Visual chemotaxis assays are essential for a better understanding of how eukaryotic cells control chemoattractant-mediated directional cell migration. Here, we describe detailed methods for: 1) real-time, high-resolution monitoring of multiple chemotaxis assays, and 2) simultaneously visualizing the chemoattractant gradient and the spatiotemporal dynamics of signaling events in neutrophil-like HL60 cells.
Eukaryotic cells sense and move towards a chemoattractant gradient, a cellular process referred as chemotaxis. Chemotaxis plays critical roles in many physiological processes, such as embryogenesis, neuron patterning, metastasis of cancer cells, recruitment of neutrophils to sites of inflammation, and the development of the model organism Dictyostelium discoideum. Eukaryotic cells sense chemo-attractants using G protein-coupled receptors. Visual chemotaxis assays are essential for a better understanding of how eukaryotic cells control chemoattractant-mediated directional cell migration. Here, we describe detailed methods for: 1) real-time, high-resolution monitoring of multiple chemotaxis assays, and 2) simultaneously visualizing the chemoattractant gradient and the spatiotemporal dynamics of signaling events in neutrophil-like HL60 cells.
Eukaryotic cells sense and move toward the higher concentration within a chemoattractant gradient, a cellular process referred as chemotaxis. Chemotaxis plays critical roles in many physiological processes, such as embryogenesis1, neuron patterning2, metastasis of cancer cells3, recruitment of neutrophils to sites of inflammation4, and the development of the model organism Dictyostelium discoideum5. In general, eukaryotic cells sense chemoattractants using G protein-coupled receptors5. The engagement of chemoattractants with these receptors promotes dissociation of the heterotrimeric G proteins Gα and Gβγ, which in turn activate downstream signal transduction pathways that ultimately regulate the spatiotemporal organization of the actin cytoskeleton to drive cell migration5-9.
Cell biologists have been developing and improving chemotaxis assays to examine how G protein-coupled receptor (GPCR) signaling mediates directed cell migration. The Boyden chamber or transwell migration assay was developed in 1960 by Boyden10. The assay works by creating a gradient of chemoattractant compounds between two wells that are separated by a microporous membrane. Its simplicity and ease of use make it the most widely used chemotaxis assay to date. However, the assay does not enable the migrating process of the cells to be visualized. The Zigmond chamber is the first visual microfluidic device that allows clear imaging of cell migration on a coverslip across a narrow constriction toward a source chemoattractant11. Dunn12 and Insall13 modified and improved the high-resolution and long-term imaging capability of the Zigmond chamber chemotaxis assay. Because of the highly predictable, diffusion-dominant characteristics of fluid flow, microfluidics has been providing solutions for next-generation chemotaxis assays such as EZ-TAXIScan (a cell mobility analysis device).
With the stability of the gradient ensured, the device allows six chemotaxis assays to be carried out simultaneously (Figure 1A). In contrast to the directionally fixed gradients generated in the various chamber assays above, the needle or micropipette assay developed by Guenther Gerisch generates a gradient with a movable source14. In the assay, chemoattractant is released from a movable micropipette to generate a stable gradient. With this needle assay, researchers found that different cells generate pseudopods with fundamentally different characteristics. Applying fluorescent microscopy, we were able to visualize the gradient to facilitate its quantitative measurement throughout15. In this study, we describe detailed methods for preparing chemotactic HL60 (human promyelocytic leukemia) cells, simultaneously monitoring multiple chemotaxis assays with the cell mobility analysis device, and visualizing the GPCR-mediated spatiotemporal dynamics of signaling molecules such as protein kinase D1 in single live cells in response to visible, spatiotemporally controllable chemoattractant stimuli. Our advanced imaging methods can be applied to general chemotaxis studies, and are especially suitable for mammalian cell systems.
1. Culture and Differentiation of Human Neutrophil-like HL60 Cells
2. Coating the Cover Glass Surface of the 4well Chamber
3. Chemotaxis Assay Using Cell Mobility Analysis Device
4. Transfection with Electroporation
5. Monitoring GPCR-mediated Membrane Translocation of PKD1 by Multi-channel Fluorescent Microscopy
6. Imaging Chemotaxing Cells in Visible and Controllable Chemo-attractant Stimuli
Simultaneous imaging of chemotaxis of multiple HL60 cells using cell mobility analysis device
Based on the principle of microfluidics16, the manufacturer has provided simulated profiles of gradients: a gradient is generated within 1 min, stabilized within 5 min, and maintained over 2 hr. The highly predictable profiles of the stable gradients generated by microfluidics allow multiple chemotaxis assays to be carried out simultaneously. In the present study, we observed three simultaneous chemotaxis assays (Figure 2A and Movie 1). We found that HL60 cells started chemotaxing immediately after the chemoattractant was injected into the well of the chemoattractant, and kept chemotaxing in a straight path for the following 60 min, consistent with the simulation results for gradient stability. Tracing the travel path and morphology of the cells allows quantitative measurements and subsequent comparison of the chemotaxis behaviors using a chemotaxis index that includes total path length, speed, directionality, and roundness of the cells (Figure 2B). Total path length is the sum of the lengths of the line segments connecting the centroids of the path. Speed is obtained by dividing the total path length by the time. Directionality is measured upward and is defined as: (Y coordinate of the end of the path minus Y coordinate of the beginning) divided by total path length. This gives 1.0 for an object moving directly upward. The roundness of the cell is a measure (in percent) of how efficiently a given amount of perimeter encloses area. A circle has the largest area for any given perimeter and has a roundness parameter of 100%. A straight line encloses no area and has a roundness parameter of 0%. We show the quantitatively measured chemotaxis behavior as described by the selected chemotaxis parameters (Figure 2C).
Monitoring PKD subcellular localization in HL60 cells under a spatiotemporally visible and controllable fMLP stimulus
It is a great technical advance to apply fluorescently labeled and controllable chemoattractant stimulation to an experimental system. Historically, we have applied either homogeneous (also called uniform) stimulation or gradient stimulation to observe cell response and behaviors. However, "blind" stimulation not only provides no spatiotemporal information on how the stimulus reaches the cells, but also casts doubt on any "abnormal" observations of cell response to stimulation, simply because we do not see the stimulus. We have previously shown that fluorescent dye (Alexa594) can be applied with chemoattractant to establish a linear relationship between chemoattractant concentration and monitored fluorescent dye intensity15. With an acquisition configuration of green fluorescent protein (GFP), a red emission of fluorescent dye (Alexa594), and transmitted light, we are able to monitor the adhering cells, the application of the stimulus, and the cell response to the stimulus (Figure 3A). Protein kinase D is a family of serine/threonine kinases that play essential roles in directed cell migration9,17. In response to uniformly applied fMLP (red) stimulation, HL60 cells mediate a robust membrane translocation of GFP-tagged protein kinase D1 (green) (Figure 3B and Movie 2). In an fMLP gradient (red) (Figure 4A), HL60 cells actively recruit PKD1 to the leading edge (Figure 4B and Movie 3). A close comparison of the subcellular localization of GFP in the protrusion of the leading edge indicates that PKD1 localizes at the rear of the leading edge (Figure 4C).
Figure 1. Cell mobility analysis device allows up to 6 simultaneous chemotaxis assays. (A) Scheme shows the design of a cell mobility analysis device chip for simultaneous monitoring of 6 independent chemotaxis assays. Red shows chemoattractant added to the wells. (B) Introduction of HL60 cells to the wells of cells while the chemoattractant diffuses to establish a steady fMLP gradient. Please click here to view a larger version of this figure.
Figure 2. Simultaneous monitoring of multiple chemotaxis assays with HL60 cells. (A) Montage shows images of cell mobility analysis device chemotaxis assay to examine the inhibitory effects of PKD-specific inhibitors on chemotaxis at times of 0 and 12 min after gradient application. Chemotactic HL60 cells were pre-treated with PKD inhibitor 1 μM CID755673 for 30 min. HL60 cells with or without the treatment of PKD inhibitor were allowed to chemotax in either RPMI1640 starving medium or 100 nM fMLP gradients for 12 min. (B) Scheme shows the travel path length and morphology of traced HL60 cells. (C) Quantification of chemotaxis as total path length, speed, directionality, and roundness. Mean ± SD is shown; n = 10, 12, or 11 for no gradient, fMLP gradient without CID755673 treatment, and fMLP treatment with CID755673 treatment, respectively. Please click here to view a larger version of this figure.
Figure 3. GPCR-mediated robust membrane translocation of PKD1 in response to uniformly applied fMLP stimuli. (A) Multichannel monitoring of PKD1-GFP (green), chemoattractant (1 μM fMLP mixed with 0.1 μg/ml fluorescent dye Alexa 594, red), and DIC (differential interference contrast) to identify the adhering HL60 cells in a well of a 4well chamber coated with 0.2% gelatin in RPMI 1640 medium. Scale bar = 10 μm. (B) Montage shows that uniformly applied fMLP (red) induces robust membrane translocation of PKD1-GFP (green). Please click here to view a larger version of this figure.
Figure 4. Leading edge localization of PKD1 in chemotaxing HL60 cells. (A) Channel mode acquisition configuration facilitates the visualization of the fMLP gradient and the spatiotemporal dynamics of PKD1. In A–C, HL60 cells transiently expressed GFP-tagged PKD1; to visualize the fMLP gradient generated from a micropipette (DIC), 100 nM fMLP (Red) was mixed with 0.1 μg/ml fluorescent dye Alexa 594. (B) Enriched localization of PKD1 at the leading edge of the chemotaxing cell. Scale bar = 10 μm. (C) Merged images show that PKD1 localizes at the rear of the leading edge in HL60 cells. Green shows PKD1 cellular localization, and the DIC image shows the protruding area of the leading edge. Scale bar = 5 μm. Please click here to view a larger version of this figure.
In the present study, we show two examples of chemotaxis assays: first, simultaneous monitoring of multiple chemotaxis assays by the cell mobility analysis device; and second, visualization of the chemoattractant gradient and the spatiotemporal dynamics of signaling events in the same cells in real time.
Cell mobility analysis device for multiple simultaneous chemotaxis assays
In the present study, we introduced a detailed protocol to perform multiple simultaneous chemotaxis assays using a cell mobility analysis device. This device allows user to observe cellular chemotaxis behavior with a 10X objective lens in conventional bright-field observation. Because of the diffusion-dominant characteristics of the fluid flow, the cell mobility analysis device generates highly predictable, stable gradients and allows up to six chemotaxis assays to be carried out simultaneously. The critical steps of the protocol are to obtain reliable gradients and to align the cells on the terrace line. The user should strictly follow the manufacturer's instructions for holder assembly and injection of chemoattractants and cells. Detailed instructions are also available online. Compared to alternative chemotaxis methods11-13,15, this device significantly improves the reliability and efficiency of chemotaxis assays. Four sizes of cell mobility analysis device chip in sizes of 4, 5, 6, and 8 μm are available to accommodate different types and sizes of cells. We found that a 4 or 5 μm cell mobility analysis device chip is suitable for the HL60 and D. discoideum cells, which are about 10-15 μm in diameter. However, one limitation is that this device is not suitable for all types of cells. We had little success using the cell mobility analysis device chemotaxis assay with Raw267.4 cells. The reason may be that Raw267.4 cells migrate too slowly. The time required for efficient chemotaxis of Raw267.4 cells might be much longer than the time a gradient is maintained by the device. Instead, a transwell migration assay worked well for Raw267.4 cells9. Another limitation is that fluorescent observation is not possible with the current device. A future direction is to monitor fluorescent imaging with higher magnification. This is possible with the improved cell mobility analysis device, which is equipped with fluorescent detection and a 100X objective lens. In addition, the number of simultaneous assays is also increased to 12. All these improvements facilitate the enhanced throughput of chemotaxis assays and the observation of subcellular dynamics in migrating cells.
High transfection efficiency of HL60 cells to express fluorescent protein-tagged protein
HL60 cells are an actively dividing leukemia cell line and grow in suspension. As previously reported18-20, HL60 cells are resistant to gene transfer. Both lipid and electroporation gene transfer have been tested, and higher transfection efficiency was obtained with electroporation, as described in detail in section 4. Obtaining high viability after electroporation is critical for obtaining high transfection efficiency because of the severe damage that results from electroporation. Subsequently, thorough and gentle cell handling are required, especially after electroporation. All media have to be pre-warmed and gently added to the cells in any steps after electroporation. It is also critical to minimize the exposure time of cells to the electroporation reagent. To avoid cell death, after electroporation, RPMI 1640 electroporation recovery medium must be added to the cells immediately. We found that 20% FBS in the recovery medium gives a much higher cell recovery rate than 10% FBS. After electroporation, incubation in RPMI 1640 electroporation recovery medium for 30 min is critical for greater viability and transfection efficiency. To further increase transfection efficiency, we also used 4 μg plasmid DNA per transfection, which is almost the twice amount of plasmid recommended by the manufacturer.
There are two major limitations on electroporation transfection: the transiency of the protein expression and the cell number limit (2 x 106 cells per transfection). In undifferentiated cells, expression is detectable only during the first couple of cell divisions, since the vector plasmid is diluted by half after each cell division. For The differentiated HL60 cells survive no more than 48 hr. As a result, any experiment requires fresh transfections 6 hr prior to the experiment. The cell number for one electroporation merely meets the minimum requirement for any biochemical assays. For the purpose of repetitive use or large quantity, it is strongly recommended that an undifferentiated HL60 cell line be established that stably expresses the protein of interest which has been tagged with a fluorescent protein by a viral vector, if a viral vector is available or can be constructed.
The authors have nothing to disclose.
This work is supported by the intramural fund of NIAID, NIH.
RPMI 1640 Medium GlutaMAX | Life technologies | 61870-036 | |
Sodium pyruvate | Thermo Fisher Scietific | 11360-070 | |
Fetal bovine serum | Gemini Bio-Products | 100-106 | |
1M HEPES sterile solution, pH7.3 | Quality Biological Inc. | A611-J848-06 | |
Penicillin streptomycin solution | Fisher Scientific | 15140122 | |
NucleofectorTM 2b | Lonza | AAB-1001 | |
AmaxaTM Cell Line NucleofectorTM Kit V including NucleofectorTM Solution, Singe use pipettes, AmaxaTM certified 100 ml aluminum electrode cuvettes | Lonza | VCA-1003 | |
Lab-Tek chambered #1.0 Borosilicate Coverglass | Nalge Nunc International Inc | 155383 | |
2 % Gelatin solution | Sigma-Aldrich | G1393 | |
Fibronectin | Sigma-Aldrich | F1141 | |
HBSS (Hanks’ Balanced Salt Solution) | Life technologies | 14025-076 | |
Bovine serum albumin | Sigma-Aldrich | A3803 | |
Single well Lab-Tek II coverglass chambers | Nalge Nunc International Inc | 155361 | |
Four-well Lab-Tek II coverglass chambers | Nalge Nunc International Inc | 155383 | |
Alexa 594 | Thermo Fisher Scientific | A-10438 | |
fMLP | Sigma -Aldrich | F3506-5MG | |
Cover glass thickness 2 22 x 22 mm | Corning | 2855-22 | |
EZ-TAXIScan | Effector Cell Institute, Inc. | MIC-1001 | |
EZ-TAXIScan chip (5 mm) | Effector Cell Institute, Inc. | EZT-F01-5 | |
1701RN 10ul syringe | Hamilton | 80030 | |
Femtotips II Injection tips | Eppendorf | 5242956003 | |
Femtotips II | Eppendorf | 930000043 | |
TransferMan NK2, including motor module, X head with angle adjuster, and Positioning aids. | Eppendorf | 5188900056 | |
DIAS software | Solltech Inc. | ||
LSM 780 META or equivalent confocal microscope with a 40X 1.3 NA or 60X 1.4 NA oil DIC Plan-Neofluar objective lens | Carl Zeiss |