Keratose Hydrogel Drives Differentiation of Cardiac Vascular Smooth Muscle Progenitor Cells: Implications in Ischemic Treatment

Peripheral artery disease (PAD) is a common vascular disorder in the extremity of limbs with limited clinical treatments. Stem cells hold great promise for the treatment of PAD, but their therapeutic efficiency is limited due to multiple factors, such as poor engraftment and non-optimal selection of cell type. To date, stem cells from a variety of tissue sources have been tested, but little information is available regarding vascular smooth muscle cells (VSMCs) for PAD therapy. The present study examines the effects of keratose (KOS) hydrogels on c-kit+/CD31− cardiac vascular smooth muscle progenitor cell (cVSMPC) differentiation and the therapeutic potential of the resultant VSMCs in a mouse hindlimb ischemic model of PAD. The results demonstrated that KOS but not collagen hydrogel was able to drive the majority of cVSMPCs into functional VSMCs in a defined Knockout serum replacement (SR) medium in the absence of differentiation inducers. This effect could be inhibited by TGF-β1 antagonists. Further, KOS hydrogel increased expression of TGF-β1-associated proteins and modulated the level of free TGF-β1 during differentiation. Finally, transplantation of KOS-driven VSMCs significantly increased blood flow and vascular densities of ischemic hindlimbs. These findings indicate that TGF-β1 signaling is involved in KOS hydrogel-preferred VSMC differentiation and that enhanced blood flow are likely resulted from angiogenesis and/or arteriogenesis induced by transplanted VSMCs.


Introduction
Peripheral artery disease (PAD) is a common vascular disorder caused by atherosclerosis or thrombosis, which either restricts or completely blocks blood perfusion to organs or extremities [1]. The risk of PAD increases with age and approximately 4-8 million people are afflicted in the United States. Among this group, an estimated 50,000-80,000 people experience critical limb ischemia (CLI) that comes with a mortality rate of 20-25% [2]. As of today, there are no effective treatments for patients with PAD or CLI. Conventional approaches include antithrombotic medications and surgical angioplasty with or without stenting [3], but these "symptomatic" treatments are unable to fully cure patients with the late stage CLI, often necessitating limb amputation [3,4].
Stem cell treatment is an emerging approach to overcome degenerative diseases, such as PAD. To date, a range of stem cells and stem cell-derived somatic cells Benjamin T. Ledford and Miao Chen contributed equally to this work.

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have been investigated as therapeutics for PAD and CLI. Peripheral blood mononuclear cells [5], mesenchymal stem cells (MSCs) [6], embryonic stem cell (ESCs)/ induced pluripotent stem cells [7], vascular progenitor cells (VPCs) (e.g., endothelial progenitor cells [8] and vascular smooth muscle progenitor cells (VSMPCs) [9]), cord blood stem cells [10], and dental pulp stem cells [11] have all been tested in pre-clinical animal models (e.g., mouse or rat hindlimb ischemia) or in human patients with PAD or CLI, but the therapeutic efficiencies are marginal or even controversial. The reasons are not completely understood, and likely relate to the non-optimal selection of cell type, cell viability/engraftment, and administration timing/frequency/ routes. However, to our knowledge, vascular smooth muscle cells (VSMCs) -one of the major cell types comprising blood vessel walls -have been scarcely investigated to date. As yet, we have found only one study that examined the effects of VSMCs in a mouse ischemic hindlimb model [9].
Resident c-kit + (CD117 + ) cardiac stem cells (c-kit + CSCs) were originally isolated for the treatment of myocardial infarction, but their ability to generate new cardiomyocytes (CMs) in vivo has been controversial [12,13], while these cells have not been tested for treatment of PAD and CLI. Emerging evidence demonstrates that c-kit + CSCs are vasculogenic progenitor cells that promote vascularization through differentiation into endothelial cells (ECs) and VSMCs [14,15]. This finding is supported by our previously published study, in which we found that the majority (~ 72%) of human c-kit + CSCs cultured on keratose (KOS, a keratin-derived hydrogel) spontaneously differentiate into VSMCs and express α-smooth muscle actin (α-SMA) [16]. However, the mechanisms underlying this KOS-related phenomenon and whether the resultant VSMCs have potential to rescue an ischemic hindlimb in vivo are unknown. Since most of these cells become VSMCs and they lack expression of CD31, we define them in this study as c-kit + / CD31 − cardiac vascular smooth muscle cell progenitor cells (cVSMPCs).
A variety of in vitro methods have been described to induce VSMC differentiation from various types of stem cells, including small molecules (e.g. ascorbic acid [17]), extracellular matrix (ECM) proteins [18], mechanical strain [19], and growth factors (e.g., platelet-derived growth factor (PDGF)-AA, PDGF-BB, transforming growth factor beta-1 (TGF-β 1 )) [20,21]. Among these, TGF-β 1 is probably the most commonly reported differentiation inducer in VSMC differentiation from ESCs and MSCs [22,23]. It is well documented that TGF-β 1 is translated as an inactive form of pro-TGF-β 1 complex consisting of the latency-associated peptide (LAP) and TGF-β 1 , which is then packaged and processed in the rough endoplasmic reticulum and Golgi apparatus to form the small latent complex (SLC) composed of LAP + TGF-β 1 bound by non-covalent bonds [24]. The SLC is further covalently bound to the latent TGF-β binding protein (LTBP) to form the large latent complex (LLC). After being excreted out of cells, the LLC binds to fibrillin and fibronectin in the ECM [25]. The active form of free TGF-β 1 can then be released through cell-ECM mechanical stretch and/or ECM proteolysis. Subsequent binding of free TGF-β 1 to the TGF-β receptor I/TGF-β Receptor II complex (TGFBRI/TGFBRII) [25,26] activates the downstream response of Sma and Mad protein2/3 (SMAD2/3) co-SMAD canonical TGF-β 1 signaling pathway [25].
The biomaterial substrate on which the stem cells are cultured and expanded in vitro can greatly influence cell activity (e.g., quiescence, differentiation) and thus has potential to enhance stem cell-based therapies [27]. Although various natural (e.g., collagen), synthetic (e.g., polycaprolactone, polyethylene glycol), and hybrid (e.g., collagen-polyethylene glycol, chitosan-polyethylene glycol) biomaterials have been utilized in basic research, drug/cell delivery in animal models, and human clinical trials [27], keratin-based biomaterials have been attracting more attention in recent years due to their intrinsic biocompatibility, cellular regulatory capabilities, and broad availability [28][29][30], thus, providing distinct advantages over other biomaterials. For instance, KOS is not susceptible to collagenase digestion [31] and contains cell binding motifs that are not present in polysaccharide-based biomaterials (e.g., alginate or chitosan) [32]. On the other hand, compared to native keratin, KOS has a lower thiol content following oxidative chemistry [32,33]; thus, chemical crosslinking agents (e.g., genipin) are required to achieve stable hydrogels [16].
In the present study, we examined in vitro differentiation of human c-kit + /CD31 − cVSMPCs into the VSMC lineage on KOS hydrogels and to determine whether transplantation of the differentiated VSMCs could improve blood flow and rescue ischemic hindlimbs using mouse model of PAD. Specifically, we determined the effects of different basal medium [fetal bovine serum (FBS) vs the well-defined Knockout serum replacement (SR)] on cVSMPC differentiation on KOS hydrogel, collagen hydrogels, and tissue culture polystyrene (TCPS) in the presence and absence of the differentiation inducers and/or antagonists specific for TGF-β signaling. The levels of free TGF-β 1 and TGF-β 1 macromolecular complex were also evaluated prior to and post differentiation. In the animal study, cells differentiated on KOS hydrogels were compared to the cells differentiated on TCPS in an ischemic hind-limb model of immunodeficient mice and phosphate-buffered saline (PBS) vehicle was used as control. Limb blood flow, vascular network densities, arteriole size, and engraftment were analyzed before and after cell transplantation.

Isolation and Culture of c-kit + /CD31 − cVSMPCs
Human cardiac tissue samples (i.e., right atrial appendages) were originally collected as the discarded medical wastes by local hospitals under the written agreement of consent from patients. Donor confidentiality was kept only at the hospital and no patient identification information was collected. All procedures for human subject or tissue research were approved by the Institutional Review Board (IRB) and the Institutional Biosafety Committee (IBC) at Virginia Tech. The isolation and culture of c-kit + /CD31 − cVSMPCs was performed as previously described [34], except for an additional step of CD31 antibody-based magnetic activated cell sorting (Bio-Rad) followed by a collection of the CD31 negative fraction (i.e., c-kit + /CD31 − cVS-MPCs). Cells were cultured in the stem cell maintenance mediums consisting of Ham's F12 with 10% SR (SR-based medium) or 10% FBS (FBS-based medium), both with 200 µM L-glutathione, 10 ng/ml recombinant human bFGF (rhbFGF), 0.005 U/ml recombinant human EPO (rhEPO), and 1% P/S. The differentiation mediums (SR-and FBS-based) contained the same components and concentrations as maintenance mediums, but without L-glutathione, rhbFGF, and rhEPO. Depending on the purposes of each experiment, the differentiation media may contain an agonist (5 ng/ml rhTGF-β 1 ), an antagonist (500 nM A83-01 or 25 µg/ml TGF-β 1 neutralizing antibody (TGF-β 1 nAb)), and/or a differentiation inducer (10 µM 5-AZ).

Keratin Purification and KOS Hydrogel Formation
Lyophilized KOS powder was supplied by Dr. Mark Van Dyke's lab at Virginia Tech. The technical details to isolate and purify keratin/KOS from human hair can be found in our previous publications [16,33,35]. Briefly, KOS was obtained from human hair fibers following a series of extraction and purification steps. The hair fibers were washed with detergent, thoroughly rinsed, and dried. To oxidize the protein disulfide bonds, a 2% (w/v) peracetic acid solution was prepared using 2,000 ml of the solution per 100 g of fibers. The mixture was heated to 37 °C and subjected to gentle rotary stirring for 14 h. Subsequently, the fibers were recovered using a sieve and subjected to two extractions with 4,000 ml of 100 mM Tris base solution. After separating the fibers with a sieve, the extract solutions were combined, cleared of suspended particulates through centrifugation, and filtered using No. 4 Whatman filter discs (Thermo Fisher). The resulting protein solution was then purified and high-molecular-weight keratin nanomaterial was isolated through membrane filtration using a custom ultrafiltration system. A membrane cutoff of 100 kDa was employed, and a proprietary buffer solution facilitated the separation of the desired material. Following the removal of low molecular-weight contaminants, the keratin nanomaterial solution was concentrated against a dilute buffer using the same 100 kDa membrane. Finally, the solution was frozen and lyophilized to obtained as a solid product of KOS powder [16,35]. Based on sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and mass spectrometry analysis of three batches, lyophilized KOS generated with this method is highly purified (~ 99%) with a minimum variability (~ 1%) between batches [33] 1 3 KOS hydrogels were fabricated by dissolving the UVsterilized KOS (4.5% w/w) in PBS in the presence of genipin (1.5% w/w) and casting the mixture into culture plates or onto plastic coverslips to a thickness of ~ 0.5 mm [16]. Hydrogel were incubated at room temperature (RT) for 12 h to allow genipin to cross-link, washed three times with PBS (5 min each) and then incubated in PBS at RT for 48 h to remove residual genipin. Cells were plated on the top of KOS hydrogels and cultured with the stem cell maintenance medium or differentiation medium. Hydrogels used for photomicrographs were produced by making a ''sandwich'' between para-film and glass coverslips, so that removing the glass coverslip after 12 h would produce a flat surface that was ideal for cell imaging.
VSMC Differentiation from c-kit + /CD31 − cVSMPCs VSMC differentiation from cVSMPCs was conducted on two different platforms. In the TCPS-based platform, cVS-MPCs collected from the FBS-based maintenance medium were plated at a density of 6.3 × 10 3 cells/cm 2 and cultured in SR-based maintenance medium for 4 days with medium changes every other day. To induce VSMC differentiation, the SR-based maintenance medium was switched to the SRbased differentiation medium containing 5 ng/ml rhTGF-β 1 on Day 4 and maintained for an additional 28 days with medium changes every 2 days.
In the KOS hydrogel-based platform, cVSMPCs collected from the FBS-based maintenance medium were plated at a density of 1.2 × 10 4 cells/cm 2 and cultured with the SR-based maintenance medium for 4 days with medium changes every other day. To induce VSMC differentiation, the medium was changed to SR-based differentiation medium omitting rhTGF-β 1 on Day 4 and maintained for an additional 28 days with medium changes every 2 days.

Immunocytochemical Analysis of Cultured Cells
Five primary antibodies were used to identify cVSMPCs and VSMCs before and after differentiation: c-kit (also known as CD117), CD31 (also known as PECAM-1), alpha smooth muscle actin (α-SMA), calponin, and smooth muscle myosin heavy chain (SMMHC, also known as myosin 11) ( Table 1).
Due to strong autofluorescence of KOS hydrogels, the cells cultured on KOS had to be first detached from the hydrogels with elastase or TrypLE (ThermoFisher), then washed and reattached to coverslips for 2 h or directly cytospun onto slide. This step was omitted for cells cultured on TCPS surfaces. Thereafter, cells from TCPS or KOS hydrogels were fixed in 2% PFA at RT for 30 min, followed by blocking in 1.5% BSA buffer containing 0.2% Tween-20 for 30 min at RT. The resultant cells were incubated with 1:200 c-kit, 1:300 CD31, 1:300 α-SMA, 1:1000 calponin, or 1:30 SMMHC primary antibody overnight at 4 °C on a plate shaker. Cells were then washed three times with PBS (5 min each) and stained with a secondary antibody of goat anti-rabbit Alexa Fluor 488 or goat anti-mouse Alexa Fluor 594 (Table 1) at a concentration of 1:2000 in blocking buffer. An additional set was stained only with a secondary antibody as a negative control. All groups were counter-stained with Hoechst 33342 to label cell nuclei. Fluorescent images were taken using an Olympus IX73 microscope equipped with DP70 CCD camera and the software DP-BSW Version 3.0 (Waltham, MA). The fluorescent images were analyzed using NIH ImageJ (Version 2.1) to obtain the mean intensities of positive cells against the total number of nuclei. Statistical analysis and figure production were conducted using Microsoft Excel 2019 (Redmond, WA) and GraphPad Prism 5 (San Diego, CA).

Collagen Lattice Contraction Assay
Mouse collagen was prepared in the lab using the modified protocols [36]. Briefly, a total of 40 tails (~ 1 cm long in each) of non-obese diabetic SCID gamma (NSG) mice at ages of 6-12 week (Jackson Lab, Bar Harbor, ME) were collected, sectioned, and immersed in 70% ethanol for 15 min for disinfection. Tendons were extracted, transferred to ice cold 0.1% acetic acid solution at a ratio of about 100 ml of acetic acid solution per gram of tendon, and incubated for 2 days at 4 °C. Once tendons were dissolved, the mixture was centrifuged at 8800 × g for 90 min at 4 °C. The supernatant was then frozen at -20 °C for 2 h before being transferred to -80 °C freezer. On the following day, the frozen samples were lyophilized until dry.
The collagen lattice contraction assay was modified from the method published previously [22]. Briefly, cells that were differentiated either on TCPS plates in SR-based differentiation medium containing 5 ng rhTGF-β 1 or on KOS hydrogels in SR-based differentiation medium without rhTGF-β 1 were mixed with 500 µl of 3% collagen solution to get a final cell concentration of 3.75 × 10 5 cells/mL and placed in 24-well plate to allow 1 h crosslinked at RT to form stable hydrogels before being transferred into a 37 °C cell culture incubator. Additionally, collagen gels alone or gels loaded with cells from TCPS wells in the absence of TGF-β 1 were used as controls. Gel diameters were measured using ImageJ at 6, 24, and 48 h of incubation. The gel contractility was calculated as the ratio of diameter at each time point to the initial diameter (at 0 h).

TGF-β 1 ELISA Assay
Two separate experiments were performed to characterize the concentrations of TGF-β 1 in the culture medium. The first was to determine if KOS hydrogels deplete TGF-β 1 from the culture medium. To accomplish this, SR-based differentiation medium with 5 ng/mL rhTGF-β 1 (5 ng/ml) was added to KOS hydrogels and TCPS surfaces in the absence of cells. Medium was collected at 0, 12, 24, and 48 h and the levels of free rhTGF-β 1 were measured using Legend Max Frey Active TGF-β 1 ELISA Kit according to the manufacturer's protocol.
The second was to examine the rate of TGF-β 1 secretion. cVSMPCs were seeded on TCPS plates or KOS hydrogel surfaces and cultured in an SR-based maintenance medium with medium changes every 2 days. On day 4, spontaneous differentiation was induced by replacing the maintenance medium with SR-based differentiation medium in the absence of exogenous rhTGF-β 1 (denoted as Day 0), and cells were cultured for an additional 28 days with a medium change every 2 days. Samples of conditioned medium were collected prior to changing fresh medium on: 1 day before and 1 day after differentiation induction, and then every 7 days (i.e., Days -1, 1, 8, 15, 22, and 28) and TGF-β 1 was measured by ELISA.

Western Blotting
cVSMPCs were cultured on TCPS or KOS hydrogels and spontaneous differentiation was induced using SRbased differentiation medium in the absence of exogenous rhTGF-β 1 with medium changes every 2 days. For the TCPS plates, cells were washed with the cold PBS and cell lysates were collected following treatment with RIPA buffer. For the KOS hydrogel plates, cells were first detached using 7 U/ml elastase for ~ 1 min, washed once with the cold PBS to remove residual hydrogel, and then treated with RIPA buffer.
Total protein concentration in cell lysates was quantified using the Bradford assay, and then samples were mixed with 4X loading buffer with or without β-mercaptoethanol (10% β-mercaptoethanol in Laemmli buffer) at a 1:4 ratio of dilution and incubated for 5 min at 95 °C. Samples were then loaded into a 4-20% gradient SDS-stacking gel and run in a Mini Protean Tetra Cell (Bio-Rad Lab, Hercules, CA) at 100 V for 15 min, followed by 150 V for ~ 60 min. Thereafter, proteins were transferred onto PVDF membranes using a Mini Protean Tetra Cell in an ice bath at 90 mA for 16 h. PVDF membranes were blocked for 1.5 h at RT in 3% milk solution in TBST buffer on a plate rocker, and then immunoblotted with specific primary antibodies (1:100 LAP, 1:500 LTBP-1, Table 1). Beta-actin (β-actin, 1:5000) was used as an endogenous loading control. All antibodies were diluted in 1% milk solution and incubated with PVDF membranes overnight on a plate rocker at 4 °C, then washed 3 times (5 min each) with TBST buffer. The resulting membranes were then incubated with mouse or goat secondary antibodies conjugated with HRP at 1:10,000 for 2 h at RT on a plate shaker. Films were washed and treated with ECL and images were taken immediately on a Kodak Image Station 400MM (Kodak, Rochester, NY).

Inhibition of TGF-β 1 Signaling
cVSMPCs were differentiated on TCPS or KOS hydrogels using SR-based differentiation medium in the presence of 5 ng/ml TGF-β 1 (on TCPS) or in the absence of TGF-β 1 (on KOS hydrogels) and then treated with either of two antagonists specific for a TGF-β 1 signaling. The first, 500 nM A83-01 (a TGF-β 1 receptor inhibitor with IC 50 of ~ 12 nM [37]), was included in the differentiation medium throughout 28-day differentiation with medium changes every 2 days. The second, 25 µg/ml TGF-β 1 nAb, was included in the differentiation medium from Day 0 to 21 with medium changes every 2 days. Thereafter, cells were maintained in a differentiation medium in the absence of TGF-β 1 nAb with medium changes every 2 days. After 28-day differentiation (Day 28), immunocytochemistry was performed to probe for α-SMA, calponin, and SMMHC.

Surgically Induced Mouse Hindlimb Ischemia and Cell Transplantations
Forty-five adult NSG mice, ages 8-12 weeks, were used in this study. These mice were randomly divided into three groups (15 mice per group): cells differentiated on TCPS for 28-day without TGF-β 1 , cells differentiated on KOS hydrogels for 28-day without TGF-β 1 , and PBS only (control). Animals were kept in a bio-secured, environmentally controlled animal facility at the College of Veterinary Medicine of Virginia Tech. Each mouse was labeled via ear tattoos for identification [38]. The animals had access to food and water ad libitum. Cages, water, and bedding were changed weekly. Ambient temperature was kept at 22-23 °C with humidity at 50-70%. Illumination was kept at a 12 h/day-night cycle. All procedures for mouse research were approved by the Institutional Animal Care and Use Committee (IACUC) at Virginia Tech.
The surgically induced mouse hindlimb ischemic model was similar to that previously described [38,39]. Briefly, mice were anesthetized with initial 3-4% isoflurane using an EZ-7000 anesthetic system (Euthanex, Palmer, PA) and then maintained at 1-2% for the duration of the procedure. The mouse body temperature was maintained at 37 °C using a water-heated pad and monitored with a TC-1000 temperature controller (Euthanex). The left femoral artery (FA) was dissected and separated from the femoral vein and nerve, and followed by a unilateral ischemia through ligation of FA inferior to the pudendal epigastric trunk and superior to the popliteal bifurcation on the left limb using a sharp-tip Electrocoagulator (Fine Science Tools, Foster City, CA). About 2 mm of FA between both ends were removed for complete excision. Following the ligation, 10 μl of PBS (Group 1), or 10 μl of PBS containing 4 × 10 5 cells detached from TCPS (Group 2), or 10 μl of PBS containing 4 × 10 5 cells detached from KOS hydrogel (Group 3) were injected into each of 3 sites of the ischemic limbs along the sagittal plane in the middle of the gastrocnemius muscle with ~ 2 mm depth from the surface of muscle and ~ 2 mm distances between each site. The injection sites were then marked by slightly staining with green animal tattoo ink (green) to aid localization of injection sites for post-analysis. Skin incisions were then closed with a 5-0 silk suture and mice were returned to the animal facility after recovery from anesthesia. The right limb of the same surgical mouse remained intact and used as an internal control to calculate the blood flow ratios of the ischemic limb by Laser Doppler imaging [39].

Laser Doppler Imaging
A non-invasive MoorLD12-HIR Laser Doppler imager (Moor Instruments, Wilmington, DE) was used to simultaneously measure the blood flow of both limbs before (basal level) and on Days 0, 14, and 28 after surgery, as described previously [38,39]. Under anesthesia, animals were kept at 37 °C on a water-heated pad for at least 15 min before and during the imaging scanning to minimize the potential impact of body temperature on limb blood flow. A string aligned with the fourth metatarsal pad of each paw was placed across both paws as a reference line to select the region of interest (ROI) in post-analysis. The same region, size, and mean pixel intensities of all three animal groups were measured using Laser Doppler Imager Software (v 5.3, Moore Instrument). Three images were collected from the same area of hindlimb of a mouse and the measurements were averaged for statistical analysis. The blood flow was calculated as a function of image intensity and normalized to the control limb for each mouse.

Immunohistochemical Analysis of Vascular Densities in the Gastrocnemius Muscles
At the end of 28-day post-injection, animals were euthanized with CO 2 , and then processed for immunostaining and vessel painting. For immunostaining against CD31 (for ECs) and α-SMA (for VSMCs), the gastrocnemius muscles harvested after euthanasia were immediately fixed with 4% PFA, equilibrated to 30% sucrose solution, embedded in OCT freezing medium, and cryo-sectioned at 5 µm thickness. About 120 serial sections were cut and 1 out of every 18-24 sections was used for staining. Briefly, sections were blocked with 3% BSA in PBST (PBS with 0.1% Tween-20), stained with goat-anti mouse CD31 antibody (1:200) or goat-anti-mouse α-SMA (1:300) overnight at 4 °C followed by 3 washes with the cold PBS, and then incubated simultaneously with Alexa Fluor 488-conjugated chicken anti-goat secondary antibody (1:1000) and Hoechst 33342 (1:5000) for 1 h at RT.
For both immunostaining and vessel painting, sections were mounted onto slides with Vectashield anti-fade medium and coverslips were placed atop sections and sealed with nail polish. Fluorescent imaging and analysis were the same as described above.

Statistical Analysis
All experiments were repeated at least three times with over three replicates per group. All data are shown as mean ± standard error of mean (Mean ± SE) unless otherwise stated. Student's t-test with a two-tailed distribution was used to compare the two groups. Two-way analysis of variance (ANOVA) followed by the Bonferroni test was used to compare three or more groups. In all analyses, p < 0.05 was considered statistically significant. Microsoft Excel 2019 and GraphPad Prism 5 were used for statistical analysis and plotting.

KOS Hydrogel-Preferred Differentiation of VSMCs from cVSMPCs is Independent of both FBS and Differentiational Inducer (5-Aza), but is Significantly Inhibited by TGF-β 1 Antagonists
Our previous results demonstrated that the majority (72%) of c-kit + /CD31 − cVSMPCs cultured on KOS gel differentiated into the VSMC lineage following induction with the FBS-based differentiation medium containing 5-Aza [16]. To identify the factors that promoted VSMC differentiation, we first differentiated cVSMPCs on KOS hydrogel in FBS-based differentiation medium with and without 5-Aza, and compared those cells to cVSMPCs on KOS hydrogel in SR-based differentiation medium without 5-Aza (Fig. 1A). Three days after induction, cells were switched to FBS-or SR-based differentiation medium without 5-Aza and maintained additional 28 days. As shown in Fig. 1A, percentages of α-SMA + cells among the three groups were similar (p > 0.05): FBS with 5-Aza (71.6 ± 2.2%, n = 5), FBS without 5-Aza (70.5 ± 3.3%, n = 4), and SR without 5-Aza groups (71.7 ± 3.3%, n = 4). These data suggest that KOS hydrogels can induce VSMC differentiation in a defined SR medium without addition of a differentiation inducer.

Majority of VSMCs Differentiated on KOS Hydrogel are Immature, but Functionally Active
The previous data indicate that differentiation of cVSMPCs into VSMCs is preferred on KOS hydrogels. To determine the phenotype and maturity of those differentiated cells, cVSMPCs were cultured on TCPS and KOS hydrogels for 3 days in SR-based cell maintenance medium and then differentiated for 28 days in SR-based differentiation medium without any inducers (e.g., 5-Aza and rhTGF-β 1 ). Cells were stained one day (Day -1) before differentiation, and then on Days 14 and 28 of differentiation for immature (α-SMA) and mature (calponin, and SMMHC) markers of differentiation [22,[42][43][44]. The mean florescent intensities of each marker were normalized to the number of total nuclei and divided by the florescent intensity of TCPS before differentiation. The mean fold changes were plotted against the differentiation time. As presented in Fig. 3, the expression levels of all three markers were not different for cells on TCPS and KOS hydrogels prior to differentiation; however, following 14 days of differentiation, the expression levels of all markers were significantly increased, with calponin statistically different for cells on TCPS (3.24 ± 0.34) and KOS hydrogels (7.54 ± 0.61) (Fig. 3B, p < 0.001, n = 4). By Day 28, the expression levels of all three markers were significantly increased (relative to Day 14) on KOS hydrogels, but not on TCPS plates. In addition, differences in expression levels for cells on KOS and TCPS were statistically significant: 19.37 ± 1.17 on KOS vs 2.26 ± 0.61 on TCPS for α-SMA (p < 0.001, n = 4-5), 9.33 ± 1.12 on KOS vs 2.44 ± 1.39 on TCPS for calponin (p < 0.001, n = 4), and 10.86 ± 1.99 on KOS vs 1.58 ± 0.21 on TCPS for SMMHC (p < 0.001, n = 4). Together, these data suggest that the differentiated VSMCs on KOS hydrogels are still a heterogeneous mixture of mature and immature VSMCs.
Next, we employed the Collagen Lattice Contraction Assay [22] to examine the functional contractility of VSMCs. To do so, the differentiated VMSCs on TCPS with or without rhTGF-β 1 and on KOS hydrogels without rhTGF-β 1 were counted, mixed with collagen, and replated in 24-well plates. Gel diameter for each well was measured at 0, 6, 24, and 48 h, normalized to the control group (collagen gel without cells) at 0 h, and plotted as ratios against time. It was found that gel diameter in all wells gradually decreased over 48 h (sFig. 2). VMSCs developed on either TCPS with rhTGF-β 1 or on KOS hydrogels without rhTGF-β 1 resulted in reductions in the gel diameters of ~ 25% (p < 0.05, n = 6), ~ 50% (p < 0.001, n = 6), and ~ 60% (p < 0.001, n = 6) after 6, 24, and 48 h, respectively. However, the pair-wise differences between TCPS with TGF-β 1 and KOS hydrogel without TGF-β 1 were not statistically significant (sFig. 2B). In addition, although collagen gels seeded on TCPS without cells also shrank, gel diameters were significantly larger than those for cells developed on either TCPS with rhTGF-β 1 or KOS hydrogels without rhTGF-β 1 (sFig. 2B), indicating that functionally contracting VSMCs can be generated by differentiating cVSMPCs on TCPS with rhTGF-β 1 or on KOS hydrogels without addition of extraneous rhTGF-β 1 .

Collagen Extracellular Matrix Failed to Mimic KOS Hydrogel to Induce VSMC Differentiation
The capacity for KOS hydrogels to promote VSMC differentiation in the absence of inducers (e.g., 5-Aza, rhTGF-β 1 ) might stem from its ability to recapitulate the properties of the ECM. To test this, we compared VSMC differentiation on KOS hydrogels to differentiation on collagen hydrogels, collagen hydrogels cross-linked with genipin, and TCPS (in the SR-based differentiation medium without inducers). Twenty-eight days after differentiation, cells were stained for α-SMA, calponin, and SMMHC. The results showed that within 4 groups, only cells cultured on KOS hydrogels demonstrated a significant increase in expression of all 3 markers compare to three other groups (Fig. 4B): α-SMA with a ~ 8.2fold increase, calponin with a ~ 4.5-fold increase, and SMMHC with a ~ 6.7-fold increase (Fig. 4, p < 0.001, n = 4-5). In contrast, the differences among TCPS, collagen, and collagen with genipin groups were not statistically significant (Fig. 4B).

KOS Hydrogel Modulates TGF-β 1 Induction
Given the fact that the SR medium used in the present study does not contain TGF-β 1 or other growth factors [45] and that many types of cells are known to secrete TGF-β 1 [46,47], we hypothesized that the preferred VSMC differentiation of cVSMPCs on KOS hydrogels may depend on TGF-β 1 synthesis and secretion from these cells before and during differentiation. Three experiments were performed to test the possible mechanisms. First, to determine the affinity of TGF-β 1 for TCPS and KOS hydrogel surface, SR-based differentiation medium with 5 ng/ml rhTGF-β 1 was added to cell-free wells. The concentrations of free TGF-β 1 in wells were measured at different time points. It was found that the initial levels of TGF-β 1 were identical for TCPS and KOS wells (i.e., 6288 ± 35 pg/ml, n = 4) (Fig. 5A). However, within 12 h the levels decreased to around 250 pg/ml for KOS wells. Conversely, in TCPS wells, TGF-β 1 levels dropped to 944 ± 47 pg/ml within 12 h, but then gradually increased (1996 ± 40 pg/ml at 24 h and 3716 ± 38 pg/ml at 48 h), consistent with the Vroman effect [48]. Nevertheless, the significant differences in TGF-β 1 concentrations between TCPS and KOS wells (p < 0.001, n = 4) suggest that KOS exhibits a higher affinity for TGF-β 1 .
Second, to confirm that cVSMPCs and its derivatives synthesize and secrete TGF-β 1 , cVSMPCs were differentiated for 28 days on TCPS and KOS hydrogels without exogenous TGF-β 1 , and TGF-β 1 concentrations were measured by ELISA as described above. The results show that the concentrations of free TGF-β 1 in TCPS wells gradually increased over 28 days and were significantly higher than in KOS wells at each time point (Fig. 5B, p < 0.01 to 0.001, n = 4). However, the concentrations of free TGF-β 1 in KOS wells  (Fig. 5B). Together, these data confirm not only that cVS-MPCs and its derivatives secrete TGF-β 1 during differentiation but that KOS hydrogels have a higher affinity for TGF-β 1 , consistent with the cell-free rhTGF-β 1 experiment above.
Third, to verify synthesis and secretion of TGF-β 1 , we examined levels of LAP and LTBP-1 (components of the inactive TGF-β 1 complex [47]). To do so, cells cultured on TCPS and KOS hydrogels were collected and lysed on Days -1 [pre-differentiation (Pre-Diff)], 14, and 28, together with its conditional medium (that may contain secreted TGF-β 1 complex), were probed for LAP and LTBP-1 by Western blot (Fig. 5C). The blots show that the overall expression levels of both LAP and LTBP-1 decrease over time for cells on both TCPS and KOS hydrogels. However, the expression levels of LTBP-1 was significantly higher on KOS than on TCPS plates at Day -1 (Fig. 5C, p < 0.001, n = 4) and Day 14 (p < 0.05, n = 4). Collectively, these data imply that KOS might influence the dynamics of VSMC differentiation by enhancing expression and secretion of TGF-β 1 complex.

Transplantation of KOS Hydrogel-Derived VSMCs Significantly Increased Blood Flow and Vascular Densities of Ischemic Hindlimb
Based on the preceding data, it appears that KOS hydrogels facilitate preferential differentiation of cVSMPCs toward VSMCs in vitro. To examine whether these differentiated cells can modulate vascularization in vivo, we injected cells collected from KOS hydrogels into an ischemic hindlimb model and compared the extent of functional recovery to treatment with VSMCs developed on TCPS plates or no cells Laser Doppler Imager was used to evaluate the blood flow of both ischemic and contralateral control hindlimbs before [pre-ischemia (Pre-I)] and immediately after [post-ischemia (Post-I)] surgery-induced ischemia, and then after 14 and 28 days of recovery. As shown in Fig. 6, the mean ratios of blood flow (ischemic to control limb) were comparable among all three groups, both prior to and immediately following surgery. However, over the next 2-4 weeks, blood flow rates gradually recovered in the ischemic limbs for all groups. Specifically, on Day 14, the mean ratios were 0.29 ± 0.03 (n = 15) for cells on TCPS and 0.34 ± 0.04 (n = 13) for cells on KOS hydrogel, which were both significantly greater than for those limbs treated with PBS (0.21 ± 0.02, p < 0.05, n = 14). By Day 28, the mean ratios for the cells on KOS group had increased ~ 0.5 (or 50%) while the ratios for the cells on TCPS group and PBS groups increased by ~ 0.35 (or 35%) and ~ 0.25 (or 25%), respectively. The differences between cell groups (cells on TCPS vs cells on KOS) or cell groups vs PBS group were statistically significant (Fig. 6B, p < 0.001, n = 14-15).
At the end of animal protocol (28 days post-ischemia), the gastrocnemius muscles were extracted and histologically examined for evidence of revascularization by probing for α-SMA (a smooth muscle cell marker) and CD31 (an endothelial cell marker) antigens with immunostaining, and by painting functional capillaries with DiI. In the alpha (α)-SMA antibody assay, no differences were found in the control limbs of all three animal groups among three different arteriole sizes: 10-25 µm, 25-50 µm, and > 50 µm (Fig. 7, ns, n = 4-5) while in the ischemic limbs, the mean densities of α-SMA were higher for both cell groups (33.5 ± 5.85, n = 5 for cells on TCPS and 32.9 ± 6.34, n = 4 for cells KOS) relative to PBS group (15.0 ± 0.94, n = 4) in the smallest arterioles (Fig. 7, 10-25 µm, p < 0.01, n = 4 to 5). Nevertheless, differences between the two cell groups were not statistically different (Fig. 7B).
Separately, the ratio of CD31 + cell intensities between the ischemic and control limbs differed among the three groups (Fig. 8A). Specifically, while two cell groups resulted in higher ratios (i.e., more CD31 + cell intensities in the ischemic limb), only the cells on KOS group (1.82 ± 0.1, n = 4) was statistically different from the PBS group (Fig. 8A, 0.98 ± 0.13, p < 0.05, n = 5). To confirm the increase in CD31 + cells, Dil -a lipophilic fluorescent dye known to stain endothelial cell membranes specifically [49] -was perfused into the mouse hearts at the end of 28 days post-ischemia. The gastrocnemius muscles were sectioned and histologically analyzed with fluorescent microscopy. Corroborating the CD31 findings, the results show higher intensities for both cell groups (0.79 ± 0.1, n = 4, for cells on Finally, to verify engraftments of injected cells, histological sections were co-immunostained for human mitochondria (hMt) and mouse α-SMA. Similar to other published studies [50], we also found low incorporation rates for both cells on TCPS (0.07 ± 0.02%) and cells on KOS (0.09 ± 0.01%) groups. There was no statistical significance between the two groups (sFig. 3, ns, n = 4), and it was noted that few human VSMCs (hMt in green) colocalized with mouse VSMCs (α-SMA in red) (sFig. 3A). Altogether, these data suggest that the enhanced blood flow following transplantation of the cells differentiated on KOS hydrogel was probably through mediating neovascularization and revascularization of the host vascular network (especially capillaries) rather than the engraftments of transplanted cells.

Discussion
In the present study, we investigated the role of TGF-β 1 signaling in differentiation of c-kit + /CD31 − cVSMPCs on KOS hydrogels in vitro and then examined the ability of KOS hydrogel-preferred VSMCs to recover the blood perfusion of ischemic hindlimbs using immunodeficient mice. Overall, the results show that KOS hydrogel was able to induce a majority of cVSMPCs to differentiate into functionally contractile VSMCs in the defined SR medium without addition of any exogenous growth factors or chemical differentiation inducers. This unique effect on KOS hydrogels could not be replicated on collagen hydrogels, but could be significantly inhibited by TGF-β 1 antagonists. Together these indicate that KOS hydrogel may modulate the availability and activation of free TGF-β 1 to promote VSMC differentiation. Importantly, KOS-preferred VSMCs significantly increased the blood flow of ischemic hindlimbs, but the mechanisms underlying this beneficial effect remain unknown, probably relating to the paracrine effects of transplanted VSMCs on angiogenesis and/or arteriogenesis (as opposed to engraftment into developing vessels). To the best of our knowledge, this is the first study to demonstrate the unique biological property of KOS hydrogel in promoting VSMC differentiation from cVSMPCs through involvement of TGF-β 1 signaling and the therapeutic potential of VSMCs for the treatment of ischemic limbs.
Stem cell-derived somatic cells, such as cardiomyocytes, have shown a great promise for basic research, drug screening/toxicity testing, and cell-based therapies for various diseases, such as myocardial infarction [51]. In the past two decades, regarding to generation of VSMCs, several protocols have been developed to produce VSMCs [52,53] from various types of stem cells, such as mesenchymal stem cells [54] and pluripotent stem cells [55]; however, all these methods utilized mediums containing growth factors (e.g., EGF, TGF-β 1 ), chemical inducers (e.g., 5-Aza), and/or differentiation facilitators (e.g., Vitamin C). To date, only one study has shown progenitor cell (c-kit + /Scal1 + ) differentiation into VSMCs without inducers [56], but they performed their culture on decellularized vascular scaffolds.
In contrast, the present study demonstrates a simple but unique approach to directly differentiate cVSMPCs into VSMCs on KOS hydrogel in the defined SR medium without addition of any growth factors or differentiation inducers (Figs. 1, 2, and 3). The resultant VSMCs exhibited VSMC markers (Fig. 3) and the ability of functional contraction (sFig. 2), typical phenotype of VSMCs documented by other laboratories [57,58]. While it is true that only ~ 72% of cells on the current KOS hydrogel culture system expressed the standard markers of VSMC phenotype, higher yields and/or purities can be achieved by antibody-based cell sorting or optimization of the protocol in future study. Despite this, our method is simple, cost-effective, and suitable for scale-up to produce large numbers of clinical-grade VSMCs using xenofree medium, thus offering a new avenue in manufacturing VSMCs for research and clinical communities.
TGF-β 1 is a multifunctional cytokine involved in the regulation of proliferation, differentiation, migration, and survival of various types of cells [59,60]. For VSMCs, TGF-β 1 is known to play a pivotal role in modulating differentiation and controlling phenotypic switching between contractile and synthetic states [61]. In the present study, we found that 1) addition of exogenous TGF-β 1 to defined SR medium induces VSMC differentiation on TCPS plates to a similar extent as the cells on KOS hydrogels without inducers (TGF-β 1 or 5-Aza) (Figs. 1 and 2); 2) VSMC differentiation on KOS hydrogels (without inducers) is significantly inhibited by TGF-β 1 antagonists ( Fig. 2 and sFig. 1); 3) cVSMPCs and/or their derivatives (i.e., VSMCs) secrete TGF-β 1 , LAP, and LTBP-1 (components of the inactive TGF-β 1 macromolecular complex) (Fig. 5); and 4) KOS hydrogel can bind free TGF-β 1 to activate TGF-β 1 signaling (Fig. 5). These data indicate that KOS hydrogel is able to promote VSMC differentiation in the absence of differentiation inducers, but the underlying mechanism remains to be investigated. It is possible that the affinity of KOS to TGF-β 1 not only affects availability of free TGF-β 1 but also influences the release of active TGF-β 1 from the latent complex.
TGF-β 1 -LAP is believed to be stored in the ECM together with LTBP-1 as a macromolecular complex following secretion by cells [62]. Two mechanical activation models of TGF-β 1 complex have been proposed: cell contractiontriggered integrin activation and ECM-strain-dependent activation [63]. The former model postulates that the actin/ myosin-mediated cell contraction can exert a strain on the arginine-glycine-aspartate (RGD) (integrin) binding site of LAP to induce a conformation change in the TGF-β 1 complex, thus releasing free TGF-β 1 . In contrast, the latter model proposes that the tensile strain of the ECM has to be induced (probably by cells) to trigger release of the active TGF-β 1 , which explains why the TGF-β 1 complex must bind to the ECM to be activated [63]. However, it is unknown how KOS interacts with the TGF-β 1 complex and the integrin adhesion receptors involved in contracting the ECM. Future studies are necessary to examine these potential interactions.
In parallel with the above, numerous studies have found that the ECM composition and structure also affect TGF-β 1 activation, and that the TGF-β 1 complex mainly binds to the fibrillar proteins of ECM [63]. Being one of the major components in the fibrillar ECM, the type I collagen (the primary protein in rat tail tendons)-based hydrogels with or without a cross-linker of genipin were directly compared with KOS hydrogels to examine their ability to promote VSMC differentiation. The rat tail collagen hydrogels were selected because the collagen type I exists in the ECM of vascular tissue and plays a critical role in VSMC differentiation by concentrating TGF-β 1 at the cell surface [58]. Genipin was added to increase the mechanical properties of collagen hydrogels, and omitted to eliminate the possibility of covalently crosslinking growth factors to the hydrogels, which might create locally high concentrations of growth factors like TGF-β 1 [64,65]. Nevertheless, our results demonstrated that KOS (but not collagen) hydrogels possess the appropriate biological and/or mechanic properties to promote VSMC differentiation.
One of the ultimate goals of cell therapy is to restore the normal cellular structure and functionality of impaired organs. As such, we examined the therapeutic effects of transplanted cells from KOS hydrogel in a mouse hindlimb ischemia model compared to cells from TCPS and PBS vehicle. We found that 1) VSMCs differentiated on KOS hydrogels significantly increased blood flow to the ischemic limb as compared to other groups (Fig. 6); 2) vascular densities evaluated by VSMC (α-SMA) and EC markers (CD31, Dil dye) were also higher for the cells on KOS group than for other groups (Figs. 7 and 8). However, the engraftments of transplanted cells in both cell-treated groups were very low and there was no statistically significant difference between two cell-treated groups (sFig. 3). The causes for low engraftments are not completely understood; but it is commonly believed to be related to acute inflammation, immune response, lack of local blood supplies and appropriate exchanges of O 2 /CO 2 and of nutrients/metabolic wastes [66,67]. Nevertheless, given that too few implanted cells were identified to demonstrate a process of engraftment, we postulate that the enhanced blood flow was likely due to increased angiogenesis/arteriogenesis trigger by paracrine factors from the injected cells.
ECs and VSMCs are the two main cell types of the vascular wall. Notwithstanding, in the past decade, ECs and endothelial progenitor cells have been widely studied in the hindlimb ischemic models of rodents [5,68] while VSMCs (the focus of this study) have barely been investigated [69,70]. For example, Foubert et al. reported that coadministration of both endothelial progenitor cells and smooth muscle progenitor cells dramatically increased capillary and arteriolar densities and foot blood perfusion in mouse hindlimb ischemia [9], but administration of smooth muscle progenitor cells alone did not show a significant increase of blood flow compare to coadministration of both cell types. The present study only injected the differentiated VSMCs, and the outcomes were significant not only in blood flow (Fig. 6) but also in vascular densities (Figs. 7 and 8). Together, these data suggested that the beneficial effects of transplanted KOS cells could not be simply ascribed to the paracrine mechanism alone [71] given the administration of cells from TCPS group.

Conclusion
In summary, we demonstrated that KOS hydrogels supported VSMC differentiation of c-kit + /CD31 − cVSMPCs in the defined SR medium without addition of exogenous growth factors or differentiation inducers. The underlying mechanism is not completely understood, but at least partially involves TGF-β 1 synthesis and signaling. Adhesive interactions between KOS hydrogel and cells and affinity of TGF-β 1 for the KOS may also play important roles in stimulating differentiation. More importantly, the resultant functional and contractile VSMCs, which can be developed by simply culturing on KOS hydrogels, increased the blood flow of mouse ischemic hindlimbs following transplantation. The increased angiogenesis and/or arteriogenesis observed in vivo may be triggered by paracrine effects of the transplanted cells, which need to be further investigated. In addition, a number of other critical questions remain to be answered: Does KOS hydrogel alone or together with other cells types (e.g., ECs) have any significant beneficial or adverse effects in ischemia? Will these effects (if any) take place by promoting or inhibiting angiogenesis/arteriogenesis or blocking local microcirculation? In the long-term, the potential translational significance from the present findings could be high. The dramatic effect of KOS hydrogel in defined SR medium on VSMC differentiation suggests a possibility that KOS-promoted VSMC differentiation may be triggered probably by a single signaling pathway that does not require multiple growth factors and cytokines existing in FBS? This novel KOS hydrogel-based cell culture method (without addition of various growth factor and FBS) can replenish an additional new but simple platform to produce a large amount of VSMCs, which can be used not only in basic research, drug screening, and toxicity testing, but also in vascular cell-based therapies to treat patients with ischemic diseases, PAD and heart failure.