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PRINTING BIOLOGICAL LIQUID ON HYDROPHOBIC 3D ELECTRODES
S. Chu1, M.J. Lerman3, J.N. Culver4, J.P. Fisher2, and R. Ghodssi1,2,3*
1Institute for Systems Research, Department of Electrical and Computer Engineering,
2Fischell Department of Bioengineering, 3Department of Materials Science and Engineering,
4Institute for Bioscience and Biotechnology Research, Department of Plant Science and Landscape Architecture,
University of Maryland, College Park, Maryland, USA
ABSTRACT
This paper presents for the first time a programmable and
scalable 3D electro-bioprinting (3D-EBP) process for patterning
bionanoreceptors, cysteine-modified Tobacco mosaic virus
(TMV1cys), on high-density micropillar array electrodes. The
structural hydrophobicity in high aspect ratio geometries of
micro/nano devices poses a critical challenge for assembling 3D
biomaterial-device interfaces. Here, we have successfully integrated
electrowetting principles with a modified state-of-the-art bioprinter
for automated, high-throughput, and large-scale patterning of
TMV1cys particles on hydrophobic 3D electrodes. The 3D-EBP
processed bionanoreceptors maintained both structural and
chemical functions as characterized via SEM and fluorescence
microscopy. Overall, the innovative 3D biomanufacturing process
creates excellent opportunities for advancing on-demand bio-
integrated devices including multiplexed biosensors and bioenergy
harvesting devices.
INTRODUCTION
The convergence between biochemistry and micro/nano
manufacturing technologies has brought revolutionary
advancements in the development of miniaturized devices and
biomaterial integrated systems [1], [2]. Technology for bio-device
integration has rapidly evolved over the past two decades based on
biochemistry principles, processing methodologies learned from the
micro/nano fabrication industry, and instrumental
science/engineering. The diversity of biofabrication technologies
have led to greater understanding of biomolecular activities through
on-chip characterizations, and is enabling rapid advancement of
high-density, multiplexed bioarrays necessary for biosensing and
diagnostic applications.
Building upon the variety of methods, many researchers have
demonstrated integration of biological materials with three-
dimensional transducers in an effort to achieve enhanced
bioelectronics performance at a miniaturized system scale [3], [4].
Three-dimensional structures at micro- and nano- scales offer a
number of beneficial properties compared to their two-dimensional
counterparts, allowing for an extensive range of applications
including micro energy storage/harvesting device [5], micro/nano
actuators [6], micro thermal management devices [7], water-
repellent surfaces [8], optical modulators [9], 3D VLSIs [10], etc.
The primary attractive feature of these approaches is the high
density of physical interfaces between system components and
materials within the restricted surface area budget, which can
directly translate into large enhancements in performance. Moreover,
precise control over the arrangement of the structures through
microfabrication or emerging 3D printing technologies further
highlights the utility of three-dimensional components with
increased surface-to-volume ratio, functional uniformity and
tunability.
One of the interesting characteristics present in small scale 3D
structures is their limited wetting property known as structural
hydrophobicity [11]–[13]. When micro- or nano- scale structural
components are densely arranged, the wetting of 3D cavities with
liquids is limited by surface tension at the solid-liquid, liquid-air,
and solid-air interfaces. Considering most biological materials are
stored in buffered aqueous solutions due to the narrow biological
stability windows, the structural hydrophobicity can be a significant
limiting factor when attempting to introduce biological materials
into the 3D cavities for device functionalization. In our recent report,
we have provided experimental evidence to highlight the potential
limitation using cysteine-modified Tobacco mosaic virus
(TMV1cys) and Au-coated Si micropillar array (μPA) electrodes. In
addition, the electrowetting principle - which controls surface
wettability with applied electric potential - has been introduced as
an enabling technique for uniform and localized immobilization of
the bionanoreceptors on densely-arranged μPA electrodes [14], [15].
Here, we have successfully integrated the electrowetting-
assisted biofabrication technique with a customized bioprinter for
automated, high-throughput, and large-scale printing of the
bionanoreceptors on 3D electrodes. The simple system components,
optimized printing process parameters including extrusion
pressure/time and printing distance, combined with the
electrowetting process determined from our previous work, allows
a novel 3D bioprinting technique with excellent repeatability and
consistency, ultimately offering a unique function to existing
bioprinting systems to bring an on-demand programmable, scalable,
and readily adaptable 3D biofabrication technology.
MATERIALS AND METHODS
Hydrophobic 3D Electrodes and Biological Ink
Figure 1: Au-coated μPA electrodes and their wettability. (a)
Fabrication process and dimensions for the μPA electrodes. A
comparison of TMV1cys solution droplet (10 μL) contact angles on
(b) planar Au and (c) Au-coated μPA (SAR=10). (d) A plot of TMV
1cys solution droplet contact angles on Au-coated μPA at different
SARs. The relationship between the contact angle/hydrophobicity
and SARs follows the trend anticipated by Cassie-Baxter equation.
978-1-940470-03-0/HH2018/$25©2018TRF 144 Solid-State Sensors, Actuators and Microsystems Workshop
Hilton Head Island, South Carolina, June 3-7, 2018
DOI 10.31438/trf.hh2018.41
The Si micropillar array (μPA) electrodes have been fabricated
through a standard microfabrication process as described in Figure
1a. A Si wafer is etched down via DRIE using a
photolithographically patterned negative photoresist (NR9-1500PY,
Futurrex) as the etch mask. The resulting Si μPAs are passivated by
500 nm PECVD SiO2 followed by sputtering of Cr (30 nm)/Au (200
nm). The pillar aspect ratio was held constant at 10 (height: 70 μm,
diameter: 7 μm) while the spacing aspect ratio (SAR) varied from
3.3 to 10 to investigate the effect of the structural hydrophobicity
level on the final pattern size/resolution.
The biomaterial used in this work is genetically-modified TMV
expressing cysteine residues on its outer surface (TMV1cys, the
details of genetic modification and purification protocols can be
found in previous report [16]). A TMV1cys solution of 0.2 mg/ml
concentration in 0.1 M, pH 7 phosphate buffer (PB) solution is used
as the bio-ink for all experiments. The enhanced metal binding
properties of TMV1cys, such as self-assembly onto Au surfaces and
surface metallization with Ni, provide effective means for
characterization of functionalization morphology after the
electrowetting assisted bioprinting process.
As an initial step, the structurally hydrophobic properties of the
μPA electrodes were characterized via contact angle measurements
of 10 μL sessile drops of the TMV1cys solution on Au-coated planar
(Figure 1b) and μPA (Figure 1c) electrodes. While the planar Au
surface exhibits a hydrophilic nature with a 65° contact angle, the
μPA surface yields more hydrophobic characteristics as indicated
by the 132° contact angle (SAR=10). This is close to the
theoretically expected value (136º), calculated based on the Cassie-
Baxter equation (Eq. 1), where θ* is the apparent contact angle on
the μPAs, ϕs is the fraction of the solid in contact with the TMV1cys
solution, and θ0 is the contact angle on the planar Au substrate [12].
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ሻ (1)
Figure 1d plots both theoretically expected (derived from
Cassie-Baxter equation (Eq. 1) with θ0=65º, and ϕs calculated from
the pillar geometry) and experimentally acquired contact angles (θ*)
indicating that the hydrophobicity of the μPAs decreases with an
increase in SAR. The disparity between the two can be attributed to
the gentle pressure applied when loading the droplet onto the μPAs
via syringe tips and hydrophilic surface (Au) inducing liquid
pinning onto the pillar tips [14]. Such wetting state is only valid
when there is no external disruption force due to the hydrophilicity
of the Au surface. Once the Cassie-Baxter state is disrupted with an
external force (e.g. mass loading), a faster transition of the wetting
state - indicated by the droplet spreading into the cavities
(transitioning to the Wenzel state) - is observed with the lower SAR
μPAs (data not shown). It becomes a significant challenge to
introduce the solution into narrower cavities (SAR >= 7) that do not
easily transition to a Wenzel state, which can be attributed to less
pressure loading per spacing segment. In other words, the wider
spacing reduces ϕs which increases the fraction of the liquid droplet
that is supported by the micropillars [11]. The TMV1cys droplets on
such high SAR μPAs are readilly movable with no traceable liquids
remaining on the electrode surface.
Integration of 3D-EBP with a Bioprinter
The 3D electro-bioprinting (3D-EBP) technique has been
integrated with a state-of-the-art commercial 3D bioprinter (3D
Bioplotter, Envision TEC) by simply connecting a function
generator (Agilent 33220A) to the printing nozzle and the substrate
as described in Figure 2a. The built-in graphical user interface
allowed a highly automated and programmable 3D-EBP with
precise control over nozzle pressure, printing distance, array
size/density, and solution temperature. As the 3D Bioplotter is
mainly designed for printing materials carried in hydrogels (e.g.
gelatin, alginate, corn starch, etc.), the controllable pressure and
time ranges for ink extrusion at the nozzle are optimal for higher
viscosity fluids compared to the buffer solution used in this work.
Through multiple iterations at the lowest extrusion time and
pressure ranges, the optimal process parameters have been
determined as 0.3 second extrusion at 0.1 bar with a distance from
the nozzle to the substrate set at 100 μm. It should be noted here that
the optimization of the process parameters strongly depends on the
nozzle size and the viscosity of the ink solution.
Figure 2: (a) Overview of the programmable 3D-bioprinting system
integrating 3D-EBP with a commercial bioprinter. (b) Illustration
of TMV solution impinging onto the μPA from teflon-modified
needles. The solution impinging onto the μPA surface without
electrowetting (left) keeps a spherical shape without further
access/spreading into the microcavities (Cassie-Baxter state), while
the droplet impinging with electrowetting (right) selectively wets the
underlying cavities (Wenzel state).
In this initial demonstration, a 32-gauge stainless steel needle
(inner diameter: 100 μm, outer diameter: 260 μm) was utilized with
a hydrophobic modification at the tip via dip-coating with Teflon;
the tip was dipped into Teflon solution (TeflonTM AF 400S1) and
cured at 180°C for 3mins. A syringe was connected during the
curing step to continuously flow air through the tip to prevent
blockage during the curing process. The hydrophobic coating at the
tip was critical to isolate the droplet forming towards the substrate
during extrusion (Figure 2b); otherwise the extruded droplets would
roll-up to the outer metallic surface causing inconsistency in droplet
size and printing failures.
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The electrowetting voltage was continuously applied during
the printing process by seamlessly integrating the function generator
to the metallic -nozzle and -printing bed without hindering the
printing operation. Based on our previous work, a sinusoidal voltage
signal of 1.2 Vrms at 10 kHz was applied to induce uniform and
localized wetting of the TMV1cys ink into the μPAs [14]. All
processes were conducted at room temperature with ambient
humidity. Both the print-bed and the bio-ink cartridges were kept at
10°C for extended stability of TMV1cys particles throughout the
multiple printing processes. The bio-printed electrodes were
incubated overnight in a humid chamber for self-assembly of
TMV1cys particles onto the electrode surface through thiol-Au
binding [15].
RESULTS AND DISCUSSIONS
Bio-ink Printing on 3D μPA Electrodes
Figure 3 compares the printing processes on μPA electrodes
(SAR: 10) under two different conditions emphasizing
electrowetting as an enabling method. When the printing process is
conducted without applying the electrowetting voltage (Figure 3a),
the extruded droplets at the printing nozzle do not settle onto the
μPA surface, and collectively accumulate at the nozzle until falling
onto the surface due to gravity (the droplets loaded onto the μPA
surface does not penetrate into the cavity). However, with the
application of the electrowetting technique (Figure 3b), the 3D-EBP
allowed consistent dispensing of ~1 uL of the bio-ink on the μPA
electrodes throughout the programmed array patterns by
successfully introducing the bio-ink into the designated micro-
cavities. The spacing between the droplets has been set to 1 mm to
prevent adjacent droplet from merging. Particularly for creating
discrete patterns, the liquid merging between the neighboring
droplets becomes problematic for lower SAR μPA electrodes as they
are more susceptible to lateral spreading of liquid during the
electrowetting process (further discussed in the next section).
However, the merging event can be utilized for creating continuous
patterns on the 3D substrate.
Figure 3: Optical images comparing the TMV1cys solution droplet
printed μPA electrodes (SAR=10). (a) Hydrophobic nature of the
μPA electrodes repels the droplets being loaded onto the surface
resulting in failure in printing/patterning the droplets. (b)
Continuous application of electrowetting voltage during the
printing process allows successfully patterning of uniform droplet
volumes on the μPA electrodes.
Functionalization Morphology and Pattern Sizes
In order to characterize the TMV1cys printed on the μPA
electrodes using SEM, an electroless Ni coating was performed after
the overnight incubation step. As shown in Figure 4a, all μPAs
featuring different SARs/densities were successfully coated with the
TMV1cys at the printed spots. The top-down SEM images taken at
the droplet boundaries confirm the localized patterning of the
TMV1cys within the 3D substrate with more surface functionalized
particles identified on the printed side (left from the light-blue line)
compared to the non-printed area (right from the light-blue line).
However, it should be noted that the TMV1cys density on the μPA
electrodes with higher SAR is noticeably less compared to that
observed from the lower SAR electrodes. This can be attributed to
limited TMV1cys diffusion from top to bottom of the electrodes for
dense microstructures. Comparing with the μPAs without
electrowetting (Figure 4b), the nanoparticles assembled on the lower
bottom of the pillar surfaces in Figure 4a (SAR: 10) confirm that the
ink solution has successfully reached the bottom surface with
electrowetting, indicating that the use of higher concentration ink
can help increase the biofunctionalization density on high SAR μPA
electrodes.
Figure 4: Process evaluation using SEM. (a, cross-section view)
TMV1cys particle coating on all sidewalls of the μPAs has been
achieved (observed as surface textures). (a, top-down view) Also,
clear functionalization boundary has been observed at the wetting
edges (blue-dotted line) of each of the μPAs highlighting the pattern
fidelity enabled by structural hydrophobicity. (b) The clean surface
of the μPAs processed without electrowetting further emphasizes the
electrowetting as an enabling technique for bio-printing. (c) The
pattern sizes ranged from 0.89mm2 to 0.37mm2 (N=5) following the
hydrophobicity levels of the μPAs. (scale bars: 10μm).
The dependence of the pattern/spot sizes on the SARs/pillar
densities have been characterized by estimating semi-circular
pattern boundaries observed from top-down SEM images. Figure 4c
compares the pattern sizes measured from the μPA electrodes with
different SARs at the respective contact angles - representative of
their hydrophobicity levels. The lower SAR μPA electrodes resulted
in a larger pattern size due to the facile spreading of the bio-ink into
the microcavities with the electrowetting (under the identical
process settings, which is in a good agreement with our previous
investigation. The statistical analysis using five different locationss
of a printed array show a high consistency in pattern size per SARs,
emphasizing the excellent uniformity and repeatability of the 3D-
EBP process.
Scalable Biofunctionalization Density
Biochemical activity of the printed TMV-1cys particles is
evaluated using a sulfhydryl (-SH on cysteine) specific fluorescent
labeling reagent, Fluorescein-5-Maleimide (Thermo Fisher
Scientific). As our previous report confirmed that the electrowetted
TMV1cys on μPAs enhances bionanoreceptor density by surface
area enhancement (SAE) factors anticipated from the 3D electrode
geometries [14], this work has focused on achieving scalable
nanoreceptor density per different SARs of the μPAs. Figure 5a
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compares top-down fluorescent microscopy images taken from the
printed spots on different electrode geometries. Under an identical
exposure time, TMV1cys printed on μPA electrodes resulted in a
significantly higher fluorescence compared to the planar electrodes
with an excellent patterning fidelity (Figure 5a, bottom-right), due
to the underlying microstructure. However, there is a noticeable
discrepancy between the change in increment factor of the
fluorescence intensity and the SAE factor calculated based on the
μPA geometry (Figure 5b). This is attributed to the lower density of
TMV1cys on the high SAR μPAs as supported by the SEM
characterizations in Figure 4a, indicating that control over
concentration of bio-ink may need to be optimized to obtain highly
controllable functional scalability of the resulting nano/micro/bio-
integrated components.
Figure 5: Characterization of fluorescence intensity from (a)
TMV1cys functionalized on planar Au and μPA electrodes
displaying SARs ranging from 3.3 to 10 (scale bars: 100μm). (b) A
significant increase in functionalization density is achieve with the
μPAs as reflected in the increase in fluorescence intensity (N=5).
CONCLUSIONS
This work offers a unique function to existing bioprinting
systems bringing a simple, innovative, and technology enabling 3D
biofabrication through integration of the core manufacturing
technique based on the electrowetting principle. The presented
system is fully automated, scalable, and readily adaptable for a wide
range of applications. While the presented demonstration utilizes a
particular biomaterial of interest to our work (TMV1cys), this
manufacturing technology can be applicable for integrating other
micro/nanomaterials (DNA, proteins, cells, carbon nanotubes,
graphene, etc.), stored in aqueous media, onto conductive
hydrophobic electrodes. Possible applications include areas
developing energy storage/harvesting devices, biochemical sensors,
optical metamaterials, heat management devices, superhydrophobic
surfaces, and more. Overall, the innovative 3D biomanufacturing
process generates excellent opportunities for advancing on-demand
bio-integrated devices opening up unprecedented possibilities in
micro/nano/bio integrated fabrication technologies and platforms.
ACKNOWLEDGEMENTS
This work was funded by the Biochemistry Program of the
Army Research Office (W911NF-14-1-0286). The authors would
like to acknowledge the support of the Maryland Nanocenter and its
FabLab for support in fabrication and imaging processes.
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CONTACT
*R. Ghodssi, tel: +1-301-405-8158; ghodssi@umd.com
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