3D Printing of PDMS Improves Its Mechanical and Cell Adhesion


Dec 21, 2017 - The results demonstrate that 3D printing of PDMS improved the mechanical properties of fabricated samples up to three fold compared to ...
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3D Printing of PDMS Improves its Mechanical and Cell Adhesion Properties Veli Ozbolat, Madhuri Dey, Bugra Ayan, Adomas Povilianskas, Melik C Demirel, and Ibrahim Ozbolat ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.7b00646 • Publication Date (Web): 21 Dec 2017 Downloaded from http://pubs.acs.org on December 22, 2017

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3D Printing of PDMS Improves its Mechanical and Cell Adhesion Properties Veli Ozbolat1,2,3, Madhuri Dey2,4, Bugra Ayan1,2, Adomas Povilianskas1, Melik C. Demirel1,2,6 and Ibrahim T. Ozbolat1,2,5,6,* 1

Engineering Science and Mechanics Department, Penn State University, University Park, PA

16802, USA, 2

The Huck Institutes of the Life Sciences, Penn State University, University Park, PA 16802,

USA, 3

Mechanical Engineering Department, Ceyhan Engineering Faculty, Cukurova University,

Adana 01950, Turkey, 4

Chemistry Department, Penn State University, University Park, PA 16802, USA,

5

Biomedical Engineering Department, Penn State University, University Park, PA 16802, USA,

6

Materials Research Institute, Penn State University, University Park, PA 16802, USA,

* Corresponding author, Email: [email protected]

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ABSTRACT Despite extensive use of polydimethylsiloxane (PDMS) in medical applications, such as lab-ona-chip or tissue/organ-on-a-chip devices, point-of-care devices and biological machines, the manufacturing of PDMS devices is limited to soft-lithography and its derivatives, which prohibits the fabrication of geometrically complex shapes. With the recent advances in threedimensional (3D) printing, use of PDMS for fabrication of such complex shapes has gained considerable interest. This research presents a detailed investigation on printability of PDMS elastomers over three concentrations for mechanical and cell adhesion studies. The results demonstrate that 3D printing of PDMS improved the mechanical properties of fabricated samples up to three folds compared to that of cast ones due to decreased porosity of bubble entrapment. Most importantly, 3D printing facilitates the adhesion of breast cancer cells, whereas cast samples do not allow cellular adhesion without the use of additional coatings such as extracellular matrix proteins. Cells are able to adhere and grow in the grooves along the printed filaments demonstrating that 3D printed devices can be engineered with superior cell adhesion qualities compared to traditionally manufactured PDMS devices. Keywords: 3D printing, PDMS, cell adhesion, soft lithography 1. Introduction Polydimethylsiloxane (PDMS) has been extensively used in medical applications including lab-on-a-chip machines

7,8

1,2

or tissue/organ-on-a-chip devices

3,4

, point-of-care devices

5,6

, biological

and two-dimensional (2D) or three-dimensional (3D) cell culture. It is

biocompatible, transparent, gas permeable and economical 9–11. PDMS is widely used in medical research and technology, and there are a wide array of manufacturing techniques used for forming PDMS including soft-lithography and its derivatives, molding, dip casting, spin coating

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and many others

12–15

. Most of these techniques yield geometrically simple structures with

limited complexities in 3D. In addition, degassing is generally required as bubble formation is observed. Such bubble formation may generate some shortcomings such as surface roughness and imperfections, and inferior mechanical properties. As PDMS is hydrophobic, cells in general poorly attach on PDMS without any surface modifications such as treating with charged molecules such as extracellular matrix proteins (i.e., collagen, fibronectin) 16–18 With the recent advances in 3D printing and innovative development of ink materials, fabrication of highly complex shapes made of plastics, metals and even ceramics is feasible 19,20. Femmer et al. demonstrated the 3D printing of PDMS using the digital light processing (DLP) approach for the first time in the literature, where 100 µm resolution was attained 21. Later, the same group demonstrated the fabrication of complex architectures (i.e., triple periodic minimal surfaces with substantial overhangs) by DLP

22

, where a sacrificial mold was first 3D printed

followed by casting PDMS and removing the sacrificial mold thereafter. Synthetic spider webs 23 and fluidic chambers 24,25 have been recently 3D printed with translucent and non-flowing PDMS elastomers. Also, PDMS were printed within an oil-based granular gel 26 with high resolution (80 µm) and mechanical properties (700% strain) and in a hydrophilic support bath

27

to enable true

freeform fabrication of complex structures. In this paper, we performed a comprehensive analysis of 3D printability of PDMS ink, which are blends of two PDMS elastomers, SE 1700 25,28

and widely used Sylgard 184

29,30

. We demonstrated that the 3D printed samples possess

improved mechanical and cell adhesion properties compared to traditionally manufactured samples using casting process. Various complex models of organs were also 3D printed to demonstrate the printability of highly intricate geometries.

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2. Experimental section 2.1. Ink preparation Viscoelastic polydimethylsiloxane (PDMS) inks were prepared by blending two silicone elastomers: a shear-thinning PDMS material SE 1700 (Dow Corning, Auburn, MI, USA) and a low-viscosity PDMS material Sylgard 184 (Dow Corning), which is used to dilute SE 1700 for desired rheological properties. Both SE 1700 and Sylgard 184 base materials were first mixed with their curing agents in a 10:1 (base:curing agent) ratio by weight before blending. Prior to mixing SE 1700 and Sylgard 184, both solutions were placed in a vacuum desiccator for degassing for 15 minutes just after mixing the base and agent. Final inks were loaded into a 3 cc syringe barrels syringe (Nordson EFD, USA) at room temperature and centrifuged in Sorvall Legend X1R Centrifuge (Thermo Fisher Scientific, Waltham, MA, USA) at 5,000 rpm for 5 minutes to remove any air bubbles. Finally, PDMS inks were prepared by mixing SE 1700 and Sylgard 184 in five different ratios (10:0, 9:1, 8:2, 7:3 and 0:10). 2.2. Rheological measurement Rheological measurements were performed using a MCR 302 rheometer (Anton Paar, Ashland, VA, USA) with a 25-mm diameter parallel-plate geometry measuring system. The storage modulus (G') and loss modulus (G'') were recorded from amplitude sweep of Inks 10:0, 9:1, 8:2, 7:3 and 0:10 at a constant frequency of 1 Hz at a strain range from 0.01% to 100%, where Inks 10:0 and 0:10 were used as control groups. In addition, the frequency sweep was carried out to get G', G'' and complex viscosity (∣η*∣) at a constant strain of 0.01%, which was within the linear viscoelastic range at a frequency range from 100 Hz to 0.1 Hz. The flow and viscosity curves were obtained from the rotational test, which was carried out at shear rates ranging from 0.01 to 1000 s-1. The duration of a data point was decreased logarithmically from

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30 to 2 s. Lastly, structural breakdown and recovery (3ITT) test was conducted. In this test, a constant frequency of 1 Hz and a constant strain of 0.05% (within the linear viscoelastic range) were chosen in an interval of 1 to 3. In interval 2, the sample was sheared for 2 s at a shear rate of 100 s-1. Constant temperature of 25 ˚C was chosen for all experiments. 2.3. Printability analysis 3D printing was performed using an INKREDIBLE 3D bioprinter (Cellink, Sweden). PDMS inks were loaded into syringe barrels at room temperature, which were then capped with 20 GA (610 µm), 21 GA (510 µm) and 22 GA (410 µm) deposition tips (Nordson EFD). The extrusion pressure was controlled with the pressure control unit equipped with a built-in air pump. The printing speed was controlled with a (G-code) program that was sent to the printer using the Repetier software. Widths of printed filaments were measured and analyzed for three inks (Inks 9:1, 8:2 and 7:3), three different sizes of nozzle tips (410, 510 and 610 µm), four different extrusion pressure levels (150, 200, 250 and 300 kPa) and different printing speeds (30-300 mm/min). The gap between the nozzle tip and printing platform was fixed to 0.6 mm in all experiments. The widths of printed filaments were measured using EVOS FL Auto microscope (Thermo Fisher Scientific, Waltham, MA, USA) at 4X magnification. Printability experiments were performed for Inks 9:1, 8:2 and 7:3 with different tip diameters (410, 510 and 610 µm) at a constant extrusion pressure (250 kPa) and printing speed (180 mm/min). Dual-layer constructs were printed with the same distance between adjacent filaments both in x- and y-directions. This distance was kept larger for 610 and 510 µm nozzle tips for Ink 7:3 due to its large extrusion volume. We first define the circularity (C) in order to define printability (Pr):

C=

4πA L2

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(1)

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where, L and A are the perimeter and area of the pores, respectively. The shape of the enclosed area is closest to a circle when an C is equal to 1. Since highest circularity for a square shape is equal to π/4, the Pr of ink based on a printed square shape is defined as follows 31:

Pr =

π 1 ⋅

4 C

=

L2 16 A

(2)

For an ideal mixing ratio of an ink, the pores exhibit a square shape, which has a Pr value of 1. The optical images of 3D printed constructs were taken, then the area and perimeter of the pores were measured using the EVOS FL Auto microscope at 4X magnification. 2.4. 3D Printing of models The mesh files of human hand, ear and femur were downloaded from National Institutes of Health (NIH) 3D Print Exchange (https://3dprint.nih.gov/) and the print paths were generated after scaling down mesh models using a slicer package Slic3r (http://slic3r.org/). The print paths for the human nose and blood vessel were taken from Cellink (Sweden), and the print path for the bifurcated vessel was generated in-house. 3D complex models were printed using Ink 8:2 with a 510 µm nozzle tip at a constant printing speed and extrusion pressure of 160 mm/min and 300 kPa, respectively. 2.5. Shape fidelity analysis Shape fidelity analysis of the printed nose model was performed after curing in an oven at 80 ˚C. The nose model was chosen due to its complex shape which includes curved contours, cavities and overhangs. Three-dimensional structural magnetic resonance imaging (MRI) was conducted on a 14.1 Tesla micro-imaging system (Agilent, Santa Clara, CA, USA). The specimen was placed inside a 30 mL syringe and inserted into the magnet. To avoid imaging artifacts, a standard 3D spin echo sequence with an echo time of 5.8 ms and a 150 µm isotropic

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resolution were used to image the sample. The data was zero-filled by a factor of two in each direction before Fourier transformation resulting in a 75 µm isotropic pixel resolution (Matlab, The MathWorks, Inc., Natick, MA, USA). Next, image segmentation was conducted using Avizo software (Thermo Fisher Scientific, Hillsboro, OR, USA). The MRI data were then reconstructed into a 3D mesh file using Avizo, which was compared with the original design model. Using Geomagic Control X (3D Systems Inc, Rock Hill, SC, USA), 3D heat map was obtained in order to show the surface deviation. The deviation threshold of 100 µm was chosen as it represented 10-20% of the average filament diameter. 2.6. Mechanical testing Specimens were prepared in accordance with ASTM D412 Type C standard. The specimens were modeled and scaled down to 1:2.5 ratio using computer-aided design (CAD) software (Autodesk Inventor 2017, USA). The print paths (longitudinal and transverse filament directions) were generated using Slic3r and specimens were prepared for Inks 9:1, 8:2 and 7:3. Dogbone specimens were printed in both longitudinal and transverse filament directions as well as they were cast into a 3D printed mold for mechanical property evaluation. The samples were cured at 80 °C in an Isotemp 282A vacuum oven (Thermo Fisher Scientific, Waltham, MA, USA) for two hours. An Instron 5966 tensile testing device (Instron, Norwood, MA, USA) was used to perform the uniaxial tensile testing. Test specimens were fixed on the gripers at both ends and rigidly held by a 1 kN load-cell platform. Test specimens were manually loaded until positive tension was reached. Initial thickness, width and length of the reduced section were measured with a digital caliper before loading. Tensile loads were applied at a loading rate of 500 mm/min until a failure was observed. Tensile stress, strain, peak stress and strain at break were recorded. Young’s modulus was calculated based on the slope of the tensile stress/strain curve.

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2.7. Porosity analysis The porosity analysis was performed according to ISO 15901-1using a MicroActive AutoPore V 9600 mercury porosimeter (Micromeritics Instrument, Norcross, GA, USA). Samples were first chopped into 3-5 mm thick pieces and then loaded into the porosimeter (1.5 to 2 g in weight), where they were subjected to a pressure cycle starting at approximately 0.1 psi, increasing to 61,000 psi in predefined steps to determine the porosity and average pore diameter of the samples. The contact angle and temperature of mercury was set at 130° and ∼19 °C, respectively. 2.8. Optical profilometer images 3D printed PDMS surfaces were measured using a Nexview™ optical surface profiler (Zaygo, Middlefield, CT, USA) with a 50X lens. In order to image adjacent filaments, a composite of seven stitch fields were collected and post-processing was applied. 2.9. Cell culture and Reagents Green fluorescent protein (GFP)-transfected metastatic breast cancer (MDA-MB-231) cells were grown in RPMI media (RPMI 1640, Corning, Manassas, VA, USA) supplemented with 10% fetal bovine serum (FBS) (Corning, Manassas, VA, USA), 2mM Glutamine (GlutaMAX 100X, Gibco, Life Technologies, Grand Island, NY, USA) and 1% Penicillin-Streptomycin (Gibco). Cells were cultured in tissue culture flasks and maintained at 37 ˚C and 5% CO2 flow in an incubator. Confluent flasks were washed with phosphate-buffered saline (PBS) and trypsinized (0.25%Trypsin, 2.21mM EDTA, 1X, Corning, Manassas, VA, USA) to harvest the required number of cells. 2.10.

Cell adhesion study

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3D printed and cast surfaces with Inks 9:1, 8:2 and 7:3 were freshly prepared for cell adhesion study. The distances between filaments were varied while keeping the filament width at 760 µm. The distance between the centers of adjacent filaments were determined to be 760, 740, 720 and 700 µm for Sample 1, 2, 3 and 4, respectively. Half of the samples were coated with 10 µg/ml fibronectin from bovine plasma (Sigma Aldrich, St. Louis, MO, USA). MDA-MB-231 cells were seeded at a population of 10,000 cells per 200 µL on both fibronectin-free and -coated surfaces for a comparative study. Samples were all cultured in RPMI culture media, supplemented with 10% FBS and 2 mM glutamine. After three days of cell culture, the samples were carefully rinsed with PBS and fixed with 4% formaldehyde solution overnight for further analysis. 2.11.

Imaging and quantification of cell attachment

The nuclei of the fixed cells adhered on the PDMS surface were fluorescently stained with Hoechst nucleic acid stain (Life Technologies), with a 1:200 dilution in PBS. About 200 µl of this diluted Hoechst solution was added to each PDMS sample and the cells were incubated in a dark room for 10 minutes. Following this, the samples were carefully rinsed with PBS and then imaged on the EVOS FL Auto microscope. Since the printed PDMS surface had an uneven topography, Z-stacks were obtained from 15 different regions at a 20X magnification for each sample to quantify the average number of nuclei per millimeter square for a particular sample. The images were further processed using ImageJ software (NIH, USA). The images were then carefully thresholded to highlight the region of interest and then analyzed for the number of nuclei present per millimeter square. Further, using the same magnification, images were also taken on the green fluorescent channel to observe the overall cell morphology. Cell area was obtained by outlining each cell along its boundary and then generating the area it occupies by

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using the manufacturer’s software. The average cell area was calculated and cells which spread out more occupied a higher area as compared to those were rounded.

2.12.

Visualization of cell cytoskeleton

The actin cytoskeleton of adhered MDA-MB-231 cells were stained with phalloidin Alexa Fluor 568 (Life Technologies,Camarillo, CA, USA). Samples were fixed in 4% formaldehyde solution overnight. Then, they were washed twice with PBS, permeabilized with 0.1% TritonX100 (Sigma Aldrich) for 15 minutes. After carefully rinsing the samples twice with PBS, phalloidin was diluted 1:100 in PBS and added to the samples in order to stain the actin cytoskeleton of adhered cells. The samples were incubated at room temperature for 30 minutes. The cell cytoskeleton was then viewed under the EVOS FL Auto microscope. 2.13.

Fourier transform infrared spectroscopy

Infrared spectra of cured SE 1700 and Sylgard 184 PDMS and their printed and non-printed inks (Inks 9:1, 8:2 and 7:3) were recorded in a Nicolet 6700 FTIR spectrometer (Thermo Fisher Scientific, Waltham, MA, USA) using an attenuated total reflection mode. The spectra were recorded with 4 cm-1 resolution from 400 to 4000 cm-1. For each spectrum 256 scans were performed. 2.14.

Statistics

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All data are presented as the mean ± standard deviation (SD) and were analyzed by Minitab 17.3 (Minitab Inc., State College, PA, USA) using one-way analysis of variance (ANOVA) to test for significance when comparing the data. Post-hoc Tukey’s multiple comparison test was used to determine the individual differences among the groups. Statistical differences were considered at p