- Email: [email protected]
DNA ligation using a disposable microfluidic device combined with a micromixer and microchannel reactor
Sensors and Actuators B 157 (2011) 735–741
Contents lists available at ScienceDirect
Sensors and Actuators B: Chemical journal homepage: www.elsevier.com/locate/snb
DNA ligation using a disposable microfluidic device combined with a micromixer and microchannel reactor Yong-Jun Ko a,1 , Joon-Ho Maeng b,2 , Yoomin Ahn c,∗ , Seung Yong Hwang d a
Department of Mechanical Engineering, Graduate School, Hanyang University, Seoul 133-791, Republic of Korea Department of Biochemistry, Graduate School, Hanyang University, Ansan, Gyeonggi 426-791, Republic of Korea Department of Mechanical Engineering, Hanyang University, 1271 Sa-3-dong, Sangnok-gu, Ansan, Gyeonggi 426-791, Republic of Korea d Division of Molecular and Life Science, Hanyang University, and GenoCheck Co. Ltd., Ansan, Gyeonggi 426-791, Republic of Korea b c
a r t i c l e
i n f o
Article history: Received 25 June 2010 Received in revised form 7 January 2011 Accepted 10 May 2011 Available online 17 May 2011 Keywords: DNA ligation Micromixer Micropillar Micronozzle Serpentine microreactor PDMS–glass microbiochip
a b s t r a c t A novel PDMS and glass-based microfluidic device consisting of a micromixer and microreactor for DNA ligation is described in this article. The new passive type planar micromixer is 10.33 mm long and composed of a straight channel integrated with nozzles and pillars, and the microreactor is composed of a serpentine channel. Mixing was enhanced by convective diffusion facilitated by the nozzles and pillars. The performance of the micromixer was analytically simulated and experimentally evaluated. The micromixer showed a good mixing efficiency of 87.7% at a 500 L/min flow rate (Re = 66.5). DNA ligation was successfully performed using the new microfluidic device, and ligation time was shortened from 4 h to 5 min. When used for on-chip ligation, this new micromixer offers advantages of disposability and portability. © 2011 Elsevier B.V. All rights reserved.
1. Introduction DNA ligation binds DNA fragments and is an essential element in genetic recombination. Generating large quantities of recombinant DNA requires the following steps. First, the DNA containing the gene of interest and the plasmid acting as a vector are cut into fragments using the same restriction enzyme. Next, the genecontaining DNA fragments are combined with the cut plasmids using DNA ligase, resulting in recombinant DNA plasmids. Host cells receive the recombinant DNA plasmids by transformation. As the cells reproduce, they clone the genes of interest carried by the recombinant DNA plasmids. The cloned recombinant DNA plasmids may be cut by restriction enzymes, and DNAs carrying the gene of interest separated from the plasmids by gel electrophoresis. DNA ligation performed in a general laboratory typically requires long processing times and is a tedious, labor-intensive procedure. Therefore, the development of a biochip platform with a miniaturized analysis device, with which ligation can be performed simply and quickly, is desirable. A miniaturized ligase-detection reaction
∗ Corresponding author. Tel.: +82 31 400 5281; fax: +82 31 406 5550. E-mail address: [email protected] (Y. Ahn). 1 Current address: R&D Center, LG Innotek Ltd., Ansan 426-791, Republic of Korea. 2 Current address: Gachon Bionano Research Institute, Kyungwon University, Seongnam 426-701, Republic of Korea. 0925-4005/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.snb.2011.05.016
chip was demonstrated for the detection of low-abundance DNA point mutations [1]. To execute DNA ligations with an integrated biochip such as a lab-on-a-chip, micromixing ligation reagents with DNA fragments is necessary. These can be products from previous lab-on-a-chip processes. A droplet technique based on dielectric electrowetting (EWOD) has been used to mix reagents in microfluidic DNA ligation techniques [2]. In typical EWOD systems, reagents stored in their own reservoirs are separated into droplets, and the droplets are rapidly and independently moved in the microchannels, and mixed by digital electrode control. Reagent quantities are diminished to sub-micro scale by droplet microfluidics using the incidental coplanar electrode microsystem. However, the moving reagents usually contaminate the channel surface, so expensive EWOD devices are difficult to reuse. Another simple device used in the mixing of reagents on a biochip is the micromixer [3], which can effectively mix ligation reagents. Using a serial processing chip for PCR and ligase reaction detection, a Y-shaped passive micromixer was used to mix ligations with PCR products [4]. However the Y-shaped mixer requires a very long flow length for effective mixing. Micromixers are generally classified into two types: active and passive. Active-type micromixers require an external power source to operate the actuators, such as micropumps [5,6], or to generate the electric or magnetic field [5]. Active micromixers usually provide rapid mixing. However, the fabrication process is not simple because of the moving components of the mixer. Also, integrating
736
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
the active mixer with the other components of the totally integrated biochip is difficult. Compared to the active-type micromixer, fabrication of a passive-type micromixer is not complicated, operation of the mixer is easier, and an external power source is not required. In addition, integrating the passive micromixer with other fluidic components is easier [7,8]. In a passive micromixer, the diffusion between the laminar flow fluids or chaotic advection is used to create a mixed flow. For rapid mixing, the interfaces between the two fluids are increased by splitting the fluid flows to substreams and recombining them in the microchannel [7]. In this manner, three-dimensional micromixers were developed [9]. However, these micromixers consist of multiple layers and their complicated structures require a longer fabrication process with a higher manufacturing cost. To remedy the shortcomings of these micromixers, mixers with micropillars in the microchannel have been recently developed [10,11]. However these previously reported passive planar micromixers have not yet been tried to real biomedical microfluidic systems. In this article, a new passive planar mixer was generated to perform DNA ligation for the first time in a disposable microfluidic device without bubble generation. This work extends our previous efforts using micronozzles and micropillars to enhance the mixing performance [12]. The performance of the micromixer was analytically estimated using CFD-ACE+ software and measured
experimentally. The proposed micromixer was integrated with a serpentine channel-type microreactor to generate a novel DNA ligation biochip. DNA ligations were performed with the integrated biochip, and the ligation products were evaluated. 2. DNA ligation biochip 2.1. Design and fabrication of the biochip A schematic of the proposed micromixer, which is combined with a channel type microreactor, is shown in Fig. 1. The micromixer is composed of micronozzles and micropillars, inspired by the modified Tesla-mixer [10] and obstruction micromixer [11]. In the modified Tesla-mixer, the angled surface is used to guide the fluid so that it collides. Here, the micronozzle was used in a novel micromixer. Asymmetric layout of the obstacles is known to have a greater effect on mixing in obstruction micromixers than the number of obstacles, and two or three obstacles are sufficient to enhance the mixing efficiency [11]. Hence the asymmetric layout of four obstacles with a nozzle was used in the channel. The first obstacle is located close to the nozzle exit in order to break up the injected fluids from the nozzle effectively. The stirring the fluids will create the lateral mass transport. The diameter of the inlet and outlet was 1.5 mm, with length 10.33 mm, width 400 m, and height 100 m. The precise dimensions of micromixer are in Fig. 2.
Fig. 1. Schematic diagram of key features and dimensions of the micromixer and microchannel type reactor. Mixers with (a) nozzles only, (b) pillars only, and (c) nozzles and pillars. Dashed lines represent the mixing zone sections.
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
737
Fig. 2. Layout of pillar and nozzle configuration of micromixer. Units: m.
The microreactor was designed as a serpentine channel because this produces very few or no bubbles [13]. The microreactor channel had 20 mm length, 400 m width, and 100 m height and spacing between the microreactor channel walls was 200 m. Generally, the Reynolds number (Re) of fluid flow in a microchannel is very low and fluid flow is laminar. Hence, mixing is by molecular diffusion between the laminar flows. A very long flow length is needed for complete mixing. In addition to molecular diffusion, mass transport in the laminar flow is generated by convection. Convection within the flow would enhance mixing. For transversal mixing, a convective velocity that is consistent with the direction of diffusion should be generated. Therefore, micropillars and micronozzles were formed in the microchannel. These cannot generate turbulent flow at low Re, but they can stir the flow [10]. Therefore, the mixing efficiency should be improved by transversal mass transport. In our previous study, a micromixer was designed with both pillars and nozzles. In this study, microchannel mixers with only pillars and only nozzles were designed to evaluate the effects of the pillars and nozzles on the mixing, as shown in Fig. 1a and b. Micromixers were fabricated using photolithography and micromolding techniques. The micromixer chip was of biocompatible materials with good optical properties. A PDMS molding technique was used to fabricate the micropillars and micronozzles, as described [12]. The inlet and outlet were made in the PDMS layer by mechanical punching. Finally, the PDMS layer and glass substrate were bonded after O2 plasma treatment. Bubbles can be caused by the hydrophobicity of PDMS [14], so this was prevented by modifying the PDMS surface to be hydrophilic. A PVP solution was used as a dynamic coating to prevent DNA adhesion to the PDMS surface in the PDMS–glass polymerase chain reaction (PCR) chip [15]. A PVP solution was used as surface treatment to make the PDMS surface hydrophilic. In our previous study [16], we performed a contact angle experiment using concentrations of PVP ranging from 2.5 × 10−9 to 2.5 weight% (wt%) in de-ionized water, to determine the optimal concentration required for surface treatment. The measured contact angle decreased as the concentration increased, and was saturated at about 20◦ at a concentration of over 0.25 wt%. This demonstrated that 0.25 wt% is sufficient for the passivation of the PDMS surface. Hence, the PDMS chip surface was coated with a 0.25 wt% PVP solution.
2.2. Simulation of micromixer The performance of the newly designed micromixers was quantitatively simulated using CFD-ACE+ software (ESI Group, USA). Water and ethanol were used as mixing fluids. The simulation was performed at steady state at 20 ◦ C. Densities of water and ethanol were assumed to be 997 and 789 kg/m3 , and dynamic viscosities
9.998 × 10−4 and 1.2 × 10−3 kg/m s−1 . The diffusion coefficient was assumed to be 1.2 × 10−9 m2 s−1 . Fluids were assumed to be Newtonian and incompressible. An analytical solution for the mixing of two different fluids does not yet exist, so a mixing simulation was performed for the mixing of two identical fluids, and the physical properties of the fluid were assumed to be the average values of water and ethanol. Fluid flow in the channel and the diffusion by the concentration gradient were determined using “flow” and “user scalar” modules. The mixing area in the micromixer was modeled by a finite volume method and three-dimensional unstructured grids. The number of meshes was about 210,000. A second-order upwind scheme and conjugate gradient squared with preconditioning were used to analyze the velocity field. An algebraic multigrid was used for pressure field calculations. At both inlets, a fixed velocity was assumed. At the outlet, fixed pressure was assumed. No-slip conditions were applied to the wall of the micromixer.
3. Experiment 3.1. Mixing rate Test fluids for measuring the mixing rate were deionized water and an ethanol solution blended with a blue ink as a dye. The two fluids were injected into each inlet with a syringe pump (KDS200, KD Scientific Inc., USA) at a constant flow rate. The flow rate of each inlet was the same and increased from 50 L/min to 500 L/min. These flow rates correspond to Reynolds number (Re) from 6.65 to 66.5. Mixing phenomena of two fluids in the micromixer at a steady-state flow were observed with an optical microscope (OPTITHOT300, Nikon Co., Japan). Optical images at the end of the mixing zones were measured collected with a digital camera (COOLPIX950, Nikon Co., Japan) connected to a microscope. Mixing efficiency was numerically calculated as flowing. Fluid flow images from the middle area in the vertical direction of the channel, obtained from the CFD simulation and experimental observation, were converted to a portable gray map (pgm) format by the PaintShop Photo Pro XI (Corel Corp., Canada) software. For converted images, bright intensities in the width diffusion direction in the channel were measured pixel-by-pixel which were located an equal distance away. The intensities (Ii ) were measured for 50 pixels from each diffusion line image. Pure deionized water was injected into the micromixer and the standard intensity of water (IW ) was obtained. The standard intensity of ethanol with dye was also obtained as IE . The intensity of each pixel was normalized as: IiN =
Ii − IE IW − IE
(1)
738
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
where IiN is the normalized intensity of a pixel. Finally, the mixing efficiency, E, was calculated as:
⎛ E = ⎝1 −
50
(1/50)
50
(1/50)
i
N,per.mix 2
(IiN − Ii
(IiN,unmix i
)
N,per.mix 2 − Ii )
⎞ ⎠ × 100 (%)
(2)
where IiN,unmix is the initial normalized intensity before mixing and N,per.mix
is the normalized intensity when two fluids were perfectly Ii mixed [17]. 3.2. DNA ligation and cloning For DNA ligation and cloning, the DNA Ligation Kit Version 1.2 (Takara, Japan) was used as the DNA ligase. Hind III (Roche, Switzerland) was used as the restriction enzyme, which cleaves the DNA sequence 5 - AAGCTT-3 , resulting in sticky ends. The T&A cloning vector (Real Biotech Corp., Taiwan) of 2728 bp was used as the plasmid vector. HV1 of 440 bp was used as the DNA insert. Echerichia coli strain DH5a competent cells (Real Biotech Corp., Taiwan) were used as the host cells. Ligase buffer obtained was from Invitrogen (USA). HV1 is in the hypervariable region of the mitochondrial DNA. The number of mitochondrial DNA molecules in one cell is 1000–3000 times greater than that of the nuclear DNA, and the mitochondrial DNA is relatively safe from external environmental factors. Hence, it is used for personal identification in medical research. Plasmid and HV1 were cut by a restriction enzyme (Hind III). The fabricated biochip was filled with 10 L ligase buffer. Two micropipette tips were put into the two inlets of the biochip. One of the tips was filled with 4 L ligase and 4 L HV1 and the other was filled with 4 L ligase, 1–3 L plasmids, and 1–3 L deionized water. The exact ratios of the ligase:plasmids:water in the tip were 4:1:3, 4:2:2, and 4:3:1. The volume of the deionized water was controlled to make up a total mixed volume of ligase, plasmid, and deionized water of 8 L, so that equal amounts of mixtures were injected at each inlet onto the chip. Buffer in the biochip was ejected from the outlet of the chip by micropipette, and reagents in the tips were injected into the chip. Hence, using exactly 10 L ligase buffer with the micropipette, we controlled for samples to reach the microreactor and remain for a specific amount of time. Reagents were mixed during injection. After mixing, reagents filled the reactor channel of the chip and DNA ligation including incubation with ligase was for 1–5 min at room temperature. Finally, the product was ejected from the outlet with a micropipette. The products were transformed into host cells, and cells were cultured on agar for over 12 h. Colonies with recombinant plasmids were white, while colonies with non-recombinant plasmids were blue. White colonies were selected and re-cultured in Luria–Bertani medium. Plasmid DNA was isolated and was cut with Hind III. Cut fragments were separated by gel electrophoresis to determine if HV1 was successfully inserted into the plasmid vector in the newly developed microfluidic device. 4. Results and discussion 4.1. Micromixer performance The mixing rate was simulated and measured for micromixers with pillars only, with nozzles only, and with both. The effect of the flow rate on the mixing efficiency of micromixer at 10.32 mm downstream is shown in Fig. 3. The mixing rates were generally lower than the expectations predicted by the simulation, probably because the dimension and shape of the fabricated micromixer were not exactly the same the same as those of the design. Also, the
Fig. 3. Measured and simulated mixing efficiency as function of flow rate.
microscope was not focused precisely at the center line in the channel when images were collected to measure the intensity. However, the experimental results show that mixing rate tended to be similar to the simulation. When the mixer was composed of only nozzles, the mixing efficiency was independent of the flow rate and almost constant at 31.6–32.5%. When only nozzles were in the mixer channel, the mixing of convective diffusion caused by the nozzles was not significant because of the distance between the nozzles and the boundary layer of the two fluids. Mixing occurred only by diffusion between different concentrations through the interface of the two fluids, even though the flow rate increased. When only micropillars were in the channel, mixing efficiency increased with the flow rate, from 59.2% to 87.8%. As the flow rate increased, the fluid velocity, in direction Y in Fig. 1, which was toward the interface between the two fluids, increased because of the pillars, causing convective diffusion. Hence, the diffusion from the concentration differences at the interface of the two fluids was amplified by the convective diffusion. The convective diffusion is dependent on flow speed, so mixing rate grows with the flow rate. Our results showed that the mixing efficiency of the micromixer composed of nozzles and pillars increased from 71.0% to 88.2% as the flow rate increased from 50 L/min to 500 L/min (Re increased from 6.65 to 66.5). When the nozzles and pillars were combined, the fluid flow vertical to the main flow direction along the channel was produced at first by the nozzles and then affected by the pillars. As a result, the vertical component of the fluid flow was enhanced toward the interface of the fluids. Therefore, the convective diffusion became strong enough to increase the mixing efficiency. Since the vertical velocity of the fluid flow increased with flow rate, the efficiency increased as well. Fig. 4 shows the simulated mixing pattern in the micromixer composed of nozzles and pillars. The laminar flows of the two injected fluids are split and recombined by the pillars and nozzles, produce more interfaces between the two fluids, and enhancing the diffusion mixing. Therefore, mixing efficiency is expected to increase by this process as well as by convective diffusion. Improved mixing efficiency was obtained by the proposed mixer but not by mixers with only nozzles or only pillars. When the flow rate was greater than 300L/min (Re = 39.9), the mixer with only pillars showed almost the same efficiency as the mixer with nozzles and pillars. However, the proposed mixer was more efficient at low flow rates than the mixers with only nozzles or pillars. The mixing efficiency at each section as 1–6 units as shown in Fig. 1 was simulated and experimentally measured when flow rate is 50, 300, and 500 mL/min for the pillar and nozzle combination
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
739
Fig. 4. Simulated mixing patterns along the channel of the micromixer.
mixer. The simulated and measured mixing efficiency is shown in Fig. 5. In Fig. 5, the measurements for each units from 1 to 6 were performed at downstream of 1.72, 3.44, 5.16, 6.88, 8.6, and 10.32 mm, respectively. Compared to the simulated results, measured mixing efficiency showed a sudden increase at the first mixer unit, however the increasing rate of the efficiency as the fluids pass the mixer channel was less than that of simulated mixing efficiency. When the flow rate was 50 L/min, the measured mixing ratio increased gently as fluids passed through each unit of the micromixer. This is because the convection is not dominant when the flow rate is low. The mixing is accomplished only by diffusion. When fluids are mixed by diffusion through the interface of the two fluids, mixing ratio increases linearly in proportion to the length of the interface. Therefore, the mixing progresses slowly as the fluids flow through the micromixer. However, when the flow rate is 300 or 500 L/min, the mixing ratio rapidly increase more than that of 50 L/min as shown in Fig. 5. This is because the convection becomes significant when the flow rate is high. The mixing accelerates as the fluid flow through the mixer channel. However the mixing efficiency does not increase linearly. The increasing ratios slow down with the mixer channel length.
Results from a previous study showed that the proposed mixer had an improved mixing performance compared to a typical modified tesla mixer and obstruction mixer [12]. The modified tesla mixer achieved ∼78% mixing at 5 mm downstream from an entrance for 100 m wide channels at Re = 0.05 [11], while we observed an ∼83% mixing rate at about 5 mm downstream, for Re = 66.5 flow. The obstruction mixer is capable of a maximum of ∼56% in the first unit of the mixer set for a 300-mm wide channel [10], while the new micromixer reached ∼64% mixing after the first unit for Re = 66.5. At higher Reynolds number (Re > ∼1), the mixing performance of the modified tesla mixer [18] and the obstruction mixer [19] is substantially improved. Simulation results of the diamond-shaped obstruction micromixer showed 70–90% mixing performance at 5 mm from the channel entrance for the 10 < Re < 100 range [19]. Recently reported passive micromixers have a mixing efficiency of 80–98% at a flow rate of 5–1200 L/min [7,20]. In comparison, the mixing efficiency of our novel micromixer was measured as 88.2% at a 500 L/min flow rate (Re = 66.5). This shows that the micromixer can be efficient even with only a simple two-dimensional structure of nozzles and pillars instead of a complex three-dimensional structure. In addition to increasing the flow rate, the mixing efficiency of the new mixer could be increased by increasing the length of the mixer channel with more nozzles and pillars. 4.2. DNA ligation by biochip
Fig. 5. Measured and simulated mixing efficiency at each unit of the pillar and nozzle combination mixer.
The minimum required ligation time in the microfluidic device was observed with 3 L of plasmids. After ligation reactions of 1, 3, and 5 min on the biochip, ligation was examined. When the ligations were performed for 1 min, a few white colonies could be found with the biochip products. However, white colonies appeared after transformation with products from ligation times of 3 or 5 min. Fig. 6 shows gel electrophoresis of plasmids from white colonies produced with products from 3 and 5 min ligation times. When the reaction time was 5 min, the 2728-bp plasmid and 440-bp HV1 insert appeared (lanes 4 and 5, markers in the left lane). However, the HV1 insert did not appear in lanes 2 and 3, and another insert appeared in lane 1. In these colonies, DNA fragments other than HV1 were inserted in the plasmid during the ligation process. Because the size of the contaminant DNA fragments was very large or very small, the bands of these fragments were not seen by gel electrophoresis. 3 min was not sufficient for ligation. Based
740
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
Fig. 6. Gel electrophoresis; lanes 1–3 are results from a 3-min reaction, lanes 4 and 5 are results from a 5-min reaction, and left column is DNA markers.
on these our results, the minimum required ligation time with the biochip was about 5 min. This is similar to other on-chip DNA ligation processes based on droplet microfluidic systems [2]. Ligation by a traditional laboratory method requires about 4 h [21]. These results show ligation of specific DNA fragments can be completed within 5 min using the biochip presented here. Some techniques, such as proximity ligation, require only 5 min for ligation with T4 DNA ligase without microfluidic mixers [22]. In the other microfluidic chip, the ligation detection was performed with a ligation mixture premixed with a polymerase chain reaction (PCR) product, and did not use flow through a mixer. This reaction was completed in 6.5 min [1]. In an advanced version of this chip, a straight channel- type of micromixer was used for mixing and the ligase reaction could be detected after 8.1 min [4]. These results imply that additional 1.6 min was spent for mixing. The total ligation time (8.1 min) is longer than our results (5 min), probably because the mixing performance of the straight channel was inferior to our micromixer. Therefore, mixing could affect and enhance the DNA ligation reaction in a totally integrated lab-on-a-chip. The ligase reaction with the same mixing procedure could be shortened to 4.1 min by optimization of reagent selections and reaction conditions [23]. The performance of the novel microfluidic device was compared to the conventional laboratory method. Products from the conventional ligation process and from the biochip with a 5-min ligation time and volume of 3 L plasmid were obtained and transferred into host cells. The cells were cultured and white colonies selected. Gel electrophoresis with plasmids from the selected cells was performed before enzymatic cutting, and the results are shown in Fig. 7a. The products (lanes 6–10 in the figure) from the biochip show the same results as the conventional method (lanes 1–5). The selected cells were cultured again in Luria–Bertani medium. Plasmid DNA was obtained from five white colonies and cut with Hind III. Gel electrophoresis was used to visualize the DNA fragments. Fig. 7b shows the results. Plasmid vectors and HV1 inserts were detected from all five white colonies. However, an unknown DNA fragment of about 800 bp was detected in one, probably from contamination during the ligation or cell culturing after the transformation. This can be minimized by carefully performing the ligation procedure. For example, after the plasmid vectors and HV1 inserts were are cut, gel-electrophoresis of the cut products and extraction of the desired DNA fragments is advised. As shown in Fig. 7b, one of the five ligations by the conventional method also failed. These results confirmed that DNA ligation was successfully performed using the novel microfluidic device. We attempted to minimize the volume of the vector in ligations with the new biochip. We tested 1, 2, and 3 L of plasmid vectors with the biochip ligation. The results are shown in Fig. 8. When 1 and 2 L vector volumes were used, other DNA fragments appeared
Fig. 7. Electrophoresis (a) after transformation and (b) after cutting with restriction enzyme; lanes 1–5 are ligation by the conventional laboratory method and lanes 6–10 are ligation by the biochip.
Fig. 8. Electrophoresis: lanes 1 and 2 are from 1 L of vector, lanes 3 and 4 are from 2 L, and lane 5 is from 5 L of vector.
in the gel electrophoresis, as shown in lanes 1–4 of Fig. 7. Reactions with 1 or 2 L plasmid volumes failed, but the ligation with 3 L produced good results when the ratio of ligase:plasmids:water in the second inlet was 4:3:1. Therefore, at least a 5-min reaction time and a 3-L vector volume are needed to perform DNA ligation with the proposed new biochip. The total volume of reagents for the ligation in the reservoirs of the new biochip was 16 L, and this was approximately the same as the 15 L of the traditional ligation process [2]. Reagent usage efficiency of the new biochip was not as high as the droplet microfluidic device, which used only 2.1 L of reagents with no waste. These results reveal a disadvantage of the PDMS–glass biochip, which is its high surface area-to-volume ratio [24]. This disadvantage is caused by the adhesion of DNA and ligase to the PDMS, and is not completely avoided by surface passivation with PVP additive. However, a novel ligation chip has been fabricated with only a simple soft-lithography technique. Hence, the chip can be made very cheaply on a mass production basis. We expect that the novel ligation chip is most suitable as a disposable unit and might be effectively applied as a point-of-care lab-on-a-chip system. 5. Conclusions A DNA ligation biochip consisting of a micromixer and microchannel reactor was inexpensively fabricated with a PDMS replica molding technique. The PDMS chip surface was coated
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
with a 0.25 wt% PVP solution to prevent bubble generation. The micromixer had a simple structure of micronozzles and micropillars. Mixing efficiency was enhanced by convective diffusion and fluid splitting caused by the micronozzles and micropillars, and increased with flow rate. The micromixer was 10.33 mm long and had a maximum mixing rate of 87.7% at a flow rate of 500 L/min (Re = 66.5). DNA ligation was successfully accomplished in the integrated microfluidic device, with time shortened from about 4 h for the conventional laboratory technique to only 5 min using the proposed method. The ligation used 8 L ligase, 4 L HV1 insert, and 3 L plasmid vector. Although the PDMS–glass biochip rapidly completed the ligation, the biochip did not require lower amounts of reagents than conventional ligation devices. To improve the performance of the PDMS-based ligation biochip, future studies are needed to determine a technique that inhibits the interaction of the ligation reagents with the PDMS chip surface. The obtained results proved the usefulness of the in-plane passive microfluidic mixer for practical biomedical analysis. Furthermore, because this DNA ligation platform can be fabricated using polymer replica molding technologies, inexpensive disposable on-chip microfluidic systems with this PDMS-based ligation platform are also practicable. The new disposable ligation chip is expected to be useful as a totally integrated lab-on-a-chip for genetic recombination. References [1] M. Hashimoto, M.L. Hupert, M.C. Murphy, S.A. Soper, Y.-W. Cheng, F. Barany, Ligase detection reaction/hybridization assays using three-dimensional microfluidic networks for the detection of low-abundent DNA point mutations, Anal. Chem. 77 (2005) 3243–3255. [2] Y.-J. Liu, D.-J. Yao, H.-C. Lin, W.-Y. Chang, H.-Y. Chang, DNA ligation of ultramicro volume using an EWOD microfluidic system with coplanar electrodes, J. Micromech. Microeng. 18 (2008) 045017. [3] V. Hessel, H. Löwe, F. Schönfeld, Micromixers – a review on passive and active mixing principles, Chem. Eng. Sci. 60 (2005) 2479–2501. [4] M. Hashimoto, F. Barany, S.A. Soper, Polymerase chain reaction/ligase detection reaction/hybridization assays using flow-through microfluidic devices for the detection of low-abundant DNA point mutations, Biosens. Bioelectron. 21 (2006) 1915–1923. [5] L.H. Lu, K.S. Ryu, C. Liu, A magnetic microstirrer and array for microfluidic mixing, J. Microelectromech. Syst. 11 (2002) 462–469. [6] J.H. Tsai, L. Lin, Active microfluidic mixer and gas bubble filter driven by thermal bubble micropump, Sens. Actuator A: Phys. 97–98 (2002) 665–671. [7] F. Schönfeld, V. Hessel, C. Hofmann, An optimized split-and-recombine micro mixer with uniform ‘chaotic’ mixing, Lab. Chip 4 (2004) 65–69. ´ H.A. Stone, G.M. Whitesides, [8] A.D. Stroock, S.K.W. Dertinger, A. Ajdari, I. Mezic, Chaotic mixer for microchannels, Science 295 (2002) 647–651. [9] M. Elwenspoek, T.S.J. Lammerink, R. Miyake, J.H.J. Fluitman, Towards integrated microliquid handling systems, J. Micromech. Microeng. 4 (1994) 227–245. [10] H. Wang, P. Iovenitti, E. Harvey, S. Masood, Optimizing layout of obstacles for enhanced mixing in microchannels, Smart Mater. Struct. 11 (2002) 662–667. [11] A.A.S. Bhagat, I. Papautsky, Enhancing particle dispersion in a passive planar micromixer using rectangular obstacles, J. Micromech. Microeng. 18 (2008) 085005. [12] Y.-J. Ko, S.-M. Ha, H.-J. Kim, D.-H. Lee, Y. Ahn, Development of a PDMS–glass hybrid microchannel mixer composed of micropillars and micronozzles, J. Solid Mech. Mater. Eng. 2 (2008) 445–454.
741
[13] C.-H. Cho, W. Cho, Y. Ahn, S.-Y. Hwang, PDMS–glass serpentine microchannel chip for time domain PCR with bubble suppression in sample injection, J. Micromech. Microeng. 17 (2007) 1810–1817. [14] W. Cho, Y.-J. Ko, Y. Ahn, J.-Y. Yoon, N.G. Cho, Surface modification effect of wetability on the performance of a PDMS-based valveless micropump, Key Eng. Mater. 326–328 (2006) 297–300. [15] B.C. Giordano, E.R. Copeland, J.P. Landers, Towards dynamic coating of glass microchip chambers for amplifying DNA via the polymerase chain reaction, Electrophoresis 22 (2001) 334–340. [16] D.-Y. Kang, Y. Ahn, S.Y. Hwang, A study about biochip combined with micro mixer and reactor for DNA ligation, Trans. Korean Soc. Mech. Eng. A 32 (2008) 624–632. [17] N.L. Jeon, S.K.W. Dertinger, D.T. Chiu, I.S. Choi, A.D. Stroock, G.M. Whitesides, Generation of solution and surface gradients using microfluidic systems, Langmuir 16 (2000) 8311–8316. [18] C.-C. Hong, J.-W. Choi, C.H. Ahn, A novel in-plane passive microfluidic mixer with modified Tesla structures, Lab. Chip 4 (2004) 109–113. [19] A.A.S. Bhagat, E.T.K. Peterson, I. Papautsky, A passive planar micromixer with obstructions for mixing at low Reynolds number, J. Micromech. Microeng. 17 (2007) 1017–1024. [20] R.H. Liu, M.A. Stremler, K.V. Sharp, M.G. Olsen, J.G. Santiago, R.J. Adrian, J.H. Aref, D.J. Beebe, Passive mixing in a three-dimensional serpentine microchannel, J. Microelectromech. Syst. 9 (2000) 190–197. [21] J. Sambrook, D.W. Russell, Molecular Cloning, vol. 1, third ed., CSHL Press, 2001, pp. 1.84–1.87. [22] M. Gullberg, S.M. Gústafsdóttir, E. Schallmeiner, J. Jarvius, M. Bjarnegård, C. Betsholtz, U. Landegren, S. Fredriksson, Cytokine detection by antibody-based proximity ligation, Proc. Natl. Acad. Sci. U.S.A. 101 (2004) 8420–8424. [23] M. Hashimoto, F. Barany, F. Xu, S.A. Soper, Serial processing of biological reactions using flow-through microfluidic devices: coupled PCR/LDR for the detection of low-abundant DNA point mutations, Analyst 132 (2007) 913–921. [24] C. Zhan, J. Xu, W. Ma, W. Zheng, PCR microfluidic devices for DNA amplification, Biotechnol. Adv. 24 (2006) 243.
Biographies Yong-Jun Ko received a Ph.D. in mechanical engineering in 2009 from Hanyang University. He is now working at R&D Center, LG Innotek Ltd., Ansan, Korea. His main research interests include microfluidic devices, microbiosensors, microbiochip for protein and cell, plastic/glass microfabrication, and nanotechnology. Joon-Ho Maeng received his M.S. (2006) and Ph.D. degree (2009) in the Department of Biochemistry, Hanyang University, Korea. He is now a postdoctoral research fellow in the Kachon Bionano Research Institute, Kyungwon Unversity, Korea. His main research interests include biosensors, bioMEMS, point-of-care tests, nanotechnology and biomedical devices. Yoomin Ahn received his B.S. in 1984 and M.S. in 1986 from Seoul National University, Korea. He received his Ph.D. in 1992 from Purdue University, USA. He is a professor in the Department of Mechanical Engineering at Hanyang University, Korea. His present research interests include microelectromechanical systems, microfluidic biochip, and lab-on-a-chip. Seung Yong Hwang is a full professor at the Laboratory “Integrated Genomics”, Division of Molecular and Life Science at Hanyang University in Korea. He received his B.Sc. from Hanyang University in 1989 and then finished his Ph.D. in 1995 at Monash University in Australia. Later, he worked at the Stanford University in the USA as a postdoctoral fellow. He is also running a biotech company, GenoCheck Co. Ltd. His main research interests are toxicogenomics and developing biochips for various screening systems.
Contents lists available at ScienceDirect
Sensors and Actuators B: Chemical journal homepage: www.elsevier.com/locate/snb
DNA ligation using a disposable microfluidic device combined with a micromixer and microchannel reactor Yong-Jun Ko a,1 , Joon-Ho Maeng b,2 , Yoomin Ahn c,∗ , Seung Yong Hwang d a
Department of Mechanical Engineering, Graduate School, Hanyang University, Seoul 133-791, Republic of Korea Department of Biochemistry, Graduate School, Hanyang University, Ansan, Gyeonggi 426-791, Republic of Korea Department of Mechanical Engineering, Hanyang University, 1271 Sa-3-dong, Sangnok-gu, Ansan, Gyeonggi 426-791, Republic of Korea d Division of Molecular and Life Science, Hanyang University, and GenoCheck Co. Ltd., Ansan, Gyeonggi 426-791, Republic of Korea b c
a r t i c l e
i n f o
Article history: Received 25 June 2010 Received in revised form 7 January 2011 Accepted 10 May 2011 Available online 17 May 2011 Keywords: DNA ligation Micromixer Micropillar Micronozzle Serpentine microreactor PDMS–glass microbiochip
a b s t r a c t A novel PDMS and glass-based microfluidic device consisting of a micromixer and microreactor for DNA ligation is described in this article. The new passive type planar micromixer is 10.33 mm long and composed of a straight channel integrated with nozzles and pillars, and the microreactor is composed of a serpentine channel. Mixing was enhanced by convective diffusion facilitated by the nozzles and pillars. The performance of the micromixer was analytically simulated and experimentally evaluated. The micromixer showed a good mixing efficiency of 87.7% at a 500 L/min flow rate (Re = 66.5). DNA ligation was successfully performed using the new microfluidic device, and ligation time was shortened from 4 h to 5 min. When used for on-chip ligation, this new micromixer offers advantages of disposability and portability. © 2011 Elsevier B.V. All rights reserved.
1. Introduction DNA ligation binds DNA fragments and is an essential element in genetic recombination. Generating large quantities of recombinant DNA requires the following steps. First, the DNA containing the gene of interest and the plasmid acting as a vector are cut into fragments using the same restriction enzyme. Next, the genecontaining DNA fragments are combined with the cut plasmids using DNA ligase, resulting in recombinant DNA plasmids. Host cells receive the recombinant DNA plasmids by transformation. As the cells reproduce, they clone the genes of interest carried by the recombinant DNA plasmids. The cloned recombinant DNA plasmids may be cut by restriction enzymes, and DNAs carrying the gene of interest separated from the plasmids by gel electrophoresis. DNA ligation performed in a general laboratory typically requires long processing times and is a tedious, labor-intensive procedure. Therefore, the development of a biochip platform with a miniaturized analysis device, with which ligation can be performed simply and quickly, is desirable. A miniaturized ligase-detection reaction
∗ Corresponding author. Tel.: +82 31 400 5281; fax: +82 31 406 5550. E-mail address: [email protected] (Y. Ahn). 1 Current address: R&D Center, LG Innotek Ltd., Ansan 426-791, Republic of Korea. 2 Current address: Gachon Bionano Research Institute, Kyungwon University, Seongnam 426-701, Republic of Korea. 0925-4005/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.snb.2011.05.016
chip was demonstrated for the detection of low-abundance DNA point mutations [1]. To execute DNA ligations with an integrated biochip such as a lab-on-a-chip, micromixing ligation reagents with DNA fragments is necessary. These can be products from previous lab-on-a-chip processes. A droplet technique based on dielectric electrowetting (EWOD) has been used to mix reagents in microfluidic DNA ligation techniques [2]. In typical EWOD systems, reagents stored in their own reservoirs are separated into droplets, and the droplets are rapidly and independently moved in the microchannels, and mixed by digital electrode control. Reagent quantities are diminished to sub-micro scale by droplet microfluidics using the incidental coplanar electrode microsystem. However, the moving reagents usually contaminate the channel surface, so expensive EWOD devices are difficult to reuse. Another simple device used in the mixing of reagents on a biochip is the micromixer [3], which can effectively mix ligation reagents. Using a serial processing chip for PCR and ligase reaction detection, a Y-shaped passive micromixer was used to mix ligations with PCR products [4]. However the Y-shaped mixer requires a very long flow length for effective mixing. Micromixers are generally classified into two types: active and passive. Active-type micromixers require an external power source to operate the actuators, such as micropumps [5,6], or to generate the electric or magnetic field [5]. Active micromixers usually provide rapid mixing. However, the fabrication process is not simple because of the moving components of the mixer. Also, integrating
736
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
the active mixer with the other components of the totally integrated biochip is difficult. Compared to the active-type micromixer, fabrication of a passive-type micromixer is not complicated, operation of the mixer is easier, and an external power source is not required. In addition, integrating the passive micromixer with other fluidic components is easier [7,8]. In a passive micromixer, the diffusion between the laminar flow fluids or chaotic advection is used to create a mixed flow. For rapid mixing, the interfaces between the two fluids are increased by splitting the fluid flows to substreams and recombining them in the microchannel [7]. In this manner, three-dimensional micromixers were developed [9]. However, these micromixers consist of multiple layers and their complicated structures require a longer fabrication process with a higher manufacturing cost. To remedy the shortcomings of these micromixers, mixers with micropillars in the microchannel have been recently developed [10,11]. However these previously reported passive planar micromixers have not yet been tried to real biomedical microfluidic systems. In this article, a new passive planar mixer was generated to perform DNA ligation for the first time in a disposable microfluidic device without bubble generation. This work extends our previous efforts using micronozzles and micropillars to enhance the mixing performance [12]. The performance of the micromixer was analytically estimated using CFD-ACE+ software and measured
experimentally. The proposed micromixer was integrated with a serpentine channel-type microreactor to generate a novel DNA ligation biochip. DNA ligations were performed with the integrated biochip, and the ligation products were evaluated. 2. DNA ligation biochip 2.1. Design and fabrication of the biochip A schematic of the proposed micromixer, which is combined with a channel type microreactor, is shown in Fig. 1. The micromixer is composed of micronozzles and micropillars, inspired by the modified Tesla-mixer [10] and obstruction micromixer [11]. In the modified Tesla-mixer, the angled surface is used to guide the fluid so that it collides. Here, the micronozzle was used in a novel micromixer. Asymmetric layout of the obstacles is known to have a greater effect on mixing in obstruction micromixers than the number of obstacles, and two or three obstacles are sufficient to enhance the mixing efficiency [11]. Hence the asymmetric layout of four obstacles with a nozzle was used in the channel. The first obstacle is located close to the nozzle exit in order to break up the injected fluids from the nozzle effectively. The stirring the fluids will create the lateral mass transport. The diameter of the inlet and outlet was 1.5 mm, with length 10.33 mm, width 400 m, and height 100 m. The precise dimensions of micromixer are in Fig. 2.
Fig. 1. Schematic diagram of key features and dimensions of the micromixer and microchannel type reactor. Mixers with (a) nozzles only, (b) pillars only, and (c) nozzles and pillars. Dashed lines represent the mixing zone sections.
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
737
Fig. 2. Layout of pillar and nozzle configuration of micromixer. Units: m.
The microreactor was designed as a serpentine channel because this produces very few or no bubbles [13]. The microreactor channel had 20 mm length, 400 m width, and 100 m height and spacing between the microreactor channel walls was 200 m. Generally, the Reynolds number (Re) of fluid flow in a microchannel is very low and fluid flow is laminar. Hence, mixing is by molecular diffusion between the laminar flows. A very long flow length is needed for complete mixing. In addition to molecular diffusion, mass transport in the laminar flow is generated by convection. Convection within the flow would enhance mixing. For transversal mixing, a convective velocity that is consistent with the direction of diffusion should be generated. Therefore, micropillars and micronozzles were formed in the microchannel. These cannot generate turbulent flow at low Re, but they can stir the flow [10]. Therefore, the mixing efficiency should be improved by transversal mass transport. In our previous study, a micromixer was designed with both pillars and nozzles. In this study, microchannel mixers with only pillars and only nozzles were designed to evaluate the effects of the pillars and nozzles on the mixing, as shown in Fig. 1a and b. Micromixers were fabricated using photolithography and micromolding techniques. The micromixer chip was of biocompatible materials with good optical properties. A PDMS molding technique was used to fabricate the micropillars and micronozzles, as described [12]. The inlet and outlet were made in the PDMS layer by mechanical punching. Finally, the PDMS layer and glass substrate were bonded after O2 plasma treatment. Bubbles can be caused by the hydrophobicity of PDMS [14], so this was prevented by modifying the PDMS surface to be hydrophilic. A PVP solution was used as a dynamic coating to prevent DNA adhesion to the PDMS surface in the PDMS–glass polymerase chain reaction (PCR) chip [15]. A PVP solution was used as surface treatment to make the PDMS surface hydrophilic. In our previous study [16], we performed a contact angle experiment using concentrations of PVP ranging from 2.5 × 10−9 to 2.5 weight% (wt%) in de-ionized water, to determine the optimal concentration required for surface treatment. The measured contact angle decreased as the concentration increased, and was saturated at about 20◦ at a concentration of over 0.25 wt%. This demonstrated that 0.25 wt% is sufficient for the passivation of the PDMS surface. Hence, the PDMS chip surface was coated with a 0.25 wt% PVP solution.
2.2. Simulation of micromixer The performance of the newly designed micromixers was quantitatively simulated using CFD-ACE+ software (ESI Group, USA). Water and ethanol were used as mixing fluids. The simulation was performed at steady state at 20 ◦ C. Densities of water and ethanol were assumed to be 997 and 789 kg/m3 , and dynamic viscosities
9.998 × 10−4 and 1.2 × 10−3 kg/m s−1 . The diffusion coefficient was assumed to be 1.2 × 10−9 m2 s−1 . Fluids were assumed to be Newtonian and incompressible. An analytical solution for the mixing of two different fluids does not yet exist, so a mixing simulation was performed for the mixing of two identical fluids, and the physical properties of the fluid were assumed to be the average values of water and ethanol. Fluid flow in the channel and the diffusion by the concentration gradient were determined using “flow” and “user scalar” modules. The mixing area in the micromixer was modeled by a finite volume method and three-dimensional unstructured grids. The number of meshes was about 210,000. A second-order upwind scheme and conjugate gradient squared with preconditioning were used to analyze the velocity field. An algebraic multigrid was used for pressure field calculations. At both inlets, a fixed velocity was assumed. At the outlet, fixed pressure was assumed. No-slip conditions were applied to the wall of the micromixer.
3. Experiment 3.1. Mixing rate Test fluids for measuring the mixing rate were deionized water and an ethanol solution blended with a blue ink as a dye. The two fluids were injected into each inlet with a syringe pump (KDS200, KD Scientific Inc., USA) at a constant flow rate. The flow rate of each inlet was the same and increased from 50 L/min to 500 L/min. These flow rates correspond to Reynolds number (Re) from 6.65 to 66.5. Mixing phenomena of two fluids in the micromixer at a steady-state flow were observed with an optical microscope (OPTITHOT300, Nikon Co., Japan). Optical images at the end of the mixing zones were measured collected with a digital camera (COOLPIX950, Nikon Co., Japan) connected to a microscope. Mixing efficiency was numerically calculated as flowing. Fluid flow images from the middle area in the vertical direction of the channel, obtained from the CFD simulation and experimental observation, were converted to a portable gray map (pgm) format by the PaintShop Photo Pro XI (Corel Corp., Canada) software. For converted images, bright intensities in the width diffusion direction in the channel were measured pixel-by-pixel which were located an equal distance away. The intensities (Ii ) were measured for 50 pixels from each diffusion line image. Pure deionized water was injected into the micromixer and the standard intensity of water (IW ) was obtained. The standard intensity of ethanol with dye was also obtained as IE . The intensity of each pixel was normalized as: IiN =
Ii − IE IW − IE
(1)
738
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
where IiN is the normalized intensity of a pixel. Finally, the mixing efficiency, E, was calculated as:
⎛ E = ⎝1 −
50
(1/50)
50
(1/50)
i
N,per.mix 2
(IiN − Ii
(IiN,unmix i
)
N,per.mix 2 − Ii )
⎞ ⎠ × 100 (%)
(2)
where IiN,unmix is the initial normalized intensity before mixing and N,per.mix
is the normalized intensity when two fluids were perfectly Ii mixed [17]. 3.2. DNA ligation and cloning For DNA ligation and cloning, the DNA Ligation Kit Version 1.2 (Takara, Japan) was used as the DNA ligase. Hind III (Roche, Switzerland) was used as the restriction enzyme, which cleaves the DNA sequence 5 - AAGCTT-3 , resulting in sticky ends. The T&A cloning vector (Real Biotech Corp., Taiwan) of 2728 bp was used as the plasmid vector. HV1 of 440 bp was used as the DNA insert. Echerichia coli strain DH5a competent cells (Real Biotech Corp., Taiwan) were used as the host cells. Ligase buffer obtained was from Invitrogen (USA). HV1 is in the hypervariable region of the mitochondrial DNA. The number of mitochondrial DNA molecules in one cell is 1000–3000 times greater than that of the nuclear DNA, and the mitochondrial DNA is relatively safe from external environmental factors. Hence, it is used for personal identification in medical research. Plasmid and HV1 were cut by a restriction enzyme (Hind III). The fabricated biochip was filled with 10 L ligase buffer. Two micropipette tips were put into the two inlets of the biochip. One of the tips was filled with 4 L ligase and 4 L HV1 and the other was filled with 4 L ligase, 1–3 L plasmids, and 1–3 L deionized water. The exact ratios of the ligase:plasmids:water in the tip were 4:1:3, 4:2:2, and 4:3:1. The volume of the deionized water was controlled to make up a total mixed volume of ligase, plasmid, and deionized water of 8 L, so that equal amounts of mixtures were injected at each inlet onto the chip. Buffer in the biochip was ejected from the outlet of the chip by micropipette, and reagents in the tips were injected into the chip. Hence, using exactly 10 L ligase buffer with the micropipette, we controlled for samples to reach the microreactor and remain for a specific amount of time. Reagents were mixed during injection. After mixing, reagents filled the reactor channel of the chip and DNA ligation including incubation with ligase was for 1–5 min at room temperature. Finally, the product was ejected from the outlet with a micropipette. The products were transformed into host cells, and cells were cultured on agar for over 12 h. Colonies with recombinant plasmids were white, while colonies with non-recombinant plasmids were blue. White colonies were selected and re-cultured in Luria–Bertani medium. Plasmid DNA was isolated and was cut with Hind III. Cut fragments were separated by gel electrophoresis to determine if HV1 was successfully inserted into the plasmid vector in the newly developed microfluidic device. 4. Results and discussion 4.1. Micromixer performance The mixing rate was simulated and measured for micromixers with pillars only, with nozzles only, and with both. The effect of the flow rate on the mixing efficiency of micromixer at 10.32 mm downstream is shown in Fig. 3. The mixing rates were generally lower than the expectations predicted by the simulation, probably because the dimension and shape of the fabricated micromixer were not exactly the same the same as those of the design. Also, the
Fig. 3. Measured and simulated mixing efficiency as function of flow rate.
microscope was not focused precisely at the center line in the channel when images were collected to measure the intensity. However, the experimental results show that mixing rate tended to be similar to the simulation. When the mixer was composed of only nozzles, the mixing efficiency was independent of the flow rate and almost constant at 31.6–32.5%. When only nozzles were in the mixer channel, the mixing of convective diffusion caused by the nozzles was not significant because of the distance between the nozzles and the boundary layer of the two fluids. Mixing occurred only by diffusion between different concentrations through the interface of the two fluids, even though the flow rate increased. When only micropillars were in the channel, mixing efficiency increased with the flow rate, from 59.2% to 87.8%. As the flow rate increased, the fluid velocity, in direction Y in Fig. 1, which was toward the interface between the two fluids, increased because of the pillars, causing convective diffusion. Hence, the diffusion from the concentration differences at the interface of the two fluids was amplified by the convective diffusion. The convective diffusion is dependent on flow speed, so mixing rate grows with the flow rate. Our results showed that the mixing efficiency of the micromixer composed of nozzles and pillars increased from 71.0% to 88.2% as the flow rate increased from 50 L/min to 500 L/min (Re increased from 6.65 to 66.5). When the nozzles and pillars were combined, the fluid flow vertical to the main flow direction along the channel was produced at first by the nozzles and then affected by the pillars. As a result, the vertical component of the fluid flow was enhanced toward the interface of the fluids. Therefore, the convective diffusion became strong enough to increase the mixing efficiency. Since the vertical velocity of the fluid flow increased with flow rate, the efficiency increased as well. Fig. 4 shows the simulated mixing pattern in the micromixer composed of nozzles and pillars. The laminar flows of the two injected fluids are split and recombined by the pillars and nozzles, produce more interfaces between the two fluids, and enhancing the diffusion mixing. Therefore, mixing efficiency is expected to increase by this process as well as by convective diffusion. Improved mixing efficiency was obtained by the proposed mixer but not by mixers with only nozzles or only pillars. When the flow rate was greater than 300L/min (Re = 39.9), the mixer with only pillars showed almost the same efficiency as the mixer with nozzles and pillars. However, the proposed mixer was more efficient at low flow rates than the mixers with only nozzles or pillars. The mixing efficiency at each section as 1–6 units as shown in Fig. 1 was simulated and experimentally measured when flow rate is 50, 300, and 500 mL/min for the pillar and nozzle combination
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
739
Fig. 4. Simulated mixing patterns along the channel of the micromixer.
mixer. The simulated and measured mixing efficiency is shown in Fig. 5. In Fig. 5, the measurements for each units from 1 to 6 were performed at downstream of 1.72, 3.44, 5.16, 6.88, 8.6, and 10.32 mm, respectively. Compared to the simulated results, measured mixing efficiency showed a sudden increase at the first mixer unit, however the increasing rate of the efficiency as the fluids pass the mixer channel was less than that of simulated mixing efficiency. When the flow rate was 50 L/min, the measured mixing ratio increased gently as fluids passed through each unit of the micromixer. This is because the convection is not dominant when the flow rate is low. The mixing is accomplished only by diffusion. When fluids are mixed by diffusion through the interface of the two fluids, mixing ratio increases linearly in proportion to the length of the interface. Therefore, the mixing progresses slowly as the fluids flow through the micromixer. However, when the flow rate is 300 or 500 L/min, the mixing ratio rapidly increase more than that of 50 L/min as shown in Fig. 5. This is because the convection becomes significant when the flow rate is high. The mixing accelerates as the fluid flow through the mixer channel. However the mixing efficiency does not increase linearly. The increasing ratios slow down with the mixer channel length.
Results from a previous study showed that the proposed mixer had an improved mixing performance compared to a typical modified tesla mixer and obstruction mixer [12]. The modified tesla mixer achieved ∼78% mixing at 5 mm downstream from an entrance for 100 m wide channels at Re = 0.05 [11], while we observed an ∼83% mixing rate at about 5 mm downstream, for Re = 66.5 flow. The obstruction mixer is capable of a maximum of ∼56% in the first unit of the mixer set for a 300-mm wide channel [10], while the new micromixer reached ∼64% mixing after the first unit for Re = 66.5. At higher Reynolds number (Re > ∼1), the mixing performance of the modified tesla mixer [18] and the obstruction mixer [19] is substantially improved. Simulation results of the diamond-shaped obstruction micromixer showed 70–90% mixing performance at 5 mm from the channel entrance for the 10 < Re < 100 range [19]. Recently reported passive micromixers have a mixing efficiency of 80–98% at a flow rate of 5–1200 L/min [7,20]. In comparison, the mixing efficiency of our novel micromixer was measured as 88.2% at a 500 L/min flow rate (Re = 66.5). This shows that the micromixer can be efficient even with only a simple two-dimensional structure of nozzles and pillars instead of a complex three-dimensional structure. In addition to increasing the flow rate, the mixing efficiency of the new mixer could be increased by increasing the length of the mixer channel with more nozzles and pillars. 4.2. DNA ligation by biochip
Fig. 5. Measured and simulated mixing efficiency at each unit of the pillar and nozzle combination mixer.
The minimum required ligation time in the microfluidic device was observed with 3 L of plasmids. After ligation reactions of 1, 3, and 5 min on the biochip, ligation was examined. When the ligations were performed for 1 min, a few white colonies could be found with the biochip products. However, white colonies appeared after transformation with products from ligation times of 3 or 5 min. Fig. 6 shows gel electrophoresis of plasmids from white colonies produced with products from 3 and 5 min ligation times. When the reaction time was 5 min, the 2728-bp plasmid and 440-bp HV1 insert appeared (lanes 4 and 5, markers in the left lane). However, the HV1 insert did not appear in lanes 2 and 3, and another insert appeared in lane 1. In these colonies, DNA fragments other than HV1 were inserted in the plasmid during the ligation process. Because the size of the contaminant DNA fragments was very large or very small, the bands of these fragments were not seen by gel electrophoresis. 3 min was not sufficient for ligation. Based
740
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
Fig. 6. Gel electrophoresis; lanes 1–3 are results from a 3-min reaction, lanes 4 and 5 are results from a 5-min reaction, and left column is DNA markers.
on these our results, the minimum required ligation time with the biochip was about 5 min. This is similar to other on-chip DNA ligation processes based on droplet microfluidic systems [2]. Ligation by a traditional laboratory method requires about 4 h [21]. These results show ligation of specific DNA fragments can be completed within 5 min using the biochip presented here. Some techniques, such as proximity ligation, require only 5 min for ligation with T4 DNA ligase without microfluidic mixers [22]. In the other microfluidic chip, the ligation detection was performed with a ligation mixture premixed with a polymerase chain reaction (PCR) product, and did not use flow through a mixer. This reaction was completed in 6.5 min [1]. In an advanced version of this chip, a straight channel- type of micromixer was used for mixing and the ligase reaction could be detected after 8.1 min [4]. These results imply that additional 1.6 min was spent for mixing. The total ligation time (8.1 min) is longer than our results (5 min), probably because the mixing performance of the straight channel was inferior to our micromixer. Therefore, mixing could affect and enhance the DNA ligation reaction in a totally integrated lab-on-a-chip. The ligase reaction with the same mixing procedure could be shortened to 4.1 min by optimization of reagent selections and reaction conditions [23]. The performance of the novel microfluidic device was compared to the conventional laboratory method. Products from the conventional ligation process and from the biochip with a 5-min ligation time and volume of 3 L plasmid were obtained and transferred into host cells. The cells were cultured and white colonies selected. Gel electrophoresis with plasmids from the selected cells was performed before enzymatic cutting, and the results are shown in Fig. 7a. The products (lanes 6–10 in the figure) from the biochip show the same results as the conventional method (lanes 1–5). The selected cells were cultured again in Luria–Bertani medium. Plasmid DNA was obtained from five white colonies and cut with Hind III. Gel electrophoresis was used to visualize the DNA fragments. Fig. 7b shows the results. Plasmid vectors and HV1 inserts were detected from all five white colonies. However, an unknown DNA fragment of about 800 bp was detected in one, probably from contamination during the ligation or cell culturing after the transformation. This can be minimized by carefully performing the ligation procedure. For example, after the plasmid vectors and HV1 inserts were are cut, gel-electrophoresis of the cut products and extraction of the desired DNA fragments is advised. As shown in Fig. 7b, one of the five ligations by the conventional method also failed. These results confirmed that DNA ligation was successfully performed using the novel microfluidic device. We attempted to minimize the volume of the vector in ligations with the new biochip. We tested 1, 2, and 3 L of plasmid vectors with the biochip ligation. The results are shown in Fig. 8. When 1 and 2 L vector volumes were used, other DNA fragments appeared
Fig. 7. Electrophoresis (a) after transformation and (b) after cutting with restriction enzyme; lanes 1–5 are ligation by the conventional laboratory method and lanes 6–10 are ligation by the biochip.
Fig. 8. Electrophoresis: lanes 1 and 2 are from 1 L of vector, lanes 3 and 4 are from 2 L, and lane 5 is from 5 L of vector.
in the gel electrophoresis, as shown in lanes 1–4 of Fig. 7. Reactions with 1 or 2 L plasmid volumes failed, but the ligation with 3 L produced good results when the ratio of ligase:plasmids:water in the second inlet was 4:3:1. Therefore, at least a 5-min reaction time and a 3-L vector volume are needed to perform DNA ligation with the proposed new biochip. The total volume of reagents for the ligation in the reservoirs of the new biochip was 16 L, and this was approximately the same as the 15 L of the traditional ligation process [2]. Reagent usage efficiency of the new biochip was not as high as the droplet microfluidic device, which used only 2.1 L of reagents with no waste. These results reveal a disadvantage of the PDMS–glass biochip, which is its high surface area-to-volume ratio [24]. This disadvantage is caused by the adhesion of DNA and ligase to the PDMS, and is not completely avoided by surface passivation with PVP additive. However, a novel ligation chip has been fabricated with only a simple soft-lithography technique. Hence, the chip can be made very cheaply on a mass production basis. We expect that the novel ligation chip is most suitable as a disposable unit and might be effectively applied as a point-of-care lab-on-a-chip system. 5. Conclusions A DNA ligation biochip consisting of a micromixer and microchannel reactor was inexpensively fabricated with a PDMS replica molding technique. The PDMS chip surface was coated
Y.-J. Ko et al. / Sensors and Actuators B 157 (2011) 735–741
with a 0.25 wt% PVP solution to prevent bubble generation. The micromixer had a simple structure of micronozzles and micropillars. Mixing efficiency was enhanced by convective diffusion and fluid splitting caused by the micronozzles and micropillars, and increased with flow rate. The micromixer was 10.33 mm long and had a maximum mixing rate of 87.7% at a flow rate of 500 L/min (Re = 66.5). DNA ligation was successfully accomplished in the integrated microfluidic device, with time shortened from about 4 h for the conventional laboratory technique to only 5 min using the proposed method. The ligation used 8 L ligase, 4 L HV1 insert, and 3 L plasmid vector. Although the PDMS–glass biochip rapidly completed the ligation, the biochip did not require lower amounts of reagents than conventional ligation devices. To improve the performance of the PDMS-based ligation biochip, future studies are needed to determine a technique that inhibits the interaction of the ligation reagents with the PDMS chip surface. The obtained results proved the usefulness of the in-plane passive microfluidic mixer for practical biomedical analysis. Furthermore, because this DNA ligation platform can be fabricated using polymer replica molding technologies, inexpensive disposable on-chip microfluidic systems with this PDMS-based ligation platform are also practicable. The new disposable ligation chip is expected to be useful as a totally integrated lab-on-a-chip for genetic recombination. References [1] M. Hashimoto, M.L. Hupert, M.C. Murphy, S.A. Soper, Y.-W. Cheng, F. Barany, Ligase detection reaction/hybridization assays using three-dimensional microfluidic networks for the detection of low-abundent DNA point mutations, Anal. Chem. 77 (2005) 3243–3255. [2] Y.-J. Liu, D.-J. Yao, H.-C. Lin, W.-Y. Chang, H.-Y. Chang, DNA ligation of ultramicro volume using an EWOD microfluidic system with coplanar electrodes, J. Micromech. Microeng. 18 (2008) 045017. [3] V. Hessel, H. Löwe, F. Schönfeld, Micromixers – a review on passive and active mixing principles, Chem. Eng. Sci. 60 (2005) 2479–2501. [4] M. Hashimoto, F. Barany, S.A. Soper, Polymerase chain reaction/ligase detection reaction/hybridization assays using flow-through microfluidic devices for the detection of low-abundant DNA point mutations, Biosens. Bioelectron. 21 (2006) 1915–1923. [5] L.H. Lu, K.S. Ryu, C. Liu, A magnetic microstirrer and array for microfluidic mixing, J. Microelectromech. Syst. 11 (2002) 462–469. [6] J.H. Tsai, L. Lin, Active microfluidic mixer and gas bubble filter driven by thermal bubble micropump, Sens. Actuator A: Phys. 97–98 (2002) 665–671. [7] F. Schönfeld, V. Hessel, C. Hofmann, An optimized split-and-recombine micro mixer with uniform ‘chaotic’ mixing, Lab. Chip 4 (2004) 65–69. ´ H.A. Stone, G.M. Whitesides, [8] A.D. Stroock, S.K.W. Dertinger, A. Ajdari, I. Mezic, Chaotic mixer for microchannels, Science 295 (2002) 647–651. [9] M. Elwenspoek, T.S.J. Lammerink, R. Miyake, J.H.J. Fluitman, Towards integrated microliquid handling systems, J. Micromech. Microeng. 4 (1994) 227–245. [10] H. Wang, P. Iovenitti, E. Harvey, S. Masood, Optimizing layout of obstacles for enhanced mixing in microchannels, Smart Mater. Struct. 11 (2002) 662–667. [11] A.A.S. Bhagat, I. Papautsky, Enhancing particle dispersion in a passive planar micromixer using rectangular obstacles, J. Micromech. Microeng. 18 (2008) 085005. [12] Y.-J. Ko, S.-M. Ha, H.-J. Kim, D.-H. Lee, Y. Ahn, Development of a PDMS–glass hybrid microchannel mixer composed of micropillars and micronozzles, J. Solid Mech. Mater. Eng. 2 (2008) 445–454.
741
[13] C.-H. Cho, W. Cho, Y. Ahn, S.-Y. Hwang, PDMS–glass serpentine microchannel chip for time domain PCR with bubble suppression in sample injection, J. Micromech. Microeng. 17 (2007) 1810–1817. [14] W. Cho, Y.-J. Ko, Y. Ahn, J.-Y. Yoon, N.G. Cho, Surface modification effect of wetability on the performance of a PDMS-based valveless micropump, Key Eng. Mater. 326–328 (2006) 297–300. [15] B.C. Giordano, E.R. Copeland, J.P. Landers, Towards dynamic coating of glass microchip chambers for amplifying DNA via the polymerase chain reaction, Electrophoresis 22 (2001) 334–340. [16] D.-Y. Kang, Y. Ahn, S.Y. Hwang, A study about biochip combined with micro mixer and reactor for DNA ligation, Trans. Korean Soc. Mech. Eng. A 32 (2008) 624–632. [17] N.L. Jeon, S.K.W. Dertinger, D.T. Chiu, I.S. Choi, A.D. Stroock, G.M. Whitesides, Generation of solution and surface gradients using microfluidic systems, Langmuir 16 (2000) 8311–8316. [18] C.-C. Hong, J.-W. Choi, C.H. Ahn, A novel in-plane passive microfluidic mixer with modified Tesla structures, Lab. Chip 4 (2004) 109–113. [19] A.A.S. Bhagat, E.T.K. Peterson, I. Papautsky, A passive planar micromixer with obstructions for mixing at low Reynolds number, J. Micromech. Microeng. 17 (2007) 1017–1024. [20] R.H. Liu, M.A. Stremler, K.V. Sharp, M.G. Olsen, J.G. Santiago, R.J. Adrian, J.H. Aref, D.J. Beebe, Passive mixing in a three-dimensional serpentine microchannel, J. Microelectromech. Syst. 9 (2000) 190–197. [21] J. Sambrook, D.W. Russell, Molecular Cloning, vol. 1, third ed., CSHL Press, 2001, pp. 1.84–1.87. [22] M. Gullberg, S.M. Gústafsdóttir, E. Schallmeiner, J. Jarvius, M. Bjarnegård, C. Betsholtz, U. Landegren, S. Fredriksson, Cytokine detection by antibody-based proximity ligation, Proc. Natl. Acad. Sci. U.S.A. 101 (2004) 8420–8424. [23] M. Hashimoto, F. Barany, F. Xu, S.A. Soper, Serial processing of biological reactions using flow-through microfluidic devices: coupled PCR/LDR for the detection of low-abundant DNA point mutations, Analyst 132 (2007) 913–921. [24] C. Zhan, J. Xu, W. Ma, W. Zheng, PCR microfluidic devices for DNA amplification, Biotechnol. Adv. 24 (2006) 243.
Biographies Yong-Jun Ko received a Ph.D. in mechanical engineering in 2009 from Hanyang University. He is now working at R&D Center, LG Innotek Ltd., Ansan, Korea. His main research interests include microfluidic devices, microbiosensors, microbiochip for protein and cell, plastic/glass microfabrication, and nanotechnology. Joon-Ho Maeng received his M.S. (2006) and Ph.D. degree (2009) in the Department of Biochemistry, Hanyang University, Korea. He is now a postdoctoral research fellow in the Kachon Bionano Research Institute, Kyungwon Unversity, Korea. His main research interests include biosensors, bioMEMS, point-of-care tests, nanotechnology and biomedical devices. Yoomin Ahn received his B.S. in 1984 and M.S. in 1986 from Seoul National University, Korea. He received his Ph.D. in 1992 from Purdue University, USA. He is a professor in the Department of Mechanical Engineering at Hanyang University, Korea. His present research interests include microelectromechanical systems, microfluidic biochip, and lab-on-a-chip. Seung Yong Hwang is a full professor at the Laboratory “Integrated Genomics”, Division of Molecular and Life Science at Hanyang University in Korea. He received his B.Sc. from Hanyang University in 1989 and then finished his Ph.D. in 1995 at Monash University in Australia. Later, he worked at the Stanford University in the USA as a postdoctoral fellow. He is also running a biotech company, GenoCheck Co. Ltd. His main research interests are toxicogenomics and developing biochips for various screening systems.