The overall goal of this experiment is to determine the absolute protein copy number of a target protein with single-cell resolution. Single cell techniques will enable a better understanding of the behavior of a protein as it relates to a central dogma and help toward accurate quantitation of the controlled gene expression. Working at the single-cell level can capture the rich heterogeneity of the cell population that would otherwise be lost by the averaging that occurs with traditional biochemical techniques.
Generally, individuals new to this method will struggle because of the sensitivity of single-cell assays or a limited amount of protein in each cell. We first had the idea for this method when we wished to perform a single-cell protein assay that could be easily sampled for further analysis down the stream. Begin this procedure by making chips and printing antibody microarrays as described in the text protocol.
To prepare the apparatus, feed a 100-millimeter length of 150-micron inner diameter 360-micron outer diameter fused silica tubing to a 40-millimeter length of 1-millimeter inner diameter 1/16-inch outer diameter PFA tubing until it protrudes by two millimeters. This will form the concentric nozzle. Apply a thin layer of cyanoacrylate glue to the end of a 10-microliter pipette tip and insert it into the 40-millimeter piece of PFA tubing.
If required, reposition the fused silica tubing to maintain a 2-millimeter protrusion at the nozzle before the adhesive sets. Insert the other end of the fused silica capillary into the end of a 200-millimeter length of 014-inch inner diameter 062-inch outer diameter PTFE tubing. Connect this to a 100-microliter Hamilton syringe filled with four percent bovine serum albumen in phosphate-buffered saline.
Next insert a 400-millimeter length of 1-millimeter inner diameter 2-millimeter outer diameter FEP tubing into a 200-microliter pipette tip until it forms a seal. Apply a thin layer of cyanoacrylate glue to another 200-microliter pipette tip, then push this into the first tip to fix the tubing in place. Then connect the open end of the FEP tubing to a 1-milliliter Hamilton syringe filled with mineral oil.
Insert the 200-microliter pipette tip assembly into the 10-microliter pipette tip of the concentric nozzle. Next, fill the 100-microliter syringe with 4-percent PBSA blocking solution. Reattach and flush the aqueous tubing with the blocking solution.
Repeat twice for a total of three flushes. Then fill the 100-microliter syringe with 0.125-micrograms-per-liter detection antibody in 4-percent PBSA and reattach the tubing. Replace the blocking solution in the tubing by dispensing 25 microliters of detection antibody solution.
Flush the oil tubing with mineral oil until all tubing and the nozzle fills with oil. Then refill the 1-milliliter syringe with mineral oil and reattach the tubing. Finally, secure the tubing and nozzle assembly to an XYZ manipulator.
To form addressable droplets, secure the chip on the microscope stage. Record the microscope's stage coordinates of each spot in the array using the automated microscope control software. Next, set the XYZ manipulator on the microscope stage.
Place the syringes in separate syringe pumps as they will need to be operated independently. Place the nozzle at an angle of 50 to 60 degrees. Then set the aqueous syringe pump to dispense 10 nanoliters of 100 microliters per minute, and set the oil syringe pump to disperse 5 microliters at 100 microliters per minute.
Dispense the aqueous solution until a bead of fluid is visible at the head of the nozzle. Dab the bead with a dust-free wipe to remove it. Using the automated microscope control software, set the microscope stage coordinates to an antibody spot in the array and focus on the cover slip surface.
When aligning the nozzle, be careful to not disturb the antibody spot. Take extra care and make sure you observe the approach of the glass capillary tip through a microscope. Being careful not to disturb the spot, use the XYZ manipulator to align the glass capillary tip of the concentric nozzle to the side of the antibody spot.
Once aligned, dispense 10 nanoliters of aqueous solution. Without moving the stages, dispense five microliters of oil to cap the aqueous solution. Slowly raise the nozzle clear of the droplet and move to the next well.
Repeat this step for all antibody spots in the array. After 30 minutes, image all spots in the array using the automated microscope control software to determine the background of single molecules bound to each antibody spot prior to loading cells. To load the addressable droplets with cells, we suspended the cells in a solution of 0.125-micrograms-per-milliliter detection antibody in L-15 media with 10 percent FDS.
Filter the cell solution through a 40-micron-pitch cell strainer to ensure a single-cell suspension. Load the single cells into the addressable droplets from the cell reservoir using the micromanipulator and microcapillary. Observing through a 10x objective, use the microinjector to aspirate a single cell into the microcapillary from the cell reservoir.
Working manually with a joystick, pour automatically using the eject feature of an electronic manipulator stage. Reject the micropipette by translating it upwards to clear the 1-millimeter height of the chip. Set the stage coordinates to that of an addressable droplet using the automated microscope control software.
Inject the micropipette by returning it to the stored Z position. Perform this manually with the joystick or automatically if using an electronic manipulator stage. The mechanical processes due to laser-induced lysis can create an emulsion at the droplet edge that higher-pulse energies can completely disrupt the droplets.
Ensure proper lysis pulse energies are kept to a minimum and cells are not deposited at the immediate edge of the droplets. Dispense the cells in the addressable droplet using the microinjector. Image the cells in droplets using bright-field microscopy, including any fluorescence imaging.
Also image all spots in the array using single-molecule TIRF microscopy. Focus on the cell in an addressable droplet. Achieve complete optical lysis by focusing a single 6-nanosecond laser pulse close to the location of the cell.
Note the time at which each cell is lysed. Acquire single molecule images using TIRF microscopy of all spots every ten minutes for the first 30 minutes, then every 20 minutes for a further 60 minutes. If only interested in the amount of protein bound at equilibrium, image all spots after incubating the chip for 90 minutes at room temperature.
To perform single-molcule counting for any non-congested antibody spot, load the background image acquired before lysis. In Fiji, select the background image. Under the image menu, select duplicate to duplicate the background.
Then select the duplicate image and under the process filters tab select Gaussian blur and specify a 50-pixel radius. Navigate to process and select image calculator. There specify the operation divide, the images required, then select the boxes create new window and 32-bit result.
Subtract each pixel in the image by one. Then select a 50-pixel-by-50-pixel area in any of the four corners in the background flat image. Measure the pixel intensity standard deviation by selecting analyze and measure.
Under image adjust, select threshold and set the lower threshold level to three standard deviations. In the segmented image SM mask set the pixel intensity values to zero for any objects that do not have a size of four to nine square pixels and a circularity of 0.5 to 1. To do so, select analyze followed by analyze particles and specify the size and circularity.
For the same antibody spots, load and repeat the data analysis steps for all image frames captured as a time-result series acquired post-lysis. In representative results of single-molecule images, signal in the background images shows the lower level of nonspecific binding of the fluorescently labeled detection antibody to the surface and antibody spot. The magnified inset image shows highlighted single molecules with red arrows.
Also shown is a typical image of a pulldown of p53 in BE human colon carcinoma cells. This sequence of images shows accumulation of p53 protein on the antibody spot. A calibration curve obtained using known concentrations of the combinant protein is used to convert single-molecule counts on each spot to the number of target proteins in the analysis volume and hints at single-cell copy number.
The horizontal red-dashed line indicates the level of nonspecific binding. The basal p53 protein expression in BE cancer cells shows a long-tailed gamma-like distribution where p53 protein expression in some cells is significantly higher than the modal value. Here, p53 protein expression is plotted as a function of cell volume as calculated from the measured cell diameter prior to lysis.
The results indicate that a minimum concentration of p53 protein is maintained in these cells. After watching this video, you should have a good understanding of how to measure the abundance of a target protein with single-cell resolution. While working with biological cells, it's important to remember that the proteins can degrade.
Once mastered, this technique can be completed within four hours if it is performed properly, and can be incorporated into protocols analyzing primary patient samples. Single-cell techniques can help provide insight into the cellular function and behavior as well as identify rare events that would otherwise be missed by both methods. Though this method can provide insight into the biology of p53 and breast cancer cell lines, it can also be applied to other studies of diseases without the need for labeling, which is beneficial in studying primary cells from patient samples.
Following this procedure, other methods like quantum dot PCR may be performed to answer additional questions such as determining whether target RNA and protein expression is correlated. Measuring the absolute copy number of a protein will be attractive to biomathematicians and computational scientists in supporting their biological models.
