High-Pressure Compression-Molded Porous Resorbable Polymer

Article pubs.acs.org/journal/abseba

High-Pressure Compression-Molded Porous Resorbable Polymer/ Hydroxyapatite Composite Scaffold for Cranial Bone Regeneration Jin Zhang,†,‡,§ He Liu,‡,⊥,§ Jian-Xun Ding,*,‡ Jie Wu,∥ Xiu-Li Zhuang,‡ Xue-Si Chen,‡ Jin-Cheng Wang,*,⊥ Jing-Bo Yin,∥ and Zhong-Ming Li*,† †

College of Polymer Science and Engineering, State Key Laboratory of Polymer Materials Engineering, Sichuan University, Chengdu 610065, Sichuan, P. R. China ‡ Key Laboratory of Polymer Ecomaterials, Changchun Institute of Applied Chemistry, Chinese Academy of Sciences, Changchun 130022, P. R. China ⊥ Department of Orthopedics, Second Hospital of Jilin University, Changchun 130041, P. R. China ∥ Department of Polymer Materials, Shanghai University, Shanghai 200444, P. R. China S Supporting Information *

ABSTRACT: Fabricating porous scaffolds with sufficient mechanical properties is a challenge for healing bone defects. Highpressure compression-molded (HPCM) porous composite scaffold comprising poly(L-lactide) (PLLA), poly(lactide-co-glycolide) (PLGA), and hydroxyapatite (HA) was prepared and showed upregulated mechanical properties due to a solid network structure and a highly ordered crystalline architecture. The compressive yield strength and modulus of the HPCM scaffold molded at 1000 MPa and 180 °C were 0.91 and 6.84 MPa, respectively. The HPCM scaffold also exhibited an interconnected porous architecture with porosity greater than 80%, an appropriate degradation rate, and enhanced cell proliferation. Moreover, the HPCM scaffold supported the healing of a rat calvarial defect in vivo. KEYWORDS: high-pressure compression molding plus salt leaching, porous composite scaffold, high modulus, cranial bone regeneration μm in size.5 Guarino et al. reinforced poly(ε-caprolactone) (PCL) scaffolds with tricalcium phosphate and polylactide (PLA) fibers, and they discovered that the maximum elastic modulus of the scaffold with a porosity of 80% and pore size near 200 μm was 2.21 ± 0.24 MPa.6 Although the researchers attempted to improve the mechanical properties of porous scaffolds, the enhancement was always unobvious and deficient. This problem has caught the attention of the United States National Committee on Biomechanics, which has spearheaded an initiative to increase awareness about the mechanical requirements for repairing bone defects.7 It was reported that potential scaffolds must have a mechanical strength as close as possible to that of the substituted bone;8 otherwise, the integrity of the porous structure is likely to be destroyed during the tissue regeneration. In this context, sacrificing overall porosity is a viable approach to improve the scaffold strength for matching some site-specific requirements. However, with the decrease in pore number and size, the internal flow required for cell colonization

1. INTRODUCTION Porous scaffolds, seeded cells, and bioactive factors are now recognized as three basic components of tissue regeneration.1 Compared with traditional organ implantation, scaffolds can serve as the structural support to provide enough space for the formation of extracellular matrix (ECM), effectively overcoming the problems of immune rejection and the lack of available donors. It is well-established that the scaffold architecture plays an important role in determining the degree of tissue ingrowth. An ideal engineering scaffold should meet several criteria, such as having an interconnected porous structure and a controlled degradation rate commensurate with tissue remodeling.2 Additionally, the mechanical strength is another crucial property of scaffolds, especially for the replacement of load-bearing bone. Most recent studies have focused on the optimization of scaffold design from a biochemical point of view3 but have ignored a severe challenge: currently available porous composite scaffolds have poor mechanical properties, including insufficient compressive strength and low elastic moduli.4 Ma et al. once reported that the compressive modulus of poly(Llactide) (PLLA) scaffolds were within the range of 23.7−50.8 kPa, as the porosity is about 95%, and the pores were 250−420 © XXXX American Chemical Society

Received: April 16, 2016 Accepted: July 26, 2016

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DOI: 10.1021/acsbiomaterials.6b00202 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering and nutrient transportation might be hindered.9 Extending this point of view, some experimental investigations using particulate inclusions to strengthen polymer matrices have been actively pursued, such as the addition of tricalcium phosphate, high-strength fiber, and other reinforcements.10−13 In all cases, the interface adhesion between fillers and polymer matrix is the major factor affecting the mechanical properties of porous scaffolds. Thereby, surface modification of the reinforcements is required to promote compatibility and the load transfer from the polymer matrix to the fillers. These challenges in preparing a high modulus scaffold for tissue repair have motivated investigation of high-performance scaffolds that comprehensively promotes the regeneration of load-bearing bone tissue. High-pressure molding of polymers is able to drastically compact the stacking structure and optimize the crystalline architecture and, accordingly, eliminate intrinsic structure defects and optimize mechanical performance.14 For example, high-pressure-crystallized PLA has an approximately 16 times the storage modulus of the low-pressure compression-molded sample.15 Therefore, we attempt to structure a porous poly(Llactide) (PLLA)/poly(lactide-co-glycolide) (PLGA)/hydroxyapatite (HA) composite scaffold using high-pressure molding with the goal of enhancing the mechanical performance. As expected, the high-pressure compression-molded (HPCM) sample has denser architecture and more impressive mechanical performance than the low-pressure compression-molded (LPCM) scaffold. Scaffolds that have excellent mechanical performance can be used in the reconstruction of calvarial defects, which is an imperative but very challenging mission. Significantly, the results of in vitro cell culture and in vivo cranial bone repair adequately demonstrate that the HPCM composite scaffold enhances the cranial reparation much better than the LPCM scaffold, based on the mineralization measurements, mechanical properties, and synthesis of proteins that are involved in osteogenesis. Taken together, this study reveals the important benefits of a potential candidate for tissue engineering scaffolds, especially in the field of cranial bone regeneration.

samples were held at this temperature and pressure for an established time (10, 45, 90, or 180 min). Finally, the sample was cooled to room temperature at a rate of 2.5 °C min−1, and the pressure was released. The interconnected porous structure was acquired by leaching the molded samples with distilled water for more than 24 h until the weight remained constant. The overall properties of the porous composite scaffolds were absolutely optimal when the mass ratio of PLLA/PLGA/HA was 5:5:2.5. This optional condition was determined primarily by the degradation behavior, hydrophilicity, thermal properties, and the mechanical performance (Figures S1−S4). 2.3. Physical Properties of Porous PLLA/PLGA/HA Composite Scaffolds. The compressive modulus and yield strength of the porous PLLA/PLGA/HA scaffolds were determined using an Instron universal test instrument (model 5576, Instron Instruments, USA) in ambient atmospheric condition (25 °C and 50% relative humidity, RH) using a 1000N load cell. A constant crosshead speed of 0.5 mm min−1 was used. The porous cylinders were cut and trimmed to obtain an approximate diameter of 10.0 ± 0.5 mm and thickness of 15.0 ± 0.5 mm. The modulus values of the scaffolds were calculated as the slope of the stress−strain curves within the linear regions (strain range, 3− 10%). Compressive yield strength was defined as the stress carried at the yield point (10% strain). At least five specimens were tested for each condition, and the results were averaged. Morphologies of the porous scaffolds were examined by field emission scanning electron microscope (SEM, Inspect-F, FEI, Finland), operating at high vacuum and with an accelerating voltage of 20 kV. The specimens were frozen in liquid nitrogen for approximately 15 min and were then quickly fractured to expose the interior structures. Prior to SEM examination, the freshly fractured surfaces were sputtered with a thin layer of gold. The average pore sizes of the porous scaffolds were calculated based on the SEM results using the image analysis software Nano Measurer 1.2 (Fudan University, Shanghai, P. R. China). A quantitative estimation of other pore-related parameters was performed using a gravimetric method,16,17 as shown in the Supporting Information. The melting behavior was studied using a TA-Q2000 V7.3 differential scanning calorimeter (DSC, TA Instruments, USA). Approximately 5.0 mg of samples were enclosed in aluminum pans, and then heated from 40 to 190 °C at a constant speed of 10 °C min−1. The degradation behavior of the porous PLLA/PLGA/HA scaffolds was tested in a vial containing 10.0 mL of phosphate-buffered saline (PBS, 0.01 M, pH 7.4). Afterward the vial was incubated at 37 °C with constant shaking rate of 75 rpm. The incubation media was replaced every week, and the degradation studies were lasted for 12 weeks. At selected time points, the samples were carefully withdrawn from the medium and thoroughly washed with running distilled water, and dried in vacuum for 24 h to remove the residual water. The measured weights of the samples were normalized against their initial mass to assess the fraction of weight loss, which was calculated using eq 1.

2. MATERIALS AND METHODS 2.1. Materials. PLLA (viscosity-average molecular weight (Mη) = 7.9 × 104 Da) and PLGA (LA:GA = 8:2, Mη = 10 × 104 Da) were provided by Changchun SinoBiomaterials Co., Ltd. (Changchun, P. R. China). HA particles was obtained from Chengdu Xinjin Longma Chemical Co., Ltd. (Chengdu, P. R. China). Through the laser diffraction particle size analyzer (Mastersizer 2000, Malvern Instruments Ltd., UK), it was revealed that HA formed some small agglomerates with mean diameter of 6.5 μm. Dichloromethane (CH2Cl2), ethyl alcohol (C2H5OH), and commercial sodium chloride (NaCl) particles sieved in a specific range from 100 to 200 μm were purchased from Chengdu Kelong Chemical Reagent Factory (Chengdu, P. R. China). 2.2. Fabrication of Porous PLLA/PLGA/HA Composite Scaffolds. Solution coagulation was employed to achieve a good dispersion of HA in the PLLA/PLGA matrix. Then, the weighted NaCl particles with diameters of 100−200 μm as the porogen were added into the melted PLLA/PLGA/HA mixture (9:1, w/w) in an internal mixer at 180 °C and 50 rpm. The obtained mixture was melt compression molded using a homemade high-pressure compression molding apparatus.15 At first, a given amount of the PLLA/PLGA/ HA/NaCl mixture was placed inside the channel of the mold, heated to a predetermined temperature (170, 180, 190, or 200 °C), and kept for 15 min. Then, a pressure profile of 5, 300, 640, or 1000 MPa was applied to the sample within 2 min. To achieve a steady state, these

Weight Loss =

W0 − Wt × 100% W0

(1)

where W0 and Wt are the weights of the scaffolds before and after degradation, respectively, for a specific time interval. The weight loss, which was averaged for at least three specimens, was recorded. 2.4. Cytocompatibility of Porous PLLA/PLGA/HA Composite Scaffolds. Osteoblast-like MC3T3-E1 cells were cultured in αmodified Eagle’s medium (α-MEM) containing 10% (v/v) fetal bovine serum (FBS; Gibco BRL, 30 Gaithersburg, MD, USA), which was renewed every other day during the experiment. The sterilized scaffolds were transferred into 12-well tissue culture plates and immobilized in a humidified incubator at 37 °C with 5% (v/v) carbon dioxide (CO2). The morphologies of the cells cultured on the scaffolds were observed using SEM. After incubation for 3 days, the cells were fixed with 2.5% (w/v) glutaraldehyde in PBS for 2 h at 4 °C. After being thoroughly washed with PBS, the samples were dehydrated in a series of aqueous ethanol solutions of increasing concentrations. Finally, the scaffolds were dried overnight, coated with a thin layer of gold, and examined using SEM. B


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Figure 1. Compression testing results for porous scaffolds produced under different (A) pressures, (B) temperatures, and (C) time points. The statistical data are presented as the mean ± standard deviation (n = 3; *p < 0.05, **p < 0.01, and ***p < 0.001). The quantitative amounts of MC3T3-E1 cells were evaluated by adding CCK-8 solution (Dojindo, Japan) to each well. After incubation for 4 h, the absorbance of the resultant medium at 450 nm was measured using a Bio-Rad microplate reader (model 550, Hercules, CA, USA). The absorbance value at 600 nm was utilized for baseline correction. The relative cell proliferation was calculated with respect to the outcome of the first day. 2.5. In Vivo Evaluation of Porous PLLA/PLGA/HA Composite Scaffolds. All animal experiments were performed in compliance with the principles of Jilin University of Medicine and the National Institutes of Health (P. R. China). A total of 18 male Sprague−Dawley (SD) rats weighting 300−400 g were randomly assigned into one of three groups: HPCM scaffold group, LPCM scaffold group, and a control group that received no treatment after creating a full thickness calvarial defect. The normal tissue refers to the uninjured cranial bone. The rats were anesthetized using an intraperitoneal injection of chloral hydrate at a dosage of 30.0 mg per kg body weight. A sagittal incision of approximately 4 cm was then made on the skull to expose the cranial bone. Then, a 6 mm circular hole was created in the front midline of the skull. The porous PLLA/PLGA/HA scaffolds with a diameter of 6 mm and thickness of 1 mm were carefully implanted into each full-thickness calvarial defect without damaging the blood vessels or brain. The animals were euthanized at 6 or 12 weeks postoperation, afterward the bone specimens with a size of 8.0 × 8.0 mm2 were harvested and photographed. The thickness of the parietal was measured using a micrometer, and the cranial bone density was calculated by measured mass and volume. All samples were examined by X-ray (1.2 s, 50 kV, 5.4 mA, 50 cm projection) to detect the density of the regenerated tissues. To determine the critical element contents (i.e., Ca and P) involved in the bone, a nitric acid solution containing the cranium was heated to 60 °C and evaluated using inductively coupled plasma-optical emission spectrometer (ICP-OES).

To estimate mechanical stability of the regenerated skull, bone specimens with an average size of 2.0 × 2.0 mm2 at the sixth or 12th week were measured using nanoindentation (Tribo indenter, Hysitron TI-900, USA). The loading tip used in this work was a diamond Berkovich tip that had a three-sided pyramidal shape as shown in Figure S9. The depth of the indentation ranged from several hundred nanometers to a maximum of 4 μm. Typically, the radius across the indentation was 1 to 5 μm. The elastic modulus and hardness were determined from the unloading force/displacement slope at the maximum load and the projected contact area at this load. For each type, at least five samples were tested to obtain the average modulus values. For the histological examinations, samples from each group were randomly selected and fixed with 4% (w/v) PBS-buffered paraformaldehyde for 48 h. The fixed cranial bones were decalcified for 21 days in 10% EDTA/HCl at pH 7.2, dehydrated through a graded series of ethanol, and then embedded in paraffin. Then, 5 μm thick sections were cut, and assessed using hematoxylin and eosin (H&E) staining and Masson’s trichrome staining. Histological features of the regenerated bones were observed under an optical microscopy (Sichangyue Optical Instrument Co., Ltd., Shanghai, P. R. China). The immunohistochemical staining was performed to evaluate the expression of type I collagen, bone morphogenetic protein-2 (BMP-2), and vascular endothelial growth factor (VEGF). First, the frozen sections were deparaffinized with xylene and hydrated. Blocking of endogenous peroxidase activity was performed by incubation with 3% (v/v) hydrogen peroxide for 15 min. Immunohistochemical studies were conducted using primary antibodies (Santa Cruz, CA) against PBS-0.1% bovine serum albumin (1:200 dilution). After 12 h of incubation at 4 °C, the tissue slides were subsequently treated with horseradish peroxidase (HRP)-conjugated biotinylated secondary antibody (DAKO, Carpinteria, CA) for 30 min at a dilution of 1:100. Immunoreactivity was visualized with diaminobenzidine tetraC


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Figure 2. (A) SEM morphologies, (B) DSC heating scans, and (C) weight loss trends of (a, a′) HPCM and (b, b′) LPCM scaffolds. hydrochloride (DAB). The histological sections were also observed and photographed under an optical microscope. 2.6. Statistical Analysis. All data are reported as the mean ± standard deviation. SPSS 13.0 was used to analyze the data using oneway and two-way analysis of variance with Tukey’s posthoc test. A probability value of less than 0.05 was considered significantly different, and a probability value below 0.01 or 0.001 was considered highly significantly different.

tissue-engineered constructs, we attempt to achieve a highly densified porous structure by regulating the pressure, temperature, and exposure time of the high-pressure molding, as shown in Figure 1. With increasing pressure, a noteworthy increase in the mechanical properties of the scaffolds was achieved. The high pressure of 1000 MPa resulted in a considerable improvement in the compressive yield strength and the modulus, which changed from 0.23 and 1.19 MPa to a maximum of 0.75 and 5.91 MPa, respectively. These values were approximately three to five times greater than those of the low-pressure compression-molded samples, suggesting a prominent effect of high pressure on enhancing the structural

3. RESULTS AND DISCUSSION 3.1. Mechanical Performances and Microstructures of Porous Composite Scaffolds. To preserve the structural integrity of the scaffolds and to improve the functionality of the D


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particle content and are thus almost invariable despite changes in the processing pressure. Figure 2B shows the DSC heating scans of the porous composite scaffolds. Because of the relatively slow crystallization kinetics of PLA,19 an evident cold-crystallization peak at approximately 91.5 °C appears in the LPCM samples. This exothermic peak shifts toward a lower temperature of 86.3 °C in the HPCM sample, suggesting the excitation of cold crystallization by pressure-induced-primary nuclei.20 Notably, the melting point of the high-pressure crystallized PLA (170.3 °C) is approximately 9 °C higher than that of the low-pressure molded PLA. This result arises from the formation of some thicker lamellae in the HPCM scaffold.21 Correspondingly, the melting enthalpy of the high-pressure densified PLA (13.52 J g−1) is almost 2-fold greater than that of the LPCM (6.49 J g−1), indicating the improved structural tacticity. It is general knowledge that the specific structure of the macromolecules and their orderliness in the solid phase are two key factors influencing the mechanical properties of polymer materials.22,23 For instance, the crystallinity of high-pressure crystallized polyethylene can be as high as 90%. The closely packed molecular chains and some extended chain crystals result in a 2fold increase in the dynamic Young’s modulus.24 Overall, the high-pressure treatment facilitates the rearrangement of the polymer chains into crystals, and this more stable and ordered structure helps the porous scaffold acquire strong antideformation capability and excellent mechanical performance. Furthermore, the scaffold degradation rate must be similar to the rate of tissue formation, so that the 3D space can be replaced by new tissue. The weight losses of the HPCM and LPCM scaffolds in an aqueous PBS environment were tracked every week, as shown in Figure 2C. The PLLA/PLGA/HA composite scaffold formed under the low-pressure condition exhibits a weight loss of 77.38 ± 3.38% after 12 weeks of incubation, and the HPCM sample undergoes a relatively slower degradation rate with a weight loss of 67.21 ± 2.99% during the same immersion time. On one hand, in contrast to the stable and intact porous structure of the HPCM, the inconsecutive polymer skeleton of the LPCM scaffold facilitates the diffusion of the buffered solution, which results in the hydrolysis of the polymer matrices. On the other hand, it is widely accepted that a preferential hydrolysis takes place in the free amorphous region rather than in the restricted crystalline region,25 thus a faster biodegradation rate is observed in the LPCM specimen. Considering the existence of various enzymes, the complex environment of the human body and the autocatalytic effect of the accumulated acidic degradation products, it is reasonable to predict that the in vivo degradation rate is faster than the in vitro rate.26 In summary, both types of scaffolds exhibit appropriate degradation behaviors with more than 65% weight loss, which is an ideal rate that complies with the biomedical application requirements. 3.2. Cell Adhesion and Proliferation on Porous Composite Scaffolds. To elucidate the effect of high modulus on the attachment and growth of cells within the scaffolds, SEM and Cell Counting Kit-8 (CCK-8) assays were used to qualitatively and quantitatively evaluate the cell viability, respectively. As illustrated in Figure S7, after incubation for 24 h, MC3T3-E1 cells cultured in the HPCM and LPCM scaffolds are densely presented with a continuous layer of fibrous ECM. Additionally, both of the scaffolds exhibit high cell adhesion rates, verifying a good adhesion of MC3T3-E1 cells on the surface of porous composite scaffolds. The in vitro

stability of the scaffolds. A moderate processing temperature and time were best. The low temperature of 170 °C and a short duration of 10 min failed to provide sufficient time for the melting of the polymers along with the increased chain mobility. Conversely, high processing temperature greater than 190 °C and a long residence time beyond 90 min introduced problematic thermal degradation of the aliphatic ester structure, which severely impaired the mechanical properties of the scaffolds. Overall, the above data indicate that the variations in the processing parameters do have a significant influence on the mechanical properties of the composite scaffolds to a certain degree. The highest compressive performance was achieved when the scaffold was molded at 1000 MPa and 180 °C for 45 min, after which the compressive yield strength and modulus were 0.91 and 6.84 MPa, respectively. The compressive modulus here is comparable with that of the cranial bone of a 43-day-old adult rat (∼6 MPa at 25 °C),18 which highlights its great potential for repairing cranial bone. The porous composite scaffold reported below, i.e., the HPCM scaffold, was fabricated under such optimal processing conditions, and the LPCM specimen (5 MPa/180 °C/45 min) was determined for comparison. The cross-section morphologies of the porous composite scaffolds were observed using a high-resolution SEM, as shown in Figure 2A. It can be seen that the HPCM scaffold exhibits a rugged network structure and a high architecture integrity. Many interconnected open pores with an average pore size of approximately 145 μm are homogeneously interpenetrated in this three-dimensional (3D) skeleton. Such a compact and intact porous architecture not only has the ability to provide enough space for cell proliferation and nutrient transportation, but is also dependable with respect to load bearing due to its interconnected stable pores. In contrast, LPCM scaffold has a fragile polymer skeleton with some microcracks, suggesting a bad stability and weak combination of the cocontinuous structure. Specifically, in order to acquire the highly porous structure, the compression-molded mixtures are immersed in the distilled water for a predetermined duration of time. During the course, dissolution of the continuous NaCl phase destroys the integrity of PLLA/PLGA/HA skeleton in some degree. Especially for the LPCM scaffold with relatively weaker interaction force, the leaching process finally leads to the cracked edges of the pores, which is prone to collapse upon encountering external force. Quantitative estimation of the pore-related parameters for the HPCM and LPCM scaffolds was performed using gravimetric measurements,16,17 and the values of corresponding porosity, interconnectivity, and bulk density are shown in the table inserted in Figure S5. It is worth noting that the porosities of the composite scaffolds maintain a high level greater than 80%, which meets the requirement for tissue engineering. The interconnectivity value is approximately 97%, suggesting that the porogen within the mixtures is almost fully continuous and is desirable for permitting cell infiltration throughout the scaffolds. The density fluctuates at approximately 0.25 g cm−3, which is nearly the lowest density reported so far for PLLA or PLGA scaffolds with similar porosity and connectivity. These findings endow the scaffold a prominent superiority in terms of being lightweight. Additionally, variation in the processing pressure has no detectable influence on these pore parameters because the pores of the solid scaffolds are often created by the exclusion of the porogen in an aqueous environment. The pores are mainly a function of the salt E


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matrix remodeling are hence significantly enhanced by the rigidity of the scaffolds. 3.3. In Vivo Evaluation of Porous Composite Scaffolds for Cranial Bone Regeneration. The high-modulus scaffold fabricated herein offers a potential possibility for cranial bone regeneration. Figure 4A shows the gross examination of the

proliferation of MC3T3-E1 cells on the HPCM and LPCM scaffolds, Termanox coverslip, and cell culture plate were examined using CCK-8 tests, as shown in Figure 3. The cell

Figure 3. Relative cell proliferation of MC3T3-E1 cells after culture on HPCM and LPCM scaffolds, Termanox coverslip, and cell culture plate at predetermined time points. Statistical data are presented as the mean ± standard deviation (n = 3; *p < 0.05, **p < 0.01, and ***p < 0.001).

viabilities in both the HPCM and LPCM scaffold groups remain high, implying low scaffold toxicity toward MC3T3-E1 cells. The HPCM scaffold shows noticeable cell proliferation, which results in an approximately 4-fold increase in the cell number in relative to that of the first day. This is distinctly higher than the increase observed for the LPCM scaffold and even better than the increase observed for the Termanox coverslip. The efficient cell proliferation of the HPCM scaffold is primarily caused by the highly porous structure and the sufficient mechanical properties ensuring structural integrity.27 Additionally, it is well-known that cells generally have a favorable growth state on cell culture plates;28 the results, which are comparable to those of the control sample, fully demonstrate that the HPCM scaffold possesses high cytocompatibility and has great potential to effectively accelerate cell infiltration. The mechanical performance of the scaffold has already been demonstrated to have a significant influence on cellular physiology. Discher et al. reported that the stiffness of a substrate could regulate the differentiation, morphology, and spreading of various cell types.29 Likewise, other studies have shown that the enhanced mechanical strength of scaffolds helped direct cell activity and even accelerated tissue regeneration in vivo.30 Cell adhesion and migration in the scaffold always generate cell-exerted contractile forces by pulling the cytoskeleton toward the adhesion points. Some of these forces are transmitted to the substrate, which may lead to a deformation of the substrate and, in turn, elicit some changes in the protein composition of the focal adhesions.31 In terms of the LPCM sample with poorer mechanical performance, the original structure is easily damaged, and the dimensions often do not remain intact under the cellular contractile forces; thus, there is a resulting lack of cell attachment points and inadvertent impairment cell responsiveness. In sharp contrast, the high-modulus HPCM scaffold serves as a firm structural framework to support cell growth, differentiation, and metabolism, which are highly beneficial to the activities of encapsulated cells. Specifically, the osteoblasts are able to sense the mechanical signals imparted by their 3D scaffolds, and then the cells are allowed to exert their force on the surrounding ECM.32 Taken together, the cell spreading and migration, and

Figure 4. (A) Gross images and (B) X-ray images of regenerated skull in defects at 6 and 12 weeks postimplantation. The dotted circle indicates the unhealed bone tissue, and the scale bar represents 6 mm.

regenerated skull in the defects at 6 and 12 weeks postoperation. In the untreated control group, a scant amount of mineralization and bone healing were observed along the defect margins, but the center of the calvarial defect was only filled with some transparent fibrous tissue and did not form a complete defect closure. In direct contrast to the control group, the implantation of the LPCM scaffold into the defect enhanced cranial bone formation in vivo. The new skull with lamellar structure appeared at the edge of the defect at the sixth week, and the area of the remolded cranial bone expanded into the middle of the calvarial defect at 12 weeks postoperation. Most attractively, the HPCM scaffold displayed superior efficacy for stimulating the bone defect healing. After 6 weeks, new bone formation was observed over almost the entire area of the defect, and there was no significant difference between the repaired cranial tissue and the surrounding uninjured skull. As degradation proceeds, the weight loss of these scaffolds was accelerated, and they were almost completely absorbed after 12 weeks without any remnants or local complications. It must be stressed that both scaffolds have highly suitable degradation behavior, with an ideal rate typically F


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Figure 5. At 6 and 12 weeks postoperation, (A) tissue thicknesses, (B) bone mineral densities, and (C) calcium and (D) phosphorus contents of regenerated cranial bones in HPCM and LPCM scaffolds, and control groups (n = 5; *p < 0.05, **p < 0.01, and ***p < 0.001). The related parameters in the normal tissue are compared as a reference.

rising to 78.55 ± 1.66 mm and 0.91 ± 0.03 mg mm−3, respectively. These results indicate that the HPCM scaffold promotes osteogenesis in vivo. As is well-known, calcium (Ca) and phosphorus (P) are the main components of the skull, and the amounts of Ca and P represent the degree of the calvarial defect repair. The deposition of Ca and P within the regenerated cranial bone was detected by ICP-OES, as shown in Figure 5C, D. The HPCM scaffold group does promote the ingrowth of mineralized matrix into a bone defect, where the concentrations of Ca and P increase to the highest value of 1.24 ± 0.05 and 42.37 ± 2.41 mg g−1, respectively; these values represent almost 80% of the values observed in normal cranial tissue. More interestingly, the molar ratios of calcium to phosphate in the bones for the natural and experimental groups are listed in Table S1. The table shows that the ratios of calcium to phosphate exhibit a decreasing trend with the postimplantation time. This result is especially true for the HPCM group at the 12th week, in which the ratio decreases to 1.46. The value is similar to the normal tissue ratio of 1.48. On the basis of the calcium to phosphate ratios of the regenerated skull, it is reasonable to conclude that the HPCM scaffold highly mimics the mineral phase of natural bone, suggesting excellent osteogenesis during the calvarial defect healing. A nanoindentation test was conducted to evaluate the biomechanical performance of the regenerative skull, which is an essential indicator of bone healing in the presence of a

being greater than 65% at 12 weeks, which complies with the requirements of bone tissue engineering. To visualize and quantify the formation of a mineralized matrix within the defect area, X-ray microcomputed tomography was performed, and the radiographic images are shown in Figure 4B. Any radiopaque area in the defect indicates the formation of new skull.33 Both HPCM scaffold introduction and LPCM scaffold introduction sufficiently promoted ingrowth of mineralized matrix within the cranial bone defect, and the restoration began from the edge of the defect and progressed toward the center. At 6 weeks after implantation in vivo, the HPCM scaffold group demonstrated a considerable reduction in the size of the calvarial defect, whereas little mineralized matrix could be observed in the LPCM scaffold group and the untreatedcontrol group. The scan on week 12 clearly demonstrated that the HPCM scaffold possessed the highest density of mineralization compared to the other two groups, indicating the nearly complete healing of the cranial bone defect. The superiority of the HPCM scaffold can be further demonstrated by the tissue thickness and the bone mineral density, as shown in Figures 5A, B. By taking the results at the 12th week as an example, the tissue thickness and mean bone density in the control group are 52.85 ± 2.88 mm and 0.65 ± 0.04 mg mm−3, respectively, and these parameters increase to 65.57 ± 2.85 mm and 0.82 ± 0.01 mg mm−3 in the LPCM scaffold group, respectively. Intriguingly, the application of the HPCM scaffold results in an approximately 150% increase, G


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Figure 6. At 6 and 12 weeks postoperation, (A, B) load−distance curves of nanoindentation tests, (C) hardness, and (D) elasticity moduli of regenerated cranial bones in HPCM and LPCM scaffolds, and control groups (n = 5; *p < 0.05, **p < 0.01, and ***p < 0.001). The related parameters in the normal tissue are compared as a reference.

scaffold. By quantitatively analyzing the load-distance curves of the nanoindentation test, the hardness and elasticity modulus of the restored cranial bone for each group were precisely obtained. Figure 6A, B shows the load−distance curves at 6 and 12 weeks postimplantation, respectively. The hardness and elasticity modulus of the newborn skulls are calculated from the projected contact area at maximum load and the unloading force/displacement slope at this load, which are quantitatively presented in Figure 6C, D, respectively. As expected, the skull regenerated with the HPCM scaffold exhibited the highest hardness and elastic modulus values. This was followed by the LPCM scaffold group, and the lowest values were found for the control specimen. Specifically, the hardness for the HPCM and LPCM scaffolds and the control groups at week 12 were, on average, 0.51 ± 0.07, 0.25 ± 0.06, and 0.18 ± 0.01 GPa, respectively. In agreement with the variation in the hardness trend, the modulus elasticity was significantly higher in the HPCM scaffold group than in the other two groups and showed approximately 1.60 and 2.64 times more elasticity than the LPCM scaffold and control groups, respectively. Such excellent biomechanical performance in the HPCM scaffold group can be reasonably ascribed to the enhanced cell proliferation, the increased bone density, and the high amount of mineralized tissue. The histological evidence further supported the radiographic findings. Figure 7A shows the H&E staining results of the regenerated skull at the center of the 6 mm defects. In the case of the control group, the H&E staining at 6 weeks

postoperation indicated that the bone defects were largely filled with a cell layer and a fibrillar matrix, whereas a minimal amount regenerated cranial bone was observed after 12 weeks. In contrast, the LPCM scaffold induced the appearance of new cranial bone at 6 weeks after implantation. The regenerated skull was mixed with some sparse islands of cartilage plates at week 12, which is indicative of an endochondral ossification process. The poor integration of regenerative tissue could be ascribed to the lack of osteoblasts and significant fibrous tissue infiltration prior to the union of the implanted bone tissue with the native bone. Interestingly, the HPCM scaffold group demonstrated a considerable amount of organized and regularly mineralized tissue in the repaired area, which had a typical lamellar bone morphology and was similar to native bone as well as mature bone marrow. By 6 weeks, some relatively thick and dense wovenlike new bones had already formed. The entire hole in the cranial bone was almost covered by the compact bone after 12 weeks. Such mineralized tissue is closely related to the structural integrity of HPCM scaffold. The wellmaintained porous structure helps cells migrate and secrete signals that induced mineralization, which finally guided tissue regeneration. The above histological observation suggests that the HPCM scaffold readily facilitates the osteogenesis in vivo and calvarial defect healing. The Masson’s trichrome staining of the repaired cranial bone is shown in Figure 7B. Consistent with the results of the H&E staining, neither necrosis nor a foreign body reaction is observed in any of the groups at any time points. The tissue H


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Figure 7. (A) Histological morphologies of calvarial defect parts by H&E staining at 6 or 12 weeks postoperation: HPCM and LPCM scaffolds, and control groups. Magnification of the images is 100×. (B) Histological sections stained using Masson’s trichrome showing representative tissue reparation at defect sites at 6 or 12 weeks postimplantation. Collagenous fibers appear green, and the maturing bone is in red. The dotted line defines the region of the cranial bone defects. The magnifications of these images are 10× and 40×, respectively.

and the type I collagen staining is more diffuse. Type I collagen is the primary component of the extracellular matrix in bone tissue and is responsible for the remodeling phase. It is essential to ensuring tissue resistance. As a result, it is reasonable to conclude that the HPCM scaffold promotes a stronger production of osseous tissue by stimulating more type I collagen expression. BMP-2 and VEGF are integral in bone formation and fracture healing. The osteogenic factor BMP-2 can significantly improve osteogenic ability, whereas VEGF can stimulate angiogenesis and increase nutrient availability as is the best characterized angiogenic factor.34 Figure 8B, C shows the expressions of BMP-2 and VEGF in the regenerated skull. For each group, the expression of both growth factors achieves a maximum 12 weeks postoperation. The results herein are consistent with those of previous studies. As reported by Deckers et al., the transcription levels for BMP-2 and VEGF always achieve peak expression during the final stages of osteoblast differentiation.35 Noticeably, immunohistochemistry displays intensive BMP-2 and VEGF positive staining in the HPCM scaffold group, whereas endogenous staining is present in the LPCM scaffold and control groups but is much weaker. VEGF-mediated angiogenesis is a basic requirement for bone growth and can promote the recruitment of osteoclasts and improve the activity of osteoblasts.36 Transforming growth

healing improved with time in the calvarial defect model, and the ossification procedure always began from the periphery of the hole and progressed toward the center. However, the amount and structure of the new bone differed for each scaffold. Only a very small amount of bone along with a thin layer of new bone was formed in the control group at 12 weeks postoperation, and the cavity was only filled with the abundant collagen fibers. In clear contrast, healing of the bone defect was significantly increased in the experimental groups due to the implantation of scaffolds. The HPCM scaffold showed superior efficacy for remodeling the cranial cavity compared with the LPCM scaffold. Observation from the magnified images, which are the insets in Figure 7B, reveal more advanced bone formation in the HPCM scaffold group as evidenced by the larger area of the newly regenerated skulls. Figure 8A shows the immunohistochemical results for type I collagen in the regenerated cranial bone. With respect to all groups evaluated at 12 weeks, the deposition of type I collagen is greater than that at 6 weeks, suggesting that the state of collagen maturation is progressively advanced. More importantly, in the HPCM scaffold group, type I collagen is intensively stained and maintains a high level at 12 weeks postoperation. Conversely, the expression of type I collagen is predominantly moderate in the LPCM scaffold group and even discrete in the control group, where many defects are observed, I


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Figure 8. Immunohistochemistry of type I collagen (A), BMP-2 (B), and VEGF (C) expression in HPCM and LPCM scaffold, and control groups evaluated at 6 or 12 weeks postimplantation. Magnification of these images are 10× and 40×, respectively.

development potential of this scaffold for healing cranial bone defects. Therefore, further attempts are made to understand the contribution of a high-modulus scaffold to cranial bone regeneration. By implanting HPCM and LPCM scaffolds into the calvarial defects of rats for 6 or 12 weeks, it is demonstrated that the HPCM scaffold provides an interconnected porous network with sufficient mechanical strength. Compared with the LPCM scaffold, a significant increase in X-ray radiopacity, bone thickness and density, biomineralization, and mechanical properties is observed for the HPCM scaffold. More interestingly, the increased expression of BMP-2, VEGF, and type I collagen evaluated by histological staining further demonstrates that the HPCM scaffold can efficiently induce osteoconduction and osseointegration in vivo. Overall, a high-

factor of BMP-2 results in cell mineralization and the increased secretion of VEGF can also enhance the differentiation of MSCs to osteoblasts.5 Considering the critical influence of BMP-2 and VEGF on bone tissue remodeling, we tentatively conclude that the HPCM scaffold exhibits the best osteogenesis capacity and bone repair ability, which is due to the upregulation of the expression levels of BMP-2 and VEGF.

5. CONCLUSIONS A high-modulus PLLA/PLGA/HA composite scaffold is fabricated using a high-pressure compression molding/salt leaching method at 1000 MPa and 180 °C for 45 min. The compressive yield strength and modulus of HPCM scaffold are 0.91 and 6.84 MPa, respectively. The results fully reveal the J


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modulus composite scaffold represents encouraging progress in bone tissue reparation.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsbiomaterials.6b00202. Additional figures and details (PDF)

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. Tel./Fax: +86 431 8526 2116. *E-mail: [email protected]. Tel./Fax: +86 431 8879 6222. *E-mail: [email protected]. Tel./Fax: +86 28 8540 5731. Author Contributions §

J.Z. and H.L. contributed equally. All authors contributed to the execution of study. All authors revised the manuscript and approved to the final version. J.Z. and Z.M.L. were responsible for the research conception and provision of the study materials. J.X.D., X.L.Z., and X.S.C. contributed to the experimental design and manuscript revision. H.L., J.W., J.C.W., and J.B.Y. were involved in the execution of animal experiments. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors gratefully acknowledge the financial support provided by the Program of the National Natural Science Foundation of China (Grants 51120135002, 51421061, 51303174, 51273196, 51533004, and 51321062), the Polylactide Biodegradable Materials Innovation Center of Jilin Province (Grant 20160623026TC), and the Training Program of Outstanding Doctoral Students of Jilin University (Grant YB201501).

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REFERENCES

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High-Pressure Compression-Molded Porous Resorbable Polymer

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