A Rapid Microfluidic Mixer for High-Viscosity Fluids To Track Ultrafast

Sep 28, 2012 - ABSTRACT: Tracking the folding kinetics of macromole- cules under molecular crowding conditions represents a tremendous challenge due t...
0 downloads 0 Views 2MB Size
Download PDF
Article pubs.acs.org/ac

A Rapid Microfluidic Mixer for High-Viscosity Fluids To Track Ultrafast Early Folding Kinetics of G‑Quadruplex under Molecular Crowding Conditions Ying Li, Youzhi Xu, Xiaojun Feng, and Bi-Feng Liu* Britton Chance Center for Biomedical Photonics at Wuhan National Laboratory for Optoelectronics-Hubei Bioinformatics & Molecular Imaging Key Laboratory, Systems Biology Theme, Department of Biomedical Engineering, College of Life Science and Technology, Huazhong University of Science and Technology, Wuhan 430074, China S Supporting Information *

ABSTRACT: Tracking the folding kinetics of macromolecules under molecular crowding conditions represents a tremendous challenge due to the high viscosity of the solution. In this paper, we report a unique T-type microfluidic mixer with seven consecutive ω-shaped baffles for fast mixing of high-viscosity fluids. Numerical simulations and experimental characterizations proved that the micromixer could achieve a mixing time of 579.4 μs for solutions with viscosities about 33.6 times that of pure water. Over a 1000-fold improvement in mixing dead time was accomplished in comparison to those reported previously. We further used this highly efficient micromixer to track the early folding kinetics of human telomere Gquadruplex under molecular crowding conditions. Results indicated an exponential process in the initial folding phase of Gquadruplex, and the G-quadruplex formed a more compact structure under higher degrees of molecular crowding conditions. reaching a mass concentration of 300−400 g·L −1 .14,15 Researchers commonly use 40% (w/v) PEG200 (corresponding to a concentration of 400 g·L−1) to mimic the intracellular environment.16,17 To characterize the folding kinetics of macromolecules under molecular crowding conditions in a micromixer, it is necessary to mix two 40% PEG200 solutions or an 80% PEG200 solution with a dilute sample solution to reach a final concentration of 40% PEG200.17−19 The viscosities of 40% PEG200 and 80% PEG200 are about 4 times and 20 times that of pure water at 20 °C, respectively. Thus, it is necessary to develop a fast micromixer for highviscosity fluids. However, the Reynolds number (Re) is small in the microfluidic channel, and the flow is laminar.20 The absence of turbulence makes mixing depend mainly upon molecular diffusion, leaving it a great challenge to achieve rapid mixing, especially for viscous fluids.21−24 Kane et al.21 developed a serpentine micromixer and achieved a mixing time of 200 μs. This micromixer achieved a mixing efficiency of 0.99 for dilute solution but only 0.92 for a 10% (w/w) glycerol solution with a viscosity 1.3 times that of pure water at 20 °C. Egawa et al.22 demonstrated an alcove-based micromixer with a mixing time of 22 μs. The mixing efficiency of this micromixer for dilute solution was 0.99. However, the mixing efficiency decreased to 0.9 for a 3 M guanidine solution with a viscosity 1.17 times that of pure water at 25 °C. The two mixers could achieve rapid

C

haracterization of folding kinetics is essential for obtaining insight into conformational changes of biomacromolecules.1−3 Rapid mixing can initiate reactions in a short period of time and has been recognized as an attractive approach to analyze the kinetics of fast reactions.4,5 Traditionally, kinetics investigations of protein folding are performed in stopped-flow instruments that trigger the reaction by mixing the protein solution with a new medium.6,7 Coupled with means of optical detection such as fluorescence imaging8 and circular dichroism,9 stopped-flow methods have provided valuable information on the mechanisms of protein folding. This technique is simple and reliable; however, the millisecond-scale dead time restricts its application in analyzing the kinetics of fast reactions.10 To meet the requirements of investigating folding reactions within times less than milliseconds, microfluidics-based continuous-flow methods have been proposed. 4,5,11This approach relies on the conversion between space and time performed by the fluid flow.3,10 Previously, Bilsel et al.12 demonstrated a simple continuous-flow mixer that could monitor folding reactions in the time regime of 35−1000 μs. Matsumoto et al.13 designed a micromixer with a dead time of 11 μs and observed the rapid collapse of cytochrome c induced by pH jump. In the above methods, the experiments were conducted in dilute solutions containing low concentrations of solutes, which could not represent the actual intracellular conditions. The aqueous environment in living cells is highly crowded, with biomolecules occupying 30−40% of the cellular volume and © 2012 American Chemical Society

Received: May 15, 2012 Accepted: September 28, 2012 Published: September 28, 2012 9025

dx.doi.org/10.1021/ac301864r | Anal. Chem. 2012, 84, 9025−9032

Analytical Chemistry

Article

Chip Fabrication. The micromixer was fabricated according to the rapid prototyping method previously reported.28 In brief, the mold for the microstructure was fabricated by using the standard soft-lithography technique with a negative photoresist SU-8 2100 (Gersteltec Sarl, Switzerland) on a silicon wafer n-type ⟨100⟩. The poly(dimethylsiloxane) (PDMS) layer, made from a mixture of 10:1 (m/m) PDMS and curing agent (Sylgard 184, Dow Corning, USA), was obtained by molding the SU-8 structure. The PDMS sheet was then cut and peeled from the mold. The patterned PDMS sheet was punched at the end of each microchannel to produce 0.7 mm through-membrane holes. After treatment with oxygen plasma, the PDMS was irreversibly bonded to a glass slide with holes drilled by a numerically controlled machine. The microchip was then assembled on an organic glass clamp to complete the instrument. (It was found that the assembled instrument used here was more stable and durable than the microchip fabricated in the conventional way.28) Materials and Sample Preparations. Chemicals such as methanol, KI, Na2B4O7·10H2O, fluorescein, sulforhodamine B, HCl, PEG200, EGTA, Tris, hydroxyethylcellulose (HEC), and glycerol were purchased from Sinopharm Chemical Reagent (Shanghai, China). The oligonucleotide d(TTAGGG)4 , purchased from TaKaRa Biotech (Dalian, China), was labeled with 5′-fluorescein (FAM) and 3′-tetramethylrhodamine (TMR) and purified by HPLC. Stock solutions of 1 mM fluorescein and 1 mM sulforhodamine B were dissolved in methanol and water, respectively. Low concentrations of fluorescein and sulforhodamine B were diluted with 0.1 mol·L−1 borate buffer (pH 11.0) from the stock solutions before experiments. Solutions of 0.5 M KI, various concentrations of HEC (containing 1 μM fluorescein), and glycerol (containing 1 μM fluorescein) were dissolved in 0.1 M borate buffer (pH 11.0). Various concentrations of PEG200 were dissolved in the buffer containing 10 mM Tris-HCl, pH 7.4, 1 mM EDTA. The oligonucleotide d(TTAGGG)4 was dissolved in the Tris-HCl buffer with a concentration of 1 μM (oligonucleotide concentrations were determined from their absorbance at 260 nm) and then heated at 95 °C for 5 min and slowly cooled to room temperature (20 °C) before analysis. All solutions were prepared with water purified by the DirectQ system (Millipore, Bedford, MA) and filtered with 0.45 μm syringe filters before use. Optical Imaging System and Operation Procedures. A laser scanning confocal microscope (LSCM, FV1000, Olympus, Japan) was used as the detection platform. An argon laser (λ = 488 nm) and a He−Ne laser (λ = 543 nm) were used as light source. A 10×/NA 0.3 objective was used with a spatial resolution of 1 μm along the z-axis. The exposure time of each pixel was 10 μs. Fluorescein was excited with 488 nm and the emission fluorescence was collected from 510 to 550 nm, while sulforhodamine B was excited with 543 nm and the emission fluorescence was collected from 575 to 615 nm. The oligonucleotide with 5′-FAM and 3′-TMR was excited with 488 nm, and the emission fluorescence was collected at 510− 550 nm for FAM and 575−615 nm for TMR. The mixing efficiency of the ω micromixer was first evaluated by mixing of two dilute solutions (aqueous solutions of 1 μM sulforhodamine B and 1 μM fluorescein). The mixer was then characterized by mixing a low-viscosity solution (1 μM sulforhodamine B aqueous solution) and a high-viscosity solution (various concentrations of glycerol, HEC, and

mixing, but the results indicated that the mixing efficiency decreased drastically for solutions with viscosities of less than 1.5 times that of pure water. As mentioned in both previous papers, high-viscosity solutions make it hard to achieve good mixing, and the mixing efficiency may fall below 0.9 (a mixing efficiency of 0.9 is considered to be complete mixing22,25) for solutions with higher viscosity. To enhance the mixing efficiency of high-viscosity fluids, Wang et al.24 demonstrated a micromixer with acoustically induced bubbles. Their results showed that the water−glycerol solutions with various viscosities could achieve good mixing efficiency. However, several seconds was needed to reach complete mixing for the water−50% (w/w) glycerol solution with a viscosity of 5.69 mPa·s (∼6.4 times that of pure water at 25 °C). Xia et al.23 reported a mixer for fluids with large viscosity contrast using splitting-and-recombination structures. The mixer was composed of two layers and contained an interconnected multichannel network. This mixer worked well for solutions with viscosity ratio of up to the order of 104, but the mixing time was relatively long (>6 s). The mixing times of those reported micromixers are not rapid enough to meet the requirements of tracking the early folding kinetics of nuclear acids or proteins under high-viscosity conditions. It is an urgent task to develop new microfluidic mixers that can resolve the initial kinetics of ultrafast biochemical reactions. In this paper, we report a new continuous-flow micromixer with seven ω-shaped baffles in the T-channel. The mixing efficiency of this ω micromixer was first evaluated using numerical simulation. Experimental characterization of the micromixer further demonstrated that complete mixing of solutions with various viscosities could be realized. For a solution with viscosity of up to 35.25 mPa·s (∼33.6 times that of pure water), the micromixer achieved complete mixing in 579.4 μs, which represented a mixing speed about 1000-fold faster than those reported previously. Based on this advancement, the folding kinetics of human telomere G-quadruplex under molecular crowding conditions was investigated. It was found that the denatured d(TTAGGG)4 sequence formed a more compact structure under higher degree of molecular crowding conditions. An exponential process on the folding pathway of the G-quadruplex under molecular crowding conditions was discovered. The proposed ω micromixer offers unique advantages of low sample consumption and extremely high mixing efficiency for high-viscosity fluids, making it an ideal tool for characterizing folding kinetics of macromolecules under molecular crowding conditions.

■

EXPERIMENTAL SECTION Computational Fluid Dynamics (CFD) Simulations. Numerical simulations were carried out to optimize the structure of the ω micromixer by using the simulation software Fluent 6.1. The mixing efficiency was investigated using a 3D finite volume model. The simulation procedures were performed by solving the continuity equation, the Navier− Stokes equation, and the diffusion−convection equation, similar to that described by Mengeaud et al.26,27 For the two inlets, one fluid was pure water and the other fluid was glycerol with various concentrations. The properties of the two solutions were set as follows: density of water (ρ1), 103 kg·m−3; viscosity of water (μ1), 10−3 Pa·s, density of glycerol (ρ2), 1.26 × 103 kg·m−3; viscosity of glycerol (μ2), 0.799 Pa·s. The criterion for convergence was for the increment in each variable to fall below 1 × 10−5. 9026

dx.doi.org/10.1021/ac301864r | Anal. Chem. 2012, 84, 9025−9032

Analytical Chemistry

Article

Figure 1. Simulation result of the ω micromixer. (a) Mixing of dilute solutions at flow rate of 0.22 mL/min (Re = 64.48). The micromixer was composed of two inlets, a mixing channel, and an observation channel. The white dotted lines of f1−f8, the yellow dotted rectangular box of the mixing channel, and the red dotted rectangular box of the region of interest (ROI) show the positions used for later calculations. (b,c) Velocity vector distribution at the third and fourth chambers. (d) Mixing efficiency of water and various concentrations of glycerol at f1−f8 at a flow rate of 0.25 mL/min (Re = 73.27).

PEG200 containing 1 μM fluorescein). The G-quadruplex formation was further monitored by fluorescence resonant energy transfer (FRET) through the mixing of 1 μM oligonucleotide solution and various concentrations of PEG200. The 5′-FAM was used as donor and 3′-TMR as acceptor. In the experiment, a dilute solution and the other relatively viscous solution were injected into the microfluidic mixer through the two inlets, respectively. The two solutions were supplied with identical volumetric flow rate by a syringe pump (KDScientific, USA). Acquired images were further analyzed using Image Pro Plus 6.0, Matlab 7.0, and Origin 7.5.

baffles were positioned at the two sides of the channel, and the other five baffles were positioned in the center of the mixing channel (this design could achieve a better mixing efficiency than the structure with all seven ω-shaped baffles in the center of the mixing channel, see Figure S2). To demonstrate the mixing efficiency of the optimized micromixer, a dilute solution (water) and a series of glycerol solutions (concentrations from 0% to 100% (w/v), where 0% glycerol is simply water) were used. The Reynolds number is defined as

Re = uD ̅ h /v where u̅ is the average velocity in the channel, Dh the hydraulic diameter, and v, the kinetic viscosity. The result of mixing two dilute solutions (water and water) at a flow rate of 0.22 mL/min (Re = 64.48) is shown in Figure 1a. When the two solutions collided at the intersection of the T-channel, a clear fluid interface could be seen. After the solutions flowed through the first ω-baffle, mixing began to occur and the interface became foggy. After flowing through the third and forth mixing chamber, the fluid got more and more homogeneous. At the end of the sixth mixing chamber, the result showed that the two solutions had achieved complete mixing here. Figure 1b,c illustrates the velocity vector

■

RESULTS AND DISCUSSION CFD Simulations. A numerical study was conducted in this work to search for an optimal design for fast mixing, and we arrived at the configuration shown in Figure 1a, with two inlets (inlet a and inlet b), a mixing channel (containing seven mixing chambers), and an observation channel. There were seven asymmetric ω-shaped baffles in the mixing chambers (a symmetric structure in our former design could not achieve good mixing efficiency, see Figure S1). The first two ω-shaped 9027

dx.doi.org/10.1021/ac301864r | Anal. Chem. 2012, 84, 9025−9032

Analytical Chemistry

Article

Figure 2. Manufacture of the micromixer. (a) SEM images of the fabricated PDMS structures. The left inset shows the height of the channel (27 μm), and the right inset shows as magnification of the fifth and sixth chambers of the channel. (b) Schematic of the assembled micromixer instrument: 1, inlet a; 2, inlet b; 3, outlet; 4, set screws; 5, top clamp; 6, seal washers; 7, slide glass; 8, PDMS sheet; 9, bottom clamp.

Figure 3. Evaluation of the mixing efficiency of dilute solutions (fluorescein and sulforhodamine B). (a−d) Fluorescence distribution of the middle layer given by LSCM along the Z-axis at flow rates of (a) 0.003, (b) 0.03, (c) 0.1, and (d) 0.22 mL/min. The insets are the reconstructed cross sections at position f8. (e) Fluorescence distribution across the channel at position f8 under various flow rates. (f) Mixing efficiency at position f8 under various flow rates.

distribution at the third and fourth chambers. Two solutions were forced to flow through a narrow channel with a high velocity, which decreased the diffusion distance efficiently. The solutions then flowed into an expanded chamber embedded in an asymmetrical ω-baffle. Such structure could change the flow direction, disturb the fluid, and generate sufficient fluid mixing. Figure 1d shows the mixing efficiency (Cm) of various concentrations of water−glycerol at a flow rate of 0.25 mL/ min (Re = 73.27). Here Cm was calculated according to26,29

Cm = 1 −

∑ (Xi − X̅ )2 /N X̅

where Xi is the intensity of each point in the cross section, N the total number of points, and X̅ the average intensity of all the points. The value of Cm ranged between minimum 0 and maximum 1, and larger Cm indicated better mixing efficiency. As shown in Figure 1d, two dilute solutions achieved the best mixing efficiency (Cm = 0.965). When the concentration of 9028

dx.doi.org/10.1021/ac301864r | Anal. Chem. 2012, 84, 9025−9032

Analytical Chemistry

Article

Figure 4. Evaluation of the mixing efficiency for viscous solutions. (a) Mixing efficiency of buffer−glycerol solutions at various flow rates. (b) Mixing efficiency of buffer−HEC solutions at positions f1−f8 at a flow rate of 0.25 mL/min. (c) Mixing efficiency of buffer and various concentrations of PEG200 at a flow rate of 0.25 mL/min. (d) Surface plot of the mixing efficiency of buffer−70% PEG200 as a function of flow rate and the position along the outlet channel.

disturbance of the fluid was enhanced tempestuously, and complete mixing was achieved at the exit of the mixing channel. The insets in Figure 3a−d show the reconstructed cross sections (y−z plane) at position f8. This result indicated similar fluorescence distribution from one slice to the next along the zaxis. Figure 3e,f shows the fluorescence distribution and mixing efficiency at position f8 under various flow rates, which also demonstrated that homogeneous fluorescence in the observation channel was obtained as the flow rate increased to 0.22 mL/min. To further examine the mixing efficiency for high-viscosity fluids, three viscous solutions (glycerol, HEC, and PEG200) were used as samples. The viscosities of each solution are shown in Table S1. Glycerol with concentrations of 63% and 80% (w/v) was respectively mixed with the borate buffer at various flow rates. The mixing efficiency is shown in Figure 4a. This result indicated that both of the glycerol solutions achieved complete mixing at a flow rate of 0.25 mL/min, and the 63% glycerol reached a better mixing efficiency than the 80% glycerol, which fits well with the simulation result in Figure 2d. HEC solutions with viscosities of 8.95 and 35.25 mPa·s were respectively mixed with the borate buffer at a flow rate of 0.25 mL/min. The mixing efficiency at positions f1−f8 is shown in Figure 4b, which demonstrates that Cm could reach about 0.91 for the solution with viscosity of up to 35.25 mPa·s (∼33.6 times that of pure water at 20 °C).

glycerol increased from 40% to 89%, Cm decreased from 0.955 to 0.908 (still above 0.9). While the mass fraction of glycerol was higher than 90%, Cm fell below 0.9, but the solution of 100% glycerol could still reach a Cm of 0.84. This result indicates that the mixing efficiency is inversely proportional to the viscosity of the solutions (higher glycerol concentration means higher viscosity). Evaluation of the Micromixer. To examine the performance of the new ω micromixer, a microfluidic chip was fabricated according to the optimal simulation structure. Scanning elecrtron microscopy (SEM) images of the fabricated PDMS structures and a schematic of the final assembled instrument are shown in Figure 2. Two dilute solutions (sulforhodamine B and fluorescein) were mixed in the chip with various flow rates from 0.003 mL/min (Re = 0.88) to 0.25 mL/min (Re = 73.27). The fluorescence distribution of the middle layer, given by LSCM along the Z-axis, is shown in Figure 3a−d. When the flow rate was 0.003 mL/min (Re = 0.88), the two solutions flowed through the observation channel with a clear fluid interface. As the flow rate increased to 0.03 mL/min (Re = 8.80), the embedded asymmetrical ω structure disturbed the fluid, and the two solutions began to mix. After the flow rate further increased to 0.1 mL/min (Re = 29.32), the ω-baffles agitated the fluid and many interfaces arose between the two solutions, which decreased the diffusion distance efficiently and evidently accelerated the mixing. When the flow rate rose to 0.22 mL/min (Re = 64.48), the 9029

dx.doi.org/10.1021/ac301864r | Anal. Chem. 2012, 84, 9025−9032

Analytical Chemistry

Article

Figure 5. Mixing time of the ω micromixer (the far left wall of the microchannel shown in Figure 1a was defined as time zero for mixing). (a) Mixing time for dilute solutions (fluorescein and sulforhodamine B). Complete mixing was achieved at position f6 under a flow rate of 0.25 mL/min, and the mixing time was calculated to be 502.1 μs for dilute solutions. (b,c) Mixing time for viscous solutions: (b) 80% glycerol and (c) 2% HEC. Complete mixing was achieved at position f8 under a flow rate of 0.25 mL/min, corresponding to a mixing time of 579.4 μs.

t = Vmix /F

A series of 40%, 50%, 60%, 70%, and 80% (w/v) PEG200 solutions were respectively mixed with Tris-HCl buffer at a flow rate of 0.25 mL/min. The mixing efficiency at position f8 is shown in Figure 4c. The mixing profile of 70% PEG200 is shown in Figure 4d. These results indicate that complete mixing was achieved for each PEG200 solution. Mixing Time of the ω Mixer. The mixing time was calculated using the method reported by Egawa et al.22 The sample flow rate was 0.25 mL/min (Re = 73.27) for both of the inlets. Time zero for mixing was set at the position where the two solutions met (the far left wall of the microchannel as shown in Figure 1a). According to the previous report,22,25 Cm = 0.9 was defined as the position where the solutions achieved complete mixing (the mixing end point). As shown in Figure 5a, the dilute solutions (fluorescein and sulforhodamine B) achieved complete mixing at position f6, while solutions with relatively high viscosities (80% glycerol and borate buffer, 2% HEC and borate buffer) showed homogeneous fluorescence at position f8. Thus the mixing end point was set at positions f6 and f8 for dilute solutions and viscous solutions, respectively. The mixing time was determined as22

where t is the mixing time, Vmix the volume of the mixing channel from the time zero for mixing to the mixing end point (Vmix = 4.184 × 10−12 and 4.828 × 10−12 m3 for dilute solutions and viscous solutions, respectively), and F the total flow rate of the two inlets (0.5 mL/min). The mixing time of this ω micromixer was calculated to be 502.1 μs for dilute solutions and 579.4 μs for viscous solutions, respectively. Thus, the device achieved a sub-millisecond mixing time for such highviscosity fluids (>30 mPa·s), an improvement of about 3 orders of magnitude over the mixers previously reported.23,24 Quench Reaction. The quench reaction of a fluorescent dye by iodide ions is often used as a characterization technique to investigate the mixing efficiency.22,29 To confirm the mixing efficiency of the ω micromixer, HEC (containing 1 μM fluorescein) with viscosity of 8.95 mPa·s and KI were injected into the two inlets with a flow rate of 0.25 mL/min. The fluorescence quenching was analyzed using the Stern−Volmer equation,30 9030

dx.doi.org/10.1021/ac301864r | Anal. Chem. 2012, 84, 9025−9032

Analytical Chemistry

Article

Figure 6. Folding kinetics analysis of human telomere G-quadruplex. (a) Folding kinetics of the G-quadruplex under molecular crowding conditions (the final concentrations of PEG200 were 0, 20%, 30%, and 40%, respectively). The data were exacted from the ROI as shown in Figure 1a. No data were recorded during the mixing dead time of 579.4 μs. The red, green, and blue lines are fits of the data to single-exponential curves. (b) FRET efficiency of the G-quadruplex under various concentrations of PEG200.

I0/I = 1 + KSV[Q]

I=

where I0 and I are the fluorescence intensities in the absence and in the presence of quencher, respectively, KSV is the Stern− Volmer constant (9.608 ± 0.273), and [Q] is the concentration of iodide in the solution. According to the Stern−Volmer equation, 70% quenching of the fluorescence from fluorescein was expected upon complete mixing with 0.5 M KI applied.22,30 For the presented micromixer, the homogeneous fluorescence at position f8 (I = 4108.5) was about 30% of the initial fluorescence (I0 = 13752.5) at the inlet, which confirmed that the two solutions achieved complete mixing after flowing through the mixing channel. Folding Kinetics of Human Telomere Oligonucleotide d(TTAGGG)4. The G-rich strand of human telomere DNA sequence d(TTAGGG)4, an structure essential to aging and cancer therapy,31,32 could fold into a G-quadruplex structure under molecular crowding conditions in the absence of salt16,17 like in dilute solutions with monovalent cations.33,34 The previous work was mostly conducted in 40% (w/v) PEG200 (corresponding to a mass concentration of 400 g·L−1) to mimic the crowding conditions inside cells.16,17 However, that work mainly focused on finding new conformations or confirming the existing conformations of G-quadruplex.16,17,35 Few studies about the folding kinetics of G-quadruplex under molecular crowding conditions have been reported. Xue et al.18 studied the kinetics under molecular crowding conditions through manual mixing, but this strategy could only investigate the kinetics of G-quadruplex formation on the time scale of minutes to hours. Thus, characterization of the early folding kinetics (within the time scale of microseconds to milliseconds) of G-quadruplex under molecular crowding conditions still remains a challenge. To investigate the early events of the G-quadruplex folding process, the Tris-HCl buffer and various concentrations (40%, 60%, and 80%) of PEG200 were mixed with 1 μM oligonucleotide d(TTAGGG)4 in the ω mixer at a flow rate of 0.25 mL/min. Thus, the final concentration of PEG200 after mixing was 0, 20%, 30%, and 40%, respectively. Fluorescence images of the donor and acceptor were snapped simultaneously using LSCM. The FRET efficiency was expressed as proximity ratio I:36

Ia Ia + Id

where Ia and Id are the fluorescence intensities of the acceptor (TMR) and the donor (FAM), respectively. To track the kinetic process of the G-quadruplex folding, the fluorescence of the central 10 pixels of the observation channel (the region of interest, ROI, as shown in Figure 1a) was extracted and averaged from the analyzed data. The FRET efficiency was plotted versus the Lagrangian time coordinate, converting from the Eulerian space coordinate,3 which is shown in Figure 6a. Since the flow lines were chaotic in the mixing region, no data were recorded during the mixing dead time of 579.4 μs. The first measured point was the entrance of the observation channel (position f8 shown in Figure 1a). In the Tris-HCl buffer, the FRET efficiency was very low and remained at roughly the same value, indicating an unfolded oligonucleotide in this condition. In the presence of PEG200, the FRET efficiency increased as the concentration increased from 20% to 40% (Figure 6b), which revealed that the Gquadruplex formed a more compact structure under higher degrees of molecular crowding (support for this conclusion is shown in Figures S3 and S4), as previously reported,35 because the FRET efficiency of the sample was inversely dependent on the distance between the two fluorophores. Previous kinetic studies revealed that telomere DNA in K+/ Na+ solution can fold into a G-quadruplex on a microsecond to millisecond time scale,2,18,37,38 which is often followed by additional isomerization on a time scale ranging from seconds to hundreds of seconds. As shown in Figure 6a, a variational FRET efficiency during the folding process was monitored, and the plotted curves could be divided into a lag phase and an exponential rise phase. The exponential phase represented the folding process of the oligonucleotide. The presence of the lag phase before the exponential phase seemed very interesting. The lag phase could sometimes be observed kinetically in the early folding stages of biomacromolecules, such as proteins,39 although this phenomenon was rarely reported previously. We inferred the lag phase might represent the adjustment of oligonucleotides in emerging molecular crowding conditions. Higher concentration of PEG200 may increase the collision probability between molecules and help the adjustment process. Thus, the DNA molecular could achieve the favorable shape in less time (resulting in a shorter length of lag phase, as shown in 9031

dx.doi.org/10.1021/ac301864r | Anal. Chem. 2012, 84, 9025−9032

Analytical Chemistry

Article

(7) Russell, R.; Millett, I. S.; Tate, M. W.; Kwok, L. W.; Nakatani, B.; Gruner, S. M.; Mochrie, S. G.; Pande, V.; Doniach, S.; Herschlag, D.; Pollack, L. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 4266−4271. (8) Christensen, S.; Sottrup-Jensen, L.; Christensen, U. Biochem. J. 1995, 305, 97−102. (9) Borucki, B.; Otto, H.; Rottwinkel, G.; Hughes, J.; Heyn, M. P.; Lamparter, T. Biochemistry 2003, 42, 13684−13697. (10) Roder, H.; Maki, K.; Cheng, H. Chem. Rev. 2006, 106, 1836− 1861. (11) Shastry, M. C. R.; Sauder, J. M.; Roder, H. Acc. Chem. Res. 1998, 31, 717−725. (12) Bilsel, O.; Kayatekin, C.; Wallace, L. A.; Matthews, C. R. Rev. Sci. Instrum. 2005, 76, 0143021−0143027. (13) Matsumoto, S.; Yane, A.; Nakashima, S.; Hashida, M.; Fujita, M.; Goto, Y.; Takahashi, S. J. Am. Chem. Soc. 2007, 129, 3840−3841. (14) Zimmerman, S. B.; Trach, S. O. J. Mol. Biol. 1991, 222, 599− 620. (15) Ellis, R. J. Trends Biochem. Sci. 2001, 26, 597−604. (16) Miyoshi, D.; Nakao, A.; Sugimoto, N. J. Biochem. 2002, 41, 15017−15024. (17) Kan, Z. Y.; Yao, Y. A.; Wang, P.; Li, X. H.; Hao, Y. H.; Tan, Z. Angew. Chem., Int. Ed. 2006, 45, 1629−1632. (18) Xue, Y.; Liu, J. Q.; Zheng, K. W.; Kan, Z. Y.; Hao, Y. H.; Tan, Z. Angew. Chem., Int. Ed. 2011, 50, 8046−8050. (19) Xue, Y.; Kan, Z. Y.; Wang, Q.; Yao, Y.; Liu, J.; Hao, Y. H.; Tan, Z. J. Am. Chem. Soc. 2007, 129, 11185−11191. (20) Stroock, A. D.; Dertinger, S. K. W.; Ajdari, A.; Mezic, I.; Stone, H. A.; Whitesides, G. M. Science 2002, 295, 647−651. (21) Kane, A. S.; Hoffmann, A.; Baumgartel, P.; Seckler, R.; Reichardt, G.; Horsley, D. A.; Schuler, B.; Bakajin, O. Anal. Chem. 2008, 80, 9534−9541. (22) Egawa, T.; Durand, J. L.; Hayden, E. Y.; Rousseau, D. L.; Yeh, S. R. Anal. Chem. 2009, 81, 1622−1627. (23) Xia, H. M.; Wang, Z. P.; Koh, Y. X.; May, K. T. Lab Chip 2010, 10, 1712−1716. (24) Wang, S. S.; Huang, X. Y.; Yang, C. Lab Chip 2011, 11, 2081− 2087. (25) Chung, C.; Shih, T. Microfluid. Nanofluid. 2008, 4, 419−425. (26) Lin, Y. C.; Chung, Y. C.; Wu, C. Y. Biomed. Microdevices 2007, 9, 215−221. (27) Mengeaud, V.; Josserand, J.; Girault, H. H. Anal. Chem. 2002, 74, 4279−4286. (28) Duffy, D. C.; McDonald, J. C.; Schueller, O. J.; Whitesides, G. M. Anal. Chem. 1998, 70, 4974−4984. (29) Li, Y.; Zhang, D.; Feng, X.; Xu, Y.; Liu, B. F. Talanta 2012, 88, 175−180. (30) Albani, J. R. Biochim. Biophys. Acta 1998, 1425, 405−410. (31) Jing, N.; Li, Y.; Xiong, W.; Sha, W.; Jing, L.; Tweardy, D. J. Cancer Res. 2004, 64, 6603−6609. (32) Chang, C. C.; Kuo, I. C.; Ling, I. F.; Chen, C. T.; Chen, H. C.; Lou, P. J.; Lin, J. J.; Chang, T. C. Anal. Chem. 2004, 76, 4490−4494. (33) Ambrus, A.; Chen, D.; Dai, J.; Bialis, T.; Jones, R. A.; Yang, D. Nucleic Acids Res. 2006, 34, 2723−2735. (34) Balagurumoorthy, P.; Brahmachari, S. K.; Mohanty, D.; Bansal, M.; Sasisekharan, V. Nucleic Acids Res. 1992, 20, 4061−4067. (35) Zhou, J.; Wei, C.; Jia, G.; Wang, X.; Tang, Q.; Feng, Z.; Li, C. Biophys. Chem. 2008, 136, 124−127. (36) Green, J. J.; Ying, L.; Klenerman, D.; Balasubramanian, S. J. Am. Chem. Soc. 2003, 125, 3763−3767. (37) Gray, R. D.; Li, J.; Chaires, J. B. J. Phys. Chem. B 2009, 113, 2676−2683. (38) Ida, R.; Wu, G. J. Am. Chem. Soc. 2008, 130, 3590−3602. (39) Walkenhorst, W. F.; Green, S. M.; Roder, H. Biochemistry 1997, 36, 5795−5805.

Figure 6a), and the folding reaction was initiated (with an exponential rise FRET efficiency) more rapidly. The rise FRET efficiency in the plotted curves could be fitted by single exponentials as expected for the pseudo-first-order kinetics. The observed kinetics in 20%, 30%, and 40% PEG200 solutions were (1.796 ± 0.061) × 104, (2.009 ± 0.046) × 104, and (2.823 ± 0.045) × 104 s−1, respectively. Due to its good mixing efficiency for high-viscosity fluids, this micromixer could rapidly initiate the folding reaction of G-quadruplex under molecular crowding conditions, allowing the investigation of the kinetics at the sub-millisecond level.

■

CONCLUSIONS In this paper, we designed a ω micromixer and confirmed the mixing efficiency using numerical simulation and experimental evaluation. This mixer achieved complete mixing within 579.4 μs for a solution with viscosity of up to ∼33.6 times that of pure water. Over 1000-fold improvement in the mixing time was accomplished in comparison to previous reports. Further, we utilized this micromixer to examine the folding kinetics of human telomere G-quadruplex d(TTAGGG)4 under molecular crowding conditions. The result indicated that the Gquadruplex formed more compact structures under higher degrees of molecular crowding conditions and there existed an exponential process in the initial folding phase of G-quadruplex. As the proposed ω micromixer has advantages of simple structure, ease of fabrication, and short mixing time for highviscosity fluids, it will open a new window for analyzing biomolecules’ folding kinetics under molecular crowding conditions. Additionally, wide application of this micromixer in the fields of chemical synthesis and polymer formulations can also be expected.

■

ASSOCIATED CONTENT

S Supporting Information *

Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: bfl[email protected]. Tel.: +86-27-87792203. Fax: +86-27-87792170. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors gratefully acknowledge financial support from the National Basic Research Program of China (2011CB910403) and the National Natural Science Foundation of China (30970692, 21075045).

■

REFERENCES

(1) Gambin, Y.; Simonnet, C.; VanDelinder, V.; Deniz, A.; Groisman, A. Lab Chip 2010, 10, 598−609. (2) Gray, R. D.; Chaires, J. B. Nucleic Acids Res. 2008, 36, 4191− 4203. (3) Yao, S.; Bakajin, O. Anal. Chem. 2007, 79, 5753−5759. (4) Hertzog, D. E.; Ivorra, B.; Mohammadi, B.; Bakajin, O.; Santiago, J. G. Anal. Chem. 2006, 78, 4299−4306. (5) Gambin, Y.; VanDelinder, V.; Ferreon, A. C.; Lemke, E. A.; Groisman, A.; Deniz, A. A. Nat. Methods 2011, 8, 239−241. (6) Kiefhaber, T. Proc. Natl. Acad. Sci. U.S.A. 1995, 92, 9029−9033. 9032

dx.doi.org/10.1021/ac301864r | Anal. Chem. 2012, 84, 9025−9032