Engineered Cartilage Covered Ear Implants for Auricular Cartilage

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Biomacromolecules 2011, 12, 306–313

Engineered Cartilage Covered Ear Implants for Auricular Cartilage Reconstruction Sang Jin Lee, Christopher Broda, Anthony Atala, and James J. Yoo* Wake Forest Institute for Regenerative Medicine, Wake Forest University Health Sciences, Medical Center Boulevard, Winston-Salem, North Carolina 27157, United States Received July 27, 2010; Revised Manuscript Received November 28, 2010

Cartilage tissues are often required for auricular tissue reconstruction. Currently, alloplastic ear-shaped medical implants composed of silicon and polyethylene are being used clinically. However, the use of these implants is often associated with complications, including inflammation, infection, erosion, and dislodgement. To overcome these limitations, we propose a system in which tissue-engineered cartilage serves as a shell that entirely covers the alloplastic implants. This study investigated whether cartilage tissue, engineered with chondrocytes and a fibrin hydrogel, would provide adequate coverage of a commercially used medical implant. To demonstrate the in vivo stability of cell-fibrin constructs, we tested variations of fibrinogen and thrombin concentration as well as cell density. After implantation, the retrieved engineered cartilage tissue was evaluated by histo- and immunohistochemical, biochemical, and mechanical analyses. Histomorphological evaluations consistently showed cartilage formation over the medical implants with the maintenance of dimensional stability. An initial cell density was determined that is critical for the production of matrix components such as glycosaminoglycans (GAG), elastin, type II collagen, and for mechanical strength. This study shows that engineered cartilage tissues are able to serve as a shell that entirely covers the medical implant, which may minimize the morbidity associated with implant dislodgement.

Introduction Cartilage tissue is often required for reconstructive surgical procedures that are performed to treat congenital anomalies or traumatic injury. The standard treatment method for auricular (ear) reconstruction uses autologous costal cartilage as a graft material. However, autologous costal cartilage is limited in supply, provides inadequate dimensions, and is progressively absorbed after implantation.1-3 Currently, alternative approaches utilize alloplastic ear implant devices composed of silicon or polyethylene.4-8 These implants are approved by the Food and Drug Administration (FDA), and they are nontoxic, cause minimal foreign body reactions, and possess adequate mechanical properties for use in non-load-bearing tissues of the craniofacial region.4,6 Although alloplastic ear implants are able to effectively eliminate the morbidity associated with the costal cartilage graft, the use of these implants is often related to other complications that include inflammation, infection, erosion, and dislodgement.7-9 As a result, implant extrusion occurs frequently because of limited vascularization and constant abrasion against the surrounding tissues. Cell-based approaches have been considered as a new concept in cartilage tissue reconstruction. Whereas some biodegradable materials serve as a scaffolding system that promotes the body’s ability to regenerate toward healing, a tissue engineering approach uses cells seeded on a scaffold to achieve neocartilage tissue formation.10,11 We have previously demonstrated that cartilage tissue can be engineered using a patient’s own cells and applied in various reconstructive procedures, including injectable cartilage for the treatment of vesicoureteral reflux and incontinence12 and as a penile prosthesis.13-15 However, scaffolds for each application a designed and configured differently * To whom correspondence should be addressed. Fax: +1 336 713 7290. E-mail: [email protected].

because they play a major role in guiding tissue formation to achieve desired function. In cartilage tissue engineering, hydrogel systems have shown to be effective in providing 3D configurations while maintaining chondrocyte viability. Furthermore, the combined high water content and elasticity of hydrogels are considered to be essential for improved tissue formation. Typical hydrogels for cartilage tissue reconstruction are made from naturally derived polymers such as fibrin, alginate,12,16 chitosan,17 agarose,18 collagen,19 and hyaluronic acid20 as well as synthetic polymers such as poly(ethylene oxide),21 Pluronics,22 and poly(propylene fumarate).23 Among the many derivatives of hydrogels, fibrin hydrogels have been used for cell delivery in cartilage tissue engineering in recent years.24-29 This is due to the easy fabrication process and their ability to maintain viability of cells during the gelation process. Engineering of cartilage tissue de novo for total auricular reconstruction requires an extensive cell expansion process and a lengthy wait time while the new cartilage forms. More importantly, the current limitations of engineered cartilage tissues for clinical application are primarily the size reduction resulting from lack of mechanical strength and long-term stability in vivo. To overcome these limitations, we have developed a novel engineering system in which cartilage tissue entirely covers an alloplastic medical implant. Coverage of the alloplastic implant with cartilage tissue may prevent implant exposure and extrusion while maintaining appropriate mechanical strength and morphological stability. In addition, the creation of cartilage tissue using a patient’s own cells would also minimize the morbidity associated with implant dislodgement and immunogenicity. The objectives of this study were to investigate the feasibility of using a chondrocyte-fibrin system to engineer auricular cartilage tissue and to demonstrate the integrity of the interface between the engineered cartilage tissue

10.1021/bm100856g  2011 American Chemical Society Published on Web 12/23/2010

Engineered Cartilage Covered Ear Implants

Biomacromolecules, Vol. 12, No. 2, 2011

Table 1. Summary of Variation of Fibrinogen and Thrombin Concentration and Initial Cell Density of the Constructs (n ) 6) fibrin hydrogels experimental groups

fibrinogen (mg/mL)

thrombin (U/mL)

cell density (cells/mL)

1 2 3 4 5 6

25 50 80 80 80 80

10 10 10 10 10 100

1 × 107 1 × 107 1 × 107 5 × 106 4 × 107 1 × 107

and a commercially available ear implant for clinical applications. In this study, we examined the morphological stability, mechanical strength, and phenotypic characteristics of the chondrocyte-fibrin constructs in vivo.

Materials and Methods Fabrication and Characterization of Fibrin Hydrogels. Fibrin hydrogels were used as a cell carrier for auricular cartilage tissue formation. To evaluate initial properties of fibrin hydrogels, we tested various concentrations of fibrinogen and thrombin (both derived from bovine plasma). A range of 25-80 mg/mL fibrinogen was dissolved in a 0.9% sodium chloride aqueous solution. Fibrinogen solution was mixed with the same volume of thrombin at a concentration range of 2.5-200 U/mL in a 40 mM CaCl2 aqueous solution. All reagents were obtained from Sigma-Aldrich (St. Louis, MO) and used as received unless stated otherwise. Fibrin hydrogels were formed by casting the thrombin and fibrinogen solutions into a silicon mold (1.0 × 2.0 × 0.3 cm3) using a double syringe applicator (Fibrijet, Micromedics, St. Paul, MN) at room temperature. After fibrinogen and thrombin were mixed, the solution lost its fluidity and was transformed into a hydrogel. The various fibrin formulations were analyzed by measuring gelation time and stiffness (Young’s modulus). We obtained mass swelling ratios of fibrin hydrogels by dividing each construct’s swollen weight by its dry weight. In brief, freshly prepared fibrin hydrogels were incubated in a 24-well plate with 2 mL of PBS for 24 h. Hydrogels were then removed, blotted for excess fluid, and weighed. To determine dry weight, fibrin constructs were frozen and lyophilized for 48 h, then weighed. Isolation and Culture of Auricular Chondrocytes. Auricular chondrocytes were obtained from fresh ear cartilage tissue biopsies from New Zealand White rabbits (Charles River Laboratories, Wilmington, MA). We dissected cartilage from the external ear after carefully removing the perichondrium. Cartilage tissues were minced into 1 mm3 pieces. These were suspended in Ham’s F12 medium containing 0.2% type II collagenase (Worthington Biochemical, Freehold, NJ) and incubated at 37 °C for 4 h. After enzymatic digestion, we passed the resulting cell suspension through a 100 µm filter to collect cells. The chondrocytes were grown and expanded in low-glucose Dulbecco’s modified Eagle’s medium (DMEM), 10% fetal bovine serum (FBS), 1% L-glutamine, and 1% streptomycin penicillin at 37 °C, 5% CO2 until sufficient numbers of cells were obtained. All reagents for cell culture were purchased from Invitrogen (Gibco Cell Culture, Carlsbad, CA). After isolation, chondrocytes were characterized using phenotypic expression and proliferation assays. In Vivo Evaluation of Chondrocyte-Fibrin Constructs. To determine the in vivo morphological stability and neocartilage tissue formation, constructs were implanted subcutaneously under the dorsal skin of athymic mice (NU/NU Nude), Charles River Laboratories). Six experimental groups were implanted with constructs containing varying concentrations of fibrinogen and thrombin and various cell densities (n ) 6, Table 1). The chondrocytes were suspended in the fibrinogen solutions; subsequently, we prepared cell-fibrin constructs by adding the same volume of thrombin solution to the silicon mold (1.0 × 2.0 × 0.3 cm3) using a double syringe applicator prior to implantation.

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All animal procedures were approved by the Institutional Animal Care and Use Committee (ACUC) at Wake Forest University. Under general anesthesia, a dorsal longitudinal incision was made, and a subcutaneous pocket was created. The chondrocyte-fibrin constructs were placed into the subcutaneous space, and the wound was closed in a routine fashion. All animals were observed for mortality and signs of overt toxicity once daily after implantation until sacrifice. The constructs were retrieved at 4, 8, 12, and 24 weeks after implantation for analyses. A total of 72 animals were used in this study (6 groups × 4 time points × 3 animals per time point). Two constructs were implanted per animal. To evaluate the dimensional changes of cell-fibrin constructs after implantation, we measured the samples prior to implantation and at retrieval using a caliper. We calculated the percent change in dimension at each time point by comparing the total dimension (thickness, width, and length) of the retrieved samples with the preimplantation values. Compression Testing. Mechanical properties of the retrieved cell-fibrin constructs were assessed using a mechanical tester with a compression interface diameter of 2 in. (Instron model 5544, Canton, MA). A 100 N load cell at a crosshead speed of 0.4 mm/min was used for compression testing. The compressive strength was determined from the maximum load recorded. Stiffness (Young’s modulus) was calculated from the initial slope of the stress-strain curve. At least three specimens were tested for each time point, and the average and the standard deviation were calculated. Histological and Immunohistochemical Analyses. The retrieved chondrocyte-fibrin constructs from each experimental group were prepared for histological analysis by fixation in 10% phosphate-buffered formalin at room temperature for 24 h. Subsequently, the samples were embedded in paraffin and sectioned into 5 µm sections. Deparaffinized sections were stained with hematoxylin and eosin (H&E), alcian blue, and safranin-O, which confirmed the cellular morphology and glycosaminoglycans (GAG) production. We performed immunohistochemical staining to confirm the production of type II collagen. In brief, samples were deparaffinized, rehydrated, and subjected to pepsin antigen retrieval at 37 °C for 20 min. After returning to room temperature, the samples were blocked in 10% rabbit serum in phosphate-buffered saline (PBS) for 30 min, followed by incubation with 1:30 goat antitype II collagen-UNLB (Southern Biotech, Birmingham, AL) for 1 h. Samples were enzymatically blocked for 10 min. The samples were then incubated in the secondary biotinylated rabbit antigoat antibody (BA-5000, Vector Laboratories, Burlingame, CA) for 30 min. Streptavidin-conjugated horseradish peroxidase (SA-5704, Vector Laboratories) was added for 30 min, and the samples were stained with 3,3′-diaminobenzidine (DAB, SK-4100, Vector Laboratories). Finally, the samples were counterstained with Gill’s hematoxylin. All histological samples were visualized and photographed with a Zeiss microscope (M1, Zeiss Axio Imager) using the Axiovision Software (Carl Zeiss, Jena, Germany). Water Content Measurement. The retrieved constructs were weighed before and after lyophilization. The difference between the weights before and after lyophilization was calculated as the water content of the constructs. Native rabbit ear cartilage served as a control. The samples were then subjected to biochemical analyses to determine the DNA and elastin contents, which are reported as percentages of the weight of wet tissue. Biochemical Analyses. DNA was isolated by DNeasy Tissue kit (QIAGEN, Valencia, CA). The construct samples were digested and incubated in a proteinase K solution (10 mg/mL) at 65 °C for 16 h until tissues were completely solubilized. The DNA content of each sample was determined using the PicoGreen dsDNA Assay kit (Molecular Probes, Carlsbad, CA). Calf thymus DNA served as a standard control. Values obtained for DNA content were normalized to wet sample weight. Elastin was extracted using 0.25 M of oxalic acid at 95 °C for 1 h. The elastin concentration in the extract was measured using the Fastin assay kit (Biocolor). Aliquots of samples (1/100 µL) and standards were

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Figure 1. Physical properties of fibrin hydrogels: (a,b) gelation time, (c,d) mass swelling ratio, and (e,f) stiffness of fibrin hydrogels with varying concentrations of fibrinogen and thrombin (without cells). *P < 0.05, n ) 6.

read using a microplate reader (ELX 800, Bio-Tex Instruments, Winooski, VT) at 513 nm. Engineered Cartilage Covered Ear Implants. Commercially available ear implants (MedPor) and base materials were kindly provided by Porex Surgical Products Group (Newnan, GA). To improve surface hydrophilicity, we modified the implants (high-density polyethylene, HDPE) by surface oxidation techniques.30 In brief, an oxidation solution [H2SO4/H2O/CrO3 (29/42/29, w/w/w)] was prepared in a chemical hood. The implants (1.5 × 1 × 0.11 cm3) were immersed in this solution at 72 °C for up to 3 min and then rinsed with an excess of distillated water three times. The rinses were followed by an acetone wash at room temperature for 10 min. The surface-modified ear implants were dried and stored in a desiccator until use. To analyze the surface characteristics of the implant, we measured water contact angle and fluid uptake ability. The water contact angle, an indicator of the wettability of surfaces, was measured by a sessile drop method at room temperature using an optical bench-type contact angle goniometer with the compact digital video camera, which is used to obtain images for optical determination of contact angles (CAM100, KSV Instruments LTD, Helsinki, Finland). A fluid uptake test was used to determine the hydrophilicity of the implant. For the evaluation of fluid uptake capacity, the implant was immersed in PBS at pH 7.4 for 30 s. Percent fluid uptake was calculated according to the following equation: fluid uptake (%) ) (Ws - Wo)/Wo × 100, where Wo is the dry sample and Ws is the wet sample. Auricular chondrocytes were suspended in a solution containing fibrinogen (80 mg/mL) and 0.9% sodium chloride to produce a cell density of 8 × 107 cells/mL. Thrombin was dissolved in a 40 mM CaCl2 aqueous solution to produce a concentration of 10 U/mL. Equal volumes of both solutions were mixed using a dual syringe system, yielding a chondrocyte-fibrin suspension with a final cell density of 4 × 107 cells/mL. The cell-fibrin suspension was then placed directly onto the surface modified ear implants. The cell-fibrin-covered implants (1.5 × 1 × 0.21 cm3) were implanted subcutaneously under the dorsal skin of athymic nude mice using the surgical procedure described above (n ) 6). Unmodified MedPor implants served as controls. At 24 weeks postimplantation, the implants were retrieved and prepared for histology by fixation in 10% phosphate-buffered formalin at room temperature for 24 h. The

sample was embedded in paraffin and sectioned into 5 µm sections. Deparaffinized sections were stained with H&E, alcian blue, and safranin-O. Statistical Analyses. One-way ANOVA, followed by Tukey’s post hoc test, was used to determine significant differences between experimental groups in all. Differences were considered to be significant at P < 0.05. All values were reported as the mean ( the standard deviation.

Results Fabrication and Characterization of Fibrin Hydrogels. The concentrations of fibrinogen and thrombin, cell density, Ca2+ concentration, and pH can be modified for optimizing fabrication conditions for hydrogel systems used in tissue engineering.21 To evaluate the initial physical properties of a hydrogel system for use in auricular cartilage reconstruction, fibrin hydrogels with varying concentrations of fibrinogen and thrombin were prepared. Variation of the fabrication parameters can affect the initial properties of cell-fibrin constructs, including the gelation time, mass swelling ratio, and mechanical strength (Figure 1). In this study, the gelation time of the fibrin hydrogels was delayed when high concentrations of fibrinogen were used (Figure 1a). The gelation time was significantly accelerated by increasing the concentration of thrombin (Figure 1b). The degree of swelling of fibrin hydrogels is dependent on the concentrations of fibrinogen and thrombin, which influence the interaction between the gel material and solvent. The mass swelling ratio was significantly decreased with increasing concentrations of fibrinogen (Figure 1c). However, the increase in the concentration of thrombin (up to 100 U/mL) significantly enhanced the mass swelling ratio of the fibrin hydrogels (Figure 1d). Compression testing indicated that the stiffness of fibrin hydrogels containing 10 U/mL thrombin and 25 mg/mL fibrinogen was significantly different from the stiffness of hydrogels containing 80 mg/mL fibrinogen (Figure 1e). In addition, when the concentration of fibrinogen in the gels was maintained constant at 80 mg/mL and the thrombin concentration was varied, the stiffness of a hydrogel containing 10 U/mL thrombin was



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Figure 2. (a) Gross appearance of chondrocyte-fibrin constructs before and after implantation; (left) the initial shape of the construct (1 × 2 × 0.3 cm3) and (right) retrieved constructs at 24 weeks after implantation. Concentrations of fibrin and thrombin, as well as cell numbers, used in constructs 1-6 are listed in Table 1. (b-d) Percentage of maintaining dimension and stiffness (Young’s modulus) of the retrieved chondrocyte-fibrin constructs at 4, 8, 12, and 24 weeks after implantation. (b,e) 10 U/mL thrombin and 1 × 107 cells/mL initial cell density; (c,f) 80 mg/mL fibrinogen and 1 × 107 cells/mL; and (d,g) 80 mg/mL fibrinogen and 10 U/mL thrombin. *P < 0.05, n ) 3.

significantly different from that of a gel containing 100 U/mL of thrombin (Figure 1f). In Vivo Evaluation of Chondrocyte-Fibrin Constructs. Primary auricular chondrocytes isolated from rabbit ear cartilage were expanded until a sufficient numbers of cells were obtained. The phenotype of these cells was confirmed by immunocytochemistry for type II collagen (data not shown). The cells were then mixed with hydrogels containing varying amounts of fibrinogen and thrombin at different cell densities (Table 1). These chondrocyte-fibrin constructs were implanted subcutaneously into athymic mice and harvested 4, 8, 12, and 24 weeks after implantation (n ) 6). By 24 weeks, the constructs containing 80 mg/mL fibrinogen, 10 U/mL thrombin, and 4 × 107 cells/mL were the most similar to native auricular cartilage in color (opaque white), and they maintained their initial shape and size (Figure 2a). To evaluate in vivo dimensional stability of cell-fibrin constructs in all experimental groups after implantation, we

analyzed dimensions of the retrieved constructs (Figure 2b-d). We calculated the percent change in dimensions by comparing the total volume (thickness × width × length) of each of the constructs with their initial volume. Dimensions of the constructs gradually reduced when the fibrinogen concentration in the hydrogel was reduced (Figure 2b). However, there were no statistically significant differences in the dimensional changes between the various thrombin concentrations (Figure 2c). In addition, the initial cell density significantly influenced the dimensional maintenance of the constructs over time. At 24 weeks, the dimensions of the constructs composed of 80 mg/ mL fibrinogen, 10 U/mL thrombin, and 4 × 107 cells/mL, which contained the highest cell density, were ∼83% of the initial size (Figure 2d). Using compression testing, the stiffness (Young’s modulus) of the retrieved chondrocyte-fibrin constructs was measured at each time point. Stiffness of the constructs was increased by decreasing fibrinogen concentration. This was due to fast

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Figure 3. Histological evaluations of the retrieved chondrocyte-fibrin constructs with varying concentrations of fibrinogen and thrombin and cell density (Table 1) after 12 and 24 weeks in vivo. Sections of the constructs were stained with safranin-O staining for GAG and immunohistochemistry for collagen type II. Scale bar indicates 200 µm. Variation of fibrinogen and thrombin concentration did not affect the GAG production. However, the constructs initially seeded with 4 × 107 cells/mL produced a more homogeneous GAG distribution up to 24 weeks.

absorption of fibrin hydrogel, followed by gel contraction (Figure 2e). However, thrombin concentration did not affect the stiffness of the constructs (Figure 2f). These results show that fibrinogen concentration and initial cell density (Figure 2g) significantly affected the mechanical strength of the engineered cartilage tissue. In addition, the stiffness of the constructs in all experimental groups gradually increased over time. Histological and Immunohistochemical Analyses. Histological staining of the retrieved samples showed the production of a new cartilaginous matrix within the chondrocyte-fibrin constructs at all time points in all experimental groups (Figure 3). The chondrocytes in the newly formed tissue demonstrated similar morphological characteristics to those in native cartilage, with cells located within typical chondrocyte lacunae, surrounded by cartilaginous matrix. The newly formed matrix generated in all experimental groups stained intensely with safranin-O, demonstrating the presence of sulfated proteoglycans at all time points. Immunohistochemical staining indicated the presence of type II collagen at all time points with increased reactivity at 12 and 24 weeks after implantation. In addition, variation of fibrinogen and thrombin concentration did not affect the GAG production. However, the constructs initially seeded with 4 × 107 cells/mL produced a more homogeneous GAG distribution up to 24 weeks. Water Content and Biochemical Analyses. We determined the water content of the constructs by comparing wet and dry weights (Figure 4a-c). All experimental groups exhibited a slight decrease in water content with time, most likely due to gel contraction or matrix production. The constructs containing 25 mg/mL fibrinogen, 10 U/mL thrombin, and 1 × 107 cells/ mL had the lowest water content (79.67 ( 1.13%) compared with the other groups because of high gel contraction. This indicates that fibrinogen is the component that contributes most to the structural integrity of a fibrin hydrogel after gelation. Figure 4c indicates that the water content also increased with decreasing initial cell density. The total DNA content within the constructs was determined using the PicoGreen dsDNA Assay (Figure 4d-f). The results

indicate that initial cell density was maintained over time (Figure 4f). However, all experimental groups exhibited a slight decrease in the DNA content,followed by increased ECM production with time. Elastin content per wet weight also increased over time with increasing fibrinogen concentration (Figure 4g) and initial cell density (Figure 4i). However, thrombin concentration did not affect the elastin content (Figure 4h). In general, the constructs exhibited an increase in cartilaginous matrix production with increasing initial cell density. Engineered Cartilage-Covered Ear Implants. Figure 5a-c shows the base material, which is used an ear implant (MedPor). The procedure of developing cartilage to cover this ear implant consisted of two main steps: (1) surface modification by oxidation to achieve hydrophilicity and (2) attachment of the chondrocyte-fibrin hydrogel to the ear implant. After surface modification by oxidation, the water contact angle of the ear implants was decreased (data not shown). The fluid uptake ability of ear implants increased with increasing treatment time for surface oxidation. This indicates that implants can be made more hydrophilic by surface oxidation. When the optimized fabrication conditions described above were used, the cell-fibrin constructs were able to cover the MedPor implant. At 24 weeks after implantation, the histomorphological evaluations consistently showed neocartilage formation on the implants. H&E staining revealed the presence of evenly dispersed triangular and ovoid-shaped chondrocytes that inhabited normal-appearing lacunae, and these were surrounded by perichondrium (Figure 5e,f). In addition, safranin-O (Figure 5g) and alcian blue (Figure 5h) staining confirmed the presence of sulfated GAG, indicating that mature cartilage tissue had formed. However, Figure 5d shows that there is minimal cellular interaction between the control implant and host tissue.

Discussion To date, there is no ideal material that can be used for total auricular reconstruction for congenital microtia or traumatic



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Figure 4. Water contents, DNA contents, and elastin contents of the retrieved chondrocyte-fibrin constructs at 4, 8, 12, and 24 weeks after implantation: (a,d,g) 10 U/mL thrombin and 1 × 107 cells/mL initial cell density; (b,e,h) 80 mg/mL fibrinogen and 1 × 107 cells/mL; and (c,f,i) 80 mg/mL fibrinogen and 10 U/mL thrombin. *P < 0.05, n ) 3.

amputation. The porous MedPor ear implant, which is the most widely used material, is relatively resistant to collapse and allows for tissue ingrowth due to mechanical stability and pore structure. However, erosion and infection often occur, especially in grafts covered by thin and scarring soft tissues.9 In contrast, engineered cartilage formed using cells is able to produce tissues that possess physical, histological, and biochemical properties similar to normal cartilage tissue. However, the problem of structural integrity remains unsolved. To overcome these limitations, we designed an auricular prosthesis consisting of a MedPor implant covered with engineered cartilage tissue. This implant possesses the proper characteristics including good biocompatibility, mechanical strength, and long-term stability. This study addressed two important issues. First, we described the fabrication and optimization of chondrocyte-fibrin constructs that could be used in auricular reconstruction. Second,

we achieved coverage of a commercially available ear implant with this engineered cartilage tissue. The engineered cartilagecovered MedPor implant was evaluated, and the biological and physical properties of the implant were found to be appropriate for use in total auricular cartilage reconstruction. This system could be more stable and more compatible with host tissue compared with the alloplastic ear implant alone because the cartilage-covered ear implant is able to improve cellular interactions and possesses enhanced structural stability. This approach needs to be further confirmed using an autologous model. In this study, we selected a fibrin hydrogel as a biological scaffold for cartilage engineering because it is a naturally derived material that offers several advantages over other common scaffold materials. It is well known that by varying the fabrication parameters, such as the concentrations of fibrinogen

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Figure 5. Commercially available ear implant (MedPor) base material: (a) gross appearance and SEM images of (b) surface and (c) crosssection of the implants. MedPor surgical implants are manufactured from linear HDPE, which has a long history of use in surgical implants. The porosity of implant is maintained large with average pore sizes of 100 µm and a pore volume in the 50% range. Scale bar indicates 500 µm. (d) Control implants (MedPor only) and (e) engineered cartilage-covered implants at 24 weeks after implantation. With histological staining, the implants were covered by auricular neocartilage tissue inhabiting typical appearing lacunae and perichondrium as stained by (f) H&E, (g) safraninO, and (h) alcian blue. Scale bar indicates 100 µm.

and thrombin in the gel, the initial properties of fibrin hydrogels can be easily altered. These include gelation time, swelling ratios, mechanical properties, and structural stability.25,31 Fibrin hydrogels were found to be stable over a long term when Ca2+ is used instead of fibrinolysis inhibitors such as aprotinin or tranexamic acid.24,26 As such, we have tested various concentrations of fibrinogen (range of 25-80 mg/mL) to improve longterm stability of these hydrogels. With regard to variation in thrombin concentration, concentrations in the range of 10-200 U/mL did not influence hydrogel appearance or mechanical strength, but the gelation time was accelerated. The gelation time of the hydrogel system is a critical factor for the injection approach. Morphological stability of engineered cartilage is an important factor in successful total auricular cartilage reconstruction. Although we have optimized the fabrication conditions of fibrin hydrogels (80 mg/mL of fibrinogen, 10 U/mL of thrombin, and 4 × 107 cells/mL initial cell density), the hydrogels maintained approximately 82.96 ( 15.88% of their original size up to 24 weeks implantation. This may be due to the relatively rapid degradation characteristic of fibrin hydrogels. However, we show the evidence of increased ECM projection in the cell-fibrin constructs, which likely prevented the loss of massive volume. The dimensions of these gels may be maintained completely when they are used to produce engineered cartilage tissue that totally covers an alloplastic ear implant. MedPor medical devices have been widely used as alloplastic implants for auricular reconstruction. The ear implant is made

from porous, HDPE, which is nontoxic and produces only minor foreign body reactions in vivo. The porosity of MedPor implant is large, with average pore sizes greater than 100 µm and a pore volume in the 50% range (based on mercury intrusion porosimetry measurements). Therefore, the implant allows for tissue ingrowth because of this interconnecting, open pore structure. However, the interface between the implant and host tissue is fragile because of the hydrophobic nature of the MedPor surface. To achieve a hydrophilic surface, a surface oxidation technique (H2SO4/H2O/CrO3, 29/42/29 weight ratio) was used. This procedure results in the creation of hydrophilic carboxylic acid and ketone groups on an oxidatively functionalized surface.30 This demonstrates that the implant surface can be made more hydrophilic, which facilitates penetration of the fibrin hydrogel with cells into the implant and improves the interface between engineered cartilage and the implant. In this study, we successfully developed an engineered cartilage covering for the MedPor implant that can improve the interface between the implant and surrounding host tissue, and provide long-term (up to 24 weeks) stability without causing morphological changes. This may overcome the limitations of the use of alloplastic implants for auricular cartilage reconstruction. Further studies are required to demonstrate the clinical applicability of this device. In vivo studies demonstrating total auricular cartilage reconstruction using an ear-shaped implant are currently being performed.


Conclusions This study demonstrates that cartilage tissues can be engineered to serve as a biological cover for a commercially available ear implant. The chondrocyte-fibrin constructs successfully formed neocartilage tissue with adequate mechanical strength and the long-term stability required for successful clinical application. This system may improve the structural and functional interactions between the implant and recipient tissue, and this in turn may enhance the outcome of auricular cartilage reconstruction by eliminating many of the problems associated with the use of the current ear implant alone. Acknowledgment. We would like to thank Mr. Greg Swords (Porex Surgical Products Group, Newnan, GA) for kindly providing the MedPor ear implants and base materials. We would also like to thank Dr. Jennifer Olson for editorial assistance and Mr. Dong Joon Lee and Dr. Se Heang Oh for technical assistance. This study was partially supported by grants from the Department of Defense (Army Forces Institute of Regenerative Medicine (AFIRM, W81XWH-08-2-0032) and Soldier Treatment and Regeneration Consortium (STRaC, W81XWH-04-1-0848)).

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