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Indong Jun, Taufiq Ahmad, Seongwoo Bak, Joong-Yup Lee, Eun Mi Kim, Jinkyu Lee, Yu Bin Lee, Hongsoo Jeong, Hojeong Jeon, and Heungsoo Shin* since it can provide a myriad of signals to cells due to its physiologically relevant concentration level similar to that of native tissue.[1,2] Thus, the distinct benefits of coculture have been reported to modu late cellular proliferation, differentiation, extracellular matrix (ECM) production, and protein secretion.[3–6] For example, Yamamoto et al. reported that coculture of stem cells and nucleus pulposus cells induced the secretion of growth fac tors associated with cellular metabolism. Another study found that a culture of stem cells combined with endothelial cells had highly angiogenic and vasculogenic capabilities.[6,7] However, conventional coculture systems often involve mixed cultivation of various cell types, and it remains difficult to recapitulate spatially controlled cell–cell and cell–ECM inter actions similar to structural characteristics in certain tissues.[4,5,8] In addition, in vivo transplantation of cocultured cells while preserving cell–cell interactions, cell–ECM interactions, and hierarchical structure has proven extremely challenging. Over the last decade, a great deal of attention has been paid to cell sheet technology due to its ability to deliver cells as a monolayer while preserving cell–cell contact and cell– ECM communications for improved therapeutic outcomes.[9] Poly(N-isopropylacrylamide) (PIPAAm)-grafted surfaces have been mainly employed for harvesting cell sheets by exploiting the thermo-rheological properties of PIPAAm; these polymer chains are aggregated, allowing cell attachment at 37 °C while being extended and hydrated below the lower critical solution temperature (LCST) of PIPAAm (≈32 °C); thus, the cell sheet is detached.[10] However, this technique involves the use of cell shifter membranes or gelatin plungers to form a multilayered cell sheet through a stacking process, which requires multiple steps for the translocation of the engineered cell sheet to the target.[10,11] Conversely, we have previously reported on a ther mosensitive hydrogel system with cell-instructive cues that can be expanded in response to the temperature change required for transfer stamping of cell sheets in a single-step proce dure.[12–14] With our system, a confluent monolayer of cells can be harvested on the hydrogels and then rapidly transferred
Although the coculture of multiple cell types has been widely employed in regenerative medicine, in vivo transplantation of cocultured cells while maintaining the hierarchical structure remains challenging. Here, a spatially assembled bilayer cell sheet of human mesenchymal stem cells and human umbilical vein endothelial cells on a thermally expandable hydrogel containing fibronectin is prepared and its effect on in vitro proangiogenic functions and in vivo ischemic injury is investigated. The expansion of hydrogels in response to a temperature change from 37 to 4 °C allows rapid harvest and delivery of the bilayer cell sheet to two different targets (an in vitro model glass surface and in vivo tissue). The in vitro study confirms that the bilayer sheet significantly increases proangiogenic functions such as the release of nitric oxide and expression of vascular endothelial cell genes. In addition, transplantation of the cell sheet from the hydrogels into a hindlimb ischemia mice model demonstrates significant retardation of necrosis particularly in the group transplated with the bilayer sheet. Collectively, the bilayer cell sheet is readily transferrable from the thermally expandable hydrogel and represents an alternative approach for recovery from ischemic injury, potentially via improved cell–cell communication.
1. Introduction The coculture of multiple cell types has been widely employed in cancer biology, organoid culture, and regenerative medicine Dr. I. Jun, T. Ahmad, S. Bak, J.-Y. Lee, E. M. Kim, J. Lee, Y. B. Lee, Prof. H. Shin Department of Bioengineering Institute for Bioengineering and Biopharmaceutical Research Hanyang University Seoul 04763, Republic of Korea E-mail:
[email protected] Dr. I. Jun, H. Jeong, Dr. H. Jeon Center for Biomaterials Biomedical Research Institute Korea Institute of Science and Technology Seoul 02792, Republic of Korea T. Ahmad, S. Bak, J.-Y. Lee, E. M. Kim, J. Lee, Y. B. Lee, Prof. H. Shin BK21 Plus Future Biopharmaceutical Human Resources Training and Research Team Hanyang University Seoul 04763, Republic of Korea
DOI: 10.1002/adhm.201601340
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Spatially Assembled Bilayer Cell Sheets of Stem Cells and Endothelial Cells Using Thermosensitive Hydrogels for Therapeutic Angiogenesis
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within 10–15 min directly to the target substrate by reducing the temperature from 37 to 4 °C. Cell-based therapies have drawn significant attention for therapeutic angiogenesis. Transplanted cells not only form the backbone of new blood vessels, but stimulate secretion of var ious proangiogenic cytokines, which enhance vascularization through recruitment of endothelial progenitor cells (EPCs) to the ischemic site.[15,16] Various reports have revealed that the administration of endothelial cells (ECs) and EPCs improved blood perfusion recovery, resulting in reduced limb necrosis and amputation. However, blood vessels regenerated solely from the administration of ECs are unstable and demonstrated regression, implying that the interaction of transplanted ECs with primary pericytes or vascular smooth muscle cells is vital to the regeneration of a stable and mature vasculature.[16,17] Similarly, coculture of mesenchymal stem cells (MSCs) with ECs induced the expression of proangiogenic markers related to vascular maturation, and in vivo transplantation led to enhanced therapeutic angiogenesis.[17–19] However, the thera peutic effect of a spatially assembled bilayer of MSCs with ECs on ischemic tissue has not yet been fully achieved. Given these findings and current issues, the objectives of our study were as follows: (1) to fabricate a spatially assembled bilayer cell sheet with human MSCs and ECs on the previously developed thermally expandable and cell-interactive hydrogel,
(2) to confirm the harvest and transferability of the cell sheet while maintaining spatial assembly, (3) to investigate the effect of the bilayer cell sheet on in vitro proangiogenic functions, and (4) to assess the effect of the bilayer cell sheet on the recovery of ischemic tissue using an in vivo athymic mice model.
2. Results and Discussion Studies have found that stem cells have a close functional asso ciation with pericytes and endothelial cells involved in angio genesis.[20,21] Therefore, several studies have been conducted on the cell-based therapeutic effect of stem cells or endothelial cells on angiogenesis.[22,23] We hypothesized that a cocultured system composed of stem cells and endothelial cells would improve proangiogenic function and may have positive effects on therapeutic angiogenesis. In order to deliver the cell sheet, we utilized a thermally expandable hydrogel incorporating cell adhesive domains, which was reported in our previous study.[13] The cell-interactive molecule fibronectin (FN) was covalently conjugated with the synthesized thermosensitive polymer based on Tetronic during hydrogel formation. As illustrated in Figure 1, the hydrogel incorporating FN (FN-hydrogel) can sup port the cell sheet formation and rapidly expand in response to a temperature change from 37 to 4 °C within a short time. As
Figure 1. A) Schematic illustration of the spatially assembled bilayer of hMSCs and HUVECs on the thermally expandable, cell-interactive hydrogel, and harvest and delivery of the bilayers to a target in response to the reduced temperature. hMSCs were first cultured on the surface of the FN-hydrogel, and then HUVECs were further cultured on the hMSC layer to form the bilayer. To transfer the cell sheets onto a target substrate, we changed the temperature from 37 to 4 °C, which rapidly induced the expansion of the hydrogel and subsequently disrupted the interaction between the cell sheet and hydrogel while maintaining cell–cell junctions with the ECM structure. B) Schematic diagram of the transplantation of the cell sheet from the hydrogel to mice with hindlimb ischemia injury. The precultured bilayer on the FN-hydrogel was placed over the defective tissue (after injury to the femoral artery). Cold sterilized saline (4 °C) was applied on the top of the hydrogel for 15 min, allowing the cell sheet to detach from the hydrogel and reattach to the ischemic target tissue. After 15 min, the hydrogel was carefully detached from the tissue, which successfully delivered the cell sheet to the injured tissue.
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2.1. Characterization of Hydrogels Containing Fibronectin The fluorescent image of the FN-hydrogel exhibits the pres ence of FN within the hydrogel (Figure 2A). The chemical structure and reaction mechanism between FN and the syn thesized Tetronic-tyramine polymer precursors are shown in the Figure S2 (Supporting Information). The viscoelastic properties of the hydrogels showed that the elastic modulus (storage modulus, G′) was 2.15 and 1.43 kPa for the FN-0 and FN-50 hydrogels, respectively. Previously, we reported a decrease in the surface stiffness as the FN content increased in FN-hydrogels using atomic force microscopy (AFM).[14] Consistent with these results, the elastic modulus of the FNhydrogel itself slightly decreased with a FN concentration of 50 µg mL−1. X-ray photoelectron spectroscopy (XPS) was used to investigate the surface chemical composition of the lyophi lized FN hydrogels. As shown in Figure 2C,D, the lyophilized FN-hydrogels presented a similar intensity for the carbon (C1s) and oxygen (O1s) peaks, while there was a considerable
difference in the high-resolution nitrogen peak (N1s, 399.8 eV) in FN-hydrogels compared with pristine hydrogels (FN-0).
2.2. Cell Adhesion on FN-hydrogels The human mesenchymal stem cells (hMSCs) demonstrated adequate spreading and adhesion on FN-50 hydrogels, while their adhesion on FN-0 hydrogels was diminished, and the cells displayed an aggregated morphology (Figure 3A). Likewise, human umbilical vein endothelial cells (HUVECs) showed sim ilar adhesion and spreading on the FN-50 hydrogel. Our results suggest that both hMSCs and HUVECs were well-attached to the hydrogel containing FN, while HUVECs appeared to require less FN for adhesion compared to hMSCs (Figures S3 and S4, Supporting Information). We then examined bilayer forma tion on FN-hydrogels by using a relatively high seeding density to achieve confluency quickly in each cell layer. As shown in Figure 3C,D, hMSCs and HUVECs maintained their unique morphology on the FN-hydrogels. However, we observed a change in morphology after seeding HUVECs on the layer of hMSCs. During the prolonged incubation period of 12–36 h, cell attachment was enhanced due to an increased number of cell– cell interactions. Additionally, 12 h after seeding the HUVECs, the number of cells in the bilayer significantly increased from 1.51 × 103 cells cm−2 (only hMSCs) to 7.72 × 103 cells cm−2 for
Figure 2. Characterization of surface and rheological properties of the hydrogels. A) Representative fluorescent images of the hydrogels reacted with defined concentrations of FITC-tagged FN (0 and 50 µg mL−1). B) Rheological properties of the hydrogels prepared with different fibronectin concentrations comparing the storage modulus (G′). C) Survey XPS spectra and D) high-resolution N1s spectra of the lyophilized samples of the FN-hydrogels. The scale bars indicate 50 µm.
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a result, binding between the cells and surface of the hydrogel, which is mediated by the adhesion of FN from the hydrogel with integrin receptors in cells, is disrupted, while the cell–cell junctions with the ECM structure are maintained in the cell sheet and harvested or transferred to the target site.
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Figure 3. The cytoskeletal structure and morphology of A) hMSCs and B) HUVECs on FN-50 hydrogels cultured for 24 h. Representative phase contrast images of C) hMSCs on HUVECs and D) HUVECs on hMSCs on FN-50 hydrogels. E) Quantification of adhesive cells cultured as a bilayer on the FN-50 hydrogels for 96 h. Proliferation of cells on the FN-50 hydrogels under conditioned media. Proliferation was evaluated using the WST-1 assay. The scale bars indicate 25 µm. * indicates significant differences compared to only hMSCs cultured on the FN-50 hydrogel (p < 0.05).
HUVECs cultured on the hMSC monolayer on FN-hydrogels. HUVECs seeding on an hMSCs monolayer of FN-50 Hydro gels showed significant increase in the cell number indicating successful formation of bilayer structure. However, there was a little change in cell number after bilayer formation, which may be due to limited space available for further proliferation of HUVECs on hMSCs. Consistent with these results, the absorb ance from hMSCs cultured on the FN-50 hydrogels (1.01 ± 0.32) was significantly increased (3.86 ± 0.58) after HUVECs seeding onto hMSCs on FN-50 hydrogels. These results suggest that FN-hydrogels supported the adhesion of hMSCs and HUVECs as well as the spatially assembled cell layer. Although both cell types readily attached to the surface of the hydrogel, hMSCs cul tured on a HUVEC monolayer did not form a bilayer. Previous studies have corroborated the poor attachment of MSCs on the surface of precultured ECs and highlighted the involvement of cell adhesion molecules in the binding of HUVECs to hMSCs, which was enhanced after activation of ECs with tumor necrosis factor-α (TNF-α).[24,25] TNF-α-treated ECs showed elevated expression of vascular cell adhesion molecule-1 (VCAM-1), a molecule primarily involved in the interaction and adhesion of MSCs with ECs.[25] Our results suggest that ECs may not serve as a feeder layer to support the stable adhesion of MSCs without activation. Alternatively, the culture of HUVECs on the 1601340 (4 of 12)
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hMSC layer was well maintained, and a significant increase in cell number was found 12 h after bilayer fabrication. hMSCs can express multiple receptors for cell–cell interactions and secrete ECM biomolecules including collagen type I, collagen type IV, fibronectin, and laminin, which may support the stable adhesion of HUVECs onto hMSCs.[26]
2.3. Thermal Expansion of the Hydrogels As shown in Figure 4A, high-resolution particle image veloci metry (PIV) for visualizing the thermo-stimuli pattern was performed by tracing the diffusive spread of microscale fluo rescent particles. The fluorescence intensity profile of the bead distribution in FN-50 hydrogels and the scatter diagram of hori zontal velocity vectors were produced at 60 s intervals after the change in temperature from 37 to 4 °C. We demonstrated that the movement of the fluorescent beads within the hydrogels exhibited relatively high velocity until 180 s, while the velocity decreased between 240 and 420 s. After 480 s, all fluorescent particles embedded in the hydrogels were in a steady state. The results from the PIV analysis were consistent with the thermoexpandable change in the size of hydrogels. When the tem perature was reduced from 37 to 4 °C, the relative size of the
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FULL PAPER Figure 4. Characterization of the thermal expansion of FN-50 hydrogels. A) Color maps of fluorescent particles embedded within the hydrogels, visualized with confocal laser scanning microscopy for 60 s intervals after decreasing the temperature from 37 to 4 °C. The color scale is shown in the left column (unit: µm s−1). B) Reversible changes in the size and C) kinetic profiles of the expandable FN-50 hydrogels in PBS in response to the temperature change from 37 to 4 °C. Scale bar indicates 100 µm.
hydrogels increased by 1.33 ± 0.06. We also confirmed that the hydrogel returned to its original size when the temperature was increased from 4 to 37 °C as shown in Figure 4B. We confirmed that the thermally expanded hydrogels reached a plateau after 9 min (540 s). Tetronic-based hydrogels possess polymer pre cursors with four polymeric arms composed of poly(ethylene oxide, PEO) and poly(propylene oxide, PPO). At 37 °C, the PPO chain can be arranged to decrease the size of the hydro gels because of its strong hydrophobic interactions. In con trast, the hydration between PEO domains becomes dominant at 4 °C, which increases the size of the hydrogel. Moreover, it has been confirmed that the thermosensitive characteristics can be changed (increased by 1.16–1.33) depending on the final polymer concentration and thickness of hydrogels, as reported in the previous study.[13,14] The transfer of the cell sheet from the hydrogel is delicately regulated by interactions between FN and the cells. The strong binding of cells on the FN-hydrogel is required for stable formation of the cell sheet, which should be rapidly detached by the expansion of the hydrogel. If the expan sion rate is too high, the cell sheet can be disrupted.
2.4. Characterization of the Bilayer Cell Sheet The in vitro detachment of cell sheets from FN-hydrogels and their subsequent transfer to glass substrates was performed at 4 °C for 15 min. As shown in Figure 5A, some dead cells were observed in the HUVEC cell sheets, while predominantly live cell signals were seen in the hMSC and bilayer cell sheets (the survival rate (%) was 99.53 ± 0.05%, 98.01 ± 0.09%, and 98.04 ± 0.21% for transferred hMSC, HUVEC, and bilayer cell sheet, respectively). We immunostained the transferred cell sheets to confirm maintenance of the ECM structure, in which FN assembly was clearly demonstrated on monolayer and bilayer sheets (Figure 5B). In contrast, as shown in Figure 5C, CD31 (platelet endothelial cell adhesion molecule (PECAM)) was not expressed in transferred hMSC cell sheets, while HUVEC cell sheets displayed thoroughly dispersed CD31 signals. The coexistence of these signals was observed in the bilayer, and
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CD31 expression in the bilayers was similar to its expression in HUVECs. These results suggest that the transfer process under reduced temperature did not have any detrimental effect on cell viability with an apparent enhancement in cell viability in the bilayer cell sheet. This improvement in viability of HUVECs in the bilayer may be due to their interaction with hMSCs in the coculture.[1] More importantly, the bilayer cell sheet showed highly developed ECM components, which play critical roles in cell adhesion, migration, and survival, suggesting that bilayer cell sheets maintained their ECM after transfer.[5,27] We also confirmed the distribution of heterogenic cell layers in the bilayer cell sheet by prelabeling hMSCs with the green fluorescent dye DiO and HUVECs with the red fluorescent dye DiD. As shown in Figure 5D, each cell type was uniformly dis tributed in two distinct dye layers where HUVECs (appearing at a depth of 6 µm) were located below hMSCs (appearing at a depth of 9 µm) after transferring to the target. A projected view of the bilayer cell sheet prepared from the fluorescently labeled hMSCs and HUVECs clearly demonstrated two distinct adherent layers of cells (Figure 5E). In addition, as shown in Figure 5F, cocultured bilayer cell sheets exhibited the coex istence of two different types of cell-specific markers, in par ticular CD31, implying the formation of spatially segregated layers, which was further confirmed using confocal laser scan ning microscopy (CLSM). CLSM revealed CD31 signals up to a depth of 6 µm, while the CD31 signal disappeared completely at 14 µm, indicating the presence of the hMSC layer on top of the HUVEC layer, which was converted by the transfer process. Collectively, these results suggest that bilayer cell sheets pro vide improved cell–cell interaction in a 3D environment, which appears to be consistent with previous reports.[5,28]
2.5. Evaluation of In Vitro Angiogenic Functions of Cell Sheets To investigate the effect of cocultured bilayer cell sheets on the expression of proangiogenic growth factors, the superna tant media were collected from each group (hMSCs, HUVECs, and the bilayer cell sheet) and added to HUVECs cultured
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Figure 5. Representative fluorescent images of the cell sheets transferred from the FN-50 hydrogels to the target substrates. A) Live/Dead staining of the cell sheets and immunofluorescent staining for B) anti-fibronectin and C) anti-PECAM. D) Orthogonal images of the bilayer at depths of 6 and 9 µm and E) cross-sectional view of the transferred bilayer cell sheet. F) To visualize the cell–cell junctions between the cell layers, immunofluorescent staining for PECAM (green) was carried out and analyzed using CLSM at depth intervals of 4 µm. The scale bars represent 500 µm in (A) and 100 µm in (B)–(F).
on a GF-reduced Matrigel for 12 h. The conditioned media from the bilayer culture resulted in a tubular network that was thoroughly spread and interconnected with a significant increase in length. Although tubules in the hMSC group were not entirely dispersed, the tubule length in the group treated with media from the hMSC was significantly increased com pared to that from the HUVEC group. As shown in Figure 6A, the total tubule length was 8228 ± 1584, 5551 ± 791, and 12 320 ± 3451 µm for media from hMSCs, HUVECs, and the bilayer, respectively. In addition, VEGF secretion was sig nificantly higher in the bilayer (1.25 ± 0.31 pg mL−1 µg−1 pro tein) compared to hMSCs (0.74 ± 0.10 pg mL−1 µg−1 protein) and HUVECs (0.22 ± 0.09 pg mL−1 µg−1 protein) (Figure 6B). Similarly, consistent with the tubule formation assay results, 1601340 (6 of 12)
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VEGF secretion was significantly enhanced in the hMSC group compared to the HUVEC group. Nitric oxide (NO) release from the cell sheets was similar in the three groups: hMSC sheets (1.26 ± 0.16 × 10−6 m cm−2), HUVEC sheets (1.12 ± 0.23 × 10−6 m cm−2), and bilayer cell sheets (1.38 ± 0.16 × 10−6 m cm−2) at 12 h (Figure 6C). However, NO release from the bilayer cell sheets was significantly higher (5.52 ± 0.34 × 10−6 m cm−2) compared to that from hMSC sheets (3.17 ± 0.17 × 10−6 m cm−2) and HUVEC sheets (3.05 ± 0.17 × 10−6 m cm−2) at 48 h. NO has been widely investigated and has important functions in vascular systems, especially in main taining endothelial homeostasis and inhibiting platelet adhe sion and smooth muscle cell hyperplasia while encouraging endothelial cell growth.[29] Moreover, decreased NO production
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FULL PAPER Figure 6. A) Representative images of tubules formed from HUVECs cultured on the GFR-Matrigel exposed to the collected media, and the total tube length in each group. B) The amount of VEGF secreted from each cell sheet group. C) NO production from the transferred cell sheets was measured by the Griess reagent kit for 72 h. The relative expression level of proangiogenic genes from cell sheets cultured for D) 3 and E) 5 d. Quantification analysis of VE-cadherin, vWF, and Tie-2 using real-time RT-PCR. *, #, and $ indicate significant differences compared to the transferred hMSC and HUVEC cell sheet, respectively (p < 0.05). The scale bar indicates 200 µm.
is associated with endothelial dysfunction.[30] Analysis of gene expression in the cell sheets showed interesting results. The HUVEC cell sheet exhibited similar or higher expression of various types of proangiogenic genes (VE-cadherin, vWF, and Tie-2) compared to hMSCs and the bilayer cell sheet at 3 d, as shown in Figure 6D. However, after 5 d, the bilayer cell sheets showed significantly higher levels of expression of proangio genic genes (Figure 6E). Several reports have shown that cocul tured MSCs and ECs have upregulated expression of proangi ogenic cytokines and junction proteins (such as connexin 43) and enhanced in vitro tubule formation and in vivo neovascu larization compared to monocultures.[17,19,31] The upregulation and significantly increased expression of proangiogenic genes found in our results is consistent with these previous studies.
2.6. Effect of Cell Sheet Transplantation on a Mouse Hindlimb Model of Ischemia We evaluated the therapeutic effect of cell sheets in an athymic mouse model of hindlimb ischemia. The no treatment group showed rapid necrosis of ischemic hindlimb tissue after 3 d, resulting in complete limb loss after 21 d (Figure 7A,B). Com paratively, the transferred cell sheet groups demonstrated a diminished rate of tissue necrosis. Additionally, we observed an apparent difference between the different cell sheet groups. For example, the hMSC cell sheet group exhibited 37.5%, 12.5%,
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and 50% limb salvage, foot necrosis, and limb loss, respec tively, at 21 d, while the HUVEC cell sheet group exhibited 33.3%, 16.6%, and 50% limb salvage, foot necrosis, and limb loss, respectively, at 21 d. In contrast, the transferred bilayer cell sheet group had 50% limb salvage, 25% foot necrosis, and 25% limb loss (Figure 7C,D). Consistent with physiological status, hematoxylin and eosin (H&E) and Masson’s trichrome (MT) staining showed consider able numbers of granulocytes in the muscle tissue and fibrosis in the no treatment group (Figure 8A,B). The bilayer group exhibited diminished inflammation and fibrosis compared to the other groups. We observed fibrotic areas of 8.01 ± 2.58, 4.05 ± 1.67, 4.67 ± 1.50, and 2.44 ± 0.85 (all values expressed as percentages) for the no treatment, hMSC, HUVEC, and bilayer groups, respectively. The number of arterioles was sig nificantly increased in the bilayer group (21.76 ± 7.01 mm−2) compared to the no treatment (6.06 ± 4.06 mm−2), hMSC (14.89 ± 5.73 mm−2), and HUVEC (11.78 ± 5.49 mm−2) groups, as shown in Figure 8E. For therapeutic angiogenesis, most approaches have used cells directly or systemically delivered to a target tissue after enzymatic treatment, often resulting in low cell engraftment efficiency, inhomogeneous distribu tion of cells, and elevated cell doses for a therapeutic effect (≈107 cells).[32] Our bilayer system accumulated relatively low cell numbers (about 6 × 104 cells for the bilayer cell sheet), but significantly enhanced angiogenesis of ischemic tissue. HUVECs and hMSCs have also been shown to be a therapeutic
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Figure 7. Representative photographs of mice treated with saline (no treatment) or transplanted with hMSCs, HUVECs, and the bilayer at A) day 3 and B) day 21 after surgery. Physical status of the ischemic hindlimb at C) day 3 and D) day 21 after the surgical operation, which was divided into three levels of limb salvage, foot necrosis, and limb loss.
cell source in the treatment of peripheral arterial disease (PAD).[5,12,33] It is widely accepted that MSCs possess immunemodulatory nature and site-specific affinity.[33] In this study, our primary goal was to investigate the effect of bilayer delivered from our thermosensitive hydrogels on angiogenic effect, and thus, we used nude mice. Although numerous studies using stem cell have been known to use immunocompetent mice, considerable number of researches still use nude mice for eval uation of therapeutic and functional efficiency.[22,34] It should also be noted that HUVECs used in our study may be valid as a physiological model, but may not be used in the clinical setting and endothelial progenitors derived from human peripheral blood mononuclear cells (hPBMNCs) or umbilical cord bloodmononuclear cells (UCB-MNCs) may represent a potential therapeutic source. Therefore, modification of our experimental protocols including cell sources may be required for translation of our approach into practical therapeutic angiogenesis treating patients with ischemic diseases. Our studies showed a diminished therapeutic effect with both the hMSC-only cell sheet and HUVEC-only cell sheet compared to the cocultured bilayer cell sheet group, suggesting that transplantation of a single cell type is not sufficient for therapeutic angiogenesis. Previous studies have reported that the transplantation of only endothelial cells often gives rise to an unstable vasculature that can undergo rapid degenera tion. Interaction of ECs with pericytes or SMCs is critical for the regeneration of a stable vasculature.[16,17] Similarly, trans planted MSCs have a limited role in direct neovascularization as fewer hMSCs can differentiate into ECs or VSMCs after 1601340 (8 of 12)
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transplantation.[35] Endogenous angiogenesis begins with the migration, proliferation, and differentiation of ECs, which ulti mately gives rise to the new vasculature.[18] However, deposi tion of ECM, recruitment of mural cells, and the presence of pericytes are essential for stable vascularization.[18,21] In our bilayer cell sheet system, HUVECs may have been preacti vated with hMSCs, which are functional homologues of peri cytes. In support of this hypothesis, it has been reported that MSCs functionally resemble pericytes and can differentiate into SMCs.[20,21] Similarly, the coculture of MSCs with ECs induced a quiescent EC phenotype and promoted the expres sion of biomarkers related to vascular maturation.[31] Therefore, the bilayer cell sheets not only provide the benefits of robust vascularization by HUVECs, but also confer stabilization of the vasculature by hMSCs. Secretion of various cytokines from hMSCs, including VEGF, may also have enhanced indigenous vascularization. Taken together, these results indicate that our transfer stamping approach might overcome the obstacles of the current cell therapy strategy including low cell engraft efficiency, inhomogeneous distribution of cells, and high cell dosage requirement.
3. Conclusions Our study is based on the development of a transferable, bilayer cell sheet with cocultured hMSCs and HUVECs. We prepared hydrogels with FN to promote cell adhesion, fabri cated the bilayer cell sheets, and transferred the cell sheets to
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FULL PAPER Figure 8. Histological analysis of the transplanted cell sheets in the recovery of a severely ischemic hindlimb. A) H&E and B) Masson’s trichrome (MT) staining of a histological ischemic hindlimb section 21 d after the surgical operation treated with saline (no treatment), a transplanted hMSCs layer, a transplanted HUVECs layer, or an hMSC/HUVEC bilayer. The blue stain for collagen indicates fibrosis by MT staining. C) Immunofluorescent staining of smooth muscle α-actin in ischemic hindlimb tissue 21 d after surgery. D) Quantification of the fibrosis area (fibrosis area (blue)/muscle area (red) × 100) and E) quantitative analysis of the number of stained arterioles within the field (1 mm2). The scale bars represent 200 µm for (A) and (B) and 100 µm for (C). *, #, and ψ represent significant differences in comparison with no treatment, hMSCs, and HUVECs, respectively (p < 0.05).
a target substrate by changing the temperature. Specifically, our cocultured bilayer cell sheet was successfully transferred to the target substrate within 15 min at 4 °C. After the transfer, the bilayer cell sheets not only maintained extracellular matrix and inherent cell markers, but also showed higher angiogenic functions in comparison with monolayer cell sheets. An in vitro study confirmed that the bilayer sheet secreted significantly greater amounts of VEGF and stimulated more in vitro tubule formation compared to the monolayer cell sheets. In addition, the bilayer sheet had significantly increased NO release and expression of proangiogenic genes. Finally, the bilayer cell sheet transplant group demonstrated significantly slower necrosis compared to the monolayer cell sheet groups. Immunohis tochemistry revealed a significant number of arterioles in the bilayer cell sheet transplant group compared to the nontreated and monolayer cell sheet groups. Taken together, the findings demonstrate that this system can be utilized as a coculture model system for the study of cell–cell interactions and as a tool for direct transplantation of cocultured cells with a spatially assembled structure.
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4. Experimental Section Materials: Tetronic-tyramine was synthesized as previously described.[14,36] Human plasma FN and purified antifibronectin were purchased from BD Biosciences (Franklin Parks, NJ, USA). Anti-CD31 was purchased from Cell Signaling Technology Inc. (Danvers, MA, USA). Hoechst 33258, Vybrant DiD and DiO cell-labeling solutions, Live/Dead viability/cytotoxicity kits, and NucBlue Live ReadyProbes Reagent were purchased from Molecular Probes (Eugene, OR, USA). The antimouse IgG biotin conjugate, Mayer’s hematoxylin, horseradish peroxidase (HRP), and hydrogen peroxide (H2O2) were purchased from SigmaAldrich (St. Louis, MO, USA). Fetal bovine serum (FBS) was obtained from Wisent (St.-Bruno, QC, Canada), and Dulbecco’s modified Eagle’s medium with low glucose (DMEM), Dulbecco’s phosphate buffered saline (PBS), trypsin/EDTA, and penicillin–streptomycin (p/s) were purchased from Gibco BRL (Carlsbad, CA, USA). Rhodamine-phalloidin was purchased from Invitrogen Corp. (Carlsbad, CA, USA). Cell Culture: HUVECs (CC-2519, Lonza, Basel, Switzerland) and hMSCs (PT-2501, Lonza, Basel, Switzerland) were cultured in EGM-2 and DMEM containing 10% FBS, respectively, supplemented with 1% p/s under standard culture conditions (37 °C, 5% CO2). For all experiments, both cell types were used between passages 4 and 6. Media composed of DMEM and EGM-2 mixed in a 1:1 ratio was used for the
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bilayer coculture of hMSCs and HUVECs. Seeding densities of hMSCs and HUVECs were 2.5 × 104 and 1 × 105 cells cm−2, respectively. The cocultured bilayer was formed by seeding HUVECs on a layer of hMSCs at the same seeding density as was used in the HUVEC monolayer. Preparation of FN-Hydrogel: Hydrogels were prepared using the previously described method with minor modifications.[13] Two separate solutions of 7.63 wt% Tetronic-tyramine were prepared with either 0.01 wt% H2O2 in PBS or 0.0025 mg mL−1 of HRP in PBS. The solution with HRP also contained defined concentrations of fibronectin (0 and 50 µg mL−1 for FN-0 and FN-50 hydrogels, respectively). Each solution was loaded into a dual syringe and injected between two glass plates lined with a Teflon mold (0.5 mm in thickness). Hydrogels underwent gelation, and hydrogel disks were subsequently prepared via a biopsy punch (8 mm diameter). Hydrogel disks were washed several times with PBS. Characterization of Cell-Interactive FN-Hydrogels: The incorporation of fibronectin within the FN-hydrogel was confirmed using the reactive dye fluorescein isothiocyanate (FITC). The hydrogels were immersed in 1 mL of a FITC solution (2 mg mL−1 in EtOH) at room temperature for 4 h and then continuously washed with EtOH for 2 h, followed by PBS for 6 h. FITC-labeled FN hydrogels were visualized using a fluorescence microscope (Carl Zeiss, Oberkochen, Germany). Rheological experiments of the FN-hydrogels were performed using an Advanced Rheometer GEM-150-050 (Bohlin Instruments, USA) in the oscillatory mode. For measurements, 100 µL of Tetronic-tyramine dissolved in 0.01 wt% of H2O2 and 100 µL of Tetronic-tyramine dissolved in 0.0025 mg mL−1 of HRP with different fibronectin concentrations (0 and 50 µg mL−1) were placed on the plate of the rheometer. The elastic modulus (G′) was recorded at a frequency range of 0.1–10 Hz and a temperature of 37 °C. For oscillatory shear rheological measurements, parallel-plate geometry with a plate diameter of 25 mm, a gap of 0.5 mm, and a stress of 10 Pa was used. XPS (Versa Prove, Physical Electronics, Inc., Chanhassen, MN, USA) was used to investigate the surface chemical composition of lyophilized FN hydrogels. To lyophilize samples, FN-hydrogels were transferred into a deep-freezer (IL Shin Bio. Base, Korea) for 1 h and subsequently freeze-dried (Fisher Scientific, Boston, MA, USA) for an additional 24 h. Adhesion of hMSCs and HUVECs on FN-Hydrogels: The adhesion of hMSCs and HUVECs on FN-0 and FN-50 hydrogels was first evaluated. Both cell types were seeded separately on each hydrogel, with densities of 5 × 103 cells cm−2 (hMSCs) and 2 × 104 cells cm−2 (HUVECs). The cytoskeletal structures of both cells were observed using confocal laser scanning microscopy (LSM 700, Carl Zeiss, Oberkochen, Germany), and images were captured after 24 h. Next, the hMSC and HUVEC monolayer cell sheets and the bilayer cell sheet of the HUVEC monolayer on the hMSC monolayer (hMSCs/HUVECs) were fabricated. The seeding densities of hMSCs and HUVECs used to form the cell layer were 2.5 × 104 and 1 × 105 cells cm−2, respectively. For fabrication of the bilayer on FN-hydrogels, an initial layer of hMSCs was formed on the hydrogel. The cells were allowed to form the monolayer for 12 h, after which a specified number of HUVECs were seeded for bilayer construction. The morphology of cells and attachment of each cell type cultured on the other cell type were observed for up to 36 h under a light microscope. The number of adherent cells on FN-hydrogels was quantified using the LUNA Automated Cell Counter (Logos Biosystems, Korea). Proliferation of cells on the FN-hydrogel was evaluated for 96 h with a 4-[3-(4-iodophenyl)-2-(4-nitrophenyl)-2H-5-tetrazolio]-1,3-benzene disulfonate (WST-1) EZ-cytox cell viability assay kit (ITSBIO, Seoul, Korea). Briefly, hMSCs on hydrogels were replenished with a working solution after 24 h and further incubated for 2 h. HUVECs were also refreshed onto the hMSC layer cultured on hydrogels at designed assay points (12, 24, and 36 h) with a working solution. The enzymatic activity was measured at an absorbance of 440 nm using a spectrophotometer. All data were calibrated using the absorbance value of the cell-free hydrogel treated with the same working solution. Characterization of the Thermal Expansion of Hydrogels Using PIV: PIV was used to measure the thermal expansion of hydrogels by monitoring the displacement of fluorescent particles. Briefly, the FN-50
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hydrogel precursor solution containing fluorescent beads (FluoSpheres Fluorescent Microspheres, Invitrogen Corp., Carlsbad, CA, USA) was spin-coated (DONG AH Trade Corp., Korea) onto a small confocal dish at 1000 rpm for 60 s. The temperature was then reduced to 4 °C, and the dynamic particle movement in the FN-50 hydrogels was acquired by a confocal laser scanning microscope over 660 s at 60 s intervals. The acquired images were processed using a PIV lab package and a personalized program from MATLAB (Mathworks, Natick, MA, USA). The thermal expansion of FN-hydrogels was also characterized by measuring their diameters. Briefly, an FN-hydrogel sample (diameter 8 mm) was immersed in 1 mL PBS at 37 °C and sequentially incubated at 4 °C for 30 min. The change in size of the FN-hydrogels based on the change in temperature was then measured every 30 min for 4 h using a vernier caliper (Mitutoyo Corp., Japan). The size kinetics of the FN-hydrogels at 4 °C was recorded every 1 min for 15 min. Harvest and Transfer of Cell Sheets from the FN-Hydrogels: Cell sheets of hMSCs, HUVECs, and the hMSC/HUVEC bilayer were fabricated on FN-50 hydrogels. To characterize the cell sheets, cells confluently cultured on the hydrogels were faced and contacted to a glass surface and incubated for 15 min at 4 °C. After incubation, hydrogel disks were carefully peeled from the glass slide. Each transferred cell sheet was analyzed using the Live/Dead viability/cytotoxicity kit, and images were captured using fluorescence microscopy. For clearer imaging of bilayered cell sheet, a cocktail of nuclei staining reagent (NucBlue Live ReadyProbes Reagent, Thermo Scientific, R37605) with the Live/Dead reagents was made, which was treated to the transferred cell sheet, and quantified the number of cells positive for ethidium homodimer-1 (Red) to total nuclei number (Blue). For immunostaining, cell sheets were fixed with 4% paraformaldehyde for 20 min, permeabilized with cytoskeleton buffer (50 × 10−3 m NaCl, 150 × 10−3 m sucrose, 3 × 10−3 m MgCl2, 50 × 10−3 m Trizma base, and 0.5% Triton X-100, pH 6.8) for 10 min, and treated with a blocking buffer (5% FBS in PBS) for 1 h. Subsequently, transferred cell sheets were incubated with antifibronectin (1:100) and anti-PECAM (1:100) primary antibodies for 1 h. Samples were then incubated with an antimouse IgG biotin conjugate (1:100) and subsequently with FITC-conjugated streptavidin (1:100) for 1 h. Cell nuclei and F-actin were counterstained with Hoechst 33258 and rhodamine-phalloidin, respectively. For observation of the bilayer cell distribution after transfer, hMSCs and HUVECs were labeled with Vybrant DiD (red fluorescence) or Vybrant DiO (green fluorescence) for 30 min before seeding on FN-50 hydrogels. After 24 h of incubation, the bilayer cell sheets were transferred to glass and fixed with 4% paraformaldehyde. Similarly, the transferred bilayer cell sheet was examined for the distribution of endothelial cell-specific markers. A separate, nonlabeled transferred bilayer was fixed and immunostained for PECAM. Transferred bilayer cell sheets were observed using confocal laser scanning microscopy to examine cell distribution and endothelial cell-specific markers. In Vitro Angiogenesis Analysis of Cell Sheets: HUVEC, hMSC, and bilayer cell sheets were cultured for an additional 3 d, and the supernatant was then collected. For the in vitro tubule formation assay, 100 µL cm−2 of growth factor reduced Matrigel (GFR-Matrigel) was coated on a 24-well plate overnight at 37 °C, on which HUVECs were cultured for 24 h. The media of the HUVECs cultured onto the Matrigel was then replaced with conditioned media collected from hMSC, HUVEC, and bilayer cell sheet cultures. Finally, tubule formation was observed using light microscopy (Olympus, Tokyo, Japan), and images were captured after 12 h. Images were analyzed using the WimTube Formation image analysis platform (WIMASIS GmbH, Munich, Germany). Secretion of vascular endothelial growth factor (VEGF) from cells into the media was analyzed using enzyme-linked immunosorbent assay (ELISA) kits (R&D Systems, Minneapolis, MN, USA) according to manufacturer’s instructions. To detect the total NO production of the cell sheets, colorimetric analysis was used using the Griess reagent kit (Molecular Probes, Eugene, OR, USA). Briefly, the supernatant from each cell sheet culture was collected at various assay points (6, 12, 24, 48, and 72 h) and incubated with a working solution of the Griess reagent for an additional 30 min. The optical density was immediately measured at 548 nm using a
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Primer
Forward
Reverse
GAPDH
TTCGACAGTCAGCCGCAT
GCCCAATACGACCAAATC
VE-cadherin
CCTTCCCTTCCCTTCCTCTT
TGGGAGCATACAGACTGGGA
Tie-2
AGGATGGAGTGAGGAATGGC
AGAGGGCATCAAAAGGCTGT
vWF
AGCCGCATGACTGTTCTTTG
AGTTTGCAGGAAAAGGGGAA
spectrophotometer (Versa Probe, Physical Electronics, Inc., Chanhassen, MN, USA). Nitrite concentrations were determined using the Griess method with NaNO2 as a standard. The supernatant from the working solution of the Griess reagent was used as the control. Total RNA from the transferred cell sheets was extracted, concentrated, purified, and used to synthesize cDNA using a Maxime RT premix kit (Intron Biotechnology, Korea) according to the manufacturer’s instructions. The real-time RT-PCR data were analyzed using the comparative threshold cycle (Ct) method. The relative expression of each gene was calculated by comparison with the expression level of GAPDH and subsequently normalized to the values for the transferred hMSC cell sheet. The oligonucleotide primer sequences are listed in Table 1. Real-time RT-PCR using SYBR green master mix was performed with an AB 7500 sequence detection system (Applied Biosystems, Foster City, CA, USA) with 40 cycles of melting at 95 °C for 15 s with annealing and extension at 60 °C for 60 s. Mouse Model of Hindlimb Ischemia: Nude mice (BkNbt:BALB/c/nu/ nu males, four weeks old with 20–25 g body weight) were obtained from Narabiotec (Seoul, Korea) and housed in accordance with the guidelines of the Institutional Animal Care and Use Committee of Hanyang University (HY-IACUC-11-054). The mouse model of hindlimb ischemia was prepared as described previously.[12] Mice were randomly divided into three experimental groups (n = 10). Three transplant groups of hMSC (2.5 × 104 cells cm−2), HUVEC (1 × 105 cells cm−2), and bilayer (hMSCs/ HUVECs with same seeding density as in the monolayers) cell sheets on the FN-50 hydrogel (1.0 × 0.5 cm size) were prepared and then placed on the ischemic tissue. The tissue was allowed to stabilize for 5 min, and then three drops of precooled (4 °C) saline solution were applied on the top of the hydrogels for 15 min. Subsequently, the hydrogel was carefully peeled off, leaving the transferred cell sheet on the ischemic tissue. The no treatment group without cell sheet transplantation was designated as the negative control group. Photographs of the limbs were visually analyzed to determine the severity of ischemia. The physiological status of limbs was categorized as follows: limb loss (necrosis or loss of tissue above the knee), foot necrosis (necrosis of a toe or below the knee), or limb salvage (normal status without any sign of ischemia) at designated time points (3 and 21 d). Histological and Immunohistochemical Analyses: Ischemic limb tissue was retrieved after three weeks of treatment, frozen in OCT compound at −20 °C, and sectioned using a Cryocut microtome. Specimens were stained with H&E to examine tissue morphology and degeneration. Tissue sections were also stained with Masson’s trichrome (Histoperfect, Masson’s Trichrome Staining Kit, BBC Biochemical, Stanwood, WA, USA) to examine fibrosis in the ischemic regions. Captured images were selected for calculation of the fibrotic area. The fibrosis percentage was calculated as the ratio of muscle (red) to fibrosis (blue) using Nikon imaging software (NIS-Elements AR, Nikon Corp., Japan). Arterioles in the ischemic tissue were immunofluorescently stained with an antismooth muscle α-actin (SMA, Abcam, Cambridge, UK) antibody. The sections were counterstained with DAPI and examined using fluorescence microscopy. The arteriole density was determined by calculating the number of vessels mm−2. Statistics: All quantitative results were obtained from triplicate samples. Data are expressed as the mean ± standard deviation and were analyzed using SPSS 21.0 software (SPSS Inc., Chicago, IL, USA). Statistical analyses were performed using ANOVA and Tukey’s HSD test. P-values less than 0.05 were considered statistically significant.
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Supporting Information is available from the Wiley Online Library or from the author.
Acknowledgements I.J. and T.A. contributed equally to this work. This research was supported by a grant of the Korea Health Technology R&D Project through the Korea Health Industry Development Institute (KHIDI), funded by the Ministry of Health and Welfare, Republic of Korea (HI15C3049) and Technology Innovation Program (10050526), and Development of disposable diaper based on biomass-oriented biodegradable super absorbent polymers funded by the Ministry of Trade, industry & Energy (MI, Korea). Received: November 20, 2016 Revised: January 27, 2017 Published online:
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Supporting Information
Table 1. Primer sequences for real-time RT-PCR.
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