Investigating the Potential of Amnion-Based Scaffolds as a BarrierMembrane for Guided Bone RegenerationWuwei Li,*,† Guowu Ma,† Bryn Brazile,‡ Nan Li,† Wei Dai,§ J. Ryan Butler,‡ Andrew A. Claude,‡

Jason A. Wertheim,∥ Jun Liao,*,‡ and Bo Wang*,†,∥

†Department of Oral and Maxillofacial Surgery, School of Stomatology, Dalian Medical University, Liaoning 116001, China‡Department of Biological Engineering and College of Veterinary Medicine, Mississippi State University, Mississippi State, Mississippi39762, United States§Department of Operation Room, The First Affiliated Hospital, Dalian Medical University, Liaoning 116001, China∥Comprehensive Transplant Center and Department of Surgery, Feinberg School of Medicine, Northwestern University, Chicago,Illinois 60611, United States

ABSTRACT: Guided bone regeneration is a new concept of large bone defect therapy, which employs a barrier membrane toafford a protected room for osteogenesis and prevent the invasion of fibroblasts. In this study, we developed a novel barriermembrane made from lyophilized multilayered acellular human amnion membranes (AHAM). After decellularization, theAHAM preserved the structural and biomechanical integrity of the amnion extracellular matrix (ECM). The AHAM also showedminimal toxic effects when cocultured with mesenchymal stem cells (MSCs), as evidenced by high cell density, good cell viability,and efficient osteogenic differentiation after 21-day culturing. The effectiveness of the multilayered AHAM in guiding boneregeneration was evaluated using an in vivo rat tibia defect model. After 6 weeks of surgery, the multilayered AHAM showedgreat efficiency in acting as a shield to avoid the invasion of the fibrous tissues, stabilizing the bone grafts and inducing themassive bone growth. We hence concluded that the advantages of the lyophilized multilayered AHAM barrier membrane are asfollows: preservation of the structural and mechanical properties of the amnion ECM, easiness for preparation and handling,flexibility in adjusting the thickness and mechanical properties to suit the application, and efficiency in inducing bone growth andavoiding fibrous tissues invasion.

1. INTRODUCTION

Barrier membranes have been wildly used in dental peri-boneimplanting1,2 and other bone fractures with large bone loss orwith poor healing potential.3 The main role of applying thebarrier membrane in bone surgery is to prevent the invasion offibroblasts, provide stability for bone grafts and blood clots, andensure a protected room for osteogenesis (also known asguided bone regeneration (GBR)).4,5 As an example in thedental field, patients with serious periodontal defects,dehiscences, and fenestrations around the region of implantor patients requiring bone augmentation generally need peri-bone implanting treatment with sufficient high-quality bone

grafts as well as a barrier membrane covering on top of theimplanting site.6−8

Two major types of the barrier membranes that arecommonly used in clinical treatment are resorbable andnonresorbable membranes. Nonresorbable membranes arebetter at space maintaining when compared with the resorbablemembranes, but these materials have high risk of infection andneed a second removal operation.9,10 Collagen barriermembrane, a type of natural resorbable membrane that iswidely used as a commercial product, has a bilayered structure

Received: March 13, 2015Published: July 9, 2015

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© 2015 American Chemical Society 8642 DOI: 10.1021/acs.langmuir.5b02362Langmuir 2015, 31, 8642−8653

consisting of a porous layer and a dense layer.11 Clinically indental application, after filling the bone grafts into the defectregion, the collagen barrier membrane is usually placed directlyover the grafted material, with the porous layer facing the bonegrafts to permit bone ingrowth, and the dense layer facing themucosa to avoid the invasion of fibrous tissues.12 However,there are still some drawbacks existing in the application of thecollagen barrier membrane in bone implantation.13 Collagenmembranes are animal-derived biomaterials; thus, they face therisk of disease transmission from animals to humans.14

Moreover, during the surgical and postoperative healing phases,this approach is still faced with challenges such as inflammatoryresponse, weak mechanical strength, and control of thedegradation rate.15,16

An ideal barrier membrane should have the advantages ofpreventing the fibrous tissue invasion, promoting boneregeneration, maintaining the bone defect margins, reducingthe associated complications and healing time, and beingabsorbed in an appropriate manner.17 Human amnionmembrane (HAM) is the interior part of human fetalmembranes, which consists of an epithelial layer, a basementmembrane, and a collagen stromal layer.18 The extracellularmatrix (ECM) components of the HAM include collagen,elastin, laminins, nidogen, fibronectin,19 proteoglycans,20,21 andnumerous growth factors such as EGF, KGF, HGF, andbFGF.22 HAM is found to have favorable biological propertiessuch as antimicrobial, anti-inflammatory, scar inhibiting, lowimmunogenicity, stimulating epithelialization, and woundhealing.19,23 In regenerative medicine, HAM has been used inocular surface reconstruction24−26 and partial-thickness burnwounds covering27−29 and as scaffold materials in tissueengineering.30,31 Moreover, HAM has recently been reportedas a suitable platform in facilitating osteogenic differentiationfor both stem cells32 and apical papilla cells.33 When coveringover the defects on maxillary and mandibular bone, the acellularHAM was found to promote injury healing and improve boneinduction.34

In this study, we created a barrier membrane made fromlyophilized multilayered acellular human amnion membranes(AHAM) and evaluated the potential of AHAM barrier

membrane in assisting bone implanting treatment. AHAMwas characterized to understand its ECM composition andbiomechanical properties, in vitro interaction with bonemarrow mesenchymal stem cells (MSCs), and osteogeniccapability. The effectiveness of the AHAM barrier membrane inguiding bone growth was evaluated via an in vivo rat tibia defectmodel.

2. MATERIALS AND METHODS2.1. HAM Decellularization and AHAM Barrier Membrane

Preparation. Fresh full-thickness HAM was collected after caesariandelivery from uncomplicated singleton pregnancy in the First AffiliatedHospital of Dalian Medical University (Liaoning, China). All motherswere seronegative for human immunodeficiency virus types I and II,human hepatitis B and C, and syphilis. Informed consents wereobtained from the mothers, and the IRB protocol was approved by theResearch Ethics Board of Dalian Medical University. The HAM wasdissected from the periplacental region, trimmed into 1.5 cm × 1.5 cmsquare pieces, and treated with 1% Triton X-100 (200 mL) for 2 h,distilled water (200 mL) for 15 min two times, and 0.1% sodiumdodecyl sulfate (SDS) (200 mL) for 10 h on a horizontal shaker(Thomas Scientific Inc., Swedesboro, NJ), followed by thorough PBSwashing.

To prepare single-layered lyophilized AHMA for structuralcharacterizations and in vitro cell culture studies, a piece of AHMA(1.5 cm × 1.5 cm) was spread and fixed with surgical sutures on acustom-made, square-shaped plastic frame (Figure 1C) and completelydehydrated with a Freeze-Dryer System (Cole-Parmer, Vernon Hills,IL) at −54 °C. To prepare the multilayered lyophilized AHAM for thein vivo rat experiments, eight pieces of the AHAM scaffolds werestacked into a multilayered patch. We then removed the air bubblesthat were trapped among the membrane layers with a roller. Similarly,the multilayered patch was sutured onto the custom-made, square-shaped plastic frame and completely dehydrated with the Freeze-DryerSystem.

2.2. Histology, Immunohistology, and SEM. Samples forhistology staining were fixed in 4% paraformaldehyde, subjected tosectioning, and stained with hematoxylin and eosin (H&E) andMovat’s Pentachrome. For immunofluorescence staining, afterdeparaffin, rehydration, and antigen retrieval, sample sections wereincubated with anticollagen type I, anticollagen type IV, andantifibronectin primary antibodies (1:200, Abcam, Cambridge, MA)at 4 °C overnight. Secondary antibodies of goat antimouse IgG-FITC(1:500, Santa Cruz Biotechnology, Santa Cruz, CA) were incubated at

Figure 1. (A) Native HAM, (B) AHAM, (C) lyophilized AHAM, (D) rehydrated AHAM, (E) H&E, and (F) Movat’s pentachrome staining of thenative HAM and AHAM (collagen, yellow; elastin, black; proteoglycans, light blue); (G) SEM images reveal the cross-sectional view, stromal layerview, and epithelial layer view of the HAM and AHAM.

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room temperature for 60 min. The sections were then stained with 1μg/mL DAPI (Invitrogen) for cell nuclei. The immunofluorescenceslides were observed with a fluorescence microscope (OlympusBX43). For scanning electron microscopy (SEM), the samples werefixed in 2.5% glutaraldehyde, dehydrated, critical point dried (PolaronE 3000 CPD), and sputter coated with gold−palladium for SEMobservation (JEOL JSM-6500 FE-SEM).2.3. In Vitro Cell Culture. Well-characterized bone marrow MSCs

(third passage) derived from Sprague−Dawley (SD) rats (product no.RASMX-01201, Cyagen Biosciences Inc., China) were resuspendedand seeded in 75 mm flasks at a density of 2 × 103 cells/cm2 withMSC medium (L-DMEM, 10% FBS, 100 U/mL penicillin and 100μg/mL streptomycin). MSCs were expanded, and the cells at the fifthpassage were used for AHAM recellularization and osteogenicdifferentiation studies. All cell culture was performed in the incubatorat 37 °C and 5% CO2 atmosphere by following the instructionsprovided by Cyagen Biosciences Inc.2.3.1. Osteogenic Differentiation in MSCs-AHAM Scaffold

Complex (AHAM Group). Singe-layered lyophilized AHAM scaffoldwas trimmed into a circular shape (1 cm in diameter), sterilized withUV light for 10 min, and pressed onto the bottom of the 24-well cellculture plate (BD Biosciences, San Jose, CA). The AHAM scaffoldswere rinsed with MSC medium to allow uniform attachment to thebottom of the wells and air dried for 10 min at room temperature.Then the scaffold was gently pipetted with 1 × 105 MSCs mixed in 0.1mL of MSC medium and maintained under a dynamic culturecondition to allow cell attachment; 0.4 mL of MSC medium was addedinto each well after 0.5 h. At day 3, the medium in each well waschanged into 0.5 mL of osteogenic differentiation medium (DMEM,10% FBS, 1% penicillin, streptomycin and dexamethasone (0.1 uM),β-glycerophosphate (10 mM), and ascorbic acid (50 uM)). Thereseeded scaffolds were then cultured under static culture conditionuntil day 21.2.3.2. Osteogenic Differentiation on 2D Cell Culture Plate (2D

Differentiation Group). MSCs (1 × 105) were seeded onto thebottom surface of the 24-well cell culture plate and cultured with 0.5mL of MSC medium for 3 days. At day 3, the medium was switched to0.5 mL of osteogenic differentiation medium and cultured until day 21.The cells were cultured under static condition during the 21 day cellculture.2.3.3. MSCs on 2D Surface without the Delivery of Osteogenic

Medium (2D Control Group). MSCs (1 × 105) were seeded in the 24-well cell culture plate and cultured with 0.5 mL of MSC medium for21 days. The cells were cultured under static condition during the 21-day cell culture. The medium of all three groups was changed on thesecond day after cell reseeding and every other day afterward until day21.2.4. DNA Assay, Cell Viability, Cell Proliferation, ALP

Activity, and Mineralization Assessment. 2.4.1. DNA Assay.DNA assay was performed to evaluate the extent of decellularization.Native HAMs and AHAMs (n = 6 for each group) were lyophilized,followed by dry weight measurement. DNA was extracted and purifiedusing a standard kit (Qiagen, Gaithersburg, MD). DNA concentrationwas then quantified by reading absorbance at 260 nm, and the resultswere expressed in ng DNA/mg dry weight of the sample.2.4.2. Cell Viability, Proliferation, and ALP Activity. Cell viability

was quantified with a Filmtracer LIVE/DEAD Biofilm Viability Kit(Invitrogen). To analyze cell proliferation and ALP activity of thethree in vitro culture groups, the MSCs−AHAM scaffold complex andthe dissociated cells in both the 2D differentiation group and 2Dcontrol group were treated with 1 mL of lysis buffer (10 mM Tris, 1mM EDTA, and 0.2% (v/v) Triton X-100) for 0.5 h on ice along with10 s of sample vortexing every 5 min. DNA within the resultant celllysate solution was then measured using a Quant-iT PicoGreen kit(Life Technologies, Grand Island, NY). ALP activity in the resultantcell lysate solution was determined via a QUANTI-Blue kit(InvivoGen, San Diego, CA) by reading optical density (OD) at620−655 nm, and the results were expressed in OD ALP/μg DNA ofthe tested sample. The time points subjected to analyses were day 3, 7,14, and 21 (n = 4 for each group).

2.4.3. Mineralization Assessment. The samples for alizarin redstaining (n = 4 for each group) were fixed in 4% paraformaldehyde for15 min and stained with 1 × alizarin red S solution (Millipore,Billerica, MA) for 30 min at room temperature. Nonspecific stainingwas removed by 4 × washes with distilled water. Images of the sampleswere then captured with a light microscope (Olympus BX43).

2.5. qRT-PCR. Total RNA of MSCs cultured in the three in vitroculture groups at day 7 and day 21 (n = 4 each) was extracted withTRIzol RNA isolation reagents, and reverse transcription (RT) wasperformed using the High Capacity RNA-to-cDNA Kit (LifeTechnologies, Grand Island, NY). Real-time polymerase chain reaction(qPCR) was performed with iQ SYBR Green Supermix and detectedwith iQ5 Optical System (Bio-Rad, Des Plaines, IL). Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as the house-keepinggene. Primer sequences design (5′-3′) was as follows: GAPDH: F-CTGGGAATCTGTCCCGTTAAG; R-CAGGAAGTCTCTGGGAA-GAATG; ALP: F-GACACGTTGACTGTGGTTACT; R-GCAGGGTCTGGAGAGTATATTTG; Collagen type I: F-CGGAC-TATTGAAGGAGCCTAAC; R-TGATGCAGGACAGAGAGAGA;Osteocalcin (OCN): F-GGGCAGTAAGGTGGTGAATAG; R-CCGTTCCTCATCTGGACTTTAT; Runx2: F-CCAAGAAGGCA-CAGACAGAA; R-GTAAGTGAAGGTGGCTGGATAG. The ther-mal profile was 50 °C 2 min, 95 °C 10 min, and then 40 amplificationcycles consisting of 95 °C 60 s and 58 °C 60 s. Relative quantificationof genes was calculated using the 2 ̂(Ct gene 2−Ct GAP 2)‑(Ct gene 1−Ct GAP 1)

equation, where “Ct gene 1” represents the threshold cycle (Ct) of thetarget gene in 2D control group and “Ct gene 2” is the Ct of the targetgene in either AHAM group or the 2D differentiation group. “Ct GAP1” and “Ct GAP 2” are for the GAPDH house-keeping gene in each ofthe respective conditions.

2.6. Biomechanical Evaluation of Membrane Materials. Theuniaxial mechanical properties of the multilayered AHAM (8 layers)and collagen membrane (Bio-Gide, porcine derived, Geistlich) (15mm × 3 mm, n = 4 for each) were characterized by a uniaxial testingmachine (Mach-1, Biosyntech, MN) along the fiber-preferreddirection. After 10 cycles of preconditioning, the sample was elongatedto failure at the ramp speed of 0.1 mm/s. The stress was calculated bynormalizing the force to the initial cross-sectional area, and the strainwas calculated by dividing the displacement with the initial grip-to-gripdistance (gauge length at 1 g preload). The biaxial mechanicalproperties were assessed with a custom-made biaxial mechanicaltesting system.35 The membranes (n = 4 for each) were cut into squaresamples with one edge of the sample aligned along the fiber-preferreddirection and the other edge along the cross fiber-preferred direction(XD) (15 mm × 15 mm). After 10 cycles of preconditioning, thesample was subjected to an equibiaxial tension at TPD:TXD = 30:30 N/m. Membrane extensibility was characterized by the maximumstretches along PD (λPD) and XD (λXD) at an equibiaxial tension of30 N/m. All samples were tested in a PBS bath.

2.7. Animal Experiments. The animal experiment protocol wasapproved by the Animal Welfare Committee of Dalian MedicalUniversity. Thirty female Sprague−Dawley (SD) rats (9 week old)weighing at 240 ± 20 g (Laboratory Animal Center of Dalian MedicalUniversity) were used for the animal experiments. Rats wereanesthetized by intraperitoneal injections with pentobarbital sodium(35 mg/kg body weight). Then a 2 cm long full-thickness incision wasmade on the right leg to expose the cranio-medial portion of the tibia.The rats were randomly selected and assigned as the followingexperimental groups (n = 6 for each group).

Group I (control group). After the periosteum was removed fromthe surgical site, a cylinder hole (2 mm in diameter, 2.5 mm in length)was drilled in the middle portion of the tibia below the knee jointusing a surgical micromotor (600 rpm) while irrigated with cold 0.9%sterile saline solution. Then a sterilized threaded cylindrical titaniumscrew (2 mm in diameter, 2.5 mm in length, Northwest Institute forNonferrous Metal Research, China) was implanted into the cylinderhole.

Group II (defect group). A cuboid defect (2 mm width × 2 mmlength × 2.5 mm depth) was created adjacent to the cylinder hole and

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reaching to the external edge of the tibia. Only the titanium screw wasimplanted into the cylinder hole, and the cuboid defect was untreated.Group III (Bio-oss only group). After the cuboid defect creation and

the titanium screw implantation following the same proceduresdescribed in Group II, the cuboid defect region was fully filled withBio-oss bone particles (bovine derived, diameter 0.25−1.00 mm,Geistlich).Group IV (Collagen group). After the implantation of the titanium

screw and the Bio-oss bone particles following the same proceduresdescribed in Group III, a collagen membrane (6 mm × 10 mm, Bio-Gide, Geistlich) was used as the barrier membrane to cover the top ofthe surgical site; note that the edges of the membrane were at least 2mm beyond the borders of the surgical site.Group V (AHAM group). The same procedure was performed as

that described in group IV, except our lyophilized multilayered AHAM(8-layer, 6 mm × 10 mm) was used as the barrier membrane. Theincision was closed after the above-described surgical procedures.2.8. X-ray, Micro-CT, and Histological Analyses. 2.8.1. X-ray

Assessment. Tibias retrieved from the rats 6 weeks postsurgery wererandomly selected and subjected to X-ray assessment (n = 4 for eachgroup). The digital radiographs of the tibias were obtained with aSOREDEX DIGORA Optime X-ray machine (InstrumentariumDental Inc., Finland). The imaging parameters were set at 60 kV, 8mA, and 0.06 s, and the gray level of the bone was analyzed with themachine software. The greater mineralization density was representedby the higher degree of imaging gray level.2.8.2. Micro-CT Imaging. After X-ray assessment, tibias (n = 4 for

each group) were scanned with a high-resolution microcomputedtomography (micro-CT) system (50 kV, 220 μA, Xradia MicroXCT-400) at a section interval of 18.25 μm, 3 s integration time, and anisotropic resolution of 18.676 μm. Moreover, the Bio-oss bone graftswere scanned separately to get the original graft density. Evaluation ofnew bone formation was based on the difference between the densityof the Bio-oss grafts (as the baseline) and the bone density in thecuboid defect site. The region of interest was chosen in adjacent to 3−6 screw threads in the cuboid defect site (length 2.5 mm × width 1mm × height 0.8 mm), and the total area of the defect, trabecular bonevolume/total defect volume (BV/TV), newly formed trabecularthickness (Tb.Th), and trabecular number (Tb.N) were measuredwith the CTAN software (Skyscan, Bruker, Belgium).2.8.3. Histology. After micro-CT scanning, tibias were fixed with

10% neutral buffered formalin for 48 h, subjected to dehydration, andthen embedded in methacrylate. Cross sections of the tibia (200−300μm) were then cut using a diamond-precision parallel saw

(EXAKT300CP, Norderstedt Germany). The region of theimplantation and cuboid defect site was then reduced to a thicknessof 10−15 μm by microgrinding and polishing. The sections werestained with the toluidine blue and Methylene blue-basic fuchsin(Sigma, St. Louis, Missouri). The total percentage of the new bonecoverage along the titanium screw surface and the total percentage ofthe matured bone inside the surgical site were measured with ImageJsoftware and are listed in Table 3.

2.9. Statistical Analysis. The experimental data were presented asmean ± standard deviation. One-way analysis of variances (ANOVA)was used for statistical analysis, with the Holm−Sidak test being usedfor posthoc pairwise comparisons and comparisons versus the controlgroup; the Student’s t test was applied for two-group comparison(SPSS, Chicago, IL). The differences were considered statisticallysignificant when p < 0.05.

3. RESULTS

3.1. Morphological, Histological, and UltrastructuralCharacteristics of AHAM. HAM showed the morphology ofa typical collagenous membrane with a relatively uniformthickness and a semitransparent appearance (Figure 1A). It wasknown that the HAM thickness varied from region to region,and the HAM we harvested for this study was found to have athickness ranging from ∼70 to 100 μm. The overall structurewas preserved after the decellularization (Figure 1B), and thethickness of AHAM ranged from 50 to 70 μm. Afterlyophilization, the dried AHAM scaffold showed an opaqueappearance due to loss of water (Figure 1C); however, afterrehydration the AHAM could regain the semitransparency andthe same handling properties before lyophilization (Figure 1D).H&E staining (Figure 1E) showed that the native HAM had

a layered structure consisting of a single epithelial layer and anamnion layer formed by a thick basement membrane and astromal layer. After decellularization, the cellular components inboth epithelial and amnion layers were found to be removed.Movat’s pentachrome staining (Figure 1F) also demonstratedthe lyophilized AHAM scaffold preserved the major ECMincluding collagen (yellow color) and proteoglycans/glyco-saminoglycans (blue color). Quantitative DNA analysisdemonstrated that, when compared with the native HAM, theAHAM had a 98.59% reduction in DNA content, decreasing

Figure 2. Immunofluorescence staining for cell nuclei, fibronectin, and collagen type I and IV (green) in HAM and AHAM.

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from 2117.9 ± 80.5 ng/mg dry weight of the HAM to 29.7 ±5.5 ng/mg dry weight of the AHAM (p < 0.01).The SEM cross-sectional view, stromal layer view, and

epithelial layer view of the HAM and AHAM (Figure 1G)further showed that the epithelial layer lining was fullyremoved, leaving a layer of fine collagen network. Also,collagen fibers in the AHAM stromal layer exhibited a reticularnetwork, and the overall ultrastructural integrity was wellpreserved. We noticed that the collagen fibers in the stromallayer of the HAM were more tortuous than the collagen fibersin the AHAM, implying collagen fibers experienced a degree ofstraightening; this observation also echoed with the thicknessthinning we mentioned above. Immunofluorescence stainingfurther confirmed the preservation of collagen types I, IV, andfibronectin in the AHAM (Figure 2). DAPI staining of theAHAM once again verified that no cell nuclei or DNAfragments remained in the AHAM (Figure 2).3.2. Cell Viability, Calcium Deposition, Cell Prolifer-

ation, and Osteogenic Differentiation. The cell viability inthe AHAM group, 2D differentiation group, and 2D controlgroup remained high after 21 day MSC culture (Figure 3A).Calcium deposition of different groups was verified by thealizarin red staining (Figure 3A). When examined at day 21, the

mineralized nodules were found in both the AHAM group andthe 2D differentiation group, which confirmed the occurrenceof osteogenic differentiation. Moreover, the AHAM grouppresented much stronger alizarin red staining than the 2Ddifferentiation group.Total DNA content of the AHAM group (Figure 3B)

showed a significant increase from day 3 to day 7 (p < 0.01)and a decrease from day 14 to day 21 (p = 0.037). From day 7to day 21, the total DNA amount of the AHAM group was allsignificantly higher than both the 2D differentiation group andthe 2D control group (p < 0.01). While ALP activity of the 2Dcontrol group was undetectable during the 21 day cell cultureperiod (Figure 3C), the ALP activity in both the 2Ddifferentiation group and the AHAM group showed asignificant increasing trend along with the cell culture time.We also noticed that the ALP activity of the AHAM group wasmuch higher than the 2D differentiation group for both day 7(p = 0.032) and day 21 (p = 0.017).At day 7 (4 days osteogenetic medium treatment), mRNA

expressions of collagen type I of the AHAM group was 1.61-fold higher than the 2D differentiation group (p = 0.035), andRunx2 expression of the AHAM group was 1.59-fold higher (p= 0.062) than the 2D differentiation group (Figure 3D). At day

Figure 3. (A) Live/dead staining and Alizarin red staining of the 2D control group, 2D differentiation group, AHAM group (21 days culture withreseeded rat mesenchymal stem cells), and AHAM without any cell culture (negative control). (B) DNA content and (C) ALP activity in the 2Dcontrol group, 2D differentiation group, and AHAM group throughout 21 days culture. (D) qRT-PCR analysis of the selected gene expressions: 2Ddifferentiation group and AHAM group at day 7 and day 21 in vitro culture (*p < 0.05; **p < 0.01).

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21 (18 days osteogenetic medium treatment), the mRNAexpression level of collagen type I and Runx2 decreasedremarkably, but the expression of ALP and OCN of both the2D differentiation group and the AHAM group increasedgreatly when compared with day 7. The expressions of ALP andOCN in the AHAM group were all higher than the 2Ddifferentiation group (ALP 1.54-fold, p = 0.066; OCN 2.41-fold, p = 0.027).3.3. Biomechanical Comparison between AHAM and

Collagen Membrane. We found that the thickness of 8-layerAHAM was approximately one-half of the thickness of the Bio-Gide collagen membrane (Figure 4A) (8-layered AHAM 0.218± 0.022 mm vs Bio-Gide collagen membrane 0.478 ± 0.017mm, p < 0.01). The stress−strain curve of the uniaxialmechanical properties testing showed that the 8-layer AHAMwas stiffer than the Bio-Gide collagen membrane along the fiberpreferred direction (Figure 4B); this trend was furtherevidenced by the maximum tensile modulus data and failurestrain data (Table 1). From the biaxial mechanical propertiesdata (Figure 4C), we found that, due to location specific fiberstructure, both the 8-layer AHAM and the Bio-Gide collagenmembrane exhibited a degree of anisotropy in which the PD

direction was stiffer than the XD direction. The extensibilitiesalong both the PD and the XD directions demonstrated thatthe 8-layer AHAM had a stiffer stress−strain behaviorcompared to the Bio-Gide collagen membrane (Table 1).

3.4. X-ray and Micro-CT Analyses of the Rat TibiaRepairing. The pictures for in vivo rat tibia procedures for thecontrol, defect, Bio-oss only, collagen, and AHAM groups arepresented in Figure 5A. Six weeks postsurgery, the plain X-rayof the defect group (Figure 5B) showed less mineralizationdensity at the cuboid defect region (yellow arrow) whencompared with the control group. Quantification of themineralization density revealed that the average gray level ofthe unrepaired defect region was lower than the correspondingregion of the control group (defect group 135.83 ± 1.47;control group 144.67 ± 7.42, p = 0.01). The Bio-oss onlygroup, the collagen group, and the AHAM group all showed ahigh degree of mineralization due to the existence of the Bio-oss grafts, of which the gray level of the AHAM group (172.17± 3.31) and the collagen group (172.33 ± 2.66) were found tobe higher than the Bio-oss only group (169.50 ± 2.43, p < 0.01)(Figure 5B, Table 2).

Figure 4. (A) Morphology and thickness of the 8-layer AHAM (left) and the Bio-Gide collagen membrane (right). (B) Uniaxial mechanical behaviorof the 8-layer AHAM and the Bio-Gide collagen membrane. (C) Biaxial mechanical properties of the 8-layer AHAM and the Bio-Gide collagenmembrane.

Table 1. Membrane Thickness and Biomechanical Parameters for 8-Layer AHAM and Collagen Membrane

biaxial mechanical properties uniaxial mechanical properties

thickness (mm) maximum stretch maximum stretch (XD) maximum tensile modulus (MPa) failure stress (MPa) failure strain (%)

8-layer AHAM 0.219 ± 0.020 1.010 ± 0.002 1.031 ± 0.014 10.513 ± 0.961 1.701 ± 0.482 25.608 ± 9.706collagen membrane 0.482 ± 0.021 1.022 ± 0.012 1.064 ± 0.003 5.637 ± 0.492 1.612 ± 0.143 48.318 ± 5.526p value p < 0.010b p < 0.010b p < 0.010b p < 0.010b p = 0.773 p = 0.045a

ap < 0.05. bp < 0.01.

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Micro-CT analysis (Figure 5C) of the defect group showednoncontinuous cortical bone regeneration in the cuboid defectsite. For the micro-CT images, we quantified the total area ofthe bone tissue inside the yellow rectangle, finding that thebone tissue area of the defect group (1.71 ± 0.12 mm2) issignificantly smaller than that of the control group (3.92 ± 0.18mm2, p = 0.0057). In the Bio-oss only group, we observedcomplete cortical bone regeneration and noticed the newlyformed bone mass originating from the periphery of the defectand reaching toward the marrow cavity. The morphology ofBio-oss bone particles was visible in the defect region,confirming the particles were imbedded in the newly formedbone tissue. However, a small gap was noticed in the regionadjacent to the titanium screw−defect interface. Both thecollagen group and the AHAM group showed an extensiveossification throughout the defect site, including the corticalbone region and the marrow cavity region. In addition, most ofthe bone particles were fully embedded within the newlyformed bone tissues, and no obvious gap was found around thetitanium screw−defect interface region, implying minimumbone absorption in those regions. Overall, the titanium screwimplants of the collagen group and AHAM group were rootedin more dense bone tissues when compared with the Bio-ossonly group.

Quantitative micro-CT evaluation of new bone regenerationin three defect repairing groups was compared with the controlgroup and shown in Table 2. We found that the total bonetissue area of the AHAM group showed no significantdifference when compared with the control group (p = 0.12),but the total bone tissue area of the Bio-oss only group and thecollagen group was smaller than the control group (p < 0.01 forboth groups). Moreover, when compared with the Bio-oss onlygroup (12.71 ± 8.12%), the AHAM group and the collagengroup exhibited a 2.19-fold (27.86 ± 5.83% for the AHAMgroup) and 1.66-fold (21.07 ± 8.09% for the collagen group)increase in BV/TV, respectively. The BV/TV, Tb.Th, and totalTb.N of the AHAM group were all close to the control group(p = 0.065 for BV/TV; p = 0.28 for Tb.Th; p = 0.16 for Tb. N).

3.5. Histology of the Repaired Rat Tibias. Methyleneblue-basic fuchsin staining of the defect group showedincomplete cortical bone regeneration with ingrowth of fibroustissues at the center of the defect site (Figure 6B). In the Bio-oss only group, the cortical bone regeneration was continuous,and Bio-oss bone particles could be seen imbedding inside thenew bone tissues (Figure 6C); moreover, inside the bonemarrow cavity a few osteoclasts (Figure 6M, red arrow) andosteoblasts (Figure 6M, green arrow) could be seen along theouter periphery of new bone tissue. In the defect group (Figure

Figure 5. (A) In vivo animal experiment pictures, (B) X-ray, and (C) micro-CT images for the control group, defect group, Bio-oss only group(cuboid defect repaired with Bio-oss bone grafts only), collagen group (cuboid defect repaired with Bio-oss bone grafts and covered with Bio-Gidecollagen membrane), and AHAM group (cuboid defect repaired with Bio-oss bone grafts and covered with lyophilized multilayer AHAMs).

Table 2. Morphometric Parameters (Radiology Imaging) Calculated at the Surgical Site

control Bio-oss collagen AHAM

gray level 56.82 ± 1.69 169.50 ± 2.43b 172.17 ± 3.31b 172.33 ± 2.66b

bone tissue area mm2 3.92 ± 0.18 3.01 ± 0.12a 3.15 ± 0.07a 3.69 ± 0.09BV/TV % 45.82 ± 1.01 12.71 ± 8.12b 21.07 ± 8.09b 27.86 ± 5.83Tb.Th mm 0.12 ± 0.01 0.23 ± 0.01 0.10 ± 0.02b 0.12 ± 0.02Tb.N 1/mm 3.68 ± 0.08 1.09 ± 0.59b 2.48 ± 0.49 2.47 ± 0.38

ap < 0.05. bp < 0.01 compare with the control group.

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6G), the Bio-oss only group (Figure 6H), and the collagengroup (Figure 6I), new bone tissues could be observed alongthe titanium screw surface, but these bone tissues were notconnected to each other or with the bone tissues in the marrowcavity. Fibrous tissues (FT) were also observed with amorphology that separated the NB and shielded the titaniumimplant.In the AHAM group (Figure 6E), the established osseous

tissues were widespread around the grafted Bio-oss particles.Osteocytes (Figure 6O, black arrow) were found to be evenlydistributed in the newly formed thick bone matrix, andosteoblastic cells (Figure 6O, green arrow) were lining thesurface of new bone, indicating that the active bone formationprocess was ongoing. A better bone−implant connection wasalso evidenced by much less fibrous connective tissues betweenthe new bone tissues and the titanium implant (Figure 6J,Table 3). We also compared the cortical bone formationunderneath the barrier membrane (Figure 6P−T). In the Bio-oss only group without barrier membrane covering, the edge ofthe new cortical bone showed a ragged outlook, and some boneparticles could be seen imbedding inside the cortical bone(Figure 6R). In both collagen (Figure 6S) and the AHAM

group (Figure 6T), the edge of new cortical bone formedunderneath the barrier membrane showed a smoother outlook,and the thicknesses of the new cortical bone were also thickercompared with the Bio-oss only group. Furthermore, the newlyformed cortical bone in the AHAM group showed higher bonedensity and cellularized compared to the collagen group. Intoluidine blue histology (Figure 7), mature bone appeared aslight purple, new bone as dark purple or blue, osteoid as paleblue, and mineralized front as light blue. In the AHAM group(Figure 7J and 7O), the newly formed bone tissues were highlycellularized and osteoid lined along the outer layer of the newbone tissue. The newly formed bone tissues showed differentstages of maturation, with up to 86.69 ± 6.32% of the newlyformed bone tissues showing light blue or purple, indicating amore mature stage of mineralization (Table 3).

4. DISCUSSIONGBR is a new concept of dental implant therapy, whichemploys a barrier membrane to seclude a protected bone-fillingspace around the dental implant surface, retard apicalepithelialization, and eliminate gingival connective tissueforming in the wound, thereby restoring the functional support

Figure 6. Methylene blue-basic fuchsin staining: (A−E) Lower magnification of the rat tibia 6-weeks postsurgery; (F−J) higher magnification of theimplant−bone connection at the 4-5th thread of the titanium screw (yellow arrow in A−E); (K−O) higher magnification of the inner site of thesurgery tibia (blue rectangle in A−E); (P−T) higher magnification of the cortical bone of the surgery tibia (green rectangle in A−E). Red arrow,osteoclasts; green arrow, osteoblasts; black arrow, osteocytes. Note that methylene blue-basic fuchsin stains the new bone and cortical bone in red,fibrous tissue (FT) in dark blue, titanium implant (DI) in black, and the Bio-oss particles (BP) in gray.

Table 3. Morphometric Parameters (Histological Imaging) Calculated at the Surgical Site

staining control, % Bio-oss, % collagen, % AHAM, %

bone coverage along the titanium screw surface methylene blue-basic fuchsin 56.82 ± 1.69 30.71 ± 10.91b 54.26 ± 9.36 56.99 ± 5.24matured bone in the surgical region toluidine blue 91.32 ± 4.56 68.68 ± 4.18b 78.08 ± 2.66a 86.69 ± 6.32

ap < 0.05. bp < 0.01 compare with the control group.

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to the implanted teeth.36,37 In this study, we developed a barriermembrane from the lyophilized AHAM, characterized itsstructural and biomechanical properties, performed in vitrocell culture studies to assess its toxic effects and osteogeneticpotential, and eventually assessed its capability in guiding boneregeneration with an in vivo rat tibia model.For clinical applications, the barrier membrane needs to be

properly prepared in order to have no ethical concerns,preserve beneficial biological properties, be easily handled, andbe free from antigenicity. Since the HAM is the discardedbiomaterial after parturition and has no harm to either mothersor babies, the ethical issues associated with the use of HAM arelimited, and the sufficient availability is able to meet the largedemand in both dentistry and bone defect treatments. Due tothe less ethical concerns, sufficient availability, and favorablebiological properties, HAM has been routinely applied inclinical treatments for years.38 Furthermore, after decellulariza-tion, the AHAM showed near-complete removal of cellular/DNA content and preserved the structural and biomechanicalintegrity of amnion membrane matrix. SEM indicated areticular structure with long collagen fibers tightly arrangedwithout disruption in the AHAM. The preservation of essentialECM components in the AHAM, such as collagen type I andIV, fibronectin, and proteoglycans/glycosaminoglycans, wasalso beneficial for later bone tissue regeneration. As we know,the preservation of the basement membrane and themaintenance of ECM macromolecules and structures areessential for cell attachment, proliferation, and migration.19,24

We also noticed that the lyophilized AHAM had a tactility ofdried membrane tissue; however, after rehydration the AHAMrestored the properties of a wet AHAM. The lyophilizedAHAM enables the long-term preservation and easy handling ofthe HAM for future clinical applications.We believe that the favorable in vivo GBR of the lyophilized

AHAM is the result of the scaffold with minimal toxic effects tocells and its capability in enhancing osteogenesis. Therefore, we

tested the toxic effects and osteogenesis capabilities of thelyophilized AHAM with an in vitro cell culture study. As aresult, the AHAM greatly promoted MSCs viability, prolifer-ation, and osteogenic differentiation. The early stage of cellosteogenic differentiation happened from day 5 to 14 of the cellculture,39 and our qRT-PCR results verified that the AHAMup-regulated the mRNA expression levels of collagen type I andRunx2 at culture day 7. Runx2 and collagen type I are the earlycell fate markers that are generally highly expressed at thebeginning phase of osteogenic differentiation.40,41 Runx2 alsoplays an important role in regulating downstream genes forosteoblast differentiation and maturation, such as ALP andOCN.40,41 After these peak expressions at day 7, the RNAexpression levels of both Runx2 and collagen type I started todecline. The final phase of osteogenesis is from day 14 to 28 ofthe cell culture, which results in a high osteocalcin andosteopontin expression and calcium deposition.42 In our results,at day 21, the AHAM caused an increasing expression of ALP,an early marker for osteogenic differentiation and ECMmineralization, and an increasing expression of OCN, a markerfor late stage of osteoblast differentiation and bone tissuematuration,43,44 suggesting that the AHAM can improve theearly stage of osteogenic differentiation of the MSCs andfurther promote the later stage of in vitro bone regenerationwhen compared to the normal 2D cell culture environment.The osteogenic property of AHAM in vitro was last

confirmed by highly efficient calcium deposition of the MSC-AHAM culture (visualized by alizarin red staining). Altogether,in vitro cell studies indicates that the lyophilized AHAMprovides a suitable environment for osteogenicity by preservingthe essential ECM composition, 3D ECM architecture, andimportant biologically active components.45

To achieve the proper thickness that is comparable with theBio-Gide collagen membrane (∼0.45−0.5 mm thickness), wecreated the 8-layer AHAM for this study based on thickness ofthe fresh HAM we harvested for this study (∼50−70 μm for a

Figure 7. Toluidine blue staining: (A−E) Lower magnification of the rat tibia 6-weeks postsurgery; (F−J) higher magnification of the implant−boneconnection at the 4-5th thread of the titanium implant (yellow arrow in A−E); (K−O) higher magnification of the inner site of the surgery tibia (redrectangle in A−E). In toluidine blue staining, mature bone (MB) appeared in light purple, newer bone in dark purple or blue, osteoid (Os) in paleblue, and mineralized front in light blue.

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single layer, ∼0.40−0.56 mm for 8-layer stack). However, afterlyophilization, we noticed that the 8-layer AHAM wassignificantly thinner compared to the Bio-Gide collagenmembrane, but thickness recovered after rehydration and the8-layer AHAM demonstrated a stiffer behavior under bothuniaxial and biaxial loading conditions. As pointed out byRakhmatia et al., sufficient structural strength is one of theimportant factors to determine the choice of the barriermembrane since stronger mechanical properties can prolongthe ability to maintain the secluded space and stabilize the bonegraft beneath during bone healing.46 One advantage of themultilayered design is the potential to achieve the desiredthickness and mechanical properties by modifying the numberof AHAM layers, which can suit various clinical applications.Various animal models, such as rat, rabbit, pig, goat, and dog,

have been used in implant dentistry and bone tissueregeneration studies.47,48 To test the capability of the 8-layerlyophilized AHAM as a barrier membrane in guiding boneregeneration for dental implant application, we chose the rattibia defect model. The advantage of the rat tibia model is thesmooth and slightly convex surface of the cranio-medial portionof the tibia, which is easy for bone exposure and defect creating.We noticed that the width of the cranio-medial portion rangedfrom 6 to 8 mm, which provides slightly limited space toaccommodate both the titanium implant and a very largecuboid defect. Thus, in our study a cuboid bone defect with 2 ×2 × 2.5 mm dimension was created adjacent to the implantinsertion. According to the previous study, a 3 mm diameterbone defect in rat tibia is considered as the critical size bonedefect that cannot heal spontaneously for up to 7 weekspostoperation.49 The defect size created in our rat tibia model isslightly smaller than the above-mentioned critical size; however,we demonstrated the incomplete cortical bone healing in thedefect group 6 weeks postsurgery, the mineralization density ofthe defect group based on the X-ray was significantly lower thanthe control group, the total area of the bone tissue in the defectgroup was obviously smaller than the control group based onmicro-CT, and the inflammatory fibrous tissues invasion in thecenter of the defect site from histology results. The incompletedefect healing confirms that the rat tibia defect modelestablished in our study serves the purpose of assessing variouslevels of bone regeneration under different experimentalconditions.Six weeks postsurgery, the Bio-oss only group exhibited an

insufficient bone growth, which is characterized by less boneingrowth near the titanium screw−defect interface and thepresence of the fibrous soft tissues. In contrast, the AHAMgroup exhibited massive bone growth and minimum Bio-ossparticle absorption. The newly formed bone tissues were highlycellularized and mature with evenly arranged osteocytes and theosteoid and osteoblasts lining along the outer layer of the newbone tissue, which indicated ongoing active bone formation.The bone−implant connection was also found beingestablished in the AHAM group, while few fibrous connectivetissues existed between the new bone tissue and the titaniumimplant. Compared to the AHAM group, the collagen groupexhibited less mature new bone tissues and some extent offibrous tissues invasion.

5. CONCLUSIONSOur results demonstrated that both the collagen membrane andthe lyophilized 8-layer AHAM acted effectively as a shieldinglayer to prevent the invasion of the fibrous tissue and promoted

bone-to-implant connection when compared with the Bio-ossonly repairing (no membrane coverage). Moreover, thelyophilized AHAM barrier membrane further induced themassive bone growth and maturation when compared with thecollagen membrane. In short, we developed a new barriermembrane produced from lyophilized multilayered AHAM.The advantages of the AHAM barrier membranes are theirexcellent biomechanical properties, preservation of naturalECM structure and composition, easiness in preparation andhandling, flexibility in adjusting the thickness and mechanicalproperties to suit the application, and efficiency in inducing themassive bone growth and avoiding fibrous tissues invasion.

6. LIMITATIONS AND FUTURE STUDIESThe limited width of the rat tibia restricted the accommodationof both the cylindrical threaded titanium implant and a largesize neighboring cuboid defect, which would cause a high riskof fracture. Hence, the tibia defect size in our study is slightlysmaller than the critical bone defect size (3 mm). However, aswe pointed out above, we confirmed that the used cuboiddefect size (2 mm × 2 mm × 2.5 mm) did not experiencespontaneous defect healing and served the purpose of our studyto assess the defect repairing under various conditions. Thecurrent findings in the rat tibia model have shed light on thepotential of the AHAM as a barrier membrane in assistingdefect regeneration and dental implanting. Another limitationof this study is that the biocompatibility, immune responses,and membrane degradation of the AHAM after in vivoimplantation were not assessed. Future histology study isrequired to give evidence if there is any effective immuneresponse following AHAM application inside the surgical site.In future studies, we will assess the time for membraneresorption and the potential of the AHAM barrier membrane inassisting dental implantation and bone regeneration byrepairing critical sized oral osseous defects in a large animalmodel, as well as adding more time points. Other barriermembrane enhancement approaches we are consideringinclude constructing the barrier membrane by addingosteoinductive growth factors.

■ AUTHOR INFORMATIONCorresponding Authors*Phone: 086-13998698869. E-mail: liwuwei96@163.com.*Phone: (662) 325-5987. E-mail: jliao@abe.msstate.edu.*Phone: (312) 503-3698. E-mail: bo.wang@northwestern.edu.NotesThe authors declare no competing financial interest.

■ ACKNOWLEDGMENTSThis study was supported by the Natural Science Foundation ofLiaoning, China (2013010206-401) and in part by NSF EPS-0903787.

■ ABBREVIATIONSAHAM, acellular human amnion membrane; GBR, guided boneregeneration; ECM, extracellular matrix; MSCs, mesenchymalstem cells; SD, Sprague−Dawley; SDS, sodium dodecyl sulfate;H&E, hematoxylin and eosin; PBS, phosphate-buffered saline;SEM, scanning electron microscopy; qRT-PCR, alkalinephosphatase, quantitative real-time reverse transcription poly-merase chain reaction; GAPDH, glyceraldehyde-3-phosphatedehydrogenase; OCN, osteocalcin; Ct, threshold cycle; XD,

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fiber preferred direction, cross preferred direction; Bio-oss,Geistlich Bio-Oss Granulate cancellous bone granules; micro-CT, microcomputed tomography; BV/TV, bone volume pertotal volume; Tb.Th, trabecular thickness; Tb.N, trabecularnumber; ANOVA, one-way analysis of variances

■ REFERENCES(1) Wetzel, A. C.; Stich, H.; Caffesse, R. G. Bone apposition onto oralimplants in the sinus area filled with different grafting materials. Ahistological study in beagle dogs. Clin Oral Implants Res. 1995, 6 (3),155−163.(2) Haas, R.; et al. Bovine hydroxyapatite for maxillary sinus grafting:comparative histomorphometric findings in sheep. Clin Oral ImplantsRes. 1998, 9 (2), 107−116.(3) Claffey, N.; et al. Surgical treatment of peri-implantitis. J. Clin.Periodontol. 2008, 35 (8 Suppl), 316−332.(4) Becker, B. E.; Becker, W. Regeneration procedures: graftingmaterials, guided tissue regeneration, and growth factors. Curr. Opin.Dentistry 1991, 1 (1), 93−97.(5) Dahlin, C.; et al. Healing of bone defects by guided tissueregeneration. Plastic and reconstructive surgery 1988, 81 (5), 672−676.(6) Skoglund, A.; Hising, P.; Young, C. A clinical and histologicexamination in humans of the osseous response to implanted naturalbone mineral. Int. J. Oral Maxillofac. Implants 1997, 12 (2), 194−199.(7) Smiler, D. G.; et al. Sinus lift grafts and endosseous implants.Treatment of the atrophic posterior maxilla. Dent. Clin. North Am.1992, 36 (1), 151−186 discussion 187−188.(8) Young, C.; Sandstedt, P.; Skoglund, A. A comparative study ofanorganic xenogenic bone and autogenous bone implants for boneregeneration in rabbits. Int. J. Oral Maxillofac. Implants 1999, 14 (1),72−76.(9) Machtei, E. E. The effect of membrane exposure on the outcomeof regenerative procedures in humans: a meta-analysis. J. Periodontol.2001, 72 (4), 512−516.(10) Wang, H. L.; Carroll, M. J. Guided bone regeneration usingbone grafts and collagen membranes. Quintessence Int. 2001, 32 (7),504−515.(11) Zitzmann, N. U.; Scharer, P.; Marinello, C. P. Long-term resultsof implants treated with guided bone regeneration: a 5-yearprospective study. Int. J. Oral Maxillofac. Implants 2001, 16 (3),355−366.(12) Marinucci, L.; et al. In vitro comparison of bioabsorbable andnon-resorbable membranes in bone regeneration. J. Periodontol. 2001,72 (6), 753−759.(13) Soost, F.; et al. Validation of bone conversion inosteoconductive and osteoinductive bone substitutes. Cell TissueBanking 2001, 2 (2), 77−86.(14) Dori, F.; et al. Effect of platelet-rich plasma on the healing ofintra-bony defects treated with a natural bone mineral and a collagenmembrane. J. Clin. Periodontol. 2007, 34 (3), 254−261.(15) Ling, L. J.; et al. The influence of membrane exposure on theoutcomes of guided tissue regeneration: clinical and microbiologicalaspects. J. Periodontal Res. 2003, 38 (1), 57−63.(16) Grad, S.; et al. Chondrocytes seeded onto poly (L/DL-lactide)80%/20% porous scaffolds: a biochemical evaluation. J. Biomed. Mater.Res. 2003, 66 (3), 571−579.(17) Gielkens, P. F.; et al. Vivosorb, Bio-Gide, and Gore-Tex asbarrier membranes in rat mandibular defects: an evaluation bymicroradiography and micro-CT. Clin Oral Implants Res. 2008, 19 (5),516−521.(18) Danforth, D.; Hull, R. W. The microscopic anatomy of the fetalmembranes with particular reference to the detailed structure of theamnion. Am. J. Obstet Gynecol. 1958, 75 (3), 536−547 discussion 548−550.(19) Niknejad, H.; et al. Properties of the amniotic membrane forpotential use in tissue engineering. Eur. Cells Mater. 2008, 15, 88−99.(20) Parry, S.; Strauss, J. F., 3rd Premature rupture of the fetalmembranes. N. Engl. J. Med. 1998, 338 (10), 663−670.

(21) Murdoch, A. D.; et al. Primary structure of the human heparansulfate proteoglycan from basement membrane (HSPG2/perlecan). Achimeric molecule with multiple domains homologous to the lowdensity lipoprotein receptor, laminin, neural cell adhesion molecules,and epidermal growth factor. J. Biol. Chem. 1992, 267 (12), 8544−8557.(22) Koizumi, N. J.; et al. Growth factor mRNA and protein inpreserved human amniotic membrane. Curr. Eye Res. 2000, 20 (3),173−177.(23) Tseng, S. C. Amniotic membrane transplantation for ocularsurface reconstruction. Biosci. Rep. 2001, 21 (4), 481−489.(24) Azuara-Blanco, A.; Pillai, C. T.; Dua, H. S. Amniotic membranetransplantation for ocular surface reconstruction. Br. J. Ophthalmol.1999, 83 (4), 399−402.(25) Tsubota, K.; et al. Surgical reconstruction of the ocular surfacein advanced ocular cicatricial pemphigoid and Stevens-Johnsonsyndrome. Am. J. Ophthalmol. 1996, 122 (1), 38−52.(26) Chen, H. J.; Pires, R. T.; Tseng, S. C. Amniotic membranetransplantation for severe neurotrophic corneal ulcers. Br. J.Ophthalmol. 2000, 84 (8), 826−833.(27) Gajiwala, K.; Gajiwala, A. L. Evaluation of lyophilised, gamma-irradiated amnion as a biological dressing. Cell Tissue Banking 2004, 5(2), 73−80.(28) Park, M.; et al. Healing of a porcine burn wound dressed withhuman and bovine amniotic membranes. Wound Repair Regen. 2008,16 (4), 520−528.(29) Mohammadi, A.; Johari, H. G. Amniotic membrane: a skin graftfixator convenient for both patient and surgeon. Burns 2008, 34 (7),1051−1052.(30) Ishino, Y.; et al. Amniotic membrane as a carrier for cultivatedhuman corneal endothelial cell transplantation. Invest. Ophthalmol.Visual Sci. 2004, 45 (3), 800−806.(31) Tsai, S. H.; et al. Characterization of porcine arterial endothelialcells cultured on amniotic membrane, a potential matrix for vasculartissue engineering. Biochem. Biophys. Res. Commun. 2007, 357 (4),984−990.(32) Lindenmair, A.; et al. Osteogenic differentiation of intact humanamniotic membrane. Biomaterials 2010, 31 (33), 8659−8665.(33) Chen, Y. J.; et al. The effects of acellular amniotic membranematrix on osteogenic differentiation and ERK1/2 signaling in humandental apical papilla cells. Biomaterials 2012, 33 (2), 455−463.(34) Samandari, M. H.; Adibi, S.; Khoshzaban, A.; Aghazadeh, S.;Dihimi, P.; Torbaghan, S. S.; Keshel, S. H.; Shahabi, Z. Humanamniotic membrane, best healing accelerator, and the choice of boneinduction for vestibuloplasty technique (an animal study). TransplantRes. Risk Manage. 2011, 3, 1−8.(35) Wang, B.; et al. Myocardial scaffold-based cardiac tissueengineering: application of coordinated mechanical and electricalstimulations. Langmuir 2013, 29 (35), 11109−11117.(36) Nyman, S.; et al. The regenerative potential of the periodontalligament. An experimental study in the monkey. J. Clin. Periodontol.1982, 9 (3), 257−65.(37) Nyman, S.; et al. New attachment following surgical treatmentof human periodontal disease. J. Clin. Periodontol. 1982, 9 (4), 290−296.(38) Lindenmair, A.; et al. Mesenchymal stem or stromal cells fromamnion and umbilical cord tissue and their potential for clinicalapplications. Cells 2012, 1 (4), 1061−1088.(39) Aubin, J. E. Regulation of osteoblast formation and function.Rev. Endocr. Metab. Disord. 2001, 2 (1), 81−94.(40) Franceschi, R. T.; et al. Transcriptional regulation of osteoblasts.Ann. N. Y. Acad. Sci. 2007, 1116, 196−207.(41) Komori, T. Regulation of skeletal development by the Runxfamily of transcription factors. J. Cell. Biochem. 2005, 95 (3), 445−453.(42) Hoemann, C. D.; El-Gabalawy, H.; McKee, M. D. In vitroosteogenesis assays: influence of the primary cell source on alkalinephosphatase activity and mineralization. Pathol. Biol. 2009, 57 (4),318−323.

Langmuir Article

DOI: 10.1021/acs.langmuir.5b02362Langmuir 2015, 31, 8642−8653

8652

(43) Aubin, J. E. Regulation of osteoblast formation and function.Rev. Endocr. Metab. Disord. 2001, 2 (1), 81−94.(44) Graneli, C.; et al. Novel markers of osteogenic and adipogenicdifferentiation of human bone marrow stromal cells identified using aquantitative proteomics approach. Stem Cell Res. 2014, 12 (1), 153−165.(45) Macchiarini, P.; et al. Clinical transplantation of a tissue-engineered airway. Lancet 2008, 372 (9655), 2023−2030.(46) Rakhmatia, Y. D.; et al. Current barrier membranes: titaniummesh and other membranes for guided bone regeneration in dentalapplications. J. Prosthodont Res. 2013, 57 (1), 3−14.(47) Stubinger, S.; Dard, M. The rabbit as experimental model forresearch in implant dentistry and related tissue regeneration. J. InvestSurg 2013, 26 (5), 266−282.(48) Pellegrini, G.; et al. Pre-clinical models for oral and periodontalreconstructive therapies. J. Dent. Res. 2009, 88 (12), 1065−1076.(49) Lewandrowski, K. U.; et al. Effect of a poly(propylene fumarate)foaming cement on the healing of bone defects. Tissue Eng. 1999, 5(4), 305−316.

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