Stiffness of aligned fibers regulates the phenotypic ... - ACS Publications

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Biological and Medical Applications of Materials and Interfaces

Stiffness of aligned fibers regulates the phenotypic expression of vascular smooth muscle cells Bingcheng Yi, Yanbing Shen, Han Tang, Xianliu Wang, Bin Li, and Yanzhong Zhang ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.9b00293 • Publication Date (Web): 24 Jan 2019 Downloaded from http://pubs.acs.org on January 25, 2019

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ACS Applied Materials & Interfaces

Stiffness of Aligned Fibers Regulates the Phenotypic Expression of Vascular Smooth Muscle Cells Bingcheng Yi a, Yanbing Shen a, Han Tang a, Xianliu Wang a, Bin Li d,e*, Yanzhong Zhang a,b,c,e*

a College

of Chemistry, Chemical Engineering & Biotechnology, Donghua University, Shanghai 201620, China

b State

Key Laboratory for Modification of Chemical Fibers and Polymer Materials, Donghua University, Shanghai 201620, China

c Key

Lab of Science & Technology of Eco-Textile, Ministry of Education, Donghua University, Shanghai 201620, China d Orthopaedic

e China

Institute, Medical College, Soochow University, Suzhou 215006, China

Orthopedic Regenerative Medicine Group (CORMed), Hangzhou 310058, China

* Corresponding author Yanzhong Zhang, Ph.D., Professor of Biomaterials, College of Chemistry, Chemical Engineering & Biotechnology, Donghua University, 2999 North Renmin Road, Shanghai 201620, China. Tel/Fax: +86 21 6779 2374, Email: [email protected] Bin Li, Ph.D., Department of Orthopaedics, The First Affiliated Hospital, Orthopaedic Institute, Soochow University, Suzhou, Jiangsu 215006, China. Email: [email protected]

1

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Abstract Electrospun uniaxially aligned ultrafine fibers show great promise in constructing vascular grafts mimicking the anisotropic architecture of native blood vessels. However, understanding how the stiffness of aligned fibers would impose influences on the functionality of vascular cells has yet to be explored. The present study aimed to explore the stiffness effects of electrospun aligned fibrous substrates (AFSs) on phenotypic modulation in vascular smooth muscle cells (SMCs). A stable jet coaxial electrospinning (SJCES) method was employed to generate highly-aligned ultrafine fibers of poly(L-lactide-co-caprolactone)/poly(L-lactic acid) (PLCL/PLLA) in shell-core configuration with a remarkably varied stiffness region from 0.09 to 13.18 N/mm. We found that increasing AFS stiffness had no significant influence on cellular shape and orientation along fiber direction with the cultured human umbilical artery smooth muscle cells (huaSMCs), but inhibited cell adhesion rate, promoted cell proliferation and migration, especially enhanced the F-actin fiber assembly in the huaSMCs. Notably, higher fiber stiffness resulted in significant down-regulation of contractile markers like alpha-smooth muscle actin (-SMA), smooth muscle myosin heavy chain (SM-MHC), calponin and desmin, whereas up-regulated the gene expression of pathosis associated osteopontin (OPN) in the huaSMCs. These results allude to the phenotype of huaSMCs on stiffer AFSs being miserably modulated into a proliferative and pathological state. It consequently affected adversely the proliferation and migration behavior of human umbilical vein endothelial cells (huvECs) as well. Moreover, stiffer AFSs also revealed to incur significant up-regulation of inflammatory gene expression, such as interleukin-6 (IL-6), monocyte chemoattractant protein-1 (MCP-1) and intercellular adhesion molecule-1 (ICAM-1), in the huaSMCs. This study stresses that while electrospun aligned fibers are capable of modulating native-like oriented cell morphology and even desired phenotype realization or transition, it might not always direct cells into correct functionality. The integrated fiber stiffness underlying is thereby a critical parameter to consider in engineering structurally anisotropic tissue-engineered vascular grafts to ultimately achieve long-term patency.

Keywords: Vascular smooth muscle cells, electrospun aligned fibers, stiffness, shell-core structure, contractile phenotype, macrophage-like phenotype 2


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1. Introduction

Vascular diseases, specifically the blockage by thrombosis or atherosclerosis occurred in small-diameter (< 6 mm) coronary and peripheral vessels, is one of the leading causes of mortality worldwide due to the lack of appropriate functional substitutes for revascularization.1 Tissue-engineered blood vessels (TEBVs) have been proposed as most promising alternatives to realize the functional recovery of diseased vascular tissues.2,3 However, complications such as thrombogenesis and intimal hyperplasia after long-term implantation, are still frequently reported in small-diameter TEBVs4,5, even if well-exercised luminal surface modification techniques (e.g., heparin- or RGD-conjugation6,7) for improving antithrombogenicity were prior applied. One of the major reasons recognized for the implantation failure is the mechanical stiffness (or compliance) mismatch between engineered grafts and native arteries, which is highly susceptible to induce chronically intimal hyperplasia.8,9 Designing and fabricating three-dimensional vascular scaffolds that ultimately mimic the structural, biochemical and biomechanical characteristics of the extracellular matrix (ECM) of native blood vessels, remains a challenge in engineering ideal TEBVs with desired long-term patency and regeneration capability.

As a unique ultrafine fiber producing technique, electrospinning has been extensively adopted to fabricate biomimicking fibrous scaffolds for tissue regeneration applications. With a nano/micro scale fineness that closely resembles the fibrous morphology of constituents within the native ECM, electrospun fibers also offer advantages of high specific surface area for binding biomolecules, and mechanically a sufficiently lowered softness enabling cells to play with for efficacious remodeling10,11, which thereby are well-suitable for uses in engineering soft tissues (such as blood vessels, nerves, cornea, etc.). In vascular tissue engineering12, recognizing the fact that native vascular tissue mainly comprises of three layers of structure (i.e., intima, media, and adventitia) with orderly arranged collagen and elastic fibers13, recently some attempts have been made to use electrospun aligned fibers for constructing bi-layered14-16 or tri-layered8,17,18 scaffolds towards ultimately recapitulating the natural anisotropic architecture of the vasculature wall.

The use of aligned electrospun fibers for constructing of vascular scaffolds is indeed of considerable interest and importance. It has been well-documented that oriented electrospun 3



nanofibers in 2-D planar form can not only guide the vascular cells to elongate and align along the fiber direction5,8,19,20, but also promote directional cell movement, desired phenotypic switch and aligned ECM deposition.5,21 Moreover, electrospun aligned fibers with the surface decoration of hyaluronan can be fabricated into circumferentially aligned tubular structure, that was demonstrated to conducively encourage regeneration of contractile vascular SMCs in the media layer and formation of endothelial cell monolayer in the lumen.5 In terms of fiber stiffness effects, stiffness of scaffolding fibers in random orientation was recently reported to be a more critical parameter than surface chemistry in directing the contractile behavior of SMCs7, and an appropriate stiffness could enhance SMC contractility, reduce cell proliferation and guide mesenchymal stem cells to express SMCs-specific phenotypes.7,22,23 For instances, within certain limits of substrate stiffness (0.38-12.70 N/mm2), lower stiffness of random electrospun-fibrous substrate has been proved to promote the contractile phenotype of vascular SMCs.7 SMCs can organize their cytoskeleton and focal adhesions much more on “rigid” glass or “stiff” nanofiber hydrogel matrix than on “soft” gels, further promoting SMC spreading, polarization, and random motility.23,24 However, the above researched substrates were mainly based on homogeneous matrices of fibers; relatively little is known regarding whether the underlying stiffness of electrospun aligned fibers can influence overall cellular behavior and specific functionality, particularly the vascular SMCs, which are a principle cellular component responsible for controlling the contraction/dilation dynamics in the blood vessels.

The objective of this study was to examine the function alteration of vascular SMCs affected by the stiffness of highly-aligned electrospun fibers in shell-core structure. Coaxial electrospinning permits to produce ultrafine fibers whose stiffness could be controlled by varying the shell-core structure with no intervention in surface chemistry of a scaffold architecture (obviating a potential confounding effect).25,26 In our previous studies stable jet electrospinning (SJES)27,28 had been demonstrated to be effective in generating highly-aligned fiber arrays. Therefore, a method combining coaxial electrospinning with SJES (termed stable jet coaxial electrospinning, SJCES) was developed to fabricate highly-aligned fibrous substrates (AFSs) consisting of shell-core structured fibers with tailored stiffness, by varying compositional ratios of the opted rigid PLLA-core and pliable PLCL-shell layers. A series of biological tests including cell adhesion, 4


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proliferation, morphology, migration and expression of specific phenotypic makers, were subsequently carried out to reveal the response of vascular SMCs to the fiber stiffness. This study will provide insightful understanding on fiber stiffness effects associated with both vascular pathophysiology and the design of vascular scaffolds for viable TEBVs.

2. Materials and methods

2.1 Fabrication of aligned PLCL/PLLA shell-core fibers

Using

1,1,1,3,3,3-hexafluoro-2-propanol

as

the

common

dissolving

solvent,

poly(L-lactide-co-caprolactone) (PLCL, LA:CL 50:50, IV 2.5 dl/g, Jinan Daigang Biomaterials) at a concentration of 12% w/v and poly(L-lactic acid) (PLLA, MW 100,000, Jinan Daigang Biomaterials) at 10% w/v concentration were prepared as the core and shell solutions, respectively. To enable facile collection of highly-aligned fiber arrays, a tiny amount (5% w/w) of high molecular weight poly(ethylene oxide) (PEO, Mw > 5,000,000, Avocado, UK) were respectively introduced into the PLCL and PLLA spinning dopes for eliminating the jet whipping observed in conventional electrospinning.27,28 Then SJCES was performed to generate shell-core fibers of PLCL/PLLA using the following parameters: applied voltage 5-7 kV, needle tip-to-collector gap distance 20 cm, drum (10 cm in diameter) collecting speed 1000 rpm, ambient conditions (20-25 °C, 25-30% humidity). Flow rates of the PLLA-core and PLCL-shell were concurrently altered within a total flow rate of 0.5 mL/h to generate 4 groups of shell-core fibers as follows: 0.5-0 (0.5 mL/h for PLCL and 0 mL/h for PLLA as S1 group, and so on), 0.35-0.15 (S2 group), 0.2-0.3 (S3 group) and 0.05-0.45 (S4 group). After SJCES, all the produced AFSs were placed in vacuum for 2 weeks of drying. 2.2 Characterization of the AFSs 2.2.1 Fiber morphology Morphology of AFSs was observed by a Hitachi TM-1000 scanning electron microscope (SEM) at an acceleration voltage of 8-10 kV. All AFSs were sputter-coated for 60 s with gold to increase conductivity prior to SEM imaging. Fiber diameter and fiber alignment degree (n > 50) were 5



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analyzed using the ImageJ software (NIH, Bethesda, Maryland, USA) based on obtained SEM images.

2.2.2 Shell-core structure

Shell-core structure of the generated PLCL/PLLA fibers via the SJCES was examined using transmission electron microscopy (TEM, JEM-2100F, JEOL) at 80 kV. All the composite fibers for TEM observation were prepared by directly depositing fibers onto a rectangle frame, then transferring the as-electrospun fibers to copper grids, and finally allowing to dry in a vacuum oven for 48 h at room temperature. Shell-core structure of the PLCL/PLLA fibers were also characterized using an inverted fluorescence microscope (Eclipse Ti-S, Nikon, Japan) by prior doping the PLLA solution with a 5 μg/mL rhodamine (Sinopharm Chemical Reagents, China). Images from TEM and fluorescent microscopy were analyzed using the ImageJ for determining shell/core thickness of the fibers (n >5). To estimate the core radius (Rc) and shell thickness (Ts) theoretically, the overall fiber radius of a group of shell-core fibers (Ra) was firstly measured from its SEM images; then the Rc of the shell-core fibers was reckoned through the following formula:

π  Ra2 - π  Rc2 V  s 2 π  R0.5 0 V0.5 0

(1)

Where Vs is the flow rate of the PLCL solution in the PLCL/PLLA group during SJCES, and V0.5-0 and R0.5-0 are the flow rate (0.5 mL/h) and the fiber radius (0.7 m) in the 0.5-0 group, respectively. The Ts could thus be calculated as Ra minus Rc of the PLCL/PLLA shell-core fibers. 2.2.3 Tensile properties Tensile properties of the AFSs, collected at a drum rotation speed of 1000 rpm for 2 h, were determined using a tabletop tensile tester (HY-940FS, Hengyu Instrument, China) equipped with a 50 N load cell at ambient conditions. All samples were stretched at a cross-head speed of 10 mm/min. Elastic modulus was derived from the generated stress-strain curves, while the fiber stiffness was calculated by multiplying the elastic modulus with cross-sectional area and dividing a gauge length of 3 cm.7,29 At least 8 samples were tested for each type of the AFSs. 6


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2.3 Cellular responses of huaSMCs to the varied AFS stiffness

2.3.1 Cell culture and seeding

Human umbilical arterial smooth muscle cells (huaSMCs) and human umbilical vein endothelial cells (huvECs) were obtained from the ScienCell Research Laboratories (ScienCell, USA). For cell culturing, huaSMCs were maintained in smooth muscle cell medium (ScienCell, USA) supplemented with 2% fetal bovine serum (FBS), 1% penicillin/streptomycin and 1% SMCGS, and huvECs were maintained in endothelial cell medium (ScienCell, USA) supplemented with 5% fetal bovine serum (FBS), 1% penicillin/streptomycin and 1% ECGS.

For cell seeding, all the AFSs were prior sterilized by ultraviolet (UV) light irradiation for 12 h, and 75% ethanol treatment for 2 h followed by washing three times with phosphate buffered saline (PBS). Then, the cells were placed in 24- or 6-well culture plates and incubated with specific cellular medium at 37 °C overnight to deposit a layer of proteins.

2.3.2 Cell adhesion and proliferation

For cell adhesion, huaSMCs were stained with 5 M Celltracker Green CMFDA (Yeasen, China) for 15 min according to the manufacturer’s instruction. Then, 500 L of 1 105 CMFDA-labeled huaSMCs were seeded on the AFS scaffolds in 24-well culture plates and incubated at 37 °C, 5% CO2, and 95% relative humidity for 6 h. Thereafter, the AFS scaffolds were washed by PBS to remove the unattached huaSMCs and further fixed by using paraformaldehyde (4%) for fluorescence microscopy.

For cell proliferation, 500 L of 5  104 cells were incubated on the AFS scaffolds in 24-well culture plates for 1, 3 and 5 days and the cell number was determined by the CCK-8 assay (Beyotime, China). Briefly, each well of the plate was added with 20 µL of CCK-8 and incubated at 37 °C for 4 h. Then 100 µL volumes of the solution were transferred to a 96-well plate reader (MK3, Thermo, USA) for absorbance measurements at 450 nm.

2.3.3 Cell morphology analysis 7



huaSMCs at a density of 5  104 cells for cell alignment analysis or 1  104 cells for cell shape analysis were seeded onto the AFS scaffolds in 24-well culture plates. After culturing the cell-scaffold constructs for 3 days, the cellularized constructs were washed and fixed by using 4% paraformaldehyde for 30 min. These constructs were then permeabilized using 500 L of 0.2% (v/v) Triton X-100 (Aldrich, USA) in PBS for 10 min. Thereafter, the stress fibers of huaSMCs were stained with 200 μL of Alexa Fluor™ 568 Phalloidin (1:40 dilution in PBS, Invitrogen™, USA) for 30 min at room temperature in the dark. The cell nuclei were stained using 0.8 mg mL1 of 4′,6-diamidino-2-phenylindole (DAPI, Invitrogen™, USA) in PBS for 10 min at room temperature. After washing, morphology of the huaSMCs and single huaSMC was observed using the inverted fluorescence microscope. Morphological characteristics, including cell alignment defined as the degree of cell nuclei deviated from the fiber direction, average area of cell spreading, elliptical form factor (EFF) defined as the major axis divided by the minor axis, and shape factor defined as 4πA/P2 where A is the area and P is the perimeter of the cell, were analyzed via ImageJ based on the captured microscope images.

2.3.4 Cell mobility

Cell migration assay by prior producing a cell-free wound-like gap area (defined as "artificial wound") in the middle of a confluent cell layer was used to investigate cell mobility. In brief, custom-made stainless steel rings ( = 15 mm) with 1 mm barrier in the middle were placed on the prepared AFS scaffolds in 24-well culture plates. Then huaSMCs were seeded at a density of 5  105 cells/well with 500 L of smooth muscle cell medium. After 24 h of culture, the rings were removed to initiate cell migration to the cell-free regions within a time frame of 48 h. Finally, the cells were fixed and stained with Alexa Fluor™ 568 Phalloidin for mobility observation using inverted fluorescence microscope.

2.4 Immunofluorescence staining

huaSMCs at a density of 5  104 cells per 500 L were seeded on each of the AFS scaffolds in 24-well culture plates. After culturing for 3 days, immunofluorescence was performed to examine the phenotype protein (α-SMA, SM-MHC) and migration-related protein (CD44) expression.30,31 8


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Briefly, the cellularized constructs were washed with PBS three times, fixed using 4% paraformaldehyde for 30 min and further permeabilized using 0.2% Triton X-100 in PBS for 10 min. Then the non-specific binding sites were blocked using 10% goat serum for 30 min. Thereafter, rabbit polyclonal anti-human α-SMA antibody (1:200 dilution, Abcam, UK), mouse monoclonal anti-human SM-MHC antibody (1:250 dilution, Abcam, UK) or mouse monoclonal anti-human CD44 antibody (10 g/mL, Abcam, UK) were incubated overnight at 4 °C. Then a hybrid solution of the secondary antibodies, including goat anti-rabbit IgG H&L secondary antibodies (Alexa Fluor 568) (1:200 dilution, Abam, UK) for α-SMA, goat anti-mouse IgG H&L (FTIC) (1:1000 dilution, Abam, UK) for SM-MHC and CD44, were added for 90 min at room temperature followed by cell nucleus DAPI-staining. Finally, the samples were observed using a laser confocal scanning microscope (LCSM, Carl Zeiss, Germany) for the detection of phenotypic proteins.

2.5 Transwell co-culture of huaSMCs and huvECs

A confluent huaSMCs monolayer on AFSs was used to co-culture with huvECs using Transwell device (=15 mm, pore size 8 m, Corning, USA) for determining the AFS stiffness-related influences of huaSMCs on huvECs by performing proliferation and migration assays. Briefly, huaSMCs at a density of 1  105 per 800 μL were seeded onto each of the AFS scaffolds and incubated overnight to form a confluent huaSMCs monolayer in the lower part of the Transwell chamber. Then huvECs (1  105 cells per 200 𝜇L) were seeded into the upper part of the Transwell chamber to co-culture with the confluent huaSMCs monolayer on AFSs. After 24 h of incubation, the upper part of the Transwell chamber was taken out and huvECs migrating downward through the Transwell membrane were stained by crystal violet (Mesgen, China) and observed using the inverted fluorescence microscope. Likewise, proliferation of the huvECs in the upper part of the Transwell chamber at 3 days was assayed by the CCK-8 method. For comparison, the group cultured with huvECs only in the upper part of the Transwell chamber (i.e., without huaSMCs and AFSs in lower part of the Transwell chamber) was chosen as a blank control.

2.6 Gene expression analysis 9



huaSMCs at a density of 5  105 cells per 2 mL were seeded onto each of the AFS scaffolds in 6-well culture plates for 5 days of culture. Quantitative real-time polymerase chain reaction (qRT-PCR) was then performed to investigate the mRNA expression of α-SMA, calponin, OPN, desmin, IL-6, MCP-1 and ICAM-1. In brief, mRNA was isolated from the huaSMCs using a total RNA isolation kit (Tiangen, China) according to the manufacturer’s instruction. The cDNA was reverse-transcribed using FastKing RT Kit (with gDNase) (Tiangen, China) following the manufacturer’s protocol. Then, qRT-PCR was performed using the Super Real PreMix Plus (SYBR Green) reagent Kit (Tiangen, China) with an ABI Prism 7500 Sequence Detection System (Applied Biosystems, USA). All PCR results were normalized to the expression levels of GAPDH prior to further analysis. All primer sequences (Sangon Biotech, China) used for the real-time reverse transcription-PCR are listed in Table 1. Table 1 Primer sequences of related gene in huaSMCs Genes

Forward primer sequence (5’-3’)

Reverse primer sequence (5’-3’)

GAPDH

TATTCTCTGATTTGGTCGTA

ATGGCAACAATATCCACT

-SMA

AGACTTCCGCTTCAATTC

CTGTTAGGACCTTCCCTC

ACACAACTACTACAATTCC

TCTCTCCAAACTCTAACC

ATGGAAAGCGAGGAGTTGAATG

TGCTTGTGGCTGTGGGTTT

AGGAGAGCCGGATCAATCTCC

TCCCGTGTCTCGATGGTCTT

CTGGATTCAATGAGGAGAC

AATCTGTTCTGGAGGTACT

MCP-1

TTCCTCTTGAACCACAGT

CTTGCAAAGACCCTCAAA

ICAM-1

GACTAAGCCAAGAGGAAG

CTCAGCATACCCAATAGG

Calponin OPN Desmin IL-6

2.7 Statistical analysis

Data are presented as means ± standard deviation. Statistical analysis was carried out using the Origin 8.0 software, and Tukey’s HSD post hoc tests were used to make pair-wise comparisons between groups. A value of *p< 0.05 or **p< 0.01 was considered statistically significant based on one-way analysis of variance (ANOVA). All experiments were carried out at least thrice.

3. Results

3.1. Fabrication of highly-aligned shell-core fibers of PLCL/PLLA 10


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SJCES was proven to be effective in producing highly-aligned composite fibers of PLCL/PLLA, where the elastic PLCL (Tg = 31 C) and the rigid PLLA (Tg = 56 C) were used as the shell and core components in the electrospinning process, respectively (Fig. 1A). SEM observation revealed that all the AFSs with various couples of shell-core feeding rates during SJCES present a unidirectionally aligned and smooth morphology (Fig. 1B). Average diameters for the AFSs measured (PLCL/PLLA: 0.50-0, 0.35-0.15, 0.2-0.3 and 0.05-0.45) were in the ranges of 1.40 ± 0.10, 1.40 ± 0.16, 1.34 ± 0.20 and 1.28 ± 0.17 m, respectively. There were no significant differences in fineness (p > 0.05) between these shell-core fibers. Moreover, from the quantified fiber orientation (Fig. 1C), no significant difference in orientation of the aligned PLCL/PLLA fibers was detected as well.

Fig. 1 Fabrication of highly-aligned composite fibers of PLCL/PLLA in shell-core structure via the SJCES method: (A) Schematic of the SJCES process; (B) Fiber morphology and measured diameters; (C) Quantified degree of fiber orientation.

Furthermore, to confirm the formation of shell-core structure with the SJCES approach, both TEM and fluorescence imaging were performed (Fig. 2). Increasing flow rates of the PLLA-core solution during electrospinning could generate theoretically coaxial fibers with larger core diameters.25 As expected, clear shell-core structure with the PLLA-core fully sheathed by the 11



PLCL-shell was detected for all the AFSs (Fig. 2A-B); and there was no any observable polymer mixing between the core and shell components. Similar to the TEM images, fluorescence images of the PLCL/PLLA fibers with the PLLA-core solution for electrospinning being prior-doped with a tiny amount of fluorescent dye, qualitatively depict a tendency of progressively augmented core diameter along with increasing the PLLA-core flow rates. Such a varying tendency in core radius and shell thickness by increasing the inner flow rates was also quantitatively affirmed (Fig. 2C). Collectively, results shown in Figs. 1 & 2 demonstrated the SJCES a facile and effective method for producing highly-aligned fiber arrays with similar substrate topography and consistent surface chemistry.

Fig. 2 Characterization of the formed shell-core structure with the PLCL/PLLA fibers produced via SJCES: (A) TEM images; (B) Fluorescence micrographs (red represents rhodamine-doped PLLA core); (C) Measured core radius and shell thickness of the PLCL/PLLA fibers from the TEM and fluorescence images.

3.2. Stiffness tuning of the PLCL/PLLA AFSs

To modulate the gross coaxial fiber-substrate stiffness, two polymeric components, i.e., the elastic PLCL and the rigid PLLA with drastically different mechanical properties, were judiciously 12


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chosen as the shell and core components, respectively. As expected, a progressive distending in diameter of the PLLA-core significantly improved the elastic moduli of AFSs, varying from 14.68 ± 1.09 MPa (corresponding stiffness: 0.09 ± 0.01 N/mm) of the pure PLCL to 2141.72 ± 63.74 MPa (corresponding stiffness: 13.18 ± 0.39 N/mm) of the 0.05-0.45 PLCL/PLLA (Fig. 3B-C). That is, the stiffness of AFSs could be increased along with encapsulating more component of the stiffer PLLA-core, which accordingly gave rise to stiffness tuning of the composite fibers in a span range of 145 times. As a result, the percentage elongation at failure of the AFSs decreased with increasing the PLLA-core mass fractions, showing a significantly decrease to about 115% from 480% of the pure PLCL AFSs (Fig. 3A). These mechanical data reveal that tailoring the fiber stiffness could be easily achieved through altering the core dimension with the SJCES method.

Fig. 3 (A) Typical stress-strain curves of the AFSs with varied PLCL/PLLA ratios in the shell-core structure; the insert is an enlarged view of the initial portions of these curves. (B-C) Derived elastic moduli and stiffness of the different AFSs.

3.3. Cell adhesion and proliferation on AFSs

Shell-core structure enabled variation in AFS stiffness was used to firstly investigate the effect of fiber stiffness on cell adhesion by culturing CMFDA-labeled huaSMCs for 6 h (Fig. 4A). Distinct differences in cell adhesion were observed among these AFSs. That is, compared to the smooth surface of tissue culture polystyrene (TCPS) plate, a commonly used reference or control in cell culture, huaSMCs adhered on the aligned fibrous substrata showed elongated cell morphology 13



along the fiber direction and an inclination of descending cell adhesion on stiffer AFSs (Fig. 4A-B). This suggests that huaSMCs prefer to adhere quickly on the soft fibrous substrata compared with the stiffer ones. When cell proliferation was quantified at 1, 3, and 5 days, despite significant differences noted between all AFS groups and the TCPS control enhanced cell proliferation of huaSMCs was observed for cells cultured on stiffer fibrous substrates (Fig. 4C). This shows that huaSMC proliferation positively correlates with the fiber stiffness.

Fig. 4 (A) Schematic diagram and fluorescence images of CMFDA-labeled huaSMC adhered on AFSs with increased fiber stiffness. (B) Quantified huaSMC adhesion on AFSs from the fluorescence images in (A). (C) Cell proliferation of huaSMCs affected by the stiffness of AFSs.

3.4. Cell morphology on AFSs

Morphology of the huaSMCs on AFSs was analyzed using a fluorescence microscope after 3 days 14


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of culture (Fig. 5A). The highly-aligned fibrous substrates appeared to support a better cell elongation and preferential orientation with individual cells exhibiting a unidirectionally oriented spindle-shape when compared to the TCPS control. Quantified results on cell orientation (θ) indicate no significant difference between the AFS groups (Fig. 5B), suggesting an impervious cell-orientation by the varied stiffness of AFSs with comparable topographical and chemical properties. A further scrutinization delineated an intensified organization of the cytoskeleton with increasing the fiber stiffness. This was attributed to the formed higher density of F-actin stress fibers within the individual cells (Fig. 5C). The stress fibers in huaSMC mainly located in the peripheral region of cell body when cultured on softer S1 AFSs. With increasing the AFS stiffness, more neonatal stress fibers showed up around central region of the cell body and elongated along the direction of substrate fibers. The AFS stiffness-modulated subtle differences in organization of the F-actin stress fibers reflect remodelable response in huaSMCs, which are closely associated with variation in cell functionality (e.g., cell proliferative and migratory states).32,33

To assess the cell spreading and polarization as a function of substrate stiffness in AFSs, we quantitatively analyzed cell spreading, shape and elongation by measuring cell area, EFF and shape factor (Fig. 5D-F). To one's surprise, no significant difference in the cell morphological parameters (i.e., similar cell area, EFF and shape factor) was observed on the AFSs with varied stiffness, whereas the huaSMCs on the TCPS control exhibited higher cell area and lower cell elongation and polarization. Our findings suggest that topological cues may be more influential than stiffness cues in directing the cell morphology in current span range of elastic moduli 14.68-2141.72 MPa.

15



Fig. 5 (A) Morphology of the huaSMCs on AFSs with different stiffness. (B) Quantified orientation of huaSMCs affected by AFS stiffness. (C) Cytoskeletal F-actin stress fibers assembled in single huaSMC anchored on different AFSs. (D-F) Quantitative analysis on huaSMCs spreading and elongation affected by the stiffness of AFSs, in terms of mean cell area, elliptical form factor (EFF = dr/ds) and shape factor (= 4A/P2) of individual huaSMCs. In above experiments, huaSMCs were all cultured for 3 days.

3.5. Cell motility on AFSs 16


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Motility of the huaSMCs on the different AFSs was evaluated by observing the cells migrated into the defined “artificial wound” area (Fig. 6A) for 48 h and then analyzing the cell-free region in fluorescent images of cytoskeleton. Aligned fibrous structure of scaffolds is able to display directional contact guidance effect,5,34 in which the cell motility in the direction parallel to the alignment of fibers is usually higher than that in the perpendicular direction. This was further confirmed in our current study (Fig. 6B-C). The higher cell motility in the direction parallel to the alignment of fibers promoted the forefront huaSMCs (around the “artificial wound” edges) to migrate inward the gap area to form a gradually fused cell confluence, while in the perpendicular direction all AFSs still exhibited obvious, uncataclysmic “artificial wound”. Of note, the cell motility in the directions of parallel and perpendicular to the alignment of fibers, was both enhanced on AFSs with higher fiber stiffness.

To probe the expression of migration-related protein for the observed difference in cell migration, the steady state assembly of CD44 as a function of AFS stiffness was assessed qualitatively (Fig. 6D). Immunofluorescent microscopy revealed that CD44 was increasingly expressed as substrate stiffness increases. Relatively lower fluorescence is observed in huaSMCs cultured on the softest fiber substrate (i.e., the S1 group). This demonstrated that migration-related protein expression of CD44 in the huaSMCs was functionally enhanced on AFSs with higher fiber stiffness, thereby promoting the motility of huaSMCs.

17



Fig. 6 (A) Time sequence of steps in performing cell mobility test. (B) Fluorescent staining images of huaSMCs on AFSs after activating cell migration for 48 h, in the directional of parallel (//) or perpendicular () to the fiber direction. (C) Statistical results of the average area of cell-free region in images from (B). (D) Assembly of CD44 influenced by the fiber stiffness.

3.6. Phenotype expression of the huaSMCs on AFSs

Since the expression of multiple huaSMC-specific markers can attest to the phenotype of huaSMCs, we examined the effect of AFS stiffness on protein and gene expression of typical markers. huaSMCs cultured on AFSs all expressed -SMA (an early differentiated smooth muscle marker35) and SM-MHC (a late differentiated smooth muscle marker36) proteins. Increasing the stiffness of fibrous substrata resulted in decrease in SM-MHC protein expression (Fig. 7A), 18


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whereas the difference in -SMA protein expression seemed to be not significant. At the genetic level, it similarly shows higher gene expression of -SMA and calponin (a mid-differentiated smooth muscle marker37) in huaSMCs on the soft fibrous substrata than on the stiff counterparts (Fig. 7B-C). The combined results suggest that huaSMCs could have undergone into de-differentiated phenotype in response to the higher stiffness of AFSs. However, de-differentiated phenotype of SMCs is usually found during normal vascular development or in the pathological process (e.g., atherosclerosis, restenosis after angioplasty or bypass, and hypertension38,39), in which contractile capabilities in vascular SMCs could not be observed in such a pathological status.7,40 To clarify, OPN as an important phenotype marker of de-differentiated SMCs41, also well regarded as a proinflammatory and proatherogenic molecule in the context of atherosclerosis42,43, neointimal hyperplasia44 and vascular calcification45, was used to check the de-differentiated state of the huaSMCs. Significant increase in OPN gene expression was found on stiffer AFSs (Fig. 7D), consequently indicating the pathological phenotype of the de-differentiated huaSMCs. Similarly, expression of desmin, which is a major constituent of the network in SMCs46 and reflects the normal functional activity of the SMCs in terms of structural and mechanical integrity linking myofibrils to the cell membrane47 and extracellular matrix48, was examined. It shows that desmin gene expression was significantly lower on the stiff AFSs than on the soft AFSs (Fig. 7E), which also proved the aberration in SMC normal functionality resulted from the abnormal stiffness of AFSs. These results manifest that excessive fiber stiffness tended to develop the huaSMCs into a de-differentiated, non-contractile, and pathological phenotype.

19



Fig. 7 (A) Immunofluorescence images of differentiation-related protein markers in huaSMCs for -SMA (red), SM-MHC (green) and DAPI (blue) after 3 days of culture. (B-E) qRT-PCR analyses of the phenotype-related genes in huaSMCs cultured on different AFSs for 5 days.

3.7. Coordinated influence of huaSMCs on huvECs

Considering the collaborative role of SMCs and ECs in maintaining the normal vascular structure and function and the inherent interaction between SMCs and ECs49,50, huvECs’ behavior affected by the huaSMCs monolayer cultured on the AFSs with varied stiffness were examined using a Transwell method (Fig. 8A). Stiffer fibrous substrates were firstly found to weaken the inducive role of huaSMCs in promoting huvEC proliferation (Fig. 8B). The proliferation of huvECs cultured in the co-culture system with huaSMCs monolayer on AFSs with the highest stiffness 20


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(i.e., the S4 group) was compromised by approximately 21.4% compared to that of the S1 group. As for the trans-membrane cell migration (Fig. 8C), there were much more huvECs migrating downward to the reverse side of the Transwell insert membrane in softer AFS groups (e.g., S1 and S2). huvECs co-cultured with huaSMCs on stiffer AFSs showed a decreasing tendency in migration, but still exhibiting positive effect as compared to huvECs cultured alone (control group). These results reveal that the presence of huaSMCs on stiffer AFSs could concomitantly affect the proliferation and migration of huvECs as well. The adverse effects on huaECs from the pathological huaSMCs on stiffer AFSs indicate the mutually implicated interactions between the two primary vascular cells. huaSMCs monolayer cultured on soft AFSs were prone to promote better huvEC proliferation and migration than that on stiff AFSs.

Fig. 8 (A) Schematic diagram of huvECs co-cultured with huaSMCs in Transwell device. (B) Proliferation of huvECs affected by the huaSMC monolayer cultured on different AFSs. (C) Optical micrographs showing the migration of huvECs to the reverse side of the Transwell insert when having the huvECs (upper insert) co-cultured with the huaSMC monolayer (bottom of the lower chamber) on AFSs, where purple represents huvECs stained with crystal violet. Control group is the mono-culture of huvECs with no huaSMCs and AFSs in the lower chamber. 21



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3.8. Macrophage-like phenotypes of huaSMCs

To monitor whether huaSMCs exhibit any macrophage-like phenotypes40,51 or characteristics on stiffer AFSs, the expression of macrophage-related genes such as IL-6, MCP-1 and ICAM-1 was detected (Fig. 9). It is interesting to note that enhancing the stiffness of AFSs led to increased gene expression of typical inflammatory mediators concerning IL-6, MCP-1 and ICAM-1, which are already known to be upregulated in hypertensive and atherosclerosis individuals whose artery elasticity is substantially worsen (become stiffer).40,52 These results support the previous observation of macrophage-like phenotypic expression in the vascular SMCs and implicate a potential pathological evolution for the TEBVs with higher stiffness.

Fig. 9 qRT-PCR analyses of inflammatory markers secreted by huaSMCs cultured on the AFSs

4. Discussion

Creating clinically acceptable TEBVs as alternatives to treat blocked or damaged blood vessels in small-diameter caliber is currently the subject of intense research.2,3 Amongst, addressing the frequently reported thrombosis and intimal hyperplasia problems, which are recognized to be two of the most major causes responsible for the low long-term patency of TEBVs, remains a paramount challenge. This situation urgently calls for the development of novel scaffolds resembling the natural architecture of the vessel wall, from which enable cells to grow and remodel their living microenvironment effectively.53 Electrospun aligned fibers offer a tremendous possibility in recapitulating the anisotropic tri-layer architecture of native blood vessels.

While

rendering

the

aligned

fibers

anti-thrombgenic

(i.e.,

achieving

rapid

endothelialization or native-like hemocompatibility) by applying different biochemical surface 22


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modifications54,55 have been extensively demonstrated, impacts from the biomechanical attributes in particular fiber stiffness as a critical underlying parameter, have not been paid due attention. Given that inappropriate scaffold stiffness have a close connection with the functionality of vascular SMCs (e.g., phenotypic alteration from mature contractile to de-differentiated synthetic phenotype, and SMC migration into the lumen with rapid proliferation leading to intimal hyperplasia56), an insightful understanding on the stiffness effects of electrospun aligned fibers, despite of being in ‘sufficient’ softness due to the extremely delicate nano/micro scale fineness in diameter, on SMCs functionality will benefit for rational design of the electrospun aligned fibers-based vascular scaffolds to achieve native-like cell-matrix interactions and remodeling, ultimately leading to long-term success of such tissue-engineered vascular grafts.

SJES has been demonstrated to be an effective approach in producing highly-aligned ultrafine fibrous architecture for vascular tissue engineering.5,34 It is also known that by regulating the proportion of fiber composition constituted of two components with drastically different stiffness (e.g., the elastic PLCL and the rigid PLLA), stiffness tuning of fibers without inflicting interference from chemistry can be achieved via coaxial electrospinning approach.25 Going a step further, in the present study SJCES by combining the SJES and coaxial electrospinning permitted the generation of highly-aligned ultrafine fibers with the same surface chemistry, consistent topographic features (i.e., the fiber fineness and orientation) but varied stiffness. By judiciously selecting two of the most intensively utilized vascular scaffolding biodegradable polymers, i.e., the elastic PLCL and the rigid PLLA5,57, as the representative constituents, we demonstrated that simply varying the injection rates of the PLCL-shell and PLLA-core enables systematic control over their shell-core configuration (Fig. 2), without jeopardizing the overall morphological features (Fig. 1) and surface chemical attribute of the fibers. The non-significant difference in fiber diameter of the four groups actually reflects a gross effect contributed by the polymer type (viscoelasticity), solution concentration, and flow rate in electrospinning. Elastic moduli of the coaxial electrospun scaffolds could be varied nearly 145 times when the mass/volume fraction of PLLA in the shell-core fibers was increased from 0 to 90% (Fig. 3). The shell-core structured fibers of PLCL/PLLA may offer a fundamental platform to mechanistically study interactions of the AFS stiffness with different types of cells in engineering tissues with anisotropic architecture, 23



such as blood vessel, tendon, nerve, and so on.

huaSMCs showed different responsive behaviors when being cultured on the AFSs with varied stiffness. First of all, increasing the stiffness of fiber substrate inhibited to a certain extent the adhesion of huaSMCs but promoted cell proliferation (Fig. 4). The reduced cell adhesion reflects that excessive stiffness could affect the adhesion rate of SMCs on AFSs. Whereas the upregulating tendency on cell proliferation agrees with previous studies using nanopatterned substrata33, fibrous mat7 and hydrogel systems58. Meanwhile, all the highly-aligned fibrous substrates allowed the huaSMCs to grow and spread along the direction of fiber orientation. And no significant difference in cell orientation and cell shape was observed in huaSMCs cultured on the different AFSs (Fig. 5), although there were some noted evidences of increased cell spreading if increasing substrate stiffness of hydrogel or randomly orientated fibrous system.7,58 These results suggest that topographical cues in the present case with aligned fibrous texture may be more influential than matrix stiffness in affecting the cell morphology, which should be considered as an important regulating modality in designing functional vascular scaffolds; as displaying appropriated cytomorphology is essential in identifying the shape, structure, and function of cells.59

But, topographical cues-governed specific morphology in cells may not necessarily reflect the cells in a state of the correct functionality. The dynamic organization of the F-actin stress fibers in vascular SMCs cultured on different substrata plays a prominent role in dictating the cellular behavior, especially proliferation and migration.46,59,60 High cytoplasmic volume fraction of myofilaments in vascular SMCs was usually expressed to maintain the contractility in differentiated phenotype; nevertheless, under pathological conditions like injury or atherogenesis, it would be decreased with the cells being modulated towards a less differentiated phenotype and lost the ability to contract.61 That is to say, the key feature of F-actin stress fibers is to contract for healthy vascular SMCs. In our experiments the dependence of the F-actin stress fibers assembled in huaSMCs on stiffness of the AFSs is somewhat different as the content of F-actin bundling increased slightly with increasing the substrate rigidity (Fig.5 A, C), despite of being in a good agreement with previous observation.7,58 Such unexpected phenomenon (i.e., increased density of F-actin stress fibers) could be associated with the enhanced capacity in cell migration of the 24


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huaSMCs in synthetic or noncontractile phenotype (Fig. 6), as confirmed by the lower expression of contractive makers -SMA, SM-MHC and desmin, and the higher expression of pathosis-related gene OPN and CD44 protein when cultured on AFSs with excessive stiffness (Figs. 6 & 7). This illustrates that F-actin stress fibers here were no longer the key indicator of contractility in the case of SMCs being pathological, but are greatly responsible for the enhanced proliferation and mobility of huaSMCs cultured on stiffer AFSs. High proliferation and mobility are well-known to be two of the prominent characteristics of vascular SMCs responsible for the formation of intimal hyperplasia.

In natural blood vessels, ECs are in close proximity to and communicate with SMCs via heterocellular junctions and signaling molecules.62 These interactions between SMCs and ECs play an important role in maintaining the normal vascular structure and function, such as contractility of SMCs and antithrombogenicity of ECs.49,63 A healthy SMC surrounding can maintain the endothelial cell function and facilitate the remodeling of damaged endothelium within self-healing ability.50 ECs attach, spread and form a confluent monolayer better on quiescent SMCs than on proliferating SMCs. ECs on healthy SMCs showed enhanced capacities in adhesion, proliferation and migration.64,65 Likewise, engineering scaffolds with a biomimicking architecture of medial layer to regenerate a healthy native-like tunica media could also protect and support endothelial cell growth and maturation.5 Conversely, functionality of the ECs will be affected by the unhealthy status of vascular SMCs. Our data demonstrated that the inducive role of huaSMCs monolayer in promoting the proliferation and migration of the huvECs was undermined to some extent on stiffer AFSs (Fig. 8). This not only confirmed the existence of mutual interaction between SMCs and ECs, but also manifested the significant impact of pathologic vascular SMCs, as a result of extensive environmental (substrate) stiffness of the AFSs, on ECs’ proliferation and migration pertaining to endothelialization formation.

Inflammation plays a critical role in the vascular diseases relating to thrombus formation and neointimal hyperplasia.38,39 Thus we were interested in gauging how the stiffness of AFSs for vascular tissue engineering could alter inflammatory signatures of the huaSMCs. Interestingly, the primary macrophage-related genes (IL-6, MCP-1, and ICAM-1) of the huaSMCs on all the fibrous 25



substrates were detected. This revealed that higher stiffness of AFSs also promoted the SMCs to more-likely transit into a macrophage-like phenotype, although the relatively compliant PLCL fiber substrate also expressed inflammatory cytokines in the huaSMCs, indicative of activation of potential inflammatory response in vivo (Fig. 9). As a result, the macrophage-relevant proinflammatory cytokines presented in the vascular SMCs, would initiate recruitment and infiltration of leukocytes and macrophages66, and activate them to express more proinflammatory molecules. This ultimately disrupted the intact endothelial monolayer for thrombus formation39, upregulated the SMCs proliferation for neointimal hyperplasia38,39, and influenced ECM composition for arteriosclerosis.67 It is thus conjectured that the excessive stiffness associated inflammatory reaction could contribute to the malignant cell remodeling-resultant low long-term patency problem in the small-caliber TEBVs.

To summarize, to the best of our knowledge, this is the first aligned fibrous scaffolds model with tunable stiffness while keeping consistent surface chemistry and topographical features, to study purely the effects of aligned fiber stiffness on vascular SMCs. Although electrospun PLLA5,15 and PLCL19,68 fibers have been widely used in vascular tissue scaffolding, our results, as schematically delineated in Fig. 10, demonstrated that increasing stiffness of the AFSs had no significant influence on cell morphology, but modestly enhanced the density of internal F-actin fibers assembled in huaSMCs, which consequently led to improved capacity in cell proliferation and migration. Synthetic or pathologic characteristics of the huaSMCs were confirmed by the significant down-regulation of -SMA, SM-MHC, calponin and desmin and up-regulation of OPN genes. Meanwhile, higher fiber stiffness controlled the phenotypic alteration of huaSMCs from contractile to macrophage-like, as evidenced by the up-regulation of inflammatory gene expression. Presence of inflammatory mediators could have the inflammatory cells (e.g., leukocytes and macrophages) recruited around and consequently disrupted endothelial cell-cell junctions in favor of thrombus formation39,69 and upregulated SMC proliferation for neointimal hyperplasia38,39 in TEBVs, even if they are prior well-functionalized at surface. From a material selection perspective, it is plausible to suggest that electrospun fibers of the rigid PLLA compared to its counterpart of the PLCL, despite of being very soft in its micro/nano scale fineness, is not an ideal biodegradable for vascular tissue scaffolding. 26


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Fig. 10 Stiffness effects of AFSs on functional modulation in the huaSMCs

5. Conclusions

In this study, we successfully developed highly-aligned fibrous scaffolds with tunable stiffness by combining coaxial electrospinning and the SJES approaches. Through altering the flow rates of the PLLA-core and PLCL-shell solutions, modulus/stiffness of the aligned fibers was tuned in a wide range of up to 145 times, without imposing adverse influence to the fiber topography and surface chemistry. Higher stiffness of the aligned fibrous substrates was found to significantly encourage the proliferation and migration of huaSMCs, while morphologically had no significant effect on cell shape. More importantly, excessive fiber stiffness directed the huaSMCs into a de-differentiated, non-contractile, and pathological phenotype, which not only affected the proliferation and migration of huvECs but also promoted the expression of macrophage-related makers in the huaSMCs. This work exemplifies that fiber stiffness is an important parameter to consider in using electrospun aligned fibers for rational design of vascular grafts towards attaining cooperative effects of topological (alignment of electrospun fibers) and biomechanical (e.g., stiffness) features. In addition, our SJCES approach for generating highly-aligned fiber substrates with tunable stiffness can not only be applied to construct biomimetic 3D tubular vascular scaffolds, but also be utilized for establishing cardiovascular disease model for investigating 27



Page 28 of 34

pathological mechanism (e.g., atherosclerosis), or as a biomimetic platform to evaluate the cellular responses to fiber stiffness in other anisotropic tissue engineering researches.

Acknowledgements

This work was supported by the National Key Research and Development Program of China (2016YFC1100203),

the

Fundamental

Research

Funds

for

the

Central

Universities

(CUSF-DH-D-2018066) and the National Natural Science Foundation of China (31570969 and 31771050). Professor Yue Zhou from the College of Biomedical Engineering, Shanghai Jiaotong University is acknowledged for kindly providing the ECs and SMCs for our current study.

Disclosures

There

are

no

conflicts

of

interest

28


to

declare.

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