Engineering Tissues That Mimic the Zonal Nature of Articular Cartilage

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Engineering tissues that mimic the zonal nature of articular cartilage using decellularized cartilage explants seeded with adult stem cells Lu Luo, Johnnie Chu, Rajalakshmanan Eswaramoorthy, Kevin Mulhall, and Daniel Kelly ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.6b00020 • Publication Date (Web): 08 Mar 2016 Downloaded from http://pubs.acs.org on March 18, 2016

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ACS Biomaterials Science & Engineering

Engineering tissues that mimic the zonal nature of articular cartilage using decellularized cartilage explants seeded with adult stem cells Lu Luo, PhD,1,2 Johnnie Y.J. Chu, MAI,1,2 Rajalakshmanan Eswaramoorthy, PhD,3 Kevin J. Mulhall, MCh FRCSI,4 Daniel J. Kelly, PhD1,2,5,6,*

1

Trinity Centre for Bioengineering, Trinity Biomedical Sciences Institute, Trinity College Dublin, 152-160

Pearse Street, Dublin 2, Ireland. 2

Department of Mechanical and Manufacturing Engineering, School of Engineering, Trinity College

Dublin, College Green, Dublin 2, Ireland. 3

Department of Biomedical Sciences, Sri Ramachandra University, No.1, Ramachandra Nagar, Porur,

Chennai, Tamil Nadu 600116, India 4

Department of Orthopaedic Surgery, Mater Misericordiae University Hospital, Eccles Street, Dublin 7,

Ireland 5

Department of Anatomy, Royal College of Surgeons in Ireland, 123 St Stephen's Green, Dublin 2, Ireland.

6

Advanced Materials and Bioengineering Research Centre (AMBER), Naughton Institute, Royal College

of Surgeons in Ireland and Trinity College Dublin, College Green, Dublin 2, Ireland. *Corresponding author E-mail address: [email protected] Address: Department of Mechanical and Manufacturing Engineering School of Engineering Trinity College Dublin Dublin 2, Ireland Telephone: +353-1-896-3947 Fax: +353-1-679-5554

ACS Paragon Plus Environment


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Abstract Articular cartilage (AC) possesses uniquely complex mechanical properties; for example its stiffness increases with depth through the tissue and it softens when compressed. These properties are integral to the function of AC and can be attributed to the tissue’s collagen network and how it interacts with negatively charged proteoglycans. In this study, scaffolds containing arrays of channels were produced from decellularized AC explants derived from skeletally immature and mature pigs. These scaffolds were then repopulated with human infrapatellar fat pad derived stem cells (FPSCs). After 4 weeks in culture, FPSCs filled channels within the decellularized explants with a matrix rich in proteoglycans and collagen. Cellular and neo-matrix alignment within these scaffolds appeared to be influenced by the underlying collagen architecture of the decellularized cartilage. Repopulating scaffolds derived from decellularized skeletally mature cartilage with FPSCs led to the development of engineered cartilage with depth-dependent mechanical properties mimicking aspects of native tissue. Furthermore, these constructs displayed the characteristic strain softening behavior of AC. These findings highlight the importance of the collagen network to engineering mechanically functional cartilage grafts.

Keywords: Cartilage tissue engineering, Biomimetic, Zonal, Collagen network structure, Extracellular matrix, Recellularization.


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1. Introduction Tissue engineering holds great promise for creating viable grafts to repair cartilage defects. In general, however, most attempts at engineering this tissue in vitro have resulted in the development of homogeneous constructs that do not recapitulate the complex depthdependent properties of native skeletally mature articular cartilage. Such spatially homogenous tissues are more akin to the cartilage that covers our joints at birth, however in skeletally mature articular cartilage the cell morphology and arrangement, extracellular matrix (ECM) composition and structure, and mechanical properties all vary through the depth of the tissue

1-5

. During postnatal development the size and orientation of the

collagen fibrils changes through the depth of articular cartilage, as does the biochemical composition, leading to dramatic increases in the mechanical properties of the tissue and the creation of a tissue with a depth-dependent stiffness and Poisson’s ratio

6-10

. Notably

the apparent compressive modulus of the deep zone of the tissue can increase by an order of magnitude during skeletal development 9, which can be to attributed to the realignment and stiffening of the collagen network which is put into a state of pre-stress by the swelling pressure generated by negatively charged proteoglycans. This leads to the development of a tissue where the compressive modulus increases through its depth

3, 9

. This complex

collagen network is also critical for the capacity of the tissue to generate fluid load support and to articular cartilage lubrication

11-12

. The acquisition of these depth-dependent

properties in tissue engineered grafts is highly desirable and should lead to better integration into the surrounding native cartilage and long-term mechanical and phenotypic stability following implantation, by providing a depth-dependent composition, structure and mechanical properties that matches the surrounding native tissue 1-2, 13.



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To date, some progress has been made in engineering cartilage grafts with a depthdependent composition mimicking aspects of the native tissue

14-20

. These strategies have

varied from modifying the depth-dependent biochemical composition 19 or stiffness 16-17 of a cell laden hydrogel, to the use of a bioreactor to module the local oxygen levels and the mechanical environment through the depth of mesenchymal stem cell laden constructs 20. However, no strategy has been developed to recapitulate the collagen structure of skeletally mature articular cartilage in tissue engineered grafts. In skeletally mature articular cartilage, the collagen fibrils are densely packed and horizontally aligned in the superficial zone but perpendicularly aligned in the deep zone, with the fibrils in the middle zone displaying a more random or arcading organization Benninghoff

21

1-2

, features of the tissue first described by

. Furthermore, while it has been possible to engineer cartilage grafts with

bulk mechanical properties approaching that of the native tissue

22-23

, previous tissue

engineering strategies have failed to recapitulate the complex depth-dependent mechanics of articular cartilage that is associated with the pre-stressed collagen network of skeletally mature tissue. These unique mechanical properties are integral to the functionality of articular cartilage, and the failure to engineer such complexity into cell laden constructs may contribute to the field’s consistent failure to regenerate true hyaline cartilage in damaged synovial joints.

One approach to tissue engineering cartilaginous grafts with such functionality might be to decellularize native articular cartilage in such a way that the depth-dependent collagen structure of the tissue is at least partially maintained

24

, and to then repopulate this

decellularized cartilage explant with chondro-progenitor cells. It is known that cells can


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sense their substrate architecture, align themselves, and potentially synthesize and orient their extracellular matrix in a manner mimicking the architecture of the underlining substrate

25-27

. Furthermore, there is strong emerging evidence suggesting that cartilage

extracellular matrix (ECM) itself is chondro-inductive 28, with numerous studies using this as a biomaterial to create porous scaffolds for cartilage tissue engineering

29-33

. However, a

potential limitation with such approaches is that the inherent structure of the tissue is lost in processing the ECM to produce porous scaffolds. Therefore, decellularized cartilage explants that retain their native structure may represent the ideal template for chondroprogenitor cells to generate a tissue that recapitulates the complex collagen structure and mechanics of articular cartilage. It is hypothesized that cells seeded onto such a graft would act to replace key matrix components (e.g. proteoglycans) lost during the decellularization process, thus restoring the depth-dependent mechanical properties of the tissue.

We have previously shown that porcine articular cartilage can be effectively decellularized whilst maintaining the collagen content and the collagen architecture of the tissue. The decellularization led to a ∼90% removal of the porcine DNA, however, this was accompanied with a dramatic reduction in the mechanical properties of the graft

24

. The

overall objective of this study was to explore if re-populating such decellularized cartilage explants with human stem cells can lead to the development of engineered grafts with a native ‘Benninghoff’ collagen architecture

21

and hence depth-dependent mechanical

properties mimicking the native adult tissue. Decellularized porcine cartilage explants, herein termed scaffolds, were seeded with human infrapatellar fat pad derived stem cells (FPSCs) and maintained in chondrogenic conditions for 4 weeks before extensive biochemical, histological and mechanical assessment. To demonstrate the importance of the



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collagen network to engineering cartilaginous tissues with depth-dependent mechanics mimicking the native adult tissue, we seeded FPSCs into scaffolds derived from decellularized skeletally mature and immature articular cartilage grafts. As the Benninghoff collagen architecture is not fully developed in skeletally immature articular cartilage, it is hypothesized that only by repopulating scaffolds derived from skeletally mature articular cartilage with stem cells is it possible to engineer tissues with depth-dependent mechanical properties mimicking aspects of the native adult tissue.


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2. Materials and methods 2.1. Preparation of decellularized cartilage scaffolds Porcine articular cartilage samples were harvested fresh from skeletally immature (4 months old) and mature (18 months old) animals obtained from the local abattoir (Large white pigs, Female) as previously described 24. Briefly, osteochondral plugs (Ø 6 mm, 10 mm in thickness) were taken from the medial and lateral trochlear ridges of the patellar groove of the knee joints using a Osteochondral Autograft Transfer System (Athrex, Naples, FL, USA) and stored in phosphate buffered saline (PBS; Sigma-Aldrich, Dublin, Ireland) at -85 ◦C. Upon requirement, samples were thawed. The sub-chondral bone of the sample was then removed and the remaining articular cartilage was trimmed to a thickness of approximately 1 mm using a vernier caliper and microtome blades to produce cartilage disks at a uniform size (Ø 6 mm, ∼1 mm in thickness).

These cartilage disks were then decellularized using a previously described protocol

24

. All

decellularization and sterilization procedures were carried out under gentle agitation at room temperature (RT) unless stated otherwise. Briefly, cartilage disks were first subjected to two freeze-thaw cycles. Channels (Ø 400 µm) were then introduced into the disks using flat-ended needles (Fig. 1A) to facilitate decellularization and subsequent recellularization of the samples. The channeled disks were then incubated in hypotonic buffer (10 mM TrisHCl/2 mM EDTA, pH 8.0) containing 100mM KCl, 5 mM MgCl2 and 100 mM dithiothreitol (DTT; all Sigma-Aldrich) for 24 hours, which was followed by another two cycles of incubation in hypotonic buffer containing 0.5% (w/v) sodium dodecyl sulfate (SDS; Sigma-



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Aldrich) for 18 hours at 30 ◦C, with 100 mM DTT added in the first cycle. After this, samples were rinsed in PBS twice (3 hours each time) and treated with 800 U/ml hyaluronidase (Sigma-Aldrich) in PBS for 12 hours at 37 ◦C. Samples were again rinsed in PBS twice, subjected to another freeze-thaw cycle, and then incubated in hypotonic buffer containing 25 U/ml deoxyribonuclease, 25 U/ml ribonuclease (both Sigma-Aldrich), 5 mM MgCl2 and 150 mM NaCl for 16 hours at 37 ◦C. After another two rinses in PBS and one more freezethaw cycle, samples were treated with 0.5 M NaOH in a sonication bath for 5 hours, with the aim of increasing the porosity of the decellularized tissue. Finally, following 3 rinses in PBS and one more freeze-thaw cycle, samples were left swelling in diH2O for 6 hours at 4 ◦C and then freeze-dried at -45 ◦C to produce a scaffold that is suitable for long-term storage.

Prior to cell seeding, scaffolds were sterilized by three cycles of incubation in PBS containing penicillin (500 U/mL)-streptomycin (500 µg/mL), 0.5 µg/ml amphotericin B (both GIBCO, Biosciences, Dublin, Ireland) for 24 hours in total. Scaffolds were then rinsed three times in PBS for a total period of 24 hours to remove any of the chemical residues.

2.2. Isolation and expansion of human stem cells Human infrapatellar fat pad derived stem cells (FPSCs) were isolated from the infrapatellar fat pad of a patient (male, 19 years old) undergoing anterior cruciate ligament reconstruction at Mater Misericordiae University Hospital as previously described 34. Ethical approval was obtained by the ethics board of the Mater Misericordiae University Hospital. Cells were plated at a density of 5x103 cell/cm2 and expanded using high-glucose Dulbecco’s modified Eagle’s medium (hgDMEM) GlutaMAXTM supplemented with 10% fetal bovine


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serum, penicillin (100 U/mL)-streptomycin (100 µg/mL), 0.25 µg/ml amphotericin B (all GIBCO, Biosciences) and 5 ng/ml Fibroblast-growth factor-2 (ProSpec-Tany TechnoGene Ltd, Rehovot, Israel) in a humidified atmosphere of 5% CO2 at 37◦C.

2.3. Cell seeding and construct culture At the end of passage two, cells were trypsinized and seeded onto both sides of the decellularized cartilage scaffolds at a density of 250,000 cells/side (10 x 106 cells/ml, 25 µl/side). Each cell seeded scaffold was cultured in 2 ml chondrogenic medium, consisting of hgDMEM GlutaMAXTM supplemented with penicillin (100 U/ml)-streptomycin (100 µg/ml), 100 µg/ml sodium pyruvate, 40 µg/ml L-proline, 4.7 µg/mL linoleic acid, 50 µg/ml L-ascorbic acid-2-phosphate, 1.5 mg/ml bovine serum albumin (BSA), 1 × insulin–transferrin–selenium, 100 nM dexamethasone (all from Sigma-Aldrich) and 10 ng/ml recombinant human transforming growth factor-β3 (TGF-β3; ProSpec-Tany TechnoGene Ltd), for a total period of 4 weeks. Care was taken to ensure constructs do not flip over throughout the culture period, with the articular surface side of the scaffold facing upward, in order to mimic the decreasing oxygen gradient through the depth of the native tissue 35. Medium was changed twice per week. In the first two weeks, constructs were cultured on a rotational platform (Sigma-Aldrich) to facilitate cell migration into the scaffolds 24. In the last 2 weeks of culture, rotation was stopped to provide a static culture condition for the seeded cells. Constructs were assessed on day 1, 14 and 28.



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2.4. Live/dead staining The viability of cells within the constructs was assessed (n=2 per group) using a live/dead assay kit (VWR International, Dublin, Ireland) as previously described 24. Briefly, constructs were cross sectioned into halves along the channels (see Fig. 1A) and rinsed 5 times in PBS before incubated in PBS containing 2 mM Ethidium Homodimer-1 and 4 mM Calcein for 1 hour at 37 ◦C. Excessive dye was then removed by rinsing in PBS. Samples were visualized immediately under an Olympus FluoView™ FV1000 confocal microscope, with the sectioned plane facing down towards the laser. A series of images were then taken and stacked, starting from the sectioned plane to a depth of 200 µm into the construct, in order to cover the cell distribution within the radius of channels of the scaffold. Calcein stains live cells (shown green) while ethidium Homodimer-1 stains the nucleus of dead cells (shown red).

2.5. DAPI and actin staining The cellular alignment within the scaffolds was assessed (n=2 per group) using a protocol adapted from Steward et al.36. Briefly, constructs were cross sectioned into halves along the channels, fixed in 4% paraformaldehyde (PFA; Sigma-Aldrich) overnight at 4 ◦C and rinsed with PBS. After this, samples were permeabilized in PBS containing 0.5% (v/v) Triton X-100 (Sigma-Aldrich) for 45 min at RT, rinsed again with PBS, and then incubated in a PBS solution containing 1.5% (w/v) BSA, 10 µg/ml 4',6-diamidino-2-phenylindole (DAPI) and 5 U/mL rhodamine phalloidin (both from VWR) for 1.5 hour at RT. Excessive dye was then removed by rinsing in PBS. Samples were visualized immediately under an Olympus FluoView™ FV1000 confocal microscope, with the sectioned plane facing down towards the laser. A


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series of images were then taken and stacked, starting from the sectioned plane to a depth of 200 µm into the construct, as described above. DAPI stains the nuclear of the cells (shown blue) and rhodamine phalloidin stains the F-actin of the cytoskeleton of the cells (shown red).

2.6. Quantitative biochemical analysis The biochemical content of the decellularized scaffolds and cell seeded constructs was also assessed (n=3-4 per group). The wet weight and/or dry weight of the sample was obtained and samples were then digested with 125 µg/ml papain in 0.1 M sodium acetate, 5 mM Lcysteine HCl, 0.05 M EDTA, pH 6.0 (all Sigma-Aldrich) under constant rotation for 18 hours at 60 ◦C. The DNA content of the sample was measured using the Hoechst bisBenzimide H33258 dye assay with calf thymus DNA as a standard

37

. Sulphated glycosaminoglycan

(sGAG) content was assessed using the dimethylmethylene blue dye-binding assay (Blyscan, Biocolor Ltd., Carrickfergus, Northern Ireland), with a glycosaminoglycan standard. Collagen content was determined through measurement of the hydroxyproline content

38

and

calculated using a hydroxyproline-to-collagen ratio of 1:7.69 39.

2.7.

Histology,

polarized

light

microscopy

(PLM)

and

Immunohistochemistry All samples were perpendicularly cut into halves along the channels, fixed in 4% PFA, wax embedded and cross sectioned at 5 µm for histological and immunohistochemical analysis (n=2 per group). Sections were stained with haematoxylin and eosin (H&E) for cellular



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content, and 1% alcian blue 8GX in 0.1 M HCL counter-stained with nuclear fast red for sGAG content (all Sigma-Aldrich). The collagen content and fiber orientation within the sample was assessed by pre-incubating sections in PBS containing 1000 U/ml bovine testicular hyaluronidase for 18 hours at 37 ◦C before staining with picro-sirius red (SigmaAldrich) and visualizing stained sections under normal bright field and polarization. Hyaluronidase treatment was included to remove all sGAG in the sections and thus to enhance the visualisation of the collagen architecture

40

. Sections were also

immunohistochemically stained for type II collagen as previously described

41

. Briefly,

samples were treated with 0.25 unit/ml chondroitinase ABC (Sigma-Aldrich) in a humidified environment at 37 ◦C for 1 hour for antigen retrieval. Non-specific sites were blocked by incubating sections in blocking buffer containing 1% (w/v) BSA and 10% (v/v) goat serum (all Sigma-Aldrich) for 1 hour at room temperature. Sections were then incubated with a mouse monoclonal type II collagen antibody (1:100; 1 mg/mL; Abcam, Cambridge, UK) at 4 ◦C over night. Sections were then blocked for peroxydase activity using 3% (v/v) hydrogen peroxide (Sigma–Aldrich), followed by incubation for 1 hour in the presence of an anti-mouse IgG biotin secondary antibody produced in goat (1:133; 2 mg/mL; Sigma-Aldrich). Color was developed using the Vectastain ABC kit followed by exposure to DAB peroxidase substrate (both Vector Laboratories, Peterborough, UK). Sections of porcine cartilage and ligament were included as positive and negative controls respectively.

2.8. Test of depth-specific mechanical properties Constructs were test for depth-specific mechanical properties (n=4 per group) at day 1 and day 28 using a protocol adapted from Gannon et al. 9. Cores 4 mm in diameter obtained


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from the centre region of the engineered constructs, where tissues were relatively flatter, were used for testing. Next, samples were cross sectioned into halves along the channels and placed in a PBS solution containing 1 µg/ml Hoechst 33342 dye (Sigma-Aldrich) for 1.5 hour, which stained the nuclei of the cells seeded into the scaffolds. Samples were then placed into a custom-built unconfined compression rig with the cross section of the sample being monitored by a fluorescent microscope, which traced the displacement of cells within the channels of the construct during the whole testing period. Due to the sensitivity required in this test (particularly for testing at low strain levels), any outgrowth of neotissue on both sides of the construct was carefully removed prior to testing and a preload of 0.4 N was applied to ensure an even and complete contact between the sample surface and the platen. After the preload, the sample thickness of each group was 0.87 ± 0.20 mm (day1 immature), 0.75 ± 0.09 mm (day1 mature); 0.80 ± 0.14 mm (day28 immature) and 0.79 ± 0.05 mm (day28 mature) respectively. Samples were then compressed such that offset strains of 2.5, 5.0, 7.5, 10.0, 12.5% and 15.0% strains were achieved, where a relaxation period of up to 30 minutes was implemented between each bulk strain increment. The intra-tissue displacement field at each offset strain was calculated using the digital image correlation software VIC 2D 2009 (Correlated Solutions, Columbia, SC, USA). The average axial strain was computed for four consecutive zones through the depth of the constructs: the superficial zone, the middle zone, the upper deep zone and the lower deep zone, which corresponded to the top (from the articular surface) 0-6 %, 7-18 %, 19-30 % and 31-50 %, respectively, of the total construct thickness. The average equilibrium stress was also obtained at each offset strain, using a load cell connected in series with the platens to determine the applied stress. Subsequently, the equilibrium compressive modulus of each



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zone at each offset strain was calculated from the applied stress and the computed local zonal strain values.

2.9. Statistical analysis Statistics were performed using MINITAB 15.1 software (Minitab Ltd., Coventry, UK). A twosample T-test was used for data sets containing only two groups. For data sets which contain more than two groups, a general linear model for analysis of variance with Tukey’s test for multiple comparisons was used. Significance was determined at p≤0.05. All numerical and graphical results are presented as mean ± standard deviation.


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3. Results 3.1. The depth-dependent collagen architecture of skeletally immature and mature articular cartilage is maintained post-decellularization Skeletally immature (4 month old) and mature (18 month old) articular cartilage grafts were decellularized using a previously described protocol 24 that involved introducing an array of seeding channels into the resulting scaffolds (Fig. 1A). H&E staining revealed a nearcomplete removal of cell nuclei through the depth of both decellularized scaffolds, while alcian blue staining also showed a complete removal of sGAG in both groups (Fig. 1B). Both decellularized scaffolds stained intensively for collagen, as evidenced by picrosirius red staining. Importantly, polarized light microscopy (PLM) revealed that the depth-dependent collagen architecture of the skeletally immature and mature cartilage was maintained postdecellularization. Scaffolds derived from decellularized immature tissue displayed a relatively isotropic collagen architecture, except for the superficial and deep zone of the graft (previous studies have shown that the collagen fibrils are horizontally aligned in these regions of 4 month old porcine articular cartilage

42

). In contrast, scaffolds derived from

decellularized mature tissue exhibited a classical Benninghoff collagen architecture, which consists of horizontally aligned collagen fibrils in the superficial zone and perpendicularly aligned collagen fibrils in the deep zone of the tissue (Fig. 1B).

Significantly higher levels of DNA and collagen content were measured in scaffolds derived from mature tissue (p≤0.05 and p≤0.001 respectively); however when normalized to the dry



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weight of the scaffolds, these differences were no longer significant (Table 1). No sGAG was detected in either scaffold type, which is consistent with the alcian blue staining results.

3.2. Decellularized porcine cartilage derived scaffolds support the attachment, viability and proliferation of human stem cells Decellularized skeletally immature and mature porcine articular cartilage derived scaffolds were seeded with human FPSCs, and the viability of the cells inside the scaffolds was monitored over a 4 week culture period. One day after seeding, viable cells were found throughout the channels, attaching to the walls of both immature and mature scaffolds (Fig. 2A). After 14 days of culture, significant cell proliferation was observed in both groups, with cells filling the channels and covering the periphery of the scaffolds. Some dead cells were observed in the bottom regions of both scaffolds by day 28, nevertheless most cells still remained viable (Fig. 2A). The DNA content within both cell seeded scaffolds at day 1 and day 28 was measured, which demonstrated that significant cell proliferation had occurred in both scaffolds over the culture period (Fig. 2B). A near 5-fold increase in DNA content was observed in immature scaffolds, while a near 13-fold increase in DNA content was observed in mature scaffolds.


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3.3. FPSCs secreted higher levels of collagen in scaffolds derived from decellularized skeletally mature cartilage After 28 days of culture, FPSCs had secreted similar levels of sGAG in both decellularized scaffolds, but significantly higher levels of collagen in scaffolds derived from mature tissue (p≤0.05). Similar results were observed when levels of matrix synthesis were normalized to DNA levels (Fig. 3; p≤0.05). The ratio of collagen:sGAG accumulation in scaffolds that derived from immature tissue was ∼0.4, compared to ∼1.4 in mature scaffolds (p≤0.05), approaching that observed in the native tissue (Fig. 3).

Alcian blue staining demonstrated sGAG deposition within the channels of both decellularized scaffolds after 14 days of culture. This sGAG deposition continued with time in culture. By day 28, a neo-tissue staining strongly for alcian blue filled more than half of the depth of most channels in both scaffolds, as well as covering the periphery of the scaffolds (Fig. 4). However, the bottom regions of both scaffold types stained less intensely for sGAG deposition, which is possibly due to lower cell viability in these regions (as shown in Fig. 2A). Cells in the centre of the channels typically displayed a round chondrocyte-like morphology in both scaffolds (Fig. 4). There was some evidence of cell migration from the channels or periphery into the body of the scaffolds; however overall the scale of such migration appeared to be small (Fig. 4). There was little evidence of cell synthesized sGAG diffusing into the body of the scaffolds by this timepoint.

Positive staining for collagen deposition was also observed in the channels of both scaffolds on day 14. By day 28, newly deposited collagen also filled more than half of the depth of



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most channels and covered the periphery of both scaffold types (Fig. 5), with less collagen deposition observed in the bottom region of both scaffolds. Immunohistochemical staining showed this newly deposited matrix stained positively for type II collagen.

3.4. Cellular and neo-matrix alignment within scaffolds appears to be influenced by the underlying collagen architecture of the decellularized cartilage The alignment of the cells seeded within the decellularized cartilage derived scaffolds was assessed throughout the culture period. On day 1, cells displayed a spread shape in both scaffold types. In decellularized immature cartilage derived scaffolds, cells were horizontally aligned in each channel. In contrast, cells appeared generally more perpendicularly aligned in the channels of mature cartilage derived scaffolds, although this cellular alignment was predominant in some channels but less obvious in others (Fig. 6). A similar pattern of cellular alignment was found in the channels of both scaffolds at day 14. For both groups, a small portion of cells had re-organized their actin cytoskeleton, from a spread, tense structure into a more confined and round shape, resembling a chondrocyte-like phenotype. By day 28, most cells displayed this round shape, thus the cellular alignment pattern was no longer obvious. However, some cells were still found to be spread and horizontally aligned in the channels of immature cartilage derived scaffolds, but perpendicularly aligned in the channels of mature cartilage derived scaffolds, similar to that observed on day 1 (Fig. 6).


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The alignment of collagen fibrils secreted by cells within the channels was assessed using PLM. On day 14, the newly synthesized collagen fibrils appeared to be perpendicularly aligned in the channels of scaffolds derived from mature cartilage, mimicking the alignment of the native collagen fibrils (Fig. 7). Collagen fibril alignment was less obvious in the immature cartilage derived scaffolds, which may be due to lower levels of matrix deposition within these scaffolds at this time point (see Fig. 5). A more obvious and predominant pattern of collagen fibril alignment was observed by day 28 in both scaffold types. Collagen fibrils appeared to be perpendicularly aligned through the depth of the channels in the mature cartilage derived scaffolds; whereas they appeared to align horizontally in the lower region of the channels in the immature cartilage derived scaffolds (Fig. 7).

3.5. Repopulating scaffolds derived from decellularized skeletally mature cartilage with FPSCs leads to the development of engineered cartilage with native-like depth-dependent mechanical properties By day 28, the total sGAG content of the engineered grafts (including cell synthesized and the residual in the decellularized scaffold) as a percentage of wet weight was ∼1% for both scaffold types (Fig. 8A). The total collagen as a percentage of wet weight was significantly higher in the mature grafts compared to immature grafts (p≤0.001), similar to that found in native adult articular cartilage (∼10%; Fig 8B).

The equilibrium compressive modulus through the depth of FPSCs seeded constructs (at 5% global offset strain) was estimated after 1 and 28 days of culture. At day 1, both scaffolds appeared very soft (Fig. 9A). The equilibrium modulus of the constructs dramatically



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increased with time in culture, particularly in the scaffolds derived from mature cartilage, and in these engineered tissues the modulus was found to increase with depth as is observed in native adult articular cartilage 9 (Fig. 9A).

We next sought to determine if seeding the scaffolds with FPSCs would lead to the development of cartilaginous grafts demonstrating the characteristic strain softening behavior (a reduction in compressive modulus of the deep zone with increases in applied strain) observed in the native adult tissue 9. To test this, the compressive modulus in the upper deep zone of the engineered tissue was estimated for increasing levels of applied strain. There was little evidence of such strain softening behavior directly after seeding the scaffold with FPSCs (day 1). In contrast, there was a notable reduction in modulus with increasing levels of applied strain in engineered tissues after 28 days of culture (Fig. 9B). Specifically, the modulus of the mature constructs after 28 days of culture decreased from 4.42 MPa at 2.5% offset strain to 0.36 MPa at 10% offset strain (p≤0.05). Strain softening was less obvious in the immature constructs, where the modulus after 28 days of culture decreased from 0.96 MPa at 2.5% offset strain to 0.29 MPa at 10% offset strain.


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4. Discussion The overall objective of the study was to engineer functional cartilage grafts with a collagen structure and depth-dependent mechanical properties mimicking the native adult tissue, using scaffolds derived from decellularized porcine articular cartilage that are subsequently repopulated with human stem cells. As the collagen architecture within articular cartilage evolves postnatally 9, only attaining its characteristic Benninghoff architecture with skeletal maturation, it was hypothesized that only by repopulating scaffolds derived from skeletally mature cartilage with stem cells would it be possible to engineer truly functional cartilaginous grafts. Skeletally immature and mature articular cartilage tissues were decellularized using a previously developed protocol 24. The decellularization procedure led to a near complete removal of cell nuclei and sGAG in both tissue types, whilst maintaining their collagen content and architecture. Both scaffold types supported the attachment, viability, proliferation and chondrogenic differentiation of FPSCs, with significantly higher levels of proliferation and collagen synthesis observed in scaffolds derived from mature cartilage. Cellular and neo-matrix alignment appeared to depend on the underlying collagen architecture of the scaffolds. The impact of this was that the tissues engineered using decellularized mature cartilage scaffolds repopulated with human FPSCs possessed depthdependent compressive mechanical properties and a strain softening behaviour characteristic of the native adult tissue. To the best of our knowledge this is the first time such native tissue-like properties have been recapitulated in engineered cartilage grafts.

In line with previous studies 24, 43-44, the decellularization procedure led to a near complete removal of cell nuclei and sGAG in both skeletally immature and mature cartilage tissues,



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whilst maintaining their collagen content and architecture (Fig. 1B). Decellularization was found to be more effective for immature cartilage tissues, as evidenced by the lower residual DNA levels in the scaffolds (table 1). This may be due to the fact that the ECM of the mature tissue is denser and more organized, hence creating a greater barrier for the diffusion of decellularization solutions and the removal of residual DNA.

Both decellularized cartilage derived scaffolds were found to provide a substrate favoring stem cell proliferation (Fig. 2A), which is consistent with other findings showing that decellularized (or devitalized) cartilage ECM supports the attachment, proliferation and chondrogenic differentiation of stem cells

24, 29, 45-49

. This robust proliferation can be

attributed to a number of factors. First, the residual (primarily type II) collagen network within the decellularized tissue may be supporting cellular growth. Previous studies have found that type II collagen promotes cell proliferation biosynthesis compared to type I collagen

51-53

50

, and enhances chondrocyte

. Secondly, the significant increase in surface

area provided by the introduction of channels into the scaffolds likely enhances cell proliferation. It is well established in the tissue engineering literature that increases in scaffold porosity and/or surface area support stem cell proliferation 54-55. Lastly, any growth factors potentially retained in the scaffolds post-decellularization may further enhance cell proliferation. Some cell death was observed towards the bottom of the channels within the scaffolds. This is likely due to nutrient transfer limitations within such regions of the engineered tissues leading to cell death. Optimal use of bioreactor culture may overcome such challenges.


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Scaffolds derived from skeletally mature cartilage were found to provide a substrate more supportive of stem cell proliferation and collagen synthesis than those derived from immature tissue (Figs. 2B and 3). Previous studies have reported that the thickness of collagen fibrils within porcine articular cartilage increases dramatically with skeletal maturity, from approximately 20 nm in 4 week old cartilage to nearly 200 nm in 3 year old cartilage 9. It has also been demonstrated that increases in fiber diameter in electrospun scaffolds can lead to increases in cell proliferation 56, thus suggesting that the larger sized collagen fibrils in mature cartilage scaffolds may be supporting greater levels of cell proliferation. Chondrogenesis in electrospun scaffolds has also been shown to depend on the fiber diameter, with enhanced collagen type II expression observed on scaffolds with a 300 nm fiber diameter 57, similar to the collagen fibril diameter of mature articular cartilage. Other differences between scaffolds derived from immature and mature cartilage, such as the type of collagen present in the tissue and it structural alignment, may also be contributing to these findings.

The alignment of cells, and the collagen they synthesized, appeared to depend on the alignment of the underling collagen fibrils within the decellularized scaffolds (Figs 6 and 7), implying that the seeded cells sensed the collagen fibrils alignment within the channels. In mature cartilage derived scaffolds, the collagen network that formed in the channels mimicked the classic Benninghoff architecture that is characteristic of fully developed articular cartilage. This result is in line with other studies which show that the underlying substrate architecture can direct cellular and neo-matrix alignment

25-27, 58

and even the

biosynthetic activity of cells 27. For example, it has previously been demonstrated that MSCs can align and generate structurally orientated extracellular matrix within channels as wide



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as 100 µm 58, which suggests that it is not only cells in direct contact with such geometrical features that are influenced by these cues. While the channels introduced within this study are larger (400 µm), it would appear that at least the cells around of the periphery of these channels are responding to these geometrical cues. Interestingly, some newly synthesized collagen fibrils were also found to be aligned in the upper region of the channels of immature cartilage derived scaffolds. This suggests that the immature cartilage used in this study was beginning to transition towards a mature tissue, as evidenced by empty cell lacuna that arranged in a columnar form in the upper region of these decellularized tissues.

Critically, the impact of repopulating scaffolds derived from skeletally mature cartilage with FPSCs was the development of engineered constructs with mechanical properties (i.e. depth-dependent compressive modulus, strain softening) mimicking aspects of the native adult tissue after only 4 weeks of culture (Fig. 9). Following decellularization, which resulted in a complete loss in tissue sGAG content, neither the immature or mature grafts displayed such mechanical behavior and had mechanical properties dramatically lower than the native tissue. This is in agreement with previous studies which demonstrate how proteoglycans electrostatically and non-electrostatically contribute to the compressive properties of articular cartilage

59

. The mechanical properties of the engineered tissues dramatically

improved over 28 days in culture. At low offset strains, the equilibrium modulus of the deep zone of tissues engineered using mature cartilage scaffolds was estimated to be of a similar order of magnitude to our previous measurements of native mature porcine tissue 9. For example, at 2.5% offset strain the mean equilibrium compressive modulus of porcine articular cartilage is 9.4 MPa 9, compared to 4.4 MPa in these engineered tissues. Furthermore, the characteristic strain softening behavior of the articular cartilage re-


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emerged following the deposition of proteoglycans inside the scaffold. Newly deposited proteoglycans within the scaffold will promote the development of an osmotic swelling pressure within the engineered tissue, which in turn will generate a state of pre-stress or tension within the existing collagen network. High loads are initially required to relieve this pre-stress as the tissue is compressed, but once the fibrils are no longer in tension they no longer contribute to the compressive properties of the tissue and hence the engineered construct starts to appear softer with increasing levels of applied strain. Higher loads are required to overcome the osmotic pressure induced pre-stress generated by the thicker and perpendicularly aligned fibrils within the deep zone of the mature cartilage derived scaffolds, which explains the greater increases in compressive modulus with depth in these constructs compared to the grafts engineered using immature cartilage scaffolds.

One of the potential weaknesses of this study is the limited cellular infiltration into the body of scaffolds (Fig. 4), likely due to the dense nature of the cartilage ECM. This challenge has previously been reported when attempting to repopulate decellularized cartilage explants 43, 60

. For example, cellular infiltration up to only 300 µm into the body of a decellularized

cartilage-bone scaffold has been reported 3 months after implantation into nude mice

43

.

Such challenges may potentially be overcome by increasing the number of channels in the decellularized scaffolds, which may accelerate cellular infiltration and the tissue remodelling process. Even without such modifications, it is expected that such scaffolds will be slowly recellularized over time following implantation into a load-bearing orthotopic defect. In vivo validation in relevant pre-clinical models is required to test this hypothesis. Another impact of this limited cellular infiltration, and the channelled geometry of the scaffold, is that the resultant construct possesses a heterogeneous composition and hence mechanical



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properties in the radial direction. Therefore, the reported mechanical properties in the axial direction can only be considered an estimate, as the method used to calculate the depth dependent properties assumes that the tissue is otherwise homogenous in planes parallel to the loading platen. Lastly, the thickness of the scaffolds produced in this study is limited to that of the porcine adult articular cartilage (∼1mm thick) which may impact its clinical application (the thickness of human knee cartilage ranges from 1.7 ∼2.5 mm 61). However, this may be overcome by switching to an alternatively cartilage source, such as bovine articular cartilage.

5. Conclusion This study highlights the importance of collagen network architecture on engineering zonal cartilage grafts. Scaffolds derived from decellularized skeletally mature articular cartilage, which maintain the original collagen structure of the tissue, can facilitate the development of engineered cartilage grafts with a structure and depth-dependent mechanical properties mimicking aspects of the native adult tissue. The results of this study may also inspire the design of novel scaffolds derived from natural or synthetic polymers to produce biomaterials that mimic many of the structural features of the decellularized scaffolds, but which have an appropriate thickness and a higher porosity to facilitate greater cellular migration. Indeed, many novel scaffold fabrication techniques are now available that may facilitate this

62-64

. Manipulating scaffold architecture to mimic the collagen network

structure of the native adult tissue appears to be critical for engineering functional articular cartilage grafts.


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Acknowledgements Funding was provided by the European Research Council Starter Grant (StemRepair — Project number 258463). We are thankful to Mr. Simon Carroll for designing the tool to make the channels within cartilage grafts. We also thank Dr. Conor Buckley and Dr. Binulal Sathy for technical advice.

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in porcine articular cartilage during growth and maturation. Osteoarthr. Cartil. 2009, 17 (4), 448-55. DOI: 10.1016/j.joca.2008.09.004. 43. Kheir, E.; Stapleton, T.; Shaw, D.; Jin, Z.; Fisher, J.; Ingham, E., Development and characterization of an acellular porcine cartilage bone matrix for use in tissue engineering. J. Biomed. Mater. Res., Part A 2011, 99 (2), 283-94. DOI: 10.1002/jbm.a.33171. 44. Elder, B. D.; Kim, D. H.; Athanasiou, K. A., Developing an articular cartilage decellularization process toward facet joint cartilage replacement. Neurosurgery 2010, 66 (4), 722-7; discussion 727. DOI: 10.1227/01.neu.0000367616.49291.9f. 45. Kang, H.; Peng, J.; Lu, S.; Liu, S.; Zhang, L.; Huang, J.; Sui, X.; Zhao, B.; Wang, A.; Xu, W.; Luo, Z.; Guo, Q., In vivo cartilage repair using adipose-derived stem cell-loaded decellularized cartilage ECM scaffolds. J. Tissue Eng. Regener. Med. 2014, 8 (6), 442-53. DOI: 10.1002/term.1538. 46. Diekman, B. O.; Rowland, C. R.; Lennon, D. P.; Caplan, A. I.; Guilak, F., Chondrogenesis of adult stem cells from adipose tissue and bone marrow: induction by growth factors and cartilagederived matrix. Tissue Eng., Part A 2010, 16 (2), 523-33. DOI: 10.1089/ten.TEA.2009.0398. 47. Rowland, C. R.; Lennon, D. P.; Caplan, A. I.; Guilak, F., The effects of crosslinking of scaffolds engineered from cartilage ECM on the chondrogenic differentiation of MSCs. Biomaterials 2013, 34 (23), 5802-12. DOI: 10.1016/j.biomaterials.2013.04.027. 48. Almeida, H. V.; Liu, Y.; Cunniffe, G. M.; Mulhall, K. J.; Matsiko, A.; Buckley, C. T.; O'Brien, F. J.; Kelly, D. J., Controlled release of transforming growth factor-beta3 from cartilage-extra-cellularmatrix-derived scaffolds to promote chondrogenesis of human-joint-tissue-derived stem cells. Acta Biomater. 2014, 10 (10), 4400-9. DOI: 10.1016/j.actbio.2014.05.030. 49. Almeida, H. V.; Cunniffe, G. M.; Vinardell, T.; Buckley, C. T.; O'Brien, F. J.; Kelly, D. J., Coupling Freshly Isolated CD44(+) Infrapatellar Fat Pad-Derived Stromal Cells with a TGF-beta3 Eluting Cartilage ECM-Derived Scaffold as a Single-Stage Strategy for Promoting Chondrogenesis. Adv. Healthcare Mater. 2015, 4 (7), 1043-53. DOI: 10.1002/adhm.201400687. 50. Qi, W. N.; Scully, S. P., Extracellular collagen modulates the regulation of chondrocytes by transforming growth factor-beta 1. J. Orthop. Res. 1997, 15 (4), 483-490. DOI: DOI 10.1002/jor.1100150402. 51. Bosnakovski, D.; Mizuno, M.; Kim, G.; Takagi, S.; Okumura, M.; Fujinaga, T., Chondrogenic differentiation of bovine bone marrow mesenchymal stem cells (MSCs) in different hydrogels: influence of collagen type II extracellular matrix on MSC chondrogenesis. Biotechnol. Bioeng. 2006, 93 (6), 1152-63. DOI: 10.1002/bit.20828. 52. Nehrer, S.; Breinan, H. A.; Ramappa, A.; Young, G.; Shortkroff, S.; Louie, L. K.; Sledge, C. B.; Yannas, I. V.; Spector, M., Matrix collagen type and pore size influence behaviour of seeded canine chondrocytes. Biomaterials 1997, 18 (11), 769-76. 53. Veilleux, N. H.; Yannas, I. V.; Spector, M., Effect of passage number and collagen type on the proliferative, biosynthetic, and contractile activity of adult canine articular chondrocytes in type I and II collagen-glycosaminoglycan matrices in vitro. Tissue Eng. 2004, 10 (1-2), 119-27. DOI: 10.1089/107632704322791763. 54. Murphy, C. M.; Haugh, M. G.; O'Brien, F. J., The effect of mean pore size on cell attachment, proliferation and migration in collagen-glycosaminoglycan scaffolds for bone tissue engineering. Biomaterials 2010, 31 (3), 461-6. DOI: 10.1016/j.biomaterials.2009.09.063. 55. Matsiko, A.; Gleeson, J. P.; O'Brien, F. J., Scaffold mean pore size influences mesenchymal stem cell chondrogenic differentiation and matrix deposition. Tissue Eng., Part A 2015, 21 (3-4), 48697. DOI: 10.1089/ten.TEA.2013.0545. 56. Badami, A. S.; Kreke, M. R.; Thompson, M. S.; Riffle, J. S.; Goldstein, A. S., Effect of fiber diameter on spreading, proliferation, and differentiation of osteoblastic cells on electrospun poly(lactic acid) substrates. Biomaterials 2006, 27 (4), 596-606. DOI: 10.1016/j.biomaterials.2005.05.084.


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57. Noriega, S. E.; Hasanova, G. I.; Schneider, M. J.; Larsen, G. F.; Subramanian, A., Effect of fiber diameter on the spreading, proliferation and differentiation of chondrocytes on electrospun chitosan matrices. Cells Tissues Organs 2012, 195 (3), 207-21. DOI: 10.1159/000325144. 58. Chou, C. L.; Rivera, A. L.; Sakai, T.; Caplan, A. I.; Goldberg, V. M.; Welter, J. F.; Baskaran, H., Micrometer scale guidance of mesenchymal stem cells to form structurally oriented cartilage extracellular matrix. Tissue Eng., Part A 2013, 19 (9-10), 1081-90. DOI: 10.1089/ten.TEA.2012.0177. 59. Canal Guterl, C.; Hung, C. T.; Ateshian, G. A., Electrostatic and non-electrostatic contributions of proteoglycans to the compressive equilibrium modulus of bovine articular cartilage. J. Biomech. 2010, 43 (7), 1343-50. DOI: 10.1016/j.jbiomech.2010.01.021. 60. Schwarz, S.; Elsaesser, A. F.; Koerber, L.; Goldberg‐Bockhorn, E.; Seitz, A. M.; Bermueller, C.; Dürselen, L.; Ignatius, A.; Breiter, R.; Rotter, N., Processed xenogenic cartilage as innovative biomatrix for cartilage tissue engineering: effects on chondrocyte differentiation and function. J. Tissue Eng. Regener. Med. 2012. DOI: doi: 10.1002/term.1650. 61. Shepherd, D. E.; Seedhom, B. B., Thickness of human articular cartilage in joints of the lower limb. Ann. Rheum. Dis. 1999, 58 (1), 27-34. 62. Moutos, F. T.; Freed, L. E.; Guilak, F., A biomimetic three-dimensional woven composite scaffold for functional tissue engineering of cartilage. Nat. Mater. 2007, 6 (2), 162-7. DOI: 10.1038/nmat1822. 63. McCullen, S. D.; Autefage, H.; Callanan, A.; Gentleman, E.; Stevens, M. M., Anisotropic fibrous scaffolds for articular cartilage regeneration. Tissue Eng., Part A 2012, 18 (19-20), 2073-83. DOI: 10.1089/ten.TEA.2011.0606. 64. Steele, J. A.; McCullen, S. D.; Callanan, A.; Autefage, H.; Accardi, M. A.; Dini, D.; Stevens, M. M., Combinatorial scaffold morphologies for zonal articular cartilage engineering. Acta Biomater. 2014, 10 (5), 2065-75. DOI: 10.1016/j.actbio.2013.12.030.



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Figure 1. (A): Schematic representation of the transverse and cross section of the channeled cartilage explants. In total 7 channels were introduced to each cartilage disk. (B): Alcian blue, picrosirius red and H&E staining as well as PLM images of collagen architecture of decellularized skeletally immature and mature cartilage derived scaffolds prior to cell seeding. Images for alcian blue and picrosirius red were taken at 20x magnification. H&E images were taken at 200x magnification, demonstrating cell removal through the depth of the decellularized tissue, with images shown in the box region taken at 400x magnification. PLM images were taken at 100x magnification, showing the collagen architecture of decellularized cartilage scaffolds. Bright color in PLM images indicates the existence of aligned collagen fibrils. Dark regions within the scaffolds indicate a lack of collagen alignment in these areas. 146x165mm (300 x 300 DPI)


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Figure 2. (A): viability of FPSCs seeded in decellularized immature and mature cartilage derived scaffolds at day 1, 14 and 28. Live cells appear green and dead cells appear red. Magnified images on the right column show some cell death at day 28. (B): DNA content of the cells seeded in decellularized immature and mature cartilage derived scaffolds at day 1 and 28. The DNA content of the cells is calculated by subtracting the residual DNA content of the scaffolds after decellularization from the total DNA content of the constructs. *: p≤0.05; ***: p≤0.001. 185x263mm (300 x 300 DPI)



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Figure 3. sGAG and collagen synthesis by FPSCs within the immature and mature scaffolds at day 28. The levels of sGAG (or collagen, or DNA) secreted by cells is calculated by subtracting the residual sGAG (or collagen, or DNA) level within the scaffolds after decellularization from the total sGAG (or collagen, or DNA) level in the engineered constructs at day 28. *: p≤0.05. 59x27mm (600 x 600 DPI)


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Figure 4. Alcian blue staining of cell seeded immature and mature cartilage derived scaffolds at day 14 and 28. Images from first and second row were taken at 20x magnification. Images from third row were taken at 100x magnification, showing cell morphology and sGAG distribution within the channels, with inserted images taken at 400x magnification. Images from last row were taken at 200x magnification, showing cell migration into the body of the decellularized scaffolds at sites shown in box regions in images from the second row. 193x288mm (300 x 300 DPI)



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Figure 5. Picrosirius red and type II collagen staining of cell seeded immature and mature cartilage derived scaffolds at day 14 and 28. Images from first and second row were taken at 20x magnification. Images from third row were taken at 100x magnification. Images from last row were taken at 200x magnification. 193x287mm (300 x 300 DPI)


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Figure 6. DAPI and F-actin staining showing cell alignment within the channels of cell seeded immature and mature cartilage derived scaffolds at day 14 and 28. Images in first and third column represent the highest degree of cell alignment observed in the two groups at corresponding time points, while images in second and fourth column represent the average degree of cell alignment observed in the two groups. Images from first to third row were all taken at 200x magnification. Images from the fourth row show the magnified images of the box regions in third row. 140x152mm (300 x 300 DPI)



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Figure 7. PLM images of collagen alignment within the channels of the cell seeded immature and mature cartilage derived scaffolds at day 14 and 28. All images were taken at 100x magnification. 172x229mm (300 x 300 DPI)


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Figure 8. (A) total sGAG as a % wet weight and (B) total collagen as a % wet weight of cartilaginous tissues engineered using immature and mature cartilage derived scaffolds at day 28; ***: p≤0.001. 30x11mm (600 x 600 DPI)



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Figure 9. (A): The estimated equilibrium modulus (for a global compressive offset strain of 5%) of the superficial zone (SZ), middle zone (MZ), upper deep zone (DZ-U) and lower deep zone (DZ-L) of cartilaginous tissues engineered using immature and mature cartilage derived scaffolds at day 1 and day 28. (B)：The estimated equilibrium modulus of the upper deep zone of the engineered tissues under 2.5%, 5% 7.5%, 10%, 12.5% and 15% global offset strain at day 1 and day 28. *:p≤0.05; **: p≤0.01; #:p≤0.05 vs. day1; ##: p≤0.01 vs. day1. 108x90mm (300 x 300 DPI)


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Table 1. DNA, sGAG and collagen content, dry and wet weight, DNA/dry weight, collagen %dry weight, DNA/wet weight and collagen %wet weight of the decellularized skeletally immature and mature cartilage derived scaffolds. ns: not significant; *:p≤0.05; **: p≤0.01; ***: p≤0.001. 43x14mm (600 x 600 DPI)



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For Table of Contents Only 77x42mm (300 x 300 DPI)


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Engineering Tissues That Mimic the Zonal Nature of Articular Cartilage

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