単一細胞付着性細胞培養のタイムラプス蛍光顕微鏡による蛋白質分解率の定量化

The overall goal of this fluorescence imaging approach is to determine the half-life of a protein of interest in single living cells using a SNAP-tag. Cellular proteins are turned over extensively with synthesis and degradation rates being specific for each protein, and subject to physiological regulation. However, most commonly used methods to determine protein half-life are either limited to population averages from lice cells, or require the use of protein synthesis inhibitors.

In this video, we demonstrate how a SNAP-tag in combination with fluorescence time lapse microscopy can be used to measure protein half-lives in living cells. The advantage of this method is that it achieves single-cell resolution, and therefore provides an estimate for the heterogeneity of half-lives in a population of cultured cells. The experimental procedure presented here can be applied to any type of adherent cultured cells, expressing a protein of interest fused to a SNAP-tag.

This can either be achieved by endogenous protein tagging or by using over-expression of tagged proteins. The SNAP-tag is a mutant version of the DNA repair enzyme, O6-alkylguanine-DNA alkyltransferase that specifically reacts with benzylguanine, which can be coupled to molecular probes. Therefore, any SNAP-tag fusion protein can be irreversibly labeled with a florescence cell permeable dye.

Upon wash out of residual dye the labeled protein population will decay exponentially, and the half life of the corresponding protein can be inferred. In this video we use embryonic stem cells, which form tight colonies when grown classically on gelatin coated dishes. To allow them to grow as a monolayer of cells, and thereby achieve single-cell resolution for florescence imaging experiments, we cultured them on inc adhering coated plates, which reduces the cell to cell interactions mediated by cellular inc adhering expressed at the plasma membrane.

Dilute the recombinant inc adhering to five nanogram per microliter in PBS, and add 30 microliters of diluted inc adhering per valve in a 96 valve plate, suitable for imaging. Avoid extensive pipette-ing, as recombinant inc adhering is very fragile. Incubate at 37 degrees for one and one half hours.

Aspirate the inc adhering solution, wash once with 100 microliters of PBS, and add up to 200 microliters pre-wormed ESL culture medium. Use ESL lines expressing a protein of interest fused to the SNAP-tag. Wash the cells with five ml of PBS.

Then add two ml of Tripson, and incubate for four minutes at 37 degrees. Add four ml of ESL culture medium, re-suspend, and spin down at 1000 T for four minutes. Aspirate the supernatant, re-suspend in fresh medium, and count the cells.

Seat 30, 000 cells per square centimeter in the inc adhering coated imaging plate. Proceed with the SNAP dye staining 24 hours after cell seating. Dilute the cell dye in ELS culture medium to the optimal concentration previously determined.

Aspirate the medium using a pipette and carefully add 50 microliters of the diluted SNAP dye. Incubate for 30 minutes at 37 degrees. Aspirate the dye and wash three times with 200 microliters of pre-worm PBS.

Add 200 microliters of ESL culture medium, and incubate for 15 minutes at 37 degrees. Perform the washing steps by gentle pipette-ing, and do not use an automatic aspirator as cells tend to detach easily. Repeat the same washing steps twice more, then add 200 microliters of imaging medium.

Use phenol red free medium to reduce the florescence background. Use a microscope allowing controlled temperature and CO2. Set the temperature to 37 degrees, and the CO2 to five percent prior to use, and allow equilibriate for one to two hours.

Place the plate into the microscope, and select spot suitable for imaging. The bright field images show the ESL attached to inc adhering coated dish in a monolayer. Select the objective, a 20X objective is appropriate for this size of ESLs.

Select the illumination settings for the florescence channel to be recorded. Adjust the laser power and exposure to obtain a strong signal, since it will decrease over the course of the movie. Select the acquisition parameters for the time lapse experiment.

For proteins with expected half lives of two to twenty hours, choose acquisition times of 12 to 24 hours with time intervals of 15 minutes. Start the imaging experiment. Use the FIJI Software for processing and analysis of the movie.

Collect the acquired images as a stack and deform it. Use the subtract background FIJI function to remove background from all pictures in the stack. Select a cell of interest, draw a region of interest around it using the FIJI tool bar, and add the region of interest to the ROI Manager.

Proceed to the next frame and repeat the previous actions. Follow the cell of interest throughout the course of the movie and save the ROI set for each tracked cell. Click measure to obtain values for the mean intensity and the area of the ROIs.

To estimate local background add a region of interest close to the cell of interest for each time frame. In the end, click measure to obtain values for the mean background intensity. First, calculate the mean intensity of the cell by multiplying the obtained value for the mean intensity of the cellular ROI with its area.

Then, proceed to calculating the integrated background value of the cell, by multiplying the mean intensity of the background ROI with the area of the cellular ROI. Finally, to obtain a background corrected value for the integrated intensity of the cell, subtract the integrated background intensity value from the integrated cellular intensity value. To obtain an average half life of the cell population normalize the obtained values to the intensity value of the first frame for each cell.

This ensures that each cell contributes with the same weight to the average half life value independently of its absolute florescence intensity. After cell divisions track both daughter cells separately and sum their integrated intensities after background subtraction to account for protein dilution during cell division. Save the ROI sets for both daughter cells.

For the estimation of the protein half life, use the curve fitting tool in MATLAB. The protein decay follows an exponential where F(x)is the florescence intensity at the given time point, a"the initial intensity, and b"the decay rate. After obtaining the values for the decay rate b"from the fit, the half life can be calculated by dividing the logarithm of two by b"This movie shows the decay of the SNAP dye florescence in a doxycycline unusable opt four SNAP-tag fusion cell line.

The observed florescence signal will depend on the cell line used, the expression level of the corresponding protein of interest, as well as the properties of the dye being used. For protein decay experiments it is important to use an adequate SNAP dye concentration. The concentration should be high enough to yield a bright florescence signal in the beginning of the time lapse.

However, using too high dye concentrations might cause residual dye being left in the medium, even after washing. The free dye might subsequently bind to newly produced SNAP-tag molecules over the course of the movie, which would distort the decay curve. It is therefore crucial to optimize the SNAP dye concentration by testing different dilutions.

This figure shows the decay curves for eight different concentrations of a florescence SNAP dye tested on a doxycycline unusable cell line. An initial dye concentration of three micromolar was chosen according to the manufactures instructions for live cell imaging, and a serial dilution of one to three was performed until reaching a concentration of 1.4 nanomolar. For the two highest concentrations the signal does not decrease, whereas the observed decay follows an exponential curve for lower concentrations.

In this example an optimal concentration of 12 nanomolar was selected, since this concentration ensures minimal amounts of residual dye left in the medium after washing, but the florescence signal is still bright enough to observe a clear decay curve. The described protocol provides an estimate of the cell-to-cell variability and half life for any given protein fused to SNAP-tag. This figure shows the single cell decays, and the population average for three different doxycycline unusable SNAP-tag fusion cell lines.

Nanog"and Oct4"are rather short lived with average half lives of 2.9 and 4.8 hours. Whereas Srsf11"is longer lived having an average half life of 25.3 hours. The box plots show the distribution of half lives for the three proteins.

We have shown that the use of a SNAP-tag, in combination with florescence time lapse microscopy, allows for determining the half life of any protein of interest. The single cell decay measurements provide an estimate for the heterogeneity of half lives in a population of cells, which is omitted when using other methods to determine protein half lives. One of the most crucial steps when using a SNAP-tag to monitor protein decay, is to ensure that no residual unbound dye is left in medium or in the cells after washing.

As otherwise it might bind to newly produced SNAP-tag molecules later in the course of the experiment and thereby compromise the decay curve. This is on one hand achieved by carefully performing all the described washing steps. On the other hand the SNAP dye concentration should be kept as low as possible while still being in a range that allows to obtain a good signal to noise ratio.

The SNAP dye concentration should test be optimized for each experiment according to the cell line and the type of dye being used. While it is possible that the SNAP-tag might alter the half life of candidate proteins in some cases, we see no evidence for such effects, as the determined half lives of the candidate proteins test adhere closely matched published values. To further rule out an effect of the SNAP-tag on protein degradation, one option would be, to block protein synthesis with cycloheximide, and perform restroom blots at different time points, using an antibody that recognizes the indigenous protein.

And to directly compare the decay of the SNAP-tag candidate protein with the untagged indigenous version.