Measuring Protein in Biological Samples

BIO 330 Cell Biology Name ______

Spring 2017

Laboratory

Measuring Protein in Biological Samples

Measurements are often made of certain protein levels or mRNA levels in biological cell, tissue, or organ samples. However, in order to account for the amount of starting material, which can vary greatly, the end results are usually “normalized” per amount of total protein, total RNA, or total DNA. For example, in Dr. DuPriest’s research on the enzyme chymase, her results were expressed in terms of enzyme activity per mg protein used in the initial reaction.

To measure proteins, researchers typically compare their samples to a standard curve. A standard curve is something a researcher constructs using increasing KNOWN amounts of material (protein in this case). Then, after figuring out an equation that describes the relationship between protein concentration and some observed outcome (often a color concentration) obtained from the known “standards”, we can use the standard curve equation to figure out the concentration of our UNKNOWN samples.

In this activity, we will prepare a crude extract of chicken liver. We will then measure the concentrations and amounts of protein in that extract. To do that, we will need to construct a standard curve of pure protein (bovine serum albumin, BSA). Both our samples (the liver homogenate) and standards (BSA) will be read by spectrophotometry according to the protocol given below.

Protocol

1. Prepare the crude liver homogenate.

The whole class will prepare a single homogenate of chicken liver using the blender; each group of students will receive an aliquot of the homogenate to work with. It is important to keep the liver and homogenate on ice at all times, except when you MUST remove it from ice for a specific reason. The class should obtain a large piece of liver. Using a kitchen knife and cutting board, cut the liver into cubes, and measure out about ~25g (+/-2. 5 g). At the same time that some people are weighing and cutting the liver, others should measure out the appropriate amount of homogenization buffer. (Homogenization buffer contains NaCl to match salt levels in tissue; detergent to split open cells; pH buffer to keep pH around 7.4 and some other minor components.) The amount of buffer should approximately match 10X the weight of liver. (E.g., ~ 250 mL of buffer for 25g of liver). The buffer should be kept on ice until ready for homogenization. When both liver and buffer are ready, combine them in the pre-chilled blender. Put the lid on securely, and start the blender on low level for 5-10 seconds, then switch to high power for two more 10-second bursts. Pour into a pre-chilled Erlenmeyer flask and keep on ice.

Pour the homogenate into four pre-chilled 50-mL centrifuge tubes; fill the tubes ~ 2/3 full. The tubes must be weighed to ensure they are balanced. Eyeball the one that appears to be fullest; weigh it first and record its weight. Then measure the next ones in succession, adding more homogenate until it weighs the same as the first one, within 0.1g. Place the tubes on ice while you are measuring the others. Load the centrifuge tubes into the rotor in the pre-chilled table-top centrifuge. Centrifuge at ~1500g for 10 minutes. Carefully remove the tubes, and place back on ice, being careful not to jar the samples and remix them. (Make sure to turn the centrifuge off when you are finished.)

Each group should obtain one of the centrifuge tubes of liver material. Transfer 1mL of the supernatant (top layer now cleared of debris) to a new microcentrifuge tube (avoid disturbing the pellet), and place the tubes on ice. The pellet is now waste material and can be discarded. The supernatant will now be referred to as your “liver sample”. This type of sample can be used in a wide variety of applications, such as measuring enzyme activity, or to measure specific protein levels. In this activity, we will simply measure the concentrations of protein and DNA in the extracts. However, before we do that, we must prepare our protein standards.

2. Prepare your protein standards. Protein standards are simply protein solutions of known concentration. You will be making a four-point standard curve. You will start with the most concentrated standard (the stock solution, which comes as 1.52 mg/mL protein) first, then make serial dilutions to obtain your other concentrations. Transfer 50 mL of the stock solution to a microcentrifuge tube, and add 50 mL of water to the tube. Calculate the concentration of this solution, and mix thoroughly (you MUST mix completely between each step in order to obtain a linear curve). Then take 50 mL of that solution, transfer to another tube, and add 50 mL of water and mix as before. Again, calculate the concentration. Repeat this again to make a total of four solutions, including the stock solution. Label each tube clearly with the concentration you have calculated – keep track of the appropriate units. Set aside. You will have to perform an assay on these standards prior to measuring protein concentration.

3. Perform your protein assays and read output using spectrophotometry. To measure protein concentration in a biological sample (or standard), we perform a Bradford assay in which we combine our liver sample (or standard) with a dye that changes color when it binds with certain amino acids present in nearly every protein. First, you will need to make 2 separate dilutions of your sample in case your sample is too concentrated for your standard curve. Make these dilutions BEFORE putting sample into your spectrophotometer cuvettes. To make a 1:10 dilution in a microcentrifuge tube, combine one part protein with 9 parts water; to make a 1:50 dilution, combine one part protein with 49 parts buffer. (You can do the calculations yourself – make at least 100 mL total volume for each dilution).

After you have made your sample dilutions, set up your spectrophotometer cuvettes. Obtain two plastic rectangular spectrophotometer cuvettes for each concentration of your standards (8 cuvettes), plus two for each concentration of your sample (1:10, 1:50; 4 cuvettes); and one for a blank (total of 13). Don’t write on them, but keep them organized so you know which is which.

Fill your blank with 1 mL of water. Set aside. For the remaining 12 cuvettes, add 985 uL of protein assay reagent to each cuvette. You cannot label these cuvettes, but keep them lined up so you are sure of which one is which. Once all 12 cuvettes are prepared to this point, and the samples and standards are all ready, add 15 uL of your protein standard or sample. Quickly, but carefully, mix each sample: use a piece of Parafilm to cover each cuvette, seal using your index finger and invert 3-4 times to mix. Try to complete all of these within about 2 minutes. Let all the reactions incubate at room temperature for 15-30 minutes, then read on the spectrophotometer. You should read the absorbance at 595 nm, which corresponds to the blue color that develops when the red dye combines with protein. You should be able to see that the protein standards that contain high amounts of protein contain more blue color, while the standards with low concentrations contain less blue color. Open LoggerPro on the laptop. Select Experiment, then Calibrate, then Spectrometer 1 to calibrate the spectrophotometer. Allow the machine to warm up. During warm-up, insert your blank cuvette (the one with 1mL of water) into the spectrophotometer. Check with the instructor for proper orientation. After warmup is complete, click Finish Calibration, then OK. You are done with the blank, but keep it nearby in case something happens and you need to re-calibrate. Then, one at a time, read your samples. Place a cuvette in the spectrophotometer, click the green Collect button, then Stop (wait just 2-3 seconds). Scroll down the table on the left to where the closest wavelength to 595 is, and record the corresponding absorbance value. Repeat for each sample (no need to save runs on the computer). Record each sample’s reading in Table 1. After collecting your data on all standards, calculate what the concentration and amount of protein is in each of your samples. To do this, you will need to use Excel to construct your standard curve, putting your averaged protein concentration on the x-axis and the A595 values on the y-axis. The resulting plot should be linear. After you plot your points, right-click on any point, and select Insert Trendline. Make sure to select the options to show r-squared value and show equation on chart. Using the equation (you will need to rearrange it using algebra), plug in your averaged A595 value for your liver extract as y and solve for x to determine the concentration of protein in your liver extract. This concentration will be in the same units as you calculated for your standards. Do this for each of your sample dilutions as well as your undiluted sample; multiply your reading by your dilution factor (10 or 50) to get your final calculated protein concentration. You should get the same results after adjusting for dilution factor, but often, you will not. You will explain why this might be in one of the Discussion questions. You will also need to print out your Excel standard curve figure and attach to this packet.

Data

Table 1: Protein Standard Curve

Protein Concentration (mg/mL) / A595
Replicate 1 / A595
Replicate 2 / A595
Average
1.52
0.76
0.38
0.19

Linear equation relating protein concentration with A595: ______

Table 2: Liver Sample Protein Concentrations

Protein Concentration (mg/mL) / A595
Replicate 1 / A595
Replicate 2 / A595
Average / Raw
[Protein] / Corrected
[Protein]
1:10 dilution sample
1:50 dilution sample

Discussion

It is highly unlikely the two results you got for Corrected Protein Concentration were identical numbers. Using your best judgment, determine whether they are basically the same (within reasonable error), or whether there might be a reason that the values are different. If different, explain why they might be different.

______

Why are standard curves used? Do you have to do a new one every time you do an assay (measurement)? Why or why not? ______

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