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Development of Multistage Magnetic Deposition Microscopy Pulak Nath,†,‡ Joseph Strelnik,†,§ Amit Vasanji,† Lee R. Moore,† P. Stephen Williams,† Maciej Zborowski,† Shuvo Roy,†,| and Aaron J. Fleischman*,† Department of Biomedical Engineering/ND-20, Lerner Research Institute, Cleveland Clinic, 9500 Euclid Avenue, Cleveland, Ohio 44195, and Department of Biomedical Engineering, University of Cincinnati, 2600 Clifton Avenue, Cincinnati, Ohio 45221 Magnetic deposition microscropy (MDM) combines magnetic deposition and optical analysis of magnetically tagged cells into a single platform. Our multistage MDM uses enclosed microfabricated channels and a magnet assembly comprising four zones in series. The enclosed channels alleviate the problem plaguing previous versions of MDM: scouring of the cell deposition layer by the air-liquid interface as the channel is drained. The fourzone magnet assembly was designed to maximize capture efficiency, and experiments yielded total capture efficiencies of >99% of fluorescent- and magnetically-labeled Jurkat cells at reasonable throughputs (103 cells/min). A digital image processing protocol was developed to measure the average pixel intensities of the deposited cells in different zones, indicative of the marker expression. Preliminary findings indicate that the multistage MDM may be suitable for depositing cells and particles in successive zones according to their magnetic properties (e.g., magnetic susceptibilities or magnetophoretic mobilities). The overall goal is to allow the screening of multiple disease conditions in a single platform. Isolation of targeted cells from a biologically relevant sample such as peripheral blood is a common requirement in biomedical science. Generally, cell separation is a sample preparation step where complex biological samples are simplified by depleting unwanted cells, thereby enriching desired cells. Separation is generally achieved based on the differences in their physical properties, such as density and size, or their biochemical properties, such as surface antigen expression.1,2 Advanced sorting techniques, such as fluorescence-activated cell sorting (FACS)3 and magnetic cell separation,4 have evolved over the years and now are capable of separating cells with high selectivity and * Corresponding author. Phone: 1-216-445-3218. Fax: 1-216-444-9198. E-mail:
[email protected]. † Cleveland Clinic. ‡ Present address: Los Alamos National Laboratory, Los Alamos, NM 87545. § University of Cincinnati. | Present address: Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, CA 94158. (1) Recktenwald, D.; Radbruch, A. Cell Separation Methods and Applications; Marcel Dekker Inc.: New York, 1998. (2) Pertoft, H. J. Biochem. Biophys. Methods 2000, 44, 1–30. (3) Herzenberg, L. A.; Parks, D.; Sahaf, B.; Perez, O.; Roederer, M.; Herzenberg, L. A. A. Clin. Chem. 2002, 48, 1819–1827. (4) Safarik, I.; Safarikova, M. J. Chromatogr., B 1999, 722, 33–53. 10.1021/ac8010186 CCC: $40.75 2009 American Chemical Society Published on Web 12/04/2008
recovery. Several disease conditions can be detected or monitored by counting the number of disease specific cells. For example, the low concentration of helper T lymphocytes (characterized by a surface expression of cluster of differentiation 4, or CD4+ cells) and a low ratio of the number of those cells to the number of cytotoxic T lymphocytes (CD8+ cells), CD4/CD8, is a clinical measure of AIDS progression.5,6 The presence of circulating tumor cells (CTCs) is a hallmark of metastatic disease in cancer patients. The concentration of these cells in the blood has been shown recently to correlate with the prognosis following treatment of breast cancer.7,8 A count of lower than 5 tumor cells in 7.5 mL of whole blood (that is, less than one CTC per milliliter of whole blood) has been shown to correlate with a lower incidence of relapse following chemotherapy.7 Biological samples such as whole blood are a complex mixture of cells (∼5 × 109/mL of red blood cells, ∼8 × 106 /mL of white blood cells, and ∼3 × 108 /mL of platelets) suspended in plasma. As a result, detecting the disease-specific cells in a blood sample for diagnostic purposes requires their isolation from the large pool of normal blood cells. Modern tools such as FACS and magnetic cell sorting rely on the interaction between cell surface antigens and antibodies conjugated to fluorochromes or magnetic particles, and therefore, these techniques can be very specific to targeted cells.3,4,9 However, FACS machines are bulky and expensive and require special training to operate. Magnetic cell separation, on the other hand, is a simple technique that often employs a relatively small permanent magnet and a simple fluidic system to isolate magnetically tagged cells. Since virtually all untreated biological materials are diamagnetic or only weakly magnetic, magnetic cell separation can be highly specific and could be applied to cell mixtures as complex as whole blood.10 Therefore, there have been significant advances in the area of magnetic cell separations. These develop(5) Yu, L. M.; Easterbrook, P. J.; Marshall, T. Int. J. Epidemiol. 1997, 26, 1367– 1372. (6) Grodzinski, P.; Yang, J.; Liu, R. H.; Ward, M. D. Biomed. Microdevices 2003, 5, 303–310. (7) Cristofanilli, M.; Budd, G. T.; Ellis, M. J.; Stopeck, A.; Matera, J.; Miller, M. C.; Reuben, J. M.; Doyle, G. V.; Allard, W. J.; Terstappen, L. W.; Hayes, D. F. N. Engl. J. Med. 2004, 351, 781–791. (8) Cristofanilli, M.; Hayes, D. F.; Budd, G. T.; Ellis, M. J.; Stopeck, A.; Reuben, J. M.; Doyle, G. V.; Matera, J.; Allard, W. J.; Miller, M. C.; Fritsche, H. A.; Hortobagyi, G. N.; Terstappen, L. W. J. Clin. Oncol. 2005, 23, 1420–1430. (9) Shapiro, H. M. Practical Flow Cytometry, 4th ed.; John Wiley and Sons, Inc.: Hoboken, NJ, 2003. (10) Liberti, P. A.; Rao, C. G.; Terstappen, L. W. M. M. J. Magnet. Magnet. Mater. 2001, 301, 301–307.
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ments primarily focused on miniaturization,11-21 improving separation efficiencies, and separating cells based on their magnetic properties.18,22,23 Separations have been achieved in flow or a static medium by either selective immobilization and deposition12,16,21,24 or by magnetic deflections.18,19,22,23,25 Deposited cells may be difficult to recover compared to deflection-based systems but can allow direct interrogation by optical20,21,24,26 or electrochemically12 means. Both permanent magnets11,13,21 and electromagnets12,16,27,28 have been utilized as the source of magnetic gradients. However, electromagnets can be difficult to fabricate and require a power source and control for operation. This work is based on our early experience with an industrial instrument for the analysis of wear particles in lubricating oil taken from mechanical equipment. It is known as the ferrograph29 and its subsequent modifications for biomedical applications, bioferrograph30,31 and magnetic deposition microscopy, MDM.26 Ferrous and other magnetically susceptible debris from a flowing sample is deposited onto a glass slide adjacent to the high gradient region of the interpolar gap of a dipole magnet assembly.31 The slide is subsequently subjected to analysis, such as cytochemical staining and optical microscopy.31,32 The strong magnetic field and gradient at the interpolar gap is even capable of attracting and depositing certain cells on the slide based on their own, weak magnetic susceptibility (without tagging by magnetic microparticles), such as in the case of malaria parasite-infected red blood (11) Blankenstein, G. In Scientific and Clinical Applications of Magnetic Carriers; Hafeli, U., Schuett, W., Teller, J., Zborowski, M., Eds.; Plenum Press: New York, 1997; pp 233-245. (12) Choi, J.-W.; Ahn, C. H.; Bhansali, S.; Henderson, H. T. Sens. Actuators, B 2000, 68, 34–39. (13) Pamme, N.; Manz, A. Anal. Chem. 2004, 76, 7250–7256. (14) Han, K.-H.; Bruno Frazier, A. J. Appl. Phys. 2004, 96, 5797–5802. (15) Inglis, D. W.; Riehn, R.; Austin, R. H.; Sturm, J. C. Appl. Phys. Lett. 2004, 85, 5093–5095. (16) Smistrup, K.; Tang, P. T.; Hansen, O.; Hansen, M. F. J. Magnet. Magnet. Mater. 2006, 300, 418–426. (17) Yellen, B. B.; Er, R. M.; So, H. S.; Hewlin, J. R.; Shan, H.; Le, G. U. Lab Chip 2007, 7, 1681–1688. (18) Liu, C.; Lagae, L.; Wirix-Speetjens, R.; Borghs, G. J. Appl. Phys. 2007, 101, 024913-024913-024914. (19) Xia, N.; Hunt, T. P.; Mayers, B. T.; Alsberg, E.; Whitesides, G. M.; Westervelt, R. M.; Ingber, D. E. Biomed. Microdevices 2006, 8, 299–308. (20) Smistrup, K.; Kjeldsen, B. G.; Reimers, J. L.; Dufva, M.; Petersen, J.; Hansen, M. F. Lab Chip 2005, 5, 1315–1319. (21) Chang, W. S.; Shang, H.; Perera, R. M.; Lok, S. M.; Sedlak, D.; Kuhn, R. J.; Lee, G. U. Analyst 2008, 133, 233–240. (22) Schneider, T.; Moore, L. R.; Jing, Y.; Haam, S.; Williams, P. S.; Fleischman, A. J.; Roy, S.; Chalmers, J. J.; Zborowski, M. J. Biochem. Biophys. Methods 2006, 68, 1–21. (23) Pamme, N.; Eijkel, J. C. T.; Manz, A. J. Magnet. Magnet. Mater. 2006, 307, 237–244. (24) Zborowski, M.; Malchesky, P. S.; Jan, T. F.; Hall, G. S. J. Gen. Microbiol. 1992, 138, 63–68. (25) Pekas, N.; Granger, M.; Tondra, M.; Popple, A.; Porter, M. D. J. Magnet. Magnet. Mater. 2005, 293, 584–588. (26) Zimmerman, P. A.; Thomson, J. M.; Fujioka, H.; Collins, W. E.; Zborowski, M. Am. J. Trop. Med. Hyg. 2006, 74, 568–572. (27) Choi, J.-W.; Liakopoulos, T. M.; Ahn, C. H. Biosens. Bioelectron. 2001, 16, 409–416. (28) Ramadan, Q.; Samper, V. D.; Puiu, D. P.; Yu, C. J. Microelectromech. Syst. 2006, 15, 624–638. (29) Evans, C. H.; Tew, W. P. Science 1981, 213, 653–654. (30) Zborowski, M.; Malcheski, P. S.; Savon, S. R.; Green, R.; Hall, G. S.; Nose, Y. Wear 1991, 142, 135–149. (31) Meyer, D. M.; Tillinghast, A.; Hanumara, N. C.; Franco, A. J. Tribiol. 2006, 128, 436–441. (32) Zborowski, M.; Fuh, C. B.; Green, R.; Baldwin, N. J.; Reddy, S.; Douglas, T.; Mann, S.; Chalmers, J. J. Cytometry 1996, 24, 251–259.
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cells.26,33 A system comprising five flow channels capable of processing up to five samples simultaneously was used to isolate an average of one cancer cell (breast cancer carcinoma line MCF7) from 106 lymphocytes.34 The simple design of the device is suitable for miniaturization and multiplexing. In this work, a miniaturized permanent magnet assembly was designed that can provide high magnetic fields and gradients comparable to a previously tested MDM device but with better target cell recovery and improved cell detection. The number of interpolar gaps was increased from one to four in series to maximize the overall cell capture efficiency. Microfabrication techniques were used to construct enclosed microchannels. They provided better control over flow channel dimensions than the previously used medical grade rubber cutout placed between a polymer manifold and a glass34 or Mylar slide.26,33 The magnetic deposition and flow characteristics were investigated using video microscopy and digital image processing instead of the cytochemical staining used previously. The staining had required laborious channel draining and disassembly for cell deposition access, leading to cell losses from the magnetic deposit. Although the multistage MDM design was primarily developed to increase the overall capture efficiency over that of a single stage unit, it was also discovered that it might be possible to trap cells in successive zones according to their magnetic mobilities. MATERIALS AND METHODS Design of the Magnet Assembly. The multistage MDM consists of a microfabricated enclosed channel placed against a multistage permanent magnet assembly (Figure 1a). The magnet assembly consists of four permanent magnet trapezoids, made of neodymium-iron-boron (NdFeB), of staggered polarity and interspersed between high permeability, 1018 carbon steel trapezoids, referred to as pole pieces. The magnetic field lines travel in closed loops from permanent magnet, to pole piece, to the open space above the assembly, to the neighboring pole piece, and back to the permanent magnet. The fringing field in the space above and near the corners of the carbon steel pole pieces gives rise to a high gradient in the magnetic flux density B in these regions. As the force imparted on a magnetically labeled cell depends on the gradient,35 we expect the cells to deposit against the bottom wall of the channel in narrow bands centered above the permanent magnet trapezoids. We refer to these deposition bands as “zones”. The magnetic gradient and, therefore, the force decreases with distance from the magnet assembly. Thus, it was desirable to employ as thin a bottom wall as possible to maximize capture. On the basis of the availability of materials and suitability of microfabrication processes (discussed in the next section), the thickness of the channel walls (top and bottom) was set to 400 µm. 2D magnetostatic simulation software (MAGNETO version 5.1, Integrated Engineering Software, Manitoba, Canada) was used to optimize the length of the interpolar gap, given by the top segment width of the permanent magnet trapezoids, shown in (33) Karl, S.; David, M.; Moore, L.; Grimberg, B.; Michon, P.; Mueller, I.; Zborowski, M.; Zimmerman, P. A. Malaria J. 2008, 7, 1–9. (34) Fang, B.; Zborowski, M.; Moore, L. R. Cytometry 1999, 36, 294–302. (35) Zborowski, M. In Scientific and Clinical Applications of Magnetic Carriers; Ha¨feli, U., Schu ¨ tt, W., Zborowski, M., Eds.; Plenum Press: New York, 1997; pp 205-233.
Figure 1. (a) Schematic of the multistage MDM. The dimensional relationships between the parts is to scale. (b) Interpolar width optimization. The rectangle inside the channel shows the region where the vertical gradient component was calculated. The contour bands illustrate the magnitude of the vertical gradient component, with legend shown on the right (in units of T2/m), for the case of a 1.0 mm interpolar width. Everything is to scale except channel height, which is exaggerated for clarity. (c) Effect of interpolar width on the mean y gradient component in the 0.25 mm × 5 mm rectangle of Figure 1b. The values were calculated from 2D magnetostatic simulation software.
Figure 1b. The aim was to vary the gap width in order to maximize the average value of the magnetic gradient’s y component, dB2/ dy, in the region of deposition. The height and base of each permanent magnet was chosen to be 10 and 6 mm. The mean y component of the gradient was calculated for the region of interest at the deposition zone, highlighted by a rectangle in Figure 1b. The rectangle dimensions correspond to the 250 µm height of the channel and a width of 5 mm centered on the interpolar gap. The rectangle width was selected to be sufficiently broad so that
the field and gradient are negligible at the boundaries. The interpolar width that gave the maximum mean gradient was 1.0 mm (Figure 1c). The magnet assembly was built by Dexter Magnetic Technologies, Elk Groove Village, IL, according to our specifications. Gaussmeter measurements of the manufactured device were then used to calibrate the output of 3D magnetostatic simulation software (Amperes version 5.2, Integrated Engineering Software, Manitoba, Canada). Figure 1b plots contours of the y gradient component, dB2/dy, from Amperes, predicted for the central plane of the 8 mm deep magnet assembly and either of the two middle zones. In the rectangular region, the mean B is 0.370 T and the mean dB2/dy is 518 T2/m. Channel Fabrication. The channel structure was fabricated with SU-8 (MicroChem, Newton, MA), an epoxy-based negative photoresist, on 4 in. diameter, 400 or 500 µm thick Pyrex (Corning 7740, Mark Optics, Santa Ana, CA) substrates. Transparent Pyrex wafers were chosen as the substrate so that deposited cells/ particles could be investigated using optical microscopy. The fabrication process was composed of two major steps: (1) fabrication of open microchannels (250 µm high × 500- or 1000-µm wide) on Pyrex wafers and (2) wafer bonding to form enclosed microchannels. Adhesion of thick SU-8 on Pyrex was known to be less than ideal.36,37 Therefore, a highly cross-linked thin intermediate layer of SU-8 was utilized to improve adhesion.37 Wafer bonding was achievedusingtheSU-8-based,adhesivewaferbondingtechnique.38-42 The wafer bonding process was optimized to avoid common problems associated with SU-8 based wafer bonding techniques, such as clogging of the channels with SU-8 and sensitivity to film thickness variation. In this case, an intermediate layer of SU-8 was applied to a cap wafer, which was then selectively exposed such that the portion that will join with the open section of the channel was cured and hardened; whereas, the portion that will join with the opposing channel wall was left uncured. In this way, SU-8 reflow into the open channel was prevented. However, SU-8 reflow occurs for the uncured portion that opposes the channel walls, to compensate for the thickness variation of the SU-8 structural layer. Magnetic Cells. The leukemia-derived Jurkat cell line was chosen as the model to investigate magnetic deposition. The cells were cultured in RPMI 1640 medium (supplemented with 10% FBS, 292 µg/mL L-glutamine, 50 µg/mL penicillin, and 50 µg/ mL streptomycin) at 37 °C, 80% humidity, and 5% CO2. Predetermined quantities of cells were removed from the culture and washed with buffer solution by centrifugation at 390g, 4 °C for 10 min. Then supernatants were discarded and buffer solution was used to suspend the pellets. The buffer solution was composed (36) Agirregabiria, M.; Blanco, F. J.; Berganzo, J.; Arroyo, M. T.; Fullaondo, A.; Mayora, K.; Ruano-Lopez, J. M. Lab Chip 2005, 5, 545–552. (37) Carlier, J.; Arscott, S.; Thomy, V.; Fourrier, J. C.; Caron, F.; Camart, J. C.; Druon, C.; Tabourier, P. J. Micromech. Microeng. 2004, 14, 619–624. (38) Blanco, F. J.; Agirregabiria, M.; Garcia, J.; Berganzo, J.; Tijero, M.; Arroyo, M. T.; Ruano-Lopez, J. M.; Aramburu, I.; Mayora, K. J. Micromech. Microeng. 2004, 14, 1047–1056. (39) Li, S.; Freidhoff, C. B.; Young, R. M.; Ghodssi, R. J. Micromech. Microeng. 2003, 13, 732–738. (40) Svasek, P.; Svasek, E.; Lendl, B.; Vellekoop, M. Sens. Actuators, A 2004, 115, 591–599. (41) Tuomikoski, S.; Franssila, S. Phys. Scr. 2004, T114, 223–226. (42) Yu, L.; Tay, F. E. H.; Xu, G.; Chen, B.; Avram, M.; Iliescu, C. J. Phys.: Conf. Ser. 2006, 34, 776–781.
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Figure 2. Schematic diagram showing the experimental setup.
of 2 mM EDTA and 0.5% BSA in PBS solution and degassed under absolute pressure of 0.16 atm while continuously stirring for 30 min. The washing step was repeated. The cells were labeled with both magnetic nanoparticles and fluorescent labels using a twostep “sandwich” immunoassay. The washed cell pellet was combined with the primary antibody, anti-CD45-PE (Caltag Laboratories, Burlington, CA) at a concentration of 16 µL of the undiluted reagent per 106 cells. The mixture was incubated for 30 min at 4 °C. After incubation, the cells were washed, in the manner described above, to remove unbound primary antibody. The pellets were then combined with the secondary antibody, antiPE-MACS (Miltenyi Biotech Inc., Auburn, CA) using the same volume as the primary antibody (16 µL of reagent per 106 cells) and incubated for 15 min at 4 °C. The unbound antibodies were removed by another washing step. This was followed by pellet resuspension in the buffer solution. The mobility distribution was evaluated by a technique known as cell tracking velocimetry (CTV).43 The mean and standard deviation in mobility, as evaluated by tracking 1517 cells, were 5.33 × 10-4 and 1.80 × 10-4 mm3/(tesla · ampere · second) or mm3 TAs. Magnetic Particles. The particles were custom-made by micromod Partikeltechnologie GmbH, Rostock, Germany. The particles are intended to be approximate analogues, in diameter and mobility, to many of the labeled cells investigated in our laboratory. Mean diameter measured by Coulter Multisizer II was 6.24 µm, CV < 4%. Iron content was 0.45% (w/w) in the form of magnetite distributed in latex. CTV tracked 486 particles, yielding a mean and standard deviation of 7.29 × 10-4 and 2.38 × 10-4 mm3/TAs, respectively. Experimental Setup. The schematic of the experimental setup is presented in Figure 2. The microfabricated channel overlays the magnetic assembly. The channel was orientated vertically, in order to avoid the interference of gravitational sedimentation on the magnetic cell deposition. The channel/ magnet assembly was attached to an aluminum holder placed on a 2D translation stage (model 462-XY-SD, Newport Corp., Irvine, CA). The 2D translation stage was bolted to a precision laboratory jack (model 271, Newport Corp.), allowing 3D positioning. The (43) Chalmers, J. J.; Zhao, Y.; Nakamura, M.; Melnik, K.; Lasky, L.; Moore, L.; Zborowski, M. J. Magnet. Magnet. Mater. 1999, 194, 231–241.
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translation stage was adjusted in the horizontal direction for focusing the channel. The long working range of the precision jack (4 in.) allowed visualization of all four deposition zones, as well as the channel exit. A horizontally oriented fluorescence microscope (Olympus America Inc., Center Valley, CA) was mated to a 1.4 M-pixel charge-coupled device (CCD) digital video camera (Retiga EXi, QImaging Corporation, Surrey, BC, Canada), with the 5× objective focused on the microchannel such that flow/ deposition images could be acquired in real time. The video camera was interfaced to a PC via firewire (IEEE 1394) cable, to allow direct video streaming of images to RAM at up to 20 frames/ s. One of the independent drives of a dual syringe pump (model “33”, Harvard Apparatus, Holliston, MA) provides the carrier flow. Sample was injected into the channel via a six-way analytical injection valve (Rheodyne model 7725, Rohnert Park, CA) with a handmade sample loop of 100 µL volume. The sample loop was partially filled with 50 µL of sample suspension with a syringe and needle. This use of the injection valve allows pulses of precise and reproducible volumes of sample to be introduced onto a steadily flowing stream of pure carrier. In some experiments, the sample replaced the carrier in the pump’s syringe and the injection valve and sample loop were eliminated, allowing direct infusion into the channel. Image Acquisition. Real-time disk recording software was used to control the camera and acquire the images (Video Savant 4, IO Industries, London, Ontario, Canada). Images were acquired at 5-20 frames per second. Video Savant was also capable of capturing single frames, used for the image processing of deposited particles. Illumination was provided internally (epiillumination) by a 100 W mercury burner. Three different filter cubes allowed viewing by brightfield, darkfield, and fluorescence field. The florescent imaging cube contained a blue excitation filter (470-490 nm), a barrier filter transmitting from 510 nm upward, and a dichroic mirror transmitting from 505 nm upward (Olympus, model U-MNIB). Magnetic Particle Deposition. A volume of 50 µL of sample at different concentrations (105 and 106 particles/mL) were added to the sample loop and injected into the MDM channel (1 mm wide) at 10 µL/min. After the entire sample passed through the MDM, single images of all four deposition zones were captured. The top surfaces of the permanent magnet trapezoids had a darker appearance compared to the steel pole pieces and were easily visible through the transparent channel walls, thus enabling the objective to be focused on the deposition zones. In order to ensure that all the deposited particles at each zone were evaluated, the image size in the x-direction extended 1 mm beyond the upper and lower magnet-pole boundaries. The image size in the zdirection extended to the channel side walls. With the assumptions that the particles were deposited in a single layer and individual particles could be identified, they were manually counted from the prerecorded images. In some case, the particles were clustered in such a fashion that it was difficult to identify individuals. In such cases, the total area of the aggregate was divided by the area of an individual particle to obtain an approximate number. The capture efficiency was calculated by dividing the number of deposited particles by the total number of particles introduced into the system, which was calculated by multiplying the injected volume by the sample concentration. Experiments at each
concentration were repeated six times. The capture efficiency was also evaluated for different flow rates (10-200 µL/min). For these studies, the concentration of the sample was reduced to 104 particles/mL to prevent the accrual of clusters of particles on the deposition zones, and thereby simplify the manual counting. Experiments at each flow rate were repeated three times. Cell Deposition. Labeled cell samples of concentration 105/ mL were introduced at 10 µL/min into the MDM channel (1 mm wide). The cells were infused from the syringe, bypassing the loop, as it was suspected that cells sedimentation in the loop, even for a brief period, causes their aggregation. Images of the deposited cells were acquired under two lighting conditions: bright field and fluorescence field. These images were used to measure the fluorescence intensity of each deposited cell using digital image processing. Digital Image Processing. A customized macro routine within a commercial digital image processing software (Image Pro Plus, v6.1, Media Cybernetics, Silver Spring, MD) was developed to measure the fluorescence intensity of each cell deposited on the four zones. Running the macro resulted in the following sequence: (1) A region of interest (ROI) was selected within the bright-field images. (2) The appearance of cells was enhanced with edge-emphasizing, large spectral filters. (3) The resulting images were median-filtered to remove noise and thresholded by intensity and area (to remove debris and large objects not consistent with cell size). This step generated binary masks of segmented cells. (4) A watershed filter was applied to split apart the cells that were connected. (5) A product of cell mask sets and corresponding cell fluorescence image sets were used to produce images that consisted of fluorescently labeled cells that are isolated from background and nonspecific fluorescence (replaced with black pixels) but still retained their original intensity levels. Objects not consistent with average cell size were removed via size/shape exclusion. (6) Given an intensity range of 0-255 (8-bit) gray-levels, the mean intensity for each cell in the masked images was calculated and exported to a Microsoft Excel spreadsheet. The process of uploading the image to Image Pro and obtaining the data spreadsheet took less than a minute per image. Image Pro’s Plus version also features a cell counting module which counts the deposited cells automatically based on the filtered and thresholded images. RESULTS AND DISCUSSION Air-Liquid Interface. Video microscopy allowed capturing the deposition process in real time. The magnetic particle accumulation at the deposition zones was observed as it occurred. When the experiment was repeated at the same flow rate with nonmagnetic polystyrene particles, no significant accumulation of particles was observed. The retention of deposited particles with the passage of an air-liquid interface was investigated using video microscopy in a 500 µm wide channel. After particles accumulated on the deposition zones, a large bubble was introduced into the channel. Observing the real-time video of the passage of the bubble through the deposition zone revealed that a large fraction of the particles was swept away by the air-liquid interface (A real time video is presented in the Supporting Information.). This highlights a limitation of conventional MDM devices. Because the channel must be dismantled in order to recover the deposited cells for analysis, the process of draining the channel causes an
air-liquid interface to pass across the deposition zone, producing a substantial loss of deposited particles. The alternate method applied here was to work exclusively with enclosed microchannels and perform analysis of the deposited cells in situ. In a setting without a translation stage and horizontal microscope, one could run the experiment with the channel/magnet assembly in the vertical position and afterward close inlet and outlet valves, thus keeping the channel filled with buffer as it is transported elsewhere for acquiring images. Effect of Loading. The effect of particle loading on capture efficiency was investigated by injecting 50 µL of micromod particles of concentration 105 and 106 per mL, resulting in 5 000 and 50 000 particles infused. Particles were observed to deposit only at the four deposition zones. By manual counting, the total capture efficiency summed over the four zones was found to be 79.8 ± 6% and 84.6 ± 9%, respectively. As the particles were less than a monolayer deep in both cases, there could be no significant channel overloading. A single layer of particles occupies only about 2.5% of the channel height. The higher capture efficiency at higher particle number suggests a potential cooperative effect, with deposited particles helping to capture and retain subsequent particles, by a combination of magnetic and Stokes drag forces.44 Manual versus Automated Counting. We next investigated using ImagePro as an alternative to the labor-intensive method of manual counting. Five different images of deposited magnetic cells, selected from the flow rate versus capture efficiency study described above, yielded manual counts varying from 260 to 1450 cells. Image Pro counts differed from the manual counts by no more than 3.9%. Linear regression of the Image Pro counts versus manual counts yielded a slope near unity with good correlation: Image Pro counts ) 1.03(manual counts) + 1.33; N ) 6; R2 ) 0.9990, R ) 0.05. Subsequently, the automated method replaced the manual method of counting whenever possible. Cell Deposition. We performed experiments with labeled cells to observe the effect of entrapment as a function of axial position, which was related to zone number. Magnetic deposition was observed only near the four magnetic zones, and cell deposition in the space between the zones was not significant. When unlabeled cells were passed through the channel, no significant deposition was observed in any part of the channel. The total number of cells introduced was kept to less than ∼104 per experiment, as higher numbers might have caused the cells to be deposited in multiple layers, complicating the counting process. No injection loop was used, and the sample was fed from a pumpmounted syringe for approximately 10 min, at 10 µL/min, and a concentration of 105 cells/mL. To evaluate capture efficiency, the total number of cells introduced into the system was calculated by adding the total number of cells deposited to the number that escaped through the outlet. This was considered more accurate than simply evaluating the product of injection time and flow rate, due to the uncertain retention of sample in the inlet tubing and connectors. To determine the number of escaped cells, the channel outlet was continuously imaged at 10 fps during the course of the experiment. The images were subsequently played back to manually count the cells. After completion of the separation, single images of each of the four zones were acquired. (44) Mikkelsen, C.; Hansen, M. F.; Bruus, H. J. Magnet. Magnet. Mater. 2005, 293, 578–583.
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Table 1. Cell Count and Mean Cell Fluorescence Intensities (FIs) in Zones 2-4 for Different Experiments with 1 × 105 labeled Jurkat Cells/mL at 10 µL/min Flow Rate expt no. 1
expt no. 2
expt no. 3
zones
cell count
mean cell FIa
cell count
mean cell FI
cell count
mean cell FI
2 3 4
179 28 21
50.9 ± 24.0 46.8 ± 16.9 23.2 ± 18.9
189 42 25
51.3 ± 21.8 34.0 ± 20.2 22.5 ± 19.1
1018 237 96
87.6 ± 28.1 38.4 ± 23.4 49.1 ± 22.2
a The differences between the mean cell FIs in the deposition zones 2-4 were significant (p e 0.01) except between zones 2 and 3 in experiment no. 1 and zones 3 and 4 in experiment no. 3.
Figure 3. Capture efficiency (b) and fluorescence intensity measured as 8-bit gray scale value (vertical bar) as a function of zone number. The error bars show 1 standard deviation from the mean for three experiments.
Figure 4. Microphotographs of different stages of the digital image processing: (a) original bright field image, (b) segmented image after applying filter, (c) original fluorescent image, and (d) image after the background is isolated.
From these images, the capture efficiency in each zone was determined from Image Pro analysis and plotted in Figure 3. The error bars represent 1 standard deviation calculated from three trials. On the basis of three experiments, the total capture efficiency for the magnetically labeled cells was 99.46 ± 0.26%. Fluorescence Intensity Analysis. We postulated that the multistage MDM might offer the secondary benefit of separating cells by composition between the zones. The Image Pro Plus macro was used to measure the mean 8-bit gray scale value of each deposited cell. Figure 4 shows an example of an image at different stages of the image processing steps. Properties of individual cells became easier to measure after applying the macro that improved their resolution from the background. The gray scale value (pixel intensity) should be proportional to the 48
Analytical Chemistry, Vol. 81, No. 1, January 1, 2009
florescence intensity of the images, which should likewise be proportional to magnetophoretic mobility, as described earlier.22 Zone 1 was excluded from this analysis, as the line graph in Figure 3 shows almost 90% of the captured cells appeared in this zone. In spite of our efforts to avoid overloading, zone 1 was too overloaded to allow accurate florescence intensity analysis. The mean and standard deviation of the pixel intensity for the other three zone populations are listed in Table 1. The average fluorescence intensities for zones 2 to 4 are presented as bar graphs in Figure 3. They follow a trend of decreasing with increasing zone numbers. We performed a two-sample unequal variance, one-tailed Student’s t-test for a significance level of R ) 0.05 to determine if the mean florescence intensity of cells captured in different zones were significantly different. The p values from a one-tailed t-test were calculated using the function “TTEST” in Microsoft Excel. In all three experiments, when comparing zones 2 and 4 we find that the p values are
