Stability-Based Purification

 Denaturation fractionation exploits differences in the heat sensitivity of proteins. The three-dimensional (tertiary) structure of proteins is maintained by a number of forces, mainly hydrophobic interactions, hydrogen bonds and sometimes disulfide bridges. The denatured state of a protein is characterised by a disruption of some or all of these bonds, thus yielding an unfolded protein chain that is, in most cases, insoluble. One of the easiest ways to denature proteins in solution is to heat them (thermal denaturation). However, different proteins will denature at different temperatures, depending on their different thermal stabilities; this, in turn, is a measure of the number of bonds holding the tertiary structure together. If the protein of interest is particularly heat stable, then heating the extract to a temperature at which the protein is stable yet other proteins denature can be a very useful preliminary step. The temperature at which the protein being purified is denatured is first determined by a small-scale experiment. Once this temperature is known, it is possible to remove more thermo-labile proteins by heating the mixture to a temperature 5–10 °C below this critical temperature for a period of 15–30 min. The denatured, unwanted proteins are then removed by centrifugation. The presence of the substrate, product or a competitive inhibitor of an enzyme often stabilises it and allows an even higher heat denaturation temperature to be employed.

In a similar way, proteins differ in the ease with which they are denatured by extremes of pH (< 3 and > 10). The sensitivity of the protein under investigation to extreme pH is determined by a small-scale trial. The protein mixture is then adjusted to a pH not less than 1 pH unit from that at which the test protein is precipitated. More sensitive proteins will precipitate and are removed by centrifugation.

Solubility-Based Purification

 Proteins differ in the balance of charged, polar and hydrophobic amino acids that they display on their surfaces. Charged and polar groups on the surface are solvated by water molecules, thus making the protein molecule soluble, whereas hydrophobic residues are masked by water molecules that are necessarily found adjacent to these regions. Since solubility is a consequence of solvation of charged and polar groups on the surfaces of the protein, it follows that, under particular fixed conditions, proteins will differ in their solubilities. In particular, it is possible to exploit the fact that proteins precipitate differentially from solution upon the addition of species such as neutral salts or organic solvents (‘ salting out ’, solubility fractionation). It should be stressed here that these methods precipitate native (i.e. active) protein by aggregation; the insoluble protein typically does not denature during this process.

Salt Fractionation

 Salt fractionation is frequently carried out using ammonium sulfate. With increasing salt concentration, freely available water molecules that can solvate the salt ions become scarce. Water molecules that have been forced into contact with hydrophobic groups on the surface of a protein are increasingly deployed (rather than those involved in solvating polar groups on the protein surface, which are bound by electrostatic interactions and are far less easily given up). Therefore, more and more water molecules are removed from the hydrophobic surface areas of the protein, thus exposing the hydrophobic patches. The exposed hydrophobic patches cause proteins to aggregate by hydrophobic interaction, resulting in precipitation. The first proteins to aggregate are therefore those with the most hydrophobic residues on the surface, followed by those with fewer hydrophobic residues. Clearly, in protein mixtures, the aggregates formed are made of mixtures of more than one protein. Individual identical molecules do not seek out each other, but simply bind to another adjacent molecule with an exposed hydrophobic patch. However, many proteins are precipitated from solution over a narrow range of salt concentrations, making this a suitably simple procedure for enriching the proteins of interest.

As an example, Table 5.3 illustrates the purification of a protein using ammonium sulfate precipitation. As increasing amounts of ammonium sulfate are dissolved in a protein solution, certain proteins start to aggregate and precipitate out of solution. By carrying out a controlled pilot experiment where the percentage of ammonium sulfate is increased stepwise from, say, 10% to 20% to 30% etc., the resultant precipitate at each step being recovered by centrifugation, redissolved in buffer and analysed for the protein of interest, it is possible to determine a fractionation procedure that will give a significantly purified sample. In the example shown in Table 1, the original homogenate was made in 45% ammonium sulfate and the precipitate recovered and discarded. The supernatant was then made in 70% ammonium sulfate, the precipitate collected, redissolved in buffer, and kept, with the supernatant being discarded. This produced a purification factor of 2.7. As can be seen, a significant amount of protein has been removed at this step (237 g of protein) while 81% of the total enzyme present was recovered, i.e. the yield was good. This step has clearly produced an enrichment of the protein of interest from a large volume of extract and at the same time has concentrated the sample.

Table1. Example of a protein purification schedule

Organic Solvent Fractionation

Organic solvent fractionation is based on differences in the solubility of proteins in aqueous solutions containing water-miscible organic solvents such as ethanol, ace tone and butanol. The addition of organic solvent effectively ‘dilutes out’ the water present and reduces the dielectric constant. At the same time, water molecules are deployed to hydrate the organic solvent molecules, thereby removing water molecules from the charged and polar groups on the surface of proteins. This process gradually exposes more and more charged surface groups and leads to aggregation of proteins due to charge (ionic) interactions between molecules. Proteins consequently precipitate in decreasing order of the number of charged groups on their surface as the organic solvent concentration is increased.

Organic polymers can also be used for the fractional precipitation of proteins. This method resembles organic solvent fractionation in its mechanism of action, but requires lower concentrations to cause protein precipitation and is thus less likely to cause protein denaturation. The most commonly used polymer is polyethylene glycol (PEG), with a relative molecular mass in the range 6000–20 000.

Isoelectric Precipitation Fractionation

 Isoelectric precipitation fractionation is based upon the observations that proteins have a solubility minimum at their isoelectric point (pI). At this pH, there are equal numbers of positive and negative charges on the protein molecule; intermolecular repulsions are therefore minimised and protein molecules can approach each other. This therefore allows opposite charges on different molecules to interact, resulting in the formation of insoluble aggregates. The principle can be exploited to remove unwanted protein, by adjusting the pH of the protein extract so as to cause the precipitation of these proteins, but not that of the target protein. Alternatively, the target protein can be removed by adjusting the pH of the extract to the pI of the target protein. In practice, the former alternative is preferable, since some denaturation of the precipitating protein inevitably occurs.

Inclusion Body Purification

 Finally, an unusual solubility phenomenon can be utilised in some cases for protein purification from bacteria. Early workers who were over-expressing heterologous proteins in Escherichia coli at high levels were alarmed to discover that, although their protein was expressed in high yield (up to 40% of the total cell protein), the protein aggregated to form insoluble particles that became known as inclusion bodies. Initially this was seen as a major impediment to the production of proteins in bacteria, since the inclusion bodies effectively carry a mixture of monomeric and polymeric denatured proteins formed by partial or incorrect folding, probably due to the reducing environment of the cytoplasm. However, it was soon realised that this phenome non could be used to advantage in protein purification. The inclusion bodies can be separated from a large proportion of the bacterial cytoplasmic protein by centrifugation, giving an effective purification step. The recovered inclusion bodies must then be solubilised and denatured, typically by addition of 6 M guanidinium hydrochloride (as a chaotropic agent ) in the presence of a reducing agent to disrupt any disulfide bridges. For refolding, the denatured protein is then either diluted into or dialysed against a suitable buffer to attain the active, native conformation. The efficiency of the refolding step varies widely with individual proteins. In some cases, reasonably high yields of pure, refolded protein are obtained. However, there are also other cases, where the refolding of denatured protein proves challenging and/or rather inefficient.

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مواضيع ذات صلة

Chromatographic Methods for Protein Purification

Producing Recombinant Protein

Engineering Proteins for Purification

حوامض أمينية محورة Modified Amino Acids

Amplification methods

Immunoelectrophoresis

Immunoprecipitation

Antibody production

Introduction to Immunological methods

Isoelectric focusing

Pore gradient gel electrophoresis

SDS-PAGE