Sequential Array Cytometry: Multi-Parameter Imaging with a Single Fluorescent Channel
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- Gossett, D.R., Weaver, W.M., Ahmed, N.S. et al. Ann Biomed Eng (2011) 39: 1328. doi:10.1007/s10439-010-0199-8
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Heterogeneity within the human population and within diseased tissues necessitates a personalized medicine approach to diagnostics and the treatment of diseases. Functional assays at the single-cell level can contribute to uncovering heterogeneity and ultimately assist in improved treatment decisions based on the presence of outlier cells. We aim to develop a platform for high-throughput, single-cell-based assays using well-characterized hydrodynamic cell isolation arrays which allow for precise cell and fluid handling. Here, we demonstrate the ability to extract spatial and temporal information about several intracellular components using a single fluorescent channel, eliminating the problem of overlapping fluorescence emission spectra. Integrated with imaging technologies such as wide field-of-view lens-free fluorescent imaging, fiber-optic array scanning technology, and microlens arrays, use of a single fluorescent channel will reduce the cost of reagents and optical components. Specifically, we sequentially stain hydrodynamically trapped cells with three biochemical labels all sharing the same fluorescence excitation and emission spectrum. These markers allow us to analyze the amount of DNA, and compare nucleus-to-cytoplasm ratio, as well as glycosylation of surface proteins. By imaging cells in real-time we enable measurements of temporal localization of cellular components and intracellular reaction kinetics, the latter is used as a measurement of multi-drug resistance. Demonstrating the efficacy of this single-cell analysis platform is the first step in designing and implementing more complete assays, aimed toward improving diagnosis and personalized treatments to complex diseases.
KeywordsCytometry Microfluidics Single-cell Immunocytochemistry High content screening Cellomics Drug discovery Personalized medicine Point-of-care
High content screening (automated microscopy and image analysis) is a powerful tool for drug discovery, diagnostics, and biomedical research.1 Automated measurement of temporal and spatial information about targeted cellular processes can help elucidate drug–target interactions in vitro,10,22 identify the presence of rare abnormal cells in a tissue or blood,19 and enable high-throughput experimentation.1,4 Microscopy techniques are compatible with living cells, and working with biomolecules in their cellular microenvironment provides more accurate information about their function and molecular mechanisms. In contrast to microscopy techniques, bulk measurement tools such as plate readers or Western blots can produce misleading averages of populations and mask behaviors of rare abnormal cells or subpopulations.8 However, when performed manually, the throughput of single-cell microscopy measurements is low. Automation can increase the quantity of measurements and enhance reproducibility by limiting user bias. Current approaches to automation through robotics (used for high content screening) have been cost prohibitive and remain out of reach for point-of-care diagnostics, personalized medicine, and academic use. Further, the number of independent parameters that can be measured with these tools can be limited by overlap of fluorescence spectra.13 Immunophenotyping, for example indentifying T-cell subpopulations, stemness, or circulating cancer cells, often requires the identification of multiple biochemical parameters.2,3,13,19 Moreover, dynamic processes such as drug permeability through a cell monolayer may be best characterized by a temporal parameter.15,20 While there is much effort toward expanding the capabilities of the scanning microscope and other currently used cytometric methods, such as flow cytometry,3,17 several other tools and techniques are being developed which aim to bring down cost and expand access through miniaturization and simplification. High-throughput, parallel fluorescence detection has been achieved by an integrated zone-plate array.18 Fiber-optic array scanning technology can scan substrates 500 times faster than conventional scanning micrscopy.12 Wide field-of-view lens-free fluorescent imaging on a chip is another alternative to mechanical scanning lens-based systems.6 This compact technology has achieved ~10 μm spatial resolution over an 8-cm2 field of view with a single image.5 Many of these techniques are well equipped to identify rare, single-cell events. However, positive and negative identifications may require a composite overlay of several signals demonstrating colocalization and some of these techniques are currently limited to a single wavelength. In this study we propose a technique called “Sequential Array Cytometry” which will release these tools from these restrictions.
Materials and Methods
Microfluidic Channel Fabrication
Microfluidic channels were fabricated by replica molding.9,21 In brief, a two-layer mold was constructed in SU-8 50 (MicroChem Corp., Newton, MA, USA) using standard photolithographic methods. A transparent, elastomeric polymer, polydimethylsiloxane (PDMS; Sylgard 184 Silicone Elastomer, Dow Corning Corp., Midland, MI, USA), was cast over the mold and cured at 65 °C for at least 3 h. The PDMS could then be removed from the mold and inlet and outlet holes were punched with a pin vice assembly (Technical Innovations, Inc., Angleton, TX, USA). The molded side of the PDMS and a slide glass were activated with air plasma for 30 s then placed in contact to form a permanent bond. Minimal pressure was applied to ensure contact, but care was observed to not collapse raised channel features onto the glass. The latter would prevent necessary flow through the hydrodynamic cell traps. The completed channels were left to bake at 65 °C for at least 5 min prior to use. Later, the inlet and outlet of the channel could be fitted with polymeric tubing and have fluid driven through them by a syringe pump. The top surface of the PDMS was sterilized with 70% ethanol then washed with sterile water.
Solutions and Cell Lines
The fluorescent stain solutions used were 2 μM calcein AM (Invitrogen Corporation, Carlsbad, CA, USA), 1.5 μM SYTO 16 green fluorescent nucleic acid stain (Invitrogen), and 25 μg/mL FITC-conjugated Lectin from Triticum vulgaris (wheat germ agglutinin; Sigma-Aldrich Corp., St. Louis, MO, USA). 100 μM fluoxetine (Cerilliant Corporation, Round Rock, TX, USA) was used as an inhibitor of drug-resistance transporters. The HeLa cell line (epithelial; human cervical adenocarcinoma) was propagated in Dulbecco’s Modification of Eagle’s Medium (DMEM) 1× with l-Glutamine, 4.5 g/L Glucose and Sodium Pyruvate, 10% fetal bovine serum (FBS), and 1% Penicillin/Streptomycin at 37 °C and 5% CO2. The Caco-2 (epithelial; human colorectal adenocarcinoma) cell line was propagated in DMEM 1× with l-Glutamine, 4.5 g/L Glucose and Sodium Pyruvate, 20% FBS, and 1% Penicillin/Streptomycin at 37 °C and 5% carbon dioxide (CO2). For sequential array cytometry, both cell lines were released from tissue culture flasks with 0.25% Trypsin and resuspended in their culture media. For measurements of noise and saturation limit 9.9 μm fluorescent polymer microspheres (Thermo Scientific, Waltham, MA, USA) were used.
Hydrodynamic Cell Trapping and Solution Exchange
Channels (see Fig. 1) were prefilled with 1× phosphate buffered saline (PBS) by injecting PBS through the outlet (outlet during the rest of the protocol) with a syringe pump at a flow rate of 20 μL/min. The inlet was obstructed with a metal pin during this time to eject air through the air-permeable PDMS walls. Subsequently, the pin was removed, and a 20-μL drop of PBS was loaded over the inlet. The flow direction on the syringe pump was reversed such that fluid was drawn from the drop through the channel. The drop was replenished before any air could enter the inlet. For cell loading, a suspension of cells was added dropwise to the inlet and the flow rate was reduced to 10 μL/min (optimized for cell trapping). For all other solutions and wash steps 10 μL/min was used.
Image Capture Post-Processing
Channels were mounted on an inverted microscope stage and secured. The appropriate axial position and exposure times for the selected stains were chosen prior to experiments. Ambient light was minimized. Grayscale fluorescent images were captured every 30 s using an automated time-lapse function in the image acquisition software (NIS Elements; Nikon Instruments Inc., Melville, NY, USA). In-between exposures the shutter was closed to minimize bleaching. Image processing was carried out in NIS Elements and ImageJ (http://rsbweb.nih.gov/ij/). The final image of the calcein AM staining process was used to identify the address and size of every cell within the imaged portion of the array. The mean intensity at every address in defined regions of interest (ROIs), which encompass the cell area at each address, was stored over time for kinetics experiments. For multi-parameter image reconstruction an image was selected from each staining step. The images were opened in ImageJ. Using the “Image Calculator” function sequential images were subtracted from one another to obtain only the staining that occurred during that interval. These were pseudo-colored different colors using the “Lookup Tables” function and overlayed in NIS Elements.
As stated above, the mean intensity per area for each cell was extracted from the images at every time point. A two-tailed t-test assuming unequal variances was carried out comparing the mean of control and experimental conditions.
Low cost cytometric instruments for personalized medicine and point-of-care diagnostics will require simplified optics and operation speeds faster than current state-of-the-art automated microscopy and flow cytometry. Even these powerful instruments can be limited by the overlap of fluorescence emission spectra while some of the new imaging techniques for these applications are currently limited to a single wavelength. Here, we present a method which removes both of these limitations. We demonstrate the ability to create composite images, but more importantly the ability to identify colocalized parameters in a way that is not limited by spectral overlap. It remains to be shown the number of independent parameters that can be obtained by sequential array cytometry, but the theoretical limit with our current optical setup is approximately 60 components (our measured saturation number of levels divided by 4σ). It will depend on the dynamic range of the detection scheme, the specificity of fluorescent probes, the quality of solution exchange, and the length of time that cells remain intact while trapped. The photostability of fluorescent probes should be considered when designing sequential array cytometry protocols; the shutter is open only briefly (milliseconds to seconds) at each exposure time to minimize photobleaching, but over long periods of time bleaching of an unstable stain could result in inaccuracies if not taken into account. Alternatively, after imaging, bleaching could be actively used to drastically reduce the intensity and noise contribution of the previous stain, enabling a further increase in the number of stains possible.
We thank Aydogan Ozcan and Ahmet Coskun for helpful discussions and Henry T. K. Tse for assistance with automated data processing.
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