Lecture 7: Proteomics Flashcards
What is proteomics?
Proteomics is the study of the proteins present in a tissue.
• It looks at things like PTMs, expression, interactions, localisation and function.
• We can look at what goes wrong in disease and try and find drug targets.
• We can try and catalogue all the proteins in a sample.
• We can compare proteins in two or more different samples.
How can we prepare samples for proteomics?
There are multiple steps in sample prep.
• Cell disruption.
• Protection from proteolysis with inhibitors like benzamide.
• Fractionation. Sometimes the protein will come from a specific part of the cell such as the mitochondria.
• Denaturation. Performed with SDS (used in SDS-PAGE) or urea (used in 2D-PAGE and MS).
• Reduction of disulphide bonds. Performed with DTT, β-mercaptoethanol or TCEP.
• After reducing disulphide bonds, we want to cap the free thiols through alkylation. This is done using iodoacetamide.
How can we use protein precipitation?
Protein precipitation is a method used to remove contaminating species from a sample.
• For example, we want to get rid of salts, detergents, nucleic acids and lipids.
• Also used to concentrate protein samples.
• Dissolving the pellet again can be difficult.
• Trichloroacetic acid (TCA) is a very effective precipitant.
• Acetone can be used with TCA.
• Chloroform and methanol are another alternative.
How can we assay proteins?
After performing purifications, we want to assay the sample and see if protein is present.
• A280 absorbance with a nanodrop. It’s very fast. Doesn’t require reagents or prep time. A standard curve is also not required. However, we can get interference from other components like DNA that absorb UV.
• Bradford assay turns from brown to blue (absorbance at 595 nm).
• Coomassie blue. 2D-PAGE sample compatible. It is quick, but it has a poor linear range.
• Bicinchoninic acid (BCA) turns from green to purple (abs 526 nm). 2D-PAGE incompatible. It is slow, but it has a good linear range.
What is SDS-PAGE?
SDS-PAGE is a simple but powerful method used to separate proteins by molecular weight.
• Protein is boiled with SDS and a buffer containing DTT.
• Gel is loaded and run.
• The protein is then stained.
• SDS-PAGE has lower resolution than 2D-PAGE.
• However, it is better for hydrophobic proteins, high MW basic proteins and proteins which extreme pIs.
• Differentially expressed proteins can be cut out with a scalpel, digested and identified with mass spec.
What is 2D-PAGE?
2D-PAGE uses two rounds of separation, one based on pI and one based on MW.
1st dimension
• For the 1st dimension we use urea for denaturing, DTT for reduction and CHAPS (a zwitterionic molecule) detergent for solubilisation.
• Bromophenol blue or Orange G are used as a tracking dye.
• IEF separates proteins according to their pI. Proteins are amphoteric, and the pI is when the net charge of the protein is zero.
• IEF is performed at very high voltages.
• More acidic proteins move to the anode. More basic proteins move to the cathode.
• We use immobilised pH gradient strips fixed onto a plastic backing.
• The strips are easy to handle, have no gradient drift, higher protein load, better for extreme pH and they are reproducible. We can add the ampholytes.
• Various strip lengths are available with different pH ranges.
2nd dimension
• After IEF, the IPG strips are equilibrated in a buffer coating.
• Urea and glycerol increase viscosity.
• DTT preserves the reduced state and SDS denatures protein.
• Bromophenol blue is used as a tracking dye.
• 2nd dimension gel is run in the same way as SDS PAGE.
• For this and SDS-PAGE we fix the proteins with EtOH/MeOH and acetic acid solution before staining.
• We then dye. Classical Coomassie is relatively cheap, low reproducibility, poor 100 ng detection limit and compatible with mass spec.
• Colloidal Coomassie has good reproducibility, better 10 ng detection limit and is compatible with mass spec.
• Silver has a good detection limit ( <1 ng) but it is incompatible with mass spec, expensive and has low reproducibility.
• Fluorescence has good reproducibility, has a good < 1ng detection limit and is compatible with mass spec. However it has an expensive scanner and is expensive itself.
Other
• We can use computers to aid with image analysis.
• We always run at least 4 2D gels in order to get good stats.
• Software can show statistically significant changes when we are comparing different samples (e.g. normal vs disease).
What is 2D-DIGE?
2D-DIGE is 2D difference gel electrophoresis. It labels up to 3 different proteins samples with fluorescent dyes prior to 2D-PAGE.
• One problem with 2D-PAGE is the gel-to-gel variability and the challenge of aligning features.
• DIGE allows an internal standard and up to 2 further samples to be run on 1 gel.
• The samples are pre-labelled with fluorescent dyes. The dyes are mass + charge matched and spectrally resolvable. They have good photostability.
• The samples are mixed and then run in the same way as classical 2D-PAGE.
• The internal standard reduces gel-to-gel variation.
• Fewer gels saves time and it increases throughput. However, the multiplex fluorescence scanner is expensive.
How can we study post-translational modifications?
We can selectively stain SDS-PAGE or 2D-PAGE gels in order to detect PTMs.
• Once stained, they can be viewed with a fluorescent total protein stain such as SYPRO ruby.
• Oxidation with periodic acid converts cis-glycols to dialdehydes, which can then react with the hydrazide-based reagents. Periodate-oxidised carbohydrate groups can be stained by stains such as Pro-Q Emerald. This creates a bright green-fluorescent signal. Detection can be as low as 1ng on gels and 2-18ng on blots. We visualise with UV. Glycans can be analysed by HPLC or mass spec.
• We can stain phosphoproteins with Pro-Q diamond at a range of 1-16 ng.
How can we validate proteins?
A common approach to validate proteins is to use Western blotting.
• Proteins are transferred onto a membrane with electroblotting.
• We block non-specific binding with casein or milk.
• We then use a primary antibody and wash.
• We add a horseradish peroxidase (HRP) tagged secondary antibody and wash again.
• We can detect the proteins with a chemiluminescence substrate.
What is mass spectrometry?
Mass spec is used to determine the mass of molecule in a sample.
• We can find elemental composition, MW, 3D structure, PTMs, sequence and quantify the amount of a protein in a sample. We can also use it for protein identification.
• The samples are ionised in order to generate charged molecules. Their mass-to-charge ratios are measured on the mass spectrometer and displayed on a spectrum.
What are the two methods of digestion?
- We first cut expressed bands from SDS or 2D gels.
- We can perform in-gel digestion. We wash the gel and reduce the sample with DTT before alkylating with iodoacetamide. We then use trypsin and extract the peptides for mass spec.
- We can also perform in solution digestion. We add trypsin, DTT and iodoacetamide. We clean up the peptide sample by using solid phase extraction (SPE).
- We can also use filter aided sample preparation (FASP). FASP uses a centrifugal filter. Proteins above 3 kDa are retained on the filter. Trypsin can then be added to digest the proteins into peptides, so they can pass through the filter. Flow through must be kept for mass spec. Lys-C is often used in order to increase sequence coverage.
- We can remove contaminants with HPLC or C18 tips.
How do we ionise the sample? What are the two methods?
- The sample is directly inserted into the ionisation source and ionised. The ions are extracted into the analyser and separated according to m/z. The separated ions are detected with information on relative abundance and m/z ratio.
- There are many different ionisation methods such as ESI and MALDI. Most methods have the possibility of making positively and negatively charged sample ions.
- In MALDI, the sample is mixed with a matrix. A laser is fired on spotted sample/matrix. The matrix transformed the laser energy into excitation energy. the splutter of peptide and matrix ions gives an ionised sample. The ions are single charged; it is soft ionisation.
- In ESI, the peptide sample is pumped through a narrow capillary in 0.1% formic acid. High voltage is applied to the capillary.
How do we analyse the ions?
There are many different methods of mass analyser available.
• Quadrupole.
• Time of flight analysers.
• Fourier transform. For example, an FT-ion cyclotron resonance determines m/z based on cyclotron frequency of ions in magnetic field).
• Ion trap (and orbitrap).
• Magnetic sectors use static magnetic sector.
We can use more than 2 analysers.
• A tandem mass spec uses more than 2 analysers in the same mass spec.
• Tis is used for sequencing and structural work.
• If the analyser is not the same, it is called the hybrid mass spec.
How does a quadrupole mass analyser work?
A quadrupole mass analyser uses electricity to block of facilitate the passage of ions.
• There are 4 rods with DC current running through them.
• 2 are positive and 2 are negative.
• AC also runs through them at radio frequency. The rods oscillate from negative to positive.
• We change the voltage.
• Only ions of a certain m/z can pass through to the detector.
• Like charges repel and opposite charges attract.
• Positive acts as a high mass filter, while negative acts as a low mass filter.
What is a time of flight mass analyser?
Ions are accelerated by an electric field of known strength.
• Ion velocity depends on m/z.
• The m/z is determined by time taken to reach the detector.
• Linear TOF is lower resolution.
• Orthogonal TOF has multiple turns and it gives a higher resolution.
