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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<0.05 (*), p<0.01 (**), p<0.001 (***).

3. Results and discussion 3.1. Rheological evaluation of PDMS inks One of the aims in this study is to determine whether 3D extrusion based printing 32 could be used to print PDMS inks. In order to understand the printability of PDMS, we conducted a rheological study. The results from amplitude sweeps for five PDMS groups with varying mixing ratios of SE 1700 and Sylgard 184 (10:0, 9:1, 8:2, 7:3 and 0:10, henceforth referred as Inks 10:0, 9:1, 8:2, 7:3 and 0:10 respectively) were presented in Figure 1A. Inks 10:0, 9:1, 8:2 and 7:3 showed dominant storage modulus (G'), indicating these inks had a more gel-like character. The value of the linear viscoelastic (LVE) range determines the limit at which the samples’ structures are permanently deformed and the sample stability are in terms of preventing sedimentation within the sample. A LVE range of Inks 10:0 and 9:1 was observed between 0.01-0.035% strains while Inks 8:2 and 7:2 had LVE ranges between 0.1-0.18% and 0.1-0.21% strains, respectively. The structures of these groups began to permanently deform once the LVE range exceeded the yield point (Figure 1A). After the yield point, the material flowed like a liquid, seen as the flow point where G' and loss modulus (G'') cross-over. Since the structural strength of the inks is

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represented by the value of each G', Ink 10:0 had the greatest structural strength and Ink 0:10 had the lowest. Ink 0:10, which contained only Sylgard 184, behaved like a liquid at all shear rates since it had a dominating G''. Hence, this type of inks had not LVE range or yield point. As a result, the attempt to use Ink 0:10 for extrusion-based printing was unsuccessful as the shape of the printed ink was not maintained. The results of frequency sweep tests for Inks 9:1, 8:2 and 7:3 show no long-term relaxation. Due to the dominating G' values over G'' at all frequencies (Figure 1B). However, G' of Inks 9:1 and 8:2 were an order of magnitude higher than that of Ink 7:3, which could indicate their viscoelastic behavior making them more suitable for extrusion-based printing. Additionally, the complex viscosity of the inks increased with increased concentration of SE 1700 in the blend. The results from rotational tests are shown in Figures 1C-D. Inks 10:0, 9:1, 8:2 and 7:3 showed shear thinning behavior of viscoelastic solids. The viscosity of the inks decreased as shear rate increased. Viscosity (η) of the inks decreased with increased concentration of Sylgard 184 in the blend. On the other hand, Ink 0:10 expressed zero shear viscosity (see the plateau region of the curve in Figure 1C). The viscosity was constant at lower frequencies, so the ink behaved and flowed like a Newtonian fluid. After the zero shear plateaus was exceeded at 120 s1

, the ink showed shear thinning behavior illustrated by the power law region of the curve. In our

study, Inks 9:1, 8:2 and 7:3 behaved as a Herschel-Bulkley fluid as their strain rate and stress had a non-linear relationship. The yield stress is what must be taken into account during the extrusion process for the inks. A larger yield stress, as with Ink 9:1 compared to Ink 7:3, indicated a more stable structure, higher layer thickness, and more gel-like character. Prior to the yield stress, all inks exhibited elastic behavior.

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From the results of 3-interval-thixotropy-tests (see Figure 1E), the recovery processes for Inks 9:1, 8:2 and 7:3 were observed. Interval 1 details the at-rest state of the samples. Interval 2 details structure decomposition under high shear (as experienced during extrusion) and Interval 3 details structure regeneration of the ink which is observed post-extrusion, once the ink is laid. Initially all inks expressed a large G' indicating that they were gel-like in structure. In interval 2, Ink 9:1 remained gel-like as G' never crosses G''. As G' and G'' become closer with high shear and eventually cross-over, Inks 8:2 and 7:3 were progressively less elastic until they flowed like liquid, which is an important behavior during the extrusion process. All inks showed recovery to a dominating elastic state in Interval 3. Since Ink 0:10 is not appropriate for extrusion-based printing and it needs to be diluted for the given printing parameters, it was not used for the rest of the study. 3.2. 3D Printability of PDMS ink We used an INKREDIBLE bioprinter (Figure 2A) to print PDMS inks. Printability of inks was evaluated based on the integrity of dual-layer constructs (Figure 2B), where the printability (Pr) is calculated according to Equation 1, 2. Figure 2C demonstrates the general printing scheme to analyze the printability of the ink, where the pores enclosed by the adjacent filaments of consecutive layers exhibit deviations from a perfect square when the printability (Pr) is equal to 1. Optical images of dual-layer constructs printed with various mixing ratios of PDMS at 250 kPa and 180 mm/min are shown in Figure 2D.

When the ink had a higher Sylgard 184

concentration making the ink less viscous, the ink after printing spread at the cross site so it was challenging to print structurally-stable constructs. Also, due to the fusion of the adjacent filaments, the pores developed fillets. Converging to ideal mixing concentration at given printing parameters, continuous uniform filaments were extruded so the shape of the pores closed to

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square with sharp edges. An increase in SE 1700 concentration in the ink beyond the ideal concentration made the filament extrusion more difficult; filaments and pores exhibited irregular shapes and the Pr value diverged from 1. Figure 3 shows the width of printed filaments for three inks at various nozzle tip sizes, printing speeds and extrusion pressures. Ink 9:1 could not been printed in a continuous manner at higher printing speeds using a 510 µm tip. Continuity was only achieved at 30 mm/min (using a 410 µm tip) at a constant extrusion pressure of 250 kPa. Since the gap between the nozzle tip and printing stage was fixed for all experiments, the filament width became much larger at lower printing speeds and higher extrusion pressures. The filament width increased with the increased concentration of Sylgard 184. The filament widths were larger than tip diameters for a given speed interval for Ink 7:3 since 250 kPa extrusion pressure was too high for that particular ink. The filament widths using three different nozzle tips (at 250 kPa extrusion pressure) were demonstrated in Figure 3B. A continuous filament was achieved for Ink 9:1 at all speeds (using a 610 µm tip); however, a decrease in the tip size limited the printing speed necessary to achieve continuity in printing. For Inks 8:2 and 7:3, continuity could be achieved at 250 kPa for a given printing speed interval but the width of Ink 7:3 filaments exceeded the tip diameter when a 610 µm tip was used. The filament widths for various extrusion pressures (using a 510 µm tip) are presented in Figure 3C. Continuous filaments could not be achieved at higher printing speeds for Ink 9:1. In order to achieve filament continuity in higher speeds, elevated extrusion pressures were required for dispensing. As Sylgard 184 concentration in the ink increased, the width of the filaments increased; thinner filaments could be printed at higher printing speeds or lower extrusion pressures due to the lower viscosity of the ink.

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After the optimization of printing parameters, various models were 3D printed in high fidelity as shown in Figure 4A, including structural models of human hand, nose, blood vessel, ear, a branched structure and femoral heads). Characterization of the shape fidelity of the printed nose model was performed through surface deviation analysis by comparing the difference between the scanned and original models (Figure 4B). 78.45% of the surface was within ±100 µm tolerance threshold with 1.86% underprint and 19.68% overprint error. The average surface deviation of 189 µm was observed with 263 µm overprint deviation along curved surface of supra alar crease and 178 µm along the columella. Furthermore, 787 µm deviation was observed along the upper portion of the nostrils where the unsupported bridging occurred. Sharp edges of the model along the nasion resulted in underprint error of -181 µm. These results display the ability to reproduce geometrically-complex structures with relatively less accuracy along sharp edges and overhanging features. 3.3. 3D Printing enhances the mechanical properties of PDMS inks A uniaxial tensile test was performed to assess the mechanical properties of printed inks. Since printing direction affects the mechanical properties in additive manufacturing 33–35, we also evaluated the mechanical properties of samples printed with longitudinal and transverse filament directions. As shown in Figure 5, PDMS samples showed significantly increased ultimate strength over cast samples, where the ultimate strength of samples made of Ink 7:3 increased from 3.432±1.216 MPa in cast samples to 3.772±0.685 MPa in printed samples with transverse filament direction to 5.036±0.657 MPa in printed samples with longitudinal filament direction. In addition, the ultimate strength increased from 3.280±1.900 and 3.736±1.457 MPa in cast samples to 10.906±4.276 and 9.338±1.264 MPa in printed samples with longitudinal filament

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direction for Inks 8:2 and 9:1, respectively. The Young’s modulus increased from 1.672±0.515 MPa in cast samples to 2.990±0.414 MPa in printed samples with transverse filament direction to 3,156±0.248 MPa in printed sample with longitudinal filament direction for Ink 8:2 and increased from 1.902±0.341 MPa in cast samples to 2.540±0.469 MPa in printed samples with transverse filament direction to 2.823±0.280 MPa in printed samples with longitudinal filament direction for Ink 9:1. On the other hand, the Young’s modulus did not show significant differences between cast and printed samples for Ink 7:3. Mechanical test results showed that the failure strains of printed samples were higher than those of cast samples for all the inks (see Supplementary Video 1). Also, significant difference between samples that were printed with transverse and longitudinal filament directions were seen only for Ink 7:3. This can be attributed to the fact that strong adhesion was observed between filaments and curing of filaments took place long after printing. Porosity analysis (see Table 1) also confirms the mechanical test results. Control groups (cast samples) had approximately 7%, 29% and 12% more porosity, with higher average pore size, compared to printed samples of Inks 7:3, 8:2 and 9:1, respectively. Less porosity and pore size in printed samples could be attributed to the less air entrapment during the shear thinning process while the ink was extruded

36

yielding improved mechanical

properties. 3.4. 3D Printing improves cell adhesion Green fluorescent protein (GFP)-transfected metastatic breast cancer cells (MDA-MB-231) were cultured on printed surfaces of Inks 7:3, 8:2 and 9:1, and compared to that of non-printed (control) surfaces in order to study the influence of 3D printing on cell adhesion. All constructs were printed with the same filament widths but with varying distances (from 700 to 760 µm) between filament centers, as shown in Figures 6A-B. Approximately 10,000 cells suspended in

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200 µl of cell culture media were seeded on the surface of printed samples and these samples were then monitored for over a period of three days. PDMS intrinsically presents a hydrophobic surface, primarily due to the presence of methyl groups, which are nonpolar 37. Thus, cell attachment on PDMS is generally impaired. However, in this study printed PDMS substrates showed significant cell adhesion as compared to nonprinted ones, which were quantified by the number of nuclei attached per mm2, as shown in Figure. 6C. For the printed samples cells exhibited preferential attachment in the grooves or the rough areas over 24 hours of culture. Gradually, they were seen spreading along the sides towards the broader filaments over a period of 3 days (Figure 6E). Cell adhesion increased by ~90% for printed samples as compared to non-printed ones for all the types. Furthermore, it was observed that Ink 8:2 facilitated the best surface for cell adhesion. As compared to Inks 7:2 and 9:1, there were about 22% and 42%, respectively, more nuclei per mm2 observed on Ink 8:2. For the non-printed samples, cells adhered on the surface and formed aggregates and did not spread (Figure 6E). Thus, low cell adhesion was observed for non-printed samples as compared to printed ones for all inks. The distribution of individual cell area depicting cell morphology is shown in the form of box and whisker plots which depicts the range of values obtained for all types of inks. As shown in Figures 6C-D, the average cell area for all non-printed samples lie below the average of all printed samples. This denotes that cells had a higher circularity or a more rounded morphology when cultured on cast inks. Furthermore, surface treatments of PDMS with ECM proteins (e.g. fibronectin) containing Arg-Gly-Asp (RGD) motifs is known to render hydrophobic surfaces suitable for cell adhesion

38

. To compare cell adhesion, one half of the

boxes were coated with 10 µg/ml fibronectin prior to cell culture. As expected, PDMS surfaces treated with fibronectin showed higher cell adhesion as compared to non-fibronectin coated

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samples (Figure 6D). Cell adhesion increased by approximately 44% for Ink 7:3, 60% for Ink 8:2 and 75% for Ink 9:1 when the surface was coated with fibronectin. Additionally, cells cultured on flat or non-printed PDMS formed disorganized clusters on the surface and exhibited significantly lower cell adhesion for non-fibronectin coated samples as compared to coated ones. This study shows that the printed samples offered a more hospitable environment for cell attachment. Interestingly, surfaces which were not treated with surface proteins also exhibited cell adhesion but to a lesser extent. In order to understand the major cause of cell adhesion on the printed inks, we first checked if the surface chemistry was influential. In that sense, Fourier transform infrared (FTIR) spectroscopy was performed in reflective (ATR) mode to detect the surface chemistry was changed due to printing. As shown in Figure 7, no differences were observed for all ink types. Printing generated an uneven surface which aided in cell adhesion. Variation in the surface roughness adds a 3D component to the cell environment compared to flat surfaces and this, in turn, could affect cell adhesion by upregulation of various ECM adhesion-related genes39–43. It has been previously studied that cells form stress fibers by orienting their actin filaments in a convenient manner such that all internal and external forces are perfectly balanced when cells try to adhere to a substrate

44,45

. These stress fibers are connected by focal adhesion complexes and

communicate with the external environment via traction forces exerted on the surface and thus the topography of the substrate guides cell adhesion and alignment

45,46

. Incubating the PDMS

substrate with complete cell media, supplemented with FBS before and after cell seeding results in adsorption of various ECM proteins like fibronectin, vitronectin from FBS onto the PDMS surface

47–51

. This competitive adsorption of proteins from the serum demonstrates the Vroman

effect, which directly enhances cellular adhesion

52,53

as proteins being adsorbed on the surface

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contain the RGD sequence recognized by cell surface integrins 52,54. Some studies also indicated that exposing untreated PDMS to just cell culture media containing amino acids (such as arginine, glutamine, histidine, and threonine) increased the surface concentration of nitrogen and oxygen which might create a hospitable environment for cells to attach

29,55

. Moreover, for

prolonged culture time cells are known to secrete various ECM proteins such as collagen, laminin which further enable them to anchor to the matrix 48,50. 4. Conclusions This study represents a comprehensive analysis of 3D printability of blends of PDMS inks. The extrudability at sufficient resolution enabled the fabrication of highly intricate geometries. Most importantly our study reveals that the shear thinning behavior of PDMS inks yielded higher mechanical properties in 3D printing compared to cast counterparts, which could be due to decreased porosity and bubble entrapment. In addition, we showed for the first time that 3D printed surface of PDMS inks promoted better cell adhesion and spreading compared to the cast surface. With such feature, 3D printing of PDMS not only enables the generation of 3D models of tissues and organs, but also brings a new concept in surface engineering for cell adhesion studies. For future work, we will incorporate nanoclay 56 to enhance the 3D printability of highly indicated free-standing objects.

5. Acknowledgements The authors thank Dr. Jian Yang and Ethan Gerhard from Biomedical Engineering Department at Pennsylvania State University for their assistance with mechanical testing experiments. The authors also thank Bruce Perrulli and Julia-Grace Polish from Anton-Paar USA, Inc. for their assistance with the rheology experiments. The authors are greatful to Dr.

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Dino Ravnic (Department of Surgery at the Pennsylvania State University) for providing the INCREDIBLE 3D bioprinter. The authors also thank Dr. Thomas Neuberger with his assistance with the MRI scan. Veli Ozbolat acknowledges the support from the International Postdoctoral Research Scholarship Program (BIDEP 2219) of the Scientific and Technological Research Council of Turkey (TUBITAK), and Bugra Ayan acknowledges support from the Turkish Ministry of National Education. The authors also thankful to Materials Research Institute at the Pennsylvania State University in supporting the porosity experiments.

Supporting Information Figure S1: Representative stress-strain curves of dogbone samples. Figure S2: Representative microscopic images of tested dogbone samples (Ink 9:1) showing pores in their neck section. A supplemental video capturing failure strains of dogbone samples.

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Figure 1. Rheological analysis of PDMS inks. A) Measured G' and G'' from amplitude sweep of Inks 10:0, 9:1, 8:2, 7:3 and 0:10 from 0.01% to 100% strain at 25 ˚C. B) Measured complex viscosity (η*), G' and G'' from frequency sweep of Inks 7:3, 8:2 and 9:1 at 25 °C. C) Measured viscosity of Inks 10:0, 9:1, 8:2, 7:3 and 0:10 from rotational testing from 0.01 s-1 to 1000 s-1 at 25°C. D) Shear strain vs shear stress and calculated yield points for Inks 10:0, 9:1, 8:2, 7:3 and 0:10 at 25 °C. E) Measured G' and G'' from a 3 Interval Thixotropy Test (3ITT) of Inks 9:1, 8:2 and 7:3 at 25 °C.

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Figure 2. 3D printing of PDMS inks: A) Experimental setup, B) a concept figure of dual-layer printing, C) different structures of dual-layer constructs with their corresponding Pr values, D) optical microscope images for Inks 9:1, 8:2 and 7:3) and three nozzle tips (610, 510 and 410 µm) at 250 kPa extrusion pressure and 180 mm/min printing speed.

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Figure 3. Impact of printing parameters: A) Ink concentration, B) tip diameter, C) extrusion pressure on the printed filament width for ten different printing speeds.

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Figure 4. A) 3D printed organ models: A1) human hand, A2) nose, A3) blood vessel, A4) ear, A5) branched structure, A6) femoral head. B) Fidelity analysis of printed nose geometry: B1) Heat maps and B2) average percent frequency histogram representing surface deviations between printed and model geometries.

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Figure 5. Mechanical characterization of PDMS samples. A) A 3D printed dogbone-shaped test samples in transverse and longitudinal directions. Comparison of mechanical properties of 3D printed and cast inks: B) ultimate strength, C) Young’s modulus, D) failure strain (n=3) (see representative stress-strain curves in Figure S1).

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Figure 6. Cell adhesion analysis of different inks surfaces. A) An optical microscope image of 3D printed surface and B) the optical surface profile of same printed surface. C) Number of

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nuclei per mm2 and average cell area (n=200) for samples without fibronectin coating and D) with fibronectin coating. E) Immune-images of adhered cells on cast and 3D printed surfaces.

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Figure 7. Fourier transform infrared (FTIR) spectra of A) the controls and printed and nonprinted Inks B) 9:1, C) 8:2 and D) 7:3. Sylgard 184 exhibited characteristic IR bands 57. Among the most intense are those associated with –CH3 rocking between 785–815 cm−1, ≡Si–OH stretching between 825–865 cm−1, asymmetric Si–O–Si stretches 1055–1090 cm−1, symmetric – CH3 deformations ≈1245–1270 cm−1, asymmetric Si–CH3 stretches ≈2950–2970 cm−1.

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Table 1. Porosity analysis of dogbone samples (see Figure S2 for representative images of pores from dogbone samples). Ink

Porosity [%]

Fabrication technique

Average pore diameter [µm]

7:3

8:2

9:1

a)

Cast (Control)

14.7079

0.0105

Printed along transverse direction

13.7725

0.0092

Printed along longitudinal direction

13.3505

0.0088

Cast (Control)

17.2498

0.0128

Printed along transverse direction

13.3162

0.0088

Printed along longitudinal direction

13.6859

0.0092

Cast (Control)

14.2156

0.0104

Printed along transverse direction

12.7251

0.0085

Printed along longitudinal direction

12.8271

0.0092

Printing path directions are shown in Figure 5A.

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3D Printing of PDMS for medical applications 83x47mm (300 x 300 DPI)

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