Cellularized Microcarriers as Adhesive Building Blocks for Fabrication of Tubular Tissue Constructs
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- Twal, W.O., Klatt, S.C., Harikrishnan, K. et al. Ann Biomed Eng (2014) 42: 1470. doi:10.1007/s10439-013-0883-6
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To meet demands of vascular reconstruction, there is a need for prosthetic alternatives to natural blood vessels. Here we explored a new conduit fabrication approach. Macroporous, gelatin microcarriers laden with human umbilical vein endothelial cells and aortic smooth muscle cells were dispensed into tubular agarose molds and found to adhere to form living tubular tissues. The ability of cellularized microcarriers to adhere to one another involved cellular and extracellular matrix bridging that included the formation of epithelium-like cell layers lining the lumenal and ablumenal surfaces of the constructs and the deposition of collagen and elastin fibers. The tubular tissues behaved as elastic solids, with a uniaxial mechanical response that is qualitatively similar to that of native vascular tissues and consistent with their elastin and collagen composition. Linearized measures of the mechanical response of the fabricated tubular tissues at both low and high strains were observed to increase with duration of static culture, with no significant loss of stiffness following decellularization. The findings highlight the utility of cellularized macroporous gelatin microcarriers as self-adhering building blocks for the fabrication of living tubular structures.
KeywordsMacroporous microcarriersCultispherTissue engineeringIsotropicElasticBiomaterialElastinCollagenEndothelial cellsVascular smooth muscle cellsReplacement blood vessels
Limitations exist for the availability of suitable autologous vascular conduits derived from a patient’s body for vascular replacement procedures such as coronary artery bypass grafting.10 Therefore, there is a need for prosthetic alternatives to autologous vascular conduits. A variety of approaches have been developed to fabricate blood vessels.4,21,24,36,37,47 These include the use of tubular scaffolds manufactured from natural and synthetic biomaterials that are subsequently seeded with vascular cells to create living prostheses.6,16,37,44 We were motivated to explore alternative approaches that would facilitate cell-based fabrication of conduits comprised of vascular cells and extracellular matrix (ECM) constituents that they synthesize.
Microcarrier beads are 100–300 μm diameter spherical particles that allow attachment and growth of anchorage-dependent cells while in suspension culture.8,17,45 Microcarrier beads are manufactured from natural and synthetic materials, including gelatin, collagen, dextran, glass, polyethylene and polystyrene. Variant forms of microcarrier beads are macroporous, having large pores of tens of micrometers that provide additional areas for cells to attach and grow.35,38
Microcarriers have been generally used for suspension tissue culture to produce high yields of anchorage-dependent cells and their secreted products, but in recent years their utility in tissue regeneration and tissue engineering has emerged.31,32 For example, microcarriers have been used as cell delivery systems to regenerate tissue at sites of injury.32 Transplantation of skin cell-containing microcarriers onto cutaneous wounds of rodents and humans has been shown to lead to dermal regeneration15,22,29,46 and a reduction in detrimental wound contraction.12 Implantation of gelatin microcarriers loaded with bone marrow-derived mesenchymal stem cells has been shown to improve bone regeneration of craniofacial and long bone defects.28,48 An additional benefit of the gelatin microcarriers used in such applications is that they degrade over time in vivo without eliciting an inflammatory reaction.9,43
Only a few studies have explored the use of cellularized microcarriers as building blocks for three-dimensional (3D) tissue fabrication. Small disc-shaped constructs (1–2 cm in diameter × 0.1–0.8 cm in thickness) have been fabricated from dermal fibroblast-containing macroporous gelatin microcarriers.34,39 Similarly, cylindrical bone tissue constructs (2 cm in diameter × 1 cm in thickness) have been fabricated from macroporous microcarriers carrying human mesenchymal stem cells.7 In each of these studies, the cellularized microcarriers were placed into cylindrical perfusion culture chambers to facilitate cell-based joining of microcarriers into 1–2 cm-sized tissue constructs. Here we utilized vascular cell-containing macroporous gelatin microcarriers (Cultisphers) in conjunction with agarose molds to facilitate 3D tissue engineering of living tubular constructs and evaluated their histological and material properties.
Materials and Methods
Human umbilical vein endothelial cells (HUVECs, Lonza; Basel, Switzerland) were maintained in humidified 5% CO2, 95% air in Endothelial Growth Medium-2 (EGM-2; Lonza), containing 2% fetal bovine serum. Human aortic smooth muscle cells (HASMCs, Lonza) were maintained in humidified 5% CO2, 95% air in Smooth Muscle Growth Medium (SMGM; Lonza), containing 5% fetal bovine serum.
Cell Culture on Microcarriers
Gelatin CultiSpher-G cell carriers (Percell Biolytica, Astorp, Sweden), with an average particle diameter of 130–380 μm and pore size of 20 μm, were purchased from Sigma Chemical Co. (St. Louis, MO). Dry microcarriers were rehydrated, autoclaved and preincubated in cell culture medium according to manufacturer instructions. Microcarriers were then seeded with a 1:1 ratio of HUVECs:HASMCs (each at passage 4–5). Typically, a total of 8 × 106 cells were added to 50 mL of medium (1:1 mixture of EGM-2 and SMGM) containing 0.1 g microcarriers in a 125 mL siliconized Techne biological stirrer flask (R&D Systems, Minneapolis, MN). The cell-microcarrier suspension was subjected to an intermittent stirring regime (30 min at 0 rpm, 2 min at 50 rpm) on a Techne Biological Stirrer (model MCS-104S) for 24 h at 37 °C in a humidified 5% CO2 incubator. The volume of culture medium in the flask was then increased by addition of 50 mL of medium and the stirring speed switched to a continuous 50 rpm. Every 24 h, half of the medium volume was replaced with fresh medium.
Short Tube Formation in Agarose Molds
Stacked Tube Culture
After 5 days in agarose molds, tubular constructs were removed and stacked onto 2 mm diameter stainless steel center post guides mounted on an acrylic base plate in wells of 6-well plates. The stacked tubes were submerged in 1:1 mixture of EGM-2 and SMGM and cultured for varying periods of time.
Histological Staining and Immunohistochemistry
For histochemical staining, tubular constructs were fixed in 4% paraformaldehyde, PBS pH 7.4, embedded in OCT and subjected to frozen sectioning (10 μm thickness). Frozen sections were subjected to hematoxylin and eosin staining (H&E), Movat’s modified pentachrome and Masson’s trichrome staining according to standard procedures. Frozen sections of tubular constructs were also subjected to Picrosirius red staining and viewed with polarized light under dark-field optics to detect the birefringence of collagen fibers as described.5 For immunohistochemical staining, constructs were fixed, embedded in OCT and cryosectioned as above. Sections were then permeabilized in 0.02% Triton-X 100, PBS for 30 min at room temperature (RT) and then incubated with 1.0% BSA, PBS for 1 h at RT. Sections were incubated with primary antibodies i.e., rabbit anti-bovine tropoelastin (provided by Dr. Robert Mecham, Washington University, St. Louis, MO), mouse monoclonal anti-smooth muscle alpha actin (Sigma) or goat anti-VE cadherin (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) diluted in 1.0% BSA, PBS. After an 18 h incubation in a humidified chamber at 4 °C, secondary antibodies i.e., Alexa Fluor 488 conjugated donkey anti-goat IgG (Life Technologies, Carlsbad, CA) or Alexa Fluor 568 conjugated donkey anti-rabbit IgG (Life Technologies) were added. Nuclei were stained by incubation of sections with 2.5 μg/mL Hoechst 33258 (Molecular Probes, Life Technologies, Eugene, OR) in PBS for 15 min at RT. TUNEL assays were carried out as per the instructions provided with the ApopTag Fluorescein In Situ Apoptosis Detection kit by Chemicon (Temecula, CA). For en face immunohistochemical analysis, tubular constructs were opened by making a slice in the wall, along the longitudinal axis of the tube. The sliced tube was then mounted flat with the lumenal surface facing upward. The orientation of the whole mounted construct was preserved during antibody labeling and confocal imaging.
qPCR primer sequences
Target sequence ID no.
Forward primer 5′–3′
Reverse primer 5′–3′
A uniaxial ring test was used to probe the passive mechanical response of tube-shaped constructs. To initiate mechanical testing, tubular constructs were removed from agarose molds after 7, 12 and 17 days of culture and immediately secured onto horizontally-oriented 25 gauge cannulas mounted to the upper and lower arms of a uniaxial mechanical tester (Bose Enduratec 3200) (Fig. 6a). Samples were kept hydrated with culture medium while being mechanically preconditioned with three tensile displacement cycles up to 1.2 mm (20–25% strain) at a displacement rate of 0.01 mm/s. An identical fourth cycle was then immediately performed, during which load data (50 points/s) was recorded by system software (Wintest). An image-based technique was used to measure the local strain in the middle section of each sample. Immediately following sample mounting, blue tissue marking dye was applied by a fine tip applicator to create a dot pattern. A series of images was captured throughout testing using a Nikon SMZ-U light microscope and a Q-Imaging camera. Using ImagePro 5.1 to spatially calibrate the image, the vertical distances between the dots were calculated to facilitate measurement of local strain. Similar testing was performed on samples that had been decellularized using a hypotonic treatment with deionized water followed by a treatment with sodium dodecyl sulfate (SDS) in Dulbecco’s PBS as previously described.42
Cellularized Cultisphers Fuse to form Tubular Structures
Using a custom template (Fig. 1a), tubular agarose molds were made in 6-well tissue culture plates (Fig. 1b). After equilibration of the agarose molds with culture medium, cellularized Cultisphers were dispensed into the molds and cultured for varying periods. After 3–5 days in culture, the cellularized beads had joined together such that intact tubular structures (~4 mm diameter x ~ 2.5 mm long having a ~2 mm bore) could easily be removed from the molds (Figs. 1c, 1d). Low magnification microscopic examination of the tubular constructs revealed closely packed microcarrier beads and inter-bead material including evidence of material lining the lumenal and ablumenal surfaces of the tubes (Figs. 1e, 1f).
Stacked Tubes Fuse to form Elongated Tubes
Elastin and Collagen in Cultispher Tubular Constructs
Sections of the Cultispher tube constructs cultured for 7–17 days in agarose molds were also subjected to Picrosirius red staining and examined by polarized light microscopy. In 7-day constructs, the regions located between microcarrier beads were found to contain numerous fiber-like structures that show green/yellow birefringence (Figs. 4g, 4h, arrows), characteristic of newly deposited or less cross-linked collagen. Fibers having green/yellow birefringence could be found that were aligned circumferential to the lumen. Green/yellow fibers were also widely distributed throughout the day 12 and 17 constructs with relatively higher levels of green/yellow birefringence apparent in association with the epithelial layers (Figs. 4i–4l). Over time, the relative levels of red/orange birefringence, which is characteristic of mature cross-linked collagen, appeared to increase in association with the layers of cells lining the lumenal and ablumenal surfaces of the tubes (Fig. 4l). We also observed that over time the relative level of red/orange birefringence associated with construct-embedded Cultisphers diminished, concomitant with the progressive reduction in Cultispher size, perhaps reflective of a degradative process.
Immunohistological staining was performed to evaluate elastin deposition in the 7, 12 and 17-day constructs. Analysis of the constructs in cross section showed punctate deposits of elastin immunolabel in interbead regions and regions associated with the epithelial-like layers (Figs. 4m–4r). With time in culture, the relative intensity of anti-elastin immunolabeling appeared to increase. Confocal analysis of anti-elastin stained whole mounts, cut to permit en face examination of the lumenal surface, revealed fibrillar configurations, with elastin-containing fibers often oriented parallel to one another (Figs. 4s, 4t). Since control reactions showed that cell-free Cultisphers were weakly positive for elastin (i.e., diffuse staining, never fibrillar) (Fig. 4u), it is possible that the Cultispher manufacturing procedure might allow for dermal elastin to be a component in addition to porcine skin gelatin. For this reason, our interpretations of the immunohistological analysis of elastin were focused on extra-Cultispher regions of the constructs i.e., inter-Cultispher bead regions and the layers of cells lining the lumenal and ablumenal surfaces of the tubes. As evidence for the specificity of the anti-elastin immunolabeling, controls showed no reactivity by the fluorochrome conjugated secondary antibody in the absence of primary antibody, as well as positive detection of elastin in the ascending aorta (Figs. 4v–4x).
Tubular Constructs Exhibit Progressive Stiffening with Prolonged Culture Periods
Decellularized Tubular Constructs Exhibit Mechanical Behavior Similar to that of Living Constructs
Living constructs were treated with hypotonic and hypertonic rinses and SDS to obtain acellular ring constructs. The uniaxial mechanical response of decellularized 17 day constructs (inset panel of Fig. 6d) was nearly identical to that of living constructs cultured for the same amount of time (Fig. 6d). Two-way ANOVA analyses revealed that while culture time is a statistically significant factor in determining Eelastin and Ecollagen, cellularization state and the interaction between the two are insignificant factors. Moreover, a post hoc Tukey test with α = 0.05 indicates statistically insignificant differences in both moduli of the cellularized and decellularized 17 day samples.
Here we demonstrate that cellularized gelatin macroporous microcarriers (i.e., Cultisphers) can be used as self-adhering building blocks for the fabrication of tubular structures. The ability of cellularized Cultisphers to adhere to one another involved the formation of both cellular and ECM bridging between microcarrier beads. Cellular bridges included cells occupying the inter-bead spaces as well as epithelium-like cell layers lining the lumenal and ablumenal surfaces of the tubular constructs. The propensity of cellularized Cultisphers to form cellular bridges was apparent even when they were in suspension culture, as evidenced by the formation of aggregates interconnected by both VE-cadherin-positive and SM alpha actin-positive cells. While the present studies employed Cultisphers laden with endothelial cells and vascular smooth muscle cells, we also found that Cultisphers containing co-cultures of endothelial cells and the multipotent mouse cell line, C3H10T1/2, underwent inter-microcarrier bead cellular bridging (data not shown). By contrast, little to no inter-microcarrier bead cellular bridging was apparent when Cultisphers containing only endothelial cells (HUVECs) were used (data not shown). This observation is similar to other findings showing that co-culture of endothelial cells with retinal cells on microcarrier beads was required to stimulate endothelial cells to form endothelial cords that interconnect microcarrier beads.11
A potential application of the cellularized microcarrier-based fabrication approach described here is in the manufacture of replacement tubular tissue structures damaged by injury or disease. For example, each year hundreds of thousands of people undergo vein or artery replacement therapy13; however, systemic vascular disease often means that autologous replacement blood vessels are not available. Cellularized microcarrier-based fabrication is a cell-based approach to blood vessel construction that might permit synthesis of living prostheses having cellular composition and mechanical properties comparable to the walls of native blood vessels. The clinical applicability of microcarrier-based fabrication of living tubular constructs will require that the cellular components be derived from a patient’s own tissues. One source for autologous cells is adipose tissue. Indeed, vascular cells have been isolated from the vascular fraction of human adipose tissue19,27 and human adipose-derived stem cells have been used to produce endothelial cells and smooth muscle cells.1,41,51
In addition to fabrication and biomechanical testing of living microcarrier-based constructs, we also demonstrated that acellular ECM scaffolds could be generated from the living constructs using a standard decellularization protocol.42 Biomechanical testing of acellular ECM scaffolds showed that decellularization did not significantly alter the mechanical behavior of constructs, implying that ECM mediators of the mechanical response are not compromised by the decellularization process. The retention of mechanical properties increases the potential of these scaffolds to serve as clinical biomaterials (e.g., off the-shelf tissue-engineered vascular scaffolds).
Engineered tissues need to display strength and compliance necessary to sustain mechanical integrity in response to physiological mechanical forces. In tissues such as arterial blood vessels, collagens and elastin provide these properties.14,20,33 Analysis of the ECM composition of the tubes generated from Cultispher microcarriers containing vascular cells revealed the presence of collagen in inter-microcarrier bead regions and associated with the lumenal and ablumenal epithelium-like layers. Importantly, relative levels of collagen deposition in the tubes appeared to increase with duration of time in culture, which is consistent with the progressive stiffening measured by mechanical testing of 7, 12 and 17-day tubes. Elastin expression in the constructs was also demonstrated immunologically and using qPCR. Quantitative and qualitative deficiencies in the process of elastogenesis are impediments to the engineering of tissues such as blood vessels requiring functional elastin architecture.2 En face analysis of the anti-elastin labeled tubular constructs revealed extensive deposition of elastin into elongated fibers, often observed oriented in parallel configurations.
Biomechanical testing of the tubes generated from cellularized Cultisphers displayed mechanical behavior consistent with that of an isotropic, incompressible, homogeneous, elastic material. A modulus of 104.5 ± 60.4 kPa (Eelastin) was calculated from the elastin-dominated portion of the mechanical response of 17-day constructs and a modulus of 1278.1 ± 329.3 kPa was calculated from the collagen-dominated portion of the mechanical response. By comparison, incremental Young’s moduli of the intact wall (both the tunica adventitia and tunica media layers) of human carotid arteries have been reported to be equal to 160 ± 40 kPa and 900 ± 250 kPa for the low strain region and the high strain region, respectively.18 Isolated insoluble elastin and elastin-rich tissues (e.g., bovine nuchal ligament) have been shown to behave as a nearly linear elastic material with Young’s modulus of roughly 400–800 kPa, depending on the tissue source and isolation procedures.18,25,26,40 Biomaterials produced by cross-linking recombinant elastin polypeptides show similar behavior with a lower modulus of about 250 kPa.3 More recent studies50 suggest that aortic elastin shows some anisotropy in its material behavior, with axial stiffness being less than circumferential stiffness. The Eelastin modulus of the Cultispher tube constructs is consistent with the lower end of the range reported by Zou and Zhang50 for the axial tangent modulus of isolated aortic elastin at low strains.
The relatively low Eelastin of the Cultispher tube constructs may be an indication that the elastin produced in the constructs in not highly cross-linked. Previous work by the Wang group demonstrated production of mature elastin in baboon VSMC-seeded tubular scaffolds fabricated from porous poly(glycerol sebacate) and cultured in a pulsatile flow bioreactor for 3 weeks.23 Interestingly, these authors reported a significant effect of scaffold pore size on elastin production and mechanical properties; with constructs having the smallest pores (25–32 μm) attaining the highest values of both elastic modulus (~60 kPa) and ultimate tensile strength.23 In the current work, we achieved comparable elastic modulus values after 17 days in static culture, without any mechanical pre-conditioning. The macroporous gelatin microcarriers used in our study have a pore size at the low end of the range examined by Lee et al.23 Therefore, we might expect that mechanical pre-conditioning of Cultispher-based constructs (e.g., application of cyclical strain or flow) would lead to further improvement in their mechanical properties.
This work was supported by the National Science Foundation/EPSCoR Grant (EPS-0903795) and by NSF CMMI-1200358. We thank Dr. Amy Bradshaw for providing expert advice on Picrosirius red staining and polarized light microscopy. We thank Michael Gore (University of South Carolina School of Medicine) for his fabrication of templates.
Conflict of interest
No competing financial interests exist.