Anisotropic liquid crystalline hydrogels direct 2D and 3D myoblast alignment

Abstract

Tissue development and regeneration are governed by processes that span subcellular signaling, cell-cell interactions, and the integrated mechanical properties of cellular collectives with their extracellular matrix. Synthetic biomaterials that can emulate the hierarchical structure and supracellular mechanics of living systems are paramount to the realization of regenerative medicine. Recent reports detail directed cell alignment on mechanically anisotropic but stiff liquid crystalline polymer networks (LCNs). While compelling, the potential implementation of these materials as tissue engineering scaffolds may be hindered by the orders of magnitude larger stiffness than most soft tissue. Accordingly, this report prepares liquid crystalline hydrogels (LCHs) that synergize the anisotropic mechanical properties intrinsic to LCNs with the cytocompatibility and soft mechanics of PEG hydrogels. LCH are prepared via sequential oligomerization and photopolymerization reactions between liquid crystalline (LC) monomers and poly(ethylene glycol) (PEG)-dithiol. Despite their low liquid crystalline content, swollen LCH oligomers are amenable to rheological alignment via direct ink write 3D printing. Mechanically anisotropic LCHs support C2C12 myoblast culture on their surface and direct their alignment in the stiffest direction. Further, C2C12s can be encapsulated within LCH oligomers and 3D-printed, whereby mechanical anisotropy of the LCH directs myoblast polarization in 3D.

Keywords: 3D bioprinting; anisotropic hydrogel; cellular alignment; liquid crystalline; tissue engineering.

Figure 1.. Synthesis of LC-PEG hydrogels.

(a) Chemical structures and simplified illustration of two-step LC-PEG synthesis and network structure. (b) Photographs of dry (left) and swollen (right) LC-PEG-5k network under crossed polarizers. The dashed white boxes indicate the sample edges; scale bars = 1 mm. (c) Mass swelling ratio (q) of LC-PEG networks in water. (c) Photographs of strained LC-PEG-5k hydrogel under crossed polarizers. Scale bars = 5 mm.

Figure 2.. Phase behavior of LC-PEG networks.

(a) Polarized optical micrographs of dry and swollen LC-PEG networks. Scale bars = 100 μm. (b) DSC thermograms of (top) dry and (bottom) swollen LC-PEG networks. (c) (Top) WAXS intensity as a function of q for a dry LC-PEG-3.5k network with varying temperature and (bottom) 2D scattering pattern of the network at 50°C. (d) (Top) WAXS intensity as a function of q for LC hydrogels with varying temperature and (bottom) 2D scattering pattern of LC-PEG-3.5k hydrogel at 40°C. (e) (Top) SAXS intensity as a function of q for LC hydrogels with varying temperature and (bottom) 2D scattering patterns of (left) LC-PEG-1k and (right) LC-PEG-3.5k hydrogels.

Figure 3.. Mechanical properties of swollen LC-PEG oligomers and LCHs.

(a) Photographs of swollen LC-PEG oligomers molded into small balls. Scale bar = 5 mm. (b) Mass swelling ratio (q) of LC-PEG oligomers in water. (c) Viscosity of swollen LC-PEG oligomers as a function of shear rate (s⁻¹). (d) Photographs of strained LC-PEG-3.5k hydrogel under crossed polarizers. Scale bars = 5 mm. (e) Evolution of LCH shear storage modulus during in situ photorheology. Blue region indicates UV light exposure at 10 mW cm⁻². (f) Tan(δ) of LCHs as a function of frequency at 0.1% strain. (g) Shear moduli of LCHs as a function of strain at 1 rad s⁻¹. (h) Normalized stress relaxation of LCHs after application of 5% strain over 3 s.

Figure 4.. Fabrication of anisotropic LCHs by shear alignment.

(a) Photographs taken under crossed polarizers of temporal decay of shear induced LC-PEG-1k birefringence at 4°C. Scale bars = 1 cm. (b) Photographs of shear aligned LC-PEG-1k hydrogels under crossed polarizers. Scale bars = 5 mm. (c) Representative stress-strain data from tensile testing of shear-aligned LC-PEG-1k hydrogels parallel and perpendicular to alignment direction. (d) Elastic moduli of shear-aligned LC-PEG-1k hydrogels in the parallel and perpendicular directions. (e) Photographs of 3D-printed LC-PEG-1k hydrogel grid taken under crossed polarizers immediately after printing. Scale bars = 500 μm. (f) Polarized optical micrographs of 3D-printed LC-PEG-1k hydrogel grid junction after swelling to equilibrium. Scale bars = 200 μm. (g) Photograph of LCH fiber extrusion and photopolymerization on rotating mandrel. (h) Polarized optical micrographs of LC-PEG-1k fiber after swelling to equilibrium. White dashed lines indicate edge of LCH fiber; scale bars = 200 μm. (i) Representative stress-strain data from tensile testing of LCH fibers. (j) Elastic moduli of LCH fibers.

Figure 5.. Anisotropic LCHs direct muscle cell alignment in 2D and 3D.

(a) Representative images of actin-stained C2C12 myoblasts colored by cellular orientation after three days of growth on shear aligned (left) and isotropic (right) LC-PEG-1k hydrogels. Double sided arrow indicates alignment direction; scale bars = 500 μm. (b) Orientation-order parameter (S) of C2C12 myoblasts after three days of growth. Statistical analysis was performed with one-tailed Student’s t-test, with significance claimed at **P < 0.01. (c) Live-dead staining of C2C12 myoblasts in aligned LC-PEG-5k fiber 2 hours after extrusion. Dashed white lines indicate edge of hydrogel fiber; scale bar = 200 μm. (d) Polarized optical micrographs of C2C12 myoblasts (red arrowheads) in aligned LC-PEG-5k fiber after three days of culture. Scale bars = 100 μm. (e) Polar frequency histogram (10° bins) of C2C12 alignment in aligned and isotropic LC-PEG-5k fibers after three days of culture. (f) Confocal fluorescence image of a polarized cluster of C2C12 myoblasts after six days of growth in aligned LC-PEG-5k fiber. Scale bar = 20 μm.