Lecture 1: Experimental Methods Flashcards
How do proteins fold?
During the 1950s and 1960s, Anfinsen demonstrated that native conformation is determined by amino acid sequence and the totality of interatomic interactions.
• He did this by showing that denaturing and reducing ribonuclease in 8M of urea and mercaptoethanol would revert to the native state if these chemicals were removed.
• Levinthal paradox: A polypeptide chain of 100 amino acids will have 5 x 10¬47 possible conformations. This happens in seconds. There must be a kinetically defined protein folding pathway.
• There is a specific pathway that must be followed, with defined intermediates. Each stage of the process has different properties.
What are the differences between folded proteins, unfolded proteins and protein folding intermediates?
Native folded (N)
- Compact
- Functional. Enzyme activity observed.
- Stable and persistent hydrogen bonds.
- Extensive secondary structure
- Ordered hydrophobic core with specific contacts.
Unfolded (U) • Expanded ensemble of structures. • Generally, do not have native function and activity. • Lack of stable secondary structure. • Lack of specific tertiary contacts.
Protein folding intermediates (I) • Compact, but larger than native. • Known as molten globules. • A mix of N and U forms. • Native levels of secondary structure. • Generally, do not have tertiary contacts.
What is stopped and quenched flow?
Stopped flow can be used to quickly measure folding.
1) Set up two syringes. One has unfolded protein in urea. The other has refolding buffer.
2) Both syringes are depressed and put into a mixing chamber.
3) The syringes mix.
4) The flow will eventually hit a stop syringe. This activates detection and stops flow.
- Quenched flow uses a similar technique. However, it stops the reaction after a certain amount of time. This is achieved using a chemical, light or temperature.
- Both techniques can be linked to near and far UV CD, fluorescence and SAXS.
What is UV CD?
We can use far and near UV CD to study secondary and tertiary structures respectively. We can study what elements form and on what timescale.
• The chiral environment of the peptide backbone in helices and beta sheets gives rise to far UV CD ellipticity.
• This also happens in response to aromatic side chains in folded proteins with tertiary structure.
• Stopped flow far and near UV CD show tertiary and secondary structures form on different timescales.
• A lot of far UV CD is recovered in dead time, indicating a lot of secondary structure formation.
• In stopped flow we can’t measure the entire spectrum, we must select a single wavelength.
• Dead time is the length of time for sample mixing, it’s normally a few ms.
• Near-UV CD recovers a lot more slowly, indicating intermediates don’t have a lot of tertiary structure.
How can we use fluorescence?
- Trp and tyr max wavelength and intensity of fluorescence change based on environment.
- A red shift towards a lower wavelength occurs when a protein unfolds. This is due to an increase in the polarity of the Trp environment,
- We can monitor hydrophobic clustering using ANS dye. It binds to exposed hydrophobic clusters (as seen in partially folded molten globules) and enhances fluorescence.
How can we measure intermediate structure and shape?
We can measure this with SAXS or FRET.
• SAXS can show the distribution of molecular dimensions (RG¬).
• SAXS in real time is not sensitive, but by collecting many data points the kinetics can be extracted.
• Single FRET can show the distance between two parts during folding.
• One of acceptor and a donor can be a trp but the other must be a chromophore attached via a cys residue.
What is hydrogen-deuterium exchange?
We can use this technique to gain residue specific information about secondary structure formation during protein folding.
Amide protons will exchange with deuterium in a heavy water solution.
The intrinsic rate (Kin) depends on pH.
If an amide is involved in hydrogen bonds, the exchange rate (kex) will be much lower.
Protection factor=Kin/Kex
Protection factors can be many orders of magnitude.
We can use this to probe secondary structure with hydrogen bonds at various times during protein folding.
Protected amides in the N state can be used to find how amides are protected during refolding.
What is pulse-labelling hydrogen exchange?
We can use pulse labelling to probe the presence of hydrogen-bonded secondary structure at various time points (ms to sec) during protein folding.
• We denature a protein using urea or guanidine HCl in heavy water.
• All amide sites are deuterated.
• The U protein is then returned to refolding buffer for a variable time, τ (3ms-2s) which allows some areas to refold.
• The solution is then diluted again into a high pH water buffer.
• All exposed deuterons exchange but hydrogen bonded ones will be protected.
• We detect exchange in the stable native state using 2D NMR. E.g. 1H-1H COSY.
• Peak intensity is proportional to occupancy at each site.
What is the folding funnel?
The observation of multiple pathways suggests that U -> I -> N model is too simple.
• A folding funnel is used to describe the multiple pathways that exist in protein folding.
