Pipetting and Determining Protein Concentration

Transcription

1 Pipetting and Determining Protein Concentration Background Information: When performing experiments in Cell Biology, it is often necessary to use very small volumes of reagents sometimes because the reagents are incredibly expensive and other times because the products of our experiments are very easily detectible (so largevolume experiments simply are not necessary. Pipetmen are instruments used to accurately transfer small volumes (1 µl to 1 ml) of solutions. Pipetman is a brand name. The generic name for this instrument is micropipettor. In a research lab you are likely to hear these terms used interchangeably. Because of their accuracy, ease of use, and convenience in sterile techniques, they are practically universal lab tools. In this lab, you will learn how to properly use these instruments to transfer small volumes of fluid accurately. Pipetmen will be used in a number of this semester s experiments, so please be sure to feel comfortable in their proper use. Use illustration (right) and test below to become familiar with the parts of a Pipetman. Barrel - the working end of the Pipetman; a disposable tip is seated on the lower end of the barrel before each use Plunger - the plunger is pressed and released to withdraw and expel liquid Volume adjustment knob - is rotated to change the digital volume Volume display - displays the volume the Pipetman is currently set to deliver Tip ejector button - is used to remove a tip from the barrel without direct handling of the disposable tip A typical lab will use three different Pipetmen; each one is appropriate for a specific volume range. Somewhere on the micropipettor, you will see a number that represents the maximum volume, in microliters (µl) that can be transferred by that Pipetman. The minimum volume appropriate for each Pipetman is typically ten percent of the maximum. Note that not all micropipettors will have their minimum volume printed on them, but are always identified by their maximum volume. The table below shows the volume ranges, expected accuracies, and the appropriate tips for each of the pipetmen you will be using. Pipetman Volume range (µl) Accuracy Disposable tip P ± 10 µl Large (blue) P ± 1 µl Small (yellow) P ± 0.5 µl Small (yellow) Pipetman Volume Range (µl)

2 In most experiments, accuracy is important when transferring small volumes of liquid with a pipetman. Accuracy is achieved if the Pipetman has been calibrated and tested for accuracy (this is usually done on a yearly or semi-annual basis), and the instrument is being used properly. To transfer a volume accurately: 1. Select the appropriate micropipettor and adjust it using the dial on the plunger to the desired volume is indicated on the volume display 2. Place the appropriate tip on the barrel of the Pipetman, making sure it is tightly sealed 3. Depress the plunger only until you feel resistance. Note that the plunger can be depressed further; depressing it too far will result in inaccurate volume transfer. 4. Still holding the plunger, place the end of the tip just below the surface of the solution you are transferring. 5. Release the plunger slowly, making sure that the end of the tip remains under the surface of the solution being transferred. The solution should be sucked up into the tip as the plunger is released. 6. When the plunger is fully released, the tip should contain the desired volume. Place the tip into the bottom of the vessel into which the solution will be dispelled. 7. Slowly depress the plunger again until the solution is completely dispelled from the tip. To ensure that all of the fluid is dispelled, depress the plunger all the way down. Be sure to remove the tip from the fluid before releasing the plunger so as to avoid re-aspirating some of your fluid from its intended location. 8. Eject the used tip using the tip ejector button Accurate transfer of protein solutions is necessary when trying to determine the concentration of protein within a liquid sample. The Bradford protein assay is a simple and accurate procedure for determining the concentration of solubilized protein (Bradford 1976).The procedure is based on an absorbance shift observed when Coomassie brilliant blue G-250 dye binds to protein under acidic conditions. The intensity of this blue color reflects the amount of protein in the sample assayed. It is possible to use the chemical reaction described above to determine the amount of protein in any sample if one compares the intensity of the blue color produced to that of samples with known protein concentrations. This is accomplished through the generation of a standard curve. A standard curve is generated when the amount of blue color (measured as Absorbance of light at 595 nm) is plotted versus known protein concentrations. This plot enables us to determine the formula for a line; the formula can then be used to calculate the amount of protein in a sample of unknown protein concentration if you know the amount of blue color (again, by Absorbance at 595 nm).

3 Overall objectives: Become proficient in the use of micropipetters and learn to determine protein concentration using the Bradford protein assay to prepare a standard curve. Experimental Procedures: A. Pipetting 1. Familiarize yourself with the use of a Pipetman. a. Pick up a P-20 and set the volume setting for 10 µl (reading down the setting will should be 100 ). b. Push the plunger down and notice that at some point the plunger becomes more resistant and requires more effort to push further. This point is called the first stop. Notice that the plunger can be pushed well beyond the first stop until it reaches the second stop. Be sure you can feel the first stop point and notice how far the plunger travels before reaching this point. c. Reset the Pipetman to 1.0 µl ( 010 ). Again push the plunger to the first stop. You should notice that the plunger travels a much shorter distance before it reaches the first stop than it did when the Pipetman was set for 10 µl. 2. Transfer a solution using a Pipetman using the steps described in the background information above (they will be reiterated here). a. Set the volume dial to the desired volume by rotating the volume adjustment knob. Note: the volume setting should never be adjusted above the maximum volume specified for a particular Pipetman as this will RUIN the equipment. Remember that the maximum volume is the volume shown on the plunger. a. Place a disposable tip on the Pipetman by placing the end of the barrel into a tip and pressing down firmly. b. Push the plunger to the first stop and place the tip into the solution you are transferring. c. Slowly release the plunger to draw the desired volume of solution into the tip. Note: if you release the plunger too fast it will a) draw an inaccurate volume and b) splash solution into the barrel. Getting solution into the barrel can damage a Pipetman. d. Place the tip in the tube that is to receive the solution. e. Press the plunger to the first stop to expel the solution and then press to the second stop to blow out any residual fluid. f. Continue to hold the plunger down until you withdraw the tip from the tube to avoid drawing the solution back into the tip. Note: Depending on the situation, the same tip may be used again. In many cases, however, it is necessary to use a new tip for the each transfer. g. When you wish to eject the tip you are using, place the tip over the appropriate waste container and press the tip ejector button. If the tip is difficult to eject, it is likely that you are jamming the tips onto the Pipetman harder than necessary.

4 3. Check your pipetting accuracy. a. Obtain a 4 x 6-inch piece of blotting paper and a microcentrifuge tube containing blue dye. b. Following the protocol above, use a P-20 to spot the following volumes of dye directly onto the paper in a linear order: 1 µl, 3 µl, 5 µl, 7 µl, 10 µl, 15 µl, and 20 µl. c. Spot each volume twice to check your consistency. Compare the spots you made with those of the instructor. If they don t look right, check with the instructor to determine the cause. d. Obtain two microcentrifuge tubes and label them A and B. e. Transfer the volumes of solutions I through IV as indicated in the table below. Tube Solution I Solution II Solution III Solution IV A 100 µl 200 µl 150 µl 550 µl B 180 µl 230 µl 315 µl 275µl f. A total of 1000 µl was supposed to have been transferred to each of the two tubes. To verify your accuracy, set your P-1000 to 1000 µl, load it with a blue tip, and withdraw the solution mixture from each tube. The solution should just fill the tip. If there is solution left in either tube, or if there is a large portion of the tip that is left unfilled (has an air bubble), there may be a problem with our technique. Continue practicing or consult your instructor. When you are comfortable with manipulating the micropipettors, move on to Part B of the lab exercise. B. Bradford Protein Assay and Generation of a Standard Curve 1. Label 10 microcentrifuge tubes to correspond to the Known Concentrations given in the table below. 2. The stock BSA protein is at a concentration of 100 mg/ml. Add the following volumes of BSA solution and distilled water together to each tube. (NOTE: The final volume in each tube should be 800 µl.) Known Concentration of BSA and Tube Label Volume of Stock BSA Volume of dh 2 O 0 0 µl 800 µl µl 790 µl µl 780 µl µl 750 µls µl 700 µl µl 650 µl µl 600 µl µl 550 µl µl 500 µl µl 450 µl

5 3. Add 200 µl of Bradford dye reagent to each tube and vortex to mix. 4. Incubate at room temperature for at least 5 minutes. NOTE: The absorbance will increase over time so do not leave at room temperature for more than 50 minutes. 5. Pipette 250µl of each tube into each of 2 wells of the microtiter plate according to your map. 6. Read absorbance (also termed optical density or OD) at 595 nm using microtiter plate reader. Post-lab assignment: Use your data to do the following: a. Graph Absorbance (OD) at 595 nm versus protein concentration. Plot the average of your 4 OD/absorbance readings at each concentration to create a linear standard curve (this is most easily done in Excel). b. What is the R 2 value of your standard curve (if done in Excel, this information is also automatically calculated)? i. A perfectly linear standard curve has an R 2 value of 1. If your standard curve is not 1, hypothesize (provide logical reasons) why your standard curve is not perfect. c. Where does the standard curve plateau or peak (reach maximal value before the last known concentration or at the last known concentration)? d. If you had also assayed an experimental sample, you would have to backcalculate the amount of protein in that sample by using the formula (y = mx + b). i. Which of these values would the OD/absorbance of your unknown experimental sample? ii. Which of these values would be the unknown value that this formula will calculate? iii. Provide this formula for your standard curve. Note that it is likely that your graph (with all of your data points) contains some outliers points that should be discarded because they are outside of the linear range of the assay. To acquire the best line/formula, some of the data points outside of the linear range will have to be removed from your graph. To the best of your ability, determine which of these points to discard; removed them from your data and replot to produce a line with an R 2 value close to 1. Use this formula! This exercise is based on: Bradford, M.M A rapid and sensitive method for the quantitation of microgram quantities of protein using the principle of protein-dye binding. Anal. Biochem. 72: