This study was conducted to separate and visualize the protein molecules using sodium dodecyl sulfate polyacrylamide gel electrophoresis and to estimate their molecular size using gel analysis. According to Okada et al. (2011), SDS-PAGE is one of the most effective techniques used for the separation of proteins (Okada et al., 2011). In 1967, the utility of SDS-PAGE for protein molecular weight (Mw) determination was initially demonstrated by Shapiro and coworkers, who showed that a straight line could be fitted to plots of gel migration distance against the logarithm of Mw for 11 proteins (Shapiro et al., 1967).
In order to understand the strategy underlying SDS-PAGE, It is essential to know about protein structure and its organization explained in Fig1.
Fig1: The four different structures of protein
SDS is an anionic detergent with a net negative charge. It binds to most soluble protein molecules in aqueous solutions over a wide pH range. The amount of bound SDS is proportional to the size of the molecules. The SDS eliminates most of the complex secondary, tertiary or quaternary structure of proteins, which is one requirement for protein sizing by SDS-PAGE. Furthermore, it is usually necessary to reduce protein disulphide bridges before the proteins adopt the random-coil configuration necessary for separation by size. This is achieved with reducing agents such as 2-mercaptoethanol. In addition, SDS also confers a negative charge that is utilized to separate the protein in an electrical field within polyacrylamide gels. The polyacrylamide forms porous gels allowing the separation of the molecules by size (Goetz et al., 2004). The polyacrylamide gel involves an upper stacking gel and a lower resolving gel of different pH. The stacking gel has larger pores and a pH of 6.8. This will lead the protein sample to accumulate in the stacking gel then its migration to the resolving gel, which has lower pore sizes and a pH of 8.8. A polyacrylamide gel with a certain acrylamide concentration restrains larger molecules from migrating as fast as smaller molecules (Goetz et al., 2004).
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For efficient resolution by SDS-PAGE, protein molecules must be fully denatured by boiling them at 100ËšC with sample buffer, which contains the strongly anionic detergent SDS along with the reducing agent 2-mercaptoethanol (which reduces all disulfide bonds present). Both of these agents disrupt protein secondary, tertiary and quaternary structures to produce linear chains, and SDS binds to hydrophobic regions of these chains proportionally to the molecular size of the proteins regardless of differences in their amino acid sequence. So, all denatured proteins act as negatively charged polymers in the presence of SDS. Sample buffer also contains glycerol to prevent diffusion of protein sample when loaded into the gel well, and an anionic dye as Bromophenol blue to allow progress of bands migrating across the gel to be monitored.
When the proteins denatured with SDS are subjected to electrophoresis along with the polyacrylamide gel, they are separated according to their molecular sizes (Okada et al., 2011). Thus, an appropriate standard sample proteins of Mw markers was prepared with Mw range between 6.500 Da and 200,000 Da. A stock solution of unknown protein (A) was also provided. Then three tubes were prepared according to table 1.
Table 1: The formulation of protein samples for SDS-PAGE resolution
After centrifugation of the three tubes prepared along with Mw markers tubes, the samples were placed in boiling water for five minutes to allow denaturation and association of linearized protein chains with SDS present in the sample buffer.
The samples were briefly centrifuged before being loaded onto the acrylamide gel with a current of 40 mA. When the dye front has traversed the gels, the gel is removed and placed in Coomasie blue stain to bind to proteins facilitating their visualization. The gel was destained after that to visualize the blue bands for later photographs (Fig2).
Fig2: The Photograph result of the Electrophoresed proteins.
A plot of log MW versus Rf (Calibration curve in the next page) was generated from the bands in the gel shown in Fig2 to determine the MW of the unknown protein.
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Hames (1998) defines the Rf as the migration distance of the protein through the gel divided by the migration distance of the dye front that is equal to 85mm in this research. The distance should be measured from the top of the resolving gel to the band of interest (Hames, 1998).
Table2. SDS-PAGE gel analysis data
Molecular Weight (Da)
Distance travelled by protein (mm)
Myosin from porcine heart
β-Galactosidase from E. coli
Phosphorylase b from rabbit muscle
Albumin, bovine serum
Glutamic Dehydrogenase from bovine liver
Ovalbumin from chicken egg
Glyceraldehyde-3-phosphate Dehygrogenase from rabbit muscle
Carbonic Anhydrase from bovine erythrocytes
Trypsinogen Inhibitor bovine pancreas
Trypsin Inhibitor from soybean
α-Lactalbumin from bovine milk
Aprotinin from bovine lung
In order to estimate the molecular weight of protein A, the distance migrated by this unknown protein is measured. It can be seen in Fig2 the protein A has three bands that have migration distances of 11mm, 16mm and 27mm respectively from up to down. In a gel of uniform pore size, the relative migration distance of a protein (Rf) is negatively proportional to the logarithm of its Mw (Goetz et al., 2004) and the curve generated should be linear.
The linear function is considered as: Y= a X +b ; where Y is Log Mw and X is Rf
The slope a = Change in Y/ Change in X
So a = 0.226 Therefore b = 5.27
So the final equation of the linear curve is Y = 0.226 X + 5.27
From this equation, Y is calculated (see table3).
Distance travelled by protein (mm)
Log Mw from the equation of the linear curve
Table3. Calculation Data of the three bands of Protein A
Therefore the three bands of Protein A have molecular weight of 200000Da, 205116Da, and 219583Da respectively.
In conclusion, the ability of SDS gel electrophoresis to dissociate almost any protein complex and to resolve individual protein chains by size has proven applicable to a wide range of biochemical complications, and the method is crucial for any lab studying proteins (Maizel et al., 2000).