Lecture 6: Assessing Mass and Complex Formation Flashcards
What is SEC?
Size exclusion (or gel filtration) chromatography can be used to separate proteins based on size and shape.
• A porous resin is used. Molecules smaller than the pores will enter into them and be slowed down.
• Molecules bigger than the pores will pass around the beads and move faster.
• We can use different columns with different size pores in the beads.
• The sample does not bind to the resin and is not concentrated. The volume loaded onto the column affects the degree of separation of peaks emerging (resolution).
• Big columns have more separation, but they also take longer to run.
• Large columns are used for purification and small columns are used for analytical work.
• The sample is first concentrated to a small volume (less than 5% of the column volume).
• The sample is loaded onto the column in a continuous flow of a buffer in which the protein is stable.
• SEC is very useful for removing aggregated protein.
• We use a series of standards for calibration.
• A SEC has a void volume (V0¬). It is the volume outside the beads. Very large macromolecules elute here.
• The column also has a total volume (VT), where very small things elute.
• The protein will elute between these two volumes if the correct column is being used.
• We create a calibration curve and use it to estimate the mass of a macromolecule based on its elution volume.
• It is simple to use, and various resins are available for different particles.
• The disadvantages are that the sample must be concentrated to get good separation. The elution volume is also dependent on the shape of the protein.
What is multi-angle light scattering?
MALS is a method which we can use to give information about mass and size.
• MALS uses the scattering of a beam of monochromatic light.
• MALS is often carried out after SEC.
• We can use MALS to find stoichiometry.
• The scattering intensity is proportional to the MW and concentration.
• λ is larger than RG¬.
• SEC is used to separate analytes.
• MALS can show molar mass and size of proteins.
• It can also be used to determine protein-protein binding.
How can we use dynamic light scattering?
Dynamic light scattering is used to analyse the motion of macromolecules in solution.
• Static scattering is elastic. The frequency in and out are the same.
• If the scatterer moves than the frequency will be spread by the Doppler effect.
• Diffusion coefficients are measured by DLS.
• They can give information on sample size and aggregation.
• A laser is shone at a small volume. As molecules flow in and out of this volume, this causes changes in the intensity of scattered light.
• The rate of diffusion is related to the diffusion coefficients of the particles involved (DT). Large particles move more slowly.
• DT can be used to calculate the hydrodynamic radius (RH). RH depends on the shape and size of the scattering particle. It cannot give an accurate measurement of mass.
• We can use DLS to study a sample with multiple species or aggregates and get analysis of a sample with a broad distribution of species.
• Scattering intensity is proportional to mass. It can easily detect small amounts of high mass species. DLS can therefore be used to tell how homogenous a sample is by mass.
• DLS is easy and it requires little sample.
How does analytical ultracentrifugation (AUC) work?
AUC can be used to assess the mass and shape of a protein.
• Proteins are separated based on the rate of sedimentation.
• Centrifugation works based on three forces.
• Centrifugal force (F¬C) drives the molecule away from the centre of the rotor.
• The buoyant force (F¬B) comes from the displaced solution causing resistance.
• Frictional force (F¬D) comes from resistance which happens as the molecule moves.
• The particle accelerates until these 3 forces balance.
• Sedimentation rate depends on particle size, particle density, medium density, medium viscosity, angular velocity of the rotor, distance from the axis of rotation and frictional ratio (measure of shape).
• A particle will sediment faster if it has greater density, greater mass or if the centrifugal force is greater.
• A particle will sediment slower if the shape is less spherical (frictional coefficient is bigger) or if the solution is denser.
v = Sw2R
• S = sedimentation coefficient. S depends on size, density and shape. It is expressed in Svedberg units. 1 x 10-13 seconds is equal to 1 Svedberg.
• Most proteins are in the range of 2-25S.
• AUCs contain transparent cells. As the centrifuge spins, the optics allow us to follow its movement within the cell over time. We can detect changes in absorbance or the refractive index.
• When we spin a sample at a constant high speed, we see a region at the top of the tube which has been depleted of sample. A barrier separate the depleted region from the sample.
• The rate that the boundary moves (rb) allows us to determine the sedimentation coefficient.
v = Sw2R
Therefore: drb/dt = Sw2R
• Integration suggests that plotting ln
• If we centrifuge at a lower speed, the sample will not pellet, a concentration gradient will develop. This will reach an equilibrium position. This creates a smooth and stable concentration gradient. The final equilibrium position doesn’t depend on frictional forces as there is no net movement. The final gradient depends on mass and it is independent of shape.
How do pull downs work?
Pull down experiments can be used to test of molecules interact with each other?
• Use a resin that interacts with molecule A. This can be a Ni-NTA resin, or an antibody coupled resin.
• We then incubate with molecule B, wash the resin and see whether B stuck.
• A control is performed without A, in order to check for non-specific binding.
• If molecule B is only pulled down when A is there, then we predict an interaction.
How does isothermal titration calorimetry work?
ITC is a technique which can be used to study the thermodynamics of interactions between two molecules.
• The ITC machine injects molecule A into a cell containing molecule B.
• The machine then tries to maintain the temperature of this cell.
• We measure the energy needed to maintain a constant temperature.
• We can plot the change in enthalpy over time. This gives a series of spike which correspond to the injection of ligand.
• We can integrate this graph to give the total amount of power which is exchanged per injection.
• We can then plot the power for these injections against the molar ratio ligand/protein. The overall change of kcal/mol gives the change in enthalpy. The molar ratio is the midpoint of the graph on the x-axis. The gradient at this point is 1/KD.
• From enthalpy change and fitted KD we can find enthalpy change and overall free energy change.
• This method gives direct thermodynamic measurements.
• However, it does use a lot of sample.
