Engineered vascularized bone grafts

Abstract

Clinical protocols utilize bone marrow to seed synthetic and decellularized allogeneic bone grafts for enhancement of scaffold remodeling and fusion. Marrow-derived cytokines induce host neovascularization at the graft surface, but hypoxic conditions cause cell death at the core. Addition of cellular components that generate an extensive primitive plexus-like vascular network that would perfuse the entire scaffold upon anastomosis could potentially yield significantly higher-quality grafts. We used a mouse model to develop a two-stage protocol for generating vascularized bone grafts using mesenchymal stem cells (hMSCs) from human bone marrow and umbilical cord-derived endothelial cells. The endothelial cells formed tube-like structures and subsequently networks throughout the bone scaffold 4-7 days after implantation. hMSCs were essential for stable vasculature both in vitro and in vivo; however, contrary to expectations, vasculature derived from hMSCs briefly cultured in medium designed to maintain a proliferative, nondifferentiated state was more extensive and stable than that with hMSCs with a TGF-beta-induced smooth muscle cell phenotype. Anastomosis occurred by day 11, with most hMSCs associating closely with the network. Although initially immature and highly permeable, at 4 weeks the network was mature. Initiation of scaffold mineralization had also occurred by this period. Some human-derived vessels were still present at 5 months, but the majority of the graft vasculature had been functionally remodeled with host cells. In conclusion, clinically relevant progenitor sources for pericytes and endothelial cells can serve to generate highly functional microvascular networks for tissue engineered bone grafts.

Fig. 1.

Protocol timeline. Unlabeled hMSCs were seeded and allowed to attach to the bone scaffold. Medium containing osteoinductive agents was used to commit these cells to the bone lineage, after which it was replaced with a hydrogel containing labeled ECs (red) and additional hMSCs (25% labeled green) to serve as pericytes. The scaffold was continued to be cultured in vitro or implanted the following day.

Fig. 2.

hMSCs contribute to in vitro formation of a 3D vascular network within a porous scaffold. (A) The bone scaffold was synthesized of the biocompatible polymer (85∶15) PLGA, by using sucrose leaching to generate interconnected pores averaging 1000 μm. (B) Stable network containing HUVECs and hMSCs in fibronectin-containing collagen gel at 21 days postseeding. (C) Physical interaction of hMSCs with EC network formed in the PLGA scaffold at 28 days postseeding. (D) Relative transcriptional response of the smooth muscle markers smooth muscle actin (ACTA2), calponin (CNN1), SM22 (TAGLN), and smoothelin (SMTN) to the 5-day preconditioning treatment. (E–H) In vitro formation of networks by HUVECs imaged with confocal microscopy at 10 days (E) without hMSCs, (F) with standard condition hMSCs, (G) with hMSCs cultured with TGF-β, and (H) hMSCs cultured in low serum MesenPro medium. (Scale bars: A = 1 mm; ; , . ^∗P < 0.05.)

Fig. 3.

Formation of engineered vessels in vivo. (A–D) Networks formed by HUVECs imaged with confocal laser-scanning microscopy at 4 weeks (A) without hMSCs, (B) with standard condition hMSCs, (C) with hMSCs cultured with TGF-β, and (D) with hMSCs cultured in low serum MesenPro medium. For (D) only, maximum intensity projection of 30 consecutive frames (1 sec) was used to demonstrate flow of DiD-labeled blood cells. H&E and human-specific CD31 immunohistochemical staining of scaffolds explanted 4 (E–L) and 11 (M–T) days after implantation. Acellular regions (*) derive from scaffold. EC-derived tubular structures have formed by 4 days (arrowheads) but do not contain blood cells indicating that they are not yet functional. By 11 days, the structures were larger with more robust CD31 staining, and most lumens were filled with blood cells (arrows). The symbols mark only a representative sample of indicated structures. (Scale bars: ; .)

Fig. 4.

Perivascular localization of hMSCs. A seeded scaffold containing HUVECs and MesenPro-treated MSCs was explanted after 4 weeks and embedded in paraffin. Serial sections cut from a region containing several transverse vessels (two of which are marked by arrowheads) were immunohistologically stained to identify human-derived structures: a human-specific antibody for PE-CAM (CD31) stained graft-derived endothelium, vascular-associated MSCs stained positive for the mural markers smooth muscle actin (ACTA2) and calponin (CNN1). Numerous erythrocytes within lumens indicate that the vasculature has anastomosed and is functional. Positive signals are brown with hematoxylin counterstaining. (Scale bar: 25 μm.).

Fig. 5.

Assessment of functionality of the engineered vessels within the porous scaffold. At 2 weeks, tail vein injection of Evans blue was used to visualize flow through the grafts by intravital microscopy in order to verify anastomosis and assess vessel permeability (A). The assay was repeated at 8 weeks postimplantation on additional implants with a second vascular probe, Qtracker800. In this case only very minimal leakage was detected. (C) Points of anastomosis, where dye-labeled serum flowing through the graft (red) and adjacent host vasculature clearly indicate that the engineered vessels have anastomosed with the mouse vessels. (D) Vibrant DiD-labeled RBC flow rates through graft and neighboring host vessels were calculated. Vessel density (E) and average diameter (F) in grafts with indicated hMSCs 4 weeks after implanting. ^∗P < 0.05, ^∗∗P < 0.01 compared with standard medium conditions. (Scale bar: .)

Fig. 6.

Osteogenic induction. At 8 weeks, scaffolds that had been seeded with standard condition osteoinduced hMSCs were explanted and calcium deposits stained with von Kossa. Calcium deposition was extensive and restricted to the pore surfaces of the scaffolds. Vascularized scaffolds (B) in which the initial seeding for attached hMSCs had not undergone prior in vitro osteogenic induction had more extensive mineralization than avascular scaffolds (C) but less than (A). A section from a scaffold (A) also stained with Alizarin red (D). This scaffold was also immunopositive for the late marker of osteoblast differentiation, osteocalcin (brown with hematoxylin counterstaining) (E, magnified in F). (Scale bars: ; .)