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Red blood cell and white blood cell separation using a lateral-dimension scalable microchip based on hydraulic jump and sedimentation
Sensors & Actuators: B. Chemical 307 (2020) 127412
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Sensors and Actuators B: Chemical journal homepage: www.elsevier.com/locate/snb
Red blood cell and white blood cell separation using a lateral-dimension scalable microchip based on hydraulic jump and sedimentation
T
Hamidreza Shirinkami1, Gaobo Wang1, Jinhyeok Park, Joonhyang Ahn, Yeonho Choi*, Honggu Chun* Department of Biomedical Engineering, Korea University, Hana Science Hall, 145 Anamro, Seongbukgu, Seoul 02841, Republic of Korea
A R T I C LE I N FO
A B S T R A C T
Keywords: Lab-on-a-chip Point-of-care test Sedimentation Blood cells Separation
In most microfluidic techniques, channel width must be in range of few tens of micrometers for the device to work properly. With these dimensions, low-cost mass-production of microfluidic devices for use in point-of-care test is an issue. Here we present a hydraulic jump-based technique for size-selective microparticle or cell sorting in which working mechanism is not affected by channel width. Similar to converting kinetic energy to potential energy in macro scale, high-speed flow in a microfluidic channel is intentionally transformed into a flow with high potential energy by adjusting the channel height, inducing hydraulic jump. Depending on the strength of this energy transformation, particles with appropriate sizes can be trapped in a designed location as a result of sedimentation. With properly designing lengths of multiple expanded-height chambers and controlling the flow rate, it is possible to trap and release particles or cells selectively based on their size. To demonstrate application of our technique, we enriched 9.77 and 4.95 μm microbeads as well as WBCs and RBCs in separate chambers. Experimental results showed > 90 % selective enrichment of microbeads based on size. In addition, 69 % of WBCs and 80 % of RBCs were trapped in the first and second chamber, respectively.
1. Introduction Unique advantages of microfluidics has gained an immense interest among various disciplines including but not limited to biochemical analysis, clinical diagnostics and analytical chemistry [1,2]. Microfluidic devices have made the idea of point of care test (POCT) plausible by offering miniaturized, robust and sensitive analyses for diagnostic needs [3–6]. Benefited from small working volume and large surface to area, microfluidic devices allow complex procedures to be carried out in a time efficient manner. These devices are especially well-suited to deal with cells because at least one of the channel dimensions is comparable with the cell size. For this reason, cell sorting and characterization are attractive applications of microfluidic devices [7–10]. Various approaches for cell sorting and/or characterization have been applied. Microfluidic-based cell sorting methods requiring labelling of cells (for example, by fluorescent dye) generally depend on active systems, meaning that an external force (e.g. electrokinetic force, optical force, etc.) is applied once a cell is determined to be the target by analyzing its label. Although such methods offer high sensitivity, labelling requirement limits their application to the cases where pre-
treatment of sample is possible. Moreover, these methods often rely on real-time classification and rapid application of sorting mechanism thus demanding more elaborate equipment. On the other hand, label-free methods can be used in either active or passive strategies [11]. Examples of active label-free cell sorting mechanism are dielectrophoresis (DEP) [12], acoustophoresis [13] and magnetophoresis [14]. In passive methods, fluidic phenomena are harnessed to achieve the sorting usually based on inherent properties of specific cells such as size or deformability. Inertial microfluidic is one of the most notable passive techniques in separation of cells by careful tuning of inertial effects and Dean flow in curved channels [15,16]. In recent years, several examples of such label-free passive methods are presented in the literature [17–21]. One clear advantage of these techniques is that their operation only requires maintaining a certain flow rate inside the microchannel, which is in particularly attractive for POCT. However, one of the most important criteria for successful POCT applications is cost, especially in resource-limited settings [5]. In most microfluidic schemes, sorting force, whether it comes from an external source (e.g. magnetic field) or generated by a fluidic phenomenon (e.g. Dean flow), is acting in a horizontal plane parallel to the substrate. Consequently, the channel
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Corresponding authors. E-mail addresses: [email protected] (Y. Choi), [email protected] (H. Chun). 1 These authors contributed equally. https://doi.org/10.1016/j.snb.2019.127412 Received 26 July 2019; Received in revised form 3 November 2019; Accepted 12 November 2019 Available online 30 December 2019 0925-4005/ © 2019 Elsevier B.V. All rights reserved.
Sensors & Actuators: B. Chemical 307 (2020) 127412
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small, sedimentation velocity of a particle with diameter of dp and density of ρp in a fluid with density of ρf and viscosity of μ can be expressed as [35]:
width is needed to be not more than few tens of micrometers so that cell’s displacement under this force is meaningful compared to channel dimensions [2]. Additionally, the channel surface roughness is an important factor for a desirable fluidic phenomena. While this is not an issue when fabricating microchannels for research purpose using lithography process, mass production of chips with such small lateral dimensions may not be cost-effective for disposable POCT devices. Moreover, narrow channel structures are inherently prone to clogging, especially with whole blood or lipoaspirate samples. To address these concerns, a method with its working principle independent of channel width is preferable. An example of a wide-channel device which can be simply fabricated with plastic has been recently demonstrated by using acoustic force [22], however, passive methods are more suited for POCT devices owing to their simplicity and independency from external forces. The hydraulic jump phenomenon, caused by the principle of conservation of energy, is easily observed in nature, such as in fall, dam, and even sink. The core factor in this phenomenon is that kinetic energy (high velocity) is partially converted to potential energy (sudden height increase). Previously, Park etal. had theorized a technique based on hydraulic jump phenomenon to trap and release particles in an expansion chamber [23]. Later, Song etal. used the same principle to hypothesize a scheme for separating particles, however their method required implementation of several outlets at different heights to collect separated particles [24]; a requirement that demands very complicated fabrication process even in research level. Recently, few other works have been published with similar approaches yet only to obtain plasma by separating blood cells from whole blood samples [25,26]. Here we have advanced the hydraulic jump phenomena in a microchip by integrating two height-expansion chambers in order to introduce a labelfree, sedimentation-based particle/cell separation in which lateral dimension can be in millimeter range. We believe that chips with such wide lateral dimension are substantially cheaper to mass-produce for use in disposable POCT devices. When the channel width is not a concern, cost-effective fabrication methods such as laser-cutting [27], razor-cutting [28] or die cutting of polymer sheets, double-sided or transfer tapes can be used to mass-produce disposable devices suitable for POCT in resource-limited settings [29]. In such fabrication techniques, channel height is determined by the thickness of material used. A broad range of commercial products are available for different channel thickness. For instance, 3 M (MN, USA) offers transfer tapes for diagnostic microfluidic applications with thicknesses down to 25.4 ± 1.6 μm [30]. Ultra-thin double-sided tapes are available from Nitto Denko (Japan) to produce channel heights as low as 5 μm [31]. Lamination of dry-film resists (DFR) also offers a wide range of thickness down to 5 μm at low cost (< 2 USD for a chip size of 4 inch) [32,33]. Additionally, hot embossing enables low-cost mass-fabrication of low aspect-ratio channel microchip [34].
Vs=
(ρp − ρf ) 18μ
gdp2
(1)
In practice, particles are dispersed over the inlet channel height, but here for the sake of simplicity we assume that they are on average at the middle of channel height, thus sedimentation time to reach the floor is estimated by:
t S≈
h 2×VS
(2)
In the other hand, assuming that mean velocity in the inlet channel is V0, the average time it takes for particles to travel the whole length of expansion chamber, ℓ, is:
tT≈
l ×H V0
(3)
Now the criterion for a particle to reach the floor before escaping the chamber (i.e. tS < tT) is as follows:
2L >
V0 VS
(4)
where L is the length of chamber normalized by inlet channel’s height (i.e. L = ℓ/h0). Note that this simplified analysis only hints to the most important parameters that play a role in trapping of particles and cannot be regarded as a strict criterion. Interestingly, Eq. (4) shows that chamber’s height does not have a considerable effect on trapping, further simplifying requirements on microchannel fabrication with mass production for POCT applications. In fact, independency of trapping from chamber’s height has been previously demonstrated via numerical simulations in the work of Park etal., given that chamber’s height is not too close to that of the inlet channel, i.e. for H ≥ 5 condition [23]. Since trapping depends on sedimentation velocity and thus on particle size, size-selective particle trapping is possible. Once a particle reaches the surface, local flow velocity in its vicinity is much slower than the mean velocity of inlet channel due to the chamber height expansion as well as parabolic velocity profile of a laminar flow. Moreover, particle experiences an adhesion force dependent on particle size. Therefore, releasing of a trapped particle is possible only when the flow velocity is large enough to overcome the adhesion force [23]. The releasing efficiency can be improved by appropriate surface treatments preventing permanent adhesion of cells to the surface [36]. Here we designed a two-chamber configuration to separate microbeads as well as red blood cells (RBCs) and white blood cells (WBCs) by selectively trapping and releasing them in each chamber. Schematic of microchannel device is illustrated in Fig. 1b. Height of the inlet channel (h0) is chosen to be 12 μm, comparable to cell dimensions. If h0 is too low, then the inlet channel is easily clogged. On the other hand, if h0 is too high, cells follow the parabolic flow profile of the laminar flow, resulting in heterogeneous velocities and widespread sedimentation. Geometries of chambers are designed so that both sizes of cells are trapped simultaneously with a single inlet channel flow velocity, V0. The particles or cells trapped in the second chamber can be released and collected through the outlet channel by increasing flow velocity to VR2, and then, the particles or cells trapped in the first chamber can be released and collected through the outlet channel by further increasing flow velocity to VR1 (> VR2). Here, the width of microchannel was chosen to be 1 mm so that the whole width of each chamber can be imaged under a microscope with 4× objective lens, making particle counting easier. In real application, however, this width can be extended considerably in order to scale up the throughput.
2. Materials and methods 2.1. Microfluidic chip design Park etal. have theoretically shown the possibility of trapping and releasing microparticles by artificially introducing a hydraulic jump phenomenon with a sudden expansion of channel height [23]. Such expansion, shown in Fig. 1a, leads to reduction of flow velocity by a factor of height ratio (H = h/h0). The dominant force acting on particles in the inlet channel is drag force, causing particles to follow fluid streamlines. However, in the chamber (i.e. height expanded region) flow velocity drops and consequently gravity comes into play as well to result in sedimentation. In appropriate conditions settling particle will reach the channel floor before leaving the chamber thus regarded as trapped. A simplified analysis of the required condition to achieve efficient trapping is presented as follows. Assuming slow flow situation in which Reynolds number is very 2
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Fig. 1. Working principle of present separation technique (a) Simplified analysis of particle trapping mechanism in a chamber. When a particle is in the expanded chamber, the fluid velocity is small, thus gravity causes the particle to sediment. If the channel is long enough that the particle can reach floor before leaving the chamber, trapping can happen. (b) Selective trapping of blood cells in two chambers provides an opportunity for cell sorting. Chambers are designed so that at an inlet channel fluid velocity, V0, cells are trapped in their respective chambers. By increasing the fluid velocity, it is possible to sequentially release each type of cell and collect it in the outlet.
2.2. Simulation
2.3. Chip fabrication
To find proper conditions for trapping of microbeads and blood cells, we performed numerical simulation using COMSOL Multiphysics 5.3 (COMSOL Inc, MA, USA). The simulation domain, shown in Figure S1a of ESI, was consisted of lateral cross-section of an expansion chamber and inlet/outlet channels. In our device design, width of chambers was chosen to be 1 mm, which is much larger than channel heights therefore the majority of flow is not affected by side walls, and therefore 2D assumption is reasonable. We conducted the simulation with variation in chamber dimensions and flow velocity to see the effect on trapping efficiencies of particles with difference sizes. Simulation of particle trapping consisted of two steps. First, flow velocity field was obtained by solving Navier-Stokes equation with the assumption of Stokes flow. Then, particles were injected into the inlet channel and previously-solved velocity field was used to trace them with drag and gravity forces enabled. In the chamber region, a particle considered to be trapped whenever the distance between the particle center and chamber’s floor became equal to particle’s radius. This assumption simplifies simulation but ignores the fact that if local flow velocity is large (e.g. close to entrance or exit of chamber, or when H is small) a particle settled on the surface can still role over it and exit the chamber. Thus, simulation results must be considered with caution, in that the simulation estimates settling efficiency rather than actual trapping efficiency which is also dependent on particle-surface interaction. However, it provides a valuable insight before conducting chip fabrication and experiments. To asses trapping efficiency in each simulation trial, we injected 100 particles uniformly distributed over the cross section (i.e. the height) of inlet channel (Figure S1b). We defined numerical trapping efficiency as the ratio of particles trapped in the chamber to the total number of injected particles.
A SU-8 mold and PDMS chip were fabricated based on a previous protocol [37,38]. A multilayer stacked SU-8 mold was fabricated using SU-8 2025 and SU-8 2075 negative photoresists (MicroChem Corp, MA, USA) for the inlet/outlet channels and chambers respectively. First, SU8 2025 was spin-coated on a silicon wafer with target thickness of 12 μm. The microchannel pattern was exposed on the substrate with a mask and UV-LED exposure system (MU-60, Cella Biotech, Korea), then SU-8 2075 was spin-coated over the SU-8 2025 coated substrate with thickness of 110 μm and exposed to UV with the other mask to define chambers. 10:1 mixture of PDMS pre-polymer and its curing agent (Sylgard 184, Dow Corning, MI, USA) was poured onto the master mold, cured at 80 °C in an oven for 45 min. The PDMS part was then bonded to a substrate (which was either bare or PDMS-coated slide glass, depending on the surface treatment type) by oxygen plasma treatment and kept on a 120 °C hotplate for 30 min to strengthen the bond. An example of actual microchannel design and a picture of a fabricated chip is presented in Figure S2 of ESI. 2.4. Surface treatment To facilitate releasing of trapped particles and cells, we compared two different bottom layer surfaces: a) using bare slide glass as the substrate and treating it with NaOH after the channel bonding and b) coating slide glass with a thin layer of PDMS before the channel bonding and then treating the bonded channel with aqueous solution of Pluronic F-68 (Thermo Fisher Scientific, MA, USA). For the thin layer PDMS coating, the PDMS precursor was 9 times diluted by Chloroform (Sigma-Aldrich, MO, USA), and then spin-coated on a slide glass at 1200 RPM and cured at 65 °C for 3 h. For each method, we have performed surface treatment at various concentrations and compared releasing efficiency as well as size-selectivity. 3
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should be minimized. Therefore, these results must be considered together with trapping efficiency of 4.95 μm particles, shown in Fig. 2b. Additionally, this graph is used to design the second chamber for 4.95 μm particle trapping. Consequently, we chose 2 mm as the length of the first chamber for trapping WBCs. Due to computation limit, trapping efficiency was calculated for chamber lengths up to 7 mm. For 9.77 μm particles, simulation of longer channels is not even needed since with 3 mm chamber perfect trapping is observed over the whole flow velocity range. In case of 4.95 μm particles, simulation results at V0 = 400 μm s−1 suggests that 7 mm chamber is enough to achieve complete trapping. Therefore, we decided to fabricate chips with up to 7 mm long chambers for trapping RBCs after the first chamber of 2 mm. The inlet channel height (h0 = 12 μm) is the smallest dimension of the chip. A low-cost fabrication method may result in deviation in the channel height. To investigate the effect of channel height, simulations were performed to estimate trapping efficiencies of 9.77 and 4.95 μm microparticles with different inlet channel heights (h0 = 10, 12, and 14 μm). Simulation result shows that the inlet channel height has effect on the trapping efficiency at high flow rates, however, it is negligible at the experimental condition of V0 = 400 μm s−1 (Figure S5 in ESI).
2.5. Microparticle and blood sample preparation Prior to cell sorting, we carried out experiments on carboxylate modified fluorescent polystyrene beads (Bangs lab, IN, USA) of 4.95 μm and 9.77 μm diameters as the model particle for RBC and WBC, respectively. Under fluorescent microscopy, smaller beads are observed as blue (excitation 360 nm, emission 450 nm) and larger particles as green (excitation 480 nm, emission 520 nm). The beads were dispersed in PBS prior to use. The blood sample was supplied from Korea University Guro Hospital. Ten healthy males (age: 29 ± 9 years; height: 178 ± 4 cm; bodyweight: 85 ± 11 kg; BMI: 26.8 ± 3.7) between the ages of 18 and 50 years volunteered to participate in this study. This study was approved by the ethics committee of Korea University Guro Hospital, and written informed consent was obtained from all participates in accordance with the Declaration of Helsinki (IRB no. KUGH13093-001). For cell sorting experiments, 1 μL whole blood sample was diluted 100 times with 1× Dulbecco PBS buffer (Sigma-Aldrich, MO, USA) to maintain normal osmotic pressure. The WBCs were stained with DAPI (4′,6-diamidino-2-phenylindole, Sigma-Aldrich, MO, USA) following the supplier’s protocol.
3.2. Trapping of microbeads 2.6. Experimental procedure To assess trapping efficiency under several combinations of chamber length and mean velocity, we performed experiments using fluorescent polystyrene microbeads (Fig. 3a). At slow velocities, particles sediment near the chamber entrance. By increasing velocity, particles gain the chance to escape from the chamber before being trapped. In addition, Fig. 3a hints to a correlation between mean velocity and particle spread over the length of channel. Based on this observation, V0 = 200 μm s−1 is too low since it can lead to aggregation of microbeads in short range. Moreover, bead adhesion to the inlet channel was observed at low velocities, even increasing the chance of channel clogging. The number of beads entering chamber and being trapped were counted to calculate trapping efficiencies presented in Fig. 3b–c for 9.77 μm and 4.95 μm beads. By comparing these two graphs we concluded V0 = 400 μm s−1 and ℓ1 = 2 mm and ℓ2 = 7 mm as the proper condition for separating the two sizes of particles. In such combination of parameters, about 90 % of WBCs are expected to be trapped in the first chamber with only about 20 % of RBCs being trapped in the same chamber. In the second chamber, over 95 % of RBCs entering it are expected to be trapped on the surface before reaching the exit.
Sample solution was injected into the inlet channel for 3–5 min to analyze the trapping efficiency at designated flow velocities controlled by a syringe pump (Harvard Apparatus, MA, USA). Then the releasing efficiency was analyzed with increased flow velocities. The fluorescent image of microbeads and WBCs was acquired by a fluorescence microscope (Ti-U, Nikon, Japan) equipped with a CMOS image sensor (DP74, Olympus, Japan). RBCs were analyzed under bright-field condition. Both images were taken with ×4 objective lens. The trapping and releasing efficiencies were estimated by automatically counting the particle or cell number using ImageJ open-source software (National Institutes of Health, MD, USA). If a chamber had overlapped particles or cells, manual counting was performed with higher magnification (×20) objective lens. Representative images of microbead and cell counting are shown in Figure S3 of ESI. Experimental trapping efficiency was defined as the number of particles trapped in a chamber divided by the number of particles entered the chamber during the trapping phase. Similarly, releasing efficiency was defined as the number of cells escaped from the chamber after increasing the inlet channel’s flow velocity over the number of trapped cells in the chamber before increasing the flow velocity.
3.3. Releasing of microbeads
3. Results and discussion
Once particles are trapped in chambers, adhesion force temporarily immobilize them on the substrate, thus releasing requires increased flow velocity until adhesion force is overcome [23]. However, the flow velocity near the trapped particle is low due to the parabolic flow profile of a pressure driven flow. For this reason, adhesion force must be decreased to prevent unrealistically high releasing flow velocity. Surface treatment by poly(ethylene oxide) (PEO) is one of the widelyknown methods of inhibiting cell and protein adhesion in microchannels [36]. Here we tried surface treatment with Pluronic F-68, a non-ionic surfactant based on PEO, with a PDMS-coated glass substrate. Additionally, we investigated effect of 0.1, 1, and 5 N NaOH treatment with bare glass as the substrate. Applying NaOH increases negative charges (-O−) on glass surface helping to repel negatively-charged cells and particles. We compared three different concentrations of each surface treatment solution in terms of releasing efficiency of each particle size. Additionally, a control experiment without any surface treatment was conducted. Results presented in Fig. 4a suggest that Pluronic F-68 is more effective than NaOH treatment in decreasing adhesion and therefore improving releasing efficiency, especially for smaller particles. The highest releasing efficiencies were achieved with 3 % v/v concentration of Pluronic F-68, therefore it was chosen for the
3.1. Microparticle trap-and-release simulation Following the simulation procedure described previously, trapping efficiencies of 4.95 and 9.77 μm particles were estimated for a combination of channel lengths (ℓ) and mean flow velocities (V0). Note that as it was shown by Park [23] and also through our simplified analysis, H does not have a significant effect on trapping efficiency if it is large enough so that the expanded region can be reasonably called a chamber. This conclusion was also confirmed by our simulations, showing that for H ≥ 5, trapping efficiency is not a function of H (Figure S4 of ESI). As previously discussed, this simulation is not suitable when H is small, thus we did not perform it for H < 5. Fig. 2a demonstrates trapping efficiency for 9.77 μm particles versus flow velocity. As seen from the graph, when chamber is longer than 2 mm, more than 90 % of particles are trapped even at higher flow velocities. However, for 1 mm chamber there is a decreasing trend when V0 exceeds 400 μm s−1. This is because at this velocity particles are trapped close to the exit of chamber. When designing conditions for the first chamber to trap 9.77 μm particles, trapping of smaller particles 4
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Fig. 2. Trapping efficiencies vs. mean velocity in the inlet channel obtained by simulation for (a) 9.77 μm particles and (b) 4.95 μm particles. In all the cases, H = 10.
3.4. Separation of blood cells
remaining experiments. Releasing efficiency of trapped particles was investigated with a variation of inlet channel flow velocities (Fig. 4b). More than 90 % of 4.95 μm beads are released by increasing flow velocity to 2500 μm s−1. However, at this velocity less than 10 % of 9.77 μm bead leave the trapping chamber. The 9.77 μm beads can be efficiently (> 80 %) released when the inlet channel flow velocity increased to 12,500 μm s−1. The different releasing flow velocities based on particle/cell size enable us to perform size-selective particle/cell sorting or enrichment.
Through simulation and microbead experiment, we finalized chamber length (2 and 7 mm for the first and second chamber, respectively) and height (H = 10 for both chambers) as well as the inlet channel mean flow velocity (V0 = 400 μm s−1 for trapping. V0 = 2500 μm s−1 and V0 = 12,500 μm s−1 for RBC and WBC releasing, respectively) for the selective WBC and RBC trapping and releasing. A 100× diluted whole blood in PBS was injected into the chip to achieve mean flow velocity of V0 = 400 μm s−1 in the inlet channel. With inlet
Fig. 3. Experimental result for trapping of fluorescent polystyrene beads. (a) Microbeads trapped in a 2 mm chamber at three mean velocities. The chamber is marked by the white dashed line. 9.77 and 4.95 μm beads have green and blue fluorescent color, respectively. Experimental trapping efficiency of (b) 9.77 μm and (c) 4.95 μm microbeads and vs. chamber length, obtained at various mean velocities (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article). 5
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Fig. 4. Releasing of microbeads (a) Comparison of releasing efficiency for each size of microbeads under various surface treatments. Releasing velocity was 12,500 μm s−1 for 9.77 μm beads and 2500 μm s−1 for 4.95 μm beads. In case of Pluronic F-68, substrate was coated by a thin PDMS layer. (b) Releasing efficiency as a function of inlet channel flow velocity for two sizes of microbeads with 3 % Pluronic F-68 coating.
velocity (12,500 μm s−1), the abundant RBCs in the first chamber can be selectively released by increasing the inlet flow velocity to the RBC releasing flow velocity, increasing the WBC purity and number in the first chamber. In this work, we verified the trapping and releasing efficiency of the proposed cell sorting or enrichment technique with a channel width of 1 mm. The resulting throughput of 0.288 μL min−1 cannot compete with inertial microfluidics or deterministic lateral displacement (DLD) which operates at high flowrate up to 10 mL min−1 [2]. However, the throughput of the presented device can be simply upscaled by increasing the channel width. In addition, the maximum inlet flow velocity V0 showing 100 % trapping efficiency linearly increases with the chamber length, as shown in Eq. (4). A detailed simulation result showing the linear relationship between the chamber length and throughput is presented in Figure S6 of ESI. Therefore, the throughput increases with the chamber width and length. For example, a device with chip size of 2 × 3 inch slideglass (45-times wider and 6-times longer chamber) can yield same trapping efficiency at 270-times higher flow rate (∼ 78 μL min−1). Moreover, in applications isolating larger cells such as circulating tumor cells (CTCs), not only target cells are heavier and sediment faster, but also one single chamber is enough to achieve the separation goal, which allow even longer chamber and higher flowrate. When the flowrate is high, the trapping and releasing sequences should be repeated before too many cells are trapped and overlapped in a chamber.
channel having cross-section of 12 μm × 1 mm, this flow velocity was achieved by injecting sample at 0.288 μL min−1. Trapping phase was completed after injecting 1.5 μL of sample, and flow stopped to assess trapping efficiency of cells in each chamber. With the pseudo-continuous nature of our technique, it was necessary to stop flow before too many cells are trapped in a chamber. As seen in Fig. 5a, 69 % of WBCs and 80 % of RBCs are trapped in the first and second chamber, respectively. It must be noted that the total number of cells trapped in both chambers is less than the injected cell number which was estimated from injection volume and bulk sample cell concentration. The number of lost cells which escaped both chambers was negligible. Most of the discrepancy came from adhered cells on the connecting tube wall. Therefore, the summed number of trapped cells in both chambers can be considered as actual number of injected cells into the first chamber. After completing trapping and counting cells in each chamber, releasing phase was performed. More than 85 % of RBCs trapped in the second chamber were released by increasing inlet channel mean flow velocity to 2500 μm s−1 as shown in Fig. 5b. In case of WBCs trapped in the first chamber, we were able to release more than 80 % by increasing the inlet channel mean flow velocity to 12,500 μm s−1. Fig. 5a also shows the degree of cross-contamination demonstrated as the number of trapped RBCs and WBCs in the first and second chamber, respectively. It is worth mentioning that in terms of trapped cell number in the first chamber, originally designed to trap WBCs, is dominated by RBCs. This happens because sample (i.e. diluted whole blood) is predominantly occupied by RBCs (×1000 more abundant in blood). Therefore, experiment on blood sample must be interpreted as enrichment of WBCs rather than separation purity. In addition, the WBC enrichment ratio or purity can be improved by repeating the sample injection and RBC release process. Because the RBC releasing flow velocity (2500 μm s−1) is much slower than the WBC releasing flow
4. Conclusions In this work, we have introduced a technique to trap and release cells in a size-selective manner by utilizing internal hydraulic jump phenomenon. We numerically investigated trapping and releasing efficiencies with parameters including chamber length, height, and flow 6
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Fig. 5. Trapping and releasing of blood cells (a) Number of WBCs and RBCs during trapping phase. V0 = 400 μm s−1. The number of injected cells is estimated from the volume of injected blood and its concentration. (b) Number of WBCs and RBCs in each chamber before and after releasing phase. V0 = 12,500 and 2500 μm s−1 for the WBC and RBC releasing in the first and second chamber, respectively.
(S7. Low-cost chip fabrication with machining in ESI) and tape cutting (S8. Low-cost chip fabrication with tape in ESI). Experimental results showed > 99 % of trapping efficiencies of 9.77 μm microbeads for both of fabrication methods, which suggest the possibility of low-cost massproduction of disposable cell sorter for POCT. Finally, the scheme presented here can be further developed for other applications. For instance, micro-valves can be implemented into the microchannel to enable collection of released cells without additional cross-contamination. In another example, substrate can be coated by adhesion promoters to permanently immobilize trapped cells, then use the device for on-chip culture and detection of large-size stem cell or CTCs.
velocity, and then validated the result with microbead experiments. We also demonstrated an application of our technique by separating WBCs and RBCs through enriching them in different chambers. Most microfluidic cell sorting methods, whether passive (e.g. inertial microfluidic) or active (e.g. acoustophoresis), do not only require the lateral dimension of microchannel to be in same order as cell size but also very sharply defined microchannel surface. Even though such devices are shown to have excellent performances in terms of separation purity and/or throughput [2,39–42], the μm range resolution of the channel design places concerns on low-cost mass production for POCT devices as well as channel clogging with real biological samples. In contrast, working principle of our technique is independent of microchannel’s lateral dimension, not only making mm-range channels possible but also providing a way to scale up the throughput by simply increasing channel width. Moreover, the trapping and releasing efficiency is robust with the channel height variation, further simplifying the restriction on fabrication resolution. Additionally, wider channel means less chance of clogging by cells or contaminations, and this leads to a more versatile approach for use in commercial devices and with untrained users. In this work, we have used standard soft lithography to fabricate the chip with exact dimensions for quantitative analysis of experimental results. However, a SU-8 mold fabrication with conventional photolithography is not a low-cost solution. For this reason, we also fabricated the chip with low-cost methods including machining
Conflict of interest The authors declares no conflict of interest.
Acknowledgement This research was supported by the National Research Foundation of Korea (NRF) (NRF-2011-0031866 and NRF-2018M3A9D7079485).
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Appendix A. Supplementary data
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Hamidreza Shirinkami received his BS degree in Mechanical Engineering from University of Tehran, Tehran, Iran. In 2014 he joined the Department of Biomedical Engineering at Korea University, Seoul, Korea for his PhD studies. His research interest is focused on Microfluidics, Neural interface, and Medical devices. Gaobo Wang received his BS degree in Chemistry and Chemical Engineering from Henan Institute of Science and Technology, Henan, China. In 2018, he received his Master’s degree in Biomedical Engineering from Korea University, Seoul, Korea. His research interest is focused on Microfluidics. Jinhyeok Park received his BS degree in Biomedical Engineering from Korea University, Seoul, Korea. In 2017, he received his Master’s degree in Biomedical Engineering from Korea University, Seoul, Korea. His research interest is focused on micro/nanofabrication and exosome analysis. Joonhyang Ahn received his BS degree in Biomedical Engineering from Korea University, Seoul, Korea. His research interest is focused on microfluidics. Yeonho Choi received his BS degree in Mechanical Engineering from Seoul National University, Seoul, Korea. In 2009, he received his PhD degree in Mechanical Engineering from the University of California at Berkeley, CA, USA. Since 2010, he has joined the Department of Biomedical Engineering at Korea University, Seoul, Korea. His research interest is focused on Plasmonics, Biomimetics, Exosome Analysis, and Deep-learning based diagnostics. Honggu Chun received his BS degree in Electrical Engineering and Computer Science from Seoul National University, Seoul, Korea. In 2004, he received his PhD degree in Biomedical Engineering from Seoul National University, Seoul, Korea. In 2011, he joined the Department of Biomedical Engineering at Korea University, Seoul, Korea. His research interest is focused on Iontronics, Neuro-interface, Clinical diagnostics, Exosome analysis, Single cell analysis, and Medical instrument.
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Sensors and Actuators B: Chemical journal homepage: www.elsevier.com/locate/snb
Red blood cell and white blood cell separation using a lateral-dimension scalable microchip based on hydraulic jump and sedimentation
T
Hamidreza Shirinkami1, Gaobo Wang1, Jinhyeok Park, Joonhyang Ahn, Yeonho Choi*, Honggu Chun* Department of Biomedical Engineering, Korea University, Hana Science Hall, 145 Anamro, Seongbukgu, Seoul 02841, Republic of Korea
A R T I C LE I N FO
A B S T R A C T
Keywords: Lab-on-a-chip Point-of-care test Sedimentation Blood cells Separation
In most microfluidic techniques, channel width must be in range of few tens of micrometers for the device to work properly. With these dimensions, low-cost mass-production of microfluidic devices for use in point-of-care test is an issue. Here we present a hydraulic jump-based technique for size-selective microparticle or cell sorting in which working mechanism is not affected by channel width. Similar to converting kinetic energy to potential energy in macro scale, high-speed flow in a microfluidic channel is intentionally transformed into a flow with high potential energy by adjusting the channel height, inducing hydraulic jump. Depending on the strength of this energy transformation, particles with appropriate sizes can be trapped in a designed location as a result of sedimentation. With properly designing lengths of multiple expanded-height chambers and controlling the flow rate, it is possible to trap and release particles or cells selectively based on their size. To demonstrate application of our technique, we enriched 9.77 and 4.95 μm microbeads as well as WBCs and RBCs in separate chambers. Experimental results showed > 90 % selective enrichment of microbeads based on size. In addition, 69 % of WBCs and 80 % of RBCs were trapped in the first and second chamber, respectively.
1. Introduction Unique advantages of microfluidics has gained an immense interest among various disciplines including but not limited to biochemical analysis, clinical diagnostics and analytical chemistry [1,2]. Microfluidic devices have made the idea of point of care test (POCT) plausible by offering miniaturized, robust and sensitive analyses for diagnostic needs [3–6]. Benefited from small working volume and large surface to area, microfluidic devices allow complex procedures to be carried out in a time efficient manner. These devices are especially well-suited to deal with cells because at least one of the channel dimensions is comparable with the cell size. For this reason, cell sorting and characterization are attractive applications of microfluidic devices [7–10]. Various approaches for cell sorting and/or characterization have been applied. Microfluidic-based cell sorting methods requiring labelling of cells (for example, by fluorescent dye) generally depend on active systems, meaning that an external force (e.g. electrokinetic force, optical force, etc.) is applied once a cell is determined to be the target by analyzing its label. Although such methods offer high sensitivity, labelling requirement limits their application to the cases where pre-
treatment of sample is possible. Moreover, these methods often rely on real-time classification and rapid application of sorting mechanism thus demanding more elaborate equipment. On the other hand, label-free methods can be used in either active or passive strategies [11]. Examples of active label-free cell sorting mechanism are dielectrophoresis (DEP) [12], acoustophoresis [13] and magnetophoresis [14]. In passive methods, fluidic phenomena are harnessed to achieve the sorting usually based on inherent properties of specific cells such as size or deformability. Inertial microfluidic is one of the most notable passive techniques in separation of cells by careful tuning of inertial effects and Dean flow in curved channels [15,16]. In recent years, several examples of such label-free passive methods are presented in the literature [17–21]. One clear advantage of these techniques is that their operation only requires maintaining a certain flow rate inside the microchannel, which is in particularly attractive for POCT. However, one of the most important criteria for successful POCT applications is cost, especially in resource-limited settings [5]. In most microfluidic schemes, sorting force, whether it comes from an external source (e.g. magnetic field) or generated by a fluidic phenomenon (e.g. Dean flow), is acting in a horizontal plane parallel to the substrate. Consequently, the channel
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Corresponding authors. E-mail addresses: [email protected] (Y. Choi), [email protected] (H. Chun). 1 These authors contributed equally. https://doi.org/10.1016/j.snb.2019.127412 Received 26 July 2019; Received in revised form 3 November 2019; Accepted 12 November 2019 Available online 30 December 2019 0925-4005/ © 2019 Elsevier B.V. All rights reserved.
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small, sedimentation velocity of a particle with diameter of dp and density of ρp in a fluid with density of ρf and viscosity of μ can be expressed as [35]:
width is needed to be not more than few tens of micrometers so that cell’s displacement under this force is meaningful compared to channel dimensions [2]. Additionally, the channel surface roughness is an important factor for a desirable fluidic phenomena. While this is not an issue when fabricating microchannels for research purpose using lithography process, mass production of chips with such small lateral dimensions may not be cost-effective for disposable POCT devices. Moreover, narrow channel structures are inherently prone to clogging, especially with whole blood or lipoaspirate samples. To address these concerns, a method with its working principle independent of channel width is preferable. An example of a wide-channel device which can be simply fabricated with plastic has been recently demonstrated by using acoustic force [22], however, passive methods are more suited for POCT devices owing to their simplicity and independency from external forces. The hydraulic jump phenomenon, caused by the principle of conservation of energy, is easily observed in nature, such as in fall, dam, and even sink. The core factor in this phenomenon is that kinetic energy (high velocity) is partially converted to potential energy (sudden height increase). Previously, Park etal. had theorized a technique based on hydraulic jump phenomenon to trap and release particles in an expansion chamber [23]. Later, Song etal. used the same principle to hypothesize a scheme for separating particles, however their method required implementation of several outlets at different heights to collect separated particles [24]; a requirement that demands very complicated fabrication process even in research level. Recently, few other works have been published with similar approaches yet only to obtain plasma by separating blood cells from whole blood samples [25,26]. Here we have advanced the hydraulic jump phenomena in a microchip by integrating two height-expansion chambers in order to introduce a labelfree, sedimentation-based particle/cell separation in which lateral dimension can be in millimeter range. We believe that chips with such wide lateral dimension are substantially cheaper to mass-produce for use in disposable POCT devices. When the channel width is not a concern, cost-effective fabrication methods such as laser-cutting [27], razor-cutting [28] or die cutting of polymer sheets, double-sided or transfer tapes can be used to mass-produce disposable devices suitable for POCT in resource-limited settings [29]. In such fabrication techniques, channel height is determined by the thickness of material used. A broad range of commercial products are available for different channel thickness. For instance, 3 M (MN, USA) offers transfer tapes for diagnostic microfluidic applications with thicknesses down to 25.4 ± 1.6 μm [30]. Ultra-thin double-sided tapes are available from Nitto Denko (Japan) to produce channel heights as low as 5 μm [31]. Lamination of dry-film resists (DFR) also offers a wide range of thickness down to 5 μm at low cost (< 2 USD for a chip size of 4 inch) [32,33]. Additionally, hot embossing enables low-cost mass-fabrication of low aspect-ratio channel microchip [34].
Vs=
(ρp − ρf ) 18μ
gdp2
(1)
In practice, particles are dispersed over the inlet channel height, but here for the sake of simplicity we assume that they are on average at the middle of channel height, thus sedimentation time to reach the floor is estimated by:
t S≈
h 2×VS
(2)
In the other hand, assuming that mean velocity in the inlet channel is V0, the average time it takes for particles to travel the whole length of expansion chamber, ℓ, is:
tT≈
l ×H V0
(3)
Now the criterion for a particle to reach the floor before escaping the chamber (i.e. tS < tT) is as follows:
2L >
V0 VS
(4)
where L is the length of chamber normalized by inlet channel’s height (i.e. L = ℓ/h0). Note that this simplified analysis only hints to the most important parameters that play a role in trapping of particles and cannot be regarded as a strict criterion. Interestingly, Eq. (4) shows that chamber’s height does not have a considerable effect on trapping, further simplifying requirements on microchannel fabrication with mass production for POCT applications. In fact, independency of trapping from chamber’s height has been previously demonstrated via numerical simulations in the work of Park etal., given that chamber’s height is not too close to that of the inlet channel, i.e. for H ≥ 5 condition [23]. Since trapping depends on sedimentation velocity and thus on particle size, size-selective particle trapping is possible. Once a particle reaches the surface, local flow velocity in its vicinity is much slower than the mean velocity of inlet channel due to the chamber height expansion as well as parabolic velocity profile of a laminar flow. Moreover, particle experiences an adhesion force dependent on particle size. Therefore, releasing of a trapped particle is possible only when the flow velocity is large enough to overcome the adhesion force [23]. The releasing efficiency can be improved by appropriate surface treatments preventing permanent adhesion of cells to the surface [36]. Here we designed a two-chamber configuration to separate microbeads as well as red blood cells (RBCs) and white blood cells (WBCs) by selectively trapping and releasing them in each chamber. Schematic of microchannel device is illustrated in Fig. 1b. Height of the inlet channel (h0) is chosen to be 12 μm, comparable to cell dimensions. If h0 is too low, then the inlet channel is easily clogged. On the other hand, if h0 is too high, cells follow the parabolic flow profile of the laminar flow, resulting in heterogeneous velocities and widespread sedimentation. Geometries of chambers are designed so that both sizes of cells are trapped simultaneously with a single inlet channel flow velocity, V0. The particles or cells trapped in the second chamber can be released and collected through the outlet channel by increasing flow velocity to VR2, and then, the particles or cells trapped in the first chamber can be released and collected through the outlet channel by further increasing flow velocity to VR1 (> VR2). Here, the width of microchannel was chosen to be 1 mm so that the whole width of each chamber can be imaged under a microscope with 4× objective lens, making particle counting easier. In real application, however, this width can be extended considerably in order to scale up the throughput.
2. Materials and methods 2.1. Microfluidic chip design Park etal. have theoretically shown the possibility of trapping and releasing microparticles by artificially introducing a hydraulic jump phenomenon with a sudden expansion of channel height [23]. Such expansion, shown in Fig. 1a, leads to reduction of flow velocity by a factor of height ratio (H = h/h0). The dominant force acting on particles in the inlet channel is drag force, causing particles to follow fluid streamlines. However, in the chamber (i.e. height expanded region) flow velocity drops and consequently gravity comes into play as well to result in sedimentation. In appropriate conditions settling particle will reach the channel floor before leaving the chamber thus regarded as trapped. A simplified analysis of the required condition to achieve efficient trapping is presented as follows. Assuming slow flow situation in which Reynolds number is very 2
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Fig. 1. Working principle of present separation technique (a) Simplified analysis of particle trapping mechanism in a chamber. When a particle is in the expanded chamber, the fluid velocity is small, thus gravity causes the particle to sediment. If the channel is long enough that the particle can reach floor before leaving the chamber, trapping can happen. (b) Selective trapping of blood cells in two chambers provides an opportunity for cell sorting. Chambers are designed so that at an inlet channel fluid velocity, V0, cells are trapped in their respective chambers. By increasing the fluid velocity, it is possible to sequentially release each type of cell and collect it in the outlet.
2.2. Simulation
2.3. Chip fabrication
To find proper conditions for trapping of microbeads and blood cells, we performed numerical simulation using COMSOL Multiphysics 5.3 (COMSOL Inc, MA, USA). The simulation domain, shown in Figure S1a of ESI, was consisted of lateral cross-section of an expansion chamber and inlet/outlet channels. In our device design, width of chambers was chosen to be 1 mm, which is much larger than channel heights therefore the majority of flow is not affected by side walls, and therefore 2D assumption is reasonable. We conducted the simulation with variation in chamber dimensions and flow velocity to see the effect on trapping efficiencies of particles with difference sizes. Simulation of particle trapping consisted of two steps. First, flow velocity field was obtained by solving Navier-Stokes equation with the assumption of Stokes flow. Then, particles were injected into the inlet channel and previously-solved velocity field was used to trace them with drag and gravity forces enabled. In the chamber region, a particle considered to be trapped whenever the distance between the particle center and chamber’s floor became equal to particle’s radius. This assumption simplifies simulation but ignores the fact that if local flow velocity is large (e.g. close to entrance or exit of chamber, or when H is small) a particle settled on the surface can still role over it and exit the chamber. Thus, simulation results must be considered with caution, in that the simulation estimates settling efficiency rather than actual trapping efficiency which is also dependent on particle-surface interaction. However, it provides a valuable insight before conducting chip fabrication and experiments. To asses trapping efficiency in each simulation trial, we injected 100 particles uniformly distributed over the cross section (i.e. the height) of inlet channel (Figure S1b). We defined numerical trapping efficiency as the ratio of particles trapped in the chamber to the total number of injected particles.
A SU-8 mold and PDMS chip were fabricated based on a previous protocol [37,38]. A multilayer stacked SU-8 mold was fabricated using SU-8 2025 and SU-8 2075 negative photoresists (MicroChem Corp, MA, USA) for the inlet/outlet channels and chambers respectively. First, SU8 2025 was spin-coated on a silicon wafer with target thickness of 12 μm. The microchannel pattern was exposed on the substrate with a mask and UV-LED exposure system (MU-60, Cella Biotech, Korea), then SU-8 2075 was spin-coated over the SU-8 2025 coated substrate with thickness of 110 μm and exposed to UV with the other mask to define chambers. 10:1 mixture of PDMS pre-polymer and its curing agent (Sylgard 184, Dow Corning, MI, USA) was poured onto the master mold, cured at 80 °C in an oven for 45 min. The PDMS part was then bonded to a substrate (which was either bare or PDMS-coated slide glass, depending on the surface treatment type) by oxygen plasma treatment and kept on a 120 °C hotplate for 30 min to strengthen the bond. An example of actual microchannel design and a picture of a fabricated chip is presented in Figure S2 of ESI. 2.4. Surface treatment To facilitate releasing of trapped particles and cells, we compared two different bottom layer surfaces: a) using bare slide glass as the substrate and treating it with NaOH after the channel bonding and b) coating slide glass with a thin layer of PDMS before the channel bonding and then treating the bonded channel with aqueous solution of Pluronic F-68 (Thermo Fisher Scientific, MA, USA). For the thin layer PDMS coating, the PDMS precursor was 9 times diluted by Chloroform (Sigma-Aldrich, MO, USA), and then spin-coated on a slide glass at 1200 RPM and cured at 65 °C for 3 h. For each method, we have performed surface treatment at various concentrations and compared releasing efficiency as well as size-selectivity. 3
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should be minimized. Therefore, these results must be considered together with trapping efficiency of 4.95 μm particles, shown in Fig. 2b. Additionally, this graph is used to design the second chamber for 4.95 μm particle trapping. Consequently, we chose 2 mm as the length of the first chamber for trapping WBCs. Due to computation limit, trapping efficiency was calculated for chamber lengths up to 7 mm. For 9.77 μm particles, simulation of longer channels is not even needed since with 3 mm chamber perfect trapping is observed over the whole flow velocity range. In case of 4.95 μm particles, simulation results at V0 = 400 μm s−1 suggests that 7 mm chamber is enough to achieve complete trapping. Therefore, we decided to fabricate chips with up to 7 mm long chambers for trapping RBCs after the first chamber of 2 mm. The inlet channel height (h0 = 12 μm) is the smallest dimension of the chip. A low-cost fabrication method may result in deviation in the channel height. To investigate the effect of channel height, simulations were performed to estimate trapping efficiencies of 9.77 and 4.95 μm microparticles with different inlet channel heights (h0 = 10, 12, and 14 μm). Simulation result shows that the inlet channel height has effect on the trapping efficiency at high flow rates, however, it is negligible at the experimental condition of V0 = 400 μm s−1 (Figure S5 in ESI).
2.5. Microparticle and blood sample preparation Prior to cell sorting, we carried out experiments on carboxylate modified fluorescent polystyrene beads (Bangs lab, IN, USA) of 4.95 μm and 9.77 μm diameters as the model particle for RBC and WBC, respectively. Under fluorescent microscopy, smaller beads are observed as blue (excitation 360 nm, emission 450 nm) and larger particles as green (excitation 480 nm, emission 520 nm). The beads were dispersed in PBS prior to use. The blood sample was supplied from Korea University Guro Hospital. Ten healthy males (age: 29 ± 9 years; height: 178 ± 4 cm; bodyweight: 85 ± 11 kg; BMI: 26.8 ± 3.7) between the ages of 18 and 50 years volunteered to participate in this study. This study was approved by the ethics committee of Korea University Guro Hospital, and written informed consent was obtained from all participates in accordance with the Declaration of Helsinki (IRB no. KUGH13093-001). For cell sorting experiments, 1 μL whole blood sample was diluted 100 times with 1× Dulbecco PBS buffer (Sigma-Aldrich, MO, USA) to maintain normal osmotic pressure. The WBCs were stained with DAPI (4′,6-diamidino-2-phenylindole, Sigma-Aldrich, MO, USA) following the supplier’s protocol.
3.2. Trapping of microbeads 2.6. Experimental procedure To assess trapping efficiency under several combinations of chamber length and mean velocity, we performed experiments using fluorescent polystyrene microbeads (Fig. 3a). At slow velocities, particles sediment near the chamber entrance. By increasing velocity, particles gain the chance to escape from the chamber before being trapped. In addition, Fig. 3a hints to a correlation between mean velocity and particle spread over the length of channel. Based on this observation, V0 = 200 μm s−1 is too low since it can lead to aggregation of microbeads in short range. Moreover, bead adhesion to the inlet channel was observed at low velocities, even increasing the chance of channel clogging. The number of beads entering chamber and being trapped were counted to calculate trapping efficiencies presented in Fig. 3b–c for 9.77 μm and 4.95 μm beads. By comparing these two graphs we concluded V0 = 400 μm s−1 and ℓ1 = 2 mm and ℓ2 = 7 mm as the proper condition for separating the two sizes of particles. In such combination of parameters, about 90 % of WBCs are expected to be trapped in the first chamber with only about 20 % of RBCs being trapped in the same chamber. In the second chamber, over 95 % of RBCs entering it are expected to be trapped on the surface before reaching the exit.
Sample solution was injected into the inlet channel for 3–5 min to analyze the trapping efficiency at designated flow velocities controlled by a syringe pump (Harvard Apparatus, MA, USA). Then the releasing efficiency was analyzed with increased flow velocities. The fluorescent image of microbeads and WBCs was acquired by a fluorescence microscope (Ti-U, Nikon, Japan) equipped with a CMOS image sensor (DP74, Olympus, Japan). RBCs were analyzed under bright-field condition. Both images were taken with ×4 objective lens. The trapping and releasing efficiencies were estimated by automatically counting the particle or cell number using ImageJ open-source software (National Institutes of Health, MD, USA). If a chamber had overlapped particles or cells, manual counting was performed with higher magnification (×20) objective lens. Representative images of microbead and cell counting are shown in Figure S3 of ESI. Experimental trapping efficiency was defined as the number of particles trapped in a chamber divided by the number of particles entered the chamber during the trapping phase. Similarly, releasing efficiency was defined as the number of cells escaped from the chamber after increasing the inlet channel’s flow velocity over the number of trapped cells in the chamber before increasing the flow velocity.
3.3. Releasing of microbeads
3. Results and discussion
Once particles are trapped in chambers, adhesion force temporarily immobilize them on the substrate, thus releasing requires increased flow velocity until adhesion force is overcome [23]. However, the flow velocity near the trapped particle is low due to the parabolic flow profile of a pressure driven flow. For this reason, adhesion force must be decreased to prevent unrealistically high releasing flow velocity. Surface treatment by poly(ethylene oxide) (PEO) is one of the widelyknown methods of inhibiting cell and protein adhesion in microchannels [36]. Here we tried surface treatment with Pluronic F-68, a non-ionic surfactant based on PEO, with a PDMS-coated glass substrate. Additionally, we investigated effect of 0.1, 1, and 5 N NaOH treatment with bare glass as the substrate. Applying NaOH increases negative charges (-O−) on glass surface helping to repel negatively-charged cells and particles. We compared three different concentrations of each surface treatment solution in terms of releasing efficiency of each particle size. Additionally, a control experiment without any surface treatment was conducted. Results presented in Fig. 4a suggest that Pluronic F-68 is more effective than NaOH treatment in decreasing adhesion and therefore improving releasing efficiency, especially for smaller particles. The highest releasing efficiencies were achieved with 3 % v/v concentration of Pluronic F-68, therefore it was chosen for the
3.1. Microparticle trap-and-release simulation Following the simulation procedure described previously, trapping efficiencies of 4.95 and 9.77 μm particles were estimated for a combination of channel lengths (ℓ) and mean flow velocities (V0). Note that as it was shown by Park [23] and also through our simplified analysis, H does not have a significant effect on trapping efficiency if it is large enough so that the expanded region can be reasonably called a chamber. This conclusion was also confirmed by our simulations, showing that for H ≥ 5, trapping efficiency is not a function of H (Figure S4 of ESI). As previously discussed, this simulation is not suitable when H is small, thus we did not perform it for H < 5. Fig. 2a demonstrates trapping efficiency for 9.77 μm particles versus flow velocity. As seen from the graph, when chamber is longer than 2 mm, more than 90 % of particles are trapped even at higher flow velocities. However, for 1 mm chamber there is a decreasing trend when V0 exceeds 400 μm s−1. This is because at this velocity particles are trapped close to the exit of chamber. When designing conditions for the first chamber to trap 9.77 μm particles, trapping of smaller particles 4
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Fig. 2. Trapping efficiencies vs. mean velocity in the inlet channel obtained by simulation for (a) 9.77 μm particles and (b) 4.95 μm particles. In all the cases, H = 10.
3.4. Separation of blood cells
remaining experiments. Releasing efficiency of trapped particles was investigated with a variation of inlet channel flow velocities (Fig. 4b). More than 90 % of 4.95 μm beads are released by increasing flow velocity to 2500 μm s−1. However, at this velocity less than 10 % of 9.77 μm bead leave the trapping chamber. The 9.77 μm beads can be efficiently (> 80 %) released when the inlet channel flow velocity increased to 12,500 μm s−1. The different releasing flow velocities based on particle/cell size enable us to perform size-selective particle/cell sorting or enrichment.
Through simulation and microbead experiment, we finalized chamber length (2 and 7 mm for the first and second chamber, respectively) and height (H = 10 for both chambers) as well as the inlet channel mean flow velocity (V0 = 400 μm s−1 for trapping. V0 = 2500 μm s−1 and V0 = 12,500 μm s−1 for RBC and WBC releasing, respectively) for the selective WBC and RBC trapping and releasing. A 100× diluted whole blood in PBS was injected into the chip to achieve mean flow velocity of V0 = 400 μm s−1 in the inlet channel. With inlet
Fig. 3. Experimental result for trapping of fluorescent polystyrene beads. (a) Microbeads trapped in a 2 mm chamber at three mean velocities. The chamber is marked by the white dashed line. 9.77 and 4.95 μm beads have green and blue fluorescent color, respectively. Experimental trapping efficiency of (b) 9.77 μm and (c) 4.95 μm microbeads and vs. chamber length, obtained at various mean velocities (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article). 5
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Fig. 4. Releasing of microbeads (a) Comparison of releasing efficiency for each size of microbeads under various surface treatments. Releasing velocity was 12,500 μm s−1 for 9.77 μm beads and 2500 μm s−1 for 4.95 μm beads. In case of Pluronic F-68, substrate was coated by a thin PDMS layer. (b) Releasing efficiency as a function of inlet channel flow velocity for two sizes of microbeads with 3 % Pluronic F-68 coating.
velocity (12,500 μm s−1), the abundant RBCs in the first chamber can be selectively released by increasing the inlet flow velocity to the RBC releasing flow velocity, increasing the WBC purity and number in the first chamber. In this work, we verified the trapping and releasing efficiency of the proposed cell sorting or enrichment technique with a channel width of 1 mm. The resulting throughput of 0.288 μL min−1 cannot compete with inertial microfluidics or deterministic lateral displacement (DLD) which operates at high flowrate up to 10 mL min−1 [2]. However, the throughput of the presented device can be simply upscaled by increasing the channel width. In addition, the maximum inlet flow velocity V0 showing 100 % trapping efficiency linearly increases with the chamber length, as shown in Eq. (4). A detailed simulation result showing the linear relationship between the chamber length and throughput is presented in Figure S6 of ESI. Therefore, the throughput increases with the chamber width and length. For example, a device with chip size of 2 × 3 inch slideglass (45-times wider and 6-times longer chamber) can yield same trapping efficiency at 270-times higher flow rate (∼ 78 μL min−1). Moreover, in applications isolating larger cells such as circulating tumor cells (CTCs), not only target cells are heavier and sediment faster, but also one single chamber is enough to achieve the separation goal, which allow even longer chamber and higher flowrate. When the flowrate is high, the trapping and releasing sequences should be repeated before too many cells are trapped and overlapped in a chamber.
channel having cross-section of 12 μm × 1 mm, this flow velocity was achieved by injecting sample at 0.288 μL min−1. Trapping phase was completed after injecting 1.5 μL of sample, and flow stopped to assess trapping efficiency of cells in each chamber. With the pseudo-continuous nature of our technique, it was necessary to stop flow before too many cells are trapped in a chamber. As seen in Fig. 5a, 69 % of WBCs and 80 % of RBCs are trapped in the first and second chamber, respectively. It must be noted that the total number of cells trapped in both chambers is less than the injected cell number which was estimated from injection volume and bulk sample cell concentration. The number of lost cells which escaped both chambers was negligible. Most of the discrepancy came from adhered cells on the connecting tube wall. Therefore, the summed number of trapped cells in both chambers can be considered as actual number of injected cells into the first chamber. After completing trapping and counting cells in each chamber, releasing phase was performed. More than 85 % of RBCs trapped in the second chamber were released by increasing inlet channel mean flow velocity to 2500 μm s−1 as shown in Fig. 5b. In case of WBCs trapped in the first chamber, we were able to release more than 80 % by increasing the inlet channel mean flow velocity to 12,500 μm s−1. Fig. 5a also shows the degree of cross-contamination demonstrated as the number of trapped RBCs and WBCs in the first and second chamber, respectively. It is worth mentioning that in terms of trapped cell number in the first chamber, originally designed to trap WBCs, is dominated by RBCs. This happens because sample (i.e. diluted whole blood) is predominantly occupied by RBCs (×1000 more abundant in blood). Therefore, experiment on blood sample must be interpreted as enrichment of WBCs rather than separation purity. In addition, the WBC enrichment ratio or purity can be improved by repeating the sample injection and RBC release process. Because the RBC releasing flow velocity (2500 μm s−1) is much slower than the WBC releasing flow
4. Conclusions In this work, we have introduced a technique to trap and release cells in a size-selective manner by utilizing internal hydraulic jump phenomenon. We numerically investigated trapping and releasing efficiencies with parameters including chamber length, height, and flow 6
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Fig. 5. Trapping and releasing of blood cells (a) Number of WBCs and RBCs during trapping phase. V0 = 400 μm s−1. The number of injected cells is estimated from the volume of injected blood and its concentration. (b) Number of WBCs and RBCs in each chamber before and after releasing phase. V0 = 12,500 and 2500 μm s−1 for the WBC and RBC releasing in the first and second chamber, respectively.
(S7. Low-cost chip fabrication with machining in ESI) and tape cutting (S8. Low-cost chip fabrication with tape in ESI). Experimental results showed > 99 % of trapping efficiencies of 9.77 μm microbeads for both of fabrication methods, which suggest the possibility of low-cost massproduction of disposable cell sorter for POCT. Finally, the scheme presented here can be further developed for other applications. For instance, micro-valves can be implemented into the microchannel to enable collection of released cells without additional cross-contamination. In another example, substrate can be coated by adhesion promoters to permanently immobilize trapped cells, then use the device for on-chip culture and detection of large-size stem cell or CTCs.
velocity, and then validated the result with microbead experiments. We also demonstrated an application of our technique by separating WBCs and RBCs through enriching them in different chambers. Most microfluidic cell sorting methods, whether passive (e.g. inertial microfluidic) or active (e.g. acoustophoresis), do not only require the lateral dimension of microchannel to be in same order as cell size but also very sharply defined microchannel surface. Even though such devices are shown to have excellent performances in terms of separation purity and/or throughput [2,39–42], the μm range resolution of the channel design places concerns on low-cost mass production for POCT devices as well as channel clogging with real biological samples. In contrast, working principle of our technique is independent of microchannel’s lateral dimension, not only making mm-range channels possible but also providing a way to scale up the throughput by simply increasing channel width. Moreover, the trapping and releasing efficiency is robust with the channel height variation, further simplifying the restriction on fabrication resolution. Additionally, wider channel means less chance of clogging by cells or contaminations, and this leads to a more versatile approach for use in commercial devices and with untrained users. In this work, we have used standard soft lithography to fabricate the chip with exact dimensions for quantitative analysis of experimental results. However, a SU-8 mold fabrication with conventional photolithography is not a low-cost solution. For this reason, we also fabricated the chip with low-cost methods including machining
Conflict of interest The authors declares no conflict of interest.
Acknowledgement This research was supported by the National Research Foundation of Korea (NRF) (NRF-2011-0031866 and NRF-2018M3A9D7079485).
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Appendix A. Supplementary data
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Hamidreza Shirinkami received his BS degree in Mechanical Engineering from University of Tehran, Tehran, Iran. In 2014 he joined the Department of Biomedical Engineering at Korea University, Seoul, Korea for his PhD studies. His research interest is focused on Microfluidics, Neural interface, and Medical devices. Gaobo Wang received his BS degree in Chemistry and Chemical Engineering from Henan Institute of Science and Technology, Henan, China. In 2018, he received his Master’s degree in Biomedical Engineering from Korea University, Seoul, Korea. His research interest is focused on Microfluidics. Jinhyeok Park received his BS degree in Biomedical Engineering from Korea University, Seoul, Korea. In 2017, he received his Master’s degree in Biomedical Engineering from Korea University, Seoul, Korea. His research interest is focused on micro/nanofabrication and exosome analysis. Joonhyang Ahn received his BS degree in Biomedical Engineering from Korea University, Seoul, Korea. His research interest is focused on microfluidics. Yeonho Choi received his BS degree in Mechanical Engineering from Seoul National University, Seoul, Korea. In 2009, he received his PhD degree in Mechanical Engineering from the University of California at Berkeley, CA, USA. Since 2010, he has joined the Department of Biomedical Engineering at Korea University, Seoul, Korea. His research interest is focused on Plasmonics, Biomimetics, Exosome Analysis, and Deep-learning based diagnostics. Honggu Chun received his BS degree in Electrical Engineering and Computer Science from Seoul National University, Seoul, Korea. In 2004, he received his PhD degree in Biomedical Engineering from Seoul National University, Seoul, Korea. In 2011, he joined the Department of Biomedical Engineering at Korea University, Seoul, Korea. His research interest is focused on Iontronics, Neuro-interface, Clinical diagnostics, Exosome analysis, Single cell analysis, and Medical instrument.
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