Accurate quantitative measurements of proteins is crucial to comprehend the dynamic and spatial cooperation among proteins to understand the disease back to biology. The overall objective of this experiment is to provide an integrated quantitative proteomics workflow for tissue samples, combining label-free and label-based protein profiling for biomarker discovery. In this study, we have used the instrument Orbitrap Fusion to quantify the proteins using label-based and label-free approaches.
Tissue lysis is one of the most crucial steps for the success of an LC-MS-MS experiment. Take 30 mg of tissue in a bead beating tube. After washing with the PBS, add 300 microliter of urea lysis buffer containing eight molar urea, Tris, NaCl and MgCl2.
Using a probe sonicator, lyse the contents of the tissue. Repeat this for another cycle. If you observe any intact tissue at the bottom of the tube, add zirconium beads to the tube and homogenize the tissue using a bead beater for 90 seconds, with five minutes incubation on ice.
Centrifuge the samples around 6000 G for 15 minutes at four degrees Celsius to separate the cell debris from the supernatant. Collect the supernatant in a fresh tube and mix the solution uniformly. Quantify the protein concentration in the tissue lysate using Bradford reagent.
For this, measure the OD of the sample at 595 nanometer and extrapolate it using a standard graph as shown on this screen. Following the protein quantification, run 10 microgram of tissue lysate on 12%SDS-PAGE gel to check the quality of lysates. The first step in in-solution digestion involves the reduction of disulfide bonds, using TCEP.
Following the addition of TCEP, incubate the contents of the tube for one hour at 37 degrees Celsius. The next step involves alkylation to prevent the reduced disulfide bonds from re-forming. Add an alkylating agent like iodoacetamide and incubate the tube in dark for 10 minutes.
After incubation, lower the final concentration of urea to less than one molar by diluting with four to eight volumes of dilution buffer. At this point, it is advisable to check the pH. The next step involves the addition of trypsin to the tube, and incubating the tube overnight for efficient digestion.
On the next day, dry the digested peptides in a vacuum concentrator, and proceed for desalting step. To perform the desalting of peptides, use C18 stage tips. Activate the C18 stage tip by adding 50 microliter of methanol, centrifuge the tip at 1000 G for two minutes.
Now, add 50 microliter of acetonitrile in 0.1%formic acid to wash the stage tip, and again, centrifuge the tip. Reconstitute the dried digested peptides in 50 microliter of 0.1%formic acid. Add the reconstituted peptides into the activated stage tip.
Repeat this step at least four times. To wash the sample, add 50 microliter of 0.1%formic acid, and repeat the centrifugation step. For the elution of peptides, add 50 microliter of 40%ACN in 0.1%formic acid, and pass it through the stage tip by centrifugation.
Repeat this step with 50%and 60%ACN in 0.1%formic acid and collect the filtrate in a fresh tube. Dry the desalted peptides using a vacuum concentrator. Prior to LC-MS-MS run, quantify the peptides using the Scopes method.
For this, load two microliters of reconstituted sample in the well of a microdrop plate, and measure the absorbance at 205 nanometer and 280 nanometer. Calculate the concentration from the formula described on this slide. Once the desalted peptides are dried, they are ready for label-based quantification.
For iTRAQ labeling, the dried peptides must be reconstituted in 20 microliter of dissolution buffer, provided in the iTRAQ labeling kit. Reconstitute the labels by adding ethanol from the vial provided in the kit, and mix thoroughly. It is advisable that all the steps be carried out as per the manufacturer's instructions.
Add the homogeneously mixed iTRAQ labels to their respective tubes and allow for the labeling reaction to take place. At the end of the reaction, quench any excess unbound label in the tube by adding MS-grade water, and incubate for half an hour to one hour. Now, transfer all the labeled contents into a single tube and dry the labeled peptides.
After reconstituting the samples in 0.1%formic acid, open the autosampler of NanoLC, and place the samples inside the autosampler. Once the samples have been placed in the autosampler, it is important to set the right parameters for liquid chromatography and mass spectrometry. Here, we describe the parameters for both label-free quantitation and iTRAQ-based quantification using LC-MS.
In case of label-free quantitation, a 120 minutes gradient has been used as a single unfractionated sample is being injected. This duration can be increased or decreased based on the sample complexity. In this experiment, a flow of 300 nanoliter per minute has been set with the following gradient for solution B, which is 80%acetonitrile in 1%formic acid, in this case.
In the same way, we can set a gradient for an iTRAQ experiment. Since the samples are unfractionated, a 90-minute gradient has been used. The MS parameters for both label-free quantification, as well as iTRAQ label-based quantification, can be seen here.
The first step involves setting the application mode by clicking on the global parameters. Select the peptide mode, and set the method duration as per the LC gradient being used. Define the scan parameters for the experiment.
On clicking the MS-OT option, the parameters become visible. The detector type in use is set to Orbitrap. The other option available on this instrument includes an ion trap.
The resolution has been set to 60, 000, and one may choose between different options available, depending on the need and type of the experiment. Mass range is normal, and the scan range is set from 375 to 1700. The other parameters are default for MS Application.
The intensity threshold has been set, and the charge state set in the range of two to six. Dynamic exclusion has been set to 40 seconds, and the mass tolerance has been kept to 10. The next set of parameters include the MS-MS parameters.
The isolation mode has been selected with quadrupole, isolation window set at two, isolation offset has been set to off, and the activation type is usually HCD for the proteomics experiment. Collision energy can either be fixed, or it can be assisted. Labelled samples require higher collision energies, hence, for iTRAQ experiments, it is advisable that the collision energy be set to 35, while for LFQ experiments, it can be set to 30.
The detector type used here is Orbitrap. Scan range mode will be auto. The other parameters are default for iTRAQ experiments and label-free quantitative experiment.
Clicking on the summary will provide an overview of all the LC and MS parameters used for the experiment. You can have a look on each and every parameter and do the proofreading. Once that is done, then click on the file and save it as a method file.
Once you save, then you can run the method, and you will get the raw files. And then those raw files can be analyzed using Proteome Discoverer software. This figure shows the sequence coverage of BSA in three technical replicates, which is 91%in the three replicates.
This indicates the reproducibility of the instrument. Now, this figure shows the uniformity in the number of peptide spectral matches, peptides, and proteins identified in three different biological samples, which, again, gives an idea about the reproducibility of the instrument. Here, the Venn diagram represents the common and exclusive proteins identified in three different samples in label-free experiment and label-based experiment.
These bar graphs shows the number of peptide spectral matches, peptide groups, total proteins, protein groups, and protein number after 1%FDR, identified in LFQ and iTRAQ experiment. Tissue proteomics of biological samples enables us to explore new potential biomarkers associated with different stages of disease progression. The described protocol for tissue quantitative proteomic analysis provides reproducible good coverage data.
Use of technical replicates ensures good reproducibility, even if the samples are run at different time points. Here, we have shown the analysis of tissue samples using two quantitation methods:label-free and label-based proteomics. Selection of quantitative proteomic techniques may depend upon the number of samples, availability of MS-platforms, and biological question to be addressed.
