Article pubs.acs.org/ac
Compartmentalization of Electrophoretically Separated Analytes in a Multiphase Microfluidic Platform Mark C. Draper,† Xize Niu,‡ Soongwon Cho,† David I. James,† and Joshua B. Edel*,† †
Department of Chemistry, Imperial College London, Exhibition Road, South Kensington, London, SW7 2AZ, United Kingdom Engineering and the Environment, and Institute for Life Sciences, University of Southampton, Highfield, Southampton, SO17 1BJ, United Kingdom
‡
ABSTRACT: Herein, we describe the monolithic integration of a multiphase microfluidic system to a microcapillary gel electrophoresis (μCGE) architecture for the complete isolation and storage of separated analyte bands. Within this platform, analyte molecules are separated using microchannel gel electrophoresis, and the eluted bands are stored in a sequence of approximately 40−600 encapsulating microdroplets. Importantly, employing such a system allows for total control of droplet size, shape, and composition. This approach is utilized to separate, optically detect, and encapsulate two fluorescent analytes from a composite sample mixture. Further to this, we subsequently investigate the potential of the system to be used as a concentration gradient generator through analysis of the segmented analyte bands and droplet composition.
S
ince its genesis in 1937 through the work of Arne Tiselius et al.,1 capillary electrophoresis (CE) has constantly evolved and been refined through methods including but not limited to capillary zone electrophoresis (CZE),2,3 isotachophoresis (ITP),4 micellar electrokinetic chromatography (MEKC),5 and capillary gel electrophoresis (CGE).6 Furthermore, with the advent of micro total analytical systems in the early 1990s, CE was introduced to small scale, on chip applications designed to take advantage of the reduction in sample volumes and separation times required.7 This, alongside recent advances in microfabrication,8 has facilitated production of cheap, reproducible microchips with micrometer scale resolution.9 The integration of well-proven chemical detection techniques into microfluidic designs manifested a rapid growth in multidisciplinary μCE research across the fields of genomics,10 proteomics,11 chemical analysis,12 and many others.13−15 One of the main proponent factors in the growth of μCE is the systematic ability of the process to handle significantly lower sample volumes with a higher theoretical plate count. This provides more efficient separations than traditional methods such as liquid chromatography (LC) or even more conventional CE.16 In an ideal separation protocol, a sample would enter the system in a mixed form and that solution would be subsequently separated, quantitatively detected, and removed, while retaining the individual band resolution in a form suitable for further analysis. With common microscale separation sample volumes existing in the pico/femtoliter17,18 range, it has proved difficult to experimentally confine the separated sample analyte bands without effects such as Taylor dispersion becoming a dominant factor. Many novel solutions have been © 2012 American Chemical Society
employed to address this problem with varying degrees of success. For example, elastomeric valves incorporating subnanoliter chambers,19 droplet electrohydrodynamics,20 and electrokinetic sample concentration and confinement.21,22 One particularly elegant approach applied to this problem is the methodology employed by Edgar and colleagues,23 whereby separated analytes are electroosmotically compartmentalized within the separation medium. The application of a voltage gradient leads to droplets being created at an interface with an immiscible carrier flow through the pinching of droplets upon movement in the separation medium caused by the high electroosmotic flow (EOF). Following on from this, Niu et al.24 demonstrated that compartmentalized droplets from liquid chromatography eluent can be further analyzed with a second dimensional CE separation, thus demonstrating the potential for further analysis of analytes isolated within droplets. While current methods allow for the encapsulation of separated analytes from CZE, the efficiency of the separation and the throughput of the device are limited by the inherent EOF present within the system. Adopting a different approach, we apply a capillary gel matrix as a separation buffer, enabling the system to take advantage of the benefits of a common CGE system, namely, the suppression of both the systemic electroosmotic buffer flow25 and also the longitudinal molecular diffusion in the separation channel.26 The added viscosity of the gel also minimizes pressure fluctuations within the channel. The Received: April 30, 2012 Accepted: June 1, 2012 Published: June 1, 2012 5801
dx.doi.org/10.1021/ac301141x | Anal. Chem. 2012, 84, 5801−5808
Analytical Chemistry
Article
Figure 1. Schematic outline of the CGE/Droplet coupled device. Separation channel length from point of injection to electrode ≈3 cm. Channel widths vary from 25 μm (separation phase) to 50 μm (droplet phase) with the height being held constant at 40 μm. (a) Images of the 3-stage electrokinetic injection process, (b) droplet output channel, and (c) the gel-carrier flow boundary, using a 0.1× TBE carrier doped with 0.1 mM Fluorescein are shown.
(PDMS) (Dow Corning, UK) using well documented soft lithographic techniques.28,29 Briefly, the microfluidic device was designed in AutoCAD 2011 (AutoDesk) and printed as a dark field mask (Circuit Graphics, UK). This pattern was subsequently transmitted to a 4 in. silicon wafer (Active Business Company) as a negative mask using a 1:1 contact photolithography process in combination with SU8-50 negative photoresist (Microchem). PDMS was subsequently cast onto the fabricated master and cured overnight on a hot plate at 65 degrees. Before sealing to a glass coverslide, inlets and outlets are created using disposable 1 mm biopsy punches (KAI Medical). Glass slides are initially cleaned in a freshly prepared piranha solution, to remove organic contaminants, before being sonicated in a 0.5 M NaOH solution to remove any unwanted inorganic molecules. (Caution: Piranha etch solution reacts violently with organic materials and should be handled with extreme caution.) Patterned electrodes were fabricated on the cleaned slides through use of a lift-off process, whereby first a positive photoresist (AZ-4562, Microchem) of approximately 8 μm thickness is spun onto a clean glass slide. Following a short prebake, 1 min per μm of photoresist thickness, a darkfield mask, designed and used as above, is utilized to cross-link the photoresist surrounding the desired area through UV exposure. After exposure, the slide is developed (AZ 400K Developer, Microchem), before a 120 nm coating of Platinum is sputtered (Quorum) on top, with selected sections subsequently being removed via brief sonication in acetone. The PDMS channel is subsequently bonded to the glass substrate through the application of an air plasma to both bonding faces for a duration of 30 s before being aligned to the separation channel. The patterned electrode has a consistent thickness of approximately 120 nm with a width of 100 um, thus giving a surface area of approximately 2525 μm2. The remaining electrodes, held in the S, SW, and B reservoirs (illustrated in Figure 1), are fabricated using a combination of Nanoports (Upchurch) and 0.05 cm radius Pt wire.
imposed architecture enables our device to simultaneously produce regular reproducible droplets, while simultaneously facilitating the electrophoretic separation of sample analytes. More importantly, this architecture results in the droplet generation having no adverse effects on the efficiency of the separation. Such an architecture allows us to systematically tailor the composition of our droplets to the next stage in the analytical chain, while simultaneously avoiding dispersion effects or incorporation of impurities into the compartmentalized analyte bands. Alongside this, we show that the separated band factions are diluted into a wide range of concentrations over 600 individual droplets, thus allowing the droplets containing the isolated analyte to be primed for further analysis through the addition of a predetermined solution.
■
CHIP DESIGN, FABRICATION, AND EXPERIMENTAL METHODS A schematic outline of the chip design is displayed in Figure 1, consisting of an electrokinetic T-junction injector, as developed by Manz et al.27 and a droplet creation interface connected by a separation channel approximately 3.5 cm long. Channel height is maintained at 40 μm throughout the channel architecture, with width varying from 25 μm in the separation and carrier flow channels to 50 μm in the droplet channel. The carrier fluid inlet channel is interfaced with the separation channel 3 cm from the point of injection at an angle of 50°. A CCD image of the gel/carrier flow interface at this point is displayed in Figure 1c. The incident angle was determined experimentally by testing multiple designs, and an angle of 50° was found to give the most stable gel/carrier flow boundary. As Figure 1 illustrates, our departure from traditional μCGE chip design is predominantly focused around the incorporation of a droplet T-junction at the base of the separation channel. The carrier inlet is thus positioned before the Pt electrode, Figure 1c, in order to drive electrophoretically separated analytes toward the droplet junction for encapsulation and removal. All devices were fabricated using poly(dimethylsiloxane) Sylgard 184 5802
dx.doi.org/10.1021/ac301141x | Anal. Chem. 2012, 84, 5801−5808
Analytical Chemistry
■
Article
GEL LOADING AND CHIP PREPARATION To prepare the outlined device, chips were initially run with a 0.1× Tris/borate/EDTA (TBE) solution to suppress the creation of a highly charged double layer, thus inhibiting any large scale EOF later in the experiment.30 That treatment was quickly followed by the infusion of a 2.5% poly(ethylene oxide) (PEO) gel loaded onto the chip from the buffer reservoir inlet via syringe pump at 5 μL/min. Optimizing the stability of the gel boundary necessitated testing several different design iterations. Concurrently, several gel matrices with varying viscosities were applied to the system to find the most stable combination. The 2.5% PEO gel was found to have the best combination of viscosity and force distribution, thus allowing it to be delivered into the chip under low pressure, while also being capable of absorbing the slight fluctuations inherent in the backpressure during droplet generation. Lower viscosity gels were found to be too unstable at the boundary, with the gel both retreating up the separation channel under increased backpressure or moving forward creating gel droplets in the converse scenario when backpressure drops. Once the gel phase has been filled past the carrier inlet, the oil and aqueous phase are both initiated. At this point, the syringe pump is stopped; however, since the PEO gel sieving matrix is loaded into the chip under pressure, the PEO gel continues to flow until the driving force in the PEO is reduced and balanced against the backpressure from the droplet creation T-junction. The system is subsequently left under the above conditions for an hour to allow for the formation of a stable gel/carrier interface, as displayed in Figure 1c, alongside a stable droplet flow. Within this design, it is the formation of this stable boundary between the gel and aqueous phases which is the crucial operational dynamic. As such, the method outlined within is constrained to work under the paradigm where both the forward and back pressure experienced at the gel interface is balanced, thus limiting the operational time of the system before the boundary needs to be reformed and recalibrated. In practice, this time window was found to be a couple of hours in our system, allowing adequate time for most CGE analyses to be completed.
removed to obtain a slightly yellow product. The resulting product was mixed with polyethylene glycol (PEG; ED-900, Jeffamine XTJ 501, MW: 900 g/mol, Sigma, 6.93 g) and dissolved in the solvent mixture consisting of HFE 7100 (100 mL) and anhydrous dichloromethane (Sigma Aldrich, 100 mL). The reaction mixture was heated to 65 °C and left continuously stirring for 2 days under argon to form the resulting product with milky color. After solvent removal, the product was centrifuged at 8000 rpm. The resulting product was dried in a vacuum desiccator for a day and used in the experiments without further purification.
■
SAMPLE INJECTION, SEPARATION, AND DETECTION Sample injection and separation were both carried out in a sequential process on the same monolithic PDMS chip, outlined in Figure 1. The injection process employed a variation of the aforementioned electrokinetic injector,27 with individual voltage schemes altered to fit the architecture of the designed chip. An outline of this process is displayed in Table 1, Table 1. Outline of the 3-Step Electrokinetic Injection Process Highlighting the Voltages Applied at Each of the 4 Electrodesa S (V)
SW (V)
B (V)
Patterned BW electrode (V)
time (s)
0 1200 400
1200 0 400
100 0 0
300 0 1200
30 0.1 100
a
S (V) is defined as the voltage in the sample reservoir, SW (V) the applied voltage in the Sample Waste reservoir, B (V) the applied voltage in the Buffer reservoir and Patterned BW (V) as the applied voltage at the patterned "Buffer Waste" reservoir.
highlighting the voltages applied in the flow focusing and injection phases. Through applying a positive potential gradient of 1200 V/cm between the sample reservoir and the sample waste reservoir, sample analytes are electrokinetically initiated to flow between the two. With separation efficiency inversely proportional to injected plug length, a positive bias of +100 V/ cm is applied, from the remaining patterned BW electrode and the Pt wire B electrode, in order to “pinch” the flow and define the sample volume. Upon achieving a stable sample flow, the voltage configuration is switched such that the sample plug, with an approximate volume of 50 pL, is injected down the separation column. A reverse bias is simultaneously applied to the sample and sample waste reservoirs to prevent leakage of sample into the separation channel. All applied voltages were ≤1.2 kV and applied through a home-built 8-channel high voltage power supply configured through a Labview interface developed in-house. Fluorescence detection is employed in a system similar in function to that described by Srisa-Art et al.,31,11 whereby a 4 mW 488 nm diode laser (Coherent Sapphire) is aligned to the back aperture of a 20× objective (Leica). The laser spot is focused into the separation or droplet channel. For detection in the gel phase, the detection point was set at 3 cm from the point of injection and all droplet phase detection was performed directly opposite to the patterned BW electrode at a distance of approximately 80 μm from the droplet T-junction. In all cases, fluorescence readout was collected via a
■
FLUIDICS All fluids were driven by electromechanical precision syringe pumps (PHD2000, Harvard Apparatus) integrated to the PDMS chips via polytetrafluoroethylene (PTFE) tubing (Harvard apparatus) with a 1.09 mm outer diameter in order to create a seal with the 1 mm inlet. Flow rates ranged from 0.1 to 4 μL/min (approximately 25 mms−1) and were delivered using gastight syringes for both the carrier and oil inlet. Retract flow rates ranging from 3 to 14 μL/min were applied to the droplet out reservoir. The continuous oil phase used for all experiments consisted of a mixture of fluoroinert liquid FC-40 (3M) with a home synthesized fluorinated surfactant, added 2% by molecular weight, to increase the stability and regularity of the produced droplets.
■
SURFACTANT SYNTHESIS The surfactant was synthesized similar to the protocol described by Chen et al.22 Briefly, Krytox 157 (FSH, MW: ∼5000 g/mol, Dupont 100 g) was dissolved in anhydrous HFE7100 (Sigma Aldrich, 100 mL) and mixed with excess oxalyl chloride (Aldrich, 25 g). The reaction mixture was then left to stir overnight at 85 °C under argon. The solvent was then 5803
dx.doi.org/10.1021/ac301141x | Anal. Chem. 2012, 84, 5801−5808
Analytical Chemistry
Article
standard deviation of 1.27 ms, again supporting experimental observation of stable droplet production. Such a variance in droplet size is common within comparable systems.31 The above droplet width corresponds to a physical droplet length of approximately 80 μm and thus an approximate droplet volume of 700 pL, calculated from measurement of multiple high speed camera video image frames. The above analysis was further repeated in the presence of an electric field applied in the separation channel. Under such conditions, little to no distortion in droplet generation was observed.
photomultiplier tube (Hamamatsu) coupled to the output of the optical set-up.
■
DROPLET FORMATION Droplets were formed through on-chip incorporation of a Tinjector, as shown in Figure 1.29 In each experiment, the droplet flow was briefly characterized to ensure stable droplet generation prior to electrophoretic separations taking place. Figure 2 shows representative droplet generation data over a
■
INITIAL SEPARATION PHASE Preliminary calibration experiments were conducted using a mixture of two dyes comprising equal concentrations of both 5Carboxyfluorescein (Sigma-Aldrich) and Fluorescein (Fluka Biochem) in pH 8.6 buffer. Both dyes were selected for their well documented differences in mobilities. Figure 3a−c
Figure 3. Fluorescence readout of a time-domain electropherogram displaying 3 consecutive injections and subsequent separations, of a sample plug originating from a 250 μM sample of 5-Carboxyfluorescein and Fluorescein. Peak (i) represents the eluted 5-Carboxyfluorescein band and subsequently (ii) constitutes the Fluorescein component of the sample mixture.
demonstrates the sequential injection of 3 sample plugs (25pL) via the voltage sequence outlined in Table 1. Combining the suppression of inherent buffer EOF by the gel matrix and the back pressure from the interface, analyte velocities can be considered to be almost purely a function of the electrophoretic mobility since no overall EOF is observed. Peak-to-peak resolution was characterized using, R = 2 (t2 − t1)/(w1 + w2), where t2 and t1 denote the respective analyte retention times and w1 and w2 represent peak width. The resolution was found to be 3.04 for our system calculated from the data outlined in Figure 3. A peak width variance of less than 2% between runs was observed, which is comparable to existing literature employing similar molecular dyes and electric field strengths.32 Accordingly, assuming negligible EOF, the efficiency of the system can also be accurately characterized by the number of theoretical plates, N = 5.54 (tR/W0.5), where tR denotes the analyte retention time and W0.5 is the full peak width measured at half maxima (fwhm).33 In our experiments, N was calculated to be approximately 2300 (±300) theoretical plates for 5Carboxyfluorescein and approximately 2200 (±200) plates for Fluorescein, which is comparable to systems proposed in existing literature.34−36 However, due to the high viscosity of our chosen separating medium, the separation time for each injection and separation run is approximately 100 s, which is almost twice as long when compared to traditional μCE systems using standard low viscosity buffer such as 0.1× TBE.37
Figure 2. (a) Optical readout of a 1 s window for droplets containing 0.5 mM Fluorescein in a pH 8.6, 0.1× TBE buffer. Total experimental flow rate was held constant at 3 μL/min. Each peak in (a) corresponds to an individual droplet binned 25-fold from the original acquisiton resolution. (b) A time domain Fourier Transform of the data represented in (a). (c) Histogram of the droplet width for a 100 s acquisition.
period of 1 s for a 0.1× TBE buffer doped with 0.1 μM Fluorescein. The volumetric flow rate for both the aqueous and oil phase is 1.5 μL/min, giving a constant water fraction, Wf, of 0.5 (Wf = Fw/Fw + Fo, where Fw and F0 are the total aqueousand oil-phase flow rates, respectively31). Droplet uniformity is characterized in Figure 2b,c. Figure 2b demonstrates the frequency of droplet production obtained through Fourier analysis of a 100 s time-domain fluorescent readout of the same data presented throughout Figure 2. The droplet generation frequency was calculated to be approximately 38.6 Hz (±2.4 Hz) as determined from the Fourier transform (FT). The droplet generation was found to be highly stable and reproducible even in the presence of the gel phase, as can be seed by the narrow FT peak in Figure 2b. The variation in droplet size for a Wf = 0.5 is shown in Figure 2c. The mean droplet width (defined as the time a droplet spends in the detection volume), in Figure 2, was found to be 14.58 ms with a 5804
dx.doi.org/10.1021/ac301141x | Anal. Chem. 2012, 84, 5801−5808
Analytical Chemistry
Article
production during the different experimental phases present in our operational protocol. By careful observation of the eluted 5Carboxyfluorescein peak in Figure 5a(ii), a clear differentiation between the “empty” doped droplets, approximately pre 81.5 s, and the droplets comprising the sample analyte, approximately 81.5 s plus, can be seen. Conditions were altered slightly for Figure 5c,d; namely, the sample was mixed with the PEO gel buffer prior to insertion into the sample reservoir, leading to more efficient injections. While the retract rate of the system was successfully increased to 14 μL/min to increase throughput without effecting either the droplet or gel boundary. For sequential injections under the same conditions, a slight variance in peak height can be observed and explained predominantly through slight fluctuations in the electrokinetic injection process alongside pressure fluctuations in the sample and buffer reservoirs. Figure 5a−d(ii) displays an enlarged section of the leading edge of the corresponding peak under the different flow regimes, alongside an inset displaying a 0.1 s time domain plot allowing closer inspection of the segmentation and droplet character. With the water fraction and, thus, the droplet size being held constant, the figures demonstrate that a higher degree of segmentation can be achieved with higher flow rates. Analysis of this effect is shown in Table 2. Additionally, Figure 5a−d(ii), demonstrates the presence of a characteristic Gaussian peak emanating from each of the encapsulated analyte bands. The uniform nature of our droplets under the applied field necessary for separation to occur can be assessed by performing a Fourier transform (FT) of the time-domain fluorescent readout. Examples of the FT analysis performed for selected flow rates, in the range of 2−4 μL/min, can be seen in Figure 5a,b (iii). This can be compared to the increase in droplet throughput observed when a retract rate of 14 μL/min is used, Figure 5c,d(iii). The well-defined nature of each of the peaks within our FTs demonstrates regular droplet flow throughout both the separation and encapsulation phases. However, during the flow focusing stage of the electrokinetic injection, the droplet production is disturbed due the potential gradient applied from the patterned electrode opposing the direction of flow. The FT shows sharp peaks and narrow fwhm for each data set indicating high droplet stability.31 The droplet frequency, obtained from the FT, for the lower retract rate in Figure 5a,b, varied from ≈26 to ≈59 Hz with a fwhm of ≈2.5 Hz. This corresponds to the droplets having a polydispersity of approximately 6% or less which compares well with literature.31 Analysis of the corresponding FT for the higher retract rates yields characteristic throughputs of approximately 68−148 Hz for each of the outlined data sets. As Table 2 demonstrates, this allows us to have a dynamic range to choose from between approximately 40−600 droplets per eluted peak.
This translates to a reduction in the analyte velocity of approximately 75% for all samples investigated. For example, the Fluorescein velocity dropped from approximately 0.15 cm/s in a singular phase 0.1× TBE μCE system to 0.014 cm/s in our system. It is possible to reduce the procedural time frame by altering the experimental parameters, namely, by shortening the separation channel length or by increasing the applied voltage. However, experimentally it was found that such changes reduce the maximal resolution in the system, leading to overlapping analyte bands. The parameters employed throughout this work have experimentally been shown to deliver the maximal resolution possible between injected analytes, while simultaneously being complementary to the droplet creation process through absorbing back pressure from the droplet T-Junction.
■
COMPARTMENTALIZATION OF SEPARATED BANDS Upon successful characterization of the droplet t-junction and the electrokinetic separation process, focus was shifted to analyzing the compartmentalization of the analyte within the created droplets. Figure 4 shows the separation and
Figure 4. (a) Droplet phase electropherogram demonstrating the repeated encapsulation of separated analytes into droplets. (b, c) Zoomed in portion of the 5-Carboxyfluorescein and Fluorescein bands. In all cases the droplet frequency was 20 Hz.
encapsulation of two sequential injections with each injection delivering 2 peaks. The peak heights and widths for the eluted bands were found to not vary by more than 14% (±0.7%) and 37% (±10%), respectively, between runs. Importantly, no significant loss in peak height, or resolution, was observed for separated analytes transitioning from the initial separation channel to the droplet phase. Figures 4b and c outline two zoomed in sections of the isolated 5-Carboxyfluorescein and Fluorescein bands, respectively. This figure highlights the droplet stability and uniformity that is achieved during the separation process. Figure 5a,b(i) presents separated and isolated 5-Carboxyfluorescein peaks under 2 differing flow rates and retract rates. While Figure 5c,d(i) demonstrates the additional ability to increase throughput via increasing the retract rate of the system, therefore giving varying degrees of analyte band segmentation into droplets. The regular droplet character can be clearly seen in Figure 5a−d(ii) with the frequency of the droplets maintaining a consistent value throughout the two droplet scenarios (empty droplets and encapsulated sample droplets) once in the separation phase. Dopant (10 μM Rhodamine dye) was added to the carrier flow in order to quantitively analyze the variance in the droplet
■
CONCENTRATION GRADIENT OF SEPARATED BANDS The fluorescent concentration contained within each droplet was analyzed to provide a description of the dynamic concentration range that can be obtained from the segmentation of the original eluted sample plug. This analysis can be seen in Figure 6 alongside Table 2. For a droplet sequence corresponding to the separated analyte band, comparison of the initial droplet and the droplet at peak maxima yields approximately a 10-fold increase in the droplets fluorescence. Interestingly, irrespective of droplet throughput and total flow rate in the system, the maximum dynamic 5805
dx.doi.org/10.1021/ac301141x | Anal. Chem. 2012, 84, 5801−5808
Analytical Chemistry
Article
Figure 5. (a−d) (i) Separated bands encapsulated into a series of droplets under the outlined flow rate conditions. (a−d) (ii) 2 s time traces of the leading edges of the separated peaks shown in (a−d) (i). The inset in (a−d) (ii) show an enlarged 0.1 s trace of the same data outlining the droplet character in each example. (iii) Fourier transforms of the recorded time-domain fluorescent outputs for each example.
concentration range that could be achieved stayed consistent at approximately 1 order of magnitude. By comparing the droplet band fraction (band fraction = [droplet peak]/[peak droplet maxima]) between all separations, a maximal differentiation in droplet concentration of 1:0.0335 was observed, Table 2. As an example, for the 5-Carboxyfluorescein peak in Figure 6a, a dynamic concentration range of approximately 0.13 to 4 μM
spread across 56 droplets is observed. This corresponds to an approximate change in concentration of 0.138 μM between each droplet. For Figure 6c where a retract rate of 14 μL/min was used, a change in concentration of approximately 0.016 μM was observed over the same dynamic range (0.13 to 4 μM). In all cases, the dynamic range was approximated through comparing the fluorescence intensity to a known reference 5806
dx.doi.org/10.1021/ac301141x | Anal. Chem. 2012, 84, 5801−5808
Analytical Chemistry
Article
approach allows for the online separation, characterization, encapsulation, and selection of differing analytes originating from a composite sample. A droplet generation system, producing individual stable droplets with volumes of approximately 200 pL, has been successfully coupled with a μ-channel gel electrophoresis system in a manner which allows for rapid, online, reproducible characterization and compartmentalization of a wide range of samples. The ability to control both efficiency and timings of the separation and compartmentalization phases allows the system to be tailored for specific analytes and/or for further analytical methods. We are currently enlarging the scope of this work to deal with the handling and separation of DNA samples and nanoparticles while looking at incorporating a valving system to allow the complete separation of multiple analytes into separate streams for further analysis.
■
AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Notes
The authors declare no competing financial interest.
■
Figure 6. Graphical representation of the level of analyte encapsulation within individual droplets for the initial 5-carboxyfluorescein peak taken from each of the outlined separations under different flow conditions shown in Figure 5a−d.
ACKNOWLEDGMENTS This work was supported by a case studentship funded in part by the Engineering and Physical Sciences Research Council and National Physical Laboratory. J.B.E. thanks the European Research Council starting grant scheme for financial support.
Table 2. Numerical Representation of the Data Outlined Graphically in Figure 6 Alongside Complementary Data for Additional Separations and Encapsulations of the 250 μM 5Carboxyfluorescein and Fluorescein Sample with Total Flow Rate Varying in the Range of 2−4 μL/min and Total Retract Rate Differing as Outlined in Column 2a Fw + Fo (μL/min)
retract rate (μL/min)
no. droplets/ peak (Peak A)
no. droplets/ peak (Peak B)
[peakdroplet]/ [initialdroplet] (Peak A)
[peakdroplet]/ [initialdroplet] (Peak B)
2 4 2 4
2.5 4.5 14 14
56 127 503 543
48 129 408 459
1:0.0335 1:0.255 1:0.269 1:0.255
1:0.0368 1:0.159 1:0.207 1:0.253
■
REFERENCES
(1) Tiselius, A. T. Faraday Soc. 1937, 33, 524−531. (2) Jorgenson, J. W.; Lukacs, K. D. Science 1983, 222 (4621), 266− 272. (3) Mikkers, F. E. P.; Everaerts, F. M.; Verheggen, T. P. E. M. J. Chromatogr. 1979, 169, 11−20. (4) Walker, P. A.; Morris, M. D.; Burns, M. A.; Johnson, B. N. Anal. Chem. 1998, 70 (18), 3766−3769. (5) von Heeren, F.; Verpoorte, E.; Manz, A.; Thormann, W. Anal. Chem. 1996, 68 (13), 2044−2053. (6) Viovy, J. L.; Duke, T. Electrophoresis 1993, 14 (4), 322−329. (7) Harrison, D. J.; Manz, A.; Fan, Z.; Luedi, H.; Widmer, H. M. Anal. Chem. 1992, 64 (17), 1926−1932. (8) Effenhauser, C. S.; Bruin, G. J. M.; Paulus, A.; Ehrat, M. Anal. Chem. 1997, 69 (17), 3451−3457. (9) Fujii, T. Microelectron. Eng. 2002, 61−62, 907−914. (10) Chang, Y. H.; Lee, G. B.; Huang, F. C.; Chen, Y. Y.; Lin, J. L. Biomed. Microdevices 2006, 8 (3), 215−225. (11) Srisa-Art, M.; Dyson, E. C.; deMello, A. J.; Edel, J. B. Anal. Chem. 2008, 80 (18), 7063−7067. (12) Janasek, D.; Franzke, J.; Manz, A. Nature 2006, 442, 374−380. (13) Casadevall i Solvas, X.; Srisa-Art, M.; deMello., A. J.; Edel, J. B. Anal. Chem. 2010, 82, 3950−3956. (14) Huebner, A.; Sharma, S.; Srisa-Art, M.; Hollfeldr, F.; Edel, J. B.; deMello, A. J. Lab. Chip 2008, 8, 1244−1254. (15) Gielen, F.; deMello, A. J.; Edel, J. B. Anal. Chem. 2012, 84 (4), 1849−1853. (16) Manz, A.; Harrison, D. J.; Verpoote, E. M. J.; Fettinger, J. C.; Paulus, A.; Ludi, H.; Widmer, H. M. J. Chromatogr., A 1992, 593 (1− 2), 253−258. (17) Wu, N.; Peck, T. L.; Webb, A. G.; Magin, R. L.; Sweedler, J. V. J. Am. Chem. Soc. 1994, 116 (17), 7929−7930. (18) Heller, C. Electrophoresis 2001, 22 (4), 629−643. (19) Zhang, C.; Xing, D.; Li, Y. Biotechnol. Adv. 2007, 25 (5), 483− 514. (20) Zeng, J.; Korsmeyer, T. Lab. Chip 2004, 4 (4), 265−277. (21) Jacobson, S. C.; Ramsey, J. M. Anal. Chem. 1997, 69 (16), 3212−3217.
a
The two separations and encapsulations occurring at the higher retract rate were also premixed with the PEO gel prior to loading onto the chip. The above table demonstrates the higher level of compartmentalization possible with sample injection under these conditions.
concentration. Peak-to-initial droplet ratios for additional flow rates are highlighted in Table 2 and displayed in Figure 6b and d. Additionally, Table 2 elucidates the results of our droplets per peak analysis, demonstrating the ability of our system to deliver tailored band segmentation across a wide dynamic range, moving from approximately 50−600 droplets per separated analyte. Furthermore, our platform can be used as a concentration gradient generator in a similar manner to that described by Niu et al.28 Importantly, our system incorporates a separation phase which allows for a dilution series of all separated bands to be performed. In addition, the ability to dilute the droplets on chip is a significant advantage over existing duolithic, or multilayer, chip designs.38
■
CONCLUSION We have demonstrated a novel, high throughput, droplet-based microfluidic CGE band compartmentalization system. The 5807
dx.doi.org/10.1021/ac301141x | Anal. Chem. 2012, 84, 5801−5808
Analytical Chemistry
Article
(22) Chen, C. H.; Sarka, A.; Song, Y. A.; Miller, M. A.; Kim, S. J.; Griffith, L. G.; Lauffenburger, D. A.; Han, J. J. Am. Chem. Soc. 2011, 133 (27), 10368−10371. (23) Edgar, J. S.; Milne, G.; Zhao, Y.; Pabbati, C. P.; Lim, D. S.; Chiu, D. T. Angew. Chem., Int. Ed. 2009, 48 (15), 2719−2722. (24) Niu, X. Z.; Zhang, B.; Marsalek, R. T.; Ces, O.; Edel, J. B.; Klug, D. R.; deMello., A. J. Chem. Commun. 2009, 41, 6159−6161. (25) Fung, E. N.; Yeung, E. S. Anal. Chem. 1991, 63, 1354. (26) Effenhauser, C. S.; Paulus, A.; Manz, A.; Widmer, H. M. Anal. Chem. 1994, 66, 2949−2953. (27) Manz, A.; Effenhauser, C. S.; Burggraf, N.; Harrison, D. J.; Seiler, K. J.; Fluri, K. J. Micromech. Microeng. 1994, 4, 257−265. (28) Niu, X.; Gielen, F.; Edel, J. B.; deMello, A. J. Nat. Chem. 2011, 3, 437−442. (29) Niu, X.; Gulati, S.; Edel, J. B.; deMello, A. J. Lab Chip 2008, 8, 1837−1841. (30) Effenhauser, C. S.; Bruin, G. J. M; Paulus, A. Electrophoresis 1997, 18 (12−13), 2203−2213. (31) Srisa-Art, M.; deMello, A. J.; Edel, J. B. Anal. Chem. 2007, 79, 6682−6689. (32) Dossi, N.; Toniolo, R.; Pizzariello, A.; Susmel, S.; Perennes, F.; Bontempelli, G. J. Electroanal. Chem. 2007, 601, 1−7. (33) Snyder, L. R.; Kirkland, J. J.; Dolan, W. Introduction to Modern Liquid Chromatography, 2nd ed.; Wiley: New York, 2010. (34) Mellos, J. S.; Gorbounov, V.; Ramsey, R. S.; Ramsey, J. M. Anal. Chem. 2008, 80 (18), 6881−6887. (35) Zhang, T.; Fang, Q.; Du, W. B.; Fu, J. L. Anal. Chem. 2009, 81, 3693−3698. (36) Lauer, H. H.; McManigill, D. Anal. Chem. 1986, 58, 166−170. (37) Soper, S. A.; Bruch, R. C. J. Chromatogr., A 1999, 853 (1−2), 107−120. (38) Wen, J.; Guillo, C.; Ferrance, J. P.; Landers, J. P. Anal. Chem. 2007, 79 (16), 6135−6142.
5808
dx.doi.org/10.1021/ac301141x | Anal. Chem. 2012, 84, 5801−5808
