Proteomics Techniques Flashcards
What is LC-MS/MS, and how is it used in proteomics?
Liquid Chromatography-Tandem Mass Spectrometry (LC-MS/MS) is a high-resolution technique used to identify and quantify proteins and post-translational modifications (PTMs).
It combines liquid chromatography (LC) for peptide separation and mass spectrometry (MS/MS) for peptide fragmentation and analysis.
✔ Key Steps:
1️⃣ Protein digestion – Proteins are broken into peptides using trypsin.
2️⃣ Peptide separation – Peptides are separated using high-performance liquid chromatography (HPLC).
3️⃣ Mass spectrometry analysis – Peptides are ionized and detected based on mass-to-charge ratio (m/z).
4️⃣ Tandem MS (MS/MS) – Selected peptides are fragmented, generating spectra for sequence identification.
✔ Results Interpretation:
Peptide mass spectra → Identifies protein sequences.
Fragmentation patterns → Determines modifications like phosphorylation or acetylation.
Protein quantification → Measures protein abundance in different conditions.
✔ Best Used For:
✅ Global proteomics – Identifying thousands of proteins in a sample.
✅ Post-translational modification (PTM) analysis – Detecting phosphorylation, ubiquitination, glycosylation.
✅ Comparing protein expression across conditions (e.g., cancer vs. normal).
✔ Limitations:
❌ Requires highly purified protein samples.
❌ Data analysis is computationally intensive.
What are TMT and iTRAQ, and how do they work?
TMT (Tandem Mass Tags) and iTRAQ (Isobaric Tags for Relative and Absolute Quantitation) are chemical labeling techniques used to compare protein levels across multiple samples in a single mass spectrometry run.
Each sample is labeled with a different mass tag, but all fragments appear identical in MS1, with quantification occurring in MS2 fragmentation.
✔ Key Steps:
1️⃣ Protein digestion → Break proteins into peptides using trypsin.
2️⃣ Label peptides → React peptides with TMT/iTRAQ isobaric tags.
3️⃣ Pool samples together and run LC-MS/MS analysis.
4️⃣ Quantify peptides based on reporter ion intensities in MS2 spectra.
✔ Results Interpretation:
Higher TMT/iTRAQ reporter ion intensity = Higher protein abundance.
Compares protein levels across up to 10+ conditions in one experiment.
✔ Best Used For:
✅ Multiplexing (high-throughput) – Compare up to 16 samples simultaneously.
✅ Relative protein quantification – Measures changes in protein levels across conditions.
✔ Limitations:
❌ Expensive due to reagent costs.
❌ Ratio compression – Signal intensity can underestimate fold changes.
What is label-free quantification (LFQ), and how does it compare to TMT/iTRAQ?
LFQ is a mass spectrometry-based method that quantifies proteins without chemical labeling.
Instead of using isobaric tags, it compares peptide signal intensities directly between samples.
✔ Key Steps:
1️⃣ Proteins are digested into peptides using trypsin.
2️⃣ LC-MS/MS is performed on multiple samples separately and peptides are separated based on retention time.
3️⃣ Protein intensities are compared across runs using computational algorithms (MaxQuant).
✔ Results Interpretation:
Protein quantification is based on peptide peak intensities rather than label-based reporter ions.
✔ Best Used For:
✅ When multiplexing is not needed (fewer samples, lower cost).
✅ When avoiding TMT/iTRAQ ratio compression artifacts.
✔ Limitations:
❌ More variability across runs (relies on reproducibility of LC-MS/MS).
❌ Lower precision than TMT/iTRAQ.
How does 2D Gel Electrophoresis separate proteins?
2D Gel Electrophoresis separates proteins based on two independent properties:
1️⃣ Isoelectric focusing (IEF) → Separates proteins based on isoelectric point (pI) (first dimension).
2️⃣ SDS-PAGE → Further separates proteins by molecular weight (second dimension).
✔ Key Steps:
1️⃣ Extract proteins and apply to pH gradient gel for isoelectric focusing.
2️⃣ Transfer to SDS-PAGE gel for separation by molecular weight.
3️⃣ Stain gel with Coomassie Blue or Silver stain for visualization.
✔ Results Interpretation:
Each spot on the gel = a different protein.
Changes in spot intensity indicate differential protein expression.
✔ Best Used For:
✅ Protein separation & visualization (e.g., detecting isoforms).
✅ Comparing protein expression between conditions.
✔ Limitations:
❌ Low sensitivity for low-abundance proteins.
❌ Difficult to analyze complex protein mixtures.
What is Co-IP, and how is it used to study protein interactions?
Co-IP is used to isolate and study protein-protein interactions by using an antibody to pull down a target protein and its binding partners.
✔ Key Steps:
1️⃣ Incubate a cell lysate with an antibody against the protein of interest.
2️⃣ Capture immune complexes using Protein A/G beads.
3️⃣ Wash and elute proteins bound to the target protein.
4️⃣ Analyze using Western blot or LC-MS/MS.
✔ Results Interpretation:
If a co-precipitated protein appears in the Western blot or MS/MS analysis, it suggests a direct or indirect interaction.
✔ Best Used For:
✅ Studying protein-protein interactions in vivo.
✅ Identifying binding partners of a target protein.
✔ Limitations:
❌ Requires highly specific antibodies.
❌ Can capture indirect interactions (false positives).
What is the primary application of CD spectroscopy in protein analysis?
CD spectroscopy is used to analyze secondary structure of proteins, including:
✔ Alpha-helices
✔ Beta-sheets
✔ Random coil structures
What is Circular Dichroism (CD) Spectroscopy?
CD spectroscopy is an analytical technique used to measure the difference in absorbance of left- and right-handed circularly polarized light by chiral molecules, particularly proteins and nucleic acids.
