Quantitative Immunoblotting of Cell Lines as a Standard to Validate Immunofluorescence for Quantifying Biomarker Proteins in Routine Tissue Samples

This method can help answer key questions in the field of cancer pathology, by quantifying biomarker proteins from routinely processed biopsy material. The main advantages of this technique in relation to conventional immunohistochemistry, are that it is objective, has a wide dynamic range, and permits for the quantification of proteins in specific cell types. The presence or absence of certain so-called biomarker proteins, or more particularly, the relative abundance of those proteins in cancer biopsy specimens can be very useful in making a specific pathological diagnosis of specific kinds of cancers.

But it can also be useful in predicting the response to cancer treatments, so that's why this technique is relevant both to the diagnosis and the treatment of patients with cancer. This protocol is mainly about how to use protein quantification in immortalized cell lines to validate the quantitative nature of immunofluorescence based assay. But it's important to note that immunofluorescence can easily be incorporated into a multiplexed approach, and applied to primary biopsy samples.

Assisting and demonstrating this procedure will be Lee Boudreau, a technician, and Shakeel Virk, the director of operations. Both from the Queen's Laboratory for Molecular Pathology. To begin, centrifuge the previously harvested cells in a 50 milliliter conical tube at 225 Gs for five minutes.

Decant the supernatant, and resuspend the cell pellet in 10 milliliters of PBS. Then, centrifuge the resuspended cells at 225 Gs for five minutes. Suspend the cell pellet with 10 milliliters of 10%NBF.

Then incubate the cells at room temperature on a rocker at 24 RPMs overnight. After fixing the cells, centrifuge the cells at 225 Gs for five minutes. Then remove the supernatant and resuspend the cells in 500 microliters of PBS.

Transfer the cells to a 1.5 milliliter microcentrifuge tube, and pellet the cells via centrifugation. Then, aspirate the supernatant. Add approximately 500 microliters of 1%agarose solution to each tube containing cells.

Then, pipette the mixture up and down to mix. After the agarose cell solution hardens, add one milliliter of 10%NBF to the tubes. Later, remove the NBF from the samples.

Use a razor blade to remove the cell plug, and place the plug into a plastic tissue cassette. Process samples overnight with an automated tissue processor. Then, embed the samples in paraffin wax, using standard histology methods.

Using a tissue arrayer instrument, harvest duplicate 6 millimeter cores from the paraffin block, and insert them into an empty paraffin block. After this, use a microtome to prepare two histological sections of the newly created cell line, TMA. Then mount the sections on a histology slide, and deparaffinize them.

First, load the two slides of the cell line TMA into an automated staining system. Stain one side with the optimal primary antibody dilution, leaving the other as a negative control slide. To both slides, apply a nuclear counterstain, and secondary antibody.

After the staining process, add the tyramide signal amplification reagents. After this, scan the immunostained slides using the appropriate excitation and detection wavelengths for the fluorophores that were used. Use image analysis software to identify the cellular compartment of interest, whether nucleus or cytoplasm, and quantify the mean fluorescence intensity, or MFI, for each line.

To determine which cell line has the greatest abundance of the target protein, aliquot preciously prepared cell lysate into microcentrifuge tubes. Then add 2.5 microliters of 6x laemly buffer, and enough RIPA lysis buffer to bring the total volume to 15 microliters. Next, load an SDS page gel, with a protein ladder, and the previously prepared samples.

After this, load laemly buffer into the empty wells. Run the gel until the blue dye reaches the bottom of the plate. After performing a semi-dry protein transfer, use a razor blade to cut the membrane horizontally to separate the protein of interest from the control protein.

Perform blocking and antibody incubations according to the manufacturers instructions. Then place the membrane strips in a clear plastic bag. Use a P1000 pipette to cover the membranes with ECL mixture.

Then seal the bag, and incubate the membranes at room temperature in the dark for one to two minutes. Next, place the plastic bag containing the membranes in a digital imaging platform. Use chemiluminescence and colorimetric marker detection to capture various exposures of the membrane.

Using image analysis software, or visual observation, determine which cell line expresses the most target protein. After immunoblotting a serial dilution of this cell line, open the exposure images using image analysis software. Use the rectangular selections tool to select the first lane of the gel.

Then go to analyze, gels, and select first lane. Repeat this process by moving the rectangular selections tool over to the next lane, and go to analyze, gels, and select next lane. Next, go to analyze, gels, and plot lanes.

Use the straight line tool to draw lines across the bases of each peak, to remove the background noise. Then use the wand tool to select each peak, and obtain the band intensity of each peak from the results window. Create a scatterplot of band intensity versus the amount of total protein loaded for each primary antibody using the densitometry output.

Then, use a line of best fit and visual observation to determine the location of the linear dynamic range of each antibody. Select a protein concentration that yields a band intensity within the linear range, and perform an immunoblot of all cell line lysates with that protein concentration, as demonstrated previously. Next, perform densitometry on the digital scans, selecting exposures that yield signals within the previously identified linear ranges for each antibody.

After this, calculate the ratio of target protein band intensity to loading control band intensity for each cell line. Finally, use statistical software to perform a Pearson correlation test between the values obtained from image analysis of the IF staining, and those obtained from immunoblotting. This protocol was used to confirm the ability of IF to determine the relative quantity of the anti-apoptotic protein Bcl-2.

Varying dilutions of the primary antibody were tested on human tonsil tissue with an immunohistology stainer. For this experiment, a one to 50 dilution was determined to be the optimal dilution that yielded a strong signal with little background noise. This dilution was used to stain the cell line TMA by immunofluorescence, and image analysis was used to quantify the cytoplasmic Cy5 signal attributed to Bcl-2.

Based upon the immunoblot for all the tested cell lines, Granta-519 had the greatest abundance of Bcl-2. An immunoblot of the serial dilution of Granta-519 lysate was then used to find the linear dynamic range of Bcl-2, and control protein GAPDH. The dynamic range for Bcl-2 in this assay ranged from a band intensity of nearly 0, to 7500 units.

The dynamic range for GAPDH ranged from 3000 to 6500 units. The immunoblot was repeated with exposures within that linear range. And the ratio of Bcl-2 to GAPDH was calculated for each cell line.

A Pearson correlation test demonstrated that the intensity ratios from immunoblotting were strongly and positively correlated with the intensity readings from quantitative IF.While attempting this procedure, it's important to remember to expose the final membrane for multiple time points, In order to find the optimal exposure where all of the bands are within their respective linear ranges. After validating the quantitative nature of the IF protocol, it can be modified for multiplex IF, in routinely processed tissue samples in order to answer clinical questions such as where a target protein is expressed.