A biomimetic hyaluronic acid‐silk fibroin nanofiber scaffold promoting regeneration of transected urothelium

Abstract This study was designed to investigate the regulatory effect of hyaluronic acid (HA)—coating silk fibroin (SF) nanofibers during epithelialization of urinary tract for urethral regeneration. The obtained electrospun biomimetic tubular HA‐SF nanofiber scaffold is composed of a dense inner layer and a porous outer layer in order to mimic adhesion and cavernous layers of the native tissue, respectively. A thin layer of HA‐gel coating was fixed in the inner wall to provide SF nanofibers with a dense and smooth surface nano‐topography and higher hydrophilicity. Compared with pure SF nanofibers, HA‐SF nanofibers significantly promoted the adhesion, growth, and proliferation of primary urothelial cells, and up‐regulate the expression of uroplakin‐3 (terminal differentiation keratin protein in urothelium). Using the New Zealand male rabbit urethral injury model, the scaffold composed of tubular HA‐SF nanofibers could recruit lumen and myoepithelial cells from the adjacent area of the host, rapidly reconstructing the urothelial barrier in the wound area in order to keep the urinary tract unobstructed, thereby promoting luminal epithelialization, smooth muscle bundle structural remodeling, and capillary formation. Overall, the synergistic effects of nano‐topography and biophysical cues in a biomimetic scaffold design for effective endogenous regeneration.

oral mucosa tissue. 6,7 However, such method has a number of disadvantages, such as donor injury, limited donor tissue leading to multiple subsequent surgeries, and various possible treatment complications, such as urinary fistula, urethral stricture, and fibrotic scar tissue formation. 8,9 Urethral grafts prepared by traditional tissue-engineering methods have good performance, high patency, and best biocompatibility, 4,6,[8][9][10] which are consistent with our previous report that cell scaffold is a necessary condition for successful tissue regeneration of tubular structure. [11][12][13] However, the invasiveness, high cost, and long production time seriously limit their clinical translation.
In situ tissue-engineering using degradable biomaterials can utilize the regeneration potential of human body through reasonable scaffold design (including structural optimization and functionalization) and reproduce natural tissue regeneration. 14 In this field, some studies have shown that in situ tissue-engineered scaffolds produced new urethras nearly free of foreign materials in vivo. [15][16][17] However, few of them can reproduce the biological function of damaged urethra in the early stage of transplantation, and there are difficulties in subsequent lumen epithelial tissue remodeling.
Urothelium is a stratified transitional epithelium derived from the endoderm, extending from the renal pelvis to the proximal urethra, serving as a key barrier between the blood and urine. 3 It is composed of cytokeratin-5 (K5) expressing basal cells, intermediate cells, and surface cells specialized for synthesis and transport of uroplakins that assemble into the apical barrier. 5,18 The migration of urothelial cells (UCs) usually occurs during urethral regeneration in response to urethral injury and disease. [18][19][20] It has been found that urethral stem/ progenitor cells can differentiate into UCs in vitro and in vivo. 4,9,10 However, little is known about leveraging the recruitment and migration of mature UCs as a strategy to regenerate the urethral epithelium at the injury zone.
Among the potential migration enhancements for epithelial cells, hyaluronic acid (HA), a critical element of extracellular matrix (ECM), 21,22 can be used to bind with the cell surface receptors (e.g., CD44, RHAMM, and ICAM-1) [23][24][25] or to tune the mechanical properties of the ECM, 26,27 in turn, promoting cell motility and connecting tissues. With a wealth of enriching features such as high hydrophilicity, 28 immune neutrality, 29,30 and biodegradability, 31 HA and its derivatives have been widely used as suitable materials for surgical implants (e.g., cosmetic surgery) and tissue regeneration, [28][29][30] or as a drug conjugate for targeted delivery. 32,33 Previous work in our laboratory has shown that the HA-coated collagen fiber composite scaffold manufactured by coaxial electrospinning technology can induce regenerative immune response in the traumatic urethral region and can recruit and support the proliferation of urethral stem/ progenitor cells. 17 The results showed that scaffolds mimicking ECM by using protein and HA-based biomaterials may play a much more important role in guiding the development of cell behavior and the formation of functional tissue than previously thought. However, limited mechanical properties and rapid degradation restrict the use of collagen-based scaffolds in tubular geometry. [15][16][17]34 Silk fibroin (SF) is a natural protein with excellent mechanical properties, good cell compatibility, controllable degradation, and versatile process-ability in different material formats. [35][36][37] The potential of SF nanofibers in regenerative medicine has been investigated in some tissue-engineering applications, such as urethras, 5,16 skin, 38 tendon, 39 and bone regeneration. 40 However, to our knowledge, the utilization of biomimetic urethral scaffolds built with HA and SF nanofibers has not been explored in urethral reconstruction. Herein, HA was integrated on the surface of SF nanofibers by electrospinning technology to prepare biomimetic urethral scaffolds based on biomimetic principles. Degradable SF nanofibers components provide sufficient mechanical strength and appropriate peptide/protein for urethral scaffold. HA coating integrated on the surface of SF fiber imparted glycoproteins to the hybrid scaffold, supporting UCs motility and organization, contributing to epithelialization and physiological function reestablishment. To testify this, the surface topography and physicochemical properties of HA-SF nanofiber scaffolds were characterized by scanning electron microscopy (SEM), X-ray photoelectron spectroscopy (XPS), static water contact angle measurement, Fourier transform infrared spectra analysis with attenuated total reflection head (ATR-FTIR), thermo-gravimetric analysis (TGA), tensile test and atomic force microscopy (AFM). On this basis, the adhesion, proliferation, and phenotype of primary UCs on HA-SF nanofibers were studied. Finally, the urethral scaffold composed of tubular HA-SF nanofibers was constructed and implanted into the defect site of rabbit urethra for in vivo urethral regeneration assessment.

| Characterization of SF and HA-SF nanofibers
Using electrospinning, the inner wall surface nano-topography of tubular SF and HA-SF nanofibers present a smooth and porous nanofiber network, with the average diameters in the range of about (253 ± 15) nm and (222 ± 10) nm, respectively ( Figure S1). To increase architecture stability, tubular SF and HA-SF nanofibers were treated with EDC/ethanol solution. The nano-topography of tubular SF and HA-SF inner surface changed significantly after treatment (Figure 1a; Figure S1). The inner surface of SF exhibits tight interlinked nanofiber network nano-topography, and the HA-SF interconnect nanofiber network surface is decorated with gel-like concave coating morphology (Figure 1a). The surface nanotopography of the HA-SF nanofibers is very close to the microscopic morphology of the epithelial layer of native urethra. Quantitatively, the mean diameters of SF and HA-SF nanofibers were in the range of (286.3 ± 16.7) nm and (254 ± 13) nm, respectively, which was slightly larger than that (237.2 ± 16) nm of the native urethral tissue (Figure 1b).
Since the inner wall surface of HA-SF has a layer of HA gel-like coating, the obtained tubular architecture is composed of a dense inner layer and a porous outer layer, while the SF scaffold is composed of multiple porous layers ( Figure S2).
HA is a highly hydrophilic biopolymer, so it is reasonable to expect that the SF nanofibers coated by HA have high wettability, which can be determined by water contact angle measurement. As shown in Figure 1c and Table S1, the average water contact angles of SF and HA-SF nanofibers are in the range of (80 ± 2) and (65.9 ± 1) , respectively. Suggested that HA-coating can improve the hydrophilicity of SF nanofibers, probably due to the presence of hydroxyl and carboxyl groups in HA molecules.
The chemical distribution of HA-SF nanofibers was determined by ATR-FTIR (Figure 1d). In the ATR-FTIR spectrum, the characteristic absorption peaks of SF nanofibers are obvious at 1652, 1559, and 1287 cm À1 , consistent with that of amide I, amide II, and amide III, respectively. Compared with SF nanofiber film, the ATR-FTIR spectra of HA-SF nanofiber show obvious additional absorption peaks at 1646, 1556, and 1287 cm À1 . The band shifts of amide I and II were observed. Other studies have found similar band shifts. [37][38][39][40] These changes may be due to the strong interaction between SF and HA molecules, such as hydrogen bond and electrostatic interaction, ultimately indicating HA had successfully coated on SF nanofiber. In addition, XPS was used to further confirm that the presence of HA coating had no effect on the surface chemical elements of SF nanofiber (Figure 1e; Figure S3). There are no additional element peaks in the surface chemistry of SF and HA-SF nanofiber, indicating that this coating method has no obvious effect on the surface chemical element of SF and HA. The existence of HA-coating was further confirmed by TGA ( Figure S4). Compared with HA and SF nanofiber, the T d10 of HA-SF nanofiber is 366.32 C, which is between that of HA and SF nanofibers ( Figure S4 and Table S1), further confirms the existence of HA-coating.
Both SF and HA-SF nanofibers were tested in tensile mode to generate stress-strain curves and derive tensile properties ( Figure S5 and Table S1). The Young's modulus and ultimate tensile strength of HA-SF nanofibers in dry state are (0.83 ± 0.4) MPa and (1.7 ± 0.4) MPa, respectively, which are close to those of SF nanofibers. However, the elongation at break of HA-SF is (387 ± 17)%, which is slightly lower than that of SF nanofibers (407 ± 20)%. These tests showed that HA-SF and SF nanofibers are soft but tough substrate.  Figure S7). It is worth noting that compared with SF nanofiber thin films, the thin HA gel coating of HA-SF nanofibers not only contributes to the uniform distribution of UCs on its surface but also its porous SF nanofibers contribute to the ingrowth of UCs.
Meanwhile, uropakin-3 was used to visualize urothelial plaque in UC to evaluate waterproof and anti-injury functions. Figure 2c shows the typical confocal laser scanning microscope (CLSM) images of UC uropakin-3 positive in the cross section of different nanofiber films.
The density of uropakin-3 positive (green fluorescence) on HA-SF nanofibers was significantly higher than that of SF nanofibers ( Figure 2d). These uropakin-3 staining suggests that HA-SF nanofibers facilitates the urothelial barrier restoration.
To study the effects of SF and HA-SF nanofibers on the proliferation of UCs, we performed immunofluorescence staining of Ki67, a recognized proliferation marker located in the nucleus. In Figure 3a  2) ml/s was slightly lower than that (8.9 ± 0.2) ml/s of preimplants, significantly higher than that (6.6 ± 0.2) ml/s of SF graft. Furthermore, Masson's trichrome stain was used to histologically examine neo-tissue remodeling in the regenerated urethral tissue. As shown in   (Figure 6a). At 14 weeks after implantation, compared with SF graft group, the morphology of newly formed smooth muscle bundle (red) in the regenerated urethra of HA-SF graft group was close to that of normal urethral than that of SF graft group (Figure 6a,b). The percentage of smooth muscle in the HA-SF group was (21 ± )%, which was higher than that in the SF group (13.8 ± 1.6)%, and close to (23.7 ± 0.9)% of the normal urethra.
The outer layer was composed of smooth, wavy collagen fibers, quantitatively showing no significant difference in the content of collagen fibers in all samples. In addition, immunofluorescence staining confirmed that more α-SMA positive (green fluorescence) signal was observed in the regeneration zone of HA-SF graft group, which was close to the α-SMA positive area of normal urinary tract smooth muscle cells; comparatively, α-SMA positive signal in the regeneration region of SF graft was weak (Figure 6c,d). The percentage of α-SMA positive area in HA-SF graft regeneration area was (21.76 ± 1.6)%, which was close to that (24 ± 1.2)% of normal urethra (p > 0.05), and significantly higher than that (12 ± 1.2)% of SF graft group. These results suggest that HA-SF grafts can induce structural remodeling of smooth muscle cells, rather than recruitment of single cells. As the outer layer of tubular HA-SF nanofibers is mainly SF nanofibers, its biophysical clues and surface properties are exactly the same as those of pure SF nanofibers. Therefore, we speculate that the urinary epithelium on the inner wall of tubular HA-SF graft may determine the tissue remodeling pattern of SMC on its outer wall. The formation of vascular network in regenerated tissue is the key to tissue integration. 42,43 Compared with SF group, there was extensive neovascularization in HA-SF group ( Figure 6c). As shown in Figure 6c

| DISCUSSION
In this study, we developed a biomimetic tubular nanofiber scaffold that promotes urethral functional recovery by recruiting healingassociated endogenous UCs to reconstruct the urinary tract epithelial barrier. We found that by precisely controlling the biophysical and This method harnesses the regenerative potential of the human body and eliminates the need for ex vivo cell manipulation. 14,15 In addition to the limited number of endogenous stem cells and progenitor cells in cartilage, heart, and nerve tissues, in situ tissue engineering provides temporary solutions and alternatives for most of the tissue regeneration. 14 In situ tissue engineering utilizes bioresponsive scaffolds that harness the inherent regeneration ability of the body. These scaffolds are loaded with biochemical and biophysical clues to recruit endogenous cells for tissue healing. As alternatives to autografts, artificial urethra made of natural degradable biomaterials (such as collagen and SF) are promising candidates in terms of tissue integration, 5,15-17 but lack the capability to induce tissue regeneration due to the bio-inert characteristics. Hence, the manufacture process of bioactive or bio-functional urethral grafts has been widely explored through the application of surface modifications and the addition of bioactive molecules. The ECM plays an important role in regulating organogenesis, growth, function, and many human diseases. 23 As one of the key components of ECM, HA is very important for the migration, proliferation, differentiation, and intercellular communication of many cell types. 24,25 Herein, urethral graft based on HA and SF with fibrous structure was successfully prepared by sequential electrospinning. The biomi- epithelium. The main advantage of this structural design is that it successfully combines the protein with polysaccharide, which mimic the key physiological features of ECM. 44 In vitro cell scaffold co-culturing experiments showed that the gel-like coating on the inner wall of HA-SF scaffold was beneficial to the adhesion of primary UCs on its wall surface, and its hydrophilicity can provide a moisture-rich microenvironment for the growth of primary UCs. The interconnected pore networks facilitate UCs infiltration, migration, and scaffold remodeling. The proliferation ability of urethral UCs on HA-SF nanofibers is higher than that on SF nanofibers, which may be attributed to the bioactivity of HA coating in SF nanofibers. It has been reported that the combination of HA and CD44 can enhance various physiological properties, such as cell matrix adhesion and migration. [23][24][25] Moreover, the factors that affect the reaction of urethral UCs cells may include matrix stiffness and elasticity. 8 Here, the Young's modulus of HA-SF and SF nanofibers measured by our tensile test and AFM shows that HA coating has little effect on the stiffness of matrix. Therefore, the mechanical properties of SF nanofiber scaffold are maintained and matching the native urethral tissue modulus, helping the proliferation of UCs.
One of the advantages of biomimetic tubular HA-SF nanofiber scaffold is the exploitation of regenerative potential of organism, controlling cell function to realize in situ tissue reconstruction. Moreover, the use of endogenous UCs for in situ urethral regeneration can also reduce immune rejection of transplanted (exogenous) cells, one of the main risks associated with cell therapy. 15,17 In vivo experiments testify that the biomimetic tubular HA-SF nanofiber scaffold could direct endogenous UCs to the injury site aiding lumen re-epithelization in vivo. In this process, the tubular HA-SF nanofibers provide a structural framework to facilitate the attachment and migration of host endogenous UCs along its inner wall surface. In situ degradation of scaffold is desired for tissue regeneration. 14

| Preparation of electrospun SF and HA-SF nanofiber
The HA solution was prepared by dissolving 200 mg HA in 10 ml HFIP and stirring until a uniform solution was obtained. A syringe with 10 ml HA solution was fixed on the precision injection pump. The flow rate of the electrospinning device was set at 1.5 ml/h, the distance between the electrospinning nozzle and the receiver (stainless steel with an outer diameter of $2.7 mm or stainless steel rod of 10 mm) was set at 15 cm, while voltage was set at 12.3 kV. The speed of receiver was set at 1000 rpm, the temperature and humidity were 25 C and 45%, respectively. Electrospinning stopped after 2 or 6 h. A second syringe was loaded with 10 ml of fully mixed SF solution (50 mg/ml) and fixed to the precision injection pump. After connecting the electrospinning nozzle, the flow rate of the electrospinning device was set to 2.8 ml/h. The purpose of using a receiving device with 10 mm stainless steel rod was to obtain SF fiber film with HA-coating on inner surface, while stainless steel with diameter of 2.7 mm was used to obtain tubular HA-SF scaffold with inner diameter of 2.9 mm. HA-coating SF nanofiber scaffold with an inner diameter of 2.7 mm and an outer diameter of 3.5 mm was obtained after 48 h electrospinning. The electrospinning setup of pure SF nanofiber film and tubular nanofiber scaffold in this study is consistent with the HA-SF electrospinning procedure described above, but without the second HA electrospinning step.

| Characterization
The surface morphology and the diameter of the nanofibers were observed by SEM (SU8010, Hitachi) at an accelerating voltage of 5 kV. 45 The average diameter of nanofiber samples was measured from SEM images by ImageJ software. For each sample, 60 nanofibers were chosen randomly. The SF and HA-coating SF nanofibers were characterized by FTIR spectra analysis with ATR head (ATR-FTIR, thermo Nicolet, MA) in the range of 500-4000 cm À1 and scanning resolution of 2 cm À1 . 46,47 The surface chemistry elemental was assessed by XPS spectroscopy by means of an XPS Kratos Axis Ultra HSA apparatus, which uses a micro-focused monochromatic Al Kα X-ray source (1486.6 eV) covering an analyzing area of 300 Â 700 μm (900 W power). 48

| Cell growth and morphology on various nanofibers
The UCs of New Zealand rabbits were isolated using previously described methods, 11,13 and cultured in DMEM supplemented with FBS (10%, v/v), P/S (1%, v/v) in a 37 C, 5% CO 2 humidified incubator. After three passages, cells were seeded at a concentration of 10 6 cells/cm 2 on nanofibers and the medium was changed every day for the next 5 days. Prior to cell seeding, the nanofibers (diameter:

| Animal tests
All experimental procedures involving animals in this study were conducted under Institutional Guidelines for Animal Care and approved by the Animal Ethics Committee of Guangzhou Medical University (Guangdong, China). Male New Zealand rabbits aged 14 weeks were divided into two research groups: SF and HA-SF scaffold graft group.
Each animal received one graft, with nine animals in each group. The positive control was healthy urethral tissue, which was obtained from the 2 cm long urethral tissue cut during the manufacture of urethral trauma. As previously described, 11,13 under general anesthesia with pentobarbital sodium, the urethra skin was shaved and sterilized with Iodophor and 75% ethanol. Silk 5.0 fixing suture (A312; Jinhuan Medical Products Co., Ltd., China) was placed in the glans, a 6F catheter (Yiwu Medco Health Care Co., Ltd., China) was inserted into the bladder. A 2 cm skin incision was made at the proximal end of the glans, then the urethra within the cavernous tissue was separated. A 2 cm length urethral defect was made at 1 cm proximal to the base of the glans. A 2 cm length tubular SF or HA-SF nanofiber (inner diameter of 2.7 mm and outer diameter of 3.5 mm) graft was fixed on the urinary catheter, and the native urethra and the graft were anastomosed endto-end with PLGA 6.0 suture (LSP631; Jinhuan Medical Products Co., Ltd., China). Finally, the skin was sutured with PLGA 5.0 (LCC533).
Urothelial epithelial tissue (0.3 mm 2 ) was isolated from the above-mentioned healthy urethral tissue, while decellularized native urethral epithelial tissue was obtained by digestion with 0.2% collagenase IV (C2139, Sigma) at 37 C for 2 h ( Figure S6).
As previously described, 11,13 under general anesthesia, the patency of the implanted graft was monitored by digital mobile X-ray

| Histological and immunofluorescence assessment
At 6 weeks, and 14 weeks after urine flow rate experiment, the research animals were randomly sacrificed with an overdose of sodium pentobarbital (n = 3 animals per group), and the urethral tissue was dissected and separated. The retrieved graft samples were washed with normal saline to remove blood stains, fixed with 4% paraformaldehyde at RT for 24 h, dehydrated with gradient ethanol, embedded in paraffin, and cut into tissue sections (5 μm) were cut. 50 Sample were then stained with Masson's trichrome (C1006, Servicebio), VVG (GP1035, Servicebio), and Sirius red (GP1033, Servicebio) according to the protocol of the reagent manufacturer. The sample visualization was carried out using Pannoramic DESK microscope, and the representative fields were obtained through Caseviewer software (version 2.3). Three slides per graft per time point were used for histological analysis. The average urethral epithelial thickness, the percentage of smooth muscle bundles and collagen were calculated from 10 random fields per slide using the imageJ software. 13 For immunofluorescence staining of paraffin-embedded tissue sections, the sections were first de-paraffinized in xylene and then rehydrated in graded alcohol series, subjected to heat-induced epitope retrieval in 10 mM citrate buffer (pH 6.0, G1201; Servicebio), followed by the immunofluorescence staining protocol described in Section 4.5. The average K5, Uroplakin-3, CD31, or α-SMA positive expression area was measured on 10 random fields of each slide (three animals in each group) with ImageJ software.

| Statistical analysis
All experimental data were analyzed using GraphPad Prism (GraphPad Software, La Jolla) program. Where appropriate, a one-way or twoway analysis of variance (ANOVA) was performed to determine the significant difference with the Tukey post hoc test (p < 0.05). Unless otherwise stated, data and error bars are reported as mean ± standard deviation.

| CONCLUSIONS
In conclusion, we have successfully prepared biomimetic tubular HA-SF nanofibers by electrospinning and crosslinking process. The structure, morphology, and mechanical properties of the tubular HA-SF nanofibers are close to the native urethral tissue of New Zealand rabbits. In vitro cell inoculation showed that the surface of hydrophilic HA-SF nanofibers with biomimetic nanotopography is more suitable for the growth of primary rabbit UCs.
In vivo transplantation results showed that the dense gel coating at the inner wall of the tubular HA-SF nanofibers can recruit endogenous UCs from the adjacent area occupying the defect site to form new urothelia tissue. The porous outer layer is conducive to more uniform infiltration of SMC, thus achieving tissue integration and optimal regeneration.