Anal. Chem. 2004, 76, 6137-6143
Single-Cell Chemical Lysis in Picoliter-Scale Closed Volumes Using a Microfabricated Device Daniel Irimia, Ronald G. Tompkins, and Mehmet Toner*
Center for Engineering in Medicine and Surgical Services, Massachusetts General Hospital, Shriners Hospital for Children, and Harvard Medical School, Boston, Massachusetts 02114
Investigating the intracellular contents of single cells is essential for understanding physiologic and pathologic processes at the cellular level. While existing protocols for cell lysis and sample preparation work well for larger samples, scaling to a single-cell level is challenging because of unavoidable analyte dilution and losses. Thus, we are proposing a microfabricated device for the controlled handling and mixing of picoliter cell suspension and lysis solution volumes. Cells and fluids are independently isolated in two microchambers of 25-pL volumes using the geometry of the microchannels and the coordinated action of four on-chip thermopneumatic actuators. Virtual walls formed by liquid-air interfaces in the hydrophobic capillary separate the two volumes, which are subsequently allowed to mix after drawing the air out of the capillary connecting the two microchambers. Following cell lysis, a limited and stable dilution of intracellular components is achieved, simplifying the requirements for subsequent analysis. Two assays at single-cell level, one for direct estimation of the intracellular concentration of a soluble dye and the other for indirect evaluation of intracellular quantities of insoluble actin, demonstrate the use of the microfabricated device for single-cell assays. Cells are the basic structural and functional units of living organisms. Therefore, analysis of biochemical constituents and metabolism at cellular level is important for understanding the physiology and pathology of any organism. Generally, a wide range of methods can be employed to extract averaged information about the composition and processes of cells from samples containing several thousand cells. However, instances where population averages do not reflect actual events in individual cells are emerging.1-3 As a result, there is increasing interest for new strategies for sample preparation and accurate analysis at the single-cell level.4,5 Cell lysis is a major step in sample preparation, and the choice of the lysis protocol is important for the subsequent assay to be * Corresponding author. Phone: (617) 371-4883. Fax: (617) 371-4950. E-mail:
[email protected]. (1) Ferrell, J. E., Jr.; Machleder, E. M. Science 1998, 280, 895-898. (2) Levsky, J. M.; Singer, R. H. Trends Cell Biol. 2003, 13, 4-6. (3) Kaern, M.; Blake, W. J.; Collins, J. J. Annu. Rev. Biomed. Eng. 2003, 5, 179-206. (4) Zieziulewicz, T. J.; Unfricht, D. W.; Hadjout, N.; Lynes, M. A.; Lawrence, D. A. Toxicol. Sci. 2003, 74, 235-244. (5) Outlaw, W. H.; Zhang, S. Q. J. Exp. Bot. 2001, 52, 605-614. 10.1021/ac0497508 CCC: $27.50 Published on Web 09/10/2004
© 2004 American Chemical Society
performed. Different mechanisms for single-cell lysis have been demonstrated,6-13 but in the absence of definite comparisons, chemical methods are particularly attractive because extensive experience and well-established protocols for large samples are available. Interestingly enough, studies show significant differences even among chemical lysis protocols.14,15 Equally important for subsequent assays are the concentration and volume of the target molecules in the sample. Mammalian cells have an average volume of just 1 pL, and cellular analytes would be diluted 6 orders of magnitude in macroscopic samples as small as 1 µL. The combination of large dilution and minute amount of target analyte from a single cell would pose a significant challenge even for the most sensitive analytical techniques. One emerging strategy is to perform cell lysis inside microfluidic devices where intracellular contents from samples containing several16 or single cells11,12,17,18 are released in nanoliter flow streams. Although the dilution is improved 2-3 orders of magnitude compared with macroscopic samples, the combined effects of diffusion and convection with time and distance from the source determine progressive reduction in the concentration of the molecule of interest and complicate the attempts for analysis and quantification.19,20 Until now, sample volumes comparable with the cell volume could only be demonstrated using micropipets for injection of probes or aspiration of the cellular content21,22 or (6) Sims, C. E.; Meredith, G. D.; Krasieva, T. B.; Berns, M. W.; Tromberg, B. J.; Allbritton, N. L. Anal. Chem. 1998, 70, 4570-4577. (7) Lee, S. W.; Tai, Y. C. Sens. Actuators, A 1999, 73, 74-79. (8) Di Carlo, D.; Jeong, K. H.; Lee, L. P. Lab Chip 2003, 3, 287-291. (9) Han, F. T.; Wang, Y.; Sims, C. E.; Bachman, M.; Chang, R. S.; Li, G. P.; Allbritton, N. L. Anal. Chem. 2003, 75, 3688-3696. (10) Huang, Y.; Rubinsky, B. Sens. Actuators, A 2003, 104, 205-212. (11) McClain, M. A.; Culbertson, C. T.; Jacobson, S. C.; Allbritton, N. L.; Sims, C. E.; Ramsey, J. M. Anal. Chem. 2003, 75, 5646-5655. (12) Ocvirk, G.; Salimi-Moosavi, H.; Szarka, R. J.; Arriaga, E. A.; Andersson, P. E.; Smith, R.; Dovichi, N. J.; Harrison, D. J. Proc. IEEE 2004, 92, 115125. (13) Krylov, S. N.; Starke, D. A.; Arriaga, E. A.; Zhang, Z. R.; Chan, N. W. C.; Palcic, M. M.; Dovichi, N. J. Anal. Chem. 2000, 72, 872-877. (14) Maharjan, R. P.; Ferenci, T. Anal. Biochem. 2003, 313, 145-154. (15) Leimgruber, R. M.; Malone, J. P.; Radabaugh, M. R.; LaPorte, M. L.; Violand, B. N.; Monahan, J. B. Proteomics 2002, 2, 135-144. (16) Schilling, E. A.; Kamholz, A. E.; Yager, P. Anal. Chem. 2002, 74, 17981804. (17) Wheeler, A. R.; Throndset, W. R.; Whelan, R. J.; Leach, A. M.; Zare, R. N.; Liao, Y. H.; Farrell, K.; Manger, I. D.; Daridon, A. Anal. Chem. 2003, 75, 3581-3586. (18) Huang, W. H.; Cheng, W.; Zhang, Z.; Pang, D. W.; Wang, Z. L.; Cheng, J. K.; Cui, D. F. Anal. Chem. 2004, 76, 483-488. (19) Taylor, G. Proc. R. Soc. London Ser. A: Math. Phys. Sci. 1953, 219, 186203. (20) Yoshida, F.; Horiike, K.; Huang, S. P. J. Phys. Soc. Jpn. 2000, 69, 37363743.
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microvials formed at the tip of micropipets for single-cell isolation.23,24 However, these techniques are tedious and labor-intensive and require considerable experience for precise cell and fluid manipulation. Our approach to chemical lysis of single cells involves a microfabricated device that handles and mixes 25-pL volumes of two fluids, one containing a single cell and the other a lysing solution. Following the contact between the lysing solution and the cell, the outer cell membrane and the membranes separating intracellular compartments are disrupted. Soluble contents of the cell diffuse in the two chambers, and at the same time, insoluble contents of the cell are exposed to the solution in the chambers. A limited and stable dilution of all intracellular components from a single cell is therefore achieved. Two examples illustrating these features are presented. In the first, the initial intracellular concentration of a fluorescent dye preloaded in one cell is calculated from the estimated final concentration of the dye in the chambers. In the second example, the total quantity of insoluble filamentous actin is indirectly estimated from the difference in the extracellular concentration of a dye before and after binding to actin. EXPERIMENTAL SECTION Principles of Device Design. The single-cell lysis inside the device involved precise manipulation of liquids, cells, and gases and was achieved through passive and active control structures. As shown in the schematics in Figure 1, the device consists of a network of hydrophobic channels and chambers, symmetrical by the longitudinal axis. Two main parallel channels (50 µm wide and 20 µm deep) that taper to 15 × 20 µm in the vicinity of the cell lysis chamber are used to introduce fluids and cells into the device. Smaller channels (12.5 × 3 µm) connect the four air chambers and the two main channels and act as passive valves, restricting the liquid flow and trapping air inside. Within the main channel, the narrowest tapering forms a 15 × 5 µm weir that allows fluid to move through but is not large enough for cells to pass, entrapping them inside the lysis chamber (Figure 1A). In combination with the mentioned passive control structures, four thermopneumatic actuators are used to separate 25-pL volumes of fluids in each of the two lysis chambers (Figure 1C). A mixing channel (12.5 × 5 µm) connects the upper and lower sections of the lysis chambers and a sampling channel (12.5 × 5 µm) is used to evacuate the air from the mixing channel, bringing the liquids in the two sections of the lysis chambers into contact and allowing them to mix (Figure 1D). The implementation of the microfluidic network and the electric heaters into a functional device required the assembly of an elastomeric block with patterned channels of different cross-sectional area onto a glass slide with thin-film electric heaters, and it is outlined below. Photoresist Mold. Three layers (3, 5 and 20 µm) of SU-8 epoxy (Microlitography Corp., Newton, MA) were photopatterned on the same wafer. Glass slides (45 × 50 × 0.1 mm; Fisher, Pittsburgh, PA) were cleaned using a 3:1 volumetric mixture of (21) Eberwine, J.; Yeh, H.; Miyashiro, K.; Cao, Y.; Nair, S.; Finnell, R.; Zettel, M.; Coleman, P. Proc. Natl. Acad. Sci. U.S.A. 1992, 89, 3010-3014. (22) Karlsson, M.; Nolkrantz, K.; Davidson, M. J.; Stromberg, A.; Ryttsen, F.; Akerman, B.; Orwar, O. Anal. Chem. 2000, 72, 5857-5862. (23) Troyer, K. P.; Wightman, R. M. Anal. Chem. 2002, 74, 5370-5375. (24) Yasukawa, T.; Glidle, A.; Cooper, J. M.; Matsue, T. Anal. Chem. 2002, 74, 5001-5008.
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Figure 1. Schematic of the overall device functioning. (A) One cell is introduced with the fluid in the upper main channel and captured in the cell lysis chamber by a dam-like structure. (B) Lysing solution is introduced into the lower main channel. (C) Closed volume fluid compartments are formed by the coordinated action of the four thermopneumatic actuators. (D) Cell lysis is achieved by removing air from the mixing channel, allowing the contact and mixing of the fluids.
sulfuric acid (Ashland Chemical, Columbus, OH) and hydrogen peroxide (Ashland Chemical) and exposed to oxygen plasma in a parallel plate plasma asher (March Inc., Concord, CA). Chrome (50 Å) was sputtered (Lance Goddard Associates, Foster City, CA) on the glass slides and patterned using standard microfabrication techniques to form alignment marks for subsequent fabrication steps. The first layer of SU-8 (3 µm) was spin-coated on the glass slides after an additional cleaning step in the plasma asher, exposed to ultraviolet light through Mylar masks (CADArt, Poway, CA), and processed according to the manufacturer’s specifications. The other two layers of SU-8 (5 and 20 µm) were successively processed and exposed in similar conditions on top of the first one (Figure 2A). Each mask was aligned to the alignment marks in the chrome layer, and good alignment of the structures was critical for producing high-quality molds. By designing the masks such that smaller structures were extended and partially overlapped by larger structures, small errors in mask alignment could be tolerated. Thin-Film Heaters. Chrome (50 Å) and gold (1000 Å, Goddard) were sputtered on glass slides and patterned using patterned photoresist and metal etchant solutions. Circular SU-8 posts (15-µm height, 100-µm diameter) were patterned on the heater glass slides and used for aligning the microchannel network on the heater elements (Figure 2B).
Figure 2. Schematic of device fabrication. (A) Three photopolymer layers of different thickness are successively patterned on a glass slide (A1-A3) and then used as mold for PDMS casting (A4). (B) On a different glass slide, electrical heaters are etched in gold (B1) followed by the patterning of photopolymer alignment posts (B3). (C) Wells in PDMS are aligned to the posts on the heater slide, and the glass and PDMS are bonded to form the final device. (D) Scanning electron micrograph of the PDMS channels shows the upper and lower main channels, the lysis chambers, and the four air chambers.
Device Assembly and Surface Modification. Poly(dimethylsiloxane) (PDMS; Sylgard 184; Dow Corning, Midland, MI) was prepared according to the manufacturer’s instructions and cast over the photoresist mold to create complementary microchannels in PDMS. Through holes, defining the inlets and outlets, were punched using a sharpened 25-gauge needle. The bonding surfaces of the PDMS and the heater coverslips were treated with oxygen plasma (25 s, 50 W, 2% O2) produced in the parallel plate plasma asher. Precise alignment between the PDMS and the coverslip was achieved under a stereomicroscope (MZ8, Leica, Heerbrugg, Switzerland) using a lubrication layer formed by a 10-µL droplet of distilled water placed between the two pieces. The alignment posts on the heater slide and the complementary channels in PDMS were helpful not only in the alignment process but also in stabilizing the assembly during further manipulation and heating of the device on a hot plate (5 min at 70 °C) for complete bonding (Figure 2C). The surface of the microchannels was subsequently modified to achieve uniform hydrophobic characteristics. A 5 mM solution of (heptadecafluoro-1,1,2,2-tetrahydrodecyl)dimethylchlorosilane (Gelest, Morrisville, PA) in 99% toluene (Sigma-Aldrich, St. Louis,
MO) was flushed through the device at a rate of 5 µL/min for 3 min, followed by a wash with 99% toluene for 2 min at the same rate. Finally, residual toluene was removed using pressured nitrogen and heating the device to 120 °C on a hot plate for 8 h. Device Characterization. Static contact angles between phosphate-buffered saline solution (PBS), sodium dodecyl sulfate (SDS), and guanidine thiocyanate (GTC) and fluorosilane-treated or untreated glass and PDMS were measured by placing a drop of liquid (3 µL) on the surface of interest and observing it through a microscope. Dynamic contact angles between PBS and fluorosilane-treated PDMS were measured during the filling and clearing of water inside 50-µm-wide channels of the device. The activation of the thermopneumatic actuator was characterized from serial photographs of the ejected air bubble and measurements of the projected area using MetaMorph imaging software (Universal Imaging, Downingtown, PA). Volumes were normalized to the volume of the air chamber. The surface of the PDMS mold was sputter-coated with a 5-nm-thick layer of gold-palladium and examined using a scanning electron microscope (JSM5600LV, Joel Inc, Peabody, MA). Images were acquired using 5-kV acceleration voltage, and 160× and 500× magnification (Figure 2D). Cell Culture. Human lymphoblasts (MOLT-3, American Type Culture Collection, Rockville, MD) were cultured in RPMI 1640 media (GIBCO BRL Life Technologies, Rockville, MD) supplemented with 10% bovine calf serum (GIBCO) at 37 °C in an atmosphere of 10% CO2. Cultured cells were split 1:10 and subcultured every 3 days. Before the experiment, 5 mL of a cell suspension was centrifuged and the medium then removed. The pellet was resuspended in either PBS (GIBCO), or a 6 µM solution of Cell-Tracker Orange CMTMR fluorescent dye (Molecular Probes, Eugene, OR) in PBS, followed by 10-min incubation at 37 °C. Cells were then centrifuged, washed once with PBS, and resuspended into 5 mL of PBS. The final cell suspension was adjusted to a cell density of 104 cells/mL using PBS. Lysing and Calibration Solutions. GTC (Sigma) or SDS (Sigma) was dissolved in distilled water in order to prepare lysing solutions of 3 M GTC, 0.2% SDS, or 0.1% SDS. For some of the experiment,s either Oregon-green phalloidin or YOYO-1 iodide (Molecular Probes) was added to the 0.2% SDS for final concentrations of 165 and 4 nM, respectively. Serial dilutions of the CMTMR fluorescent dye (6 µM, 600 nM, 60 nM, and 6 nM) were prepared by dilution in PBS. Equal volumes (100 µL) of the dye solutions and 3 M GTC were flown through the chip and allowed to mix. Serial dilutions of 84, 42, and 21 nM Oregon-green phalloidin in 0.2% SDS were also prepared and flown through the chip. Fluorescence intensity in the chambers was calibrated using the serial dilutions. An exponential curve was fit to the data, and preexponential and exponential coefficients were calculated for each of the two dyes. Experimental Setup. Fluidic connections were made using Tygon tubing and syringe needles (Small Parts, Miami Lakes, FL). Electrical connections between the device and control electronics were made by soldering multifilament wires onto the gold pads at the periphery of the device. Four single-pulse generators were fabricated in-house, using two precision monostable circuits (74HC/HCT4538, Philips, Eindhoven, The Netherlands) and four power operational amplifiers (LM675, National Semiconductor, Santa Clara, CA) with the proper passive components. Single Analytical Chemistry, Vol. 76, No. 20, October 15, 2004
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pulses (1 V, 100 mA, 200 ms) were simultaneously delivered to each heater. Experiments were imaged using an inverted microscope (Nikon Eclipse 2000) equipped with a color video camera and recorded on a computer. After priming the upper main channel with PBS (3 × 104 N/m2 driving pressure), cells (100 µL from a 104 cells/mL cell suspension) were introduced in the upper main channel and driven slowly toward the upper lysis chamber. One cell was captured in the cell chamber by the cell capture dam (5 µm) while allowing the fluid to flow through. After cell capture, the flow was stopped and a lysing solution was introduced in the lower main channel. By the coordinated use of thermopneumatic actuators, the cell was isolated in 25 pL of PBS in the upper chamber and 25 pL of the lysing solution was separated in the lower lysing chamber. The two liquids were brought into contact by extracting the air from the mixing channel, and mixing occurred passively, by diffusion. Cell lysis was observed either directly using phase contrast microscopy or indirectly, through the release of the fluorescent dye preloaded in cells. Under phase contrast microscopy, living cells appear bright and they turn darker after complete lysis of the membrane. RESULTS AND DISCUSSION A microfabricated device was developed and tested with the objective of capturing one cell from a cell suspension, separating it in a closed volume, and mixing it with a controlled volume of lysing solution while observing cell lysis (Figure 1). Two major challenges had to be simultaneously overcome for quantitative analysis of intracellular contents, namely, capturing and lysing one cell of interest inside a closed space and keeping the intracellular components close to their original concentration. Device Design and Functioning. We approached the cell capturing and fluid mixing in closed volumes by relying on capillary effects in hydrophobic channels. Therefore, we used small changes in the pressure at the liquid-gas interfaces to manipulate small volumes of fluids inside partially filled hydrophobic capillaries. While for the majority of microfluidic devices the formation of air bubbles inside can render the device unusable, there are numerous examples of constructive uses of liquid-air interfaces for different applications. These include wall-less control of the flow of liquid streams on hydrophobic surfaces,25 passive valves that would precisely stop the liquid flow at a certain position inside a channel without the need for moving parts,26 accurate metering of nanoliter volumes of liquids in microchannels,27,28 and mixing of 600-pL volumes inside a capillary.29 In this device, we combined the use of liquid gas interfaces as both passive valves to control the flow of fluid and barriers to isolate picoliter volumes of liquids. Passive valves were formed by dimensional variation of uniformly hydrophobic channels. According to the Young-Laplace equation, the pressure needed to move a liquid through rectangular microchannels is dependent on σ, the surface tension of (25) Zhao, B.; Moore, J. S.; Beebe, D. J. Science 2001, 291, 1023-1026. (26) Duffy, D. C.; Gillis, H. L.; Lin, J.; Sheppard, N. F.; Kellogg, G. J. Anal. Chem. 1999, 71, 4669-4678. (27) Handique, K.; Burke, D. T.; Mastrangelo, C. H.; Burns, M. A. Anal. Chem. 2001, 73, 1831-1838. (28) Lee, S. H.; Lee, C. S.; Kim, B. G.; Kim, Y. K. J. Micromech. Microeng. 2003, 13, 89-97. (29) Hosokawa, K.; Fujii, T.; Endo, I. Anal. Chem. 1999, 71, 4781-4785.
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Table 1. Measured Contact Angles between Different Liquids and the Glass and PDMS Surfaces, before and after Surface Modification contact angle glass (deg)
(× PBS 3M GTC SDS 0.1% SDS 0.2%
contact angle PDMS (deg)
before after before after σ fluorosilane fluorosilane fluorosilane fluorosilane N/m) treatment treatment treatment treatment
103
72.0 67.8 60.5 49.0
15 5 10 5
87 85 78 70
100 93 92 66
99 100 97 90
Table 2. Calculated Pressure Differences Required for Driving Particular Liquids in Different Microchannels of the Device after Surface Modification (× 103 N/m2) pressure gradient (× 103N/m2)
PBS 3M GTC SDS 0.1% SDS 0.2%
main channel
lysis chamber passive valve
mixing channel
air chamber passive valve
-2.6 -1.6 -1.0 -0.3
-4.3 -2.7 -1.7 -0.6
-12.3 -7.7 -4.8 -1.7
-21.6 -13.5 -8.5 -3.0
the liquid, w and h, the depth and the height of a channel, and θ, the static contact angles between liquid and walls. For channels formed between PDMS and glass this pressure was calculated as
P ) σ cos θPDMS
(w2 + h1) + σ cos θ (h1) glass
(1)
In the absence of surface modifications, the contact angles between liquids and the PDMS or glass walls of the device were relatively small and the capillary pressure was positive, indicating capillary filling. However, following fluorosilane treatment, contact angles were measured in excess of 90° (Table 1), resulting in negative capillary pressure and repellent forces at the liquid gas interface. Therefore, small channels could act as passive valves restricting the flow of fluids to the larger channels and trapping air inside the air chambers during device priming. Following thermopneumatic actuation, air bubbles could expand from the chambers into the main channels and effectively block liquid flow through these channels. The hydrophobic nature of the liquidwall interaction was critical for stable and complete separation of picoliter volumes of fluid.30 Overall, the relatively large differences in the pressure thresholds for different valves (Table 2) assured a robust control over the fluid movement inside the device and allowed the use of low concentrations of detergents in the working fluids. A key feature of our approach to isolating small volumes was the simultaneous and symmetrical generation of small air bubbles at the ends of the chambers, through the coordinated action of four thermopneumatic actuators, without the need for external pressure sources. Electricity was applied to the heaters for only 200 ms, short enough to avoid major heating of the liquids in the channels. Upon actuation, the air in each chamber (∼100 pL) almost doubled its volume, expanding into the main channels, and (30) Ajaev, V. S.; Homsy, G. M. J. Colloid Interface Sci. 2001, 240, 259-271.
Figure 3. Thermopneumatic actuator operation. (A) Two air chambers, the corresponding thin-film heaters, and the main channel are shown using bright-field microscopy. The inset outlines the location of the actuators on the chip. The main channel is filled with PBS. Two air bubbles are formed by the expansion of the air in the air chambers after actuation, and 25 pL of liquid is separated in the lysis chamber. (B) The volume change during actuation normalized to the air chamber volume. One 200-ms electric pulse is used to expand the air from the chambers into the main channel. After actuation, some air remains sequestered in the main channel, separating the liquid segments for at least 30 min. The volumes of the air partitions are dependent on the geometry of the channel and represent 14-20% from the air chamber volume.
breaking the continuity of the liquid column (Figure 3). The pressure required for the initial outburst of air from the air chamber and through the 2 × 12.5 µm channel was estimated to 4.2 × 104 N/m2, and observations of the bubble formation in the main channel suggested that this pressure was achieved within 50 ms after the heaters were turned on. During the air expansion, we measured a dynamic contact angle of 55-60° between a PBS solution and the walls of the channel and interface advancing speeds up to 300 µm/s. The movements of newly formed liquidair interfaces were passively controlled through variations of the cross-sectional area of the main channels in the vicinity of the lysis chambers. Constrictions next to each of the lysis chambers prohibited the entrance of air, while increasing cross-sectional area of the channels away from the chambers directed the expansion of the air bubbles outward. The symmetry in speed and amplitude
of the newly formed air-liquid interfaces on both sides of the lysing chambers assured that the 25-pL fluid volumes were not ejected from the compartments. After the heaters were turned off, the air cooled fast, probably through heat conduction through the thin glass slide. Consequently, the air begun retracting and the contact angle for the moving interface increased to 105°. Because of the new pressure balance and dependency on the shape of the channel, the air did not return to its initial volume and 14-20% of the initial air volume remained trapped inside the main channel (Figure 3). This residual volume could increase to 33% if two successive pulses at a 200-ms time interval were applied (data not shown). This observation suggests that other mechanisms, such as water evaporation at the liquid-air interface, gas transport through PDMS, and hysteresis in the contact angle, may be involved in the new equilibrium state. Following cell capture and fluid compartmentalization, solutions in the two sections of the lysis chamber were forced into contact by decreasing the pressure in the mixing and sampling channels (Figure 1). After the two liquids came into contact, mixing occurred by diffusion through the mixing channel. Typically, the smaller height area of the junction between the sampling and the mixing channels prevented the liquids in the mixing channel from entering the sampling channel. If needed, supplementary mixing could be achieved by repeatedly drawing and pushing back some of the liquid from the mixing channel into the sampling channel through additional pressure changes. Although several devices for manipulating cells and small volumes of solutions have been described,31,32 none could simultaneously fulfill both challenges of isolating a single cell and mixing two fluids of volumes comparable to the cell volume. Mechanical valves have been used to separate mammalian cells in nanoliter33 and bacterial cells in hundreds of picoliter34 volumes. Such volumes were 3 orders of magnitude larger than the cell volume. Isolating cells into droplets that have all35 or part23 of their surface exposed to atmospheric air was usually complicated by liquid evaporation, which may occur within seconds for nanolitersize droplets. When droplets were isolated in oil,36 or an oil lid was used to isolate compartments,37 the use of detergents for cell lysis or the diffusion of lipo-soluble components into the oil would have become a problem. By comparison, isolating cells into small liquid compartments isolated by water-saturated air compartments had the advantages of smaller, precisely controlled volumes, easier manipulation, and larger flexibility in the choice of reagents. Single-Cell Lysis and Biochemical Assays. Cells were lysed by the surfactant action of the chemical agents on the cell membrane following contact between the solutions in the two chambers and the diffusion of lysing solution into the cell chamber (Figure 4). Cell lysis in the device occurred quickly due to the rapid diffusion of the low molecular weight lysing molecules in (31) Andersson, H.; van den Berg, A. Sens. Actuators, B 2003, 92, 315-325. (32) Reyes, D. R.; Iossifidis, D.; Auroux, P. A.; Manz, A. Anal. Chem. 2002, 74, 2623-2636. (33) Hong, J. W.; Studer, V.; Hang, G.; Anderson, W. F.; Quake, S. R. Nat. Biotechnol. 2004, 22, 435-439. (34) Thorsen, T.; Maerkl, S. J.; Quake, S. R. Science 2002, 298, 580-584. (35) Santesson, S.; Andersson, M.; Degerman, E.; Johansson, T.; Nilsson, J.; Nilsson, S. Anal. Chem. 2000, 72, 3412-3418. (36) Velev, O. D.; Prevo, B. G.; Bhatt, K. H. Nature 2003, 426, 515-516. (37) Klauke, N.; Smith, G. L.; Cooper, J. Biophys. J. 2003, 85, 1766-1774.
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Figure 4. Single-cell lysis. (A) Bright-field microscopy image shows one MOLT-3 lymphoblast captured in the upper lysis chamber and isolated in 25 pL of PBS. (B) Phase contrast of the upper lysis chamber shows the insoluble cellular structures following chemical lysis (0.2% SDS) of one cell. Scale bars are 25 µm long.
the compartments (GTC 118, SDS 288). We measured a time to cell lysis of the order of 0.5-1 s in the presence of 3M GTC, 3 s in the presence of 0.2% SDS, and 10 s in the presence of 0.1% SDS. Concentrations of SDS lower than 0.1% were ineffective in lysing the cell 10 min after exposure. One important feature of the microfabricated device was the mechanically gentle nature of cell lysis, given that the lysing agent reached the cell by diffusion in the absence of any significant convection. We noted that the size of the cells did not change significantly following the lysis, and we speculated that the insoluble cytoskeleton components and larger molecules maintain their initial position and configuration without considerable mechanical stress. Preliminary data using Oregon-green phalloidin and YOYO-1 dye fluorescent dyes suggest that at least actin and DNA retain their position for minutes after cell lysis in the device. Two assays for intracellular molecules are presented as examples of potential uses for the single-cell device. In both cases, following the lysis of the cell membrane, soluble components diffuse between the cell and the device chambers. In one assay, cells were preloaded with cell-tracker orange CMTMR fluorescent dye and the original intracellular dye concentration was calculated from the final dye concentration in the two chambers. In the second assay, cells were lysed in the presence of Oregon-green phalloidin and the quantity of intracellular insoluble actin binding the phalloidin was estimated indirectly from the change in fluorescence inside the chambers. For cells loaded with cell-tracker orange, a 50-fold dilution of the fluorescent dye after cell lysis was taken into account. This assumed that all dye released in the chambers had been initially contained only inside the cell of ∼1-pL volume and that the volume of the two chambers was 50 times larger than the cell volume. Experimentally, the fluorescence intensity versus time following cell lysis was recorded at two different locations in the chambers 6142 Analytical Chemistry, Vol. 76, No. 20, October 15, 2004
Figure 5. Serial fluorescence images of the dye diffusing throughout the two compartments after the single-cell lysis. (A) One cell, preloaded with fluorescent dye, was captured in the 25-pL upper chamber and lysed by 25 pL of GTC lysing solution, releasing the intracellular fluorescent dye into the 50-pL closed volume. The fluorescent dye diffused rapidly throughout the cell lysis chamber, then advanced slowly through the mixing channel, and diffused into the lower chamber. Pictures were taken at 3, 33, and 51 s after bridging cell suspension and lysing solution. Arrows indicate the fields where the fluorescence intensity was measured. (B) A calibration curve was generated by measuring the fluorescent signal from solutions of known dye concentration. (C) Concentrations of the released fluorescent dye in the upper (dotted line) and lower (solid line) chambers were calculated from the fluorescence signal using the calibration curve. The concentration in the upper chamber reached a peak 20 s after lysis initiation. Steady state was achieved at ∼90 s.
(Figure 5). The actual concentration of the dye in the chambers was calculated based on a calibration experiment, where several dilutions of the dye were flushed through the device, and the fluorescence intensity was measured. Calculations using the final equilibrium measurements estimated the intracellular concentration of the fluorescent dye at 2 µM (Figure 5). Some particularities of the CMTMR dye were important in interpreting this concentration value. Following initial loading, part of the CMTMR fluorescent dye was trapped inside the cells by binding to glutathione, while part could still move inside the cell and eventually be released from the cell by disrupting the cellular membrane. It was also assumed that the dye did not require enzymatic cleavage for activation and did not change its fluorescent properties upon intracellular binding. Thus, the 2 µM value for the intracellular concentration of the cell-tracker orange that was estimated from the free dye concentration inside the chambers following the cell lysis referred mainly to dye not bound to structural proteins. It is noteworthy that, apart from the microfabricated device, only standard imaging equipment was required for determining the dye concentrations inside the cell. If the cell would have been lysed in a “macro” vial, of 50-µL volume, the final concentration of the dye in the solution would have been 6 orders of magnitude
Figure 6. Fluorescence image of actin cytoskeleton after detergent cell lysis. One cell was lysed using 0.2% SDS and actin simultaneously stained by Oregon-green phalloidin. The filamentous actin remained stable for at least 30 min after cell lysis without fixation. The equilibrium concentration of unbound phalloidin in the chambers was used to evaluate the amount of filamentous actin inside the cell. Scale bar is 25 µm.
lower, in the picomolar range (2 × 10-12 M), and beyond the detection limit for regular digital cameras. Chemical lysis of the same cell inside a microfabricated channel with flow would have complicated the calculation of initial intracellular concentration because the dye release from the cell was not sudden, but a slow process developing over tens of seconds. Therefore, it would have been difficult to infer the initial intracellular concentration of the molecule of interest only from the concentration-time dependence measurement.19,20 Overall, although the confinement of intracellular molecules following cell lysis in a small space may not be critical for detecting abundant targets using sensitive techniques (e.g., capillary electrophoresis11), having a concentrated sample from a single cell appears to be beneficial for quantitative measurements using less sensitive techniques (e.g., regular fluorescence measurements). In another assay, the intracellular amount of insoluble actin filaments was estimated using the microfabricated device. One cell was captured in the upper lysis chamber and then exposed to a solution of detergent (2% SDS) and Oregon-green phalloidin (165 nM) (Figure 6). Following the lysis of the cell membrane, phalloidin dye molecules entered the cell and bonded to filamentous actin with high affinity and 1:1 stoichiometric ratio.38 Because Oregon-green fluorescent molecules are known to increase their yield upon binding to actin, direct estimation of bound actin would have been difficult.39 Instead, we used an indirect method to calculate the amount of insoluble actin from the change in unbound dye concentration in the chambers. We first generated a standard curve to correlate the fluorescent signal and the concentration of unbound dye inside the device. We then estimated the concentration of fluorescent dye in solution to 47 nM at 30 s after cell lysis. By the conservation of mass, the number of molecules bonded to filamentous actin, and consequently the total amount of filamentous actin in one cell, were estimated at 1 × 106 molecules/cell. This value was of the same order of magnitude with other values reported in the literature for the average amounts of filamentous actin per cell measured from bulk assays.39,40 Simpler calibration experiments and the capacity to perform indirect quantitative measurements at single-cell levels (38) Cano, M. L.; Cassimeris, L.; Joyce, M.; Zigmond, S. H. Cell Motil. Cytoskeleton 1992, 21, 147-158. (39) Katanaev, V. L.; Wymann, M. P. Anal. Biochem. 1998, 264, 185-190. (40) Cano, M. L.; Lauffenburger, D. A.; Zigmond, S. H. J. Cell Biol. 1991, 115, 677-687. (41) Lundell, N.; Schreitmuller, T. Anal. Biochem. 1999, 266, 31-47. (42) Bosserhoff, A.; Wallach, J.; Frank, R. W. J. Chromatogr. 1989, 473, 71-77.
are advantages of our approach. Furthermore, only standard imaging equipment is needed to perform the measurements. Various chemical and biochemical assays can potentially benefit from the new ability to mix cells and solutions in picoliter volumes. Several protocols for sample preparation involving manipulation and mixing of cells and reagents, which are currently handled on the macroscale, could be implemented at microscale and integrated in lab-on-a-chip and sensor devices. Alternatively, the single-cell lysate could potentially be transported out of the device for off-chip analysis. Although the transport may require some increase in the sample volume, resulting in dilution of the cell lysate, this would also decrease the concentration of the lysing agent. This may prove useful in some situations, for example, to avoid the use of detergent extraction procedures for SDS removal from peptide samples before mass spectrometry analysis.41,42 In addition to sample preparation, several chemical and biochemical assays exploring cellular functions could be implemented in the device. Experimental situations requiring the detection of small amounts of analyte, e.g., autocrine and paracrine signaling, fluxes of metabolites to and from cells, or low-abundance intracellular species, would benefit from limited dilution of the molecules of interest in volumes comparable to cell volume. Improved signalto-noise ratio, easier calibration, and the ability to perform direct and indirect measurements are also advantages of single-cell lysis in a closed volume compared to other sample preparation techniques. CONCLUSIONS Single-cell capture and chemical lysis inside a 50-pL closed volume is demonstrated in a microfabricated device. Passive and active control mechanisms were implemented for the manipulation, isolation, and mixing of solutions with volumes comparable to cell volume. The amount of preloaded dye fluorescent dye inside a single cell was estimated by directly measuring the steady-state concentration inside the closed volume after cell lysis. The amount of insoluble actin was indirectly calculated from the consumption and resulting decrease in concentration of fluorescently tagged phalloidin. ACKNOWLEDGMENT We thankfully acknowledge all the investigators in the Model Validation Core of the Inflammation and the Host Response to Injury Large-Scale Collaborative Research Agreement for helpful discussions. We are also thankful for the help with surface modification chemistry from Dr. Alexander Revzin, for the technical support with microfabrication procedures from Mr. Octavio Hurtado, and for the constructive feedback with reviewing the manuscript from Dr. Alexander Revzin and Mr. Kevin King. Microfabrication procedures were performed in the BioMEMS Resource Center at the Center for Engineering in Medicine, at the Massachusetts General Hospital. This work partially was supported by the National Institute of General Medical Sciences (Inflammation and the Host Response to Injury Large Scale Collaborative Project, U54 GM062119) and by the National Institute of Biomedical Imaging and Bioengineering (BioMEMS Resource Center, P41 EB002503). Received for review February 13, 2004. Accepted July 30, 2004. AC0497508 Analytical Chemistry, Vol. 76, No. 20, October 15, 2004
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