BIOL3013 Lecture 10-13 Paul Skipp: Mass spectrometry

Explain how electrospray ionisation (ESI) works

Involves applying high potential to a capillary containing the analyte

• Electrophoretic effect= influence of an io’s movement on surrounding solvent & other ions causes the solvent to be dragged along with it= movement is driven by pressure of ions dragging others through the capillary

What is MALDI?

Matrix-assisted laser desorption technique

• add the protein to a low Mw, UV absorbing matric by spotting it onto a plate

• Fire a UV laser at the matrix which gives it energy and causes vaporisation

• better for organic samples like tissue

Advantages of MALDI over ESI & vice versa

MALDI:

• tolerance to presence of salts, detergents & buffers making it great for biological samples

• Simple & fast sample preparation, can be highly-automated in a high-throughput analysis

• produces predominantly singly-charged ions leading to simpler spectra

ESI:

• amenability to online coupling= coupled with LC-MS it allows for continuous separation, ionisation & detection, MALDI is primarily an offline technique it is not as robust as ESI-LC-MS

• Higher resolution & mass accuracy (usually)

• Good for less volatile compounds due to its soft ionisation mechanism

Explain how a TOF mass spectrometry analyser woks

Acceleration: uses an electric field so all have the same energy

Flight tube: of known distance where the ions hit a detector at the end, a vacuu

What does a quadrupole analyser do?

Acts as a mass filter that can separate proteins using a combination of DC & RF fields, taking advantage of differential m/z values

• uses 4 metal rods with different potentials applied, manipulated to select for certain molecules for analysis of single proteins prior to analysis

• also measures the ms value, hence why it is an analyser

Explain steps of Q-TOF analyser

• ESI/MALDI for ion vaporisation

• Stepwave ion guide: removes anything that isn’t an ion

• Quadrupole analyser: mass filter, letting only specific m/z values thruogh

• Collision cells: use light gases like helium or argon to cause collision-induced dissociation (CID) of ions

• QuaTOF: a W-shaped TOF analsyer utilising reflectrons to measure the TOF, an MS2 scan or ms/ms

Explain the parts of an LTQ orbitrap hybrid mass spectrometer

• ESI for vaporisation of ions

• Linear ion trap: traps ions & scans them out depending on their m/z ration

• C-trap: compresses ions before injecting them into the Orbitrap analyser

• Orbitrap: a high resolution mass analyser that utilises inner & outer electrodes so that the ions oscillate back and forth, generating a sine wave. this wave can be translated into the m/z value using the oscillation frequency & instrumental constant

Explain how isotopes like C13 affect mass spectra

• 1.08% of carbon is C13

• Results in the same molecule having multiple Mw in a sample, with large Mw molecules having greater variation as a result

• REsults in multiple spectra, often like a bell shaped curve or a negatively skewed distribution for smaller molecules

Difference between top-down & bottom-up proteomic workflows

Top-down: focus on analysing complete protein structure

• separate inact poteins, direct ms/ms analysis of this

• then try to put protein back together based on intact asses & fragmentation patterns

Bottom-up: more common, involves fragmenting proteins and then putting back together into the whole one

• enzymatic digestion (trypsin) of proteins into smaller peptides before separation, then ms

• identify proteins structure by matching peptide mass spectra

What is a key issue with ESI and how can it be reduced?

Protein to produce ions with multiple charge states, complicating the mass spectra

• nanoESI reduces this by increased ionisation efficiency and reduce ionisation due to lower sample volume & smaller droplets

Benefits of native mass spectrometry & what conditions are required

Benefits:

• can provide information on protein-ligand interactions, subunit stoichiometry and topology of protein complexes utilising non-native and native spectra, as well as ligand present and ligand absent spectra

Conditions:

• aqueous solution of pH6-9

• Reduced flow rate of 10-100nL/min

• Lower voltage of 0.8-1.5kV

• Lower temperature of 20-30 degrees Celsius

How can native mass spectrometry be used to analyse membrane proteins? why are membrane proteins typically so hard to analyse

Difficulty: hydrophobic nature means that without lipid environment they tend to aggregate or lose their structure

How: utilising detergents to form micelles surrounding the protein, upon entry into the MS vacuum the detergent micelle is removed by the electric field or collision leaving the naked protein complex

• has been done for BtuC2D2 complex

How does labelling mass spectrometry work?

Before proteolytic digestion (bottom-up approach) you incubate the protein with a chemical reagent that modifies residues on the surface of the protein. Upon digestion & LC-MS/MS analysis you can compare this spectra with that of the unlabelled protein to identify modified residues, and hence residues on the exterior of the protein

• spatial information

• topology

• subunit stoichiometry

Name 2 key covalent labelling reagents used in mass spectrometry

Succinimides: bind to lysine residues exposed

Hydroxyl radicals: binds to many different residues, can react with 70% of side chains causing a mass shift of +16Da (O)

What is HRPF & how does it work?

Hydroxyl Radical protein footprinting:

• radiolysis of water generates hydroxy radicals

• these react with side chains causing oxidation modifications

• more modification on areas more solvent-exposed & vice versa

• comparisons of spectra with & without HRPF can reveal exposed peptides, providing structural information, topology, subunit stoichiometry etc

• challenge= can cause H2O2 production on accident which reversibly damages the protein

What does CL-MS do?

Cross-linking mass spectrometry: utilises chemical cross-linkers that react with residues in certain proximity, determined by the length & flexibility fo their cross-linker, keeping residues that are close together upon fragmentation and ms analysis

How can you modify CL-MS experiments?

Enrichment: can enrich cross-linkers with affinity tags (like biotin) for enrichment (using streptavidin) to increase concentration of the chemically cross-linked samples out of your whole sample

Isotope-labelled: using D or C13 in their spacer regions to allow for differentiation of cross-linked peptides in MS analysis by using light & heavy cross-linkers, characteristic double peak shows cross-linked peptide in the MS1 scan

Cleavable: linkers can have cleavable bonds that cleave upon CID resulting in less complicated MS2 due to reduced alteration in mass of fragmented peptides

What is isobaric labelling mass spectrometry primarily for?

The quantification of the relative abundance of different proteins in different samples

How do isobaric tags work?

They have the same overall mass

• reporter group: a small chemical moiety that contains heavy isotopes like C13 & N15. This is designed to be released during fragmentation in the MS2 scan, and is different weight for each protein sample

• Balancer grou: ensures the mass of the entire label is identical across all labels used

Workflow for isobaric labelling mass spectrometry

• add isobaric labels that bind to the digested proteins in the sample using their amine-reactive group (bottom-up flow)

• LC-MS/MS is performed: MS1 spectra shows 1 peak as it is all the same peptide with an identical isobaric label on it

• MS2 shows different relative abundances of the reporter group which is cleaved upon ion fragmentation

What is the purpose of thermal profiling mass spectrometry?

To measure the thermal stability of proteins within their native cellular environment. This provides information about how a protein’s stability changes when a ligand is bound, or when it forms a complex with another protein

How can isobaric labelling & thermal profiling mass spectrometry be combined? give the steps involved

1. Divide lysate into multiple aliquots and heat each to a specific temperature

2. Digestion: use trypsin to enzymatically digest the proteins

3. Isobaric labelling: add a unique lTMT label to each sample, allowing for simultaneous analysis of multiple temperature points in a single MS run

4. LC-MS/MS

5. MS1 scan: all versions of a given peptide from different temperatures appear as a single peak due to their identical mass

6. MS2 scan: fragmented peptides release reporter ions unique to each temperature point. Intensity of each one reflects the relative abundance of the protein at that temperature

7. Relative abundance of each protein across the different temperatures are used to generate a thermal denaturation curve, shifts in these curves= changes in thermal stability, indicating protein-ligand binding or complex formation

What is the purpose of HDX mass spectrometry

Probe the dynamic structure & conformational changes of proteins by measuring solvent accessibility & H-bonding networks within a protein

• identify ligand binding sites

• map solvent accessibility

• measure rate of backbone amide hydrogen exchange with deuterium, correlates with protein’s local flexibility & structural stability

What are the differences between pulse labelling & continuous labelling HDX-MS?

Continuous labelling

• extended, controlled time points= exposure to D2O

• Equilibrium solvent accessibility & stable conformations= primary focus

• Global dynamics & interaction sites= information gained

• More straightforward to implement

Pulse labelling

• bried, rapid exposure= exposure to D2O

• transient, short-lived, or partially unfolded states, and kinetic events= primary focus

• Fast-exchanging populations, folding intermediates= information gained

• More technically challenging due to rapid mixing/quenching