Adipose tissue engineering with naturally derived scaffolds and adipose-derived stem cells

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Biomaterials 28 (2007) 3834–3842 www.elsevier.com/locate/biomaterials

Adipose tissue engineering with naturally derived scaffolds and adipose-derived stem cells Lauren Flynna,b, Glenn D. Prestwichc, John L. Sempled,e, Kimberly A. Woodhousea,b, a

Department of Chemical Engineering and Applied Chemistry, University of Toronto, 200 College Street, Toronto, Ont., Canada, M5S 3E5 b Institute of Biomaterials and Biomedical Engineering, University of Toronto, 4 Taddle Creek Road, Toronto, Ont., Canada, M5S 3G9 c Center for Therapeutic Biomaterials and Department of Medicinal Chemistry, University of Utah, 419 Wakara Way, Suite 205, Salt Lake City, Utah 84108-12, USA d Division of Plastic Surgery, Department of Surgery, University of Toronto, 100 College Street, Toronto, Ont., Canada, M5G 1L5 e Women’s College Hospital, 76 Grenville Street, Toronto, Ont., Canada, M5S 1B2 Received 28 February 2007; accepted 4 May 2007 Available online 16 May 2007

Abstract A tissue-engineered adipose substitute would have numerous applications in plastic and reconstructive surgery. This work involves the characterization of the in vitro cellular response of primary human adipose-derived stem cells (ASC) to three dimensional, naturally derived scaffolds. To establish a more thorough understanding of the influence of the scaffold environment on ASC, we have designed several different soft tissue scaffolds composed of decellularized human placenta and crosslinked hyaluronan (XLHA). The cellular organization within the scaffolds was characterized using confocal microscopy. Adipogenic differentiation was induced and the ASC response was characterized in terms of glycerol-3-phosphate dehydrogenase (GPDH) activity and intracellular lipid accumulation. The results indicate that the scaffold environment impacts the ASC response and that the adipogenic differentiation of the ASC was augmented in the non-adhesive XLHA gels. r 2007 Elsevier Ltd. All rights reserved. Keywords: Adipose tissue engineering; Scaffold; Extracellular matrix; Hyaluronan; Stem cells

1. Introduction Soft tissue augmentation is a major challenge for plastic and reconstructive surgeons. Subcutaneous adipose tissue loss is associated with numerous conditions including traumatic injury, oncologic resection and congenital birth defects [1]. The resultant scar tissue formation can lead to a loss of contour, as well as functional impairment, particularly if the defect occurs in proximity to a joint [2]. In 2005, approximately 5.4 million people underwent reconstructive surgery in the United States, with 3.9 million cases associated with tumor removal [3]. In addition, over 10.2 million cosmetic procedures were performed, includCorresponding author. Department of Chemical Engineering and Applied Chemistry, University of Toronto, 200 College Street, Toronto, Ont., Canada, M5S 3E5. Tel.: +416 978 3060; fax: +416 978 8605. E-mail address: [email protected] (K.A. Woodhouse).

0142-9612/$ - see front matter r 2007 Elsevier Ltd. All rights reserved. doi:10.1016/j.biomaterials.2007.05.002

ing various forms of soft tissue augmentation with biological or synthetic fillers and implants [3]. The current treatment strategies for soft tissue reconstruction primarily involve tissue transplantation, including composite tissue flaps, or synthetic substitutes. The use of autologous tissues is associated with the creation of a donor site defect and, potentially, the need for multiple, complex and costly surgical procedures [4]. Moreover, the clinical outcome of adipose tissue transplantation is unpredictable, with graft resorption due to a lack of vascularization resulting in poor cosmesis and impaired functionality [5]. Synthetic implants are associated with immune rejection, implant migration and resorption, and a failure to integrate into the host tissues [6]. A tissue-engineered adipose substitute, that would promote regeneration, rather than repair, would be invaluable to plastic and reconstructive surgeons. The substitute should incorporate a biocompatible scaffold that

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defines the appropriate three-dimensional tissue architecture and promotes host integration and implant vascularization [7]. Ultimately, the construct should degrade as it is replaced by healthy host soft tissue. A number of different synthetic scaffolds have been investigated for adipose tissue engineering applications including polyethylene glycol diacrylate (PEDGA) [8], polyglycolic acid (PGA) [9], polyethylene terephthalate (PET) [10], poly(lactic-co-glycolic acid) (PLGA) [11] and polytetrafluoroethylene (PTFE) [12]. Naturally derived materials such as collagen [13], derivatives of hyaluronic acid [14], matrigel [15] and fibrin [16] have also been studied. There are many factors to consider when designing a scaffold including the mechanical properties, degradation characteristics, immunogenicity, cellular response to the material, ease of handling in the clinic and cost [17]. Adipose-derived stem cells (ASC) may be an ideal autologous cell source for adipose tissue engineering [18]. ASC are much more resistant to mechanical damage and ischemic conditions than mature adipocytes [19]. The cells, which can be readily harvested from excised human subcutaneous fat or liposuction samples, have been shown to proliferate rapidly and differentiate into mature adipocytes both in vitro and in vivo [20–22]. The development of methods to maintain the ASC differentiation potential in culture, while obtaining sufficient cell populations for transplantation, will be critical to the clinical application of these cells [23]. Further, the optimization of the growth and differentiation conditions to maximize stable adipose tissue formation is required [24]. With a view to develop a tissue-engineered adipose substitute, we are investigating the response of human ASC to scaffolds comprised of placental decellular matrix (PDM) and crosslinked hyaluronan (XLHA). By investigating several different scaffolds, it is possible to obtain a more thorough understanding of the impact of the matrix environment on the ASC. We previously developed a detergent and enzymatic extraction protocol to fully decellularize human placenta [25]. We believe that the PDM holds promise as a scaffold for adipose tissue engineering applications. The placenta is a rich source of human extracellular matrix (ECM) components that can be harvested without harm to the donor. Constructs derived from the ECM may mimic the native environment of the body, promoting normal cellular organization and behavior. Natural materials also have advantages in terms of ease of processing, biodegradability and biocompatibility [26]. Hyaluronan (HA) is a highly conserved glycosaminoglycan (GAG) that functions in matrix stabilization, cell signaling, adhesion, migration, proliferation and differentiation [27]. Incorporation of XLHA into the PDM scaffolds may improve the construct bulking properties and may influence cellular infiltration, differentiation and wound healing [28]. The primary objective of this research was to develop effective seeding protocols and in vitro culture conditions

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for the ASC with the PDM and XLHA. The cellular organization was characterized in each of the scaffolds. The influence of the scaffold environment on the differentiation of the ASC was also investigated. 2. Materials and methods 2.1. Materials All chemicals used in the experiments, unless otherwise stated, were purchased from Sigma-Aldrich Canada Ltd. (Oakville, Canada) and were used as received. Water was distilled and deionized using a Millipore MilliRO 10 Plus filtration system at 18 MO resistance.

2.2. Cell culture Primary cultures of human ASC were established according to the methods of Flynn et al. [25]. The ASC were isolated from freshly excised subcutaneous abdominal adipose tissue from patients undergoing elective surgery at Women’s College Hospital, Toronto, Canada, following the ethical guidelines of the University of Toronto. The cells were plated in a 1:1 mixture of Dulbecco’s Modified Eagle’s Medium and Ham’s F-12 nutrient mixture (DMEM:Ham’s F-12), supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin and 0.1 mg/mL streptomycin. The growth medium was changed every 2–3 days. To passage the cells, cultures at 90% confluence were trypsin-released (0.25% trypsin/0.1% EDTA, Gibco, Burlington, Canada), washed, counted and re-plated in new flasks at 30,000 cells/cm2. Passage 2 cells were used for the seeding experimentation.

2.3. ASC differentiation To induce adipogenic differentiation, the cells were cultured in serumfree DMEM:Ham’s F-12 supplemented with 15 mM NaHCO3, 15 mM HEPES, 33 mM biotin, 17 mM pantothenate, 10 mg/mL transferrin, 100 nM cortisol, 66 nM insulin, 1 nM triiodothyronine (T3), 100 U/mL penicillin and 0.1 mg/mL streptomycin. For the first 72 h of differentiation, 0.25 mM isobutylmethylxanthine (IBMX) and 1 mg/mL of troglitazone were added to the differentiation medium [29].

2.4. Preparation of the PDM Placentas were obtained, with informed consent, from normal-term Caesarian-section deliveries at Women’s College Hospital, Toronto, Canada and were decellularized as previously described [25]. In brief, the perfusive and diffusive decellularization protocol involved treatment with hypotonic and hypertonic solutions, enzymatic digestion, and multiple detergent extractions. Research ethics board approval for this study was obtained from Women’s College Hospital, Toronto, Canada (REB # 9918). Histological analysis was conducted on representative sections of the processed tissues to confirm the effectiveness of the extraction protocol. Following decellularization, the PDM was sectioned into samples by mass, with each scaffold consisting of a 300 mg portion of the villous tree network. The scaffolds were decontaminated by three 30 min rinses in ethanol, re-hydrated with three washes in sterile phosphate buffered saline (PBS) and stored at 4 1C in sterile PBS supplemented with 100 U/mL penicillin and 0.1 mg/mL streptomycin. The PDM scaffolds that were used in the confocal analyses were labeled with an amine reactive Alexa Fluors 350 carboxylic acid, succinimidyl ester (Molecular Probes, Burlington, Canada) to facilitate visualization. Briefly, the dye was dissolved in dimethyl sulfoxide (DMSO) at a concentration of 10 mg/mL. This stock solution was diluted with 0.15 M NaHCO3 (pH 8.3) to obtain a working concentration of 0.3 mg/mL. The PDM scaffolds were agitated in the dye solution for 1 h at room

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temperature (0.3 mg dye/300 mg PDM). To stop the labelling reaction, the PDM was incubated for an additional hour in 1.5 M hydroxylamine (pH 8.5). The scaffolds were then rinsed three times in sterile PBS and stored at 4 1C in sterile PBS supplemented with 100 U/mL penicillin and 0.1 mg/mL streptomycin. All work was conducted under minimal lighting conditions.

2.5. Thiolated HA preparation Thiol-modified HA (HA-DTPH) was synthesized according to the methods of Shu et al. [30]. GPC and 1H NMR was used to assess the purity and degree of substitution (SD) of the HA-DTPH. The HA-DTPH used in the experimentation had an SD ¼ 57%.

solution of the PEGDA was prepared in PBS. The solutions were sterilized by syringe filtration with a 0.22 mm filter. To initiate crosslinking, the PEGDA solution was added to the HA–DTPH solution in a ratio of 1:4 (v:v) and mixed thoroughly. Immediately following mixing, the ASC were resuspended in the HA–DTPH–PEGDA solution at a concentration of 3.33  106 cells/mL. To fabricate each scaffold with a seeding density of 1  106 cells, 300 mL of the cell-seeded HA–DTPH–PEGDA was added to a well in a 24-well plate and allowed to gel for 1 h (37 1C, 5% CO2). The scaffolds were then gently detached from the surface using blunt forceps and transferred into 6-well plates containing growth medium (5 mL/well).

2.6.1. PDM scaffold fabrication The ASC were suspended in growth medium at a concentration of 2  107 cells/mL. Each PDM scaffold was transferred to a 24-well plate and seeded with 1  106 ASC in 50 mL of growth medium. The scaffolds were incubated for 3 h (37 1C, 5% CO2) to facilitate cell adhesion and then transferred into 6-well plates containing growth medium (5 mL/well).

2.6.3. PDM with XLHA scaffold fabrication Two different seeding methodologies (Fig. 1) were investigated to assess whether there was a significant impact on the organization and differentiation of the ASC in the combined PDM with XLHA scaffolds. The first method was termed the PDM with XLHA ‘‘Encapsulation’’ methodology. The 300 mg PDM scaffolds were placed in a 24-well plate and an HA–DTPH–PEGDA solution was prepared, as described in the previous section. Immediately following mixing, the ASC were resuspended in the HA–DTPH–PEGDA at a concentration of 4  106 cells/mL. For each PDM sample, 250 mL of the cell-seeded HA–DTPH–PEGDA was added and allowed to gel for 1 h (37 1C, 5% CO2), to fabricate a combined construct seeded with 1  106 cells. The scaffolds were then transferred into 6-well plates containing growth medium (5 mL/well). The second seeding method was defined as the PDM with XLHA ‘‘Adhesion’’ methodology. The ASC were suspended in growth medium at a concentration of 2  107 cells/mL. The 300 mg PDM scaffolds were placed in a 24-well plate and each scaffold was seeded with 1  106 ASC in 50 mL of growth medium. The PDM scaffolds were incubated (37 1C, 5% CO2) for 2 h to facilitate cell adhesion. Following this, an HA–DTPH– PEGDA solution was prepared, as described in the previous section. A 250 mL volume of the HA–DTPH–PEGDA solution was added to each cell-seeded PDM, and the constructs were incubated (37 1C, 5% CO2) for an additional hour to facilitate gelation. The combined scaffolds were then transferred into 6-well plates containing growth medium (5 mL/well).

2.6.2. XLHA scaffold fabrication The XLHA hydrogels were fabricated according to the methods of Shu et al [31]. This methodology uses polyethylene glycol-diacrylate (PEGDA; MW 3400; Nektar Transforming Therapeutics, Huntsville AL) to rapidly crosslink the HA-DTPH to form a gel in situ. Briefly, a 12.5 mg/mL solution of HA–DTPH was prepared in DMEM:Ham’s F-12 growth medium and the pH was adjusted to 7.4 with 1.0 M NaOH. A 4.5% (w/v)

2.6.4. Scaffold culture Following seeding, the scaffolds were incubated (37 1C, 5% CO2) under agitation on a rotomix in the 6-well plates containing growth medium. The medium was changed 24 h after seeding and every 2 days thereafter. To investigate cellular differentiation within the scaffolds, after a 72-h growth period, the scaffolds were rinsed in sterile PBS and transferred into the previously defined differentiation medium. The constructs were incubated

2.6. Scaffold fabrication Four different scaffold groups were investigated to characterize the influence of the scaffold environment on the ASC population: the PDM alone, the XLHA alone and the PDM in combination with the XLHA using two different seeding methodologies. Prior to cell seeding, the PDM was incubated for 24 h in serum-free DMEM:Ham’s F-12 (37 1C, 5% CO2) supplemented with 100 U/mL penicillin and 0.1 mg/mL streptomycin. Passage 2 ASC at 90% confluence were trypsin-released, washed and counted using trypan blue exclusion. For each of the scaffold groups, the cellular seeding density was 1  106 cells/scaffold. This seeding density was selected based on our previous in vitro work that indicated that higher seeding densities promoted ASC viability on the PDM scaffolds in culture [25].

Fig. 1. The two seeding methodologies for the combined PDM with XLHA scaffolds.

ARTICLE IN PRESS L. Flynn et al. / Biomaterials 28 (2007) 3834–3842 (37 1C, 5% CO2) under agitation for up to 14 days after the induction of differentiation.

2.7. Cellular organization Cellular organization in the scaffolds and on tissue culture polystyrene (TCPS) controls was assessed at 24 and 72 h after seeding using confocal microscopy (Zeiss LSM510, FLUAR 20x/0.75 NA objective lens, excitation with an Argon laser at 488 nm and an Enterprise laser at 351 nm). The ASC were labeled prior to seeding with Cell TrackerTM Green 5-chloromethylfluorescein diacetate (CMFDA) (Invitrogen, Burlington, Canada) according to the manufacturer’s instructions. The PDM scaffolds were labeled with the Alexa Fluors 350 carboxylic acid, succinimidyl ester, as described in Section 2.4. All work was conducted under minimal lighting conditions. Samples from the XLHA scaffold group were also stained with calceinAM and ethidium homodimer-1 (LIVE/DEADs Viability/Cytotoxicity Kit, Invitrogen, Burlington, Canada) to differentiate between live and dead cells, respectively. Live/dead staining was not conducted on the scaffolds incorporating the PDM, as the ethidium homodimer-1 bound strongly to the PDM matrix, interfering with the imaging process.

2.8. Glycerol-3-phosphate dehydrogenase (GPDH) activity To assess the impact of the scaffold microenvironment on the differentiation of the cells, the cellular GPDH activity levels in each of the scaffold groups were measured at 72 h, 7 and 14 days after the induction of differentiation (n ¼ 3, N ¼ 6). The positive control was ASC differentiated on TCPS and the negative control was undifferentiated ASC on TCPS. For comparative purposes, the assay was also conducted on samples of freshly-excised human subcutaneous adipose tissue from 4 different donors. A GPDH activity measurement kit (Kamiya Biomedical Corporation) was used in the quantification. Briefly, this kit provides a substrate reagent containing NADH, which is a co-enzyme for GPDH, and dihydroxyacetone phosphate (DHAP). The activity level was quantified by spectrophotometrically measuring the decrease in the NADH concentration as the DHAP was converted into glycerol-3-phosphate by the GPDH. Within each sample, the absorbance (340 nm) was measured over a 10 min period at 25 1C using a microplate reader. The change in absorbance (DOD/min) was determined from the linear portion of the kinetic curve and each sample was assayed in duplicate. The activity levels were normalized in terms of the total cytosolic protein content within each sample, measured in triplicate using the Bio-Rad protein assay, with an albumin standard. The data was expressed in terms of mUnits/mg protein (mU/mg), where 1 unit was defined as the GPDH activity required to oxidize 1 mmol of NADH in 1 min. In preparation for the GPDH and protein assays, each scaffold or adipose tissue sample was minced in 1 mL of 0.25 M sucrose solution at 4 1C and disrupted with 3 five-s bursts of sonication, with intervals of cooling on ice. The samples were centrifuged at 16,000g for 10 min (4 1C). The supernatant was transferred into a new 2 mL tube and centrifuged for an additional hour (16,000g, 4 1C) to isolate the cytosolic protein fraction, including the GPDH. The TCPS control samples were disrupted with sonication in the enzyme extraction buffer provided in the GPDH assay kit, and centrifuged at 16,000g (4 1C) for 10 min. All supernatant samples were immediately assayed for GPDH activity and total protein content according to the manufacturers’ instructions.

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neutral buffered formalin and rinsed thoroughly with PBS. Control samples of differentiated and undifferentiated ASC on TCPS were formalin-fixed for 5 min, followed by extensive PBS rinsing. An oil red O stock solution (3 g/L) was prepared in isopropanol and was diluted 3:2 (v:v) with deionized water. This solution was agitated at room temperature for 10 min and then filtered through a type 1 qualitative filter paper. Each scaffold was incubated in 5 mL of the prepared stain for 2 h under agitation at room temperature. The TCPS control samples were stained for 5 min. All samples were rinsed thoroughly overnight in deionized water, with three solution changes to reduce non-specific staining. The scaffolds were analyzed using confocal microscopy (Zeiss LSM510, C-APO 63x/1.2 NA water immersion (DIC) objective lens, excitation with a HeNe laser at 543 nm and an Enterprise laser at 351 nm) and the control samples were visualized using light microscopy (Zeiss Axiovert 200 M).

3. Results 3.1. Macroscopic scaffold architecture The macroscopic architecture of each of the scaffold groups is shown in Fig. 2. All of the scaffolds were approximately 1.5 cm in diameter immediately following fabrication. The PDM alone scaffolds expanded during culture into a more random architecture. The XLHA alone constructs were difficult to handle and tore easily. The combined PDM with XLHA scaffolds were significantly stronger, could be easily handled with forceps and maintained their structure throughout the entire culture period. 3.2. Cellular organization Representative images of the cellular organization at 24 and 72 h post-seeding are shown in Fig. 3. On the TCPS, the cells had the expected fibroblastic morphology. On the PDM scaffolds, the cells adhered to the matrix and extended. Qualitatively, there were more adherent cells at 72 h, potentially due to cell proliferation. In the XLHA gels, the cells were rounded and formed self-aggregates. Live/dead staining revealed that the majority of these cells were alive at the 24-h time point, despite the rounded cellular morphology (Fig. 4). By the 72-h time point, there were more dead cells present, both individually and within the aggregates. In the PDM with XLHA Encapsulation samples, most of the cells could be found within the XLHA portion of the scaffold, with only a small percentage adhering to the PDM. In contrast, when the PDM with XLHA Adhesion seeding methodology was used, the

2.9. Oil red O staining Lipid accumulation within the scaffolds was visually assessed using oil red O staining at 7 and 14 days after the induction of differentiation. The PDM scaffolds were labeled prior to seeding with the Alexa Fluors 350 carboxylic acid, succinimidyl ester, as described in Section 2.4. At each time point, two scaffolds from each group were fixed for 24 h in 10%

Fig. 2. The macroscopic architecture of the PDM scaffolds, XLHA scaffolds and PDM combined with XLHA scaffolds. Bars represent 5 mm.

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Fig. 3. The cellular organization of the ASC in the scaffolds at 24 and 72 h after seeding, before the induction of differentiation. Prior to seeding, the cells were labeled with Cell TrackerTM Green and the PDM was labeled with an amine reactive Alexa Fluors 350 carboxylic acid, succinimidyl ester. Visualization was conducted using confocal microscopy (blue ¼ PDM, green ¼ cells). The PDM with XLHA Adhesion seeding methodology (PDM+XLHA Adh.) promoted greater adhesion of the ASC to the PDM scaffold than the PDM with XLHA Encapsulation methodology (PDM+XLHA Enc.). Bars represent 50 mm.

Fig. 4. LIVE/DEAD staining of the ASC in the XLHA gels at 24 and 72 h after seeding. Visualization using confocal microscopy (green ¼ live cells (calcein-AM), red ¼ dead cells (ethidium homodimer-1)). In general, more dead cells were present both individually and within the cellular aggregates at the 72-h time point. Bars represent 50 mm.

majority of the cells were located in close proximity to the PDM and many had an extended morphology. With both seeding methods, more adherent and extended cells could be visualized on the PDM at 72 h (Fig. 3). 3.3. GPDH activity The average measured GPDH activity levels are shown in Fig. 5. The values are expressed as the mean7the standard deviation. Statistical significance was determined using a one-way ANOVA with a Tukey’s post-hoc comparison of means (po0.05). Donor-specific activity levels of 20.4772.63, 30.6375.67, 34.6179.23 and 56.5376.13 mU/mg were measured in the mature adipose tissue samples, with a mean value of 33.86712.84 mU/mg. At 72 h after the induction of differentiation, the cells in the XLHA scaffolds (7.9073.85 mU/mg) and on the TCPS positive controls (8.2873.87 mU/mg) had statistically

higher activity levels than the other groups. While not statistically significant, the cells in the PDM (2.7871.04 mU/mg), the PDM with XLHA Encapsulation (3.8473.42 mU/mg) and the PDM with XLHA Adhesion (3.3472.04 mU/mg) scaffolds, all had higher mean activity levels than the non-induced TCPS negative control samples (0.8870.74 mU/mg). At 7 days after the induction of differentiation, increased activity levels, as compared to the 72-h samples, were observed in the XLHA, PDM with XLHA Encapsulation and TCPS positive control samples. The cells in the XLHA scaffolds (12.3576.80 mU/mg) and the TCPS positive controls (12.8975.19 mU/mg) had statistically higher activity levels than all of the other sample groups. The cells in the PDM with XLHA Encapsulation scaffolds (7.4773.38 mU/mg) had statistically higher activity levels than the cells in the PDM (2.5071.18 mU/mg), PDM with XLHA Adhesion (3.0671.35 mU/mg) and TCPS negative control (0.8470.54 mU/mg) samples. At 14 days after the induction of differentiation, the highest activity levels were observed in the cells from the XLHA scaffolds (23.07712.50 mU/mg), approaching the levels observed in mature adipose tissue. The XLHA and TCPS positive control (13.7478.00 mU/mg) samples had statistically higher activity levels than all of the other groups. While not statistically significant, the cells in the PDM with XLHA Encapsulation scaffolds (5.0572.49 mU/mg) had a higher mean activity level than the cells in the PDM (2.5371.98 mU/mg), PDM with XLHA Adhesion (3.0072.28 mU/mg) and TCPS negative control (0.7170.47 mU/mg) samples. 3.4. Oil red O staining Representative images of the Oil Red O stained scaffolds and controls are shown in Fig. 6. In the PDM scaffolds,

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Fig. 5. GPDH activity levels per milligram of total intracellular protein, quantified using a spectrophotometric assay (absorbance measured at 340 nm). The levels were measured in each of the scaffolds at 72 h, 7 and 14 days after the induction of differentiation. The data is presented as the mean7the standard deviation (n ¼ 3, N ¼ 6). The assay was also conducted on samples of freshly-excised abdominal adipose tissue (n ¼ 3, N ¼ 4). Statistical significance was determined at each time point by one-way ANOVA with Tukey’s post-hoc comparison of means (po0.05).

Fig. 6. Oil red O staining to detect intracellular lipid accumulation at 7 and 14 days after the induction of differentiation. Prior to seeding, the PDM was labeled with an amine reactive Alexa Fluors 350 carboxylic acid, succinimidyl ester. Visualization in the scaffolds was conducted using confocal microscopy (red ¼ lipid, blue ¼ PDM). Bars represent 10 mm. Visualization of the cells on the TCPS controls was conducted using light microscopy. Original magnification 200  . In general, at the 14-day time point, intracellular lipid accumulation was more readily detected and the cells had a more differentiated phenotype, with larger intracellular lipid droplets.

lipid accumulation could be visualized in only a very small number of individual cells after 7 days of differentiation. A greater number of cells containing small lipid droplets could be observed at 14 days. In the XLHA gels, intracellular lipid accumulation could be readily detected in both individual cells and within the aggregates at 7 days after the induction of differentiation. At 14 days, the cells had a more-differentiated phenotype, with larger lipid vesicles present within the cells. Lipid accumulation was also readily detected in the PDM with XLHA Encapsulation samples at 7 days. The cells could be visualized both

suspended within the XLHA and in close proximity to the PDM scaffold. The 14-day samples generally had a moredifferentiated phenotype. When the Adhesion methodology was used, fewer numbers of differentiating cells were visualized in the 7-day samples. All of the differentiating cells were attached to the PDM portion of the scaffold. At 14 days, a greater number of differentiating cells were generally present in the samples and some of the cells contained larger lipid droplets, characteristic of later differentiation. On the TCPS, lipid accumulation could be detected in large clusters of the cells at both 7 and 14

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days. In general, the cells had a more-differentiated phenotype at the 14-day time point. 4. Discussion Advances in obesity and metabolic research have helped to elucidate the cellular process of adipogenesis [32,33]. While this knowledge could benefit the field of adipose tissue engineering, most of these studies were conducted under two-dimensional (2D) growth conditions, with the cells grown in monolayer culture on TCPS. However, this environment does not accurately mimic the complex 3D network of the body [34]. Significant research remains to be conducted in order to understand the ideal scaffold environment for adipose tissue engineering applications. While a number of different synthetic and naturally derived materials have been studied both in vitro and in vivo, the optimal scaffold composition and architecture to promote the formation of stable adipose tissue remains unclear. Our work focuses on the development of 3D constructs incorporating scaffolds derived from the ECM. Cell adhesion to the ECM plays a critical role in the regulation of cellular behavior. Cell–matrix interactions through transmembrane receptors can induce signal transduction pathways that regulate a variety of cellular responses including survival, proliferation, differentiation, migration, and trafficking [35]. Naturally derived materials based on collagen and HA have been widely studied as scaffolds for tissue engineering applications [36]. Collagen gels support the attachment and proliferation of many types of cells, mediated by cellular integrins that bind collagen [37]. In contrast, the hydrophilic and polyanionic nature of HA does not facilitate cellular adhesion and protein adsorption [38]. Fibroblasts seeded in crosslinked HA–DTPH hydrogels were previously shown to survive and proliferate, but cellular adhesion and migration was not supported [30,39]. Similar to the results with the ASC, the fibroblasts were observed to have a rounded morphology and tended to form aggregates. A number of techniques have been developed to improve the cell adhesion to the HA–DTPH hydrogels, including the incorporation of crosslinked gelatin [40], fibronectin domains [41], and RGD peptides [42,43]. These strategies were shown to promote cellular attachment and spreading in a concentration dependent manner. The study of the ASC response in collagen-based PDM scaffolds and XLHA scaffolds has yielded interesting results. Due to the lack of an adhesive matrix, the cells seeded in the XLHA gels were rounded and tended to form self-aggregates. The live/dead staining results qualitatively indicated that there was a loss of cellular viability with time. In assaying the total intracellular protein content to normalize the GPDH expression levels, there was less protein present in the XLHA alone constructs, suggestive of a reduced cellular population (results not shown). The ASC could be observed to have an extended morphology in the scaffolds that contained the PDM. In the combined

PDM with XLHA scaffolds, it was possible to alter the cellular organization by modifying the seeding methodology. The PDM with XLHA Adhesion methodology promoted cellular adhesion to the PDM. In contrast, when the PDM with XLHA Encapsulation methodology was used, a large percentage of the seeded cells could be found within the XLHA portion of the construct. It is probable that the ASC were trapped within the XLHA and were unable to migrate unless they were in close proximity to the PDM at the time of gelation. In all of the constructs incorporating the PDM, the qualitative presence of more cells at the 72-h time point suggests that the PDM scaffolds may support the proliferation of the ASC. Similar to native adipose tissue, an engineered substitute should incorporate a mixture of mature adipocytes and undifferentiated progenitors to facilitate regeneration. The PDM scaffold may support the maintenance of this stem cell pool. We hypothesize that cellular adhesion may play an important role in the differentiation response of the ASC. This theory may be supported by the fact that mature adipocytes maintain their phenotype when cultured in suspension and have been shown to de-differentiate into a precursor state when cultured under cell-adhesive conditions [44]. In addition, a study conducted by Spiegelman and Ginty showed that adipogenic differentiation in the murine 3T3-F422A cell line was inhibited when the cells adhered and spread on fibronectin matrices [45]. The observed inhibitory effects could be reversed by disrupting the actin cytoskeleton and culturing the cells in a rounded state. In another study, McBeath et al. observed that cell shape influenced lineage commitment in human mesenchymal stem cells (hMSC) [46]. More specifically, adipogenic differentiation was observed in unspread hMSC that had a rounded phenotype (i.e. low adhesion conditions). In contrast, osteogenic differentiation was observed in culture conditions that promoted cellular attachment and spreading (i.e. high adhesion conditions). In the current work, the cells in the non-adhesive XLHA alone constructs had significantly higher GPDH activity levels, approaching the values observed for mature adipose tissue. The Oil Red O staining results support the augmentation of differentiation in the XLHA alone constructs. Cells containing intracellular lipid accumulation could be readily detected in the gels at both 7 days and 14 days. The cells in the 14-day samples had a more differentiated phenotype with larger intracellular lipid droplets, approaching the unilocular morphology of mature adipocytes. This augmentation in the differentiation response may involve the interaction of the ASC with the XLHA through the hyaluronan receptor CD44 [47]. However, our experimental findings with the two combined scaffolds suggest a role for adhesion in the modulation of the differentiation response. Higher GPDH activity levels (statistically significant at 7 days) were found when the PDM with XLHA encapsulation methodology was used. As previously discussed, the majority of the cells were suspended within the XLHA portion of the scaffold with

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this seeding strategy. The Oil Red O staining results also indicate that the encapsulation methodology promoted intracellular lipid accumulation and a more differentiated phenotype. These results clearly show that the seeding methodologies and the scaffold environment influence the ASC response. Moreover, promoting ASC adhesion may not be the best strategy for facilitating adipogenic differentiation. Overall, we believe that the PDM and XLHA scaffolds hold promise as scaffolds for adipose tissue engineering. Differentiating cells could be detected using Oil Red O staining in all of the scaffold conditions. In general, the cells seeded in the scaffolds had higher GPDH activity levels than the non-induced TCPS control samples. The large standard deviations in the activity levels can be attributed to the use of primary cells from a variety of donors. There are a number of advantages to including the PDM scaffold in the construct. The PDM significantly improves the surgical handlabililty and mechanical integrity of the scaffold. Additionally, the PDM contains a similar ECM composition to adipose tissue, including collagens type I, III, IV, V and VI [48,49]. Hence, the PDM may mimic the native adipose tissue environment and help to maintain the pool of adipogenic progenitors following implantation. The PDM may also help to promote construct vascularization following implantation, which will be critical to the success of the device. The vascular network of the placenta is preserved during the decellularization processing, and may facilitate angiogenesis by providing the appropriate 3D cues to stimulate proper cellular organization and infiltration. Vascularization could also be promoted by HA oligosaccharides produced during the degradation of the XLHA by hyaluronidases [41], and the acceleration of vascularization is possible through the addition of single or multiple growth factors within the XLHA materials [50,51]. We are continuing our investigation of the ASC response to the PDM and XLHA scaffolds in order to develop a more thorough understanding of the influence of the scaffold environment on the cellular behavior. We are currently conducting studies to quantify the cellular proliferation and metabolic activity in the different scaffolds. Our ongoing work also focuses on characterizing the changes in the ASC gene expression during adipogenic differentiation within the scaffolds using reverse transcriptase polymerase chain reaction (RT–PCR). 5. Conclusions Fabrication techniques and culture methods were developed to investigate primary human ASC in several naturally-derived scaffolds incorporating decellularized human placenta and XLHA. The results with the combined PDM with XLHA constructs indicate that cellular organization and adipogenic differentiation can be altered through variations in the cell seeding methodology. Cell adhesion may impact the ASC proliferation and differ-

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entiation responses. The PDM scaffolds facilitated cellular attachment and spreading. In contrast, adipogenic differentiation was augmented when the cells were encapsulated in the non-cell-adhesive XLHA gels. The results of our investigation clearly show that the scaffold environment greatly influences the organization and differentiation of the seeded ASC. Acknowledgments The authors would like to acknowledge Mary Boyle and the delivery suite staff at Women’s College Hospital for their assistance with the placental procurement. We thank Dr. X.Z. Shu for providing the HA–DTPH, and the Utah Centers of Excellence Program and the NIH (DC 004336) for financial support at Utah. This work was supported by funding from the Natural Sciences and Engineering Research Council (NSERC) of Canada, the Canadian Institutes of Health Research (CIHR) and the Province of Ontario through the Advanced Regenerative Tissue Engineering Centre (ARTEC). References [1] Katz AJ, Llull R, Hedrick MH, Futrell JW. Emerging approaches to the tissue engineering of fat. Clin Plast Surg 1999;26(4):587–603 viii. [2] Patrick Jr. CW. Tissue engineering strategies for adipose tissue repair. Anat Rec 2001;263(4):361–6. [3] The American Society of Plastic Surgeons. /www.plasticsurgery.orgS. [4] Shenaq SM, Yuksel E. New research in breast reconstruction: adipose tissue engineering. Clin Plast Surg 2002;29(1):111–25 vi. [5] Patrick Jr CW. Adipose tissue engineering: the future of breast and soft tissue reconstruction following tumor resection. Semin Surg Oncol 2000;19(3):302–11. [6] Patrick CW. Breast tissue engineering. Annu Rev Biomed Eng 2004;6:109–30. [7] Beahm EK, Walton RL, Patrick Jr CW. Progress in adipose tissue construct development. Clin Plast Surg 2003;30(4):547–58 viii. [8] Alhadlaq A, Tang M, Mao JJ. Engineered adipose tissue from human mesenchymal stem cells maintains predefined shape and dimension: implications in soft tissue augmentation and reconstruction. Tissue Eng 2005;11(3-4):556–66. [9] Fischbach C, Spruss T, Weiser B, Neubauer M, Becker C, Hacker M, et al. Generation of mature fat pads in vitro and in vivo utilizing 3-D long-term culture of 3T3-L1 preadipocytes. Exp Cell Res 2004;300(1):54–64. [10] Kang X, Xie Y, Kniss DA. Adipose tissue model using threedimensional cultivation of preadipocytes seeded onto fibrous polymer scaffolds. Tissue Eng 2005;11(3-4):458–68. [11] Patrick Jr CW, Zheng B, Johnston C, Reece GP. Long-term implantation of preadipocyte-seeded PLGA scaffolds. Tissue Eng 2002;8(2):283–93. [12] Kral JG, Crandall DL. Development of a human adipocyte synthetic polymer scaffold. Plast Reconstr Surg 1999;104(6):1732–8. [13] Gentleman E, Nauman EA, Livesay GA, Dee KC. Collagen composite biomaterials resist contraction while allowing development of adipocytic soft tissue in vitro. Tissue Eng 2006. [14] Halbleib M, Skurk T, de Luca C, von Heimburg D, Hauner H. Tissue engineering of white adipose tissue using hyaluronic acid-based scaffolds. I: in vitro differentiation of human adipocyte precursor cells on scaffolds. Biomaterials 2003;24(18):3125–32. [15] Kawaguchi N, Toriyama K, Nicodemou-Lena E, Inou K, Torii S, Kitagawa Y. De novo adipogenesis in mice at the site of injection of

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