P1: Basics of Proteomics and Mass Spectrometry

Why is proteomics so challenging?

1. Complexity: DNA and RNA are structurally uniform polymers of only 4 different monomers (bases), while proteins are structurally diverse polymers of 20 different monomers (amino acids). Thus, proteomics is challenging compared to transcriptomics/genomics due to the incredible diversity of proteins and their amino acid-sequences.

2. Signal strength: DNA and RNA signals can be amplified by PCR or other polymerase reactions, while proteins cannot be amplified. Additionally, they are present across a large dynamic range in biological samples (i.e. some are present in very large quantities while others are present in very small amounts).

Mass-spectrometry

Mass spectrometry (MS) is an analytical technique used to determine the mass-to-charge ratio (m/z) of molecules, primarily for identifying and quantifying compounds such as proteins, peptides, metabolites, and small molecules.

Mass spectrometry works in three main steps (and 3 corresponding components):

1. Ionization (ion source) – Converts molecules into charged ions.

2. Mass Analysis (mass analyser) – Separates ions based on their mass-to-charge ratio (m/z) by passing them through a magnetic field. Ions with the same m/z will undergo the same amount of deflection. Lighter ions are deflected to a greater degree than heavier ions.

3. Detection (detector) – Measures the abundance of ions and generates a spectrum.

Mass to charge ratio m/z

The mass-to-charge ratio (m/z) represents the mass of an ion divided by its charge. It is used to separate and identify ions in a mass spectrometer.

Bottom-up proteomics VS Top-down proteomics

Bottom-up proteomics: common method to identify proteins and characterize their amino acid sequences and post-translational modifications by proteolytic digestion of proteins prior to analysis by mass spectrometry.

Top-Down Proteomics: Analyses intact proteins without digestion, preserving information on modifications and isoforms.

Protein separation techniques

1. Isoelectric Focusing: separating proteins based on charge.

2. SDS-PAGE: separate proteins by size.

3. 2D-PAGE: separate on both charge and size.

4. Liquid chromatography:

Isoelectric focusing (IEF)

What?

Isoelectric focusing (IEF) is a technique used to separate proteins based on their isoelectric point (pI), the pH at which they carry no net electric charge.

How?

• A gel is filled with a pH gradient. This gradient is stable and ranges from a low pH (acidic) to a high pH (basic).

• The sample containing proteins is applied to the gel.

• When an electric field is applied, proteins migrate based on their charge. Each protein will stop at the point where its net charge is zero (i.e. at the pH corresponding to their pI).

• This results in focused bands corresponding to the pI of each protein.

This is often only a first basis for separation as some proteins may have the same pI and will therefore not be separated by this technique.

SDS-PAGE

What?

SDS-PAGE (Sodium Dodecyl Sulphate - Polyacrylamide Gel Electrophoresis) is a technique used for separating proteins based on their molecular weight.

How?

1. Protein denaturation with SDS: the proteins are coated with SDS, a detergent that:

   • coats the protein with negative charges. 1 SDS anion binds for every 2 amino acids, providing a negative charge proportional to the length of the protein. This allows for the separation to only be done on the basis of molecular weight rather than both weight and charge.

   • denatures the proteins, converting them into linear polypeptide chains.

2. Sample loading: the protein sample is loaded into a polyacrylamide gel (porous).

3. Electric field: an electric field is applied. Since SDS makes all proteins negatively charged, they migrate toward the positive electrode (anode). Smaller proteins move faster through the gel, while larger proteins move slower.

2D-PAGE

What?

2D-PAGE (Two-Dimensional Polyacrylamide Gel Electrophoresis) is a method for separating complex protein mixtures based on two independent properties:

1. Isoelectric point (pI)

2. Molecular weight

In other words, it combines both isoelectric focusing and SDS-PAGE into one procedure.

How?

Two electrophoresis steps performed sequentially.

1. Isoelectric focusing to separate proteins based on their isoelectric point (pI)

   • Proteins are loaded onto a pH gradient gel

   • Electric field is applied, and proteins migrate until they reach their pI (i.e. where their net charge is 0)

2. SDS-PAGE to separate proteins based on their molecular weight

   • Focused proteins from the IEF strip are laid horizontally onto a SDS-PAGE gel.

   • SDS coats the proteins, giving them a uniform negative charge.

   • An electric field is applied, and proteins migrate based on their size (smaller proteins move faster).

Final result: a 2D protein map, where each protein appears as a spot on the gel.

• The horizontal position (left to right) corresponds to pI.

• The vertical position (top to bottom) corresponds to molecular weight.

Why?

• Protein identification: specific spots can be excised and analysed in a mass-spectrometer for identification.

• Comparative analysis using “spot” comparison.

Mobile and stationary phase

In chromatography, separation of compounds occurs due to the interaction between two phases:

1. Stationary Phase – The immobile phase inside the column that interacts with the sample components.

2. Mobile Phase – The moving liquid or gas that carries the sample through the stationary phase.

Reverse-Phased Liquid Chromatography (RP-LC)

What?

Reverse-Phase Liquid Chromatography (RP-LC) is a chromatography technique which separates compounds based on their hydrophobicity using a nonpolar stationary phase and a polar mobile phase.

This is the opposite of normal-phase liquid chromatography, where the stationary phase is polar and the mobile phase is non-polar.

How?

1. Column and Phases:

   • Stationary phase: the column is packed with hydrophobic, nonpolar material, typically silica particles modified with long hydrocarbon chains (C18 is the most common as it offers strong hydrophobic interactions).

   • Mobile phase: the peptides (proteins are digested into smaller peptides before RP-LC) are dissolved in a polar solvent - the mobile phase - which is a mixture of water and an organic solvent (e.g. acetonitrile).

2. Sample injection:

   • The sample is injected into the column, and the mobile phase carries the peptides through it.

3. Separation mechanism:

   • Non-polar peptides (hydrophobic) will interact more strongly with the hydrophobic stationary phase, and will thus travel through the column slowly.

   • Polar peptides (hydrophilic) will not interact with the stationary phase as much, and will thus travel through the column faster.

4. Elution process:

   • Initially, the mobile phase has a higher percentage of water, and is thus very polar.

   • Over time, the percentage of organic solvent (acetonitrile) increases. This reduces the polarity of the solvent, thus making it easier for the hydrophobic peptides to dissolve in the mobile phase, and reducing the hydrophobic interactions between the peptides and the stationary phase. Thus, the more hydrophobic compounds will gradually elute out of the column.

   • As the mobile phase reaches a high percentage of organic solvent, even the most strongly retained compounds elute from the column.

5. Detection:

   • A detector connected to the outlet of the column monitors each eluting compound from the column.

   • This results in a chromatograph.

   • Often, the output of the column is directly connected to the mass spectrometer, allowing for separation of the proteins and direct identification.

Explain how chromatographs from RP-LC are interpreted.

In a RP-LC chromatograph:

• The x-axis corresponds to the retention time (time from injection in the column until the peak is detected.

• The y-axis corresponds to intensity.

• Each peak corresponds to a distinct protein or peptide. The position of the peak on the x-axis tells us when the compound exited the column (and thus its relative hydrophobicity), and the height of the peak tells us the abundance of the compound.

• Peaks with a shorter retention time (i.e. on the left) correspond to compounds that are more hydrophilic (polar).

• Peaks with longer retention time (i.e. on the right) correspond to compounds that are more hydrophobic (non-polar).

Basic workflow for quantitative proteomics (5 steps)

1. Protein extraction

2. Protein digestion

3. Peptide separation (e.g. RP-LC)

4. Mass analysis (with mass spectrometry)

5. Peptide identification and quantification

Substeps in protein extraction step of quantitative proteomics workflow

1. Extraction buffer

2. Denaturing agent removal

3. Extracted protein quantitation

What are the key considerations when choosing a protein extraction buffer?

• The buffer should be compatible with the enzymes used for the digestion step (i.e. they must be stable at the pH of the buffer).

• The buffer should solubilise most proteins in the biological sample.

• Choose if we want a denaturing buffer and a native buffer.

  • Denaturing buffer: linearises the proteins to make them inactive, thus limiting endogenous enzyme activity (important for proteins such as protease, phosphatase…).

  • Native buffer: leaves the proteins in their native form, allowing them to stay active and/or bound in complexes. This is useful if we want to study said protein complexes.

• If using a denaturing buffer, important that it is easily removable. This is because we later need to add proteases to break the proteins into peptides, and these will not work if they are denatured.

Safe choice of buffer for protein extraction

8M urea (denaturant) in 100mM Tris (buffering agent), pH 8.5

Additional steps to take to optimise protein extraction.

• Add reducing agents to break disulphide bridges → DTT, beta-mercaptoethanol

• Add alkylation agent to irreversibly modify free thiols after reduction (thus preventing disulphide bonds reformation) —> Iodoacetamide, chloroacetamide, N-ethylmaleimide (NEM)

• Adjust salt concentration to optimize protein solubility and prevent aggregation

• If using a weak detergent, add protease, phosphatase, deubiquitinase inhibitors to prevent protein degradation.

• Add benzoase or nucleases to free DNA/RNA bound proteins.

Detergent removal in protein extraction step

If using a denaturing buffer, it is important to remove it before we add proteases to break the proteins into peptides, as the proteases will not work if they are denatured.

One method to do this is by precipitation and buffer exchange:

1. The proteins are precipitated by adding an organic solvent such as acetone or trichloroacetic acid (TCA), thus separating them from the denaturing agent.

2. Incubation and centrifugation to pellet the proteins.

3. Protein pellet is resuspended in a native buffer.

Solid phase digestion methods

• FASP → Filter-Aided Sample Preparation

• S-Trap → Suspension-Trap

• SP3 → Single-Pot Solid-Phase-enhanced Sample Preparation

These are modern sample preparation techniques used in proteomics workflows to process proteins before mass spectrometry (MS) analysis. These methods are specifically designed to remove denaturants while efficiently digesting proteins into peptides.

Protein quantitation in protein extraction step

2 methods:

1. Bichinchonic acid (BCA) assay

2. Bradford assay

Explain how protein digestion is usually carried out

Give 2 common protocols for protein sample preparation

• FASP: Filter-Aided Sample Preparation

• SP3: Single Pot, Solid-Phase, Sample Preparation

FASP

SP3