The overall goal of this experiment is to identify and accurately quantitate over 10, 000 proteins from cell or tissue lysate across different drug treatments, time courses, or biological conditions. This method can help answer key questions in the field of medicine or biology such as what proteins are changing in response to a drug or other biological stimulus. The main advantage of this technique is that it has the ability to accurately quantitate over 70%of the expressed proteome over 10 different experimental conditions.

The implication of this technology can be extended to the diagnosis of human diseases through potential biomarker discovery. Visual demonstration of this method is critical as optimal cell lysis and fluid estimation play a key role in downstream digestion, labeling, and separation procedures. To begin this protocol, obtain cultured cells of interest.

Wash the cells twice using 10 milliliters of PBS for each wash. Scrape and collect the washed cells in a 1.5 milliliter tube containing one milliliter of PBS. Centrifuge at 600 times g and four degrees Celsius for five minutes.

Remove the supernatant and store at minus 80 degrees Celsius until ready to lyse the cells. Collect and weigh the desired tissues quickly after dissection. After this, immediately store them in liquid nitrogen and store at minus 80 degrees Celsius.

Prepare the lysis buffer on the day of the experiment as outlined in the text protocol. Add lysis buffer to the frozen sample such that the buffer to sample ratio is 10 to one and the final protein concentration is between five and 10 milligrams per milliliter. Next, add glass beads and use a blender to lyse the samples at four degrees Celsius and speed eight by blending for 30 seconds and then resting for five seconds, repeating this five times or until the samples are homogenized.

Measure the protein concentration using a standard protein quantitation assay or a Coomassie-stained SDS polyacrylamide gel with BSA as a standard. After this, add 100%acetonitrile such that the final concentration is 10%and Lys-C protease is at an enzyme to substrate ratio of one to 100. Incubate at room temperature for two hours to allow protein digestion to occur.

Then add DTT such that the final concentration is one millimolar. Incubate at room temperature for one hour. Next, add 50 millimolar HEPES until the urea concentration in the samples is diluted to two molar.

Add trypsin to each sample at a trypsin to protein ratio of one to 50. Incubate at room temperature for at least three hours. Next, add DTT until its concentration is once again one millimolar and incubate at room temperature for two hours.

Add iodoacetamide such that its final concentration is 10 millimolar. Incubate in the dark for 30 minutes at room temperature. After this, quench any unreacted iodoacetamide by adding DTT such that its final concentration is 30 millimolar.

Incubate at room temperature for 30 minutes. Check the efficiency of the trypsin digestion as outlined in the text protocol. Next, add TFA to each sample such that the final concentration is 1%Using a pH strip, measure the pH of each sample to verify it is between two and three.

Add additional TFA drop wise if needed to reach an unacceptable pH. Centrifuge the samples at 20, 000 times g and room temperature for 10 minutes. Collect the supernatant and load the samples onto prepared spin columns.

Centrifuge at 100 times g for three minutes or until the sample has passed completely through the column to bind the samples. Next, add 0.5 milliliters of 0.1%TFA to each column. Centrifuge at 500 times g for 30 seconds.

After this, add 125 microliters of a solution containing 60%acetonitrile and 0.1 TFA to each column. Centrifuge at 100 times g for three minutes to elute the peptides off the column. Reconstitute each desalted peptide sample in 50 microliters of 50 millimolar HEPES.

Using a pH strip, verify that the pH of each sample is between seven and eight. Next, dissolve the TMT reagents in anhydrous acetonitrile, add them to the samples, and then incubate at room temperature for one hour. After this, use 10 microliter pipette tips embedded with chromatography media to desalt one microgram of each sample and analyze the labeling efficiency as outlined in the text protocol.

Examine six to 10 separate peptides to ensure that unlabeled peptides are not detected in the labeled samples. Then, mix approximately two microliters of each sample together. Use 10 microliter pipette tips embedded with chromatography media to desalt this mixture.

Analyze the samples by LC-MS/MS as outlined in the text protocol. To begin, solubilize the desalted pooled TMT-labeled peptide sample in 65 microliters of buffer A.Use a pH strip to verify that the pH is approximately 8.0. If the sample is still acidic, use ammonium hydroxide to adjust the pH as necessary.

Next, fractionate the sample as outlined in the text protocol. Set the fraction collector to collect fractions every two minutes including loading time. Set the flow rate to 0.4 milliliters per minute.

Collect a total of 80 fractions. Then, use a vacuum concentrator to completely dry every other sample. Using LC-MS/MS, analyze all 80 fractions to achieve ultra deep proteome coverage.

First, pack 1.9 micrometer C18 resin into 75 micrometer inner diameter empty columns reaching a bed volume of approximately 1.3 microliters. Coil the column two to three times inside a butterfly portfolio heater to ensure the entire length is heated. Tape the column to the inside of the heater making sure not to tape over the heater's temperature sensor.

Then, heat the column to 65 degrees Celsius. Run 100 nanograms of rat brain peptides through the LC-MS/MS system to assess the system's quality. Repeat this assessment one additional time.

After this, load approximately 0.2 micrograms of reconstituted peptides on the column while flowing 5%buffer A.Elute the peptides off the column and analyze with the mass spectrometer as outlined in the text protocol. In this study, a high throughput method for the quantitation of proteins with a 10-plex isobaric labeling strategy is described. TMT labeling efficiency is examined using LC-MS/MS to analyze TMT-labeled samples and corresponding unlabeled samples.

As can be seen, the unlabeled peptide peak is found only in the unlabeled sample while the TMT-labeled peptide peak is found only in the labeled sample. Next, the effect of the MS2 isolation window on interference is evaluated. All MS2 scans are designated either as a clean or a noisy scan.

While the clean scans exhibit y1 ions of only lysine, the noisy scans exhibit them in both lysine and arginine. Representative interference removal using E.coli peptides individually labeled with three different TMT reagents is shown here. The summed relative intensities reveal that y1 ion-based correction is more accurate for TMT-based quantitation.

Once mastered, this protocol can be completed in four to five weeks if done properly. While attempting this procedure, it is very important to make sure all the quality control steps are done completely to ensure the final dataset is as accurate as possible. Adapting this procedure to focus on post-translational modifications can answer other questions such as how ubiquitination, phosphorylation, or acetylation is changing in response to disease progression or drug treatment.

After its development, the technology paved the way for researchers in the product field to explore the protein changes in cells and tissues. After watching this video, you should have a very good understanding of how to identify and accurately quantitate over 10, 000 proteins from a mammalian cell or tissue. Don't forget that working with high-pressure UPLC and few silica columns can be extremely hazardous and precautions such as wearing safety goggles should always be taken while performing this procedure.

We first had the idea of this method when we began to address the known problem of ratio compression which occurs in isobaric labeling. We have realized extensive fractionation is a great way to not only alleviate this problem, but also increases the identification of proteins and peptides which can further increase and adapts of a proteome analysis.