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Whey protein represents approximately 20% of the original milk proteins. It exhibits many important functional properties in the manufacture of food. The nutritional values and physiological benefits of whey protein receive a lot of attention. Although whey proteins have versatile functional properties and nutritional values, an estimated 70% of liquid whey is disposed as waste product. Such large volumes of disposed protein will seriously impact environmental pollution due to its high BOD (biological oxygen demand) level. The utilization of whey proteins is confined because there still exists a large variability in the composition and functional properties of commercial whey proteins.To date, the factors responsible for the variability remain poorly understood. It is possible that the incongruence of physico-chemical and functional performance of whey protein reported from different groups might result from the compositional variations arising from different whey resources or processing conditions in commercial whey products. To clarify this problem, how to obtain whey protein in its native form, free from any denaturizing, becomes of utmost importance. Many processes such as ultrafiltration, diafiltration, polyphosphate complex precipitation, heat coagulation, and ion-exchange adsorption technology have been developed for the manufacture of whey protein. By these methods, whey protein is prone to the risk of protein denaturation. For protein separation, a number of liquid chromatography methods including reversed-phase, hydrophobic interaction, and ion exchange and techniques have been employed. For the ion-exchange method, a large amount of mobile phase is usually prepared with various salts at different concentrations or at different pH to wash out the compound of interest, and for reverse-phase HPLC, some organic solvents or an acidic mobile phase have to be applied, all of which potentially denature proteins. In contrast, size exclusion chromatography is relatively amiable to protein molecules and has been frequently used in protein separation and. Yoshida used a Sephacryl S-200 column to isolated Î²-Lg and Î±-La from acid whey protein; Nakai and Al-Mashikhi isolated whey proteins using Sephacryl S-300 and TSK HW-55 columns. No single method is suitable for all whey proteins and the method of choice is usually based on one or two whey components of interest. Moreover, most previous whey-related studies are sampling from commercial products such as whey concentrates or acid whey proteins, of which properties are closely dependent on the manufacture processes.
In this work, different whey protein powder and whey protein-GOS conjugate were analyze through a simple and economical separating process using polyacrylamide gel electrophoresis.It shows that four major components (immunoglobulin, bovine serum albumin, Î²-lactoglobulin, and Î±-lactalbumin) and a small non-protein molecule can be easily collected. Protein and sugar will be analyzes to see where the GOS binds.     
Polyacrylamide gel electrophoresis was originally associated with « disc gel electrophoresis », because the system was promoted in which samples are run in individual tubes of gel, resolving into "disc" of protein zones. Although still widely used, disc gel electrophoresis is being replaced by slab gels, on which several samples can be run simultaneously with direct comparison of mobility. Thin-slab gel electrophoresis, using polyacryalmide, has been developed extensively over the past few years, and the technical problems associated with pouring and with sample application have been resolved, so that the system is as simple as, but more powerful than disc gels. A variety of commercial apparatuses are available, using gel thicknesses down to less than 1 mm. one advantage of a thin gel is that the less heat is produced per square centimeter of gel surface for a given applied voltage. Also, during staining and destaining of the protein bands diffusion of dye is more rapid into the thin gel slab.   
This involves running the sample in a buffer at a pH where the proteins remain stable and in their native form. This method was the original procedure, marking one both of differences in charges between proteins and their different sizes. The buffer chosen depends somewhat on the nature of the proteins, but generally it is slightly alkaline, in the pH range 8-9, where most proteins are negatively charged and so move toward the anode. The anode is normally at the bottom of the gel; it should be noted that there is no provision for proteins that move in the other direction in these systems. Basic proteins disappear into the cathode buffer. Of the bulk of the proteins being observed are known to be basic, then o buffer of somewhat lower pH can be employed, and the system operated with the cathode at the bottom of the gel.
The concentration of the polyacrylamide can be varied over a wide range according to the size of molecules being separated. Two variations are possible: variation in total acrylamide content and variation in cross-linker percentage (N,N'-methylene bisacrylamide). Each achieves roughly the same objective; an increase in either reduces the pore size and so slows up larger molecules more. For every large proteins, up to 106 daltons, an open, rather difficult-to-handle gel about 3-4% acrylamide and 0.1% bisacrylamidecan be made. For very small proteins around 104 daltons, 15% acrylamide and/or up to 1%bisacrylamide may be used. A high percentage of cross linker tends to make the gels more opaque; 1 part of cross linker to between 20 and 50 of monomer is the normal range. Usually gels of 7-10% acrylamide are used. Polylmerizationin initiated with freshly dissolved ammonium persulfate (1.5-2 mM) together with a free radical scavenger, TEMED (0.05-0.01% v/v, N,N,N',N'-tetramethylenediamine). Alternatively, photopolymerization using riboflavin may be used; it leaves less harmful residue in the gel. Gelation should occur within 30 minute at room temperature. Further details of suitable buffers and other parts of the system are available from apparatus manufactures.
Simple electrophoresis can also be carried out in starch gel as the medium. The quality of the starch and its behavior in forming gels can be rather variable, but before polyacrylamide systems were widely used, starch was very popular. It is still particularly useful when detecting enzymic activity after electrophoresis; a starch gel slab can easily be sliced and stained on one-half for protein, the other half for enzyme activity. Because starch gels are opaque, protein staining is a surface stain, so a larger protein amount is needed than with transparent gels in which the whole depth of the protein zones is stained.
Simple electrophoresis in agarose gels does not have the resoving power of smaller-pore gels, so an analysis of purity based on homogeneity in such an electrophoretic system is less convincing.     
This method can be used for proteins insoluble at low ionic strength; it is presently the most commonly used method for all types of proteins. It involves denaturing the protein with the detergent sodium dodecyl sulfate. Commonly known as PAGE-SDS (polyacrylamide gel electrophoresis in sodium dodecyl sulfate), this high resolution method has won most protein chemists aver, partly because one can describe the bands not just in terms of their relative mobility to each other, but also in terms of their molecular size, for PAGE-SDS separates polypeptides chains according to size. Dodecyl sulfate binds stroungly to proteins, so that only 0.1% dodecyl sulfate is suffient to saturate the polypeptide chains, with approximately 1 detergent molecule per 2 amino acid residues. Each dodecyl sulfate carries a negative charge, so a typical polypeptide of molecular weight of 40,000 acquires about 180 negative charges ; far in excess of any net charge that might exist (at neutral pH) on the polypepetide chain originally. Consequently the charge/size ratio is virtually identical for all proteins, and separation can occur only as a result of the molecular sieving through the pore of the gel. Despite the fact that the potential of separation of proteins of identical size is not possible in this system, it nevertheless appears to give the sharpest overall resolution and cleanest zones of any method. What is more, by comparison with a mixture of standard polypeptides of known molecular weight, the whole gel can be calibrated in terms of mobility against size. It is found that linear plot over a substantial range can be obtained if mobility is plotted against molecular weight.
For further dissociation incorporation of a reagent is possible such as 2_mercaptoethanol, capable of reducing disulfide bonds.   
This method is also widely used; it resembles the sodium dodecyl sulfate method in that it separates protein according to size only, not charge differences. A slab of polyacrylamide is poured in which the acrylamide concentration varies from high value at the bottom (usually about 30%) to only 3% at the top. The buffer is a high-pH one, so that the most proteins migrate into the gel toward the anode at the bottom. Electrophoresis is continued until all proteins have reached a thickness of gel which prevents them from moving any further; the small molecules may reach 25% acrylamide, large ones remaining near the top. As with the dodecyl sulfate system, molecular size (this time of the negative protein, not subunits) can be determined by comparison with standard mixture. The gradient gel system run until no further movement occurs.  
Staining and destaining
After electrophoresis protein zone must be visualized, and this is carried out by a staining procedure, normally involving an organic dye which binds tightly to protein. Many different dyes can be used, and the main objectives are sensitivity of detecting small amounts of protein and proportional staining with all types of protein. Most dyes used tend to be attracted by positively charged group on the proteins; consequently proteins with higher proportions of these generally more basic proteins tend to stain more strongly. Indeed, some acidic polypeptides have escaped detection because they bind so little dye. After the stain protein are destaining to visualize the bands correctly. 
Prebiotic GOS (galacto-oligosaccharides)
Galacto-oligosaccharides (GOS), also known as oligogalactosyllactose, oligogalactose, oligolactose or transgalactooligosacchariden (TOS), belong, because of their indigestible nature, to the group of prebiotics. Prebiotics are defined as non-digestible food ingredients that beneficially affect the host by stimulating the growth and/or activity of beneficial bacteria in the colon. GOS naturally occurs in human milk, and commercial available products are broadly used in food for both infants and adult, ranging from infant formula to biscuits to food for the critical ill. They are widely recognized as the non-digestible carbohydrates that occur naturally in human milk. Moreover they can be manufactured from lactose through enzymatic conversion
The composition of the galacto-oligosaccharide fraction varies in chain length and type of linkage between the monomer units. Galacto-oligosaccharides are produced through the enzymatic conversion of lactose, a component of milk. They occur naturally in human milk.   
GOS generally comprise a chain of galactose units that arise through consecutive transgalactosylation reactions, with a glucose unit terminal. However, where a terminal galactose unit is indicated, hydrolysis of GOS formed at an earlier stage in the process has occurred. The composition of GOS can vary quite markedly in terms of degrees of polymerization ranging from 2 to 8 monomeric units. Galacto-oligosaccharides are marketed commercially by FrieslandCampina as Vivinal GOS.  [
Glycation of whey protein with the prebiotic GOS through the maillard reaction under controlled conditions can be successfully used to improve or produce new functional and biological properties. 
Concentration of protein solution and dialysis are closely related. Conventional use of dialysis bags involves the removal of unwanted low-molecular weight solute from the sample and replacement with buffer present in the "dialysate". Osmotic forces are usually the opposite to that describe above: high concentrations of salt leaves; consequently there is an increase in volume during the early stages if the solute concentration is high. Dialysis is used both for removing excess low-molecular weight solute and simultaneously introducing a new buffer solution (it may be just water) to the sample. Dialysis tubing is available in variety of size, and conveniently does not allow molecule larger than 15,000-20,000 daltons to pass through. All low molecular-weight molecules diffuse through the tubing, and eventually the buffer composition on each size equalizes. Complete removal of endogenous salts from a sample cannot occur in on dialysis simply because at equilibrium what was originally inside the dialysis bag now is distributed throughout the buffer and the dialysis bag. If the buffer volume is 50 times that of the bag, then the best that can be achieved is a 51-fold dilution of the original salts/low-molecular-weight material. Consequently, changing the buffer and movement of the dialysis bag should occur. A well mix system can reach more than 90% equilibration in 2-3 hours. A large volume of buffer is preferable since fewer buffer changes are needed. Dialysis is a convenient step to carried out overnight as needs no attention, and equilibrium is reached by morning. However, the possibility of proteolytic degradation during dialysis may make its use undesirable.  
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Figure 1: Process of dialysis
this graphic illustrates the dialysis process. First, the concentrated protein solution is placed in dialysis bag with small holes which allow water and salt to pass out of the bag while protein is retained. Next the dialysis bag is placed in a large volume of buffer and stirred for many hours (16 to 24 hours), which allows the solution inside the bag to equilibrate with the solution outside the bag with respect to salt concentration. When this process of equilibration is repeated several times (replacing the external solution with low salt solution each time), the protein solution in the bag will reach a low salt concentration. (Figure 2) 
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Figure 2: Separation of protein from salts by semi permeable membrane.
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