Mass spectrometry Flashcards
What technology converged to result in the applicability of proteomics?
- 2D gel electrophoresis
- Mass spectrometry
- Nucleotide sequencing ESTs/genome scale
- Algorithms
- Genetic approaches
- Complex mixture analysis LC-MS(/MS)
- Chip-based approaches
Describe the history of mass spectrometry (date, event, person)
Date: 1897
Event: First mass spectrometer
-Demonstrated that electrons have mass
Person: Thomson
Date: Near 1900
Event: Mapped many isotopes with new mass spectrometer
Person: Aston
Date: 1946
Event: Time-of-flight mass spectrometer
Person: Stephens
Date: 1960s
Event: Quadruple mass spectrometer
Person: Paul and Steinweidel
Date: 1960s
Event: Gas chromatography-mass spectrometry (GC-MS)
Person: Golhke
Date: 1968
Event: Electrospray ionization (ESI)
Person: Dole et al.
Date: 1980s
Event: Fast atom bombardment (FAB)
Person: Comigarow et al.
Date: 1989
Event: Matrix Assisted Laser Desorption/Ionization (MALDI)
Person: Tanaka et al.
Date: 1989
Event: Electrospray ionisation (ESI) of biomolecules
Person: Fenn et al.
Date: 1990
Event: Postsource decay-Matrix Assisted Laser Desorption/Ionization (PSD-MALDI)
Person:Spengler et al.
Date: 2000
Event: Orbitrap
Person: Makarov
Who invented the first mass spectrometer and what in what year?
Thomson, 1897
Who mapped many isotopes with new mass spectrometer and what in what year?
Aston, near 1900
Who invented time of flight mass spectrometer and what in what year?
Stephens, near 1946
Who invented quadruple mass spectrometer and what in what year?
Paul and Steinweidel, 1960s
Who invented gas chromatography mass spectrometer and what in what year?
Golhke, 1960s
Who invented electrospray ionisation and what in what year?
Dole et al., 1968
Who invented fast atom bombardment and what in what year?
Comigarow et al., 1980s
Who invented Matrix Assisted Laser Desorption/Ionization (MALDI) and what in what year?
Tanaka et al., 1989
Who invented Electrospray ionisation (ESI) of biomolecules and in what year?
Fenn et al., 1989
Who invented Postsource decay-Matrix Assisted Laser Desorption/Ionization (PSD-MALDI) and in what year?
Spengler et al., 1990
Who invented Orbitrap and in what year?
Makarov, 2000
What are the 3 major components of a mass spectrometer
o Ion source
o Mass analyser
o Detector
What is the role of the ion source in a mass spectrometer?
Ionise atoms or molecules
What are ions?
• Ions are atoms or molecules which have been ionised, that is have a positive or negative charge
What are commonly used ionisation stems for mass spectrometry biological samples?
• Commonly used methods for biological samples
o Matrix assisted laser desorption/ionisation (MALDI)
o Electrospray ionisation (ESI)
What are less commonly used ionisation stems for mass spectrometry biological samples?
• Less frequently used methods
o Electron ionisation (EI)
o Chemical ionisation (CI)
o Fast atom bombardment (FAB)
Describe the role of the mass analyser in mass spectrometry
o Mass analyser
Measure mass
Is there only one type of mass analyser in mass spectrometry?
There are several types of mass analyser
Some mass spectrometers have more than one mass analyser that are used in tandem
What law do mass analysers rely on in mass spectrometry?
All rely on Newton’s second law: Force= mass x acceleration
If there is a constant force, degree of acceleration is proportional to the mass-can calculate mass of ions with this principle
What are the components of the time of flight mass analyser?
• Components:
o Ion source
o Electric grid to apply a pulse of acceleration voltage to accelerate ions down flight tube
o Detector
Is there air in the flight tube the time of flight mass analyser?
• The ions are accelerated down a flight tube in a vacuum
Describe how the time of flight mass analyser calculates mass using its detector
o The time taken between acceleration pulse and detector hit is proportional to mass-to-charge ratio
o Ion that is lightest will hit detector first
o Ion that is heaviest will hit detector last
What are two main potential issues with the traditional linear method of the time-of-flight analyser? Describe
o Positional spread-
Issues-
• What if ion A and B have the same m/z but are formed at different positions within the source region?
• When they reach the flight region, ion B will have a higher velocity than ion A
o Velocity spread
Issues-
• Ion A and B have the same m/z and are formed at the same position within the source region, but B has a larger initial velocity than A
What is a solution to the positional spread issue of the linear time-of-flight analyser?
• Measure ion migration at different points in time
• Want to locate detector at space-focus position
o Input time in which location is expected as per prior knowledge
What is a solution to the velocity spread issue of the linear time-of-flight analyser? Describe
• Delayed Extraction
o Delay typical nsec to usec during this time velocity spread causes the ions to spreadout
o An extraction pulse is then used to extract the ions for analysis
o The extraction pulse transmits more energy to the ions that remained in the source for longer (slower ion)
That is, the slow ion receives more energy and catches up with the fast ion
What is reflectron time-of-flight spectrometry? What is its advantage?
• Reflectron time-of-flight spectrometry
o The ions are accelerated down a flight tube
o Enter electrostatic repeller (or ion mirror) where their flight is reversed
o The ion with the higher velocity penetrates further into the ion mirror than the ion with the same mass but lower velocity and hence takes longer to be reflected by that ion mirror
o The two ions arrive at the detector at the same time
o Improves resolution
What are the advantages of the time-of-flight mass analyser?
o High ion transmission
o High sensitivity
o Unlimited mass range in theory
But has to be ionizable
o Complete mass spectrum acquired for each ionization pulse
o Ideally matched to MALDI due to the pulsed nature of the laser
o Readily coupled to continuous ion sources (e.g. ESI) via orthogonal injection
o Relatively low cost
o Easy to operate and maintain
o Very good mass resolution with delayed extraction and reflectron mode
High resolution- 10,000-80,000
Good mass accuracy- typically 1-10ppm
What are the biological applications of the time of flight mass analyser? Describe
o Protein identification PMF Tandem MS o Protein characterization Post-translational modifications o Protein quantitation Signal intensity proportional to abundance o Other biological molecules Glycomics Metabolomics Lipidomics o Mass spectrometry imaging
What is the purpose of the detector in mass spectrometry?
o Detector
Report signals of mass analyser
Converts the impact of ion to measurable electric current
The analogue signal is converted to a digital format that is process by computers into an easily interpretable format
Contrast the past detectors vs current detectors in mass spectrometry
The first detectors were photographic paper of phosphorous screens of cathode ray tubes
Replaced by variety of electronic ion detectors
What is the role of mass spectrometers and an advantage of such spectrometers?
- Mass spectrometers analyse the mass to charge (m/z) ratio of ions
- Mass spectrometers are fast, accurate and extremely sensitive
Is there only one type of mass spectrometer available?
• Wide variety of mass spectrometers available
What is the quality of the spectrum assessed by in mass spectrometers?
• The quality of the spectrum is assessed by resolution and mass accuracy
What are limitations of mass spectrometers?
• Limitations-
o Sample is consumed during analysis
o If it can’t be ionised, it can’t be analysed
o Data processing is heavily reliant on databases of known information
What is the process of matrix assisted laser desorption/ionisation and what does it form?
UV (or occasionally infrared) lasers are focused on a small (100u2m) spot containing sample/matrix
o Sample and matrix mixed together and spotted onto a metal target
The matrix absorbs the laser energy which is transferred to the analyte resulting in a mixture of analyte and matrix ions
Most of the ions are the form [M+H]+
• M= Analyte
• H= proton
What is the sample composed of in a MALDI ionization system?
• Sample a mixture of the analyte and a matrix
Are matrices specific or general to the sample?
o Different types of matrix are used depending on the sample
Which matrix is used for analysis of tryptic peptides and what is the general purpose of such an analysis?
α-cyano-4-hydroxycinnamic acid is optimum matrix for analysis of tryptic peptides for protein identification
Which matrix is used for analysis of phospho/glycopeptides and what is the general purpose of such an analysis?
2,5-dihydroxybenzoic acid is optimum matrix for analysis of phosphopeptides (for post-translation modifications) and glycopeptides
Which matrix is used for analysis of intact proteins?
Sinapinic acid is optimum matrix for analysis of intact proteins
What do all MALDI matrices have in common and why?
o Commonalities are the benzene rings as the matrix is working in conjunction with a laser (compounds have to absorb wavelength of laser)
Describe how electrospray ionisation works and how each component is fed into the system. What is the result of this type of ionisation?
o Electrospray ionisation (ESI)
Analyte solution passes through a fine, charged needle
• Black cable brings in 2000-3000V of current to box containing liquid so that voltage is carried by it
A fine mist of ions is generated
The ions desolvate (aided by sheath gas, typically nitrogen)
• Sheath gas comes in through clear cable through emitter
After emitter, short air gap (05.-1cm) before an orifice (a hole where the ions pass through and enter the instrument)
o Instrument running at high vacuum on the inside
o Removal of air befssssore analysis in mass analyser
Multiply charged ions are formed [M+nH]n+
• Analytes often flow through from an HPLC (high performance liquid chromatography) to a green tube connecting to the needle
On a MALDI data output graph, what is on the x axis (and its range compared to ESI), what is on the y axis, what does each line represent and what does the height of the peaks represent?
• Data output- MALDI
o Mass/charge (m/z) on x axis, abundance %/ion count on y axis
m/z scale goes down to 800 and up to 3500
• Higher m/z than ESI
o Each line is a different analyte
o Height of the peaks is signal intensity (normalised to 100%)
What two graphs can ESI ionization method output produce?
Spectrum-
Total ion chromatogram (TIC)
Describe what is on the x axis (and its range compared to MALDI) and Y axis of a spectrum ESI graph
• Mass/charge (m/z) on x axis, abundance %/ion count on y axis on spectrum graph
o m/z scale goes down to 350 and up to 1650
Lower m/z than MALDI
Describe what is on the x and y axis, as well as what the total ion chromatogram from an ESI is
Total ion chromatogram (TIC)
• Collecting samples as they elute off an HPLC column
• Time on x axis, ion count on y axis
What is resolution of mass spectrometry data measured by and why is resolution important?
• Resolution is measured by:
o Full width at half maximum (FWHM)
Measure how wide the peak is at half its height
o Better the resolution, the more information you can gain
Sort data output from time of flight spectrometry in order of resolution: reflection delayed extraction, linear, linear delayed extraction
o Resolution: linear
Why is resolution of mass spectrometry data important in biological samples?
o Naturally occurring isotopes are similar but it is important to distinguish between them
What is the monoisotopic mass of a peptide and how can you recognise it on a graph?
The monoisotopic mass is the mass of a peptide that is composed only of the most abundant isotopes
• The monoisotopic peak is the first in the series of peaks
When is there a higher probability that a mass spectrometry sample contains unusual isotopes? How can this be recognised?
Bigger molecule= high probability that it contains unusual isotopes
• The higher the probability that the monoisotopic peak will no longer be the highest
How is mass accuracy measured in mass spectrometry and when is it relevant?
o Relevant when searching data
ppm error= (observed mass- actual mass)/actual mass *1000000
What is the mass accuracy for time of flight mass spectrometry instruments?
o For Time Of Flight instruments, typically 0.1Da or 50 parts per million (ppm) but this depends on the size of the peptide
What are the general steps of peptide mass fingerprinting?
- Single protein spot excised from gel
- Peptides released by tryptic digestion and their masses measured using MALDI-TOF mass spectrometry
- Protein sequence databases searched for matches with theoretical masses calculated for all trypsin-released peptides using mass spectrometry data
- Identification and isolation of corresponding gene
In peptide mass fingerprinting, how is a single protein spot excised from the gel? What are prerequisites to it? What are some considerations/choices that have to be considered during the process?
a. Run and stain a 2D gel
b. Identify protein spots of interest
c. Spots cut from gel using a scalpel
d. Wear gloves and lab coat to reduce contamination (such as keratin)
e. Large spots can be cut into pieces
f. Process can be automated using a robot (reduces human error and sample contamination, increase throughput)
What is the process of preparation for peptide digestion in peptide mass fingerprinting?
a. Peptide digestion: reduction and alkylation
i. Destain: remove staining agent
ii. Reduce: remove disulfide bridges within/between protein complexes
1. E.g. DTT (dithiothretoil) and tributylphosphine
iii. Alkylate: To prevent reformation of disulfide bonds and allow proteases to get full access to protein access+ protein unfolding
1. E.g. Iodoacetamide and acrymalide
How does peptide digestion occur in peptide mass fingerprinting?
b. Peptide digestion:
i. Get protease into gel plug
1. Dry gel plugs to facilitate protease access
ii. Add solution of proteases
iii. Incubate (37oC, overnight)
1. Protease will cut up protein
2. Peptides will diffuse out into solution
a. Can use organic solvents for spots with lower abundance for higher concentration
iv. Extract peptides for analysis in the mass spectrometer
What is the process, advantages and disadvantages of automating peptide digestion in peptide mass fingerprinting?
c. Automation of peptide digestion
i. MultiPROBE liquid handling station
1. Has 8 pipette holding tips to add reagents required to sample
ii. Automates digestion and target spotting steps
iii. Pros:
1. Can set up and leave
2. Lower contamination
3. Lower human error
4. Higher reproducibility
5. Higher throughput
6. Electronic record sample location
iv. Cons:
1. Takes time to set up
2. Gel plug can be aspirated and lost
What is the ideal protease for peptide digestion in peptide mass fingerprinting?
i. Ideal protease:
1. Robust
2. Efficient
3. Quick-acting
4. Cut in predictable sites
What is the most common protease for peptide digestion in peptide mass fingerprinting? Describe its cutting patterns
- Serine endopeptidase
a. Chops in middle of sequence - Cuts at lysine (K) and arginine (R)
a. Cuts on C-terminal side of the amino acid - Won’t cut if followed by proline (P)
What are other proteases used in peptide digestion in peptide mass fingerprinting? Describe their cutting patterns
- Endoproteinase Lys-C
a. Specificity: C-terminal to lysine (K), but not if followed by P
b. Can be give bigger overall peptide segments - Endoproteinase Asp-N
a. Specificity: N-terminal to aspartic acid (D)
b. Robust enzyme - Chymotrypsin
a. Specificity: C-terminal to aromatic amino acids, but not if followed by P
b. Broad range of sites- not as specific
When is trypsin not used as the protease of choice for peptide digestion in peptide mass fingerprinting?
iii. Other proteases for in-gel digestion
1. May want to use other proteinase if:
a. If the protein is very K/R rich (or poor)
b. If you are trying to get complete sequence coverage
i. Want to use overlapping proteases
How are the digested peptides analysed during peptide mass fingerprinting?
i. Peptides produced by protease digestion analysed by mass spectrometry
1. E.g. MALDI-TOF
ii. Peptides mixed with a matrix, spotted onto a target, ionised, and a time-of-flight mass spectrum collected
iii. Once spectrum collected, the monoisotopic masses can be determined
1. Two proteins can be identified in one spot
a. However, more than two is a bit more difficult
2. Proteins can also be fragmented
Why is it important to have high quality samples during peptide mass fingerprinting and what are the properties of such a sample?
i. Important to have high quality samples
1. Better sample=better result
2. Minimal contaminants
3. No detergent present
4. Low salt concentration
5. fmole-pmole range (10-15 to 10-12)
How is low salt concentration achieved in a sample for peptide mass fingerprinting and what is the process of this/ how is it made?
a. Reverse phase chromatography clean-up
i. Peptide cleanup
1. Use micro-reverse phase columns
2. Looks like pipette tip, but small plug of C-18 resin at the end
a. Can be made using a range of resins
i. R1,R2 and oligo R3
ii. Anion exchange
iii. Graphite (glycopeptides)
iv. IMAC (phosphopeptides)
3. C-18 is hydrophobic
4. Peptides stick, salt and detergents flow through
5. Peptides eluted with organic phase e.g. acetonitrile or matrix
a. Stepwise elution or elution with matrix solution: higher sequence coverage- easier to identify
What is the difference between normal phase and reverse phase?
a. Normal phase- the solvent is less polar than the packing material
b. Reverse phase- the solvent is more polar than the packing material
When searching databases for matches with peptide results in peptide mass fingerprinting, why is it important to stick to a narrow mass range?
a. Stick a narrow mass range
i. Too low: get too many matrix ions and these results can be the result of noise
ii. Too high: The bigger the peptide->the less it ionizes-> the more the abundance goes down
When searching databases for matches with peptide results in peptide mass fingerprinting, what is considered to be good coverage?
b. Sequence coverage of more than 50% is good coverage
Does increasing mass accuracy for peptide mass fingerprinting always useful? Why/why not?
c. Number of peptides vs MS mass error
i. Curve flattens below 5ppm (around 0.05Da for a 1000 Da peptide)
1. That is, increasing mass accuracy for PMF (peptide mass fingerprinting) has limited value
Describe the 1st generation scoring algorithms used in peptide mass fingerprinting
i. Early algorithms added up the number of matching masses within a mass tolerance e.g. PepSea
1. However, this can lead to false positives, where larger incorrect proteins get more matches than smaller correct ones
i. Early algorithms added up the number of matching masses within a mass tolerance e.g. PepSea
1. However, this can lead to false positives, where larger incorrect proteins get more matches than smaller correct ones
Describe the 2nd generation scoring algorithms used in peptide mass fingerprinting
ii. 2nd generation algorithms weighted matches e.g. Molecular Weight Search (MOWSE)
1. Divide number of matches by molecular weight of hit
Describe the 3rd generation scoring algorithms used in peptide mass fingerprinting
iii. 3rd generation algorithms combined the weighted scoring with probability scoring systems (e.g. MASCOT)
1. Enable estimates to be made of random matches>thus assign significance levels (p<0.05)
Describe the properties that need to be looked for in a MASCOT search result in peptide mass fingerprinting
- MASCOT-Makes a graph of statistical significance-anything shaded in green is not statistically significant
a. Allows for comparison of mass-mass of hit vs mass of band on gel
b. Need a good score-score quality varies along search paramaters
c. Should check whether the protein that comes out is the right species (the species that was analysed)
d. How many of submitted masses got matched
i. Want 2/3rds or more to be mapped
e. Want good % coverage
f. Want expected mass accuracy to be consistent and within expectations of instrument used
g. Look at properties- want high number of matched peptides, low number of missed cleaves and oxidised methionines
What are peptide mass fingerprinting scores based on?
iv. Peptide Mass Fingerprinting scores are all effectively based on:
1. Number of matching peptides
2. % of the sequence covered by those matching peptides
What is the process of searching a database with a result from peptide mass fingerprinting?
a. Database searching
i. Create an in silico digest of the protein database
ii. Compare your observed masses to the database
iii. Increased mass accuracy decreases number of matches
iv. Increased number masses decreases number of matches
v. Modifications
Does increased mass accuracy increase or decrease number of matches?
Increased mass accuracy decreases number of matches
Does increased number masses increase or decrease number of matches?
Increased number masses decreases number of matches
What are the 3 types of databases, the number of protein sequences they have and a description of them
Name: SwissProt
Number of protein sequences: 555100
Description: High level of annotation, limited coverage
Name: TrEMBL
Number of protein sequences: 88,032,926
Description: Comprehensive, but limited annotation
Name: NCBInr (a non-redundant DB)
Number of protein sequences: 91,680,400
Description: Composite of several databases, but with duplicate sequence
What are possible reasons for getting no matches when searching a protein database with results from peptide mass fingerprinting?
i. Possible reasons for not getting an identification
1. Spectrum poorly calibrated
2. Few peptide masses (membrane proteins, small proteins, lack of tryptic sites)
a. Membrane proteins don’t solubilise well- will not work well with mass spectrometry
3. Weak spectrum (low protein abundance or large protein= lower molecular concentration)
4. Protein not in database
5. Contaminant peaks e.g. matrix, trypsin, keratin
6. Contamination with salt (ion suppression)
a. When contaminants ionise preferentially, they will supress the sample (peptides) that don’t ionise as well
7. Non tryptic protein fragments
8. Multiple protein species within a spot/gel band
What are the pros of peptide mass fingerprinting?
- High throughput
- MALDI-TOF instruments robust and easy to use
- Amount of available sequence data makes it applicable to most model organisms
- 3rd generation algorithms make the proteins assignments reliable
What are the cons of peptide mass fingerprinting?
- Very small proteins>few peptides
- Protein not in database= no id
- Post translational modifications not detected
- Hard to distinguish between closely related proteins (A problem with higher organisms)
How do beam type analysers work (in general)?
• Ions are separated in time and space under the influence of electric and/or magnetic fields
How does the quadrupole work/how is it constructed? What is their mass resolving power and their mass accuracy, and what is their greatest strength?
o Quadropole-
Ion source at one end, detector at the other
Either the ions corkscrew from the ion source to the detector, or, depending on their mass, they come off in one direction or the other
Consists of 4 parallel rods (circular or hyperbolic)
• Rods- 10 to 15 cm long
• Poles- cm in diameter
Rods generate an electrodynamic quadrupole field by the application of separate potentials to each pair of diagonally opposite rods
The field is alternating (AC) field superimposed on a constant (DC) field
Ions separated based on a stable trajectory of specific m/z values
• At a certain voltage, a certain mass/charge value will make it to the detector whereas a mass/charge value that isn’t resonant will not make it to the detector
Mass resolving power (typically 2000, lower than TOF) and mass accuracy (typically +- 0.1 Da)
• What they lack in resolving power, they make up in speed: very quick analyser
How is a tandem mass spectrometer constructed?
• Ionisation-> 1st mass analyser-> Collision cell-> 2nd mass analyser-> detector
How does a tandem mass spectrometer work/process?
o Ions separated in the first mass analyser- only allows one particular mass through
Ions of interest selected and passed onto collision cell
o Selected ion fragmented in the collision cell
o Fragments analysed in the second mass analyser and reach the detector
What are other names for tandem mass spectrometry?
o Process called:
Tandem mass spectrometry
MSMS
MS2
What are the uses of tandem mass spectrometry and its advantages?
o Tandem MS can:
Quantitative targeted proteomics/metabolomics (SRM)
Provides amino acid sequence information (de novo sequencing)
Can also be used for the characterisation of carbohydrates, phosphopeptides etc.
Not dependent of genomic information
Can be automated for high throughput protein identification
Describe the structure of a triple-quad tandem mass spectrometer
• QQQ (triple-quad) fragmentation o Triple quad is a common platform for MSMS LC-ESI as an ion source Quadrupole 1- precursor ion selection Quadrupole 2- fragmentation Quadrupole 3- fragment ion selection Detector
How does triple-quad fragmentation work?
o Parent ions are fragmented to generate product ions in a collision cell
Fragmentation induced by collision of the parent ion with an inert gas (e.g. Nitrogen or Argon)
What are other names for triple-quad fragmentation?
o Called:
Collisionally-activated dissociation (CAD)
Collision Induced Dissociation (CID)
What is collisional focusing and how does it work?
Once the product ions are formed, as they continue to pass through collision cell, as they bump into more gas molecules, refocuses the ion beam and allows them to pass onto second half of instrument
How are peptides formed? Describe their final structure
Peptides made by condensation reaction
• Amino and carboxyl group react together to form peptide bonds-> forms water as a byproduct
o Amino terminal= residue mass+ 1[H]
o Carboxyl terminal= residue mass + 17 [OH]
[OH] relative to residue mass within the peptide sequence
o Amino terminal (N-terminal)- beginning of amino acid sequence
o Carboxyl terminal (C-terminal)-end of amino acid sequence
What can cause peptides to fragment in triple-quad tandem mass spectrometry, how do they fragment and what is the charge distribution when they fragment
• With CID (collision induced dissociation), peptides fragment preferentially on the peptide bond
o Bond that fragments is relatively random chance- should get reasonably even fragmentation down the peptide chain
o When peptides are fragmented, charge remains on the carboxy terminal of the ion (y series- y will have subscript depending on how many carboxy groups it contains)
o When peptides are fragmented, charge can also remain on the amino terminal of the ion (b series-b will have subscript depending on how many carboxy groups it contains)
o The peptide back bone can fragment at other bonds resulting in x and z ion series or a and c ion series
What is the use of the HPLC system in quadrupole mass spec and what system does it work in conjunction with?
• HPLC system pumps sample into mass spec and Electrospray ion source sprays out liquid as peptides elute off HLPC system into quadrupole mass spec
What is another name for selective reaction monitoring?
• Selective reaction monitoring can also be called MRM (multiple reaction monitoring) or targeted proteomics/metabolomics
How does selective reaction monitoring work in mass spectrometry and why is it useful? What are requirements/goals for optimum selective reaction monitoring?
o Select a particular precursor ion, fragment them, reduce them and select for a particular one in 3rd quadrupole (signal intensity over time)
o Selective reaction monitoring- look for signal intensity over time as sample elutes off column
o Allows quantitation of a known protein in a complex mixture-can’t be used to discover new things
o Allows to do multiple proteins at a time (high throughput technique)
• Data of selective reaction monitoring
o Peaks are mostly 5-10 seconds wide
o Monitor a transition that is Q1 parent ion to Q3 product ion
Multiple transitions per peptide improves robustness
o Usually have multiple peptides per protein and multiple product ions per peptide
Need to determine which product ions will give strongest signal
Increasing robustness
o As peptide elutes off peak, want to measure signal intensity at least 10x across chromatographic peak
Want to identify apex/maximum signal intensity accurately (or at least 5 points across top half of peak)
• More accurate quantition of peptide
What are adjustements to selective reaction monitoring for mass spectrometry that need to be made if we want to monitor more proteins in total in one run and why?
o If want to monitor more proteins in total in one run, better to reduce the dwell time (less time on each protein and get through cycle quicker)
If dwell time is not reduced, excessive amount of proteins monitored will increase transitions, which leads to a decrease of points across the peak, and hence poor quantitation
However, as dwell time is reduced, signal intensity can decrease as well, which will mean lost sensitivity
What is the used of scheduled selective reaction monitoring for mass spectrometry, how is it perfoemd and what is a potential problem with it?
o Can schedule SRM (selected reaction monitoring) in order to monitor more proteins in one run but retain signal intensity and sensitivity
Can design experiment so that mass spectrometer only records certain peptides at certain points in time, depending on when they are expected to elute from the column
• scheduled Selected reaction monitoring (sSRM) doesn’t monitor all transitions all of the time
o Very effective if retention times are stable
• Scheduling transitions to only run when needed, so that can balance cycle and dwell time
• Potential problem: as sequential samples are injected, retention times can drift and start coming out later
What is intelligent/triggered SRM/what does it do in mass spectrometry and why it is useful?
Intelligent/triggered SRM (iSRM/tSRM): one transition is monitored, when above a certain threshold, other starts for a fixed period
• Monitors for more sensitive transition to ensure it isn’t missed, and when its been deemed to have been seen, mass spec starts looking for supplementary transitions for that peptide
o Monitors parent mass and product mass
Describe the structure and idea behind a combined quadrupole time of flight analyser
• Quad’s have lower mass accuracy and resolution than TOFs
o Combine first half of triple quad and time of flight
First half of quadruple- selects particular precursor ions
Collision cell
Quadrupole 2
Time of flight instrument
• Can have dual stage reflection for higher resolution
• Combined together and there is a winning design
o Ion source-> T-wave ion guide- > quadrupole-> Triwave (Trap, ion mobility separation and trasfer)-> QuanTOF (high field pusher, dual stage reflection, ion mirror and ion detection system)
What is the resolution and mass accuracy of a quadrupole analyser?
Resolution: About 2000
Mass accuracy: 1-500 ppm
What is the resolution and mass accuracy of a time of flight analyser?
Resolution: up to 80000
MAss accuracy: about 5 ppm
How is De novo sequencing performed with tandem mass spectrometry?
MSMS application: De novo sequencing
• Collect precursor scan (TOF MS)
• Select the precursor ion to isolate with Q1 then fragment in Q2
• Collect a product ion scan (TOF MS)
• To sequence, go through the peaks and subtract the previous peak from the current peak in order to obtain a mass that matches to an amino acid
• Databases searched using Blastp algorithm
• Settings adjusted for short-nearly-exact-matches
Describe the application of an MS1 park x MS2 park
Selected reaction monitoring:
A signal seen only when a precursor ion selected by MS1 reacts to form a specific product ion selected by MS2 (e.g. for selective quantification)
Describe the application of an MS1 park x MS2 scan
Product scan:
MS2 scans for the product ions formed when the precursor ion selected by MS1 fragments (e.g. de novo sequencing)
Describe the application of an MS1 scan x MS2 park
Precursor scan MS1 scans for the precursor ions formed when the product ion selected by MS1 fragments- screening for a class of precursor with a common product (e.g. phosphopeptides)
Describe the application of an MS1 scan x MS2 scan
Constant neutral loss scan
MS1 and MS2 both scan to maintain a constant m/z difference. Signal recorded at m/z of MS1 when a precursor of this mass loses a neutral fragment equal to the m/z difference between MS1 and MS2
What is the process of an ion trap/tandem in time?
• Process-
o Peptides stored in middle of plate for a period of time by a magnetic field
o Select a peptide for measurement
o Spin out all ions that are not the target ions
By varying the field, the ions can be spun out according to their m/z
o The target is kept and fragmentation energy is provided when there are enough of the target
One ion can be selected, fragmented and the product ions analysed
o When they are fragmented, product/fragment ions that are generated generally don’t break down further
Product ions generated by amide bond breakages
o Fragment ions then spun out to detector and their masses measured
What are the principles behind an ion trap/tandem in time?
- Method of activation
- Product ions
- Spectra interpretation
- Limitations
- Importance of multistage MS
• Principles:
o Mass selective activation
o Product ions generally do not dissociate further
o Spectra are easier to interpret
o Low mass cutoff can limit the information that is obtained (limitation)
Cut-off is around 120 to 180 mass units
o Multistage MS/MS (that is, MSn)
For MSn, a specific product (fragment ion) can be retained and subjected to another round of fragmentation
Important for analysis of signalling pathways and phosphorylation in those signalling pathways
Fragment a fragment- also important for post-translationnel modifications of molecules
What are the limitations of gel technology and mass fingerprinting and why are these limitations?
• High and low mass proteins
o Proteins more than 150 kDa not often let in
o Low mass proteins run off the end of the gel
• Acidic and basic proteins
o Some IPG buffers are not particularly good in the basic range- don’t separate in the basic range very well (after pH9, not very good separation)
Can overcome this with tightly-timed time based regulation
• Hydrophobic proteins
o May not be solubilised
o Do not enter IPG strip or precipitate during focusing
o Bind strongly with the IPG strip and do not enter the slab gel
o Cannot be visualised
• Low abundance proteins (dynamic range)
• Time, cost, reproducibility (inter and intra-laboratory comparisons)
o Data comparisons are difficult
What are the advantages and disadvantages of shotgun proteomics?
• Shotgun proteomics-
o Overcomes limitations of 2DE gels (e.g. can visualise hydrophobic proteins)
o Peptide behaviour more predictable than proteins
o Trypsin digest of whole cell proteins will generate many thousands of peptides-capability to handle complexity needs to be considered
o Disadvantage- lose the context of the protein: protein isoform information
What are other names for shotgun proteomics?
o Also known as bottom-up proteomics or MudPIT (Multi-Dimensional Proteomic Identification Technique)
What is the process of shotgun proteomics?
o Lyse cells or tissue
Subcellular fractionation
o Extract protein
o Enrich
Protein interaction-antibody purification
o Digest
o Separate
Post translational modification separations
• Ionic interaction
• Antibody-based
• Affinity ligand
o Mass spectrometry
Fragment parent ions at the rate of approximately 3-50 per millisecond/second
Make sure mass spectrometer is busy but not overwhelmed
Fragmented ions go into time of flight so that precursor ion spectrum is found
Precursor ions over a specific intensity are fragmented once again and sequenced
o Identify which proteins peptides come from
In shotgun proteomics, how is the sample input into the mass spectrometer regulated?
- Take MS scan every second and most intense ions in the MS scan are fragmented (10 peptide ions in mass spectrometer per second ideally)
- Separation of complex peptide digests (chromatography)
What are the different types of chromatography used in shotgun proteomics?
- Ultra-High-Performance Liquid Chromatography (LC)
- Ion exchange chromatography (CX)
- Reverse phase (RP) chromatography
Why is chromatography used in shotgun proteomics?
o Mass spectrometers have very high capacity (e.g. 10-100 MS/MS events per MS scan in msec-sec)
o The aim is to ensure full coverage
In shotgun proteomics chromatography, what are reproducible factors of peptides that can be used to separate them?
o Reproducible factors of peptides that can be used to separate include:
Charge (peptide isoelectric focusing, SCX (strong cation exchange) chromatography)
Mass (size exclusion chromatography)
Hydrophobicity (reverse phase(hydrophobicity) /HILIC (hydrophilic interaction liquid chromatography) chromatography)
What is the use of ultra-high-performance liquid chromatography?
• Ultra-High-Performance Liquid Chromatography (LC)-
o Where peptide separation occurs (used to fractionate complex mixtures)
What is ion exchange chromatography and how does it work? What is it often used in conjunction with and why?
• Ion exchange chromatography (CX)
o Separation based on charge
o The resin has covalently bonded negatively charged functional groups
Column chromatography (long thin columns)
Cation exchange column
o The analytes of the opposite charge are retained by charge-charge based interaction but can be eluted by slowly increasing the concentration of a similarly charged species that will displace the analyte
E.g. cation exchange (CX) retains positively charged peptides with negatively charged resin whilst those that are negatively charged are repulsed and will not be retained
In CX, positively charged analytes are displaced by the addition of positively charged sodium ions
o Salt inhibits ionization in MS
Therefore, often use in 2D LC-CX followed by RP (reverse phase)
Don’t do ion exchange chromatography on its own- often couple it with a different type of parameter
What is reverse phase chromatography in shotgun proteomics chromatography and how does it work?
o A solute molecule (analyte) binds to an immobilized hydrophobic molecule (the resin) in a polar solvent (e.g. water/formic acid)
o The solute has hydrophobic regions and binds via these to the resin
o A solvent of increasing hydrophobicity (acetonitrile) is used to dissociate the bound analyte
o At a certain point the hydrophobic interaction between the analyte and the resin is less favourable than the interaction between the analyte and the solvent and the analyte elutes
What reverse phase chromotography resin is most often used for peptides
• C-18 for peptides
What reverse phase chromotography resin is most often used for proteins
• Shorter carbon lengths (C-4) for larger molecules (proteins)
What reverse phase chromotography resin is most often used for DNA
• Diphenyl-DNA
What reverse phase chromotography resin is most often used for naturally occurring environmental compounds
• Divinylbenzene (DVB)-Naturally occurring environmental compounds and other analytes
What is offline chromatography?
o Offline-separated analytes are collected into fractions as they elute from the chromatography column
What is online chromatography?
o Online-separated analytes pass directly into the MS ion source for MS analysis as they elute in real time from the chromatography column
Always be at least one dimension of chromatography that will be online
After shotgun proteomics mass spectrometry, what is the process of the automated data analysis (how does the system perform it)
• Asks for all peptides in database that has the found peptide with very high mass accuracy
o Can use peptides with partial sequence information. Requires known:
Molecular weight
M1-M3
Enzyme specificity
• Get database sequences that match precursor peptide mass
• Generate virtual MS-MS spectra
• Compare virtual spectra to real spectrum
• Scoring
o Detect matches between theoretical b and y-ions and actual spectrum ions
o Compute correlation scores
o Rank hits
• Picks peptide sequence with highest score
What are the pros of automated data analysis after shotgun proteomics mass spectrometry?
Reduces Gb of MSMS data to list of proteins
Easy to use
Rapid (with sufficient computer power)
Can’t do it manually if have more than 100,000 MSMS spectra-> good to have computational approaches
What are the cons of automated data analysis after shotgun proteomics mass spectrometry?
Very computer intensive
Need to assess quality of MSMS spectra> limit false discovery rate
Modified peptides may not be matched
Incorrect charge assignment (MS level= peptides not matched)
What are the risks of identifying peptides with automated data analysis in shotgun proteomics?
- With so many peptides (or peptide spectrum matches PSMs) some false positives/error are likely to occur
- Spectra can be of low quality, maybe the peptide sequence is not in the database, charge state of precursor is improperly assigned
What are peptide spectrum matches?
o Peptide spectrum matches- the same peptide is matched hundreds of time
What is done to reduce error in peptide identification during automated data analysis in shotgun proteomics?
• Error is estimated based on an ID software ‘score’
• Employ a false discovery rate (FDR)- take your database and randomize or reverse and then search again- number of positive hits is the FDR
• OR can remove IDs from against the decoy database and search again
o If the spectra match the decoy, it should not be counted
Why is there a statistical advantage to shotgun proteomics?
- A single protein can be represented by many tryptic peptides
- Each identified peptide increases the confidence in the protein identification and quantification
Is the shotgun approach typically performed with ESI or MALDI?
• Shotgun approach typically performed with ESI
Is ESI or MALDI usually used with chromatography techniques? Why? What would be the advantage of combining these approaches?
• Electrospray ionisation is most generally used with chromatography techniques
• 2DLC output can be spotted onto MALDI target
• Complementary to ESI approach: increased coverage
o This is because peptides ionise differently- hence different combinations/approaches are desirable
What did Washburn et al.’s 2001 “large scale analysis of the yeast proteome by multidimensional protein identification technology” discover and how?
• MudPit was first used in “Large-scale analysis of the yeast proteome by multidimensional protein identification technology” by Washburn et al. 2001
o 5540 peptides assigned
o Corresponding to 1484 proteins
o Improved coverage of membrane proteins, basic proteins and very large proteins
o Bias against acidic (pI<4.3) and small proteins
Acidic proteins don’t have many tryptic digest sites
What is used for comparative shotgun proteomics to find protein abundance and why/how?
• Stable isotopes used to quantify changes in protein abundance
o 2H or 13C
o Isotopes have identical chemical properties
• Peptides tagged with differing isotopes will appear adjacent on a mass spectrum
• Area under each peak equate to relative abundance of the two peptides
