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Hybrid Biomimetic Interfaces Integrating Supported Lipid Bilayers with Decellularized Extracellular Matrix Components Setareh Vafaei, Seyed R. Tabaei, Vipra Guneta, Cleo Choong, and Nam-Joon Cho Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.7b03265 • Publication Date (Web): 28 Feb 2018 Downloaded from http://pubs.acs.org on March 1, 2018

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Table of Content 50x19mm (300 x 300 DPI)

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Hybrid Biomimetic Interfaces Integrating Supported Lipid Bilayers with Decellularized Extracellular Matrix Components Setareh Vafaeia,b, Seyed R. Tabaeia,b, Vipra Gunetaa, Cleo Choonga,c and NamJoon Choa,b,d,* a

School of Materials Science and Engineering, Nanyang Technological University, 50 Nanyang Avenue 639798, Singapore b Centre for Biomimetic Sensor Science, Nanyang Technological University, 50 Nanyang Drive 637553, Singapore c KK Research Centre, KK Women’s and Children’s Hospital, 100 Bukit Timah Road 229899, Singapore d School of Chemical and Biomedical Engineering, Nanyang Technological University, 62 Nanyang Drive 637459, Singapore

E-mail: [email protected]

Keywords: extracellular matrix; adipose tissue; decellularization; supported lipid bilayer; cell adhesion.

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Abstract This paper describes the functionalization of solid supported phospholipid bilayer with decellularized extracellular matrix (dECM) components, towards the development of biomimetic platforms that more closely mimic the cell surface environment. The dECM was obtained through a combination of chemical and enzymatic treatments of mouse adipose tissue that contains collagen, fibronectin and glycosaminoglycans (GAGs). Using amine coupling chemistry, the dECM components were attached covalently to the surface of a supported lipid bilayer containing phospholipids with reactive carboxylic acid headgroups. The bilayer formation and the kinetics of subsequent dECM conjugation were monitored by quartz crystal microbalance with dissipation (QCM-D). Fluorescence recovery after photobleaching (FRAP) confirmed the fluidity of the membrane after functionalization with dECM. The resulting hybrid biomimetic interface, supports the attachment and survival of the human hepatocyte Huh 7.5 and maintains the representative hepatocellular function. Importantly, the platform is suitable for monitoring the lateral organization and clustering of cell-binding ligands and growth factor receptors in the presence of the rich biochemical profile of tissue-derived ECM components.


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Introduction In response to the high complexity of native biological membranes and their microenvironments, various lipid-based model systems have been developed. The physical and chemical characteristics of these model systems are comparable to those of natural membranes 1. The lipid-based platforms, such as small 2, and giant lipid vesicles supported lipid bilayer (SLB)

6-8

3-4

, bilayer nanodiscs 5, and

, when functionalized with biological ligands, allows the

studying of membrane-associated cellular processes in simplified and controlled conditions. Amongst these lipid-based platforms, the SLB comprising of a single planar lipid bilayer associated with a solid substrate is particularly effective owing to its excellent mechanical stability while preserving the fundamental characteristics of natural membranes such as twodimensional fluidity and electrical sealing properties 9. In addition, the chemical composition of SLBs can be easily tuned to manipulate their biophysical properties such as surface charge and lipid diffusivity

11

, and their membrane organization such as phase separation

12

10

,

and

formation of lipid rafts 13. Importantly, SLBs can be functionalized with biological receptors and signaling ligands by introducing synthetic head-group modified lipids into the bilayer for direct covalent or noncovalent coupling of biomolecules

14

. Functionalized SLBs, have been used to examine range

essential biological processes that occur at the membrane interfaces including viral fusion

15

,

ligand-receptor interactions 16, enzymatic activities 17, cellular signaling and cell-cell recognition 18

. Owing to their inherent antifouling properties, SLBs are resistant to non-specific adsorption of

cells

19

and proteins

20

. This feature makes them a suitable interface for detection of specific

biomolecular interactions. In particular, they are effective tools for studying receptor dynamics during cell-cell interactions as well as triggering various intercellular reactions when interfaced


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with live cells

14, 18, 21

.

Moreover, a wide range of ultrasensitive surface-based analytical

techniques such as atomic force microscopy (AFM) dissipation (QCM-D) measurements

25

23

22

, quartz crystal microbalance with

, total internal fluorescence (TIRF) microscopy

24

, and electrochemical

, can be used to characterize the SLBs in terms of their structure and the

biochemical interactions that occur at their interface. While simplified models have been proven to be useful for many biological applications, development of models with greater compositional complexities would be the next step toward platforms that more closely mimic the cell surface environment. In this regard, functionalization of SLBs with the extracellular matrix (ECM) components holds great promise. The ECM is a tissue-specific network of extracellular bio-active molecules 26-27, that provides support and anchorage for cells and plays a key role in numerous cellular processes including cell growth and differentiation as well as intercellular communications composed of fibrous proteins such as collagens

29

, fibronectin

proteoglycans with glycosaminoglycans (GAGs) side-chains

30

32

28

. The ECM is mainly

, and laminin

31

, as well as

. These molecules provide

biomimetic adhesion sites for cell attachment, and their structural and biochemical properties make them ideal for cell and tissue engineering applications. Previously, ECM-derived molecules have been used to coat implants, functionalize surfaces, direct cell growth and differentiation, and engineer cell phenotype and behavior 33.

Numerous protocols have been developed to obtain natural ECM through decellularization of tissue via a series of physical, chemical and enzymatic treatments

34-36

. Decellularized ECM

(dECM) has shown to retain its natural 3D-ultrastructure and support the attachment and survival of cells while maintaining their phenotypic and functional characteristics. Thus, these


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biomaterials have attracted much attention as natural 3D-scaffolds for tissue engineering and regenerative medicine applications 37-45. Previously, SLBs functionalized with individual ECM components such as collagen fibronectin

47

, or peptides derived from these proteins (e.g., RGD, IKVAV)

48-53

46

, and

, have been

reported. Herein, for the first time, we demonstrate functionalization of SLBs with solubilized adipose tissue dECM and characterize their formation with QCM-D and immunofluorescence microscopy. We demonstrate that dECM functionalized bilayers sustain cell growth and maintain cell function.

Materials and Methods All chemicals used were purchased from Sigma (USA) unless otherwise stated.

1. Decellularization of White Adipose Tissue The white adipose tissue samples were extracted from cadavers of mouse (C57BL/6) and were decellularized with an in-house method using chloroform/methanol treatment. Briefly, the dissected tissue was cut into small pieces and washed several times with 1X PBS to remove all the blood and hair. During the first three days, the samples were treated with chloroform/methanol mixture at room temperature using different volumetric ratios: 2/1 (day 1), 1/1 (day 2) and 1/2 (day 3). Subsequently, the tissue samples were washed with 1X PBS several times and treated with trypsin (0.25%) for 24 h on a shaker at 37°C. Finally, the samples were washed with PBS and then treated with isopropanol for 48 h on a shaker at room temperature. The samples were stored in 1X PBS and supplemented with 1% antibiotic-antimitotic (Gibco, USA) at 4°C until further use.


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2. Solubilization of Decellularized Adipose Tissue Lyophilized ECM was crushed into powder form using a mortar and pestle. The powder (10 mg) was dissolved in 0.1 M HCl (1 ml) and mixed with 1 mg of pepsin (Sigma, Aldrich) as previously reported

54-57

for 48 h on a stirrer at 4°C. After that, the solubilized sample was

centrifuged and the supernatant was neutralized using 1 M NaOH solution. This ECM solution was then filtered through 0.2 µm syringe filter and used for integration into the lipid bilayer. 3. Adipose-Drived ECM Characterization 3.1.

DNA Content

The amount of DNA before and after decellularization was quantified using a Quant-iT™ PicoGreen® dsDNA Assay Kit (Life Technologies, USA) as per previously published protocols 58

. Briefly, lyophilized adipose tissues were treated with 1 ml 1X TE buffer (from the kit) and

0.1% Triton X-100. The samples were freeze-thawed three times (-80°C to 37°C). 100 µl of the aqueous working solution of the Quant-iT™ PicoGreen® reagent was mixed with 100 µl of the sample and was incubated for 5 min at room temperature. The fluorescence of the sample was measured at 480 and 520 nm using a plate reader (TECAN, Switzerland) and the DNA concentration was obtained from the standard curve.

3.2.

Morphology

Scanning Electron Microscopy (SEM) was utilized to study the morphology of adipose tissue sample before and after decellularization. Samples were prepared for SEM by fixing with 2.5% glutaraldehyde overnight at 4ºC. Fixed samples were subsequently dehydrated using ethanol series (20%, 40%, 50%, 70%, 90%, and 100%) for 5 min per grade. Further, the samples were immersed in 100% ethanol and and then Hexamethyldisilazane for 1 h each. The samples were loaded on a sample holder covered with carbon and copper tape, followed by coating with gold 6 ACS Paragon Plus Environment

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using a sputter coater (JEOL JFC-1600, Japan). The SEM (JSM-7600F-JEOL, Japan) was used to image the sample with an acceleration voltage of 5 kV at various magnifications.

3.3.

Total Protein Content

The total protein amount for the tissue samples before and after decellularization was determined by the bicinchoninic acid (BCA) assay using the Pierce™ BCA protein assay kit (Thermo Scientific, USA) according to the manufacturer’s manual. Briefly, lyophilized tissue was immersed in radioimmunoprecipitation (RIPA) buffer supplemented with protease and phosphatase inhibitor at 4°C for 30 min. The samples then were homogenized using a probesonicator (Qsonica, USA). After homogenization, the samples were centrifuged at 10,000 g for 20 min at 4ºC and the supernatant was transferred to a new Eppendorf for measuring the protein content. 25 μl of the protein samples were added in a 96-well microplate and mixed with 200 μl of BCA working reagent. The plate was shacked for 30 s and subsequently incubated at 37ºC for 30 min. After cooling the plate, the light absorbance of the samples were measured at 562 nm using a plate reader (TECAN, Switzerland) and the protein concentration was calculated against the standard curve. The amount of proteins was normalized to dry-weigh tissue mass.

3.4.

Collagen Content

The collagen content of intact and decellularized samples was quantified by measuring the hydroxylproline content of the samples, using hydroxylproline assay, according to the manufacturer’s manual. The hydroxylproline contents of the samples were converted to the amount of collagen. Briefly, the samples were hydrolyzed in 12M HCl at 120ºC for 3 h. The samples were dried under vacuum and mixed with equal volume of chloramine T/Oxidation buffer and incubated for 5 min. Finally, dimethylaminobenzaldehyde (p-DMBA) colorimetric solution was added to each sample and incubated for 90 min at 60 ºC. The absorbance of the colorimetric product


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was measured at 560 nm using a microplate reader (TECAN, Switzerland). The concentration of hydroxyproline was calculated based on the standard curve and converted to Col content based on the assumption that hydroxyproline constitutes 13.5% of collagen dry mass 59.

3.5.

GAG Content

The Blyscan™ GAG assay (Biocolor, UK) was used to measure the amount of total sulfated glycosaminoglycans (sGAG) in adipose tissue sample before and after decellularization according to the manufacturer’s manual. Briefly, the lyophilized tissue sample was incubated with 1 ml papain extraction reagent at 65°C overnight. The sample was centrifuged at 10,000 g for 10 min and the supernatant was separated. 100 μl of the sample was mixed with 1 ml of Blyscan dye reagent and mixed thoroughly on a plate shaker for 30 min at room temperature and subsequently was centrifuged at 10,000 g for 10 min. The colored pellet was separated from the supernatant and was dried completely. The pellet was dissociated using 0.5 mL of dissociation reagent and was vortexed. The solution was centrifuged at 10,000 g for 5 min. 200 μl of the solution was transferred inside a 96-well microplate and light absorbance was measured at 656 nm using a plate reader (TECAN, Switzerland). The amount of GAG was calculated based on the standard curve.

4. Lipid Bilayer Formation The supported lipid bilayer was prepared using solvent-assisted lipid bilayer (SALB) formation method 60. Zwitterionic lipid , 1, 2-dioleoyl-sn-glycero-3-Phosphatidylcholine (DOPC) and a head-group modified lipid, 2-dipalmitoyl-sn-glycero-3 phosphoethanolamine-N-(glutaryl) (DP-NGPE) were obtained in powder from Avanti Polar Lipid (USA). Immediately before the experiment, 10 mg DOPC and 5mg DP-NGE were dissolved in 1 ml isopropanol and ethanol respectively. To aid dissolution of DP-NGPE in ethanol, the solution was heated up to 60ºC for 8 ACS Paragon Plus Environment

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30 sec and vortexed gently. This process was repeated several times to a transparent solution was obtained. The stock solutions were diluted using isopropanol to prepare 0.5 mg/ml lipid mixture containing 80% DOPC and 20% DP-NGPE. 200 µl of the lipid mixture was injected into a microfluidic channel (sticky-slide I 0.4 Luer- ibidi) which has a clean hydrophilic glass (glass coverslip, ibidi) (25 mm × 75 mm × 170 μm) at the bottom. The lipid solution was incubated in the channel for 10 min and was gradually replaced by Tris buffer (10 mM Tris, 100 mM NaCl, pH 7.5). After complete replacement of the buffer with isopropanol, an SLB was formed on the glass surface.

5. Activation of DP-NGPE Lipids and SLB Functionalization with ECM The carboxylic group of functionalized lipid incorporated in SLBs was activated using Nhydroxysulfosuccinimide sodium (sNHS) (Alfa Aesar, USA) and 1-(3-dimethylaminopropyl)-3ethylcarbodiimide hydrochloride (EDC) (Alfa Aesar, USA) for amine coupling. A mixture of 60 mg EDC/NHS in 3 ml HEPES buffer (10 mM HEPES,100 mM NaCl, pH 5.5) was freshly prepared and introduced into the sample and incubated for 1 h at room temperature followed by injection of HEPES buffer. Solubilized ECM solution was prepared in pH 7.5 PBS (10mM Phosphate buffer, 2.7 mM KCL, and 150 mM NaCl) at a concentration 1 mg/ml. Protein solutions were introduced to the activated SLB and incubated for about two hours to form dECM functionalized SLB. Finally the substrates were washed with PBS to remove unbounded proteins.

6. Quartz Crystal Microbalance with Dissipation Monitoring The quartz crystal microbalance with dissipation (QCM-D) technique was used as a surface sensitive technique to measure and monitor the change of resonance frequency (ΔF) and energy dissipation (ΔD) caused by the adsorption of lipid and proteins on silicon oxide-coated quartz


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crystals. Before the experiment, the quartz crystals were immersed in 1% sodium dodecyl sulfate (SDS) (Sigma, USA) for 2 h and subsequently washed with Milli-Q water (Millipore). After drying the crystals with nitrogen gas, they were placed in an oxygen plasma (March Plasmod Plasma Etcher, March Instruments, California) at 180 W for 1 min. This step was done to clean the surface of the crystal from residual organic molecules and also to make the surface hydrophilic. All measurements were done using a Q-Sense E4 instrument (Q-sense AB, Gothenburg, Sweden) under continuous flow condition with a flow rate of 50 μL/min. The flow condition was conducted and controlled using a peristaltic pump (Ismatec Reglo Digital). QCMD measurements were taken at the 3, 5, 7 and 11 overtones, and the presented data was measured and normalized at the third overtone [15 MHz (ΔFn=3/3)].

7.

Fluorescence Recovery after Photobleaching (FRAP) Estimation of the diffusion coefficient of the lipid molecules in the presence and absence of

proteins and cells was carried out by the Flourescence Recovery after Photobleaching (FRAP), as per

previously

established

61

.

Briefly,

0.5

wt

%

of

1,

2-dioleoyl-sn-glycero-3-

phosphoethanolamine-N-(lissamine rhodamineB sulfonyl) (ammonium salt) (Rh-PE) (Avanti Polar Lipid) was added to the lipid mixture of 80% DOPC and 20% DP-NGPE. After formation of the bilayer on the glass surface, a 532 nm, 100 mW laser beam (Klastech Laser Technologies, Germany) was used to bleach a defined area of SLBs for 5 sec that result in the formation of a ~20 µm bleached spot. The flourescence recovery was monitored and measured using an epifluorescence Eclipse TE 2000 microscope (Nikon, Japan) combined with an Andor iXon+ EMCCD camera (Andor Technology, Northern Ireland). All images were collected at a rate of 1 image per two-second intervals. Images were analyzed using ImageJ software and diffusion coefficients were measured using the Hankel transform method . 10 ACS Paragon Plus Environment

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8. Cell Culture and Cell Seeding on SLB Platform Human hepatocarcinoma cells (Huh 7.5), purchased from Apath (USA), were cultured in a humidified atmosphere with 5% CO2 at 37°C and maintained in Dulbecco’s Modified Eagle’s Medium (DMEM) (Gibco, USA) supplemented with 10% fetal bovine serum (FBS)(Gibco, USA), 100 U/ml penicillin and 100 μg/ml streptomycin (Gibco, USA). The media was changed every three days. Cells were detached using 0.25 % trypsin (Gibco, USA) and suspended in media. The total cell number was counted by a hemocytometer. 20,000 cells were seeded onto each of the supported bilayers and cultured for three days, with media changes every 24 h.

9. Cell Surface Area The cell surface area of Huh 7.5 cells was quantified after 24 h. An inverted optical microscope (Nikon, Japan) with a 10× objective was utilized to capture the Huh 7.5 images on the bare and ECM-functionalized SLBs. All images were analyzed, and the average cell area was measured using imageJ software.

10. Cell Viability Viability of Huh 7.5 cells was observed qualitatively using LIVE/DEAD assay (Life Technologies) as per previously established protocols

62

.

Briefly, cells cultured on ECM-

functionalized SLBs were washed with PBS and treated with 2 µM clacein AM and 4 µM EthD1 in DMEM media at 24 and 72 h timepoints. Cells were incubated with the assay solution for 20 min at 37°C and subsequently washed with 1X PBS before visualization under an epifluorescence Eclipse TE 2000 microscope (Nikon, Japan) combined with an Andor iXon+ EMCCD camera (Andor Technology, Northern Ireland). The living cells were stained with calcein and emitted a strong green fluorescence that was visualized by a 494-517 nm emission


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filter. In contrast, the dead cells were stained with EthD-1 and emitted a red fluorescent that was imaged using a 517-617 nm emission filter.

11. Cell Proliferation Cell numbers were measured for Huh 7.5 cells cultured on ECM-functionalized SLB for day 1 and day 3 using cell counting kit-8 (CCK-8) (Dojindo Molecular Technologies, Japan) using previously established protocols 63. In brief, the cells were incubated with the CCK-8 solution for 1 h at 37°C. Absorbance of a range number of cells was plotted for a standard curve, which was used to calculate the number of viable cells present on the ECM-functionalized SLB on the different timepoints. The experiment was performed in triplicates and the cell proliferation was determined by normalizing the cell number at day 3 to the cell number at day 1.

12. Albumin Production The amount of human albumin secreted into the cell culture media by the Huh 7.5 cells cultured on SLB platforms was determined using an enzyme-linked immunosorbent assay (ELISA) kit (Abcam, USA), 24 h after seeding. The amount of albumin produced was normalized to the number of viable cells on the platforms at respective timepoints.

13. Visualization of ECM Proteins In order to qualitatively visualize the presence of collagen type I and fibronectin in the SLBs functionalized with ECM (dECM-SLB), the sample was stained with mouse monoclonal anticollagen type I (Abcam, USA) and mouse monoclonal IgG1 anti-fibronectin (Santa Cruz, USA), following the incubation time (~ 1 hour) at room temperature. The samples were washed with PBS properly and subsequently the secondary antibodies Alexa Fluor® 488 Goat Anti-mouse IgG (H+L) (green-fluorescent dye, LifeTechnology) and Alexa Fluor® 555 Rabbit Anti-Mouse 12 ACS Paragon Plus Environment

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IgG (H+L) (red-fluorescent dye, LifeTechnology) were added to the sample to fluorescently stain the collagen type I and fibronectin antibodies respectively. Fluorescence images were captured with an inverted epifluorescence Eclipse TE 2000 microscope (Nikon, Japan) equipped with a 60× oil immersion objectives (NA 1.49) in combination with an Andor iXon+ EMCCD camera (Andor Technology, Northern Ireland).

14. Statistical Analysis All experiments were carried out in triplicates unless otherwise stated. The one-way analysis of variance (ANOVA) was used to determine whether there are any statistically significant differences between the means of independent groups. The one-way ANOVA was performed using PRISM software. All data are represented as means ± SD. **, p < 0.01; ****, p < 0.0001.

Results and Discussion Multiple steps involved in the preparation of the dECM based supported lipid bilayer platform as a cell surface mimetic are summarized in Fig. 1. The first step is to decellularize the collagenrich ECM of adipose tissue

64

. Adipose tissue has been extensively used as a bioactive coating

material for cell studies65-67. The ECM components of adipose tissue has been comprehensively reviewed

68

; notiacably adipose tissue is enriched in collagen proteins. An effective

decellularization process allows us to achieve the maximum removal of cellular components with minimum ECM loss and damage. We used a combination of organic solvent extraction and enzymatic digestion processes. Briefly the whole-tissue biopsy from mouse adipose tissue was subjected to a sequential treatment with organic solvent for fat removal and trypsin to assist cell detachment (Refer to materials and method for detailed protocol).


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Fig. 1. Schematic illustration of various steps involved in the preparation of supported bilayer functionalized with adipose tissue dECM components for cell study applications. Adipose tissue was decellularized using a combination of chemical and enzymatic treatment. The decellularized ECM was then solubilized in acidic condition and was covalently attached via amine coupling to the surface of a supported bilayer containing lipids with head group bearing free carboxylic acid. Subsequently, cells were seeded and their attachment, spreading, growth and function were studied.

Next, we examined the effectiveness of the decellularization process by visualizing the ultrastructure of the matrix and determining its chemical composition. The matrix visualization was done by scanning electron microscopy (SEM). Fig. 2a and 2b shows the micrographs of adipose tissue before and after the decellularization process, respectively. Fig. 2b shows that no cell or cell fragments were present after decellularization, indicating efficient cell removal. In addition, the discrete organized microfiber structure of the collagen matrix (Fig. 2c), indicates that the overall ultrastructure of the dECM was preserved. The efficiency of the cell removal from the adipose tissue was also evaluated via quantification of the residual DNA. Only 7.7 ± 2.2 ng of DNA was retained in each mg of dry tissue (Fig. 2d) corresponding to about 98% reduction in the DNA level of the sample. Additionally, the dECM components; collagen and GAGs was quantified (Fig. 2e and 2f) as the amount per content of total protein in each sample. The normalized collagen content for dECM was higher than that of the native tissue (Fig. 2e), indicating that collagen was enriched during the decellularization process. This phenomenon is expected as the removal of cellular and lipid material results in the concentration of the


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collagenous ECM after decellularization (Fig. 2c). The normalized value for GAGs was almost unaltered indicating that they were preserved during the decellularization process.

Fig. 2. Characterization of decellularized mouse-derived adipose tissue. (a) Representative SEM image of native adipose tissue. (b) Low magnification SEM image of decellularized adipose tissue indicates a three-dimensional scaffold composed of fibrous ECM material that is devoid of cells and lipids. (c) High magnification SEM image of decellularized tissue demonstrates a three-dimensional architecture of the collagen fibers. Biochemical evaluation of the content of (d) DNA, (e) collagen and (f) GAGs present in the native and decellularized adipose tissue. (n = 3, mean ± SD; **: P < 0.01 and ****: P < 0.0001).

After decellularization, the remaining ECM matrix was solubilized completely by acid treatment 69. The solubilized dECM then were used for functionalization of the SLB. The dECM proteins were attached to the SLB via covalent bonding between free amine present in the protein components of the dECM matrix and free carboxylic acid present in the head group of a functional lipid (DP-NGPE) incorporated in the bilayer. A similar approach was previously used to functionalize the SLB with purified collagen and fibronectin

46-47

. Application of lipids with

reactive head groups such as amine, carboxylic acid, thiol, or maleimide are an efficient approach for ligation of lipid bilayers and have been used successfully for surface functionalization of supported bilayers and lipidic nanoparticles such as liposomes 70. 15 ACS Paragon Plus Environment

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We used the solvent assisted lipid bilayer (SALB) formation method to fabricate lipid bilayer 60

. The method involves the adsorption of lipid molecules onto a solid surface in the presence of

alcohol (e.g., isopropanol) followed by gradual replacement of the lipid-alcohol mixture with an aqueous buffer solution which leads to lipid phase transition in bulk and finally formation of lipid bilayer membrane on solid support surface

71

. The SALB method enables fabrication of

SLBs on various hydrophilic solid surfaces, including those which are not suitable for vesicle fusion method 72. In addition, the method allows the formation of SLBs with lipid compositions which cannot be formed using the vesicle fusion method (e.g., containing a high fraction of cholesterol) 11. QCM-D was used to monitor SLB formation and the kinetics of adsorption of solubilized adipose dECM to the bilayer. QCM-D is a real-time surface sensitive technique commonly used to monitor the kinetics of thin film formation and characterization of its thickness and viscoelastic properties. The method measures changes in the resonance frequency (ΔF) and energy dissipation (ΔD) of an oscillating piezoelectric quartz crystal coated with desired material. The ΔF and ΔD are related to the mass and the viscoelastic properties (rigidity and softness) of the add layer, respectively. It is widely used in soft matter research and interfacial science applications modification

75

solid supports

73-74

, particularly for studying lipid bilayer formation and it’s in situ

. The method has also been used for studying cell attachment and spreading on 76-77

. Fig. 3a, shows the QCM-D frequency and energy dissipation responses

corresponding to different steps involved in the formation of a DOPC bilayer containing 20% functional lipid (DP-NGPE) via the SALB method (arrow i-iii), activation of the free carboxylic acid via incubation EDC/NHS (arrow iv) and the subsequent addition of protein to the bilayer surface (arrow v). After complete solvent exchange (arrow iii), a ΔF and ΔD shifts of −25 ± 2


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Hz and 0.5 ± 0.3 × 10−6, respectively, were obtained, which correspond to the formation of a planar bilayer

75

. The close to zero energy dissipation indicates very low friction loss because

water molecules rest on the bilayer surface. Fig. 3b, shows an enlarged view of the QCM-D curve corresponding to the protein adsorption step (arrow v in Fig. 3a). The addition of dECM to the bilayer led to a negative frequency shift of ~-60 Hz. The negative frequency shift indicates an increase in the mass of the adlayer due to the protein adsorption. As a control, the adipose dECM was added to a bilayer containing no functional lipid (marked with blue triangles), showing almost no change in the frequency shift indicating no adsorption. Thus, adsorption of proteins to the bilayer containing functional lipids was due to the formation of covalent bond between proteins and the lipids with reactive headgroups. The relatively high energy dissipation of the protein adlayer is related to the frictional losses due to a conformation changes in the protein structure

20

. These changes are

reversible and are caused by the periodic shear motion of the QCM crystal surface

78

. Since

protein layers are not perfectly rigid, the relationship between frequency shift and adsorbed mass may not obey the Sauerbrey equation 79. Thus, we used the QCM-D experimental data collected at the 3rd, 5th, 7th, and 9th overtones and analyzed them by using the Voigt-Voinova model 80, as previously reported 81. Using this approach a surface mass density of 2120 ± 40 (ng/cm2) for dECM was obtained. It should be noted that the calculated mass density corresponds to both attached proteins and the associated water mplecules couples to the protein layer. This observation indicates that the protein layer on supported bilayer is soft and relatively high amount of water molecules are coupled to its structure. The softness and structural flexibility of protein layer are expected as they attached to the underlying bilayer surface through a limited number of anchoring points. The anchoring points are limited free amine-bearing lysine


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residues which are dispersed along the protein structure. Thus, while the attachment of proteins to the underlying bilayer is firm, the protein molecules retain their flexibility. It is noteworthy that the DOPC bilayer is zwitterionic and has an antifouling property and thus is resistant to nonspecific protein adsorption 20. Additional confirmation of the attachment of dECM components to the bilayer was achieved by immunofluorescence microscopy. Fig. 3c and d presents background-corrected fluorescence microscopy images for an ECM-coated bilayer incubated with fluorescently labeled anti-collagen type I and anti-fibronectin antibody, respectively. The results show that both collagen type I and fibronectin are present in the decellularized ECM matrix. In addition to the uniformly distributed fluorescent signal, there are randomly distributed brighter spots indicative of the presence of antibody clusters. The presence of antibody clusters suggests the presence of protein aggregates on the SLB surface. The protein aggregates might be due to the presence of non-solubilized proteins or defects in the underlying bilayer. The membrane fluidity is a crucial property of biological membranes. Importantly, despite the strong association of the supported bilayer with the solid surface, individual lipid molecules are highly mobile in the plane of the membrane

82

. To check if the underlying lipid bilayer after

functionalization with adipose-dECM retained its fluidity, FRAP experiment was carried out. Representative fluorescence micrographs, (Fig. 3e), confirm the retention of the long-range lateral fluidity of lipids with a lipid diffusivity of 2.0 ± 0.1 μm2/s after functionalization with adipose-dECM proteins.


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Fig. 3. Modification of SLBs using adipose dECM. (a) The QCM-D signal exhibited different steps of bilayer fabrication and functionalization. Frequency shift (ΔF, blue) and energy dissipation shift (ΔD, red) for the third overtone (n = 3) were measured as a function of time during the formation of 20% DP-NGPE SLB on SiO2 using SALB method. After establishing the baseline in a buffer solution, pure isopropanol was injected (arrow i). Since, the viscosity and density of isopropanol differs from that of buffer, dramatic changes in the frequency and dissipation were observed after injection. Then, 0.5 mg/ml lipid solution [DOPC:DP-NGPE (4:1)] (arrow ii) was introduced which caused a small change in resonance frequency and no change in the energy dissipation. Subsequently, the buffer was injected (arrow iii) leading to a final ΔF and ΔD of ~ -25 ± 2 Hz and 0.5 ± 0.3 × 10−6 , respectively which corresponds to a planar bilayer. In order to functionalize SLBs with ECM proteins, EDC/NHS was introduced into the chamber (arrow iv) to activate free carboxylic acid of DP-NGPE. In the last step, the dECM was injected (arrow v). The significant change in ΔF and ΔD indicates the adsorption of dECM to the SLB. (b) QCM-D characterization of the dECM binding to bilayers containing no (triangles) or 20 % DP-NGPE (rectangles). Fluorescence immunostaining micrographs of stained (c) collagen type I and (d) fibronectin present in dECM functionalized bilayer. The proteins were stained using fluorescently-labeled anticollagen type I and anti-fibronectin antibodies. (e) False-color images of fluorescence recovery after photobleaching analysis of dECM functionalized bilayer. A ~20 μm-wide circular spot was photobleached, followed by a time-lapse recording. Fluorescence was recovered nearly completely after 60 s suggesting bilayer retains its mobility.

Next, we studied the attachment and growth behavior of Huh 7.5 cells on the dECM functionalized bilayer platform. Huh 7.5 cells are a subclone of Huh 7 cells derived from hepatocyte cells in the liver tissue which shows liver specific functions such as albumin production83. 19 ACS Paragon Plus Environment

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Cell adhesion to the ECM is mediated mainly by integrins present at the cell surface. Different ECM component such as collagen and fibronectin possess specific integrin binding domains which facilitate cell attachment and subsequent intracellular signaling responsible for cell survival 84, proliferation 85, differentiation 86, and migration 87. Fig. 4a represents typical bright field image of Huh 7.5 cells seeded on bare SLB, dECM functionalized bilayer (dECM-SLB) and dECM-coated glass substrates (dECM-Glass) after 24 h. On bare supported bilayer with no dECM, only a limited number of cells attached and remained rounded and did not appreciably spread over 24 hours. This is expected and consistent with previous reports showing that the antifouling properties of the zwitterionic bilayer prevent the adsorption of proteins 20, as well as cells 19. In contrast, cells attached and spread well on dECMSLB, indicating that the functionalization of the bilayer with dECM resulted in the specific adhesion of the cells presumably due to the interaction between cells and ECM molecules. On the glass, cells attached and spread on ECM-coated surface. Quantification of images (Fig. 4b and c), shows that the number of cells attached to the dECM-SLB (131 ± 27 cells/mm2) as well as their projected area (1120 ± 521 µm2/cell) was more than twice of that of bare SLB. For cells plated on dECM-Glass, the cell number was not significantly different from that of dECM-SLB. However consistent with our previous observations 88, the projected area of cells adhered to glass substrate (1501 ± 602 µm2/cell) was slightly higher than that of its SLB counterpart (dECM-SLB). This is because dECM molecules on SLBs which are covalently linked to the lipid molecules in the fluid lipid bilayer, are less resistant to displacement when experience force from the cell 88. We have previously reported functionalization of SLBs and glass substrates with individual ECM proteins (e.g. collagen)88-89. Noticeably, the cell number and the projected area of the cells


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on dECM-SLB were similar to that of SLB functionalized with collagen fibers (see supporting Fig. S1). Consistent with our previous observations, in all studies the cell surface area was higher on glass substrates in comparison to their SLB counterparts88-89. Taken together, the dECM components support the growth and adhesion of Huh 7.5 cells on the lipid bilayer platform. We further checked if the bilayer retained its mobility after the cell seeding, by performing FRAP on membrane area underneath the cell. Fig. 4e shows that the bilayer retained its mobility underneath the cell. However, the lipid diffusivity of the bilayer underneath the cell (1.4 ± 0.05 µm2/s) was lower than that of areas with no cells (2 ± 0.1 µm2/s). Such decrease in the lipid diffusivity is expected as the adsorption of macromolecules to lipids hinders the lateral mobility of the interacting lipids in the underlying bilayer 90-91.


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Fig. 4. Huh, 7.5 Cells behavior on bare and dECM-functionalized SLBs. (a) The bright filed images of Huh 7.5 growth on bare SLB, dECM-functionalized SLB (dECM-SLB) and dECM-coated glass substrates (dECM-Glass) after 24 h, respectively. The morphology of Huh 7.5 cells cultured on bare SLB are rounded, while those cultured on dECM-functionalized SLBs and dECM-coated glass substrate show increase in spread area. (b) Quantification of cell number plated on bare SLB, dECM-SLB and dECMGlass. dECM-SLB promotes higher cell attachment (~two times) compare to bare SLB after 24 h. (c) The measurement of Huh 7.5 cell spreading size on bare SLB, dECM-SLB and dECM-Glass after 24 h. The average area of a cell was measured based on the average value of at least 200 cells. The images were taken by an optical microscope and analyzing with imageJ. The observation field was 0.5 mm2. (d) Falsecolour images of fluorescence recovery after photobleaching analysis of ECM-functionalized SLBs after cell adhesion. A 20μm-wide circular spot was photobleached under the cell, followed by a time-lapse recording. Fluorescence was recovered nearly completely after 80 s suggesting bilayer retains its mobility under ECM material and cells. The cell culture studies as well as FRAP experiments were done inside microfluidic channels. (n = 3, mean ± SD; ****: P < 0.0001).

To further investigate the biological activity of dECM on SLB, we examined the viability, proliferation, and function of the cells plated on this platform. The cell viability was quantified 22 ACS Paragon Plus Environment

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using live/dead assay combined with fluorescence microscopy. Over 90% of Huh 7.5 cells were viable within three days, as shown in Fig. 5a. The cell proliferation measured using CCK-8 assay, showed that the number of cells after 72 h was about 1.5 times the number of the cells after 24 h (Fig. 5b). We further compared the cell proliferation on dECM-SLB and bare glass as well as glass substrate coated with dECM (dECM-Glass). Overall, there was no significant difference between Huh 7.5 proliferation on these platforms (see supporting Fig. S2). The interaction between of cells and ECM component control many cellular activities including tissue-specific expressions

92

. Albumin production by hepatocyte is one such cellular

function which is affected by cell-ECM interaction 93. To check the activity of Huh 7.5 cells, we measured the amount of albumin secreted by cells grown on dECM coated bilayer. The average amount of albumin produced by the cells after 24 h was 3.1 ± 0.2 µg per 106 cells, which is in the range of our results obtained on culture dishes (Fig. 5b).


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Fig. 5. Evaluation of Huh 7.5 cells growth and functions on ECM-functionalized SLB. (a) Fluorescence image of calcein AM-stained Huh 7.5 cells with live cells fluorescing green and dead cells fluorescing red after 24 and 72 h plated on ECM-functionalized SLB. The viability of the cells was over than 90% within three days. (b) Quantification of cell viability, cell proliferation and albumin production of Huh 7.5 on ECM-functionalized SLB. Cell viability was analyzed using imageJ. Cell proliferation amount was calculated by normalization of cell number at day three to the cell number at day one. The amount of albumin was obtained by normalization of albumin secretion at day 1 to 106 cells.


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4. Conclusion In summary, a cell-surface mimetic substrate consisting of a solid-supported lipid bilayer functionalized with decellularized adipose tissue ECM components was fabricated. We demonstrated that our previously developed protocol for functionalization of SLBs with pure synthetic ECM molecules (Col I and FN)88-89 works equally as well with natural dECM. The adipose tissue was successfully decellularized using a combination of chemical and enzymatic processes. The remaining ECM components were attached to the surface of a supported lipid bilayer via covalent bonding using amine coupling methodology. The attachment process and the final surfaces, were characterized by QCM-D and immunofluorescence microscopy. The ECM components attached to the bilayer retained their biochemical functionality and promoted cell adhesion and proliferation to the otherwise non-fouling cell resistant phospholipid bilayer. The cell binding activity is known to be mainly mediated via RGD which is the principal integrin-binding domain present within ECM proteins such as fibronection and collagen type I. Therefore, our results suggest that at least RGD ligands were preserved after re-solubilized in acidic conditions. FRAP analysis revealed that the underlying bilayer retained its fluidity after ECM conjugation as well as cell anchorage and spreading. SLBs functionalized with proteins have been shown to be an enabling platform for studying cell-cell interactions, thanks to the fluidity of the underlying supported bilayers. For instance, the membrane fluidity has been exploited to examine the spatial reorganization of immune recognition receptors and cluster formation that occur during the immune signaling processes18, 21

. In this context, the fact that lipids in the system developed in this work are laterally mobile,

enables studying such process in a platform that more closely mimics the in vivo extracellular environment.


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In comparison to standard plastic and glass culture plates that only support physical attachment of adhering cells, ECM-functionalized surfaces not only provide mechanical support but can also transmit biological signals to cells through biological recognition between protein and proteoglycan receptors on cell surfaces with ECM molecules. For instance, the ECM is an important component of the stem cell niche that regulates cell differentiation and function. Therefore, while basic performance features such as cell attachment and proliferation were comparable on dECM-SLB platforms to that of other examined platforms, we envisage that the dECM-SLB platform will outperform bio-inert platforms in more biologically-demanding cases where ECM-mediated responses are essential.

Acknowledgemnnts This work was supported by the National Research Foundation (NRF-NRFF2011-01). References (1) Chan, Y.-H. M.; Boxer, S. G., Model membrane systems and their applications. Curr. Opin. Chem. Biol. 2007, 11 (6), 581-587. (2) Christensen, S. M.; Stamou, D. G., Sensing-applications of surface-based single vesicle arrays. Sensors 2010, 10 (12), 11352-11368. (3) Walde, P.; Cosentino, K.; Engel, H.; Stano, P., Giant vesicles: preparations and applications. ChemBioChem 2010, 11 (7), 848-865. (4) Furuki, T.; Watanabe, T.; Furuta, T.; Takano, K.; Shirakashi, R.; Sakurai, M., The Dry Preservation of Giant Vesicles Using a Group 3 LEA Protein Model Peptide and Its Molecular Mechanism. Bull. Chem. Soc. Jpn 2016, 89 (12), 1493-1499. (5) Nath, A.; Atkins, W. M.; Sligar, S. G., Applications of phospholipid bilayer nanodiscs in the study of membranes and membrane proteins. Biochemistry 2007, 46 (8), 2059-2069. (6) Edward T. Castellana; Cremer, P. S., Solid supported lipid bilayers: From biophysical studies to sensor design. Surf. Sci. Rep. 2006, 61, 429–444. (7) Ariga, K.; Yamauchi, Y.; Mori, T.; Hill, J. P., 25th Anniversary Article: What Can Be Done with the Langmuir‐Blodgett Method? Recent Developments and its Critical Role in Materials Science. Adv. Mater. 2013, 25 (45), 6477-6512. (8) Takeshita, N.; Okuno, M.; Ishibashi, T.-a., Molecular conformation of DPPC phospholipid Langmuir and Langmuir–Blodgett monolayers studied by heterodyne-detected vibrational sum frequency generation spectroscopy. Phys. Chem. Chem. Phys. 2017, 19 (3), 2060-2066. 26 ACS Paragon Plus Environment

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(9) Sackmann, E., Supported membranes: Scientific and practical applications. Science 1996, 271 (5245), 43-48. (10) TaeWoon Cha, A. G., y and X.-Y. Zhu, Formation of Supported Phospholipid Bilayers on Molecular Surfaces: Role of Surface Charge Density and Electrostatic Interaction. Biophys. J. 2006, 90, 1270–1274. (11) Tabaei, S. R.; Jackman, J. A.; Kim, S.-O.; Liedberg, B.; Knoll, W.; Parikh, A. N.; Cho, N.-J., Formation of Cholesterol-Rich Supported Membranes Using Solvent-Assisted Lipid SelfAssembly. Langmuir 2014, 30 (44), 13345-13352. (12) Tabaei, S. R.; Jackman, J. A.; Liedberg, B.; Parikh, A. N.; Cho, N.-J., Observation of Stripe Superstructure in the β-Two-Phase Coexistence Region of Cholesterol–Phospholipid Mixtures in Supported Membranes. J. Am. Chem. Soc. 2014, 136 (49), 16962-16965. (13) Kraft, M. L.; Weber, P. K.; Longo, M. L.; Hutcheon, I. D.; Boxer, S. G., Phase separation of lipid membranes analyzed with high-resolution secondary ion mass spectrometry. Science 2006, 313 (5795), 1948-1951. (14) Yu, C.-h.; Groves, J. T., Engineering supported membranes for cell biology. Med Biol Eng Comput 2010, 48 (10), 955-963. (15) Floyd, D. L.; Ragains, J. R.; Skehel, J. J.; Harrison, S. C.; van Oijen, A. M., Singleparticle kinetics of influenza virus membrane fusion. Proc. Natl. Acad. Sci. USA 2008, 105 (40), 15382-15387. (16) Jung, H.; Robison, A. D.; Cremer, P. S., Multivalent ligand–receptor binding on supported lipid bilayers. J. Struct. Biol. 2009, 168 (1), 90-94. (17) Tabaei, S. R.; Guo, F.; Rutaganira, F. U.; Vafaei, S.; Choong, I.; Shokat, K. M.; Glenn, J. S.; Cho, N.-J., Multistep Compositional Remodeling of Supported Lipid Membranes by Interfacially Active Phosphatidylinositol Kinases. Anal. Chem. 2016, 88 (10), 5042-5045. (18) Groves, J. T.; Dustin, M. L., Supported planar bilayers in studies on immune cell adhesion and communication. J. Immunol. Methods 2003, 278 (1-2), 19-32. (19) Andersson, A. S.; Glasmästar, K.; Sutherland, D.; Lidberg, U.; Kasemo, B., Cell adhesion on supported lipid bilayers. J. Biomed. Mater. Res., Part A 2003, 64 (4), 622-629. (20) Glasmästar, K.; Larsson, C.; Höök, F.; Kasemo, B., Protein adsorption on supported phospholipid bilayers. J. Colloid Interface Sci. 2002, 246 (1), 40-47. (21) Manz, B. N.; Groves, J. T., Spatial organization and signal transduction at intercellular junctions. Nat. Rev. Mol. Cell Biol. 2010, 11 (5), 342-352. (22) Milhiet, P. E.; Vie, V.; Giocondi, M. C.; Le Grimellec, C., AFM characterization of model rafts in supported bilayers. Single Mol 2001, 2 (2), 109-112. (23) Edvardsson, M.; Svedhem, S.; Wang, G.; Richter, R.; Rodahl, M.; Kasemo, B., QCM-D and reflectometry instrument: applications to supported lipid structures and their biomolecular interactions. Anal. Chem. 2008, 81 (1), 349-361. (24) Dewa, T.; Sugiura, R.; Suemori, Y.; Sugimoto, M.; Takeuchi, T.; Hiro, A.; Iida, K.; Gardiner, A. T.; Cogdell, R. J.; Nango, M., Lateral organization of a membrane protein in a supported binary lipid domain: Direct observation of the organization of bacterial lightharvesting complex 2 by total internal reflection fluorescence microscopy. Langmuir 2006, 22 (12), 5412-5418. (25) Navrátil, T.; Šestáková, I.; Štulík, K.; Mareček, V., Electrochemical measurements on supported phospholipid bilayers: preparation, properties and ion transport using incorporated ionophores. Electroanal 2010, 22 (17‐18), 2043-2050.


Page 28 of 32

Page 29 of 32 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

(26) Hynes, R. O., The extracellular matrix: not just pretty fibrils. Science 2009, 326 (5957), 1216-1219. (27) Frantz, C.; Stewart, K. M.; Weaver, V. M., The extracellular matrix at a glance. J. Cell Sci. 2010, 123 (24), 4195-4200. (28) Hay, E. D., Cell biology of extracellular matrix. Springer Science & Business Media: 2013. (29) Eyre, D. R., Collagen: molecular diversity in the body's protein scaffold. Science 1980, 207 (4437), 1315-1322. (30) Pankov, R.; Yamada, K. M., Fibronectin at a glance. J. Cell Sci. 2002, 115 (20), 38613863. (31) Timpl, R.; Rohde, H.; Robey, P. G.; Rennard, S. I.; Foidart, J.-M.; Martin, G. R., Laminin--a glycoprotein from basement membranes. J. Biol. Chem. 1979, 254 (19), 9933-9937. (32) Theocharis, A. D.; Skandalis, S. S.; Gialeli, C.; Karamanos, N. K., Extracellular matrix structure. Adv. Drug Del. Rev. 2016, 97, 4-27. (33) Sakiyama-Elbert, S.; Hubbell, J., Functional Biomaterials: Design of Novel Biomaterials. Annual Review of Materials Research 2001, 31 (1), 183-201. (34) Crapo, P. M.; Gilbert, T. W.; Badylak, S. F., An overview of tissue and whole organ decellularization processes. Biomaterials 2011, 32 (12), 3233-3243. (35) Elder, B. D.; Eleswarapu, S. V.; Athanasiou, K. A., Extraction techniques for the decellularization of tissue engineered articular cartilage constructs. Biomaterials 2009, 30 (22), 3749-3756. (36) Sullivan, D. C.; Mirmalek-Sani, S.-H.; Deegan, D. B.; Baptista, P. M.; Aboushwareb, T.; Atala, A.; Yoo, J. J., Decellularization methods of porcine kidneys for whole organ engineering using a high-throughput system. Biomaterials 2012, 33 (31), 7756-7764. (37) Yasui, H.; Lee, J.-K.; Yoshida, A.; Yokoyama, T.; Nakanishi, H.; Miwa, K.; Naito, A. T.; Oka, T.; Akazawa, H.; Nakai, J., Excitation propagation in three-dimensional engineered hearts using decellularized extracellular matrix. Biomaterials 2014, 35 (27), 7839-7850. (38) Baiguera, S.; Del Gaudio, C.; Lucatelli, E.; Kuevda, E.; Boieri, M.; Mazzanti, B.; Bianco, A.; Macchiarini, P., Electrospun gelatin scaffolds incorporating rat decellularized brain extracellular matrix for neural tissue engineering. Biomaterials 2014, 35 (4), 1205-1214. (39) Pati, F.; Song, T.-H.; Rijal, G.; Jang, J.; Kim, S. W.; Cho, D.-W., Ornamenting 3D printed scaffolds with cell-laid extracellular matrix for bone tissue regeneration. Biomaterials 2015, 37, 230-241. (40) Volpato, F. Z.; Führmann, T.; Migliaresi, C.; Hutmacher, D. W.; Dalton, P. D., Using extracellular matrix for regenerative medicine in the spinal cord. Biomaterials 2013, 34 (21), 4945-4955. (41) Wang, L.; Johnson, J. A.; Chang, D. W.; Zhang, Q., Decellularized musculofascial extracellular matrix for tissue engineering. Biomaterials 2013, 34 (11), 2641-2654. (42) Aamodt, J. M.; Grainger, D. W., Extracellular matrix-based biomaterial scaffolds and the host response. Biomaterials 2016, 86, 68-82. (43) Medberry, C. J.; Crapo, P. M.; Siu, B. F.; Carruthers, C. A.; Wolf, M. T.; Nagarkar, S. P.; Agrawal, V.; Jones, K. E.; Kelly, J.; Johnson, S. A., Hydrogels derived from central nervous system extracellular matrix. Biomaterials 2013, 34 (4), 1033-1040. (44) Jiang, W.-C.; Cheng, Y.-H.; Yen, M.-H.; Chang, Y.; Yang, V. W.; Lee, O. K., Cryochemical decellularization of the whole liver for mesenchymal stem cells-based functional hepatic tissue engineering. Biomaterials 2014, 35 (11), 3607-3617. 28 ACS Paragon Plus Environment

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(45) Yu, C.; Kornmuller, A.; Brown, C.; Hoare, T.; Flynn, L. E., Decellularized adipose tissue microcarriers as a dynamic culture platform for human adipose-derived stem/stromal cell expansion. Biomaterials 2017, 120, 66-80. (46) Huang, C.-J.; Cho, N.-J.; Hsu, C.-J.; Tseng, P.-Y.; Frank, C. W.; Chang, Y.-C., Type I collagen-functionalized supported lipid bilayer as a cell culture platform. Biomacromolecules 2010, 11 (5), 1231-1240. (47) Tseng, P.-Y.; Chang, Y.-C., Tethered fibronectin liposomes on supported lipid bilayers as a prepackaged controlled-release platform for cell-based assays. Biomacromolecules 2012, 13 (8), 2254-2262. (48) Garcia, A. S.; Dellatore, S. M.; Messersmith, P. B.; Miller, W. M., Effects of supported lipid monolayer fluidity on the adhesion of hematopoietic progenitor cell lines to fibronectinderived peptide ligands for α5β1 and α4β1 integrins. Langmuir 2009, 25 (5), 2994-3002. (49) Thid, D.; Bally, M.; Holm, K.; Chessari, S.; Tosatti, S.; Textor, M.; Gold, J., Issues of ligand accessibility and mobility in initial cell attachment. Langmuir 2007, 23 (23), 1169311704. (50) Ananthanarayanan, B.; Little, L.; Schaffer, D. V.; Healy, K. E.; Tirrell, M., Neural stem cell adhesion and proliferation on phospholipid bilayers functionalized with RGD peptides. Biomaterials 2010, 31 (33), 8706-8715. (51) Ochsenhirt, S. E.; Kokkoli, E.; McCarthy, J. B.; Tirrell, M., Effect of RGD secondary structure and the synergy site PHSRN on cell adhesion, spreading and specific integrin engagement. Biomaterials 2006, 27 (20), 3863-3874. (52) Svedhem, S.; Dahlborg, D.; Ekeroth, J.; Kelly, J.; Höök, F.; Gold, J., In situ peptidemodified supported lipid bilayers for controlled cell attachment. Langmuir 2003, 19 (17), 67306736. (53) Stroumpoulis, D.; Zhang, H.; Rubalcava, L.; Gliem, J.; Tirrell, M., Cell adhesion and growth to peptide-patterned supported lipid membranes. Langmuir 2007, 23 (7), 3849-3856. (54) DeQuach, J. A.; Yuan, S. H.; Goldstein, L. S.; Christman, K. L., Decellularized porcine brain matrix for cell culture and tissue engineering scaffolds. Tissue Engineering Part A 2011, 17 (21-22), 2583-2592. (55) DeQuach, J. A.; Mezzano, V.; Miglani, A.; Lange, S.; Keller, G. M.; Sheikh, F.; Christman, K. L., Simple and high yielding method for preparing tissue specific extracellular matrix coatings for cell culture. PloS one 2010, 5 (9), e13039. (56) Wolf, M. T.; Daly, K. A.; Brennan-Pierce, E. P.; Johnson, S. A.; Carruthers, C. A.; D'Amore, A.; Nagarkar, S. P.; Velankar, S. S.; Badylak, S. F., A hydrogel derived from decellularized dermal extracellular matrix. Biomaterials 2012, 33 (29), 7028-7038. (57) Hong, Y.; Huber, A.; Takanari, K.; Amoroso, N. J.; Hashizume, R.; Badylak, S. F.; Wagner, W. R., Mechanical properties and in vivo behavior of a biodegradable synthetic polymer microfiber–extracellular matrix hydrogel biohybrid scaffold. Biomaterials 2011, 32 (13), 3387-3394. (58) Chadwick, R.; Conrad, M.; McGinnis, M.; Johnston-Dow, L.; Spurgeon, S.; Kronick, M., Heterozygote and mutation detection by direct automated fluorescent DNA sequencing using a mutant Taq DNA polymerase. BioTechniques 1996, 20 (4), 676-683. (59) Neuman, R. E.; Logan, M. A., The determination of hydroxyproline. J. Biol. Chem. 1950, 184 (1), 299-306. (60) Tabaei, S. R.; Choi, J.-H.; Haw Zan, G.; Zhdanov, V. P.; Cho, N.-J., Solvent-Assisted Lipid Bilayer Formation on Silicon Dioxide and Gold. Langmuir 2014, 30 (34), 10363-10373. 29 ACS Paragon Plus Environment

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(61) Jönsson, P.; Jonsson, M. P.; Tegenfeldt, J. O.; Höök, F., A method improving the accuracy of fluorescence recovery after photobleaching analysis. Biophys. J. 2008, 95 (11), 5334-5348. (62) Guneta, V.; Loh, Q. L.; Choong, C., Cell‐secreted extracellular matrix formation and differentiation of adipose‐derived stem cells in 3D alginate scaffolds with tunable properties. J. Biomed. Mater. Res., Part A 2016. (63) Tominaga, H.; Ishiyama, M.; Ohseto, F.; Sasamoto, K.; Hamamoto, T.; Suzuki, K.; Watanabe, M., A water-soluble tetrazolium salt useful for colorimetric cell viability assay. Anal. Commun. 1999, 36 (2), 47-50. (64) Dunne, L. W.; Huang, Z.; Meng, W.; Fan, X.; Zhang, N.; Zhang, Q.; An, Z., Human decellularized adipose tissue scaffold as a model for breast cancer cell growth and drug treatments. Biomaterials 2014, 35 (18), 4940-4949. (65) Pati, F.; Ha, D.-H.; Jang, J.; Han, H. H.; Rhie, J.-W.; Cho, D.-W., Biomimetic 3D tissue printing for soft tissue regeneration. Biomaterials 2015, 62, 164-175. (66) Pati, F.; Jang, J.; Ha, D.-H.; Kim, S. W.; Rhie, J.-W.; Shim, J.-H.; Kim, D.-H.; Cho, D.W., Printing three-dimensional tissue analogues with decellularized extracellular matrix bioink. Nat. Commun. 2014, 5. (67) Luo, B.; Yuan, S.; Foo, S. E. M.; Wong, M. T. C.; Lim, T. C.; Tan, N. S.; Choong, C., From flab to fab: transforming surgical waste into an effective bioactive coating material. Adv. Healthcare Mater. 2015, 4 (4), 613-620. (68) Mariman, E. C.; Wang, P., Adipocyte extracellular matrix composition, dynamics and role in obesity. Cell. Mol. Life Sci. 2010, 67 (8), 1277-1292. (69) Singelyn, J. M.; DeQuach, J. A.; Seif-Naraghi, S. B.; Littlefield, R. B.; Schup-Magoffin, P. J.; Christman, K. L., Naturally derived myocardial matrix as an injectable scaffold for cardiac tissue engineering. Biomaterials 2009, 30 (29), 5409-5416. (70) Marqués-Gallego, P.; de Kroon, A. I., Ligation strategies for targeting liposomal nanocarriers. Biomed Res Int 2014, 2014. (71) Tabaei, S. R.; Jackman, J. A.; Kim, S.-O.; Zhdanov, V. P.; Cho, N.-J., Solvent-Assisted Lipid Self-Assembly at Hydrophilic Surfaces: Factors Influencing the Formation of Supported Membranes. Langmuir 2015, 31 (10), 3125-3134. (72) Tabaei, S. R.; Ng, W. B.; Cho, S.-J.; Cho, N.-J., Controlling the Formation of Phospholipid Monolayer, Bilayer, and Intact Vesicle Layer on Graphene. ACS Appl. Mater. Interfaces 2016, 8 (18), 11875-11880. (73) Nielsen, S. B.; Otzen, D. E., Quartz Crystal Microbalances as Tools for Probing Protein– Membrane Interactions. Lipid-Protein Interactions: Methods and Protocols 2013, 1-21. (74) Xi, J.; Chen, J.; Garcia, M.; Penn, L., Quartz crystal microbalance in cell biology studies. J Biochip Tissue chip S 2013, 5, 2153-0777. (75) Keller, C.; Kasemo, B., Surface specific kinetics of lipid vesicle adsorption measured with a quartz crystal microbalance. Biophys. J. 1998, 75 (3), 1397-1402. (76) Modin, C.; Stranne, A.-L.; Foss, M.; Duch, M.; Justesen, J.; Chevallier, J.; Andersen, L. K.; Hemmersam, A. G.; Pedersen, F. S.; Besenbacher, F., QCM-D studies of attachment and differential spreading of pre-osteoblastic cells on Ta and Cr surfaces. Biomaterials 2006, 27 (8), 1346-1354. (77) Rodahl, M.; Höök, F.; Fredriksson, C.; Keller, C. A.; Krozer, A.; Brzezinski, P.; Voinova, M.; Kasemo, B., Simultaneous frequency and dissipation factor QCM measurements of biomolecular adsorption and cell adhesion. Faraday Discuss. 1997, 107, 229-246. 30 ACS Paragon Plus Environment

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(78) Höök, F.; Rodahl, M.; Brzezinski, P.; Kasemo, B., Energy dissipation kinetics for protein and antibody-antigen adsorption under shear oscillation on a quartz crystal microbalance. Langmuir 1998, 14 (4), 729-734. (79) Sauerbrey, G., Verwendung von schwingquarzen zur wagung dunner schichten und zur mikrowagung. Z. Phys. 1959, 155 (2), 206-222. (80) Voinova, M. V.; Rodahl, M.; Jonson, M.; Kasemo, B., Viscoelastic acoustic response of layered polymer films at fluid-solid interfaces: continuum mechanics approach. Phys. Scr 1999, 59 (5), 391. (81) Patel, A. R.; Kanazawa, K. K.; Frank, C. W., Antibody binding to a tethered vesicle assembly using QCM-D. Anal. Chem. 2009, 81 (15), 6021-6029. (82) Tamm, L. K., Lateral diffusion and fluorescence microscope studies on a monoclonal antibody specifically bound to supported phospholipid bilayers. Biochemistry 1988, 27 (5), 14501457. (83) Nakabayashi, H.; Taketa, K.; Miyano, K.; Yamane, T.; Sato, J., Growth of human hepatoma cell lines with differentiated functions in chemically defined medium. Cancer Res. 1982, 42 (9), 3858-3863. (84) Stupack, D. G.; Cheresh, D. A., Get a ligand, get a life: integrins, signaling and cell survival. J. Cell Sci. 2002, 115 (19), 3729-3738. (85) Schwartz, M. A.; Assoian, R. K., Integrins and cell proliferation. J. Cell Sci. 2001, 114 (14), 2553-2560. (86) Frith, J. E.; Mills, R. J.; Hudson, J. E.; Cooper-White, J. J., Tailored integrin– extracellular matrix interactions to direct human mesenchymal stem cell differentiation. Stem Cells Dev. 2012, 21 (13), 2442-2456. (87) Hood, J. D.; Cheresh, D. A., Role of integrins in cell invasion and migration. Nat. Rev. Cancer 2002, 2 (2), 91-100. (88) Vafaei, S.; Tabaei, S. R.; Biswas, K. H.; Groves, J. T.; Cho, N. J., Dynamic Cellular Interactions with Extracellular Matrix Triggered by Biomechanical Tuning of Low‐Rigidity, Supported Lipid Membranes. Adv. Healthcare Mater. 2017. (89) Vafaei, S.; Tabaei, S. R.; Cho, N.-J., Optimizing the Performance of Supported Lipid Bilayers as Cell Culture Platforms Based on Extracellular Matrix Functionalization. ACS Omega 2017, 2 (6), 2395-2404. (90) Gilmanshin, R.; Creutz, C. E.; Tamm, L. K., Annexin IV reduces the rate of lateral lipid diffusion and changes the fluid phase structure of the lipid bilayer when it binds to negatively charged membranes in the presence of calcium. Biochemistry 1994, 33 (27), 8225-8232. (91) Yamazaki, V.; Sirenko, O.; Schafer, R. J.; Groves, J. T., Lipid mobility and molecular binding in fluid lipid membranes. J. Am. Chem. Soc. 2005, 127 (9), 2826-2827. (92) Juliano, R. L.; Haskill, S., Signal transduction from the extracellular matrix. J. Cell Biol. 1993, 120, 577-577. (93) Nagaki, M.; Shidoji, Y.; Yamada, Y.; Sugiyama, A.; Tanaka, M.; Akaike, T.; Ohnishi, H.; Moriwaki, H.; Muto, Y., Regulation of hepatic genes and liver transcription factors in rat hepatocytes by extracellular matrix. Biochem. Biophys. Res. Commun. 1995, 210 (1), 38-43.


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