43, 939–942. (2) Ahn, S. K.; Deshmukh, P.; Kasi, R. M. Macromolecules 2010, 43, 7330–7340. (3) Liu, C. D.; Chun, S. B.; Mather, P. T.; Zheng, L.; Haley, E. H.; Coughlin, E. B. Macromolecules 2002, 35, 9868–9874. (4) Liu, G. Q.; Ding, X. B.; Cao, Y. P.; Zheng, Z. H.; Peng, Y. X. Macromolecules 2004, 37, 2228–2232. (5) Cao, Y. P.; Guan, Y.; Du, J.; Luo, J.; Peng, Y. X.; Yip, C. W.; Chan, A. S. C. J. Mater. Chem. 2002, 12, 2957–2960. (6) Weiss, R. A.; Izzo, E.; Mandelbaum, S. Macromolecules 2008, 41, 2978–2980. (7) Lendlein, A.; Kelch, S. Angew. Chem., Int. Ed. 2002, 41, 2034–2057. (8) Behl, M.; Lendlein, A. Mater. Today 2007, 10, 20–28. (9) Behl, M.; Razzaq, M. Y.; Lendlein, A. Adv. Mater. 2010, 22, 3388–3410. (10) Serrano, M. C.; Chung, E. J.; Ameer, G. A. Adv. Funct. Mater. 2010, 20, 192–208. (11) Yakacki, C. M.; Gall, K. Adv. Polym. Sci. 2010, 226, 147–175. (12) Xue, L.; Dai, S. Y.; Li, Z. Macromolecules 2009, 42, 964–972. (13) Alteheld, A.; Feng, Y. K.; Kelch, S.; Lendlein, A. Angew. Chem., Int. Ed. 2005, 44, 1188–1192. (14) Ni, X. Y.; Sun, X. H. J. Appl. Polym. Sci. 2006, 100, 879–885. (15) Lee, K. M.; Knight, P. T.; Chung, T.; Mather, P. T. Macromolecules 2008, 41, 4730–4738. (16) Knight, P. T.; Lee, K. M.; Qin, H.; Mather, P. T. Biomacromolecules 2008, 9, 2458–2467. (17) Knight, P. T.; Lee, K. M.; Chung, T.; Mather, P. T. Macromolecules 2009, 42, 6596–6605.
ARTICLE
(18) Knight, P. T.; Kirk, J. T.; Anderson, J. M.; Mather, P. T. J. Biomed. Mater. Res., Part A 2010, 94A, 333–343. (19) Cai, W.; Liu, L. L. Mater. Lett. 2008, 62, 2171–2173. (20) Lu, X. L.; Sun, Z. J.; Cai, W.; Gao, Z. Y. J. Mater. Sci.: Mater. Med. 2008, 19, 395–399. (21) Migneco, F.; Huang, Y. C.; Birla, R. K.; Hollister, S. J. Biomaterials 2009, 30, 6479–6484. (22) Yamashiro, M.; Inoue, K.; Iji, M. Polym. J. 2008, 40, 657–662. (23) Gautrot, J. E.; Zhu, X. X. Macromolecules 2009, 42, 7324–7331. (24) Kim, Y. B.; Chung, C. W.; Kim, H. W.; Rhee, Y. H. Macromol. Rapid Commun. 2005, 26, 1070–1074. (25) Nijst, C. L. E.; Bruggeman, J. P.; Karp, J. M.; Ferreira, L.; Zumbuehl, A.; Bettinger, C. J.; Langer, R. Biomacromolecules 2007, 8, 3067–3073. (26) Nair, D. P.; Cramer, N. B.; Scott, T. F.; Bowman, C. N.; Shandas, R. Polymer 2010, 51, 4383–4389. (27) Metcalfe, A.; Desfaits, A. C.; Salazkin, I.; Yahia, L.; Sokolowski, W. M.; Raymond, J. Biomaterials 2003, 24, 491–497. (28) Yakacki, C. M.; Shandas, R.; Lanning, C.; Rech, B.; Eckstein, A.; Gall, K. Biomaterials 2007, 28, 2255–2263. (29) Feng, Y. K.; Lu, J. A.; Behl, M.; Lendlein, A. Macromol. Biosci. 2010, 10, 1008–1021. (30) Standard Test Method for in Vitro Degradation Testing of Hydrolytically Degradable Polymer Resins and Fabricated Forms for Surgical Implants; ASTM F1635-04a; ASTM International: West Conshohocken, PA, 2004. (31) Chen, C. H.; Lu, H. Y.; Chen, M.; Peng, J. S.; Tsai, C. J.; Yang, C. S. J. Appl. Polym. Sci. 2009, 111, 1433–1439. (32) Chen, M.; Chang, W. C.; Lu, H. Y.; Chen, C. H.; Peng, J. S.; Tsai, C. J. Polymer 2007, 48, 5408–5416. (33) Fuller, C. S.; Frosch, C. J.; Pape, N. R. J. Am. Chem. Soc. 1942, 64, 154–160. (34) Santa Cruz, C.; Balta Calleja, F. J.; Zachmann, H. G.; Chen, D. J. Mater. Sci. 1992, 27, 2161–2164. (35) Tang, Z. F.; Wang, M. H.; Zhao, Y. N.; Wu, G. Z. Wear 2010, 269, 485–490. (36) Sen, A. K.; Mukherjee, B.; Bhattacharyya, A. S.; Sanghi, L. K.; De, P. P.; Bhowmick, A. K. Thermochim. Acta 1990, 157, 45–59. (37) Gao, J. M.; Lu, Y. J.; Wei, G. S.; Zhang, X. H.; Liu, Y. Q.; Qiao, J. L. J. Appl. Polym. Sci. 2002, 85, 1758–1764. (38) Jiao, C. M.; Wang, Z. Z.; Liang, X. M.; Hu, Y. Polym. Test. 2005, 24, 71–80. (39) Vaughan, A. S.; Zhao, Y.; Barre, L. L.; Sutton, S. J.; Swingler, S. G. Eur. Polym. J. 2003, 39, 355–365. (40) Neuss, S.; Blomenkamp, I.; Stainforth, R.; Boltersdorf, D.; Jansen, M.; Butz, N.; Perez-Bouza, A.; Knuchel, R. Biomaterials 2009, 30, 1697–1705. (41) Millesi, H.; Zoch, G.; Reihsner, R. Clin. Orthop. Relat. Res. 1995, 314, 76–83. (42) Sung, H. W.; Chang, Y.; Chiu, C. T.; Chen, C. N.; Liang, H. C. Biomaterials 1999, 20, 1759–1772. (43) Zhang, C. S.; Ni, Q. Q. Compos. Struct. 2007, 78, 153–161. (44) Merline, J. D.; Nair, C. P. R.; Gouri, C.; Shrisudha, T.; Ninan, K. N. J. Mater. Sci. 2007, 42, 5897–5902. (45) Jeong, H. M.; Lee, J. B.; Lee, S. Y.; Kim, B. K. J. Mater. Sci. 2000, 35, 279–283. (46) Ivens, J.; Urbanus, M.; De Smet, C. eXPRESS Polym. Lett. 2011, 5, 254–261. (47) Chung, T.; Rorno-Uribe, A.; Mather, P. T. Macromolecules 2008, 41, 184–192. (48) Kim, B. K.; Lee, S. Y.; Xu, M. Polymer 1996, 37, 5781–5793. (49) Sangsanoh, P.; Suwantong, O.; Neamnark, A.; Cheepsunthorn, P.; Pavasant, P.; Supaphol, P. Eur. Polym. J. 2010, 46, 428–440. (50) Schep, L. J.; Slaughter, R. J.; Temple, W. A.; Beasley, D. M. G. Clin. Toxicol. 2009, 47, 525–535. (51) Boch, R.; Canaan, A. J.; Cho, A.; Dolphin, D. D.; Hong, L.; Jain, A. K.; North, J. R.; Richter, A. M.; Smits, C.; Sternberg, E. D. Photochem. Photobiol. 2006, 82, 219–224. 1320
dx.doi.org/10.1021/bm2000378 |Biomacromolecules 2011, 12, 1312–1321
Biomacromolecules
ARTICLE
(52) Rouabhia, M.; Gilbert, V.; Wang, H. X.; Subirade, M. Biomed. Mater. 2007, 2, S38–S44. (53) Hu, F. Q.; MacRenaris, K. W.; Waters, E. A.; Liang, T. Y.; Schultz-Sikma, E. A.; Eckermann, A. L.; Meade, T. J. J. Phys. Chem. C 2009, 113, 20855–20860. (54) Besenhofer, L. M.; Adegboyega, P. A.; Bartels, M.; Filary, M. J.; Perala, A. W.; McLaren, M. C.; McMartin, K. E. Toxicol. Sci. 2010, 117, 25–35.
1321
dx.doi.org/10.1021/bm2000378 |Biomacromolecules 2011, 12, 1312–1321
