Tunable Keratin Hydrogels for Controlled Erosion and Growth Factor Delivery

Abstract

Tunable erosion of polymeric materials is an important aspect of tissue engineering for reasons that include cell infiltration, controlled release of therapeutic agents, and ultimately to tissue healing. In general, the biological response to proteinaceous polymeric hydrogels is favorable (e.g., minimal inflammatory response). However, unlike synthetic polymers, achieving tunable erosion with natural materials is a challenge. Keratins are a class of intermediate filament proteins that can be obtained from several sources, including human hair, and have gained increasing levels of use in tissue engineering applications. An important characteristic of keratin proteins is the presence of a large number of cysteine residues. Two classes of keratins with different chemical properties can be obtained by varying the extraction techniques: (1) keratose by oxidative extraction and (2) kerateine by reductive extraction. Cysteine residues of keratose are "capped" by sulfonic acid and are unable to form covalent cross-links upon hydration, whereas cysteine residues of kerateine remain as sulfhydryl groups and spontaneously form covalent disulfide cross-links. Here, we describe a straightforward approach to fabricate keratin hydrogels with tunable rates of erosion by mixing keratose and kerateine. SEM imaging and mechanical testing of freeze-dried materials showed similar pore diameters and compressive moduli, respectively, for each keratose-kerateine mixture formulation (∼1200 kPa for freeze-dried materials and ∼1.5 kPa for hydrogels). However, the elastic modulus (G') determined by rheology varied in proportion with the keratose-kerateine ratios, as did the rate of hydrogel erosion and the release rate of thiol from the hydrogels. The variation in keratose-kerateine ratios also led to tunable control over release rates of recombinant human insulin-like growth factor 1.

Figure 1

(A) An idealized tissue engineering approach depicting tissue regeneration occurring in a manner inversely proportional to material erosion. (B) Structure of a wool fiber, similar to human hair as originally drawn by Fraser and Macrae (CSIRO Australia). Reprinted from Biomaterials v. 29(1), Sierpinski et al, “The use of keratin biomaterials derived from human hair for the promotion of rapid regeneration of peripheral nerves”, pages 118–128, 2008 with permission from Elsevier. Alpha keratins (circled) were used in this study. (C) Schematic of cysteine residues in keratin following oxidative (left) or reductive (right) extraction and the resulting differences in disulfide crosslinking (black lines) and chain entanglement in hydrogels of keratose (left) and kerateine (right). (D) Approach to modulate disulfide crosslinking for control of material erosion and release of therapeutic agents (black circles). Reprinted from Acta Biomaterialia v. 23, Han et al., “Alkylation of human hair keratin for tunable hydrogel erosion and drug delivery in tissue engineering applications”, pages 1189–1197 (graphical abstract), 2015 with permission from Elsevier. (E) Images of cylindrical form of each keratin hydrogel formulation (5 mm diameter): (i) KOS, (ii) 70:30 KOS:KTN, (iii) 50:50 KOS:KTN, (iv) 30:70 KOS:KTN, and (v) KTN.

Figure 2

Rheological data for each formulation. (A) G’ (elastic modulus) and (B) G” (viscous modulus) for frequency sweep from .01 to 10 Hz. Error bars indicate standard deviation and n = 3 for each formulation. The trend indicates increasing elastic modulus with increasing levels of disulfide crosslinking (increasing levels of KTN). For statistical comparisons (one-way ANOVA followed by Tukey’s post hoc test): a indicates P < 0.05 vs. KTN; b indicates P < 0.05 vs. 30:70 KOS:KTN; c indicates P < 0.05 vs. 50:50 KOS:KTN; d indicates P < 0.05 vs. 70:30 KOS:KTN; and e indicates P< 0.05 vs. KOS.

Figure 3

Compressive moduli for each keratin formulation for (A) freeze-dried scaffolds and (B) hydrogels. Error bars indicate standard deviation and n = 3 for each formulation. * Indicates P < 0.05 vs. KTN as determined by Tukey’s post-hoc test.

Figure 4

Scanning electron micrographs of keratin gels. Each image is representative of the various formulations: (A) KOS (100:0 KOS:KTN) (B) 70:30 KOS:KTN. (C) 50:50 KOS:KTN. (D) 30:70 KOS: KTN. (E) KTN (0:100 KOS:KTN). Pore size and overall structure can be discerned. Images taken at 500X and 5kV. Scale bars represent 100 pm.

Figure 5

Viability of MC3T3-E1 cells in keratin hydrogels immediately after formation (Day 0) or after 3, 5, or 7 days of incubation in the formulations indicated. Ratios indicate KOS:KTN ratios. 0:100 KOS:KTN (100% KTN) is absent due to its inability to gel in the salts present in cell culture media. Cells stained green (calcein-AM) represent live cells. Images were chosen to be representative of the amount and distribution of cells in each formulation. Scale bar represents 250 µm.

Figure 6

(A) Cumulative erosion of keratin gel mass with time, demonstrating the relative differences in erosion rate by simply changing the KOS:KTN ratios. Data are shown as percentage of the total protein (15 mg) for each hydrogel formulation. Error bars represent standard deviation and n = 4 for each formulation. Each formulation was significantly different than every other formulation by the ~12 hour time point. (B) Cumulative percent of thiol release with time. Data are shown as percentage of the total thiol content released based on the thiol content in gel starting materials (kerateine and keratose). Error bars represent standard error of the mean and n = 3 for each formulation.

Figure 7

Release of rhIGF-1 from keratin gels with time. Each sample was loaded with 100µg of rIGF-1 per mL of keratin hydrogel. Each data point represents cumulative release as a percentage of initial rhIGF-1 loading. The rates of release can be seen to correlate with the rates of erosion seen in Figure 6. Error bars represent standard deviation and n = 4 for each formulation. Each formulation was statistically different than every other formulation by the ~12 hour time point except that 30:70 and 50:50 KOS:KTN did not reach statistical significance until day 6.