Flow Cytometry and Confocal Imaging Analysis of Low Wnt Expression in Axin2-mTurquoise2 Reporter Thymocytes

NOTE: All mouse procedures were performed with the approval of the Leiden University Medical Centre (LUMC) Ethical Committee on Animal Experiments. Male and female, 6-12-week-old, wild-type (wt), which have no insertion of the Axin2-mTurquoise2 reporter construct, heterozygous (Tg/0) with one insertion of the Axin2-murquoise2 reporter construct and thus, one disrupted Axin2 gene, and homozygous (Tg/Tg) with the insertion of the Axin2-mTurquoise2 reporter construct in both alleles and thus, two disrupted Axin2 genes; Axin2-mTurquoise2 mice (B6;CBA-Axin2^em1Fstl/J mice) were used in the experiments. The animals were sacrificed by CO₂ euthanasia prior to organ isolation. Throughout the procedure, minimize exposure of the samples to light, and keep on ice or 4 °C at all times, unless indicated differently. Cover the samples with aluminum foil. All the steps should be performed in a standard laboratory with a biosafety cabinet.

1. Preparation of thymocyte cell suspension

1. Harvest the thymus from mice carefully without blood contamination by cutting open the abdomen of the mice and extracting the thymus with forceps. Store/transport temporarily in ice-cold Iscove's Modified Dulbecco's Medium (IMDM) containing 2.5% fetal calf serum (FCS).
   NOTE: To avoid blood spillage and possible thymus damage, do not sacrifice the mice by cervical dislocation.
2. Prepare a 50 mL tube with a 70 μm cell strainer, and wet the filter with 1 mL of cold IMDM/2.5% FCS medium.
3. Mash the organ with the back tip of a 1 mL syringe plunger while washing twice with cold IMDM/2.5% FCS medium (Figure 1A). If desired, add a final concentration of 50 U/mL of DNAse I to the IMDM/2.5% FCS medium to prevent dead cell clumping. Rinse the filter 2x with cold IMDM/2.5% FCS medium, and resuspend gently in the 50 mL tube.
   NOTE: Do not exceed a total end volume of 10 mL. Keep cells on ice and in the dark for subsequent steps and when not handling.
4. Centrifuge at 330 × g for 5 min at 4 °C, and gently aspirate the supernatant from the cell pellet. Resuspend the cell pellet gently in cold incomplete IMDM/2.5% FCS medium, and prepare for cell counting.
   ​NOTE: If required, freeze and store the thymocytes in liquid nitrogen in FCS-10% dimethylsulfoxide for later experimentation. Proper cell freezing and thawing will reduce excessive cell death. On average, half of the thymocytes can be apoptotic after thawing due to the natural T cell selection in the thymus, which should be considered when deciding how many thymocytes to freeze down per cryo-vial. No less than 2.5 × 10⁶ thymocytes should be frozen per vial.

2. Thymocyte flow cytometry preparation

1. Prepare thymocytes for cell surface staining procedure by preparing 2.5 × 10⁶ thymocytes per staining sample in ice-cold phosphate-buffered saline (PBS, pH 7.4) (Figure 1B). Recount the number of live cells after thawing, if thymocytes have been previously frozen. If required, add a final concentration of 50 U/mL of DNAse I to prevent dead cell clumping.
2. Use the antibody staining panels for cell surface characterization of a complete thymocyte subset.
   NOTE: Other combinations of fluorochromes can be selected. Select rare population markers for bright fluorescent fluorochromes and if possible, add a live-dead marker. Neither V450 nor V500 fluorochromes should be used in combination with the mTurquoise2 fluorescent reporter due to spectral overlap. Always check the fluorescence spectra of mTurquoise2 in combination with blue and green fluorochromes (Supplemental Figure 1A).
   1. Mix the antibodies in previously defined ratios of the lineage-negative (Lin-) panels separately in PBS/0.2% bovine serum albumin (BSA)/0.1% NaN₃ (sodium azide) buffer.
      NOTE: All unwanted cells (non-thymocytes present in the thymus) are stained in a 2-step process with a streptavidin secondary antibody (in this example, phycoerythrin (Pe)-Cy7 and allophycocyanin (APC)-Cy7) and can be excluded by using a "dump gate" in the flow cytometry analysis.
   2. Mix the antibodies in previously defined ratios of the Double Negative (DN) staining panel with the streptavidin secondary antibody Pe-Cy7 in PBS/0.2% BSA/0.1% NaN₃ buffer. Exclude the Lin- panel from this mix (Table 1).
   3. Mix the antibodies in previously defined ratios of the Immature Single Positive (ISP), Double Positive (DP), and Single Positive (SP) staining panel with the streptavidin secondary antibody APC-Cy7 in PBS/0.2% BSA/0.1% NaN₃ buffer. Exclude the Lin- panel from this mix (Table 1).
   4. First, stain the thymocytes with the unwanted non-T cell populations by using the biotin primary antibody mixes of the Lin- panels for 30 min on ice in the dark.
      NOTE: Each Lin- panel is a different set of cells and should therefore not be stained together in one sample.
   5. Spin down at 300 × g, 4 °C for 5 min and remove the supernatant. Wash the thymocytes with 150 μL of ice-cold PBS/0.2% BSA/ 0.1% NaN₃ buffer, and spin down at 300 × g, 4 °C for 5 min.
   6. Stain with the DN panel and the ISP/DP/SP panel of the corresponding Lin- staining for 30 min on ice in the dark. Spin down at 300 × g, 4 °C for 5 min, and remove the supernatant. Wash the thymocytes with 150 μL of ice-cold PBS/0.2% BSA/0.1% NaN₃ buffer, and spin down at 300 × g, 4 °C for 5 min.
3. Prepare the cells for flow cytometry measurement by homogenizing with a 35 μm cell strainer tube and taking the cells up in PBS/0.2% BSA/0.1% NaN₃ buffer. Protect cells from light, and keep on ice until and during flow cytometer measurement.
   NOTE: NaN₃ (sodium azide) is highly toxic and fatal. Special care should be taken when working with this substance. Wash hands thoroughly after handling, and immediately call a poison control center or doctor/physician if NaN₃ is swallowed.

3. Flow cytometer measurement

NOTE: Inexperienced users should first take flow cytometer training as the measurement of the mTurquoise2 signal in combination with several other fluorochromes requires experience and knowledgeable planification of the experiment. See the Table of Materials for information about the flow cytometer.

1. Start the flow cytometer according to the user manual or other established protocol. Check, and if required adjust, the bandpass filter sets in the flow cytometer for an optimal fluorescence detection strategy.
   NOTE: A recommended filter for mTurquoise2 is 470/20 nm on the ultra-violet 405 nm, 407 nm, or the less common 440 nm laser line.
2. Calibrate the flow cytometer by establishing compensation settings with commercially available compensation beads and stably transfected, mTurquoise2-expressing 293T cells.
   NOTE: Single-stained wt thymocytes can be used instead of compensation beads. In this case, compensation with beads was equally efficient and more convenient than using cells.
   1. Label the beads with each individual fluorochrome used in the experiment, and include unstained beads. Measure the beads to set up a compensation setting panel, and measure mTurquoise2-expressing 293T cells as there is no matching fluorochrome for mTurquoise2, which can be used on the beads. Save the compensation settings for use in the actual experiment.
      NOTE: The mTurquoise2-expressing cells used for the compensation setting must be as bright or brighter than the mTurquoise2-expressing thymocytes of the actual experiment. Make sure there are also mTurquoise2-negative 293T cells present.
   2. For the experiment, include fluorescence minus one (FMO) Tg/Tg-stained thymocytes, including a wt sample for the mTurquoise2 FMO and an unstained sample of thymocytes of each mouse genotype as positive controls for the analysis part.
      NOTE: These controls are important to properly set the gates for positive cells. The rest of the experimental samples are stained with the complete staining panel (Table 1).
      1. Create an experiment, add and name the number of tubes in the flow cytometer software, and create scatter plots to visualize the stained cells for the complete set of fluorochromes.
      2. Apply the previously established compensation settings to the experiment. Adjust the forward scatter (FSC) and side scatter (SSC) with unstained wt thymocytes until the complete population of cells is visible in the scatterplot. Measure the unstained Tg/Tg mTurquoise2-expressing thymocytes first to make sure the positive population is visible.
      3. Measure a Tg/Tg fully stained thymocyte sample, and check for all fluorochrome combinations. If necessary, adjust the previously established compensation values for the fluorochromes that show incorrect compensation. Measure the rest of the experimental samples, and do not adjust any settings during the measurement of the samples.
         ​NOTE: Subsequent compensation adjustment can also be performed in the analysis software package. Although the number of available cells could be a limiting factor, it is advisable to perform this step during measurement. Keep the settings equal between all samples for comparison of the mTurquoise2 intensity values.

4. Flow cytometric analysis

NOTE: Flow cytometric analysis was performed using specific software mentioned in the Table of Materials; however, other flow cytometric analysis programs are also available.

1. Gate live cells and thymocyte subsets according to FSC and SSC values.
2. Check the compensation settings in the compensation matrix dialog box found to the left of the sample to ensure no fluorochrome interference and that proper compensation has taken place to avoid spill-over or bleed-through.
   1. If the acquisition-defined matrix must be adjusted, change the values in the compensation matrix by simply increasing or decreasing the compensation value of every fluorochrome combination so that the population does not show a nearly perfect horizontal or vertical line (Supplemental Figure 1B).
3. To visually spread the mTurquoise2 intensity for proper positive gating, change the mTurquoise2 X-axis display to linear (Supplemental Figure 2).
   1. Use the mTurquoise2 FMO control as negative control for mTurquoise2 positive signal gating. Revise the correct threshold gating per cell population (Figure 2).
      NOTE: The (wt) mTurquoise2 FMO control cells do not have the mTurquoise2 marker and hence, can be used as a background threshold for Axin2 reporter activity.
4. Gate mTurquoise2-positive cells with the appropriate detection channel to define how many cells are Wnt-positive.
5. Calculate the geometric mean and median to define the amount of fluorescent intensity in the cells of interest.
   1. Click on Statistics | Add statistic within the panel showing mTurquoise2-positive cells.
   2. Define the statistic method, the population of interest, and the detection channel of mTurquoise2, and click on Add. Represent the geometric mean and the median fluorescent intensity in Arbitrary Units (AU) to plot graphs.
      NOTE: The median represents the middle value of the fluorescent intensity and hence, provides information about fluorescent intensity population shift. If required, a background correction can help to obtain clearer visualization of the dynamic range of the mTurquoise2 reporter activity. This can be done by subtracting the wt background staining frequencies from the total frequency of mTurquoise2-positive cells of the specific cell type that is gated.

5. Preparation of thymocyte cytospins for confocal imaging

NOTE: Thymocyte cytospins are recommended when working with cell suspensions of non-adhering cells. As the expression of Axin2-mTurquoise2 in thymocytes is lower than in the thymus epithelial cells, filtered thymocyte cell suspensions were used for imaging.

1. Start with a cell suspension of freshly harvested or frozen thymocytes. Suspend ~20,000 thymocytes in 100 μL of cold PBS/0.5% BSA/10% FCS per thymocyte genotype.
   NOTE: If required, thaw the cells gently to preserve maximum cell viability when working with previously frozen thymocytes. This will aid in less autofluorescence when imaging the cells. Check for cell viability to ensure the quality of the sample. The cytospin procedure employs a mechanical force that has been adapted to the fragile thymocyte; nonetheless, it requires a highly viable starting population. To ensure higher viability, it is advisable to start with freshly harvested instead of frozen thymocytes.
2. Prewet the area around the opening of the filter cards with PBS. Assemble the cytospin sample chamber holder according to the manual (Figure 1C).
   1. Place the filter card on the frost slide (smooth side against the glass slide). Place both items on the sample chamber holder. Take care in placing the filter card exactly on the sample chamber holder hole, and place the complete sample chamber holder in the rotor.
3. Carefully resuspend the thymocytes, and add 100 μL of the cell suspension in the sample chambers. Spin the thymocyte suspension for 4 min at ~350 × g onto the frost slides. Remove the filter card carefully from the frost slide without touching the cells. Air-dry the cytospins for a period ranging from 1 h up to overnight at room temperature.
   ​NOTE: When working with other cell types, test different cell densities for optimal results. Cytospins can be frozen at -20 °C in a sealed box for later experimentation. Thaw the cytospins for further handling for 1 h at room temperature.

6. Cytospin immunostaining with total β-catenin

1. Fix the cytospins for 15 min at room temperature in 100% methanol. Air-dry the slides for 10 min at room temperature. Draw a circle around the thymocyte population on the glass slide with a hydrophobic pen.
   NOTE: This fixation step is optimized specifically for β-catenin staining.
2. Place the slides in PBS/0.05% Tween-20 for 10 min at room temperature, and then, transfer them to a dark humid box during the blocking and incubation steps. Add 100 μL of PBS/10% normal mouse serum (NMS) per slide, and leave in the humid box for 10 min at room temperature. Tap the slide to remove the 10% NMS, add 100 μL of PBS/10% normal goat serum (NGS) per slide, and leave in the humid box for 30 min at room temperature.
   NOTE: Incubation with PBS/10% NMS blocks non-specific primary antibody binding (Figure 1D), while PBS/10% NGS blocks non-specific secondary antibody binding.
3. Prepare additional antibodies for cellular staining. Mix 0.5 μg of the total β-catenin antibody with the AF568-labeled fragments.
   NOTE: In this setting, a commercially available labeling kit was used for pre-labeling of the primary anti-mouse total β-catenin with secondary goat-anti-mouse IgG1 Fab fragments with Alexa Fluor 568 (AF568) fluorochrome label before adding it to the thymocytes. Perform the labeling according to the manufacturer's protocol as several concentrations might need to be tested. Use the total β-catenin-AF568-labeled antibody within 30 min.
4. Add 50 μL (0.5 μg) of the total β-catenin-AF568-labeled antibody per cytospin slide overnight at 4 °C in a humid box. Include a negative staining control according to the manufacturer's protocol or an isotype control in case of a direct labeling protocol.
5. Wash for 20 min with PBS/0.05% Tween-20 at room temperature. Then, wash for 20 min with PBS at room temperature in a jar with stirring. Perform a second fixation step to ensure the binding of the antibody to the antigen: 10 min at room temperature with 100 μL of 4% paraformaldehyde (PFA) in PBS in a humid box.
   NOTE: Keep the slides in the dark. Neither methanol nor PFA fixation will significantly affect the mTurquoise2 expression²⁷.
6. Dip the slides in PBS. Perform nuclear staining with 50 μL of TO-PRO-3 (1:1500) for 10 min at room temperature in the humid box. Wash the slides for 20 min with PBS at room temperature in a jar with stirring.
   NOTE: The TO-PRO3 concentration can be titrated depending on the use of other fluorochromes with nearby fluorescent spectra.
7. Embed the specimens with an antifade reagent according to the manufacturer's protocol, and cover with a coverslip. Air-dry for 24 h at room temperature. View the slides directly under a fluorescent or confocal microscope, or store at -20 °C for later imaging.

7. Confocal microscopic measurement

NOTE: See the Table of Materials for information about the confocal microscope.

1. Switch on the confocal microscope according to the manual or established protocol. Use negative and positive stably transfected mTurquoise2 293T cell line controls for primary adjustment of the confocal settings. Subsequently, use wt Axin2-mTurquoise2 and Tg/Tg (knock-out) Axin2-mTurquoise2 thymocytes as negative and positive controls, respectively, to ensure no underexposure of the mTurquoise2 signal.
2. Prepare the software for sequential scanning by programming the lasers and filter widths. Start with the highest wavelength laser line first, and work toward the lowest wavelength. When all sequential scanning steps are installed, load the sample on the microscope stage, focus the sample, and press Live to optimize the laser power and Smart Gain using the respective buttons on the confocal software or optional manual panel.
   NOTE: The specimen section of the slide should not be imaged for signal quantification as potential photobleaching could potentially occur. However, mTurquoise2 has high photostability²⁵.
3. In case of very low mTurquoise2 expression, increase the laser power and Smart Gain until a fluorescent signal is observed, and check with the negative control sample to ensure a true positive signal. Visualize the thymocytes with a 40x 1.4 oil lens, 63x 1.4 oil lens, or 100x 1.4 oil lens.
   NOTE: A Leica SP5 microscope was used for this study.
4. Use these confocal imaging settings on the microscope before measuring the sample.
   1. Adjust the intensity value range to a 12-bit image by clicking on Configuration | Settings | change to 12-bit in the Bit depth option to create a broader scale of intensity and thus, more distinction between low and high fluorescent signals.
   2. Adjust the imaging resolution by clicking on XY, and increase the Format to 1024 x 1024, which will also double the scanning time. Adjust the scan speed to 400-600 Hz by clicking on Speed | More to manually change the settings. Additionally, activate the Bidirectional scanning option.
   3. Adjust the sensitivity slider to reduce the background signal. Optimize the correct laser power and Smart Gain with the Quick LUT (Look-Up Table) option.
      NOTE: In the 12-bit image, the slider has a grey scale intensity value from 0 to 4095. This can also be done afterwards with the free offline Las X software. The green color will show the black background, and the blue color shows saturated pixels of the sample.
5. When all the imaging settings are optimized, measure the sample by clicking on Start, which will initiate the sequential imaging of all three channels.
   1. Measure the TO-PRO-3 nuclear fluorescent signal. Detect TO-PRO-3 with the 633 nm laser and HyD 640-750 nm.
      NOTE: In this setup, 6% laser power was used at 15% smart gain. This setting can change depending on the intensity of TO-PRO-3 staining. If very bright, it could influence lower-intensity fluorochromes when overexcited. In such a case, reduce the staining concentration.
   2. Measure the β-catenin nuclear and cytoplasmic fluorescent signals. Detect β-catenin with the 561 nm laser and HyD 580-605 nm. Accumulate up to 2 scans for one image by adjusting the setting in the XY box in the software with line average 2 in the case of a low AF568 signal.
      NOTE: In this setup, 85% laser power was used at 87% smart gain.
   3. Measure the mTurquoise2 cytoplasmic fluorescent signal. Detect mTurquoise2 with the 458 nm laser and HyD 490-600 nm. Accumulate up to 4 scans for one image by adjusting the setting in the XY box in the software with line average 4 in the case of a low mTurquoise2 signal.
      NOTE: Due to old lasers on the confocal microscope used for this protocol and the low mTurquoise2 signal, 405 nm was used at 90% laser power, along with the 458 nm and 476 nm lasers at 100% laser power with HyD 490-550 nm at 100% smart gain. A 440 nm laser is most optimal, albeit less commonly present on a confocal microscope. High laser power should be handled with care and only performed with sequential imaging. Make sure emergency settings are in place to avoid detector overexposure. The laser power on other confocal microscopes could be inferior to the ones proposed in this protocol due to more potent or newer lasers. A bleaching test could be performed to ensure no fluorescent signal is lost before imaging. In the proposed setup, the photobleaching of mTurquoise2 was acceptable.
   4. Perform brightfield imaging for total cell visualization. Detect thymocytes with the 488 nm laser and PMT Scan-DIC. Export the Lif files for image analysis.
      NOTE: In this setup, 59% laser power was used with a gain of 212 V and data offset of -4.3%. Lif files can be read in the offline LAS x software for image correction.

8. Confocal microscopy analysis

1. Analyze the images using an image processing software²⁸ (Supplemental Figure 3). Load the images in the software.
   NOTE: Multiple formats are accepted, but TIFF files with LUT or direct import of Lif files into the image processing software are recommended.
2. Measure the active β-catenin signal in the thymocyte nuclei.
   1. Select the nuclei of the TO-PRO-3-stained thymocytes in the red grey value image for the analysis of active β-catenin. Do this manually, or use automated cell selection in the software.
      NOTE: Automated cell selection might need image processing for proper thresholding and particle analysis. Manual cell selection can be laborious, but generally does not require any image processing and is recommended in this protocol.
   2. Activate manual selection with any of the selection tools in the work bar.
      1. Select the contour of the nucleus, and add it to the Region of Interest (ROI) manager. Activate the ROI manager by clicking on Analyze | Tools | ROI Manager, and when a new ROI manager window opens, click on the first option Add (t), or use the keyboard shortcut t. Repeat the previous step until all the nuclei are defined and added to the ROI manager. Use the Show all option in the ROI manager to visualize the selected cells.
   3. Select >3 background areas where no cells are present, and add these to the ROI manager.
      NOTE: Size and shape are unimportant in this case. These regions will serve as background noise measurements for the final calculation.
      1. Define the measurement on the image by clicking on Analyze | Set Measurements. In the new window that opens with different measurement options, activate Area, Integrated Density, and Mean grey value | click on OK.
      2. Activate the β-catenin grey value image by clicking on it, and visualize the selected nuclei and background areas by clicking on Show all in the ROI manager. Observe all selected areas that are now visible in the β-catenin image.
      3. Click on Measure in the ROI Manager or click on Analyze | Measure. Observe the new Results window that opens, showing the results of the β-catenin signal within the ROIs.
      4. Transfer the results to a spreadsheet calculation program by clicking on Edit | Select all; and copy/paste the list into the spreadsheet for further calculation. Save the ROI manager for future reference without having to repeat the nuclei selection by clicking on More | Save….
   4. For automated cell selection, make Duplicates (Keyboard Ctrl D) of the image to be processed as many image processing settings cannot be undone.
      NOTE: Automated nuclear labelling requires image processing, in most cases, to define the nuclei automatically and accurately. Image processing should only be done for area selection purposes. Processed images are not useful for fluorescent intensity measurement because the pixel values are altered.
      1. Perform a Gaussian filter by clicking on Process | Filters | Gaussian Blur to smoothen the image. Test multiple Sigma (Radius) values, and activate the Preview option to visualize the effect before clicking on OK.
      2. Invert the image by clicking on Edit | Invert. Check the Brightness and Contrast by clicking on Image | Adjust | Brightness/Contrast. Use the Auto option or preferably, manually change the values.
         NOTE: Do not Apply the changes as this would alter the image properties. Simply close the B&C window when the desired image has been obtained.
      3. Create a Threshold by clicking on Image | Adjust | Threshold, and define the best Threshold settings where all cells are mostly visible. Click on Apply to apply the Threshold settings.
         NOTE: If a threshold has not been applied to the cells, which show holes, click on Process | Binary | Fill Holes to fill in the gaps within the cells. If cells are fused together with the threshold settings, click on Process | Binary | Watershed to separate these cells. A fine 1-pixel line will separate any cell the program interprets as fused.
      4. Define the smallest and largest nucleus in the image by manually selecting a nucleus of choice with the Freehand selection tool, add it to the ROI manager, and measure the Area.
      5. Analyze the particles (nuclei) by clicking on Analyze | Analyze Particles, and insert the smallest Area and the largest Area in the Size (^2) box with a hyphen (–) in between. Activate the boxes Display results, Add to manager, and Exclude on edges before clicking on OK. Continue with step 8.2.3 to complete the protocol.
3. Measure the cytoplasmic mTurquoise2 and β-catenin signals in the thymocytes.
   1. Select the contour of whole thymocytes in the brightfield image following the same steps as above.
   2. Select >3 background areas where no cells are present, and add these to the ROI manager.
      NOTE: Size and shape are unessential in this case. These regions will serve as background noise measurements for the final calculation.
      1. Activate the mTurquoise2 grey value image by clicking on it, and visualize the selected total cell ROIs and background areas by clicking on Show all in the ROI manager. Observe all the selected areas that are visible in the mTurquoise2 image.
      2. Click on Measure in the ROI Manager, or click on Analyze | Measure. Observe the new Results window that opens up with the measurement results of the mTurquoise2 signal.
      3. Transfer the results to a spreadsheet calculation program by clicking on Edit | Select all, and copy/paste the list into the spreadsheet for further calculation.
      4. Activate the β-catenin grey value image by clicking on it, and visualize the selected total cell ROIs and background areas by clicking on Show all in the ROI manager. Observe all the selected areas that are visible in the β-catenin image.
      5. Click on Measure in the ROI Manager, or click on Analyze | Measure. Note the new Results window that opens up with the measurement results of the total cellular β-catenin signal.
      6. Transfer the results to a spreadsheet calculation program by clicking on Edit | Select all, and copy/paste the list into the spreadsheet for further calculation. Save the ROI manager for future reference without having to repeat the cell selection by clicking on More | Save….
4. Calculate the Corrected Total Nuclear Fluorescence (CTNF) for active β-catenin using equation 1.
   CTNF = Integrated density – (Area × average of Mean background areas) (1)
5. Calculate the Corrected total Cell Fluorescence (CTCF) for mTurquoise2 using equation 2.
   CTCF = Integrated density – (Area × average of Mean background areas) (2)
6. Differentiate between active nuclear β-catenin and cytoplasmic β-catenin by subtracting the nuclear β-catenin values obtained in step 8.2.3.3. from the total cell β-catenin values obtained in step 8.3.2.5 to obtain the cytoplasmic inactive β-catenin.
   NOTE: Make sure the measurements are done within the same cell.
   1. Calculate the average of the Mean intensity of the background areas. Calculate CTNF and CTCF using equations 1 and 2. Consider IntDen (the sum of all the pixels within the selected area) as the Integrated density and not the RawIntDen.
   2. If needed, calculate the Standard deviation of the IntDen values for plotting graphs. Consider up to 200 separate cells for statistical analysis using the Mann-Whitney U-test.
7. Plot the results in an individual data point graph, and label the y-axis with the CTNF or CTCF values as Relative Fluorescent Units (RFU).