Methods Flashcards
Protein structure (folding, unfolding, aggregation) is governed by […]
Energy landscapes
What are methods that exist to determine protein structure, folding and stability? [6]
- CPU/Bioinformatics
- Spectroscopy
- Scattering
- Calorimetry
- Electrophoresis
- Atomic structure
What are some bioinformatics tools? [5]
- BLASTp
- CLUSTAL Omega
- ProtParam
- Databases:
- PDB
- UniProt
BLASTp is used for […]
sequence search
Clustal Omega is used for […]
multiple sequence alignment
ProtParam is used for […]
physical parameters from sequence
UniProt is used for […]
Database of proteins
PDB is used for […]
Database of high-resolution protein structures
The PDB database is comprised of […]
High resolution protein structures (mostly x-ray)
How many high-resolution protein structures are on PDB?
Reasons that some proteins have no solved structure [4]
- Difficult to express/isolate
- Difficult to study by x-ray, NMR, or cryoEM
- Engineered/completely novel proteins
- No one has gotten around to it yet
What about proteins that are not in the PDB?
Proteins with no solved structurew
Example protein on UniProt.
Pepsin-like protease from Rhodotorula mucilaginosa
* psychrotrophic yeast
* growth temperature range 30 to -10 C
* isolated from permafrost soils in Antarctica
Describe the AlphaFold2 predicted structure of pepsin-like protease from Rhodotorula mucilaginosa L7.
What is AlphaFold
F. Google DeepMind
How does Alpha-Fold work?
- Data entry and database searches
- Sequence analysis
- AI analysis
- Hypothetical structure
How can you interpret Clustal Omegal search results?
(asterisk) = fully conserved residue
: (colon) = conservation of strongly similar properties
. (period) = conservation of weakly similar properties
How does x-ray crystallography work?
- Prepare protein crystals (find condition where they crystallize)
- Build protein model
- Determine positions of atoms from diffraction pattern
- Collect x-ray diffraction images (wavelength = 0.5-2.5 Angstroms) - X-rays will diffract/scatter off of electrons in a very reproducible way (do this from different angles to build a model - cannot see hydrogens, what you see is electron density (so you can tell an oxygen from a nitrogen from a carbon)
What are the main challenges of x-ray crystallography? [3]
- Growing large (mm) crystals
- Rigid, ordered structures are required
- In crystal, not in solution (protein may look different if in solution and fully hydrated as compared to its crystalline structure)
What are the benefits of x-ray crystallography?
- OK for small to large proteins as well as proteins + ligands/assemblies
- High-resolution atomic structures possible (<2Å) - (when using synchrotron sources…$$)
What is synchrotron radiation?
- Electron accelerator + storage ring
- Accelerating electrons give off electromagnetic radiation
- Very intense light source
What is cryoEM?
- pick 1000’s of 2D images
- align them + bin into groups of similar orientation
- 3D model = build from 2D images of molecules in various orientations
PDB 101 video: https://www.youtube.com/watch?v=vLo7oqfRa74
What are the main challenges of cryoEM? [3]
- Samples well dispersed (not clumped) - in order to discern what an individual particles is, they cannot be clumped
- All orientations of the protein observed (otw, get ‘preferred orientation problem’) - kind of like we never see the dark side of the moon, we can never build a 3D model in this case
- possible, but difficult to get <3 Å (due to the above two problems)
What are the benefits of cryoEM? [3]
- High resolution atomic structures possible (< 2Å)
- No size limits
- Large proteins, complexes, viruses, machines
What is nuclear magnetic resonance (NMR)?
- Certain nuclei (see footnote) change spin state in external magnetic field
- Oriented nuclei are able to absorb certain frequencies of light in the magnetic field
- Frequency of energy absorbed depends on the local magnetic field (affected by the molecular structure)
Spin active isotopes in proteins: 13C, 15N, 1H (1H is the naturally occuring isotope of carbon, but 13C and 15N are not naturally occuring, they are the rare heavier isotopes - 12C and 14N are the common isotopes - 2H (deuterium) is the heavier isotope for hydrogen but it is not spin active)
What is the difference between 1D and 2D NMR?
- 1D: one frequency axis and one nuclei type
- 2D: two frequency axes and one or two nuclei types (look for through-bond and through-space effects)
In 2D NMR, what are through-bond and through-space effects?
2D: two frequency axes and one or two nuclei types
- Through-bond = correlation spectroscopy (COSY) - within 3 bonds
- Through-space = nuclear overhauser enhancement spectroscopy (NOESY) - within 5 Å
- Measure 13C, 15N, and 1H spectra
- look for correlations and build structure model
You can tell the difference between through-bond vs. through-space interactions.
What are the main challenges of NMR? [3]
- Isotope labelling protein (13C; 15N)
- Must recombinantly produce the protein with microorganisms fed with isotope-labelled enriched carbon and nitrogen since these are not the naturally occuring isotopic forms
- Not too floppy
- Not too large (< 35,000 Da, <350 aa) - as you get larger, the spectra gets ‘messier’ and it is harder and harder to make correlations
What are the benefits of NMR? [4]
- High-resolution atomic structures possible (<2Å)
- in solution (no crystals; vary conditions) - might be more accurate because of this
- ligands
- Instrumentation is highly accessible/available
Define: spectroscopy
The study of the interaction (absorption and/or emission) of electromagnetic radiation with matter
Describe the relationships between light and energy.
- Energy is proportional to frequency and wavenumber
- Energy is inversely related to wavelength
- Wavenumber = # of wavelengths per unit length
- Frequency = # of occurences per unit time
Energy is […] to frequency and wavenumber.
proportional
Energy is […] related to wavelength.
inversely
Wavenumber is:
of wavelengths per unit length
Frequency is:
of occurences per unit time
e.g., cycles per second
What are the relevant equations regarding light and energy?
Describe the electromagnetic spectrum.
- Visible spectrum is from 700 to 420 nm
- Radio waves have very long wavelengths and very low energy
- Gamma rays; x-rays etc… have very short wavelengths and very high energy
What is ‘quantization of energy’?
- A given transition corresponds to a certain energy
- Need a photon of particular energy to cause a given transition
What are electronic, vibrational, and rotational transitions?
What wavelengths are relevant for each type of transition?
Electronic: UV-VIS (180 - 750 nm)
Vibrational: IR (0.78 - 300 um)
Rotational: Microwave (0.75 - 3.75 mm)
E stands for energy (in Joules),
v stands for frequency [in reciprocal seconds – written s-1 or Hertz (Hz)- 1Hz = 1 s-1),
h is Planck’s constant. (6.626x10^-34 J.s)
What compounds absorb 200-800 nm?
UV-VIS: electronic transitions
n → π^star transition: source of nonbonding valence electrons → N, O, S, P
π → π^star transition: source of π electrons → double, triple bonds
It takes less energy (lower wavelength) to excite conjugated (pigments!) systems. This is why we can’t see tryptophan with the naked eye, but you can see beta-carotene.
Define: wavelength
the distance between successive maxima
Define amplitude
magnitude of the electric vector at the wave maxima
Describe the UV absorption of proteins.
- 190 - 220 nm peaks are common to all proteins (backbone transitions)
- Only proteins with Trp and Try have appreciable absorption around 280 nm
- Phenylalanine has a very small absorption at 260 nm (shown on the graph it is multipled by a factor of 10 so it can be seen)
Used mainly for detecting proteins and measuring protein concentration. Not used as much for protein structural determination anymore.
Describe the Beer-Lambert Law.
- Measure absorbance at 280 nm
- Interference by aggregation (can cause scattering), DNA and other chromophores (e.g., bound groups that absorb at 280 nm)
- Valid over Abs ~0.2-1
- Extinction coefficient, absorptivity (ε M-1cm-1)
- Pathlength (l cm)
- Concentration of absorbing species (c M)
A = ε l c
Can determine extinction coefficient using bioinformatics (if the protein is known).
Longer wavelength means: [2]
lower frequency and lower energy
What is circular dichroism spectroscopy?
Absorption spectroscopy with circularly polarized light
What is the CD effect?
- Circular dichroism is due to the differential absorption of left- vs. right-handed circularly polarized light.
- Induces ellipticity, θ
Circular dichroism spectroscopy requires […]
Optically active molecules: asymmetric in structure (e.g., glycine is not chiral, therefore not asymmetric in structure and not optically active)
What is an optically active molecule?
Asymmetric in structure
Absorption of circularly polarized light is […] to secondary structure.
Inversely proportional
This can also be useful information for predicting tertiary and quaternary structure.
How can CD can be quantitative?
- Estimating secondary structure content (%)
Give examples of applications of pepsin-like proteases.
- Industry: dairy; meat; leather; detergents
- Medicine: malaria (plasmepsins); hypertension (rennin); Alzheimer’s (beta-secretase)
Native pepsin is […] at pH > 6, heat, or urea.
Irreversibly denatured
Describe how CD was used to measure pepsin secondary structure.
- To elucidate why pepsin is irreversibly denatured at pH > 6, heat, or urea.
What is fluorescence spectroscopy?
- Typically absorbed energy is dissipated as radiationless decay (e.g., energy is released as heat)
- Molecules that fluoresce (like tryptophan) will lose some energy by radiationless decay (collisions with the solvent for example), and then will give off a photon (i.e., emit) at a lower energy level (higher wavelength)
What are the properties of fluorescence molecules? [5]
- excitation (λEX) and emission (λEM) wavelengths corresponding to max intensity
- Stokes shift (λEM − λEX)
- Extinction coefficient
- Quantum yield (conversion of absorption → emission)
- Lifetime (duration of excited state)
What compounds give fluorescence?
- Conjugated systems (more conjugation = more stable excited state = more likely to emit light)
- Tryptophan is a natural tag on proteins because it fluoresces (relying on tyrosine is not as useful because the signal-to-noise ratio is too high)
How can fluorescence be used to asses protein structure using tryptophan?
- By measuring fluorescence at different pH levels we can infer informatino about the structure based on level of fluorescence.
- Tryptophan is a built-in fluoretic probe that can be used to study protein folding
Describe use of ANS in fluorescent dye-binding experiments.
- You can measure surface hydrophobicity with this method.
- The dye will binding to the hydrophobic patches of the protein, and so you can use ANS binding to look at the protein under different conditions to get a measure of how different treatments affect the protein and it’s surface hydrophobicity (an important feature for properties like emulsion ability etc.)
How can dye-binding indicate amyloid formation?
- This is often used to measure kinetics of amyloid formation under a given set of conditions through THT fluorescence
What is the difference between a fluorescence and UV-VIS spectrometer?
- Location of the detector is different.
- Fluorescence - detector is 90degrees to the sample so that only emitted light is detected (this makes sensitivity orders of magnitude (thousands) greater than absorbance)
- Absorbance - detector is right behind sample (light path is straight line)
What are molecular vibrations?
A molecule can absorb IR radiation if it vibrates such that its charge distribution (therefore, its electric dipole moment) changes during vibration
If the v of light matches v of vibrations = absorb photon (hv)
IR absorption is […]
Unique to each group of atoms
Why is IR absorption unique to each group of atoms?
- Different functional groups:
- different bond strengths (related to k)
- different masses (e.g., H, C, N, O)
- thus, vibrate at different frequencies
- thus, absorb different energies
What is functional group spectroscopy?
Mid-IR (4000-650cm^-1)
Wavenumber is inverse of wavelength (reported this way by convention).
Describe how FTIR spectroscopy can be used to study protein structure, dynamics.
Position & intensity of peaks is inversely proportional to secondary structure
Describe transmission IR.
- Used to measure liquids
- High absorptivity, thus use small very small pathlengths (0.01-1.0 mm) - hard to load samples in practice due to the small path length
- Can be used for solids if sample is ground and pelleted with potassium bromide, or dispersed in Nujol mineral oil
Describe attenuated total reflectance IR.
- measures total energy reflected from surface of sample in contact with IR transmitting crystal
- used to measure solids, pastes, and viscous liquids (e.g., peanut butter)
Easier to load samples in this method as compared to transmission IR.
What is small-angle neutron scattering?
- Where the scattered particle ends up is proportional to the size and shape of the particle it collided with.
Neutrons are particles NOT electromagnetic radiation.
The classical way of generating neutrons for this method is a nuclear reactor.
Describe use of SANS to study pepsin.
- Radius of gyration: approximates center of mass as a sphere.
Why do people bother with neutron scattering even though nuclear reactors are required to generate neutrons?
- X-rays ‘see’ electrons; x-rays will ‘see’ electron density
- Neutrons are totally different; they interact very well with hydrogens more than anything else, but they don’t interact nearly as well with deuterium (heavy hydrogen) - so in this way we can make water invisible (use heavy water) to only see the hydrogens on the protein.
How does differential scanning calorimetry work?
- Heat protein; it unfolds (protein is taking in energy to break these bonds); so the instrument has to apply more power to the sample than it does to the reference cell (usually water) to maintain the temperature
- Tm is the denaturation midpoint
- Change in heat capacity (baseline before and after unfolding) are related to exposing the buried hydrophobic amino acids - so a larger change in heat capacity would result in a greater delta-Cp.
What is the only method that directly measures enthalpy?
Calorimetry
This is the best method for determining thermodynamic stability.
Describe DSC to study pepsin stability.
- Notice how between the first and second scans the Tm is very different (i.e., pepsin does not fold back to native pepsin after being denatured)
- 2nd scan Tm is at a higher temperature but is a much shallower peak.
- From pH 5 to pH 8 the tryptophan become exposed to solvent, and there is a loss of beta-sheets and increase in disordered structure - so it becomes expanded and partially unfolded
- Native activity is lost
- In refolding: Partial recovery of secondary structure, tryptophan becomes reburied
- Refolded/denatured form may be more stable than native pepsin.
Calorimetry was essential to piecing this together - observing that the refolded state is thermodynamically stable.
Describe isothermal titration calorimetry.
- Gives information about the thermodynamics of binding (e.g., proteins to proteins; proteins to ligands; etc…)
- Each peak tells you direct information about the enthalpy of binding.
- This method gives information about stoichiometry (how many ligands bind to each protein), thermodynamics (enthalpy), and affinity (binding/association constants - how tightly the molecules bind)
Describe ITC in the study of prion proteins.
- Interested in different ligands that may prevent formation of prions (in this case, PcTS, which has metal ‘M’ in its aromatic structure)
How are so many of these PcTS molcules binding the prion? Likely via pi-pi stacking interactions (see image on the other side of this card).
DCS for […] and ITC for […].
Also mention disadvantages.
- DCS for stability. (This is the gold standard for characterizing thermodynamic stability of a protein. Disadvantage = slower than other methods like fluorescence (which can measure 96 samples at once); only one sample measured at once in DCS; also requires more sample per test)
- ITC for binding. (This is the gold standard for characterizing binding affinity. Disadvantage = screening hundreds/thousands of molecules, this would not be a good method, since only one sample can be measured at one time.)
There are faster methods, but these are the only methods that give the thermodynamis directly without use of a model.
Describe SDS-PAGE.
- Mix sample with SDS buffer
- May include reducing agent
- Heat to denature
- Proteins bind SDS in proportion to their Mw
- All proteins have similar Mw/charge ratio
- Migration distance is proportional to size and molecular weight.
Give an example application of SDS-PAGE.
- We expect protein hydrolysis under acid conditions
- Run samples as a function of time to see how quickly this hydrolysis happens.
What are the advantages and disadvantages of SDS-PAGE?
- Advantage: Very accessible, easy, simple, and affordable; cheapest thing we can do in the lab!
- Disadvantage: Can be hard to resolve proteins in gels.
