ISOLATION AND PURIFICATION OF PROTEINS

Protein purification is a series of processes intended to isolate a single type of a protein from a complex mixture. Protein purification is vital for the characterization of the function, structure and interactions of the protein of interest. The starting material can be a biological tissue or a microbial culture.

Purification process involves;

·         freeing the protein from a matrix that confines it

·         separating the protein and non-protein parts of the mixture

·         separating the desired protein from all other proteins.

Separation of one protein from all others may exploit differences in protein size, physico-chemical properties, binding affinity and biological activity.

Purification may be preparative or analytical.

Preparative purifications aim to produce a relatively large quantity of purified proteins for subsequent use. Examples include the preparation of commercial products e.g enzymes (e.g. lactase), nutritional proteins (e.g. soy protein isolate), and certain biopharmaceuticals (e.g. insulin).

Analytical purification produces a relatively small amount of a protein for research or analytical purposes, including identification, quantification, and studies of the protein's structure, post-translational modifications and function. Among the first purified proteins were urease and Concanavalin A.

Extraction

Depending on the source, the protein has to be brought into solution by breaking the tissue or cells containing it.

This can be done through cell disruption to release cellular contents by:

·         Repeated freezing and thawing

·         sonication - applying sound (ultrasound) energy to agitate particles in a sample

·         homogenization by high pressure - intensive blending of mutually related substances or groups of mutually related substances to form a constant of different insoluble phases

·         filtration - the mechanical or physical operation used to separate solids from fluids

·         permeabilization by organic solvents e.g SDS (sodium dodecyl sulfate), Triton X-100 or CHAPS(3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate)

The method of choice depends on how fragile the protein is and how sturdy the cells are.

After extraction process soluble proteins will be in the solvent, and can be separated from cell membranes, DNA etc. by centrifugation.

The extraction process also extracts proteases, which will start digesting the proteins in the solution. If the protein is sensitive to proteolysis, it is usually desirable to proceed quickly, and keep the extract cooled, to slow down proteolysis.

Precipitation and differential solubilization

This is performed by adding increasing amounts of ammonium sulfate - (NH₄)₂SO₄ and collecting the different fractions of precipitate protein.

The hydrophobic groups on the proteins get exposed to the atmosphere and attract other protein hydrophobic groups and get aggregated.

Protein precipitated will be large enough to be visible.

Ammonium sulphate can be removed by dialysis.

Ultracentrifugation

Centrifugation is a process that uses centrifugal force to separate mixtures of particles of varying masses or densities suspended in a liquid, resulting in the pellet and supernatant.

Sucrose gradient centrifugation uses a linear concentration gradient of sugar (sucrose, glycerol, or a silica based density gradient media, like Percoll). A protein sample is layered on top of the gradient and spun at high speeds in an ultracentrifuge. This causes heavy macromolecules to migrate towards the bottom of the tube faster than lighter material. Samples separated by these gradients are referred to as "rate zonal" centrifugations.

After separating the protein/particles, the gradient is then fractionated and collected.

Chromatographic methods

Different proteins interact differently with the column material, and can thus be separated by the time required to pass the column, or the conditions required to elute the protein from the column.

The eluant is usually pooled in different test tubes. Test tubes containing solution with  the protein of interest and any other similar proteins is retained.

Usually proteins are detected as they are coming off the column by their absorbance at 280 nm. Many different chromatographic methods exist:

Size exclusion chromatography

Uses porous gels. The principle is that smaller molecules have to traverse a larger volume in a porous matrix. Consequentially, proteins of a certain range in size will require a variable volume of eluent (solvent) before being collected at the other end of the column of gel.

The eluant is usually pooled in different test tubes. Test tubes containing solution with the protein to purify and any other similarly-sized proteins are retained.

Hydrophobic Interaction Chromatography

The Resins used in the column are amphiphiles with both hydrophobic and hydrophilic regions. The hydrophobic part of the resin attracts hydrophobic region on the proteins. The greater the hydrophobic region on the protein the stronger the attraction between the gel and that particular protein

Ion exchange chromatography

Separates compounds according to the nature and degree of their ionic charge. Anion exchange resins have a positive charge and are used to retain and separate negatively charged compounds, while cation exchange resins have a negative charge and are used to separate positively charged molecules.

Before the separation begins a buffer is pumped through the column to equilibrate the opposing charged ions. Upon injection of the sample, solute molecules will exchange with the buffer ions as each competes for the binding sites on the resin. The length of retention for each solute depends upon the strength of its charge. The most weakly charged compounds will elute first, followed by those with successively stronger charges. Because of the nature of the separating mechanism, pH, buffer type, buffer concentration, and temperature all play important roles in controlling the separation e.g. metal binding (nickel, cobalt)

Ion exchange chromatography is frequently used in both analytical and preparative separations.

Affinity chromatography

It’s based upon molecular conformation, which frequently utilizes application specific resins. These resins have ligands attached to their surfaces which are specific for the compounds to be separated. Most frequently, these ligands function in a fashion similar to that of antibody-antigen interactions. This "lock and key" fit between the ligand and its target compound makes it highly specific, frequently generating a single peak, while all else in the sample is unretained.

E.g. Immuno-affinity chromatography (Ab & Ag)

The protein can be eluted by changing the pH or the salinity

High performance liquid chromatography

Also high pressure liquid chromatography - applies high pressure to drive the solutes through the column faster. This means that the diffusion is limited and the resolution is improved. The most common form is "reversed phase" hplc, where the column material is hydrophobic.

The proteins are eluted by a gradient of increasing amounts of an organic solvent, such as acetonitrile. The proteins elute according to their hydrophobicity.

HPLC purification frequently results in denaturation of the purified proteins, thus not applicable to proteins that do not spontaneously refold.

Concentration of the purified protein

Lyophilization

If the solution doesn't contain any other soluble component than the protein in question the protein can be lyophilized (dried). This is commonly done after an HPLC run. This simply removes all volatile components leaving the proteins behind.

Ultrafiltration

Ultrafiltration concentrates a protein solution using selective permeable membranes. The function of the membrane is to let the water and small molecules pass through while retaining the protein.

Analytical

Denaturing-Condition Electrophoresis

A common laboratory technique that can be used both as preparative and analytical method.

The principle of electrophoresis relies on the movement of a charged ion in an electric field.

In practice, the proteins are denatured in a solution containing a detergent (SDS). In these conditions, the proteins are unfolded and coated with negatively charged detergent molecules. The proteins in SDS-PAGE are separated on the sole basis of their size.

In analytical methods, the proteins migrate as bands based on size. Each band can be detected using stains such as Coomassie blue dye or silver stain.

Preparative methods to purify large amounts of protein require the extraction of the protein from the electrophoretic gel. This extraction may involve excision of the gel containing a band, or eluting the band directly off the gel as it runs off the end of the gel.

It provides an improved resolution over size exclusion chromatography, but does not scale to large quantity of proteins.

Non-Denaturing-Condition Electrophoresis

An important non-denaturing electrophoretic procedure for isolating bioactive metalloproteins in complex protein mixtures is termed 'quantitative native continuous polyacrylamide gel electrophoresis (QPNC-PAGE).

QPNC-PAGE is a high-resolution technique applied in biochemistry and bioinorganic chemistry to separate proteins by isoelectric point (the isoelectric point (pI), is the pH at which a particular molecule or surface carries no net electrical charge).

This variant of gel electrophoresis is used to isolate active or native metalloproteins in biological samples.

The QPNC-PAGE procedure is accomplished in a special electrophoresis chamber for separating charged biomolecules. Due to the specific properties of the prepared gel and electrophoresis buffer solution (which is basic and contains Tris-HCl and NaN₃), most proteins of a biological system are charged negatively in the solution, and will migrate from the cathode to the anode due to the electric field.

MODIFICATION OF PROTEINS

Modification of proteins is done to allow manipulation and study of protein function and interactions in any environment.

Includes crosslinking, fragmenting, denaturing, reducing disulfides, or attaching various prosthetic groups (e.g. PEGylation)

Crosslinking Reagents

Use of chemical crosslinkers and bioconjugation reagents for covalent protein crosslinking techniques to conjugate antibodies, immobilize ligands, attach haptens to carrier proteins, and stabilize folded protein structures and protein interaction complexes.

Amine-to-Amine Crosslinkers

Homobifunctional amine-specific protein crosslinking reagents based on NHS-ester and imidoester reactive groups for selective conjugation of primary amines; available in short, long, cleavable, irreversible, membrane permeable and cell surface varieties.

Amine-to-Sulfhydryl Crosslinkers

Heterobifunctional protein crosslinking reagents for conjugation between primary amine (lysine) and sulfhydryl (cysteine) groups of proteins and other molecules; available with different lengths and types of spacer arms.

Carboxyl-to-Amine Crosslinkers

Carbodiimide crosslinking reagents for conjugation of carboxyl groups (glutamate, aspartate, C-termini) to primary amines (lysine, N-termini); also N-hydroxysuccinimide (NHS) for stable activation of carboxylates for amine-conjugation.

Photoreactive Crosslinkers

Aryl azide, diazirine and other photo-reactive (light-activated) chemical crosslinking reagents to conjugate proteins, nucleic acids and other molecular structures involved in receptor-ligand interaction complexes via two-step activation.

Sulfhydryl-to-Carbohydrate Crosslinkers

Crosslinking reagents based on maleimide and hydrazide reactive groups for conjugation and formation of covalent crosslinks between sulfhydryl (cysteine) and aldehyde (oxidized glycoprotein carbohydrate) groups.

Sulfhydryl-to-Hydroxyl Crosslinkers

Crosslinking reagents based on maleimide and isocyanate reactive groups for conjugation and formation of covalent crosslinks between sulfhydryl and hydroxyl groups.

Sulfhydryl-to-Sulfhydryl Crosslinkers

Sulfhydryl-specific crosslinking reagents based on maleimide or pyridyldithiol reactive groups for selective covalent conjugation of protein and peptide thiols (reduced cysteines) to form stable thioether bonds

Chemical modification Reagents

Use chemical agents to modify amino acid side chains on proteins and peptides in order to alter native charges, block or expose reactive binding sites, inactivate functions, or change functional groups to create targets for crosslinking and labeling.

Aminoethylate Reagent

Converts free sulfhydryls into primary amine groups.

Citraconic Anhydride

Citraconic Anhydride (2-methylmaleic anhydride) reversibly blocks primary amines at pH 8. Amines can be unblocked and returned to their native form.

Iodoacetic Acid

For carboxymethylation.

PEGylation Reagents

Use of activated linear and branched derivatives of polyethylene glycol (PEG) for pegylation and PEG-modification of peptides and proteins via primary amines and sulfhydryl groups to increase solubility, prolong stability and reduce immunogenicity.

Amine-Reactive PEG (Linear)

Methyl-PEG-NHS-ester reagents for PEGylation of proteins and molecules having primary amines. Also called MS(PEG)n, where n = 4, 8, 12 or 24 polyethylene glycol units.

Sulfhydryl-Reactive PEG (Branched)

A branched Methyl-PEG-Maleimide reagent for PEGylation of proteins and molecules having free sulfhydryl groups. Also called TMM(PEG)12 or (Methyl-PEG12)3-PEG4-Maleimide.

Carboxy-PEG-Amine Compounds

PEG amino acids, also called CA(PEG)n, where n = 4, 8, 12 or 24 polyethylene glycol units. The PEGylation compounds can be conjugated to molecules or surfaces using the crosslinker EDC or incorporated into AA sequences during peptide synthesis.

Proteases and Protein-Cleaving Reagents

Use of purified and agarose-immobilized proteases for enzymatic proteolysis (cleavage or digestion) of proteins to facilitate amino acid sequencing, peptide analysis and polypeptide structural characterization.

Immobilized Ficin

Immobilized ficin is used primarily to generate Fab and F(ab’)2 fragments from mouse IgG1 antibodies, and immobilization virtually eliminates autolysis of the enzyme and allows tight control of the digestion by removing the ficin at any time.

Immobilized Papain

Immobilized papain is used primarily to generate Fab and Fc fragments from antibodies, and immobilization virtually eliminates autolysis and protease contamination of the sample and allows tight control of the digestion by removing the papain at any time.

Immobilized Pepsin

Immobilized pepsin is used primarily to generate F(ab’)2 fragments from antibodies, and immobilization virtually eliminates autolysis and protease contamination of the sample and allows tight control of the digestion by removing the pepsin at any time.

Protein Denaturants and Chaotropes

Chaotropic and denaturing chemical agents, including urea and guanidine hydrochloride, to disrupt water interactions and promote hydrophobic protein and peptide solubilization, elution, refolding and structural analysis.

Guanidine-HCl and Solution

Particulate free, crystal clear, colorless solution of guanidine hydrochloride with excellent stability. Can be used for washing affinity ligand columns (non-protein ligands), solubilizing inclusion bodies, and other peptide/protein analysis methods.

Urea

A low UV absorbing protein denaturant.

Reducing Agents for Protein Disulfides

Purified powders, convenient solutions and solid-phase resins of disulfide reducing agents, including DTT, BME and TCEP, for stabilization of free sulfhydryls (cysteines) and reduction of disulfide bonds in peptides and proteins.

2-Mercaptoethanol (BME)

2-Mercaptoethanol (B-mercaptoethanol) is a mild reducing agent that is ideal for cleaving disulfide bonds and generating thiols.

2-Mercaptoethylamine-HCl

2-MEA, an reducing agent to dissociate divalent IgG to monovalent IgG by reducing disulfide bonds in the hinge region without separating heavy and light chains.

METHODS OF DETERMINING PROTEIN CONFORMATION

X-ray crystallography

This is the X-ray diffraction pattern. X rays are allowed to strike the protein crystal; the X rays are diffracted by the crystal and impinge on a photographic plate, forming a pattern of spots.

The measured intensity of the diffraction pattern, as recorded on a photographic film, depends particularly on the electron density of the atoms in the protein crystal.

This density is lowest in H atoms, and they do not give a visible diffraction pattern.

C, O and N atoms yield visible diffraction patterns but are present in great number - about 700 or 800 per 100 amino acids. This makes resolution of the structure of a protein containing more than 100 amino acids difficult.

X-ray crystallography is best for structures of rigid proteins that form nice, ordered crystals. Flexible proteins are difficult to study by this method because flexible portions of protein will often be invisible in crystallographic electron density maps, since their electron density will be smeared over a large space.

Resolution is improved by substituting into the side chains of certain amino acids very heavy atoms, (heavy metals) e.g Mercury ions, Platinum chloride.

Measures of the accuracy of a crystallographic structure include resolution, which measures the amount of detail that may be seen in the experimental data, and the R-value, which measures how well the atomic model is supported by the experimental data found in the structure factor file.

The X-ray diffraction technique cannot resolve the complete three-dimensional conformation. Complete resolution is obtained by combination of the X-ray diffraction results with the amino acid sequence analysis. In example proteins; myoglobin, chymotrypsinogen, lysozyme, and ribonuclease.

NMR Spectroscopy

Also Spectrophotometry. Involves the measurement of the degree of absorbance of light by a protein in solution within a specified wavelength.

NMR spectroscopy provides information on proteins in solution, as opposed to those locked in a crystal or bound to a microscope grid, and thus, NMR spectroscopy is the premier method for studying the atomic structures of flexible proteins

It’s useful within the range of visible light only with proteins that contain coloured prosthetic groups (the nonprotein components). Examples of such proteins include the red heme proteins of the blood, the purple pigments of the retina of the eye, green and yellow proteins that contain bile pigments, blue copper-containing proteins, and dark brown proteins called melanins.

Computational methods are used for determining protein structures from NMR data.

Electron Microscopy

A beam of electrons is used to image the molecule directly. Several tricks are used to obtain 3D images. If the proteins can be coaxed into forming small crystals or if they pack symmetrically in a membrane, electron diffraction can be used to generate a 3D density map, using methods similar to X-ray diffraction.

If the molecule is very symmetrical, such as in virus capsids, many separate images may be taken, providing a number of different views. These views are then aligned and averaged to extract 3D information.

Electron tomography, on the other hand, obtains many views by rotating a single specimen and taking several electron micrographs. These views are then processed to give the 3D information.

Electron micrographic experiments may not allow the researcher to see each atom.

Electron micrographic studies often combine information from X-ray crystallography or NMR spectroscopy to sort out the atomic details.

Optical activity

Amino acids, except glycine, exhibit optical activity (rotation of the plane of polarized light). Proteins also are optically active. They are usually levorotatory (i.e., they rotate the plane of polarization to the left) when polarized light of wavelengths in the visible range is used.

Although the optical rotation of a protein depends on all of the amino acids of which it is composed, the most important ones are C and the aromatic amino acids F, Y, and W. The contribution of the other amino acids to the optical activity of a protein is negligible.