Lab 4.pdf

LABORATORY EXERCISE 4 Measuring Protein in Solution Learning Objectives • Understand a colorimetric method to quantitate the amount of protein in a solution • Prepare and use a standard curve, understand limitations of a standard curve • Understand the limitations of colorimetric measurements Intro...

LABORATORY EXERCISE 4 Measuring Protein in Solution Learning Objectives • Understand a colorimetric method to quantitate the amount of protein in a solution • Prepare and use a standard curve, understand limitations of a standard curve • Understand the limitations of colorimetric measurements Introduction Proteins are an important part of a healthy diet. We all need protein in our diet because our bodies cannot synthesize all of the essential amino acids that are needed to make proteins in our cells. Some diets designed for weight loss recommend decreased carbohydrate and fat intake while increasing protein. The scientific notion behind these recommendations is that increased protein intake has been shown to reduce the hunger hormone (ghrelin) and also boost several appetite suppressing hormones. So how can the protein levels in food be measured? What is a protein? Proteins are one of the major groups of macromolecules in the cell. Roughly 55% of the dry weight of the cell is protein. Proteins have many different functions in the cell (motility, transporters, enzymes, intercellular communication, receptors, hormones, structural proteins). Proteins are polymers of amino acids that each have a unique 3-dimensional structure that is important for their function. The 3-D shape of a protein is based on the chemical properties of the amino acids in the protein and their order (sequence) along the polypeptide chain. Proteins are made of amino acids linked together in a chemical chain (polypeptide). The general structure of the amino acid is shown in Figure 4.1. All amino acids have a carboxyl group (COOH), an amino group (NH2), and a central or alpha carbon (Cα). Each different amino acid contains a particular chemical structure (side chain) linked to the central carbon. That unique structure is referred to as an R group. R-groups are important in determining the final structure and function of a protein. Figure 4.1 Non-ionized form of a general amino acid Amino acids form polymers when the carboxyl group of one amino acid bonds with the amino group of a second amino acid to form a peptide bond as shown in Figure 4.2. Joining two amino acids produces a dipeptide (di = two). Additional amino acids can be added to the carboxyl group of this dipeptide until a polypeptide (poly = many) is formed. This chain has a free amino group (N-terminus) at one end and a free carboxyl group at the other end (C- terminus). Figure 4.2 Two amino acids joined together into a dipeptide How can the amount of protein in food be determined experimentally? Scientists frequently need to determine the amount of protein contained in a solution. For instance, every food item you can purchase at the grocery store is required by law to list the amount of protein per serving. There are several methods available to quantify the amount of protein in a solution. The method we will use is the “biuret” method, which is a colorimetric chemical test that detects the presence of peptide bonds. Colorimetric tests are commonly used in scientific research since they provide a specific color that is relative to the amount of substance you are measuring. For example, pH strips are a colorimetric way to test the basic or acidic nature of a solution. Similarly, the biuret method measures the presence of peptides using a copper (II) ion that will form mauve-colored coordination complexes in an alkaline solution. The test is named biuret because the positive reaction of the peptide bonds is similar to the reaction observed with the biuret molecule (Figure 4.3). The biuret method can be used to assess the concentration of proteins because the amount of peptide bonds is directly related to the amount of protein. Therefore, the intensity of the color, and the absorption of light at 540 nm wavelength, is directly proportional to the protein concentration. Figure 4.3 In the biuret reaction, copper ions bind to amine groups created by peptide bonds. The resulting purple color can be quantitated with a spectrophotometer (SpectroVis Plus). When measuring the amount of protein contained in a solution, the first step is to create a standard curve. A standard curve is a graph that illustrates the linear relationship between known amounts of an X variable, and its corresponding Y variable. In this exercise, we will use known amounts of a protein called bovine serum albumin (BSA) to create a standard curve. Different amounts of the known protein will be reacted with the biuret reagent and the absorbance of the solution will be measured at wavelength 540 nm. The absorbance (at 540 nm) will increase as the amount of protein increases. By graphing the amount of protein (x axis) versus the absorbance measurement (y axis), you can determine the equation of the line for that linear relationship. That is your “standard curve”; it will be most useful to use the portion of the standard curve that is linear. The standard curve can then be used to determine the protein concentration in solutions where the amount of protein is unknown by determining the absorbance for those samples using the same reaction conditions and then reading off the standard curve. Make a Prediction: In this exercise, we will measure various solutions containing a dietary supplement protein powder and a solution of glycine amino acid. Think about how the biuret reagent is binding, do you expect to detect protein in both types of solutions? Before we begin, record your hypothesis below. Liquid handling with a micropipette In biological research, we frequently need to measure a wide range of volumes to perform a single experiment. For example, you may need to measure 1 L of volume to make a solution, but you may only need 1 mL of volume for your experimental test. This range of volume necessitates the use of different sets of tools. For instance, you would not weigh your car using a bathroom scale, nor would you weigh yourself using a highway scale. This leads us to the use of specialized liquid handling equipment termed micropipettes for measuring extremely small volumes. For much of this work we will measure reagents with micropipettes. This is essentially a precision pump fitted with a disposable tip. Depressing the plunger displaces a specific volume of air; releasing it creates a vacuum, which draws an equal volume of fluid into the tip. Each micropipettor is accurate in a certain range. The maximum capacity of the pipet is indicated on the top of the pipet. The exact range varies from one manufacturer to another, but are usually as follows: Range (µl) P20 1-20 20-200 P200 200-1000 P1000 Figure 4.4- Micropipettes with their typical volume range in microliters. The green circle shows the dial that will be used to adjust the micropipette. The yellow circle shows the tip eject button. The exact range of measurement can be found at the top of the micropipette as shown in the black circle. The P20 and P200 use yellow tips, while the P1000 uses blue tips. For best accuracy, select the smallest pipet that can measure the volume you wish to measure. Set the pipet for the desired volume with a dialing mechanism. Some pipets have a locking mechanism, which must be UNLOCKED to set the volume. The adjusting dial should NOT beyond the upper or lower range indicated on the micropipettor. Reading the set volume Last digit is red First digit is red To measure a set value: 1. Set the Pipet 2. Put a tip on the pipet. The P20 and P200 use yellow tips; the P1000 uses blue tips. 3. Depress the plunger until you feel resistance (at the FIRST stop). This first stop is called the soft stop. Put the tip in the liquid (still holding the plunger down), once the tip is in the liquid slowly release the plunger. This aspirates the liquid. To expel liquid: push plunger ALL the way down. It is good practice to touch the tip to the side of the tube that you are dispensing into. Check the tube that the droplet has been delivered and the tip that it is empty. 4. Release the tip into the waste beaker. Always use a new tip when changing reagents. BEFORE MEASURING VOLUMES be sure to practice with the soft and hard stop so that you can feel the difference. It is very easy to make an error in your experiment due to poor pipetting technique. Procedure Overview of the Procedure: Part I: Tubes containing seven different known amounts of protein (Bovine serum albumin) will be set up in duplicate (Table 4.1). After reaction with biuret reagent the absorbance readings from these tubes will be used to produce a standard curve (Table 4.3, Graph). Part II: Tubes containing different dilutions of Isopure™ a commercially available protein supplement will be set up in triplicate for measuring protein (Table 4.2). After reaction with biuret reagent the amount of protein in these tubes will be determined using the absorbance value and reading off the standard curve (Table 4.4, Graph). Back calculation from different dilutions will provide independent measures of the amount of protein in the original stock solution (Table 4.5). Part III: Once protein and saline are added into all the tubes, the same amount of biuret reagent is added to all 29 experimental tubes at the same time. The reaction needs to sit for 20 minutes to go to completion. Then the absorbance at a wavelength of 540 nm is read using the calibrated SpectroVis Plus, and the absorbance readings are recorded (Table 4.3 and 4.4). Processing the data Graphing and calculations are completed in this section. There are questions for consideration throughout the exercise and summary questions at the end. Part I Preparation of a protein standard curve with Bovine serum albumin (a blood protein) 1. You will need small amounts of several reagents to use at your table. Collect from the provided stock solutions provided in the laboratory, the amounts you will need and bring them to your table in small beakers. Label each of the beakers of to identify the contents. PLEASE TAKE WHAT YOU NEED BUT DO NOT BE WASTEFUL. a. Pour approximately 40 mL of phosphate buffered saline into a labeled beaker. Label it PBS (phosphate buffered saline). This is a buffered salt solution that is use to mix with the protein solutions. b. Pour approximately 25 mL of Biuret reagent into a labeled beaker. Label it BR. It will appear blue in color. c. Pour approximately 15 mL of BSA (Bovine serum albumin, 10.0 g/L) into a beaker. This is the protein standard of known concentration. 2. Set-up the Standard Curve You will set up fourteen test tubes with different amounts of Bovine serum albumin (BSA) solution and different amounts of saline solution (PBS) to create a set of protein standards of different concentration and duplicates of each concentration. Label the tubes 1-14. Question for thought: Before you get started, consider which tool is most appropriate for measuring volumes under 1mL. Question for thought: It will be useful to remember that 1 mL = 1000 µL. So 0.15 ml = µL Pipette the volumes of BSA and PBS as shown in Table 4.1 to correctly dilute the BSA standard to the desired concentration. First, fill each test tube with the correct volume of PBS. Second, add the correct volume of BSA to each test tube. All tubes should have a final volume of 2 mL. If any of them do not look correct, redo that tube. Mix the tubes carefully using the Vortex. Table 4.1 Creating a Standard Curve (Line) Volume of BSA stock solution (mL) Add 2nd Final concentration of BSA (mg/mL) For Reference 1-2 Volume of PBS (mL) Add 1st 2.0 0 0 3-4 1.85 0.15 0.4 5-6 1.7 0.3 0.8 7-8 1.4 0.6 1.6 9-10 1.0 1.0 2.4 11-12 0.7 1.3 3.2 13-14 0.4 1.6 4 Tube # Part II Dilution of protein samples with unknown proteinconcentration 1. You will need small amounts of five protein solutions that you will use to determine their protein concentration for this part of the experiment. As before, collect from the stock solutions provided in the laboratory in the amounts needed for your experiment and bring them to your table in small beakers. Label each of the beakers to identify the contents. PLEASE TAKE WHAT YOU NEED BUT DO NOT BE WASTEFUL. a. Pour approximately 8 mL of Solution A: Isopure™ protein supplement stock solution (3g/100mL). Label this beaker A. b. Label one beaker B and set this beaker aside for step 2. c. Pour approximately 8 mL Solution C into a small beaker. Label this beaker C. d. Pour approximately 8 mL Solution D into a small beaker. Label this beaker D. e. Pour approximately 8 mL glycine solution. Label this beaker G. 2. In the beaker labeled B, add 6.3 mL PBS. Then add 700 µL (.7 mL) of 3% Isopure™ protein supplement stock solution from Beaker A. Carefully swirl the beaker to ensure the solution is well mixed. Table 4.2. Volumes of Protein Solutions Tube # Volume of protein solution (mL) 15-17 A 2 mL stock Isopure A 18-20 B 2 mL of solution B 21-23 C 2 mL of solution C 24-26 D 2 mL of solution D 27-29 G 2 mL of glycine solution G 3. Label 15 test tubes #15-29 as shown in Table 4.2 and pipette 2 mL of the appropriate protein sample into its respective tube. Change pipet tips between the different typesof samples. Part III Setting up the Biuret reaction with all the tubes 1. Add 0.7 mL of Biuret solution to ALL 29 prepared tubes as quickly as possible. 2. **VERY IMPORTANT**. Mix all of the tubes by vortexing carefully to ensure the protein sample and Biuret solutions are well mixed. 3. Allow the protein to react with the biuret solution for 20 minutes at room temperature. Question for thought: What is happening in this time period? Question for thought: How might the results be affected if the standards and the unknown protein tubes were incubated for different amounts of time? 4. Preparing the SpectroVis Plus for measurements Near the end of the 20 minute time period, turn on the LabQuest. Calibrate the Touch screen, if necessary, as previously shown (in Exercise 2). Plug in the SpectroVis Plus. A red box that says USB:Abs should appear. Touch the red box with the stylus and select change wavelength. Change the wavelength to 540 nm. 540 nm is the maximum absorbance for Biuret solution. In the upper right-hand corner, tap the Mode window and select Events with Entry. Press OK. 5. Calibrate the spectrometer. a. Tap the USB:Abs window, choose Calibrate ►Spectrometer. The calibration dialog box will display the message: “Waiting 90 seconds for lamp to warm up.” After 90 seconds, the message will change to “Warm-up complete.” When the 20 minute incubation period is finished, complete the calibration by performing step B. b. Pour Tube 1, a 2mL Zero protein sample, into the cuvette. Place the cuvette into the spectrophotometer. Click Finish Calibration and allow the calibration to finish. Click . The spectrophotometer should display a 0.00 ABS value in the red box. Remove the cuvette from the SpectroVis Plus and pour the solution back into Tube 1. KEEP this cuvette containing the solution from Tube 1. It is your blank for the absorbance readings. Next we will measure the absorbance of the reactions. 6. You are now ready to collect absorbance data for the protein standards. Pour contents from Tube 2 into another cuvette. Wipe the outside of the cuvette with a laboratory tissue and place it in the SpectroVis Plus. Wait for the absorbance value displayed on the screen to stabilize (this may take ~ 5 seconds). Enter the absorbance value shown in the red box in the 0.00 mg/mL row. HINT – the value should be near 0.00. Do not worry if the value displayed is a negative value (eg. – 0.004). Remove the cuvette. Discard the solution in your Waste beaker. Rinse the cuvette with water. Shake out any excess water. Reread solution from Tube 1 (the Blank) and enterthe data in Table 3.3. Remove the cuvette from the SpectroVis Plus and set it aside should you need to check the blank later in the experiment. 7. You are now ready to begin reading the protein standard tubes in a sequential manner. Begin with Tube 3. Pour the contents from Tube 3 into a cuvette. Wipe the outside and place it in the SpectroVis Plus. When the absorbance value stabilizes, enter the value shown in the red box in the correct mg/mL row in Table 4.3. Remove the cuvette from the SpectroVis Plus and rinse it well with water between each new sample. Shake out any excess water. Continue the same process for Tubes 4-14. Note: The duplicate samples should give similar absorbance readings. The absorbance should increase with increasing amount of protein. If any of the readings look unusual, check that your blank is zero and that you have the correct solution in your cuvette. 8. After you have finished recording the value of the last standard curve sample (Tube 14), you are ready to collect absorbance data for your protein unknown concentration solution samples. Obtain Tube 15, pour its contents into the cuvette, and place it in the SpectroVis Plus. When the absorbance value stabilizes, record the value in Table 4.4. Remove the cuvette from the SpectroVis Plus and rinse it well with water. Shake out any excess water. Repeat step 8 for all of the remaining samples (15-29) making certain to rinse and shake dry the cuvette between each sample. After the last sample has been recorded, proceed to Processing the Data. Data and Calculations Table 4.3 Absorbance Readings for Standard Curve Protein Concentration (mg/mL) Absorbance 0.00 1. 2. 0.4 3. 4. 0.8 5. 6. 1.6 7. 8. 2.4 9. 10. 3.2 11. 12. 4.0 13. 14. Line equation for Standard Curve Average Absorbance Question for thought: Are the measurements of the duplicate samples similar to each other? Question for thought: In general, is the absorbance increasing with higher values of protein? Table 4.4 Absorbance Readings for Unknown Protein Solutions Samples Absorbance Isopure™ Stock Sol. A 15. Absorbance Absorbance 16. 17. Solution B 18. 19. 20. Solution C 21. 22. 23. Solution D 24. 25. 26. Glycine G 27. 28. 29. Average Absorbance Standard Deviation Processing the Data Determination of protein content in different samples using the Biuret Protein Assay 1. Plot the average absorbance values from Table 4.3 to make a standard curve graph. Draw a smooth-fit line (i.e. curve) to best fit the data. Good scientists critique their data to assess its validity. When you doubled the amount of protein in your standard curve, did the absorbance value approximately double? I.E. is the average ABS value for 0.8 mg/mL double that of 0.4 mg/mL? Should all of your standard curve data points be used to draw a best fit line? Explain why or why not. 2. Determine the line equation for yourstandard curve. Enter the equation in Table 4.3. 3. Calculate the average absorbance of each protein sample and the standard deviation between replicates in Table 4.4. 4. Estimate how much protein is in each unknown sample set by using the average absorbance and reading the corresponding protein concentration from your standard curve graph. Record these values in Table 4.5 (below). 5. Calculate the amount of protein (mg/mL) for each sample set by using the line equation from your standard curve graph (Table 4.3). Record these values in Table 4.5. Are the estimates and values calculated from the line equation close? Why or why not? Now let’s consider solution B. To prepare solution B, you made a dilution of the Isopure™ Stock Solution. Think of a dilution in the following manner. In Figure 4.4 shown below, the volume of the large cube can be determined by knowing the volume of the smaller cube, and multiplying by the number of small cubes that fit inside the large cube. Figure 4.4. Volume of the large cube = volume of small cube multiplied by the number of small cubes that would fit inside the large cube. In this laboratory exercise, you measured the amount of protein in various solutions. Solution A is an undiluted, original stock solution of Isopure™ protein powder (similar to the large box shown above). Solutions B, C, and D are diluted, i.e. the amount of protein in these samples is less than the original stock solution (similar to the small box shown above). Measuring diluted samples is often necessary when the original stock may have so much protein that its absorbance value may exceed the highest data point on your standard curve graph. Note that you diluted the Isopure™ Solution by a factor of 10 when preparing solution B: The total volume of Solution B was 7 mL. We added 0.7 mL of Isopure™ stock + 6.3 mL of PBS = 7 mL 7 mL / 0.7 mL = 10  This is the dilution factor. The amount of protein you measured in the diluted solution B sample only represents 1/10 of the Isopure™ stock solution. You must multiply the protein concentration that you calculated for solution B by 10 to get back to the actual protein concentration of undiluted Isopure™ solution. Solutions C and D were diluted as well, but these solutions were already prepared for you. 6. Solution C was prepared by mixing 0.5mL of Isopure™ stock solution with 9.5mL PBS. Using the same logic as above, determine the dilution factor for Solution C. Enter this value in Table 4.5. Total volume /amount of Isopure™ stock = / = the dilution factor Calculate the amount of Isopure™ protein in Solution A by multiplying the line eq. value (mg/mL Solution C) by its dilution factor value. Record these values in the space provided in Table 4.5. 7. Solution D was prepared by mixing 0.1mL of Isopure™ stock solution with 9.9mL PBS. Using the same logic as above, determine the dilution factor for Solution D. Enter this value in Table 4.5. Calculate the amount of Isopure™ protein in Solution A by multiplying the line eq. value (mg/mL Solution D) by its dilution factor value. Record these values in the space provided in Table 4.5. Table 4.5 Determining the protein concentration in Isopure solutions Samples Solution A Isopure™ Stock Solution B Protein conc. estimate by looking at your graph (mg/mL) Protein conc. determined by using the line equation (mg/mL) Dilution Factor Amount of protein in Solution A (mg/mL) XXXXX 10 Solution C Solution D Glycine G XXXXX XXXXX 8. Determine how much protein you EXPECTED to measure in Solution A by using the following information. Isopure™ unflavored protein powder contains 25 g protein per 29 g serving (there are 4 g of other stuff per serving, such as, sugar, cholesterol, salts, etc.). The Isopure™ stock solution you used (Sol. A) was prepared by mixing 3.0 g Isopure™ powder in 100mL PBS. What is the expected amount of protein (mg/mL) in the IP stock solution? You may round-off the answer. (HINT: you’ll need to convert g to mg to get started.) Figure 4.5 Nutrition information for Isopure™ unflavored protein powder. Does the amount of protein in Solution A, determined by measuring dilutions (i.e. Sol. B, C, and D) and multiplying by their respective dilution factors, equal the Expected amount as determined by the above calculation? Which measurements are closest (more accurate)? 9. Did measuring undiluted Solution A provide you with a valid answer for how much protein is contained in Solution A? Explain why or why not? 10. Finally, consider what you know about the glycine solution and how much protein it should contain. You made a prediction about this in the beginning of the exercise. Comment on whether your measurements agree with this expectation. Explain why or why not? Questions (Some of these you have already considered during the exercise, you can summarize the answers to questions here.) 1. In this experiment, you have replicates for each of your sample measurements. What does your SD value tell you about the replicates? Explain. 2. Compare the adjusted protein values that you observed for each sample to the expected protein values. How well do your adjusted values align with the expected values? Can you think of any reason why they would be different? 3. Did the biuret reagent turn a purple color in the presence of 1% solution glycine? Did the biuret reagent react with the amino acid solution even though it is not protein? Can you explain why this would or would not happen? 4. Which of the Isopure™ samples (solutions A-D) gave the best results, i.e., yielded a protein value nearest to the expected value? Explain why these results may have happened. 5. What are the limitations of a standard curve? Explain. 6. If the ABS from an unknown sample does not fall upon the linear portion of the standard curve, is it valid to use this ABS in further calculations? 7. Challenge Question: The manufacturer recommends dissolving 29g Isopure™ protein powder in 8oz liquid, which roughly translates to 12.25g/100mL. If we utilized such a 12.25% solution of Isopure as the stock solution for our next experiment, what might be a suitable dilution factor we could use to obtain useful measurements?