# Fabricating a dielectrophoretic microfluidic device using 3D-printed moulds
and silver conductive paint

Shayan Valijam  Faculty of Electrical Engineering, K. N. Toosi University of
Technology, Tehran, 1631714191, Iran  Department of Physics, Umeå University,
Umeå, 901 87, Sweden  Daniel P.G. Nilsson  Department of Physics, Umeå
University, Umeå, 901 87, Sweden  Dmitry Malyshev  Department of Physics, Umeå
University, Umeå, 901 87, Sweden  Rasmus Öberg  Department of Physics, Umeå
University, Umeå, 901 87, Sweden  Alireza Salehi  Faculty of Electrical
Engineering, K. N. Toosi University of Technology, Tehran, 1631714191, Iran
Magnus Andersson  Department of Physics, Umeå University, Umeå, 901 87, Sweden
Umeå Center for Microbial Research (UCMR), Umeå, 901 87, Sweden
<EMAIL_ADDRESS>
###### Abstract

Dielectrophoresis is an electric field-based technique for moving neutral
particles through a fluid. When used for particle separation,
dielectrophoresis has many advantages compared to other methods, providing
label-free operation with greater control of the separation forces. In this
paper, we design, build, and test a low-voltage dielectrophoretic device using
a 3D printing approach. This lab-on-a-chip device fits on a microscope glass
slide and incorporates microfluidic channels for particle separation. First,
we use multiphysics simulations to evaluate the separation efficiency of the
prospective device and guide the design process. Second, we fabricate the
device in PDMS (polydimethylsiloxane) by using 3D-printed moulds that contain
patterns of the channels and electrodes. The imprint of the electrodes is then
filled with silver conductive paint, making a 9 pole comb electrode. Lastly,
we evaluate the separation efficiency of our device by introducing a mixture
of 3 µm and 10 µm polystyrene particles and tracking their progression. Our
device is able to efficiently separate these particles when the electrodes are
energized with ±12 V at 75 kHz. Overall, our method allows the fabrication of
cheap and effective dielectrophoretic microfluidic devices using commercial
off-the-shelf equipment.

###### keywords:

DEP, LOC, lab on a chip, PDMS

##  1 Introduction

Microfluidic devices that can separate micro-particles have important
applications in a variety of fields. These include medicine, chemical
engineering, wastewater treatment, environmental assessment, forensic
identification, and cell separation [1, 2, 3, 4]. Separation of micron-sized
particles can be done using different approaches; however, for both biological
and non-biological particles, dielectrophoresis (DEP) has proven to be a
powerful method. The technique offers label-free, rapid, and controllable
manipulation, which has been shown to perform well in both low and for high-
throughput applications [5, 6]. Dielectrophoresis specifically refers to the
interaction between a dielectric (non-conductive) particle, and a non-uniform,
alternating electric field. Consequently, the DEP force depends both on the
particle characteristics, such as size, morphology, and dielectric properties
[7]; as well as the frequency of the electric field and how the electrodes are
shaped and positioned, as shown in Fig. 1. If the electrodes are positioned
next to each other, the non-uniform electric field near the electrodes results
in a force deflecting the particles away from their path allowing for particle
separation.

Effective particle separation requires an appropriate flow rate to transport
the particles. This is often achieved using microfluidic channels, which allow
for controlled and laminar flow conditions. Microfluidic devices, like those
used for DEP, can be fabricated using various methods such as lithography,
laser photoablation, hot embossing, and direct 3D printing [8, 9]. Soft
lithography is of special interest, as it is relatively cheap and easy to use.
This technique uses bio-compatible polymers like polydimethylsiloxane (PDMS)
to fabricate microfluidic structures around premade moulds. By printing these
moulds using high-precision 3D printers, such as commercial stereolithography
(SLA) printers, it is possible to produce moulds with spatial resolutions down
to 5 µm [10, 11, 12, 13]. This allows us to quickly produce small channels
with a high degree of precision.

One common technique to generate wide electric fields for electrokinetic
particle separation is by using planar comb electrodes. However, the
fabrication process of these electrodes often requires expensive techniques
like physical vapour deposition (PVD), micromachining, or inkjet printing [14,
15, 16, 17]. Previous works have shown that it is possible to integrate the
electrodes directly into the walls of a microfluidic channel using an Ag-PDMS
mix; however making the nanoparticles distribute evenly throughout the PDMS
has shown to be a challenge [18]. A more affordable and accessible technique
for creating comb electrodes is directly applying them on the microfluidic
channel. By creating a 3D-printed mould for the electrodes, it is possible to
simply fill the mould with conductive material, such as silver conductive
paint, and join the electrodes to the microfluidic channel.

In this work, we present a method for making DEP microfluidic devices for
particle separation that is quick and does not need expensive equipment or a
cleanroom. We design, optimize, and evaluate our device using finite element
method (FEM) simulations. We then fabricate the device by using 3D printing,
PDMS moulding, and conductive silver paint to create the microfluidic channels
and comb electrodes. The electrodes are energized with AC voltage. Finally, we
use high-speed imaging and particle tracking to verify and evaluate the
separation of 3 µm and 10 µm polystyrene beads.

Figure 1: The principles of dielectrophoresis on a non-conductive neutral
particle. Panel A shows that a neutral particle in an alternating uniform
field will have balanced forces and remain stationary. In panel B, the
alternating field is instead non-uniform and the resulting net force acting on
the particle will make it move. If the frequency of the field is tuned so the
particle moves toward the lower field intensity it is denoted as negative DEP.
Panel C shows that when the electrodes are positioned next to each other (in a
comb pattern), the electric field is stronger near the electrodes resulting in
a repelling force, thus pushing the particles into the center of a
microfluidic channel.

##  2 Results and Discussion

###  2.1 Designing the electrodes and determining their electrical parameter
using simulations

To make a low-cost microfluidic device that can separate micro-sized particles
using dielectrophoretic forces, we used a design-build-test approach. First,
we designed the device using FEM simulations in COMSOL Multiphysics (v5.5,
COMSOL AB). This allowed us to optimize the layout of the channels,
electrodes, and electric parameters before constructing the physical device
[19]. In our design, we took into consideration that it should be possible to
produce the channels and electrodes using a 3D printer for cheap and fast
fabrication. The general design of the device is shown in Fig. 2A, with a
detailed schematic in Fig. 2B showing the layout of the inlets, electrodes,
and outlets. We made the main channel 250 µm wide, 35 µm high and 10 mm long,
with approach channels extending out at 45∘ to the in- and outlets. These
allow us to introduce and extract particles from the main channel. Along the
side of the main channel, nine electrodes are positioned in a comb pattern.
These are 150 µm wide, placed with a center-to-center distance of 250 µm and
protruding 20 µm into the flow channel. The electrodes are made flush with the
channel floor and extend to a depth of 35 µm into the bottom layer. Both the
top and bottom layers are made in PDMS (polydimethylsiloxane), which is a
transparent, flexible and non-toxic silicone rubber. This compact design makes
it possible to fit the dielectrophoretic microfluidic device on a microscope
glass slide and observe the separation of biological samples in real-time.

Figure 2: Panel A shows the design of the dielectrophoretic microfluidic
device where the microfluidic channel (top layer) and electrodes (bottom
layer) are placed on thin layers of PDMS. In panel B, we show a compounded
(top) view with all the features labeled. The electrodes extend into the
channel and are routed back to connection pads that are energized with AC
voltage. Panel C shows the CM factor plot. The real part of the CM factor is
negative for both particle sizes for frequencies above 71.6 kHz (dotted line).
Panel D shows a simulation of the relative electric fields in the comb
electrode, when operated at 75 kHz.

We designed the device so that inlet A provides a controllable sheath flow
through the main channel. This sheath flow focuses particles from inlet B
towards the sidewall where the electrodes are positioned, enhancing the DEP-
force from the electrodes felt by the particles. To generate an electric field
between the electrodes, we connect them to +V and -V in an alternating
pattern, see Fig. 2B. Our simulations suggest that particles are efficiently
focused toward the electrodes using fluid rates of 70 µL/minute and 14
µL/minute for the sheath flow and particle flow, respectively. When particles
enter the separation stage (region of the nine electrodes), they will be
subjected to DEP forces in the y-direction, pushing them away from the
electrodes. This repulsive DEP where the particles move away from the strong
electric field is commonly referred to as negative DEP (nDEP). Depending on
the size and electrical properties of the particles, the nDEP force will
differ in magnitude, resulting in separation between the particle species. Our
aim is to separate 3 µm and 10 µm polystyrene beads into outlets C and D.
These particles are approximate size analogues to platelets and red blood
cells, respectively, making them relevant to biological applications. We
optimize this separation by tuning the layout of the device, the electrode
voltage and the driving frequency. Since the driving frequency for the
electrodes determines the nDEP force, we determined the Clausius–Mossotti (CM)
factor by solving Eq. 3. The CM factor is a complex number that describes the
polarizability of particles and its real part for our particles is shown in
Fig. 2C. A negative real part of the CM factor results in a repulsive force
from the electrodes on the particles, this occurs in both of our used particle
sizes at frequencies above 71.6 kHz. At these frequencies, it becomes possible
to separate different-sized particles using nDEP, as the nDEP force felt by
the particles is proportional to the cube of the particle radius, as seen in
Eq. 2. Increases in the driving frequency will result in a larger nDEP force;
however, high frequencies are known to cause air bubble formation, and as
such, we use a driving frequency of 75 kHz in our experiments [18].

Since our aim is to make electrodes from silver conductive paint in our final
device, we produced prototype electrodes to evaluate their electrical
properties for use in our simulations. We found the conductivity of electrodes
to be 2.5 S/m. From the simulation, we calculated the electric field strength
throughout the microfluidic system to be $\sim$0.017 V/µm in the center of the
channel and $\sim$0.450 V/µm close to the electrodes, with the full field
distributions visualized in Fig. 2D. To find the lowest voltage that yields
complete separation, we simulated a particle flow through the channel while
energizing the electrodes at different voltages. In this simulation, we
introduced 1 ml of buffer solution containing 4 000 randomly distributed
particles (2 000 of each size) at a constant flow rate. The properties of the
particles and the buffer are listed in table 1. During the simulation, we
monitored the particle motion in three parts of the channel as seen in Fig. 3;
before the separation stage (I), at the separation stage (II), and at the
separation junction (III). From snapshots of these simulations for different
applied voltages we found that ±9 V was not sufficient to separate the two
sizes (0 % of 10 µm particles flow through outlet C). By increasing the
voltage to ±10 V, the particles are partially separated (42 % of 10 µm
particles flow through outlet C), and at ±11 V, particles are fully separated
(100 % of 10 µm particles flow through outlet C). The particle motion in these
simulations can be seen in the supporting movies S1-S3, respectively. We
therefore conclude that energizing the electrodes at ±11 V (75 kHz) should be
sufficient for the complete separation of 3 µm and 10 µm particles.

Figure 3: Numerical simulation to evaluate the voltage dependency on
separation of 10 µm (red) and 3 µm (blue) particles. We test this for ±9, ±10,
±11 V and observe the particles in three key marked regions of our device: at
the focusing stage (I), at the separation stage (II), at the separation
junction (III). At 11 V, all particles are separated successfully according to
their size.

Increases in the electric field strength result in a higher separation
efficiency; however, it also causes more Joule heating. Heating the solution
will change its conductivity and permittivity, and excessive heating may be
harmful to biological cells [20]. To estimate how the temperature increases
with applied voltage, we calculated the fluid temperature as

$\Delta
T\approx\frac{\sigma_{\text{m}}\times{V_{\text{rms}}}^{2}}{2K_{\text{m}}},$
(1)

where $V_{\text{rms}}$ is the magnitude of the applied voltage, and using
$\sigma_{\text{m}}$ = 100 mS/m and $K_{\text{m}}$ = 0.60 W/m$\cdot$K for the
electrical and thermal conductivity, respectively [21]. At full separation
(±11 V) we would have an increase of 10 K in the sample fluid, suggesting that
voltages in this range will not significantly affect device performance or the
particles.

###  2.2 Building the device and testing its performance

Using the results from our simulation, we built a lab-on-a-chip device using a
3D printing and PDMS moulding approach, similar to Ho et al., 2015 [22].
First, the design of the simulated device was exported to our SLA resin
printer. Using this printer, we produced a high-resolution (50 µm) printout of
the channel, onto which 1 mm of PDMS was moulded to make the microfluidic
layer of the device, as seen in Fig. 4A. Second, a similar PDMS layer was made
with imprints of the electrodes and these were filled with silver conductive
paint, as seen in Fig. 4B. Third, holes were made in the microfluidic layer
for the in/outlets and then the two layers were bonded together, as seen in
Fig. 4C. Finally, nine copper wires were connected between the electrode pads
and a signal generator (Model 8102, Topward), set to operate at ±11 V and 75
kHz. The manufacturing process is described in greater detail in Sec. 4.3, and
a photo of the finished device is shown in Fig. 5A. To allow fluid flow
through the device, we added tubing to the inlets and outlets and connected
the device to a syringe pump (VIT-FIT, LAMBDA Laboratory Instruments). The
injection rates for inlets A and B were set to a continuous 70 µL/minute and
14 µL/minute as suggested by simulations.

Figure 4: The main steps in the procedure of making a nDEP microfluidic
device. For visualization purposes, the illustration is not to scale and cut
in half along x-axis. In panel A, we show how the top layer of the device is
made. PDMS is poured into a 3D-printed mould (and its frame). Once cured, the
PDMS is peeled off with an imprint of the flow channel. In panel B, a similar
procedure for the bottom layer is shown. The imprint for the electrodes is
filled with silver conductive paint after the PDMS is cured. In panel C, a
biopsy punch is used to make openings in the channel before both halves are
bonded together. To improve the bonding along the electrodes, we cut the
corresponding part of the top layer away and added uncured PDMS at the seam
using a syringe.

To inlet B, we connected a syringe containing a phosphate-buffered saline
(PBS) solution (0.1 S/m) and a mixture of 3 and 10 µm beads, as defined in
Sec. 4.4. The device was then placed in an optical microscope (Microphot-FX,
Nikon Instruments) with a 20X objective (427958, Nikon Instruments) for
tracking particles through the channel. Due to the high particle velocities,
we mounted a high-speed CCD camera (MotionBLITZ EoSens Cube7, Mikrotron) and
set it to record videos at 1696x1240 pixel resolution and 735 frames per
second. Particle trajectories were analyzed from the video using the free
tracking software, ToxTrac (v2.98) [23, 24]. We first analysed region I
(before the separation stage) to see if the flow ratio from the simulations
would focus the particles to the side where the electrodes were positioned.
Indeed, we observed that particles were focused closer to this side and that
particles moved with the flow and were not exposed to external forces, see
Movie S4 and Figure S1A. Next, we analysed region II (separation stage) and
region III (junction). Initially, when using the same voltage as in the
simulation (±11 V), we observed that particle separation was not satisfactory.
We therefore increased the voltage to ±12 V, at which full separation was
achieved. Since the electric field is the cause of Joule heating, we
recalculated the temperature increase using Eq. 1. At a separation voltage of
±12 V the total temperature increase is 12 K, which is only 2 K higher than at
±11 V, and still low enough to not affect the device or the sample
significantly.

Running the device at ±12 V, we analysed the trajectories of the larger (10
µm) particles at the separation stage (electrodes 4 to 7) and found that these
particles had a noticeable change in their y-directions, see Movie S5 and
Figure S1B. The gradient of the 10 µm particles was assessed to 0.042±0.0013,
by fitting a linear regression model to the trajectories. This implies that
the 10 µm particles moved about 6.3 µm in the y-direction over a 150 µm
distance (equal to the width of an electrode).

After this, we monitored the separation junction. At the separation junction,
we observed that particles were clearly separated by size, see Movie S6. To
quantify this, we used Toxtrac and set the software to track particles
according to size. Trajectories of about 30 particles from each group are
shown in Fig. 5B, in which we have plotted 3 µm and 10 µm particles in green
and blue, respectively.

When imaging the microfluidic channels, we noted a wave-like pattern along the
channel walls. These waves are caused by exposure bleeding (light leakage from
LCD pixels) during 3D printing of the moulds and can be mitigated by tuning
the printer and using resin additives. To ensure this pattern does not affect
particle separation efficiency, we simulated the impact of these artifacts, as
seen in Figure S2. The results show that the wave-like pattern does not affect
the particle trajectory as compared to channels with straight walls. Rather,
only subtle changes to the flow profile close to the channel walls were
observed. This is similar to what we observed in experiments, where we noted
that particles that move close to the surface follow the streamlines of the
wall. However, these streamlines do not significantly impact the particle
separation efficiency.

Figure 5: Panel A shows our microfluidic chip with the electrodes visible and
a cm scale bar above. The separation junction is marked with a dashed box in
Panel A and is shown in high resolution in Panel B. Particle traces are shown
for 3 µm (green) and 10 µm (blue) particles when energizing the electrodes at
±12 V. Particle traces before the separation stage and at the separation stage
are shown in Figure S1 Panel A and B, respectively.

###  2.3 Evaluating the microfluidic channels and electrodes

To evaluate the robustness of the silver conductive paint electrodes, we
tested their durability and observed their surface morphology. To test their
durability, we energized the electrodes at a higher voltage than what was used
during the experiments, ±15 V compared to ±12 V, at the same frequency of 75
kHz. The electrodes were also immersed in a buffer solution for 1 h and heated
to 100 ∘C. After these tests, we imaged the electrodes using SEM to evaluate
their surface integrity, as shown in Fig. 6. We found that none of the three
stress tests affected the surface morphology of the painted electrodes
compared to controls, either on the macro-structure as seen in Fig. 6A, or on
the microstructure as seen in Fig. 6B-C. Thus, we concluded that silver
conductive paint electrodes are sufficiently durable for the purpose of our
dielectrophoretic device.

Figure 6: EM micrographs of the comb electrode after stress-testing with 15 V
for 1 hour, followed by immersion in a buffer solution and heating to 100 °C.
In panel A, the electrodes are still showing an intact structure (scale bar is
100 µm). The microstructure of the electrodes at 5 000$\times$ magnification
is shown, before (B) and after (C) exposure, with no visual differences (scale
bars are 10 µm).

##  3 Conclusion

In this study we developed a lab-on-a-chip device that utilizes
dielectrophoresis to sort uncharged microparticles based on their size using
3D-printed microfluidic channels equipped with silver conductive paint
electrodes. We designed our device with FEM simulations to optimise the
electrodes’ frequency and voltage for sorting 3 µm and 10 µm plastic particles
and we built the device in PDMS using 3D-printed moulds that contain imprints
of the channels and electrodes. We evaluated the performance of the chip using
particle tracking, showing our device could successfully separate 10 µm from 3
µm particles. Thus, our proposed method allows for the fabrication of cheap
dielectrophoretic microfluidic devices using off-the-shelf equipment, that can
be used in various sorting applications, for example, sorting cells,
wastewater treatment, and separating bacterial spores from environmental
samples for detection.

##  4 Materials and Methods

###  4.1 Theoretical background

Dielectric (non-conductive) particles become polarized when placed in a non-
uniform electric field. When polarised, these particles experience a force,
the magnitude of which depends on the polarisability of the particle [25].
This is the dielectrophoresis (DEP) force, as shown in Fig. 1, and differences
in this force can be used to redirect and sort particles. For homogenous
spherical particles, the DEP force is given by

$F_{\text{DEP}}=2\pi
r^{3}\varepsilon_{\text{0}}\varepsilon_{\text{m}}\real[CM(f)]\divergence\absolutevalue{\vec{E}}^{2},$
(2)

where $r$ is the radius of the particle, while $\varepsilon_{\text{m}}$ and
$\varepsilon_{\text{0}}$ are the permittivity of the suspension medium and
vacuum, respectively. The gradient of the electric field is given by
$\divergence\absolutevalue{\vec{E}}^{2}$. Finally, Re$[CM(f)]$ represents is
the real part of the Clausius-Mossotti $(CM)$ factor. The $CM$ factor can vary
between -0.5 and 1.0 and is dependent on the particle’s complex permittivity
and thus the operating frequency $f$ of the applied electric field. This
factor also determines what direction the particle will move [26]. If $CM$ is
positive, the DEP force is towards the stronger electric field, referred to as
pDEP. If the $CM$ factor is negative, the particle moves away from the
stronger electric field, and is called nDEP [27]. The device we use in this
study works with nDEP, as shown in Fig. 2C. The $CM$ factor is quantified as

$CM=\frac{\varepsilon_{\text{p}}^{*}-\varepsilon_{\text{m}}^{*}}{\varepsilon_{\text{p}}^{*}+2\varepsilon_{\text{m}}^{*}}.$
(3)

where the complex permittivity of the particle and medium are given by
$\varepsilon_{\text{p}}^{*}$ and $\varepsilon_{\text{m}}^{*}$ respectively.
Thus the $CM$ factor value can be between -0.5 ($\varepsilon_{\text{p}}^{*}$
$<<$ $\varepsilon_{\text{m}}^{*}$) and 1 ($\varepsilon_{\text{p}}^{*}$ $>>$
$\varepsilon_{\text{m}}^{*}$). These two parameters are defined as

$\left\\{\begin{array}[]{lll}\varepsilon_{\text{p}}^{*}=\varepsilon_{\text{p}}-j\frac{\sigma_{\text{p}}}{w},\\\
\varepsilon_{\text{m}}^{*}=\varepsilon_{\text{m}}-j\frac{\sigma_{\text{m}}}{w},\\\
\end{array}\right.$ (4)

respectively, where the conductivity of the particle and medium are
$\sigma_{\text{p}}$ and $\sigma_{\text{m}}$, respectively. Further, $w=2\pi f$
is the angular frequency and $j=\sqrt{-1}$.

The other major force affecting the movement of the suspended particles in
microfluidic devices, is the hydrodynamic drag force. Hydrodynamic drag force
for spherical particles in laminar flow is calculated from Stokes’ law

$F_{\text{Drag}}=6\pi r_{\text{ext}}\eta(\vec{u}-\vec{u}_{\text{p}}),$ (5)

where $r_{\text{ext}}$ is the external radius of the particle, $\eta$ is the
viscosity of the medium, $\vec{u}$ and $\vec{u}_{\text{p}}$ are the flow
velocity and the particle velocity respectively. We defined the physical and
electrical properties of the particles and the medium as described in the
literature [28, 29], and the parameters used in our simulations are listed in
Table 1.

###  4.2 Numerical Simulation

We simulated the device using a three dimensional model in COMSOL Multiphysics
(v5.5, COMSOL AB). We used the laminar flow module to solve the Navier-Stokes
equation and obtain the flow field. The AC/DC module was used to simulate non-
uniform electric field by solving Laplace’s equation. The fluid (water) was
assumed incompressible, and the flow was assumed to be laminar and steady.
Both these are reasonable assumptions for flows at these low Reynolds numbers.
A non-slip boundary condition was applied to the channels, and the pressure at
all inlets and outlets was kept zero. A constant flow was assumed for the
inlets, with the injection flow rate for inlet A and B set to 70 µL/min and 14
µL/min respectively. Newton’s second law was employed to solve the particle
trajectory, given by

$\left\\{\begin{array}[]{lll}\nabla{p}+\eta\laplacian{\vec{u}}+\vec{F}_{\text{e}}=0,\\\
\laplacian{u}=0,\\\ \divergence\vec{u}=0,\\\ \end{array}\right.$ (6)

where $P$ and $\vec{F}_{\text{e}}$ are the pressure and applied forces,
respectively. The properties of the suspension medium were set to those of
water, with density 1000 kg/m3, dynamic viscosity 1.0$\times 10^{-3}$
Pa$\cdot$s, and relative permittivity 80 [30]. The Laplace equation was solved
to investigate applied electrical potential

$\laplacian{\varphi}=0,$ (7)

where $\varphi$ is the electric potential applied to the electrodes, while all
other boundaries were considered insulating. By solving Eq. 2 and Eq. 4, the
position of the particles were obtained and calculated using

${\vec{F}_{\text{DEP}}}+{\vec{F}_{\text{drag}}}=m\dfrac{d\vec{u}_{\text{p}}}{dx}$
(8)

where $m$ and $t$ are the mass of the particle and time, respectively. Our
simulation model counted 38 886 mesh elements in total.

Table 1: Physical and dielectric properties of the particles and suspension medium. Property | Particle | Medium
---|---|---
10 µm | 3 µm
Conductivity (S/m) | 2.5e-2 | 4.5e-2 | 0.1
Relative permittivity | 5.0$\times\varepsilon_{\text{0}}$ | 2.56$\times\varepsilon_{\text{0}}$ | 80$\times\varepsilon_{\text{0}}$
Dynamic viscosity (Pa$\cdot$s) | - | - | 1.0e-3
Density (kg/m3) | - | - | 1.0e+3

###  4.3 Device fabrication

We manufactured our device using 3D printing and PDMS moulding. Our design
consists of two halves, a microfluidic half, and a half containing the
electrodes, as seen in Fig. 4. 3D moulds for both these parts were made, and
then used to create the device geometry in PDMS (Sylgard 184, Dow Corning). To
make the moulds quick and cheap to manufacture, we used a desktop SLA 3D
printer (Photon Mono SE, Anycubic). The geometry of the device was designed in
COMSOL Multiphysics and prepared for the printer using a slicer software
(Photon Workshop V2.1.29, Anycubic). For printing, the channels and electrodes
were inverted (protruding) and printed with a 1 mm thick backing that will
form the bottom of the mould. To ensure accurate dimensions of the features,
print settings had to be adjusted in the slicer software. Settings of layer
thickness and exposure times are dependent on both the printer and resin. This
calibration was done first using the coverslip method, as described in [31],
and then by printing open-source calibration models.

Before we could start moulding, the printed parts had to be treated to make
them inert, as failing to do so would inhibit the curing of the PDMS. This
procedure consisted of; first, cleaning the part in isopropanol for 3 min
using a sonicator (Super RK 100, Bandelin Sonorex). Second, curing the part in
a UV box (15 W at 400 ± 10 nm FWHM) for 2 h. And lastly, degassing the part in
an oven at 70 ∘C for 6 h. When the part was ready, it was clamped down (facing
up) with a frame to complete the mould. A mixture of 10:1 PDMS base and curing
agent was then poured into the mould to a thickness of 1 mm and degassed with
a vacuum desiccator. It was then placed in an oven at 70 ∘C for 1 h to cure.
The PDMS layer containing the imprint was finished and could be peeled off.
This process was repeated for both parts of the device. In the top part,
through-holes were made for in and outlets to the flow channel using a 1 mm
biopsy punch (Integra Miltex).

In the bottom part; to make the electrodes flush with the channel floor, we
filled the imprints in the PDMS layer with 3 µl silver conductive paint
(SCP03B, Electrolube), scraping off the excess. The paint consists of 45$\%$
silver particles, has a viscosity of 70 mPa$\cdot$s (at 25∘C) and dries in
$\sim$10 minutes. Both of the PDMS layers were then treated in $O_{\text{2}}$
plasma for 1 min (0.5 mbar chamber pressure) using a plasma cleaner (Atto,
Deiner Electronic), before being aligned and bonded together under an optical
microscope. Finally, the device was placed on a hotplate for 5 min (at 90∘C)
to improve the adhesion. However, it was found that some fluid could leak out
where the electrodes bonded to the top layer. We therefore cut away the part
of the top layer in contact with the electrodes and rebonded the two halves.
Uncured PDMS was then poured on the exposed electrodes and cured, sealing the
joint. To apply a voltage to the electrodes, we connected 9 copper wires to
the extended silver pads. The channels were then washed with deionized water
(for 2 min) ahead of use.

###  4.4 Sample Preparation

To test the performance of the device, we prepared a mixture of 3 ± 0.09 µm
(C37484, Invitrogen CML Latex, Thermo Fisher Scientific) and 10 ± 1.0 µm
(C37259, Invitrogen CML Latex, Thermo Fisher Scientific) latex polystyrene
beads. The stock volume for 10 and 3 µm particles are 4.1 and 4.3 g/mL,
respectively, and were diluted 3000 times with deionized water to achieve
appropriate concentrations. To achieve a medium better suited for nDEP, we
modified the electrical conductivity of the particle and sheath flow medium to
0.1 S/m, by adding 1X phosphate-buffered saline (PBS) solution at a volume
ratio of 1:100.

###  4.5 Particle tracking

To track the particles in our microchannel, we used the free image tracking
software called ToxTrac (v2.98) [23]. Videos were first preprocessed in
UMUTracker [32] to remove the background and binarize the videos. The videos
were loaded in Toxtrac and we defined the region that we wanted to track as an
arena. In Toxtrac, we could set the tracking algorithm to track particles
depending on size. Since some particles had low contrast and were hard to
track between frames, we used the multitracking ID-algorithm to have higher
stability in the trajectories [24]. Despite this, some trajectories were
broken. However, this did not affect the final results since we could merge
broken tracks and set the tracking algorithm to identify particles based on
their sizes when they exit the device. Using this approach and monitoring the
whole separation junction, we could identify the size of particles when they
exit either outlet A or B.

To calculate the rate of change of particles in the y-direction in the
separation region, we fitted a linear regression model to individual
trajectories. We multiplied this number with the pixel to µm conversion factor
(2.5 pixel/µm), defined by the sensor’s pixel size (8 µm) and microscope
magnification (x20).

###  4.6 SEM Imaging

We used SEM to image the silver conductive paint electrodes after different
treatments. To prevent charge buildup on the PDMS surface, we coated the
sample with a 5 nm thick layer of platinum using a sputter coater (Q150T-ES,
Quorum Technologies Ltd). We then imaged the electrodes using a scanning
electron microscope (Merlin FESEM, Carl Zeiss), with its InLens imaging mode
at a magnifications of 100-50000$\times$.

## Acknowledgements

This work was supported by the Swedish Research Council (2019-04016); the
Swedish Foundation for Strategic Research; the Umeå University Industrial
Doctoral School (IDS); Kempestiftelserna (JCK-1916.2); Swedish Department of
Defence, Project no. 470-A400821.

The authors acknowledge the facilities and technical assistance of the Umeå
Core Facility for Electron Microscopy (UCEM) at the Chemical Biological Centre
(KBC), Umeå University, a part of the National Microscopy Infrastructure NMI
(VR-RFI 2016-00968).

## Author contributions statement

M.A. and S.V. conceived the experiments. S.V. and D.N conducted the
dielectrophoresis experiments. D.M. did SEM imaging. D.N., S.V. and R.Ö.
manufactured the electrodes. S.V. and D.N created the models used in the
manuscript. D.N, D.M.and M.A. analysed the results. All authors reviewed and
edited the manuscript.

## Competing interests

The authors declare that they have no known competing financial interests or
personal relationships that could have appeared to influence the work reported
in this paper.

## Data availability statement

The datasets generated during the current study are available from the
corresponding author on reasonable request.

## References

  * [1] Valijam, S. & Salehi, A.  Influence of the obstacles on dielectrophoresis-assisted separation in microfluidic devices for cancerous cells.  _Journal of the Brazilian Society of Mechanical Sciences and Engineering_ 43, 1–14 (2021).
  * [2] Xu, L., Peng, J., Yan, M., Zhang, D. & Shen, A. Q.  Droplet synthesis of silver nanoparticles by a microfluidic device.  _Chemical Engineering and Processing: Process Intensification_ 102, 186–193 (2016).
  * [3] Santana, H., Silva, J., Aghel, B. & Ortega-Casanova, J.  Review on microfluidic device applications for fluids separation and water treatment processes.  _SN Applied Sciences_ 2, 1–19 (2020).
  * [4] Verpoorte, E.  Microfluidic chips for clinical and forensic analysis.  _Electrophoresis_ 23, 677–712 (2002).
  * [5] Al-Ali, A., Waheed, W., Abu-Nada, E. & Alazzam, A.  A review of active and passive hybrid systems based on dielectrophoresis for the manipulation of microparticles.  _Journal of Chromatography A_ 463268 (2022).
  * [6] Ghomian, T. & Hihath, J.  Review of dielectrophoretic manipulation of micro and nanomaterials: Fundamentals, recent developments, and challenges.  _IEEE Transactions on Biomedical Engineering_ (2022).
  * [7] Velmanickam, L., Jayasooriya, V., Vemuri, M. S., Tida, U. R. & Nawarathna, D.  Recent advances in dielectrophoresis toward biomarker detection: A summary of studies published between 2014 and 2021.  _Electrophoresis_ 43, 212–231 (2022).
  * [8] Rogers, J. A. & Nuzzo, R. G.  Recent progress in soft lithography.  _Materials today_ 8, 50–56 (2005).
  * [9] Ferrari, E., Nebuloni, F., Rasponi, M. & Occhetta, P.  Photo and soft lithography for organ-on-chip applications.  In _Organ-on-a-Chip_ , 1–19 (Springer, 2022).
  * [10] Nilsson, D. P. _et al._ Patient-specific brain arteries molded as a flexible phantom model using 3d printed water-soluble resin.  _Scientific reports_ 12, 1–9 (2022).
  * [11] Subirada, F. _et al._ Development of a custom-made 3d printing protocol with commercial resins for manufacturing microfluidic devices.  _Polymers_ 14, 2955 (2022).
  * [12] Cheon, J. & Kim, S.  Fabrication and demonstration of a 3d-printing/pdms integrated microfluidic device.  _Recent Progress in Materials_ 4, 1–1 (2022).
  * [13] Huddy, J. E. & Scheideler, W. J.  Protocol for deposition of conductive oxides onto 3D-printed materials for electronic device applications.  _STAR Protocols_ 3, 101523, DOI: 10.1016/j.xpro.2022.101523 (2022).
  * [14] Alazzam, A., Mathew, B. & Alhammadi, F.  Novel microfluidic device for the continuous separation of cancer cells using dielectrophoresis.  _Journal of separation science_ 40, 1193–1200 (2017).
  * [15] Hajari, M., Ebadi, A., Farshchi Heydari, M. J., Fathipour, M. & Soltani, M.  Dielectrophoresis-based microfluidic platform to sort micro-particles in continuous flow.  _Microsystem Technologies_ 26, 751–763 (2020).
  * [16] Li, Q. _et al._ Review of printed electrodes for flexible devices.  _Frontiers in Materials_ 5, 77 (2019).
  * [17] Dixon, C., Ng, A. H., Fobel, R., Miltenburg, M. B. & Wheeler, A. R.  An inkjet printed, roll-coated digital microfluidic device for inexpensive, miniaturized diagnostic assays.  _Lab on a Chip_ 16, 4560–4568 (2016).
  * [18] Lewpiriyawong, N., Yang, C. & Lam, Y. C.  Continuous sorting and separation of microparticles by size using ac dielectrophoresis in a pdms microfluidic device with 3-d conducting pdms composite electrodes.  _Electrophoresis_ 31, 2622–2631 (2010).
  * [19] Valijam, S., Salehi, A. & Andersson, M.  Design of a low-voltage dielectrophoresis lab-on-the chip to separate tumor and blood cells.  _Microfluidics and Nanofluidics_ 27 (2023).
  * [20] Matsuura, Y. _et al._ Thermodynamics of protein denaturation at temperatures over 100 c: Cuta1 mutant proteins substituted with hydrophobic and charged residues.  _Scientific reports_ 5, 1–9 (2015).
  * [21] Castellanos, A., Ramos, A., Gonzalez, A., Green, N. G. & Morgan, H.  Electrohydrodynamics and dielectrophoresis in microsystems: scaling laws.  _Journal of Physics D: Applied Physics_ 36, 2584 (2003).
  * [22] Ho, C. M. B., Ng, S. H., Li, K. H. H. & Yoon, Y.-J.  3d printed microfluidics for biological applications.  _Lab on a Chip_ 15, 3627–3637 (2015).
  * [23] Rodriguez, A. _et al._ Toxtrac: a fast and robust software for tracking organisms.  _Methods in Ecology and Evolution_ 9, 460–464 (2018).
  * [24] Rodriguez, A., Zhang, H., Klaminder, J., Brodin, T. & Andersson, M.  Toxid: an efficient algorithm to solve occlusions when tracking multiple animals.  _Scientific reports_ 7, 1–8 (2017).
  * [25] Grimnes, S. & Martinsen, Ø. G.  Electrodes.  In _Bioimpedance and Bioelectricity Basics_ , 179–254, DOI: 10.1016/B978-0-12-411470-8.00007-6 (Elsevier, 2015).
  * [26] Abd Rahman, N., Ibrahim, F. & Yafouz, B.  Dielectrophoresis for Biomedical Sciences Applications: A Review.  _Sensors_ 17, 449, DOI: 10.3390/s17030449 (2017).
  * [27] Salimi, E. & Bridges, G.  Dielectric Properties of Cells.  In _Comprehensive Biotechnology_ , 585–598, DOI: 10.1016/B978-0-444-64046-8.00061-6 (Elsevier, 2019).
  * [28] Khoshmanesh, K. _et al._ Size based separation of microparticles using a dielectrophoretic activated system.  _Journal of applied physics_ 108, 034904 (2010).
  * [29] Zhu, J. & Xuan, X.  Dielectrophoretic focusing of particles in a microchannel constriction using dc-biased ac flectric fields.  _Electrophoresis_ 30, 2668–2675 (2009).
  * [30] Jia, Y., Ren, Y. & Jiang, H.  Continuous dielectrophoretic particle separation using a microfluidic device with 3d electrodes and vaulted obstacles.  _Electrophoresis_ 36, 1744–1753 (2015).
  * [31] Nilsson, D. P. G. _et al._ Patient-specific brain arteries molded as a flexible phantom model using 3D printed water-soluble resin.  _Scientific Reports_ 12, 10172, DOI: 10.1038/s41598-022-14279-7 (2022).
  * [32] Zhang, H., Stangner, T., Wiklund, K., Rodriguez, A. & Andersson, M.  Umutracker: A versatile matlab program for automated particle tracking of 2d light microscopy or 3d digital holography data.  _Computer Physics Communications_ 219, 390–399 (2017).