DETERMINATION OF PURITY AND RELATIVE MOLECULAR WEIGHT OF PURIFIED HUMAN IgG

About Authors:
¹M Prasad Naidu, ²Dr Madhu Sudan Reddy, ³T Madhu Chaithanya, ⁴N Mallikarjun Rao
¹(Medical Biochemistry) NMCH, Nellore, AP, India.
²(MD Pharmacolgy) NMCH, Nellore, AP, India.
³(Medical Pharmacology) MIMS, Vijayanagaram, AP, India.
⁴(MSc Botany). Acharya Nagarjuna University, Guntur, AP, India.
*www.prasadnaidu.com@gmail.com

Sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE)
An important technique for the separation of proteins is based on the migration of charged proteins in an electric field, a process called electrophoresis. These procedures are not generally used to purify proteins in large amounts, because simpler alternatives are usually available and electrophoretic methods often adversely affect the structure and thus the function of proteins. Electrophoresis is, however, especially useful as an analytical method. Its advantage is that proteins can be visualized as well as separated, permitting a researcher to estimate quickly the number of different proteins in a mixture or the degree of purity of a particular protein preparation. Also, electrophoresis allows determination of crucial properties of a protein such as its isoelectric point and approximate molecular weight. The polyacrylamide gel acts as a molecular sieve, slowing the migration of proteins approximately in proportion to their charge-to-mass ratio. Migration may also be affected by protein shape. In electrophoresis, the force moving the macromolecule is the electrical potential, E. The electrophoretic mobility of the molecule, µ, is the ratio of the velocity of the particle molecule, V, to the electrical potential. Electrophoretic mobility is also equal to the net charge of the molecule, Z, divided by the frictional coefficient, f, which reflects in part a protein’s shape.

REFERENCE ID: PHARMATUTOR-ART-1854

Thus: Mu = V/E = z/f

The migration of a protein in a gel during electrophoresis is therefore a function of its size and its shape.
An electrophoretic method commonly employed for estimation of purity and molecular weight makes use of the detergent sodium dodecyl sulfate (SDS).

SDS binds to most proteins in amounts roughly proportional to the molecular weight of the protein, about one molecule of SDS for every two amino acid residues. The bound SDS contributes a large net negative charge, rendering the intrinsic charge of the protein insignificant and conferring on each protein a similar charge-to-mass ratio. In addition, the native conformation of a protein is altered when SDS is bound, and most proteins assume a similar shape. Electrophoresis in the presence of SDS therefore separates proteins almost exclusively on the basis of mass (molecular weight), with smaller polypeptides migrating more rapidly. After electrophoresis, the proteins are visualized by adding a dye such as Coomassie blue, which binds to proteins but not to the gel itself. Thus, a researcher can monitor the progress of a protein purification procedure as the number of protein bands visible on the gel decreases after each new fractionation step. When compared with the positions to which proteins of known molecular weight migrate in the gel, the position of an unidentified protein can provide an excellent measure of its molecular weight. If the protein has two or more different subunits, the subunits will generally be separated by the SDS treatment and a separate band will appear for each.

The pioneering work on electrophoresis by A. Tiselius and co-workers was performed in free solution. However, it was soon realized that many of the problems associated with this approach, particularly the adverse effects of diffusion and convection-currents, could be minimized by stabilizing the medium. This was achieved by carrying out electrophoresis on a porous mechanical support, which was wetted in electrophoresis buffer and in which electrophoresis of buffer ions and samples could occur. The support medium cuts down convection currents and diffusion so that the separated components remain as sharp zone. The earliest supports used were filter paper or cellulose acetate strip, wetted in electrophoresis buffer. Nowadays these media are infrequently used; although cellulose acetate still has its use in clinical laboratories for the separation of serum proteins. In particular, for many years’ small molecules such as amino acids, peptides and carbohydrates were routinely separated and analyzed by electrophoresis on supports such as paper or thin-layer plates of cellulose, silica or alumina. Although occasionally still used, such molecules are nowadays more likely to be analyzed by more modern and sensitive techniques. While paper and or thin-layer supports are fine for resolving small molecules, the separation of macromolecules such as proteins and nucleic acids on such supports is poor.

However, the introduction of the use of gels as support medium led to a rapid improvement in methods for analyzing macromolecules. The earliest gel system to be used was the starch gel and although this still has some uses, the vast majority of electrophoretic techniques used nowadays involve either polyacrylamide gels or agarose gels. In the case of paper and cellulose acetate electrophoresis, charge on the molecule is the major determinant for its electrophoretic mobility and ultimate separation. The gels in common use, polyacrylamide and agarose, have pores of molecular dimensions whose sizes can be specified. The molecular separations are therefore based on gel filtration as well as the electrophoretic mobilities of the molecules being separated.

Components of SDS-PAGE
Acrylamide:
The chemical compound acrylamide (acrylic amide) has the chemical formulaC₃H₅NO. Its systematic name is 2-propenamide. It is a white odorless crystalline solid, soluble in water, ethanol, ether and chloroform. Most acrylamide is used to synthesize polyacrylamidewhich find many uses as water-soluble thickeners. These include use in wastewater treatment, gel electrophoresis(SDS-PAGE), papermaking, ore processing, and the manufacture of permanent press fabrics. Some acrylamide is used in the manufacture of dyes and the manufacture of other monomers.

Polyacrylamide:
It is an acrylate polymer formed from acrylamide subunits that is readily cross-linked. Acrylamide needs to be handled using Good Laboratory Practices (GLP) to avoid poisoning since it is a neurotoxin. Polyacrylamide is not toxic, but un-polymerized acrylamide can be present in the polymerized acrylamide

Sodium dodecyl sulphate: (CH₃ – (CH₂)₁₀ –CH₂-O-SO₃^-) Na⁺
Sodium dodecyl sulphate is an anionic detergent and disrupts macromolecules whose structure has been stabilized by hydrophobic associations. SDS has been shown to hydrophobically associate with the non-polar residues of a protein there by interfering with the hydrophobic interactions responsible for the protein native structure. The molecular masses of “normal” proteins can be determined to an accuracy of 5-10 % through SDS-PAGE.

Ammonium per sulphate:
It provides the free radicals that drive polymerization of Acrylamide and bisacrylamide. The polymerization is induced by free radicals resulting from the chemical decomposition of ammonium per sulphate or the photo decomposition of riboflavin.

TEMED:
It is a free radical stabilizer and accelerates the polymerization of acrylamide and bisacrylamide by catalyzing the decomposition of persulphate ion to give a free radical (i.e., a molecule with an unpaired electron).

Glycerol:
It provides high density for the samples prepared in order to prevent its mixing with the upper reservoir buffer when samples were loaded on top of the loading wells.

Bromophenol blue (Tracking dye):
The extent of migration of the dye gives an index of electrophoretic process. The dye migrates faster than all the macromolecules. Thus if the electrophoresis is stopped before or just as the dye comes out of the bottom of the gel, one can be reasonably sure that all macromolecules are still within the gel.

β-mercaptoethanol:
β-mercaptoethanol reduces any disulphide bridges present that are holding together the proteins tertiary structure. Reductive cleavage by treatment with β–Mercaptoethanol in order to expose all disulphide bonds (groups) to the reducing agent, the reaction is usually carried out under conditions that denature the protein (disrupt its native conformation).

Buffer solutions:

0.5M Tris HCl pH 6.8
Buffer pH is about 2 units lower than the electrode buffer in the upper reservoir, which shifts the equilibrium towards formation of zwitter ions. As zwitter ions do not possess a net charge, they are immobile and carry the current in this region and migrate rapidly in this strong local electric field the pH of the buffer used in the sample is the same buffer that is used in the stacking gel.

1.5 M Tris HCl pH 8.8
The faster migration of the proteins from stacking gel results in piling/stacking up of the protein sample in a tight, sharp band between the glycinate and the chloride ions.. From this point onwards the separation of protein takes place.

Electrode buffer (Reservoir buffer):
It is also an amine (Tris glycine) which is same in both upper and lower reservoirs and gel has pH usually ~9 for proteins) so it provides the proteins net negative charges. The anode, to which these negatively charged macromolecules would migrate under an electric field, is therefore placed in the lower buffer reservoir.

Sample buffer:
Samples to be run on SDS-PAGE are firstly boiled for 5min in sample buffer containing β-mercaptoethanol and SDS. The original native charge on the molecule is therefore completely masked by the negatively charged SDS molecules. The rod-like structure of protein remains because as any rotation that tends to fold up the protein chain would result in repulsion between negative charges on different parts of the protein chain, returning the conformation back to the rod shape.

The support media (polyacrylamide gels):
The components used in the formation of these gels are known to be neurotoxins and thus care has to be taken while preparing the gel. The most commonly used components to synthesize the matrix are acrylamide monomer, N', N', methyl bisacrylamide, ammonium persulphate and tetra ethylene diamine (TEMED).

Acrylamide gels are defined in terms of the total percentage of acrylamide present, and pore size in the gel can be varied by changing the concentrations of both the acrylamide and bis-acrylamide. Acrylamide gels can be made with a content of between 3% - 30% acrylamide. Formulation for separating gels depending upon the molecular weights of macromolecules is given in.Thus low percentage gels (e.g. 4%) have large pore sizes and are used. In the electrophoresis of proteins, where free movement of the proteins by electrophoresis is required without any noticeable frictional effect, for example, in flat-bed isoelectric focusing or the stacking gel system of an SDS- polyacrylamide gel. Low percentage acrylamide gels are used to separate DNA. Usually 10 - 20% acrylamide are used in techniques such as SDS-gel electrophoresis, where the smaller pore size, now introduce a sieving effect that contributes separation of proteins according to their size.Electrophoresis of proteins is generally carried out in gels made up of the cross-linked polymer polyacrylamide.

Two different porosity gels commonly used in SDS-PAGE are the stacking gel (high porosity gel) and separating or resolving gel (low porosity gel).

Separating gel
The lower separating or the resolving gel is prepared using 5-15 % acrylamide which is much higher than that used in the upper stacking gel (the percentage of acrylamide used depends upon the approximate molecular weight of the macromolecule being separated. Consequently the pores are numerous and of a smaller diameter imparting molecular sieving property to this gel. The gels in gel electrophoresis, however, retard large molecules relative to smaller ones, the reverse of what occurs in gel filtration chromatography. Because there is no solvent space in gel electrophoresis to that between the gel beads in gel filtration chromatography (electrophoresis gels are directly cast in the electrophoresis device). The movement of the large molecules in gel electrophoresis is therefore impeded relative to that of the smaller molecules as the molecules migrate through the gels i.e. it is in this gel that the macro molecules separate. The buffer used in this gel is usually an amine such as tris, which is adjusted to a pH of 8.8 using HCl. The separating gel constitutes about two-thirds of the length of the gel plate.

Stacking gel
After the separating gel had been polymerized, a second layer of gel referred to as stacking or spacer gel is casted over the separating gel. Usually 4-6% acrylamide and is consequently highly porous and devoid of any molecular sieving action. The buffer used here is also an amine, mostly tris whose pH is adjusted to 6.8 with HCl. The pH of this gel is about two pH units lower than that of the separating gel (pH 8.8). The buffer used in the sample buffer is identical to that used in the stacking gel gels are prepared.

Electrophoretic run:
Once the samples are loaded, power is switched on. When the power is switched on, a diligent observer will see that the amount of bubbles generated in the reservoir containing the anode is much less than that in the reservoir containing the cathode. The reason for this lies in the reactions that permit the flow of current from cathode to anode. They are essentially the electrolysis of water producing hydrogen at the cathode and oxygen at the anode. However, for each mole of hydrogen produced at the cathode only one-half mole of oxygen is produced at the anode leading to less number of bubbles being seen.

Electrophoretic process:
The samples to be separated are not loaded directly into the main separating gel. When the main separating gel (normally about 10cm long) has been poured between the glass plates and allowed to set, a shorter (approximately lcm) stacking gel is poured on top of the separating gel and it is into this gel that the wells are formed and proteins loaded. The purpose of this stacking gel is to concentrate the protein sample into a sharp band before it enters the main separating gel. This is achieved by utilizing differences in ionic strength and pH between the electrophoresis buffer and the stacking gel, and involves a phenomenon known as isotachophoresis. The stacking gel has a very large pore size (4% acrylamide), which allows the proteins to move freely and concentrate, or stack under the influence of the electric field. The band-sharpening effect relies on the fact that negatively charged glycinate ions (in the electrophoretic buffer) have a lower electrophoretic mobility than do the protein-SDS complex, which, in turn have lower mobility than the chloride ions (Cl^-) of the loading buffer and the stacking gel.

When the current is switched on, all the ionic species have to migrate at the same speed otherwise there would be a break in the electrical circuit. Glycine in the upper buffer reservoir exists in two forms; as zwitterions, which does not have a net charge, and as a glycinate anion with a charge of minus one.

⁺NH₃CH₂COO^- (Zwitter Ion) ---> NH₂ (Negative Ion)

When the power is switched on, chloride, protein and glycinate ions begin to migrate towards the anode. Upon entering the stacking gel, the glycinate ion encounters a condition of low pH, which shifts the equilibrium towards formation of zwitter ions. As zwitter ions do not possess a net charge, they are immobile. This immobility of glycine zwitter ions to migrate in to the stacking gel coupled with high mobility of the chloride ions creates a very high localized voltage gradient between the leading chloride and the trailing glycinate ions. Since proteins have their mobility intermediate between the trailing and the leading ions, they carry the current in this region and migrate rapidly in this strong local electric field. The proteins, however, cannot overtake the chloride ions, as the strong local field exists only between the chloride and the glycinate ions. As a result the proteins migrate quickly until they reach the region rich in chloride ions and then drastically slow down. The result is that the three species of interest adjust their concentrations so that [Cl]> [protein-SDS]> [glycinate]. There is only a small quantity of protein-SDS complexes, so they concentrate in a very tight, sharp band between glycinate and Cl^- boundaries. (i.e., the faster migration of proteins results in piling or stacking up of the protein samples into a tight sharp disc). It is in this form that the macromolecules enter the running gel. The smaller pores of the separating gel retards the movement of the sharp band of the macromolecules for a long enough time for the glycinate anions to catch up. (The larger pores in the stacking gel do not have any sieving effect therefore the macromolecules migrate faster without any hindrance however, when they reach the separating gel the pores are numerous and of a smaller diameter imparting molecular sieving property to the gel therefore the macromolecules cannot migrate with the same speed as they did in the stacking gel as a result the proteins pile or stack up into a tight sharp disc).

Once the glycinate ions reach the separating gel it becomes more fully ionized in the higher pH environment its mobility increases. (The pH of the stacking gel is 6.8 that of the separating gel is 8.8). Thus the interface between glycinate and Cl^- leaves behind the protein-SDS complexes which are left to electrophoresis at their own rates. (Upon entering the separating gel the glycinate ions encounter a condition of high pH [pH of the separating gel buffer is about 2 pH units higher than that of the stacking gel] which shifts the equilibrium towards formation of glycinate anion). The negatively charged protein-SDS complexes now continue to move towards the anode under the applied field with the same mobility. However as they pass through the separating gel the proteins separate, owing to the molecular sieving properties of the gel. Quite simply, the smaller the proteins the more easily it can pass through the pores of the gel, whereas larger proteins are successively retarded by frictional resistance due to the sieving effect of the gels. Being a small molecule, the bromophenol blue dye is totally unretarted and therefore indicates the electrophoresis front, i.e. the extent of migration of the dye gives an index of electrophoretic process. The dye migrates faster than all macromolecules. When the dye reaches the bottom of the gel, the current is turned off and the gel is removed from in-between the glass plates and shaken in an appropriate stain solution. A typical gel would take 1 to 1.5 h to prepare and set, 3h to run at 30mA. Vertical slab gels are invariably run, since this allows up to 20 different samples to be loaded on to a single gel.

Detection, estimation and recovery of proteins in gels:
Gels from a column are removed by forcing water from a hypodermic syringe around the walls of the column, allowing the gels to be extracted under gentle pressure. Slab gels are removed by introducing a thin metal plate (spatula) between the two gel plates and coaxing the plates apart. Before staining, the gels may be immersed in a fixative (5% TCA) to guard against diffusion of separated component (prevent diffusion). The most commonly used general protein stain for detecting proteins on gels is the sulphate trimethylamine dye Coomassie Brilliant Blue R-250 (CBB).

Coomassie Brilliant Blue R 250 (CBB):
Proteins are often visualized by staining with the dye Coomassie Brilliant Blue R 250. Staining is usually carried out using 0.25% (w/v) CBB in methanol: acetic acid: water (50:10:40, by volume). This acid-methanol mixture acts as a denaturant to precipitate or fix the protein in the gel, which prevents the protein from being washed out while it is being stained. Staining of most gels is accomplished in about 2h and destaining is achieved by gentle agitation in the same methanol-acid-water solution but in the absence of the dye (usually overnight). The Coomassie stain is highly sensitive; a very weakly staining band on a polyacrylamide gel would correspond to about 1µg of protein. The CBB stain is not used for staining cellulose acetate (or protein blots) because it binds to the paper. In this case, proteins are first denatured by brief immersion of the strip in 10% (v/v) trichloroacetic acid, and then immersed in a solution of a dye that does not stain the support material, for example: Procion blue, Amido black or Procion S. Although the Coomassie stain is highly sensitive, many workers require greater sensitivity and use a silver stain. Silver stains are based either on techniques developed for histology or on methods based on the photographic process. In either case, silver ions (Ag+) are reduced to metallic silver on the protein where silver is deposited to give a black or brown band. Silver stains can be used immediately after electrophoresis or alternatively, after staining with CBB. With the latter approach, the major bands on the gel can be identified with CBB and then minor bands not detected with CBB, identified using the silver stain. The silver stain is at least 100 times more sensitive than Coomassie Brilliant Blue, detecting protein down to 0.lµg amounts.

Results
Determination of purity and molecular weight of purified human IgG
To determine the purity of isolated IgG, fraction no.2 (serum samples 1, 2 and 3), were analyzed on a 12 % separating gel and 6 % stacking gel. Proteins from fraction no.2 serum samples 1, 2 and 3 resolved in to 2 bands. After determining the purity of isolated IgG, the molecular weight of isolated IgG was determined by running standard protein molecular weight markers along with fraction no.2 serum samples 1 and pure human IgG.

Two protein bands with molecular weights of 50 kD and 24 kD corresponding to heavy (50 kD) and light chain (25 kD) of immunoglobulin molecule were observed. These results clearly suggest that the IgG purified from human and serum was fairly pure.

Native–PAGE (Activity staining of the enzyme HRP)
While SDS–PAGE is the most frequently used gel system for studying proteins, the method is of no use if one is aiming to detect a particular protein (often an enzyme) on the basis of its biological activity, because the protein (enzyme) is denatured by the SDS–PAGE procedure. In this case it is necessary to use non-denaturing conditions. In native or buffer gels, polyacrylamide gels are again used (normally a 7.5% gel) but the SDS is absent and the proteins are not denatured prior to loading. Since all the proteins in the sample being analyzed carries their native charge at the pH of the gel (normally pH 8.7), proteins separate according to their different electrophoretic mobilities and the sieving effects of the gel. It is therefore not possible to predict the behavior of a given protein in a buffer gel but, because of the range of different charges and sizes of proteins in a given protein mixture, good resolution is achieved. The enzyme of interest can be identified by incubating the gel in an appropriate substrate solution such that a coloured product is produced at the site of the enzyme. An alternative method for enzyme detection is to include the substrate in an agarose gel that is poured over the acrylamide gel and allowed to set. Diffusion and interaction of enzyme and substrate between the two gels results in colour formation at the site of the enzyme. Often, duplicate samples will be run on a gel, the gel cut in half and one half stained for activity, the other for total protein. In this way the total protein content of the sample can be analyzed and the particular band corresponding to the enzyme identified by reference to the activity stain gel.

Result
The enzyme HRP was analyzed on a slab gels consisting of 12%separating gel and 6%stacking gel. After electrophoresis, the enzyme was identified by activity staining.

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