Proangiogenic scaffolds as functional templates for cardiac tissue engineering - PubMed (original) (raw)

Proangiogenic scaffolds as functional templates for cardiac tissue engineering

Lauran R Madden et al. Proc Natl Acad Sci U S A. 2010.

Abstract

We demonstrate here a cardiac tissue-engineering strategy addressing multicellular organization, integration into host myocardium, and directional cues to reconstruct the functional architecture of heart muscle. Microtemplating is used to shape poly(2-hydroxyethyl methacrylate-co-methacrylic acid) hydrogel into a tissue-engineering scaffold with architectures driving heart tissue integration. The construct contains parallel channels to organize cardiomyocyte bundles, supported by micrometer-sized, spherical, interconnected pores that enhance angiogenesis while reducing scarring. Surface-modified scaffolds were seeded with human ES cell-derived cardiomyocytes and cultured in vitro. Cardiomyocytes survived and proliferated for 2 wk in scaffolds, reaching adult heart densities. Cardiac implantation of acellular scaffolds with pore diameters of 30-40 microm showed angiogenesis and reduced fibrotic response, coinciding with a shift in macrophage phenotype toward the M2 state. This work establishes a foundation for spatially controlled cardiac tissue engineering by providing discrete compartments for cardiomyocytes and stroma in a scaffold that enhances vascularization and integration while controlling the inflammatory response.

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Conflict of interest statement

The authors declare no conflict of interest.

Figures

Fig. 1.

Fig. 1.

Analysis of bimodal scaffolds in vitro. (A and B) SEM images of bimodal scaffolds. Final scaffold design consists of 60-μm channels spaced 60 μm apart. Channel walls contain spherical pores with a 30-μm diameter and 15-μm interconnects. Note that the dehydrated structures are ∼80% of their hydrated size. (C and D) Digital volumetric imaging shows a fiber-templated scaffold (green) seeded with primary chick cardiomyocytes (red). Seeding is uniform across the channels (C), whereas the longitudinal cross-section (D) reveals a slight gradient in cell density. (E) Chick cardiomyocyte-seeded structure with positive staining for sarcomeric myosin heavy chain (Sarc; brown). Cardiomyocytes reside predominantly within the channels, whereas noncardiomyocytes migrate throughout the pores. (F) hESC-CM–seeded scaffolds cultured for 1 wk showing a high density of β-myosin heavy chain–positive (brown) cells within the channels. This scaffold measures ∼600 μm perpendicular to the page with this section taken at the midpoint, ∼300 μm into the scaffold. (Inset) The 40× magnification image shows that cardiomyocytes at the center of the construct exhibit shrunken cytoplasm, but intact nuclei indicate viability. (G) Immunolabeling against troponin T shows the presence of contractile proteins in hESC-CM seeded in the channel constructs. (Inset) The 100× magnification of the boxed area shows troponin T–positive hESC-CM oriented along scaffold channels. (H) Confocal image obtained using a live/dead assay shows the distribution of cells relative to the channel constructs (autofluorescent in red). (Scale bars: A, 100 μm; B, 20 μm; C and D, 300 μm; E, 50 μm; F, 400 μm; G, 50 μm; H, 400 μm.)

Fig. 2.

Fig. 2.

Acellular scaffolds implanted in the nude rat myocardium for 4 wk. (A) H&E overview of the implant, scar, and surrounding myocardium. (B) The 20× magnification of the boxed area in A shows the thin scar separating the implant from host tissue. Pores are filled primarily with granulation tissue including small vessels. (C) Endothelial lumens positive for the rat endothelial cell marker RECA-1 are present in the scaffold (arrows). (D) CD68+ macrophages infiltrated porous constructs (brown staining), but porosity limited fusion to foreign body giant cells. (E–H) Trichrome staining shows the collagen capsule (blue) and surrounding myocardium (red). (E) Nonporous and (F) 60-μm porous constructs had thicker and denser capsules than (G) 30-μm and (H) 20-μm pores. Insets show 20× images of boxed areas in E–H. (Scale bars: A, 100 μm; B, 10 μm; C and D, 50 μm; E_–_H, 250 μm; Insets, 50 μm.) (I) Neovascularization was assessed by quantification of RECA-1+ lumen structures (n = 3). (J) Thickness of the fibrous capsule surrounding implants was measured (n = 3).

Fig. 3.

Fig. 3.

Evaluation of vessel functionality in 4-wk myocardial implants in Sprague-Dawley rats by biotinylated lectin perfusion. (A) Lumen structures positive for biotinylated lectin (brown staining) were identified within porous implants (arrows) and host tissue (dart). (B) Perfused biotinylated lectin (streptavidin-FITC) and endothelial cells (RECA-1) were colocalized within the scaffold. Few unperfused endothelial structures were RECA-1+/lectin− (arrow). (C) Mature vessels had RECA-1+ lumens with a smooth muscle layer positive for α-smooth muscle actin. (D) Density of functional vessels was quantified over the entire implant with 40- and 80-μm porous scaffolds having significantly higher densities than nonporous and 20-μm porous scaffolds (n = 4; *P < 0.05). (Scale bars: A, 50 μm; B and C, 20 μm.)

Fig. 4.

Fig. 4.

Macrophage phenotype in response to 4-wk myocardial implants in Sprague-Dawley rats. (A) MΦ were identified with CD68+ staining. M1 and M2 phenotypes were determined by NOS2 (B) and MMR (C), respectively. (D) Overlayed images of CD68, NOS2, and MMR were analyzed to determine MΦ phenotype. A majority of MΦ in the porous implants expressed both NOS2 and MMR, although NOS2+/MMR− (darts) and NOS2−/MMR+ (arrows) MΦ could be identified (A–D). MΦ typically adhered to the material (D, brightfield Inset). (E) The fraction of each activated state was determined for CD68+ MΦ, with significant increase in NOS2+/MMR+ MΦ at all porous implant sites (n = 4; P < 0.05). There is a trend of increased NOS2−/MMR+ MΦ in 40-μm porous constructs versus nonporous (P = 0.06). (Scale bars: A_–_D, 50 μm.)

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