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Prominent vascularization capacity of mesenchymal stem cells in collagen-gold nanocomposites Shuchen Hsieh, Hui-Jye Chen, Shan-hui Hsu, Yi-Chun Yang, Cheng-Ming Tang, Mei-Yun Chu, Pei-Ying Lin, Ru-Huei Fu, Mei-Lang Kung, Yun-Wen Chen, Bi-Wen Yeh, and Huey-Shan Hung ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.6b09330 • Publication Date (Web): 07 Oct 2016 Downloaded from http://pubs.acs.org on October 9, 2016

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ACS Applied Materials & Interfaces

Prominent vascularization capacity of mesenchymal stem cells in collagen-gold nanocomposites

Shu-Chen Hsieh1,2,3,#, Hui-Jye Chen4,#, Shan-hui Hsu5,6, Yi-Chun Yang7,Cheng-Ming Tang8, Mei-Yun Chu4, , Pei-Ying Lin1, *4,12

Ru-Huei Fu9, Mei-Lang Kung1, Yun-Wen Chen10, Bi-Wen Yeh11, Huey-Shan Hung 1

Department of Chemistry, National Sun Yat-Sen University, Kaohsiung, Taiwan, R.O.C.

2

Center for Stem Cell Research, Kaohsiung Medical University, Kaohsiung, Taiwan, R.O.C.

3

School of Pharmacy, College of Pharmacy, Kaohsiung Medical University, Kaohsiung, Taiwan, R.O.C.

4

Graduate Institute of Basic Medical Science, China Medical University, Taichung, Taiwan, R.O.C.

5

Institute of Polymer Science and Engineering, National Taiwan University, Taipei, Taiwan, R. O. C.

6

Rehabilitation Engineering Research Center, National Taiwan University, Taipei, Taiwan, R. O. C.

7

Department of Neurosurgery, Taichung Veterans General Hospital, Taichung, Taiwan, R.O.C.

8 9

Institute of Oral Sciences, Chung Shan Medical University, Taichung, Taiwan, R.O.C. Graduate Institute of Immunology, China Medical University, Taichung, Taiwan, R.O.C.

10

Department of Pharmacology, National Cheng Kung University, Tainan, Taiwan, R.O.C.

11

Department of Urology, Kaohsiung Medical University Hospital, Kaohsiung Medical University, Kaohsiung,

Taiwan, R.O.C. 12

Center for Neuropsychiatry, China Medical University Hospital, Taichung, Taiwan, R.O.C.

#

These two authors contributed equally to this article.

Address correspondence to: Huey-Shan Hung, Graduate Institute of Basic Medical Science, China Medical University, Taichung, Taiwan, R.O.C. Tel: 886-4-22052121 ext 7827; Fax: 886-4-22333641; E-Mail: [email protected]

ACS Paragon Plus Environment


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Abstract The ideal characteristics of surface modification on the vascular graft for clinical application would be with excellent hemocompatibility, endothelialization capacity, and anti-restenosis ability. Here, Fourier transform infrared spectroscopy (FTIR), surface enhanced raman spectroscopy (SERS), atomic force microscopy (AFM), contact angle (θ) measurement, and thermogravimetric analyzer (TGA) were used to evaluate the chemical and mechanical properties of collagen-gold nanocomposites (collagen+Au) with 17.4, 43.5 and 174 ppm of Au and suggested that the collagen+Au with 43.5 ppm of Au had better biomechanical properties and thermal stability than pure collagen. Besides, stromal-derived factor-1α (SDF-1α) at 50 ng/ml promoted the migration of mesenchymal stem cells (MSCs) on collagen+Au material through the α5β3 integrin/endothelial oxide synthase (eNOS)/metalloproteinase (MMP) signaling pathway which can be abolished by the knockdown of vascular endothelial growth factor (VEGF). The potentiality of collagen+Au with MSCs for vascular regeneration was evaluated by our in vivo rat model system. Artery tissues isolated from implanted collagen+Au-coated catheter with MSCs expressed substantial CD-31 and α-SMA, displayed higher anti-fibrotic ability, anti-thrombotic activity as well as anti-inflammatory response than all other materials. Our results indicated that the implantation of collagen+Au-coated catheter with MSCs could be a promising strategy for vascular regeneration.

Keywords: Mesenchymal stem cells; collagen-gold nanocomposites; stromal-derived factor-1α; vascular endothelial growth factor; vascular regeneration

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Introduction Blood vessel occlusions are often treated clinically by implantation of vascular stents or grafts. Restenosis and thrombosis, however, do frequently occur after implantation of these devices1,2. To tackle this problem, novel vascular biomaterials are developed and used for the fabrication of vascular implants to prevent immune response, thrombus and intimal hyperplasia, and thus lessen the highly prevalent restenosis3. Functional vascular grafts should have good suture retention, withstand physiological forces associated with the blood flow, and maintain the structural integrity during neotissue formation4. The inherent properties of the vascular graft to interact with proteins can affect the endothelialization process within the graft5. Meanwhile, the anti-CD34-coated stents accelerate the in stiu endothelialization6. For a cardiovascular implant, endothelialization is important for the long-term success of the implant owing to the natural anti-coagulant property of the endothelial cells. On the other hand, surface modification of the vascular graft is critical for promoting the endothelialization and reducing the platelet aggregation in the graft7. Nanogold exhibits excellent biocompatibility because of its innate properties such as anti-thrombosis, low immune responses and little cytotoxicity. Literatures indicate that many polymers such polyurethane, fibronectin and collagen, when added with nanogold, can interact with the nanogold to create nanotopographical surface patterns8-12. Therefore, surface modification by the polymer nanogold composites would generate nanotopography and biomimetic interface, which consequently change the cell behavior. Our previous study indicated that synthetic polyurethane nanogold composite-coated catheter induced the differentiation of endothelial progenitor cells (EPCs) into endothelial cells that subsequently promoted endothelialization and reduced thrombosis13. The nanogold composites may trigger nitric oxide (NO)-dependent pathway [endothelial nitric oxide synthase (eNOS)/phosphoinositide 3

kinase

(PI3K)/Akt]

and

NO-independent

pathway

[α5β3

integrin/focal adhesion

kinase

(FAK)/Rho-GTPase/matrix metalloproteinases (MMP)] through cell-material interaction, leading to cell adhesion, migration and proliferation8-12. In addition, nanogold composites have been shown to induce the 2



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migration and differentiation of stem cells including mesenchymal stem cells (MSCs)14 and endothelial progenitor cells (EPCs)13 partially via stromal-derived factor-1α (SDF-1α)/CXCR4 activation. Collagen is the major composition of connective tissue and is the most popular biomaterials for medical applications because of the excellent biocompatibility15. Collagen is considered as a potential natural matrix for tissue engineering applications16. Coating collagen on the surface of synthetic vascular grafts could enhance the attachment and growth of endothelial cells17. Biodegradable polymer nanofiber meshes coated with collagen could enhance the endothelialization and stem cell differentiation into endothelial cells, which is a promising way to prevent intimal hyperplasia after implantation of small-diameter vascular grafts18. In particular, our recent in vitro study has demonstrated that collagen-nanogold composites supported the extracellular matrix cues for MSC migration and differentiation12. EPCs are a source of cells for vascular tissue engineering19. However, their proliferation ability is usually low. Meanwhile, MSCs with high proliferation ability have been considered as alternatives to EPCs. MSCs secret various growth factors and cytokines that promote tissue regeneration as well as induce recruitment and differentiation of stem cell-like progenitors20-27. For instance, SDF-1α secreted by MSCs promotes the survival and proliferation of MSCs through an autocrine or paracrine mechanism28. Vascular endothelial growth factor (VEGF) and its receptor (VEGFR) are crucial for cell growth and differentiation during the development of heart and blood vessels29. Exogenous expression of VEGF promotes myocardial repair at least in part through SDF-1α/CXCR4-mediated recruitment of MSCs30. When MSCs were seeded on electrospun poly (L-lactic acid)/collagen nanofibers that mimics the environment of native vascular extracellular matrix (ECM), they could differentiate into vascular cells. Therefore, MSCs have the potential to repair the injured vascular tissues31. So far, there is no ideal combination of material chemistry and architecture being identified to satisfy the required biocompatibility of a vascular device. Improving the long-term efficiency of a vascular device has become a major challenge32,33. Previously, our in vitro study has demonstrated that MSCs 3


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cultured on the collagen-nanogold (Col+Au) composites had a greater capacity for endothelial differentiation12. Here in this study, we aimed to create a microenvironment for the neo-vascularization in vivo through the use of this biologically compatible material. We found that MSCs filled into the Col+Au-coated catheter underwent endothelialization and repaired the damaged vascular tissue through the activation of the SDF-1α/VEGF signaling pathway.

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Materials and methods 2.1. Preparation of collagen-nanogold composites Nanogold particles (Au) were prepared from bulk gold which was atomically vaporized by an electrically gasified method under vacuum, and subsequently accumulated in a cold trap system and collected by centrifugation34. Finally, the obtained negatively charged Au is resuspended in distilled water without using any surfactant or stabilizer. The size of Au, calculated to be ~5 nm, was controlled by the evaporation time and electric current. To prepare collagen solution, type I collagen (BD Bioscience, USA) was dissolved in 5 mM of acetic acid at the concentration of 1 mg/ml at 4oC for overnight. The solution was further diluted 1:1 with 2×PBS, resulting in a 0.5 mg/ml of collagen solution at pH 7.3, and heated to 21oC in a water bath. The temperature of the bath was increased to 37oC for 30 min and then returned to 4oC for storage. To generate the collagen+Au composites, Au were added to the collagen solution so that the final collagen+Au samples contained 17.4, 43.5 and 174 ppm of Au. Collagen and collagen+Au were coated onto the culture dish, culture plate, or 15 mm round coverslips inside a dish at the amount of 20 µg/cm2 to form a thin film of collagen and collagen+Au.

2.2. Characterization of collagen-nanogold composites films Transmission Fourier transform infrared (FTIR) spectra were acquired (8 cm-1 resolution 256 scans with the sample compartment vacuum pressure set at 0.12 hPa) using a Bruker 66 v/s FTIR spectrometer. Double side polished silicon (100) wafer substrates were cut into 20×20 mm2 pieces with a diamond-tipped stylus. Spectra from a freshly plasma-cleaned silicon wafer sample were collected before each measurement to obtain the background spectrum. Raman experiments were acquired on a Raman microscope (LABRAM HR UVVIS-NIRVersion) system using a 633 nm laser with the incident laser power kept at 20 mW. A holographic grating (1800 grooves-mm-1) and a 1024×3256 pixel CCD detector with total accumulation times of 60 s were

5


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employed. The as-prepared collagen without or with Au at different concentrations (17.4, 43.5, and 174 ppm), were deposited on a silicon substrate for Raman analysis. X-ray photoelectron spectroscopy (XPS) was used to investigate the elemental composition of collagen and collagen fabricated with different concentrations of Au, by a JEOL JPS 9010 MX equipped with a monochromatic Mg Kα X-ray radiation source. The as-prepared collagen with or without Au at different concentrations (17.4, 43.5, and 174 ppm) were layered onto clean silicon substrates at room temperature for analysis. Samples were prepared for AFM (MFP-3D™, Asylum Research, Santa Barbara, CA) experiment by drop-casting 100 µl of samples onto clean Si (100) wafer substrate then air-dried. AFM experiments used a silicon cantilever (Olympus AC240TS) with a spring constant of 2.0 N/m under ambient condition and topographical images were acquired in AC mode with an image resolution of 512 × 512 pixels. The AFM results were checked for reproducibility on three different areas of the sample and at different scan sizes. A thermogravimetric analyzer (TGA2050, TA Instruments) was used for the thermogravimetric analysis (TGA). 5 mg of samples was placed in a platinum crucible and heated at a rate of 10°C/min under nitrogen. The pyrolytic temperatures (Tonset and Tp) were obtained from the TGA curves. Water contact angle of samples was measured by a PGX model instrument at room temperature by dropping 2.5 µl of Mili-Q reagent-grade (type I) water (18.2 MΩ‧cm at 25oC) onto the surface of the prepared samples. The value of contact angle was the mean of five measurements. AFM (MFP-3D™, Asylum Research, Santa Barbara) was used for the Young’s modulus analysis. Samples were prepared by casting 100 µl of samples onto clean cover glass substrate and then air-dried. For all Young’s modulus measurements in AFM, a silicon cantilever (Olympus AC240TS) with a spring constant 2.47 ± 0.36 N/m (calibrated using the thermal method) was used under ambient condition35,36. The elastic properties of samples can be derived from the AFM force curves using a loading force about 20nN. Two hundred force curve measurements were made on each sample and the average value was

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obtained. We used the DMT contact model for this experiment, where E is Young’s modulus, Ʊ is the Poisson ratio of the sample which is assumed to be 0.5, and w is the work of adhesion.

2.3. Culture and characterization of mesenchymal stem cells The mesenchymal stem cells (MSCs), kindly provided by Prof. Woei-Cherng Shyu, were isolated from Wharton’s jelly tissue of human umbilical cord37. These MSCs were cultured in Dulbecco’s Modified Eagles’s Medium (DMEM) with high glucose containing 10% FBS, 100 U/ml penicillin/streptomycin, 1% sodium pyruvate, and maintained in a humidified growth chamber with 5% of CO2 as previously described14. For the characterization of MSCs, antibodies against the stem cell markers including CD14, CD29, CD34, CD44, CD45, CD73, CD90 and CD105 were used. MSCs were stained with these antibodies that were pre-conjugated with either fluorescein isothiocyanate (FITC) or phycoerythrin (PE) (BD, Pharmingen, USA) and analyzed by flow cytometry (LSR II, Becton Dickinson, USA). FITC-conjugated IgG1 and PE-conjugated with IgG1 (BD, Pharmingen, USA) were used as the control antibodies.

2.4. VEGF siRNA transfection, MTT assay, and cellular uptake measurement MSCs (5×105/well) were plated into each well of six-well plates to reach 80%∼90% of confluences. Culture medium was replaced with OPTI-MEM I (Invitrogen) and cells were transfected with VEGF siRNA in an appropriate ratio with Lipofectamine 2000 reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s instructions. MTT assay was used to measure the cell viability. MSCs (1×104/well) were seeded into a 96-well plate and transfected with VEGF siRNAs or various concentrations of scramble siRNAs (6.25 nM, 12.5 nM, 25 nM and 50 nM) using Lipofectamine 2000 reagent

according

to

the

company’s

brochure.

After

48

h

of

incubation,

MTT [3-(4,

5-dimethylthiazol-2-yl)-2, 5-diphenyl tetrazolium bromide] (0.5 mg/ml; Sigma) solution was added into each well and incubated for additional 2 h at 37oC. Dimethyl sulfoxide was added to dissolve the 7


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formazan crystals and the absorbance was measured at 570 nm by a Microplate reader (SpectraMax M2e, Molecular Devices, USA). To quantify the cellular uptake of FITC-conjugated scramble siRNAs over time, MSCs (2×105/well) were seeded into a 6-well plate, and transfected with siRNA molecules for 2h, trypsinized and analyzed by using FACS Calibur flow cytometer (Becton Dickinson, USA). The data of FITC-fluorescein positive cells were further analyzed using the FACS software.

2.5. Immunofluorescence staining of eNOS, CXCR4, α5β3 and CD31 Cells (2×104 cells/well) were seeded onto 15-mm coverslips pre-coated with collagen or collagen+Au in a 24-well plate, and incubated in the condition medium as previously described12. MSCs were transfected with VEGF siRNAs (25 nM), and/or treated with SDF-1α (50 ng/ml). Immunofluorescent staining was described previously12. Briefly, cells were fixed, permeabilized, and incubated with eNOS antibody (1:300 dilution, Santa Cruz), CXCR4 antibody (1:300 dilution, Santa Cruz), α5β3 antibody (1:300 dilution, Santa Cruz) and CD31 antibody (1:300 dilution, Santa Cruz). After three washes, cells were incubated with the FITC-conjugated secondary antibody (1:300 dilution) for 60 min. Cell nuclei were stained with DAPI (Invitrogen) (1 µg/ml) for 10 min. After extensive washes, samples were mounted on microscope slides with the storage solution (50% of glycerol in PBS) and sealed with a synthetic mount.

2.6. Migration ability of MSCs We used the Oris(TM) Cell Migration Assay reagent kit (Platypus Technologies, Madison WI, USA) to assess the cell migration38. Collagen or collagen+Au pre-coated coverslips were placed into each well of an Oris(TM) Cell Migration Assay Tri-Coated plate. Cell seeding stoppers (2 mm in diameter) were inserted into each well on top of coated materials to prevent the cell migration into the central region of the well. MSCs (8×103 cells/well) were seeded into each well and incubated for 48 h to reach full confluence. Cells were transfected with VEGF siRNAs (25 nM) and/or treated with SDF-1α (50 ng/ml) 8



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for 48 h.

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The stopper was then removed from the test wells, but remained inside the reference wells.

The plate was further incubated at 37oC for observation of pre-migration (t=0 h) and post-migration (t=48 h) states. Cell populations at the end point were stained with Calcein AM (Sigma) (200 µl at 2 µM) in serum-free medium and the images were recorded by a fluorescence microscope (Zeiss Axio Imager A1, USA). Cells that migrated into the detection zone were quantified by measuring the fluorescence intensity and data were analyzed by Image Pro Plus 5.0 software.

2.7. MMP zymography assay Gelatin zymography was used to examine the expression of matrix metalloproteinases (MMP). MSCs were transfected with VEGF siRNA (25 nM), and/or treated with SDF-1α (50 ng/ml) for 48 h. Culture media were collected for the assay without prior thermal denaturation. Gel electrophoresis and the resulting activity assay were as described previously12. The resulting gel was then digitalized by scanning in a densitometer and analyzed by Image Pro Plus 5.0 software (Media Cybernetics). Data were normalized by the amount of total protein measured in culture medium.

2.8. Surface fabrication of collagen and collagen+Au on catheters The outer surface of intravascular catheters (22GA: 2.5 cm long) (InsyteTM AutoguardTM, BD Medical, USA) were activated by air plasma using an open air plasma system (Plasmatreat, Germany) as described previously13. Briefly, dried air (21% oxygen and 79% nitrogen) was ejected from a rotating nozzle hooked to a plasma generator. The air pressure and plasma power were set in 2.5 kg/cm2 and 1000 W. Next, catheters were put on a distance of 10 mm from the nozzle and subsequently scanned at 15 m per min. After the air plasma activation process, catheters were immediately immersed in the collagen or collagen+Au 43.5 solution for 10 s. Finally, samples were incubated and dried in an oven at 60 oC for 30 min.

2.9. Platelet activation assay 9


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Collagen and collagen+Au 43.5 were coated on coverslip glass and were put in 24-well culture plates where 0.5 ml of plasma-rich plasma (2106 platelets/ml, from Taichung Veterans General Hospital, Taichung, Taiwan, IRB-TCVGH approval number CE12164) was added in each well. Samples were removed after incubation for 1 h and the number of adherent platelets was counted. After this process, samples were fixed by 2.5% glutaraldehyde (Sigma, USA)/phosphate buffered saline (PBS) (pH = 7.4) and subsequently dehydrated in ethanol solutions (increasing concentration from 30% to 100%), critical point-dried, and sputter coated with gold. Finally, samples were examined by a scanning electron microscope (SEM) (JEOL JSM-6700 F, Japan)11.

2.10. Implantation of MSCs on collagen or collagen+Au-coated catheter To access the attachment efficiency of MSCs onto to the catheter (2.5 cm long). MSCs were labeled with 10 nM of red-fluorescent quantum dots (QDs; Qtracker 655 cell labeling kit, Invitrogen, USA) according to the manufacturer’s protocol with some modification. Labeled cells were deposited onto the external surface of catheters, and the attachment ability was examined under the fluorescence microscope. For the implantation of catheters into the animals, 3 ml of MSCs (1×106 cells/ml) were filled into the coated catheter sheath and rotated in a 3D shaker (Gene Pure, GDS100-1) at 37oC for 48 h before implantation. Prepared catheters were implanted into the femoral artery of the Sprague-Dawley rats (250 to 300 g). Experimental animals were divided into four groups: group 1 with blank catheters (catheters without any treatment); group 2 with catheter seeded with MSCs; group 3 with collagen-coated catheter seeded with MSCs (collagen-coated catheter/MSCs); group 4 with collagen+Au-coated catheter seeded with MSCs (collagen+Au-coated catheter/MSCs). The detail procedures for rat anesthetization and operation for catheter setting were described elsewhere13. All tissue specimens were obtained for analyses after 4 weeks of catheter implantation. All the protocols for animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of the China Medical University.

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2.11. Histological and immunohistochemical analyses 4 weeks after implantation, harvested specimens were fixed with 10% of formaldehyde, dehydrated in a serial ethanol solutions, and then embedded in paraffin. The specimens were sectioned (in 4 µm thickness), stained with hematoxylin and eosin (H&E) (Sigma, USA). Sections were also subjected to immune-histochemical staining using the primary antibodies including CD-31 antibody (DAKO, USA) and alpha-smooth muscle actin (anti-α-SMA) antibody (Abbiotec, LLC, USA), and sequentially with FITC- and Cyc5.5-conjuagted secondary antibodies. Native femoral artery from normal animals served as the positive controls.

2.12. Masson’s Trichrome staining We used Masson's trichrome staining to examine the fibrosis of arterial tissues due to collagen deposition. Collagen distribution and deposition were revealed as blue color precipitation in Masson's trichrome staining by using a Masson's trichrome staining kit (Sigma, USA). The area of fibrosis in arterial tissue sections was calculated by Image J 4.5 version software (Media Gybertics). For each animal, randomly selected high-power fields (HPFs) from three selected sections were quantified and analyzed. The number of pixels calculated from the three HPFs were finally summed up.

2.13. Statistical analysis Data from multiple samples (n=3∼6) were collected for a given experiment and expressed as mean ± standard deviation. All experiments were independently repeated for at least three times to assure the reproducibility of the results. Single-factor analysis of variance (ANOVA) method and the followed Bonferroni post hoc analysis were employed to evaluate the statistical significance of the results and p values less than 0.05 were regarded as significant.

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3. Results 3.1 Characterization of the prepared collagen-nanogold composites Nanogold (Au) possess excellent biocompatibility owing to its inert property. Different surface modifications on nanogold have been shown to control different biological processes such as cell attachment, growth, migration or differentiation8-12. Further surface modification of collagen on vascular implants could promote the healing as well as endothelialization and prevent intimal hyperplasia33. MSCs has been applied for the regeneration of damaged blood vessels due to their high proliferation capacity27 and ability to differentiate into endothelial cells26. In addition, MSCs seeded onto the electrospun poly(L-lactic acid) (PLLA)/collagen nanofibers that mimic the native vascular extracellular matrix (ECM) environment can be differentiated into vascular cells and be promising for the repair of vascular tissues31. Similarly we have prepared collagen-nanogold (collagen+Au)-coated catheter and expected that this fabrication would mimic the native vascular ECM environment for MSCs to perform vascular regeneration. The identity of collagen+Au was characterized by Raman microscopy, Fourier transform infrared (FTIR) spectrometry, and X-ray photoelectron spectroscopy respectively. As shown in Figure 1A, peaks at 1,255 and 1,688 cm-1, which correspond to amide III (C=N bond) and amide I (C=O bond) bands respectively were observed in the Raman spectra of pure collagen and different amounts of Au in collagen+Au composites39. The peak appeared at 3340 cm-1 is attributed to N-H stretching of collagen. New peaks at 1582 cm-1 (correspond to the phenylalanine ring vibrations) and 1350 cm-1 (attribute to the CH2 and CH3 stretching) were arisen as the Au conjugated to collagen40. The intensities of Raman signals were increased with the increase in the concentration of Au, indicating that Au induces the aggregation of collagen. The characterization of the collagen+Au composites by Transmittance Fourier transform infrared spectrometry analysis was shown in Figure 1B. The peaks at 1655 cm-1 and 1540 cm-1 correspond to the amide I and amide II bands respectively12. The peak at 3340 cm-1 comes from the NH stretching12. These data indicated the presence of collagen in the collagen+Au composites. Besides, there was a slight shift in 12



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the spectrum of the peak at 1540 cm-1 (pure collagen; N-H band) to 1535 cm-1 (collagen+Au with 174 ppm of Au) (Figure 1B), suggesting the incorporation of Au into collagen. The surface chemistry of collagen-nanogold composites analyzed by X-ray photoelectron spectroscopy was shown in Figure 2. Survey scan of the pure collagen and collagen-nanogold composites was shown in Figure 2A. Generally, the obtained gold peak is a doublet, with one peak situates around 84 eV (Au 4f7/2) and the other situates around 87 eV (Au 4f5/2) in the derived XPS spectra. However, the gold peak doublet shifted to about 87.2 eV and 91.2 eV in the derived XPS spectra of collagen + Au 174 ppm, indicating the conjugation of collagen to Au in the collagen/nanogold composite (Figure 2B). As shown in Figure 2C and Figure 2D, the XPS spectra of C1s from collagen and collagen+Au (174 ppm) peaked at 288.2, 289.4 and 290 eV which indicate the C–C, C–N and C–O bands of collagen individually41.

3.2 Biomechanical properties of collagen+Au composites To investigate the biomechanical properties of collagen and collagen+Au materials, Young’s modulus analysis was conducted. The elasticity maps of different materials were shown in Figure 3A. After calculation, the value of Young’s modulus was 181.3±71.1 MPa for collagen+Au (17.4 ppm), followed by 202.5±73.4 MPa for pure collagen, 459.9±158.2 MPa for collagen+Au (43.5 ppm) and 520.1±188.6 MPa for collagen+Au (174 ppm), indicating that the elasticity decreased with the increase in the concentration of nanogold in the collagen+Au composites. To further investigate the roughness of sample surface on different materials, atomic force microscopy (AFM) was used. A series of topographical and 3D images of collagen and collagen with different concentrations of Au were shown in Figure 3B. As compared with pure collagen, collagen containing 17.4 ppm of Au resulted in thick fibrils, which could be caused by collagen aggregation mediated by Au. Thicker fibrils as a result of more collagen aggregation by Au were observed in collagen with 43.5 ppm of Au. The surface of collagen with 174 ppm of Au appeared to be full of Au, which could be the accumulated Au multilayers on collagen. To gain insight into the 13


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wettability of pure collagen and collagen+Au composites, water contact angle was measured. As shown in Figure 3C, the contact angle of water on pure collagen was about 88.5o. The obtained average contact angles of collagen+Au (17.4 ppm), collagen+Au (43.5 ppm), and collagen+Au (174 ppm) were 68.2o, 67.6o, and 64.5o respectively, indicating that the hydrophilicity was elevated with the increase in the concentrations of Au in the collagen+Au composites. Finally, the thermogravimetric analysis (TGA) of different materials was measured by a thermogravimetritic analyzer. The pyrolytic temperatures (Tonset and Tp) were derived from the TGA curves and summarized in Table I. There is an abrupt increase in the onset temperature (Tonset) of pyrolysis when 17.4 ppm and 43.5 ppm of Au were added into collagen. The onset temperature dropped to background (that of collagen) when 174 ppm of Au were added into collagen. The peak pyrolytic temperature (Tp) increased when 43.5 ppm of Au were added into collagen, while the peak pyrolytic temperatures were comparable to collagen alone when 17.4 ppm and 174 ppm of Au were added into collagen. In summary, these data indicated that the biomechanical properties of different materials including elasticity, topographical structures, wettability, and pyrolytic temperatures were changed with the addition of different concentrations of Au.

3.3 VEGF is required for the SDF1-α -induced expression of CXCR4 and eNOS Our previous report has shown that MSCs seeded on collagen+Au 43.5 ppm displayed better cell survival, lower ROS generation, lower monocyte activation, and lower platelet activation than other treatments12. MSCs seeded on collagen+Au 43.5 ppm stimulated with VEGF or SDF-1α also expressed more α5β3 integrin, CXCR4, phosphorylated focal adhesion kinase (p-FAK), phosphorylated Akt (p-Akt), endothelial nitric oxide synthase (eNOS), and active form of matrix metalloproteinase-2 (active MMP-2) than on control material. Moreover, collagen+Au 43.5 ppm promoted the proliferation and migration of MSCs as well as induced the differentiation of MSCs into endothelial cells in the presence of VEGF or SDF-1α as compared to control material12. Therefore, collagen+Au 43.5 ppm was chosen for all the 14



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following experiments using the MSCs isolated from Wharton’s jelly tissue of human umbilical cord. The MSCs we used displayed fibroblast-like morphology (Figure 4A). Further characterization showed that these MSCs were positive for stem cell markers CD29 (99.7%), CD44 (99.6%), CD73 (99.5%), CD90 (98.6%), and CD105 (91.7%), while they were negative for markers CD14 (0.72%), CD34 (0.58%), and CD45 (1.0%) (Figure 4B), suggesting that these MSCs possess bona fide stem cell properties. It is known that both VEGF42 and SDF-1α43 signaling pathways could have a crosstalk and involve in the migration of stem cells. To explore the role of VEGF on the SDF-1α-mediated signaling pathway to induce the migration of MSCs seeded on different materials, VEGF siRNAs were used to knock down the expression of VEGF. Before their application, we tested the cytotoxicity of scramble siRNA molecules on MSCs. Different concentrations of scramble siRNAs (6.25, 12.5 and 50 nM) were transfected into MSCs for 24, 48 and 72 hrs, and the cytotoxic effects of scramble siRNAs on MSCs were assessed by MTT assay. As shown in Supplemental Figure 1A, transfection of scramble siRNAs decreased the viability of MSCs only at the highest concentration (50 nM), less toxicity or no toxicity was observed at concentrations between 0 to 25 nM of scramble siRNAs at different time points. Therefore siRNA concentration at 25 nM was used for the following experiments. To test the uptake efficiency of siRNA molecules by MSCs, different concentrations of fluorescein-conjugated scrambled siRNA molecules (0 to 50 nM) were transfected into MSCs and incubated for 5-7 h, then the fluorescence intensity was analyzed by flow cytometry. As shown in Supplemental figure 1B, the fluorescence intensity increased in a dose-dependent manner, suggesting the efficient uptake of siRNA molecules by MSCs at any concentration. To determine whether SDF-1α-induced expression of CXCR4 is through the VEGF-mediated signaling pathway, we knocked down VEGF expression by specific VEGF siRNA (25 nM) without or with SDF-1α (50 ng/ml) stimulation, and then examined the knockdown effects on the expression of CXCR4 by immunofluorescent staining. As shown in Supplemental Figure 2A, treatment of MSCs seeded on collagen with SDF-1α induced prominent CXCR4 expression, while knockdown of VEGF 15


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down-regulated this induction. Unexpectedly MSCs seeded on collagen+Au was able to induce CXCR4 expression in the absence of SDF-1α, and treatment with SDF-1α further increased the CXCR4 expression. Similarly SDF-1α-induced expression of CXCR4 can be suppressed by the knockdown of VEGF. We also quantified the CXCR4 expression based on the fluorescence intensity after treatment. As shown in Supplemental figure 2B, VEGF knockdown substantially decreased the CXCR4 protein expression of MSCs on collagen and collagen+Au after 48 h of incubation. These data suggested that VEGF is involved in regulating SDF-1α-mediated CXCR4 expression of MSCs on different materials. Previous study has shown that treatment of MSCs with VEGF and SDF-1α respectively increased the eNOS protein expression, which may account for their promotion on the cell proliferation and migration of MSCs on different materials12. To examine the contribution of VEGF on SDF-1α-induced expression of eNOS, we knocked down VEGF expression and treated with or without SDF-1α, and then the effects on eNOS expression were analyzed by immunofluorescent staining. As compared to control treatment, growth of MSCs on collagen induced the eNOS expression and induced even more eNOS expression on collagen+Au (Figure 5A). Treatment of MSCs with SDF-1α further increased the eNOS expression and this induction can be compromised by VEGF knockdown on all materials. We also quantified the eNOS expression based on the fluorescence intensity after treatment and had the same conclusion (Figure 5B).

3.4 VEGF is required for SDF-1α-induced cell migration of MSCs Previous study has shown that treatment of MSCs with either VEGF or SDF-1α can increase the cell migration on both collagen and collagen+Au12. The aforementioned study showed that VEGF was involved in SDF-1α-mediated signaling. Thus it is interesting to know whether VEGF participates in the SDF-1α-induced cell migration. To achieve this, we knocked down VEGF expression and treated with or without SDF-1α, and then examined the effects on cell migration. Cells that migrated into the boundary of each well were recorded before (t = 0 h) and after migration (t = 48 h), and the cell migration ability was determined (Figure 5C). The real time images of cell boundary determined at pre-migration (0 h) are 16



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provided in Supplemental figure 3. As quantified the migration distance compared to control treatment, growth of MSCs on collagen and collagen+Au increased the cell migration (Figure 5D). Cell migration was further stimulated after treatment with SDF-1α for 48 h, while the SDF-1α-induced cell migration was suppressed by knockdown of VEGF on all materials. In the previous study, we have shown that treatment of MSCs with VEGF or SDF-1α increased the expression of the α5β3 integrin on collagen+AuNP and suggests that α5β3 integrin/metalloproteinase-2 (MMP-2) signaling pathway contributes to the better biological performance of MSCs on collagen+AuNP12. To assess the involvement of VEGF on SDF-1α-induced expression of α5β3 integrin, we knocked down VEGF expression and treated with or without SDF-1α, and then the effects on α5β3 integrin expression were evaluated by immunofluorescent staining. As compared to collagen treatment, growth of MSCs on collagen+Au induced more α5β3 integrin expression. Treatment of MSCs with SDF-1α further increased the α5β3 integrin expression. VEGF knockdown decreased both the α5β3 integrin expression and the induced-α5β3 integrin expression on all materials (Supplemental Figure 4A). We also quantified the α5β3 integrin expression based on the fluorescence intensity after treatment and come to the same conclusion (Supplemental Figure 4B). This finding suggested that VEGF is involved in SDF-1α-induced α5β3 integrin expression for the observed biological activity. It is known that matrix metalloproteinase-2 (MMP-2) is important for stem cell migration12. To explore the role of VEGF in SDF-1α-induced cell migration of MSCs on pure collagen and collagen+Au, we knocked down VEGF and examined the enzymatic activity of secreted matrix metalloproteinase-2 (MMP-2) in SDF-1α-treated cells. As shown in Supplemental Figure 5, SDF-1α increased the enzymatic activity of MMP-2, while knockdown of VEGF suppressed this induction. Our data demonstrated that down-regulation in SDF-1α-induced cell migration after VEGF knockdown may be partially due to the reduction in the enzymatic activity of secreted MMP-2 of MSCs on collagen+Au.

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3.5 Well attachment of MSCs on collagen+Au-coated catheter After biocompatibility tests, biomechanical assays, and biological activity studies, we expected that our collagen+Au composites were suitable for vascular tissue repair. To address this idea, we stepped forward to examine the attachment of MSCs on collagen+Au coated catheters. MSCs were labeled with red fluorescence quantum dot (Qtracker 655) and allowed to attach onto catheters coated with different materials. As shown in Figure 6, best attachment of MSCs were observed on collagen+Au-coated catheters (144.3±4.73 of fluorescence intensity), followed by collagen-coated catheter (127.3±8.02 of fluorescence intensity), as compared to that of blank catheter (without any coating; 22.0±2.65 of fluorescence intensity).

3.6. Endothelial differentiation ability of MSCs on collagen+Au-coated catheter Furthermore, immunofluorescent staining showed that MSCs treated with collagen+Au 43.5 expressed more CD-31 than MSCs treated with collagen or cells incubated with control material alone after 7 day of incubation (Figure 7A). It was also quantified the CD31 expression based on the fluorescence intensity after treatment and had the same result (Figure 7B). In order to track the endothelial differentiation ability of MSCs on collagen+Au-coated catheter in vivo, cells were pre-labeled with quantum dot before implantation into femoral artery. After one month of implantation, IHC staining showed that CD31-positive cells were distributed and co-localization with quantum dot labeled-positive cells (Figure 7C). These data not only suggested that MSCs on collagen+Au material appeared to differentiate into endothelial cells, but also showed that MSCs on collagen+Au-coated catheter displayed the best ability to differentiate into endothelial cells, as compared to other treatments.

3.7 Well tissue organization, less blood clot formation and Less fibrosis occurrence of regenerated tissues on the implanted collagen+Au-coated catheter with MSCs After the adhesion test, unlabelled MSCs were seeded into uncoated catheter, collagen- or 18



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collagen+Au-coated catheters before implanting into the femoral artery of rat for the study of vascular regeneration. 30 days after implantation, catheters were isolated, and tissues were sectioned for H&E staining and histological examination. Tissues isolated from collagen+Au-coated catheter with MSCs appeared to be well-organized, as compared to those of collagen-coated catheter with MSCs, catheter with MSCs or catheter only (Figure 8A). The SEM images of platelets adhered and activated on the surface of control group (pure collagen) and collagen+Au 43.5 are shown in Supplemental figure 6. The amount of adhered and activated platelets on the control group (pure collagen) was more prominent than that on collagen+Au 43.5. Besides, less blood clots were formed on tissues isolated from collagen+Au-coated catheter with MSCs (Figure 8A), suggesting that collagen+Au-coated catheter with MSCs possessed better anti-thrombosis ability than those of other tested groups. Chronic inflammation of the vascular tissue can lead to fibrosis which is caused by over-expression and deposition of collagen. Masson's trichrome staining can be used to differentiate collagen from other fibers. We therefore examined the fibrosis of the isolated artery tissues by Mason’s trichrome staining (Figure 8B). By quantification, abundant collagen was deposited on the tissue isolated from control catheter (271.3±16.0 of intensity) (p

Prominent Vascularization Capacity of ... - ACS Publications

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