The Biomolecular Core Facility within the Centre for Biological Sciences seeks to provide researchers with a comprehensive platform for the preparation and characterisation of biological systems at the molecular level.
Absorption spectroscopy measures the absorption of light by a sample, as a function of wavelength. The most sensitive form of absorption spectrophotometry is dual wavelength spectrophotometry. Here the light from a highly stabilised lamp is divided into two beams with separate grating monochromators and shone onto a single cuvette. The difference in absorption of the two beams is then measured, allowing for accurate determinations of very small changes. Often, one beam will be at a wavelength where there is no change in absorption in the system, providing a stable baseline in a cloudy sample. The technique is about 100 fold more sensitive than normal absorption spectroscopy.
Absorption spectroscopy is widely used to measure concentrations, as the level of absorption of light is related directly to concentration.
Changes in the state of a system can lead to changes in the intensity of absorption of light or to changes in the wavelength of light that is maximally absorbed. It can therefore be used for many of the experiments for which fluorescence is also used, although generally changes in absorption are smaller than changes in fluorescence, and fluorescence requires smaller amounts of material.
Hitachi U-3010 Spectrophotometer, with 6 position sample holder and temperature control.
SLM Amino DW-2000 Dual Wavelength Spectrophotometer, with sample stirrer and temperature control: computer interfaced (pictured above).
Shimadzu UV-3000 Dual Wavelength Spectrophotometer.
Accurate determinations of absorption spectra require a sample with an OD greater than ca 0.1 at the wavelength of interest, although this is significantly lower for dual wavelength spectrophotometry.
Cuvettes are usually 1 cm x 1cm and clear on two sides. About 3.0 - 3.5 ml sample is required. For measurements below about 350 nm, quartz cuvettes are requied, but above about 350 nm disposable plastic cuvettes can be used.
Circular Dichroism (CD) measures the interaction of circularly polarised light with molecules that contain chiral chromophores. In proteins, the arrangement of the peptide bonds in secondary structures such as α-helices and β-sheets leads to characteristic CD spectra, allowing estimates to made of the secondary structure content of proteins. Nucleic acids also have characteristic CD spectra.
If a protein is correctly folded, it will have a characteristic CD spectrum. Mutants of the protein, if they fold the same as the wild type protein, should have the same CD spectra. From a CD spectrum, it is possible to estimate the proportion of α-helix, β-sheet, and random-coil.
Because the CD spectrum of a protein depends on the proportion of α-helix, β-sheet, and random-coil structure, it is possible to use CD to follow protein unfolding/refolding as a function of denaturant (eg guanidinium chloride) or temperature.
CD can be used to study drug binding to DNA and changes in DNA structure.
Jasco J720 CD Spectrophotometer
The light source is a 150W Xenon lamp, which covers the spectral range 180 - 800 nm. The optical path and sample compartment are flushed with nitrogen, as oxygen absorbs short wavelength UV light. CD cuvettes are made of quartz; a 1 cm path length cell requires a volume of 2.5 - 3.5 ml .
Typically a solution would be used with an OD of about 0.9 at the wavelength being used. This equates to a typical protein concentration of ca 0.02mg/ml with a 1 cm path length cell.
The choice of buffer is important. Many buffer components absorb strongly at low wavelengths. A phosphate buffer (10 mM) is suitable, but “Good” buffers such as MOPS should be avoided, as should DTT, imidazole, and chloride. Tris buffer is also suitable, as long as the pH is adjusted with phosphoric acid and not HCl. It is sensible to run a buffer blank first. Interference from buffer salts can be minimised by using a higher protein concentration in a shorter path length cuvette. Cuvettes with path lengths from 10 – 0.1 mm are available.
In electrospray ionisation (ESI) mass spectrometry a protein, DNA, or other molecule dissolved in a suitable solvent is sprayed from a fine electrically charged nozzle into a vacuum. The solvent evaporates from these droplets leaving the protein or other molecules as charged particles; their mass/charge ratio (m/z) is then determined in a mass analyser, from which the molecular mass of the particle can be determined.
Atmospheric pressure chemical ionisation (APCI) is an alternative ionisation technique. In this case the spray droplets are uncharged and are desolvated by passing down a heated quartz tube in a stream of nitrogen. The molecules are ionised by collision with N2 and O2 ions formed from air by a corona discharge needle.
Checking the mass of a protein following mutagenesis. Checking for post-translational modifications. Determining the level of modification of a protein after chemical labelling.
ESI mass spectrometry of proteins is most easily performed under denaturing conditions where the protein is unfolded by the presence of organic solvent and formic acid. However, using an electrospray - time-of-flight instrument with control over the vacuum in the source, it is possible to analyse proteins under native conditions that can maintain non-covalent interactions with protein subunits, ligands, cofactors etc.
Tandem mass spectrometry (MS/MS) enables a variety of experiments to be performed using two mass analysers. Ions selected by the first analyser can be collided with argon and the fragments analysed in the second. For example ‘precursor ion’ scans can be used to identify peptides containing modifications such as phosphorylation or glycosylation and ‘product ion’ scans can give information on the peptide sequence and location of the modification.
Determining the level of incorporation of deuterium after incubation with D2O gives information on accessibility of regions of the protein to solvent. This can be used to investigate protein folding, conformational changes, and protein – protein interactions.
APCI is mainly applied to polar compounds with molecular weight below 1500 Da that do not ionise well by electrospray.
VG Quattro II triple quadrupole tandem mass spectrometer equipped with ESI and APCI sources coupled to HPLC (picture above).
Micromass LCT orthogonal acceleration time-of-flight (oaToFF) mass spectrometer equipped with nano-electrospray source (picture top of page).
Ionic substances such as salts interfere with the electrospray process, as do surface active molecules like detergents. The solvent used to dissolve the sample is usually a mixture of water with an organic solvent such as methanol or acetonitrile, often containing a small amount of acid or base (eg formic acid or ammonia) to provide a suitable level of ionization of the molecule. Volatile buffers such as ammonium acetate can be used when analysing proteins under native conditions.
Non-volatile buffers should be removed by dialysis or buffer exchange into water. The more concentrated a protein sample the better, with 20 μl of a 100 μM protein solution being suitable. A variety of microscale, spin column techniques are available in the unit to desalt and concentrate samples if required. Tris-HCl < 20mM is usually tolerated but phosphate is extremely deleterious.
Some molecules give out light (fluorescence) after they have adsorbed light; the emitted, fluorescence light, is at a longer wavelength than the adsorbed light. A fluorimeter records the intensity and wavelength dependence of the emitted fluorescence light. A fluorimeter contains a powerful light source and a monochromator to select the appropriate wavelength of light to shine on the sample. The light emitted from the sample is passed through a second monochromator and then falls on a detector. To reduce interference between the excitation light beam and the emitted light, fluorescence emission is detected at right angles to the excitation beam. The quality of a fluorimeter is determined by the intensity of the light source, the resolution of the monochromators and the sensitivity of the detector. Also important is the interfacing of the fluorimeter to a computer, allowing for data manipulation.
The wavelength at which the maximum fluorescence intensity is observed, and the intensity of fluorescence, vary with environment and so can be used to detect changes in a system. It is possible to study natural fluorophores such as tryptophan, or fluorescence dyes familiar from their use in fluorescence microscopy.
Ligand binding and conformational changes. These often result in changes in Trp fluorescence for a protein, or in the fluorescence of a dye covalently attached to the protein, and can be used to determine binding constants etc.
Metal ions in cells: chelating fluorescence dyes are available that will diffuse into cells and can be used to measure ion concentrations.
Fluorescence Resonance Energy Transfer (FRET). Two fluorophores in close proximity can transfer energy so that excitation of the first fluorophore leads to fluorescence emission from the second fluorophore. The efficiency of this process depends on the distance between the two fluorophores and so can be used as a ‘molecular ruler.’ FRET can therefore be used to study protein-protein interactions, changes in DNA structure etc.
Fluorescence polarization. There is a time delay (nano seconds) between exciting a molecule and the molecule emitting fluorescence. Motion of the molecule during this time period can be detected as a loss in polarization of fluorescence. Binding of two molecules will result in an increase in effective size and so a reduction in motion, detected as an increase in fluorescence polarization. Fluorescence polarization can therefore be used to detect ligand or drug binding to a protein, protein-protein interactions etc.
SLM 8100 single photon counting fluorimeter. This instrument is equipped with a 500 watt Xenon arc lamp, with high quality grating monochromators on both excitation and emission sides, and variable slit widths. The sample is temperature controlled and stirred. Polarizers are available for measurements of fluorescence polarization. It is interfaced to a computer.
Hitachi F-2500 Fluorimeter. This is an easier instrument to use, but is less sensitive than the SLM 8100. It is equipped with a water bath for temperature control, and is interfaced to a computer.
The concentration of a fluorophore required to obtain a good fluorescence spectrum is typically 1 – 10 μM: because of a phenomenon referred to as the inner filter effect, higher concentrations of a fluorophore can lead to a lower fluorescence intensity. To avoid the inner filter effect, sample absorbances at the excitation and emission wavelengths should be kept below 0.1. Light scatter can be a problem, so filtering the sample is often an advantage. Unwanted fluorescence from impurities in the buffers etc can be a problem, and so the purest possible reagents should be used. Buffer blanks should be run to identify any problems with impurities. Drying of material onto the walls of fluorescence cuvettes should be avoided, and the cuvettes should be carefully washed.
Fluorometer cuvettes are usually 1 cm x 1cm and clear on all four sides. About 3.0 - 3.5 ml sample is normally required but on the Hitachi instruments a minimum of 0.8 ml is sufficient. For measurements below about 350 nm, quartz cuvettes are requied, but above about 350 nm disposable plastic fluorescence cuvettes can be used.
Calorimetry measures the amount of heat absorbed or evolved during a chemical or physical change in a sample. In isothermal titration calorimetry, the temperature of the sample is kept constant. Typically, small volumes of a solution of a ligand are titrated into a larger volume of a solution of protein, and the resulting heat changes are measured. Plotting the data as a function of the molar ratio of ligand to protein gives the ligand/protein binding stoichiometry and the binding constant, together with the enthalpy change for binding.
The major use of ITC is to characterize the stoichiometry of ligand binding to a protein and to determine the binding constant. ITC can also be used to measure the enthalpy change in a chemical reaction, such as the enthalpy change when ATP is hydrolyzed by an ATPase.
Microcal ITC microcalorimeter. The instrument can perform experiments between ca 10°C and 60°C. Experimental data is analyzed using the Microcal software to convert heat changes to the required binding constants.
ITC requires relatively large amounts of protein. Per run, 2.2mL of protein sample is required, at a concentration of 10-100µM. The ligand needs to be highly water soluble, so that a 300μL stock solution of 400-600µM can be prepared per run. Each experiment should be run in triplicate and have a ligand into buffer control for reference for reliable results.
The buffer used to dissolve the protein and that used to dissolve the ligand should be matched exactly (use dialysis) in terms of composition and pH, since simply mixing solutions of different pH and different compositions can lead to significant heat changes.