Proteins are biological macromolecules that perform in biological activity. The amino acid sequence primary structure of any protein is dictated by covalent bonds, but the higher levels of structure are formed and stabilized by weak, covalent interactions. Hydrogen bonds, hydrophobic interactions, electrostatic bonds and van der Waals forces are all noncovalent in nature which plays important roles in determining the structure of protein.
Hydrogen bonding are generally made wherever possible within a given protein structure. In most protein structures that have been examined to date, component atoms of the peptide backbone tend to form hydrogen bond with one another. They are bonded together to form a long chain of polypeptide which also play an important role in structure of secondary and tertiary structure of proteins. The peptide linkages, along with theÂ a-carbon atoms to which R-groups are attached, form theÂ protein backbone, with sequence NCCNCCNCCNCC...
Furthermore, there are intermolecular hydrogen bonds between protein and water molecules, and between water molecules, which are bound with the proteins, in addition to intramolecular hydrogen bonds.Â Side chains capable of forming H bonds are usually located on the protein surface and form such bonds either with the water solvent or with other surface residues. The strengths of hydrogen bonds depend to some extent on environment. The difference in energy between a side chain is usually quite small. On the other hand, a hydrogen bond in the protein interior, away from bulk solvent, can provide substantial stabilization energy to the protein.http://www.ncbi.nlm.nih.gov/books/NBK28438/bin/ch3f27.jpghttp://t1.gstatic.com/images?q=tbn:ANd9GcTgwrfnenPpJ6vLaurWTqLWQspRGn_rVqqFHDGxmU2ADFBi1yZnlQ
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Figure 3 shows the inter-molecular hydrogen between two amino groups.Figure 2 shows the intra-molecular
Ionic interactions arise either as electrostatic attractions between opposite charges or repulsions between like charges. Lysine, arginine and histidine carry positive charges and aspartate and glutamate carries negative charges. All of these may experience ionic interactions in protein structure.
Figure 4 shows an electrostatic interaction between the lysine and glutamate.
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Van der Waals force of attraction is the attractive and repulsive forces due to instantaneous dipole-induced dipole interactions arise due to fluctuations on the electron charge distributions of adjacent non bonded atoms. There are both attractive and repulsive van der Waals forces that control protein folding. Attractive van der Waals forces involve the interactions among induced dipoles that arise from fluctuations in the charge densities that occur between adjacent uncharged non-bonded atoms. Repulsive van der Waals forces involve the interactions that occur when uncharged non-bonded atoms come very close together but do not induce dipoles. It is formed between tightly packed group in the interior of protein are a major contribution to protein stability.
Has been the most widely used method to estimate the total level of protein (already in solution or easilyâ€soluble in dilute alkali) in biological samples, the Lowry Assay (Lowry et al., 1951) exhibited a color change of the biological samples in proportion to protein concentration. The color change can be then measured usingÂ colorimetricÂ techniques.Â
With sensitivity down to about 0.01 mg of protein/mL, the reaction is best used on solutions with protein concentrations ranged in 0.01-1.0 mg/mL. The Lowry assay is based on 2 reactions, namely Biuret reaction and the Folin-Ciocalteau reaction. Biuret reaction sees the peptide bonds of proteins, under alkaline reaction, reacts with copper to produce Cu+ ions which react with the Folin reagent. This is called a Biuret chromophore and is commonly stabilized by the addition of tartrate (Gornall et al., 1949).
Folin-Ciocalteau reaction is commonly poorly understood. However, in essence the second reaction is a reduction of the Folin-Ciocalteau reagent (phosphomolybdate and phosphotungstate), primarily by the reduced copperâ€amide bond complex, as well as by tyrosine and tryptophan residues. The reagent is reduced to heteropolymolybdenum blue.
The end product of the two reactions has a strong blue color, which in turn depends partially on the content of tyrosine and tryptophan. The amount of protein in the sample can then be estimated via reading the absorbance (at 750 nm) of the end product of the Folin reaction against a standard curve of a selected protein solution. An example would be Bovine Serum Albumin (BSA) solution.
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The Biuret reaction itself is not very sensitive. Lowry protein assay requires more time than other assays and is susceptible to many interfering compounds. While widely used, the Lowry procedure is less preferable an assay than some other protein assays since it is more subject to interference by a wide variety of chemicals. This is one ofÂ the major limitations of the assay as many of these interfering substances are commonly used in buffers for preparing proteins or in cell extracts. Known compounds to interfere with the Lowry assay are detergents, carbohydrates, glycerol, EDTA, potassium compounds, sulfhydryl compounds, disulfide compounds, most phenols, uric acid etcetera.
The Lowry assay is also sensitive to variations in the content of tyrosine and tryptophan residues. The assay is linear over the range of 1 to 100Î¼g protein. The absorbance can be readÂ in the region of 500 to 750nm, with 660 nm being the most commonly employed. Other wavelengths can also be used, but it may reduce the effects of contamination.
Using the Folinâ€Ciocalteu reagent to detect reduced copper makes the Lowry assay nearly 100 times more sensitive than the Biure treaction alone. Several useful modifications of the original Lowry assay have been developed to increase the dynamic range of the assay over a wider protein concentration (Hartree, 1972), to make the assay less sensitive to interference by detergents (Dulley and Grieve, 1975), and to first precipitate the proteins to remove interfering contaminants (Bensadoun and Weinstein,Â 1976).
There is also much protein-to-protein variation in the intensity of color development. Ideally, the standard should be similar to the unknown. For example, if one is measuring IgG concentrations, an immunoglobulin standard would be ideal. For serum, use bovine serum albumin as a standard since albumin is a major component of serum.
An easy and accurate alternative, based on the binding of protein to Coomassie Blue G-250 dye, is theÂ Bradford procedure. In addition, a modification of the Lowry procedure exists based on use of bicinchoninic acid (BCA) in place of the Folin-phenol reagent [Smith et al.,Â Anal Biochem.150, 76-85 (1985)]. The BCA is less prone to interference than the Lowry procedure and is more sensitive.
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Compared to Lowry Method, Bradford stands out for its simplicity by using only one reagent and greater sensitivity which is four times more than Lowry method. Bradford method is just another common colorimetric method to determine protein concentration in a sample solution. The Bradford method of protein determination is based on the binding of a dye, Coomasie brilliant Blue and shift of absorbance maximum of the dye from 495nm to 595 nm and is proportional to protein concentration when compared to a standard curve.
The Bradford assay is very fast and uses about the same amount of protein as the Lowry assay. It is fairly accurate and samples that are out of range can be retested within minutes. The Bradford is recommended for general use, especially for determining protein content of cell fractions and assessing protein concentrations for gel electrophoresis.
Assay materials include colour reagent and protein standard. It is sensitive to about 5 to 200 micrograms protein, depending on the dye quality. In assays using 5 ml colour reagent prepared in lab, the sensitive range is closer to 5 to 100 Âµg protein. Scale down the volume for the "microassay procedure," which uses 1 ml cuvettes.
The assay is based on the observation that the shift of the absorbance maximum for an acidic solution of Coomassie Brilliant Blue G-250 when binding to protein occurs. Both hydrophobic and ionic interactions stabilize the anionic form of the dye, causing a visible colour change. The assay is useful since the extinction coefficient of a dye-albumin complex solution is constant over a 10-fold concentration range.
The Bradford reagent is prepared by dissolving 100 mg Coomassie Brilliant Blue G-250 in 50 ml 95% ethanol and added 100 ml 85% (w/v) phosphoric acid. 1 liter of it is dilutes when the dye has completely dissolved and it need to be filtered through Whatman #1 paper just before use. However, there is an optional whereby 1 M NaOH can be used if samples are not readily soluble in the colour reagent). The Bradford reagent should be a light brown in color. Filtration may have to be repeated to rid the reagent of blue components. The Bio-Rad concentrate is expensive, but the lots of dye used have apparently been screened for maximum effectiveness. "Homemade" reagent works quite well but is usually not as sensitive as the Bio-Rad product.
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A standard curve of absorbance versus micrograms protein and determine amounts from the curve is plotted. The concentrations of original samples from the amount protein, volume/sample, and dilution factor are determined through the graph.
The dye reagent reacts primarily with arginine residues and less so with histidine, lysine, tyrosine, tryptophan, and phenylalanine residues. Obviously, the assay is less accurate for basic or acidic proteins. The Bradford assay is rather sensitive to bovine serum albumin, more so than "average" proteins, by about a factor of two. Immunoglogin G (IgG - gamma globulin) is the preferred protein standard. The addition of 1 M NaOH was suggested by Stoscheck (1990) to allow the solubilization of membrane proteins and reduce the protein-to-protein variation in color yield.
However, it has some limitation (Bergmeyer & Grabl 1983). The standard curves are not linear for many proteins, especially those with more than 60ug of proteins. This inherent nonlinearity is caused by the reagent itself as there is an overlap in the spectrum of the two different colour form of the dye. The background value of the reagent is continually decreasing when more proteins is bound to the dye. Absorbance may also vary with the age of the reagent. Another serious issue is the variations of the response with different proteins. When tested with 23 different proteins, the standard deviation in estimates of the proteins concentration by Bradford method was twice the value obtained by Lowry method. Hence, it is important to use a standard pprotein that gives a similar colour yield.
Absorbance Ratio Against BSA
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In Biochemistry, gel electrophoresis is a process where the molecules such as proteins, DNA, of RNA fragments are moved in an electrical field with a velocity that proportional to its overall charge density, size, and shape. Generally, there are two types of electrophoresis used in the separation of protein, that is native gel electrophoresis and sodium doecyl sulphate polyacrylamide (SDS-PAGE) gel electrophoresis.
Native gel electrophoresis is a technique used mainly to separate proteins based on their charge to mass ratio where the proteins are not denatured at the end of the process. Native gel electrophoresis is further divided into two types, which is polyacrylamide gel electrophoresis (PAGE) and agarose gel electrophoresis.
Agarose gel has a smaller resolving power as compared with polyacrylamide gel. However, agarose has a greater range of separation, that is from 200 bp to >50000b by using standard gels and electrophoresis equipment. On the other hand, polyacrylamide gel has a resolving power in the range of about 5 to 1000 bp. Moreover, they are much more difficult to handle than agarose gels. In addition, the pore size of agarose gel is not uniform but polyacrylamide gel has uniform pore size. The pore size is determined by the amount of cross-linker and the total amount of acrylamide present.
The pH of the gel is high usually around pH 9 so that all of the molecules have negative charge and will move towards the positive electrode when the current is on. In this case, the anionic system is used where the electrodes are arranged in a way that cathode is on the upper part and anode is on the lower part. The negatively charged anions are allowed to flow toward the anode under certain duration of time and the electric current will force the molecules to pass through the gel. For the molecules with a relatively homogeneous composition like nucleic acids having a constant shape and charge density, thus the velocity is depends on size. The smaller molecules will move faster through the gel than the heavier molecules and thus migrate farther in a specific time. The molecules with similar size and charge will moves through the gel as a band.
After electrophoresis, the gel may be stained by soaking in a solution of a stain that binds tightly to proteins such as Coomassie Brilliant Blue R-250 or silver stain to visualize the separated proteins. After staining, the different proteins will appear as distinct bands within the gel. The sizes of the various fragments can be determined by comparing their electrophoretic mobilities to the mobilities of fragments of known size.
However, if the proteins in a sample are radioactive, the gel can be dried and then clamped over a sheet of X-ray film. The film will developed after some time and an autoradiograph is formed. The positions of the radioactive components are shown by the dark region on the film. On the other hand, if the protein consists of an antibody, immunoblotting or Western blotting is used to detect the specific protein on a gel in the presence of many other proteins. The samples contain less than a nanogram of protein can be separated and detected by gel electrophoresis depends on the dimensions of the gel and the visualization technique used.
Sodium doecyl sulphate polyacrylamide (SDS-PAGE) gel electrophoresis is a technique use to separate proteins according to their molecular weight where the proteins are denatured at the end of the process.
In SDS-PAGE gel electrophoresis, the detergent sodium dodecyl sulfate (SDS) is used to denature protein in polyacrylamide gel electrophoresis.
Since the SDS is a detergent soap that consists of a negative charge where sulfate is attached to as well as it can dissolve the hydrophobic molecules. Therefore, in the first step, the cell is incubated with SDS to dissolve the membranes of a cell and the proteins will be solublized by the detergent as well as the proteins will be covered with the negative charges. There are two important features in this technique that are all proteins contain only primary structure and all proteins will migrate towards the positive pole when placed in an electric field since all proteins have a large negative charge. The net result is that SDS-treated proteins have similar shapes and charge-to-mass ratios.
Next, the proteins are allowed to move at different rates in a process called PAGE as mentioned before. This is because the SDS-treated proteins have similar shapes and charge-to-mass ratios, so they tend to move towards the positive pole at the same rate without any separation according to the size. The electricity is used to pull the proteins through the gel, thus the whole process is called polyacrylamide gel electrophoresis (PAGE).
The smaller size molecules will move through the polyacrylamide gel easily than the larger molecules. The protein cannot be left for electrophoresis too long, since the proteins have so many copies which are of the same sizes, thus they tend to move through the gel in the same rate. Therefore, the current have to turn off and then stain the proteins to observe how far the protein molecules moved through the gel.
SDS-PAGE separates the proteins according to their primary structure of size without consider the amino acid sequence. Following electrophoresis, the gel may be stained, most commonly with Coomassie Brilliant Blue R-250 or silver stain to visualize the separated proteins. After staining, different proteins will appear as distinct bands within the gel. In order to calibrate the gel and determine the molecular weight of an unknown protein, another lane of known molecular weight is run in the gel as a marker. Later, the molecular weight of an unknown protein is determined by comparing the distance traveled relative to the marker.