Chem 405 Biochemistry Lab I Experiment 2 Quantitation of an unknown protein solution. Introduction: The determination of protein concentration is frequently required in biochemical work. Several methods are available; each having features that suit it to a particular use. We will conduct three different protein assays in this laboratory. There are many reasons to conduct a protein assay. During a purification of a protein, you need to know how pure your sample is by determining the amount of enzymatic activity vs. protein concentration. When comparing samples on an SDS-PAGE gel or using antibody for a western blot, you need to add the same amount of protein form various preparations to the true results. Most protein assays take advantage of a reaction between a reagent dye and the protein of interest that will shift or increase the absorbance of a particular wavelength. Generally the more protein in a sample the higher the absorbance. Protein assay basics: Standard curve: The Bradford and Lowry protein assay lead to a change in absorbance when protein is present. How does this absorbance relate to the actual protein concentration? To know determine the actual concentration of a protein a standard curve is required. A standard curve is a plot of absorbance vs. a varying amount of substance in parallel with unknown protein sample. There are two methods to prepare a standard curve. Absorbance vs. total amount of standard protein, usually in mg or µg of protein. For this style of standard curve, the protein amount in the unknown sample is divided by the volume of sample added to the colormetric reagent. After taking any dilutions into account, the concentration is expressed as mg/ml. A second common method to prepare a standard curve is to prepare a set standards with various known protein concentrations. As long as the volume of the standard samples and the unknown samples are the same the final concentration of the unknown is directly calculated from the least squares line of the standard curve. Of course you still have to correct for any dilution. Don't feel bad if this is confusing, it is for most students and will take several times before you are confident at it. We will get plenty of chances to work on this throughout the semester. Samples should be run in duplicate or triplicate and the averages used for the graph. If the same cuvette or a sipper is used for reading samples, it is a good idea to read those samples with less protein content first to reduce error arising from reagent dye carryover in the cuvette as a result of incomplete rinsing. Depending on the method, proteins generally vary in the reaction with the color dye in the different protein assays. Therefore, it is important to denote what the specific control or reference protein was used to make the standard curve. Two common proteins used for standard curves are bovine serum albumin (BSA) and an immunoglobin (IgG). These two proteins have different amino acid compositions, which leads to a different standard curve and a slight difference in the final determination of the unknown protein concentration. Because the color development is dependent on the amino acid composition of the protein and the presence of a prosthetic group (especially carbohydrate) also influence the protein assay; a purified sample of
the protein being assayed or a closely related protein is a preferred standard. Once you have performed the assay a standard curve is generated and the results graphed. Before using the standard curve you ve generated you must be certain that the absorbance is a linear function of concentration which holds within the limits of the Beer- Lambert Law. A = ε c l Abs = extinction coefficient x molar concentration of The sample x pathlength of the cell Beer s law does not hold at high concentrations. A common reason is the depletion of one of the reagents necessary for color production. Thus readings should always be taken in the region where all reagents are in excess (the curve of abs vs. concentration is linear). Sample Preparation: When determining the protein concentration of an unknown sample, several dilutions should be used to ensure the protein concentration is within the range of the assay. Usually 10 fold dilutions are used to get the unknowns within the standard curve range. While this may seem redundant and a pain in the neck (or other body parts), often times your sample may be too diluted to fit within the standard curve. Without the additional dilutions, you will have to start over again. Don t forget that all dilutions must be taken into account in calculating the final concentration of the protein. Finally, with every assay a blank must be included. The blank or the tube without a standard protein is usually made up in the same buffer as in the samples. This way if the buffer has an effect on the protein assay reagent, each sample (and the blank) will have been altered the same. The blank is used to set the instrument to the 100% transmittance or 0 absorbance. When assaying protein, it is important that the volume is the same for each tube. Protein is generally added first, followed by enough buffer to bring each tube to the same volume. The tubes are mixed well after each addition. The color-producing reagent is always added last, and the reaction may need to be timed accurately. Experiment In this experiment you will be provided with a protein standard (BSA) at a known concentration (1.5 mg/ml) and an unknown sample containing protein at unknown concentration. Your objective will be to measure protein concentration by the Bradford, Lowry and Warburg-Christain methods. Compare your results with each technique and compare the different types of assays and become familiar with each technique. The first step in the analysis is to prepare dilutions of the BSA standard and to construct a standard curve for parts 2 and 3. To determine the apparent protein concentration in the unknowns, you will need to examine several dilutions. It is a good experimental design to perform the standards and the unknowns side by side and at the same time. This ensures that variables such as temperature, time, batches of reagents, etc., are internally controlled. You also should perform the assays in duplicate. This way large pipetting errors may be easily recognized and mental error ascertained. For parts 2 and 3 it will be very helpful to set up a table of volumes to add in advance. The table in the handout is only an example and is not sufficient for the whole experiment. Don t forget that this table and all calculations will be in the lab book. Also be certain to include an outline of the procedures in you lab book prior to lab.
Part 1) Warburg-Christain Method The concentration of protein can be estimated by measuring the absorbance of protein at 280 and 260 nm. Be careful that a quartz cuvette is used, glass and plastic cuvettes will not transmit about 300 nm and smaller. Most proteins have an absorbance maximum at 280 nm, owing to the presence of tyrosine, tryptophan and phenylalanine. The benefits of this method are that the sample is not destroyed and can be used for other experiments. The disadvantage is the large volume of sample required to fill the cuvette and it is less sensitive. Since the amino acid composition varies greatly between proteins, the molar absorptivity will change. Proteins that have a low aromatic amino acid content will have low or no absorbance at 280 nm. Nucleic acids often are found to be a contaminant in many protein preparations. The nucleic acids will absorb maximally at 260 nm and have a large absorbance at 280 nm. The absorbance of nucleic acids is about 10 times greater than the same concentration of a protein at 280 nm. To correct for this Warburg and Christian developed a method to adjust for interference by nucleic acids. Mixtures of pure protein and pure nucleic acids were used to come up with a correction factor for the absorbencies at 280 and 260 nm. To determine the concentration of your protein use the correction factor shown below: Protein (mg/ml) = 1.55 A280 0.76 A260 Procedure. 1. Prepare a 0.75 ml 1:4 dilution of your unknown protein. 2. Determine the absorbance of the protein at both 280 and 260 nm 3. Use the equation above to determine the concentration of the protein in your solution. Part 2) The Lowry method The Lowry method is one of the most sensitive and widely used. The Lowry procedure can detect protein levels as low as 5 µg. There are several limitations to this method. Several contaminants interfere with the assay. Most commonly: some buffers, ammonium sulfate (>0.15%) glycine and reducing agents such as dithiothreitol (DTT) or ß mercaptoethanol. The principle behind color development is an improvement on another method called the Biuret method. In alkaline conditions, copper (II) is thought to bind to the peptide nitrogen of proteins. This complex absorbs light maximally at 550 nm. This reaction is followed by the reduction under alkaline conditions of the Folin-Ciocalteu reagent. The Folin-Ciocalteu reagent is a dark blue-reduced molybdotungstates, which are reduced by tyrosine, tryprophan and polar amino acid. Materials Stock reagents Solution A: 1% w/v copper sulfate (CuSO4.5H2O) Solution B: 2% w/v Na/K tartrate Solution C: 0.2 M NaOH Solution D 4% Na carbonate Lowry reagent I: Add 24.5 ml of solution D to 24.5 ml of solution C. Then add 0.5 ml of solution A and 1 ml of solution B. This is the copper-alkali reagent (Lowry reagent I), which must be freshly prepared for each lab session Lowry Reagent II: Dilute the Folin-Ciocalteu reagent 1:2 with H2O (make 3.0 mls)
Procedure: 1. Prepare Lowery reagent I and II 2. In duplicate, prepare the standard curve using up to 6 points including the zero tube. Use small sized test tubes. The final volume of each sample will be 0.250 ml and the protein range from 0 to 250 µg of standard. Start with the stock standard BSA and create a set of dilutions using serial dilutions. How many is up to you, BUT thing about how the points will line up on the graph. Include a ZERO (a tube without any protein). 3. Prepare at least two dilutions of your unknown standard. Include a non-diluted sample. Prepare each in duplicate. 4. Prepare one additional sample (water, like the zero tube) to blank the spectrophotometer with. 5. Add 1.25 ml of Lowry reagent I to each sample. 6. Mix well and let stand at room temperature for 10 min. 7. Add 0.125 ml of Lowry reagent II 8. Immediately vortex and incubate at room temperature for 30 min. 9. Zero the spectrophotometer against a blank consisting of 0.250 ml of water as descrbed in step 4. 10. Measure absorbance at 750 nm and record in your lab book. A sample table to help you prepare your standards for the lowery protein assay is shown below: Tube # Reagent 1 2 3 BSA (µl) H2O (µl) Unknown (µl) Lowery Reagent I (ml) Lowery Reagent II (ml) Part 3) Dye-binding method (Bradford) The binding of the dye, Coomassie Brilliant Blue G-250, to proteins causes a shift in the absorbance maximum of the dye from 465 nm to 595 nm in acidic conditions. This method has several advantages over other methods and is about as sensitive as the Lowry method. Unlike the Lowry method there are few interfering substances, the only known compounds are high concentrations of the detergents triton x-100 and sodium dodecyl sulfate (SDS). This assay is quick and results observed immediately. The color is developed when the dye forms a strong, noncovalent complexes with proteins by electrostatic interactions with amino and carboxyl groups and via Van der Waals forces. There is a great deal of batch to batch variation for this assay so it is imperative to run a standard curve with each assay.
Materials 1X BioRad Dye reagent, BSA protein standard, microfuge tubes. Procedure: 1. In duplicate, prepare a standard solutions of at least 6 tubes, each with an increasing mass of protein. Use CV=CV to determine how much water and how much stock BSA solution to use in each tube. Use the microfuge tubes. Each sample should be 50 µl in volume after water and BSA have been added. Ignore the Bradford reagent, that comes later. The mass of the protein should range from 0 to 75 µg. 2. Prepare two or three dilutions of your unknown. Also assay a nondiluted sample. 3. Prepare a blank sample using 50 µl of water. 4. Add 1.5 ml of Bradford dye to each sample. 5. Vortex and incubate at room temperature for 5 min 6. Read the absorbance at 595 nm against the blank prepared in step 3. A sample table for the Bradford protein assay is shown below: Tube # Reagent 1 2 3 4 BSA (µl) H2O (µl) Unknown (µl) Bradford Reagent I (ml) Calculations: For both the Lowery and Bradford Assays - 1) Determine the mass of protein (mg) in each tube of your standard curve. 2) Average the readings for each standard. You can not conduct a standard deviation on an n=2. 3) Graph the absorbance of the standards vs. the mass of protein in each standard tube. Refer to last week s handout for determining which variable goes on which axis. The unknown absorbance can not be graphed. 4) Use a least squares line also known as a linear regression. Include the R 2 value in your graph. 5) Use the equation for the linear regression to calculate the mass of protein in your unknown. Don t forget to take into account any dilutions of the unknown. 6) Divide the mass of protein by the volume of sample of unknown you added to the test tube. 7) Report the answer in concentration (mg/ml for proteins). Lab Report: This will be a simple report. The grade will depend on: the completeness of the information requested below the calculations, observations recorded in the lab book the standard curves and the accuracy protein determinations of your unknowns.
Turn in your lab books with the following information. Tape graphs into the lab book. 1. Make certain the title and purpose is in the lab book. 2. Include the experiment in the table of contents 3. Include an outline of the procedures and reference the handout. 4. Include the table of additions for each of the three methods. Do this ahead of time. 5. Simply record the results for each section and prepare a graph of the standard curve for part 2 and 3. 6. Calculate the concentrations of your unknown (don t forget to indicate which unknown you used). The answer must be in mg/ml. 7. Finally in one very brief section discuss the differences between all three methods, any variations in the concentration of the unknown as determined by each method and your personal preferences for conducting this assay if you had a very small sample that you worked for 1 month to get.