Describe the procedures you followed by referring to the original source of the information (“Protein Quantitation in Unknowns” lab handout) instead of writing out all the steps.

After reading the “Protein Quantitation in Unknowns Lab”, please answer the two questions below. I haven’t completed the Lab but if you can answer in your own words the first two bulletin points please. This is a lab report.

Explain the purpose of this week’s exercise.

Describe the procedures you followed by referring to the original source of the information (“Protein Quantitation in Unknowns” lab handout) instead of writing out all the steps. If you made any changes, be sure to list them.

Protein Quantitation in Unknowns BIOS 312 Cell Biology Laboratory Page 1 of 7 Adapted from: Protein Assay and Measurement of Protein Content of Cells. Wolfe, Jason. Wesleyan University. ASCB Exercises in Cell Biology for the Undergraduate Laboratory, 1992. Objectives In this exercise you will: • gain additional experience in using micropipettes • use the Bradford assay and a spectrophotometer to measure protein concentration • create a standard curve that defines the relationship between two quantities • use your standard curve to calculate the protein concentrations in “unknown” samples Bradford protein assay of “unknown” samples Last week, you were introduced to the Bradford protein assay, which uses the dye Coomassie Blue G-250 to measure the relative amount of protein dissolved in a solution. Coomassie Blue forms a complex with proteins that absorbs light strongly at a wavelength of 595 nm, and the greater the protein concentration in a solution is, the greater light absorbance at 595 nm will be. Today, you will be following the same procedure to create a new standard curve, which you will then use to determine the protein concentrations in your saltwater and freshwater fish muscle cell extracts (referred to as “unknowns” since you don’t already know what their protein concentrations are.) The Excel spreadsheet will once again determine the slope and y-intercept of the linear regression trendline that describes your standard data, and you will then use these parameters to calculate the concentration of protein in a diluted sample of each cell extract based on its absorbance in the Bradford assay. IMPORTANT! There are multiple variables that can affect the outcome of the Bradford assay, including the specific micropipettes you use, the temperature of your solutions, and environmental conditions in the room at the time the assay is performed (air temperature, air pressure, humidity, etc.) To ensure that the standards and unknown samples are affected equally by these variables, you must assay them at the same time. If you create a standard curve on one day, you shouldn’t use it to determine protein concentrations of unknowns that you assay on another day. If you do, the results are likely to be inaccurate. After you determine the protein concentrations in your cell extracts, you’ll need to calculate the volume of each solution required to prepare 100 μL samples that contain 25 μg of protein in 5 μL. The protein solutions will be mixed with water and a concentrated Laemmli sample buffer that includes the detergent sodium dodecyl sulfate (SDS,) the reducing agent beta mercaptoethanol (BME,) glycerol, and the tracking dye bromophenol blue. As you learned earlier, SDS helps to denature proteins and gives them a strong negative charge that is proportional to the number of amino acids in the primary structure. BME breaks covalent disulfide bonds, which ensures that all proteins will have a uniform linear shape. Glycerol increases the density of the solution so that it will sink to the bottom of a well in a polyacrylamide gel, and the tracking dye migrates just ahead of most proteins during electrophoresis, making it easy to monitor their location in the gel. A reminder that you should always take precautions to minimize the activity of any proteases in your protein samples and avoid introducing new sources of protease. Keep your solutions cold and always wear gloves when you handle them – never touch the tubes with bare fingers! Don’t remove caps from tubes unless you need to – this allows particulates that could be contaminated with microorganisms to enter – and avoid talking or breathing over an open tube since this creates aerosols of saliva, which contains proteases.

BIOS 312 Cell Biology Laboratory Page 2 of 7 Materials Micropipettes, pipette tips Sharpie permanent marker Gloves (remember to wear these at all times!) 1.5 mL microcentrifuge tubes Clear plastic cuvettes Foam cuvette rack Ice bucket with crushed ice Set of diluted protein standards – see your TA for these Aliquot (portion) of Bradford dye reagent Aliquots of TBS (Tris-buffered saline) and dH2O (deionized water) Your saltwater (S) and freshwater (F) fish-muscle cell extracts from Week 2 Bradford assay spreadsheet for calculation of standard curve parameters Preparing samples for assay Each group will prepare ONE standard curve that will be used by the entire group. Each student in the group will perform the Bradford assay on 3 diluted samples of the S or F cell extract that they prepared on Day 1. 1. A set of BSA standards was prepared for you by diluting a 5 mg/mL BSA stock solution with TBS to the following concentrations: 1.25 mg/mL 1.0 mg/mL 0.75 mg/mL 0.5 mg/mL 0.25 mg/mL 0.125 mg/mL 2. Place 6 clean microcentrifuge tubes on ice. Label the lids 1.25, 1.0, 0.75, 0.5, 0.25, and 0.125 using your permanent marker. These will hold your protein standards. 3. Using the appropriate micropipette, one student from each group should transfer a 25 μL sample of each BSA standard to the corresponding labeled tube. Flick the standards with your index finger to mix them before pipetting; your TA will demonstrate this. Keep all materials on ice to limit protein degradation. 4. Each student should label 3 more clean microcentrifuge tubes as follows: 1/5, 1/10, 1/20. Add your initials to keep from confusing the different sets of tubes. Place these tubes on ice. They will hold diluted unknowns. 5. Retrieve your S and F cell extracts from the rack where they were stored. Thaw the tubes completely by rubbing them vigorously between your palms and mix the solutions by flicking with your index finger. Place the extracts on ice and keep the tubes closed when you are not using them.

BIOS 312 Cell Biology Laboratory Page 3 of 7 6. Each student should add 10 μL of their saltwater (S) or freshwater (F) fish muscle extract to each of their 3 dilution tubes. Add 40 μL of TBS to the “1/5” tubes (a 1/5 dilution,) 90 μL of TBS to the “1/10” tubes (a 1/10 dilution,) and 190 μL of TBS to the “1/20” tubes (a 1/20 dilution.) Mix each dilution by flicking it with your index finger and place it back on ice. Remember to keep the lids of the tubes closed. Return the tubes of undiluted cell extract to the storage rack so they can be placed back in the freezer. Bradford assay protocol You will perform the Bradford assay on your standards and diluted unknowns and then enter the absorbance readings for the standards into the Bradford Assay spreadsheet to generate a standard curve (follow your TA’s instructions.) If the R2 value (correlation coefficient) you obtain is above 0.90, then your standard curve is usable. If it does not meet this criterion, your TA will guide you in either eliminating an outlying data point or using another group’s standard curve. In this assay, as in many future exercises, pipetting technique is extremely important. If your pipetting is inaccurate, your standard curve will be too! 1. Turn on a spectrophotometer if it is not on already. Allow it to warm up for at least 10 minutes before use. Be sure it is set for “absorbance” mode (A) and a wavelength of 595 nm. 2. Put seven (7) cuvettes in your foam rack and label them near the top: 1.25 1.0 0.75 0.5 0.25 0.125 B (blank) These are for the group’s standards. 3. Each student in the group should place three more cuvettes in the rack and label them as follows (there will be a total of 12 additional cuvettes for the whole group): Initials + 1/5 Initials + 1/10 Initials + 1/20 These are for the unknowns (diluted fish-muscle cell extracts.) 4. Add 490 μL of dH2O to each cuvette in both the standard and unknown set. This will dilute the protein samples to a concentration range appropriate for testing. 5. Add 10 μL of each protein standard to the appropriate cuvette in the “standards” set. For the “B” cuvette (blank), add 10 μL of TBS (this is the buffer the BSA is diluted in.) 6. Add 10 μL of each diluted fish cell extract to the appropriate cuvette in your “unknowns” set. 7. Add 500 μL of 1X Bradford dye reagent to each cuvette. 8. Mix each sample thoroughly by gently pipetting 500 μL up and down ten times. Be sure there are no air bubbles in the light path of the cuvette. Use a Kimwipe to remove any smudges from the outer surface, and only handle the cuvettes by their top edges.

BIOS 312 Cell Biology Laboratory Page 4 of 7 9. Incubate samples at room temperature for at least 5 minutes but not longer than 30 minutes. 10. Put the blank (“B”) cuvette into the spectrophotometer with the embossed triangle facing in the same direction as the arrow by the sample holder. Close the lid and then press the zero button. The display should read “595 nm 0.000”. 11. Remove the blank from the spectrophotometer and insert the “0.125” standard cuvette in the same orientation. Give the instrument a few seconds to stabilize after you close the lid and then record the value displayed on the screen in your lab notebook. You should create a data table with one column for protein standard concentrations and one for A595 values. 12. Proceed in this way for the remaining “standard” cuvettes. You should read from the lowest to the highest protein concentration. 13. Once you have finished reading all the standards, follow the same procedures to read the absorbance of each “unknown” sample. You should NOT blank the spectrophotometer a second time. Each group member should record all dilutions and absorbance values. 14. Once you have recorded all your absorbance readings, return to your bench so that another group can use the spectrophotometer. Be sure to take your samples (cuvettes) with you! 15. Show your TA your results and then enter the absorbance readings for the standards (not the unknowns) in the indicated fields on the Bradford assay spreadsheet. DO NOT INCLUDE THE BLANK (0.00). Record the values for slope, Y-intercept, and R2 of the linear trendline in your lab notebook. 16. If the R2 value is below 0.90, you might need to discard one data point or use another group’s standard curve data.* Your TA will advise you on this. * A correlation coefficient (R2) of 1 indicates that all data points lie on a perfectly straight line. The closer the R2 value is to 1, the more reliable the standard curve is. When R2 is less than 0.90, the data are generally considered to be too scattered to permit accurate calculation of the concentration of an unknown protein solution. 17. Examine the absorbance values for your diluted unknowns. At least one of the three dilutions of each cell extract should have an absorbance value that lies between the minimum and maximum values of your standard curve. If not, your TA will advise you on preparing an additional dilution and repeating the assay for that extract. 18. For each fish-muscle cell extract, select the A595 that lies closest to the center of the standard curve and enter it below. Enter the corresponding dilution in the space provided. S extract 1: A595 Dilution S extract 2: A595 Dilution F extract 1: A595 Dilution F extract 2: A595 Dilution

BIOS 312 Cell Biology Laboratory Page 5 of 7 19. Any straight line is described by the equation y = mx + b, where x and y are the coordinates of any point on the line, m is the slope of the line, and b is its y-intercept (the point where the line crosses the y axis.) Your standard curve fits this equation. The independent variable (x) is the concentration of protein (what you controlled.) The dependent variable (y) is absorbance at 595 nm (A595), and the slope and y-intercept of the trendline are provided by the spreadsheet. We can therefore re-write the straight-line equation as: A595 = slope∙(protein concentration) + y-intercept You know the values of A595 for each diluted fish-muscle cell extract. Therefore, you can use the equation above to solve for their corresponding protein concentrations: protein concentration in units of mg/mL = (A595 – y-intercept) / slope Enter the protein concentrations of your diluted extracts below: Diluted S extract 1 = mg/mL Dilution Diluted S extract 2 = mg/mL Dilution Diluted F extract 1 = mg/mL Dilution Diluted F extract 2 = mg/mL Dilution 20. Of course, the concentrations of the diluted extracts are not what we care about, we want to know the protein concentrations of the undiluted extracts. To get these, multiply each of the concentrations above by the appropriate dilution factor, which is simply the reciprocal of the dilution that was used (e.g., 5 for 1/5, 10 for 1/10, etc.) S extract 1: mg/mL x = mg/mL Diluted extract Dilution factor concentration S extract 2: mg/mL x = mg/mL Diluted extract Dilution factor concentration F extract 1: mg/mL x = mg/mL Diluted extract Dilution factor concentration F extract 2: mg/mL x = mg/mL Diluted extract Dilution factor concentration

BIOS 312 Cell Biology Laboratory Page 6 of 7 Calculations for preparation of SDS-PAGE samples This can be done in lab today or on your own. You’ll prepare the actual samples on Day 1 next week, so be sure to have all the calculations finished before then. Now that you know the protein concentration in each of your extracts, you will be able to prepare samples for doing SDS-polyacrylamide gel electrophoresis (SDS-PAGE) analysis next week. Recall that earlier in this handout (page 1,) you were told that the samples should contain 25 μg of protein per 5 μL. What does this translate to in mg/mL? 25 μg / 5 μL = 5 μg/μL = 5 mg/mL Remember that 1 mg is 1000 μg and 1 mL is 1000 μL 1. In the space below, use the formula C1 x V1 = C2 x V2 to determine what volume of each fish-muscle cell extract you would need to dilute to give you 100 μL of solution that contains 5 mg of protein per mL.* Hint: V1 is what you’re trying to solve for. What is C1? C2? V2? Ask your TA for help if you aren’t sure. S extract 1: S extract 2: F extract 1: F extract 2: *If the concentration of your cell extract is < 6.7 mg/mL, solve for a final concentration of 1 mg/mL instead of 5 mg/mL. You will have to load a larger volume of sample onto the gel to get the same total amount of protein, but this can be done.

BIOS 312 Cell Biology Laboratory Page 7 of 7 2. In addition to 5 mg (or 1 mg) of protein per mL, your SDS-PAGE samples need to contain SDS, Tris buffer, reducing agent (beta mercaptoethanol,) tracking dye (bromophenol blue,) and glycerol at the appropriate concentrations. Fortunately for you, we have prepared a stock solution called “4x Laemmli buffer” that contains each of these components at 4x the concentration required in your complete SDS-PAGE samples. Since you are going to be preparing samples that have a total volume of 100 μL, how much of the 4x Laemmli buffer will you need for each one? Calculate this in the space below. Hint: You are trying to dilute the concentrations of the Laemmli buffer components from 4x (C1) to 1x (C2) and you know that your final volume needs to be 100 μL (V2). Again, ask your TA for help if you’re having difficulty. 3. You’re almost done. You’ve determined the volume of each cell extract and the volume of 4x Laemmli buffer that you’ll need, but you need to adjust the total volume of each sample to 100 μL. What will you add? Deionized water (dH2O)! Calculate the amount you’ll need for each sample in the space below. S extract 1 sample: S extract 2 sample: F extract 1 sample: F extract 2 sample: That’s it. Be sure to have your TA check your calculations, either today or next week before you actually prepare your samples for SDS-PAGE. Before you leave the lab, be sure that you: 1.) return your undiluted cell extracts to the designated rack for storage in the freezer, 2.) discard your used microfuge tubes in the regular trash, 3.) empty all cuvettes into the sink with running water, 4.) discard the empty cuvettes in regular trash, 5.) check your spectrophotometer to be sure no one left a cuvette in it, 6.) empty the waste beaker on your bench, 7.) return all materials to the locations you got them from, 8.) straighten up your workspace and wipe it down with 70% EtOH, and 9.) wash your hands.

BIOS 312 Lab Report 1 Guidelines • Explain the purpose of this week’s exercise. (2 pts) • Describe the procedures you followed by referring to the original source of the information (“Protein Quantitation in Unknowns” lab handout) instead of writing out all the steps. If you made any changes, be sure to list them. (2 pts) • Complete a Bradford assay data table that includes the absorbance of each standard and the absorbance of each dilution prepared from your protein extracts. Show your standard curve slope, y-intercept, and R2 value, and your calculations of the protein concentrations in the undiluted extracts. (4 pts) • Summarize your results. Did you get a high protein yield? How do you know? How will you know if your protein extracts were high-quality? Did you notice any differences in the extract appearance or protein yield for the freshwater and saltwater fish? (2 pts)

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