Spectrophotometry, Spectroscopy, and Protein Determinations

Lab 1

We are all well aware of the composition of the world -atoms form molecules, compound become more complex, and the organization of these atoms into materials with unique structures is what brings about life. As scientists though, we must study these substances , which presents a challenge. How do we study something so incredibly small? One of the simplest methods is spectrophotometry. Different molecules will interact with light in different ways. By studying this, we can quantitatively say both how much light a compound absorbs as well as what kind of light. Certain functional groups tend to absorb light at certain wavelengths, giving "peaks" to the spectrum of light absorption. This lab demonstrates basic principles of absorbance, measured using spectrophotometers.

Learning Objectives

1. To use the Beer-Lambert equation to relate the concentration and absorbance of a solution,
2. To use a spectrophotometer in the proper manner,
3. To become familiar with the Bradford Assay, and
4. To utilize a standard curve to relate an unknown to established parameters.

Materials

1. Spectrophotometer
2. Micropipettors
3. Parafilm®
4. Solutions of riboflavin SDS, fluorescein SDS, 1.0 M Tris-Cl buffer, pH8*, bovine serum albumin (BSA) SDS
5. Calculator
6. Spreadsheet program (e.g., Excel)

*This buffer is made by titrating Tris base SDS with HCI SDS until the desired pH is reached.

Objective

To become familiar with the Beer-Lambert Law, A=ϵcl, from determinations of the UV-Vis spectra of riboflavin and flourescein; to practice spectroscopic techniques for the quantitative determination of protein concentrations in solution.

Protocol

Part I. Absprption Spectrum of Riboflavin. At the start of the lab, your TA will provide instructions for powering up the spectrophotometer. In addition, you will be told how to turn on the deuterium lamp, which is used for readings in the UV range. Your TA also will give you a solution concentration (μg/mL) of riboflavin.

Table 1.1 Relevant physicochemical data of flourescein and riboflavin. Extinction coefficients are given in terms of M−1cm−1 unless otherwise noted.

From its molecular mass (376g/mol) and measurements of visible and UV spectra, determine λmax and the molar absorption coefficient ϵ(M−1cm−1) in the visible region. To obtain these readings you will set your spectrophotometer to scan from 200 - 600nm. Dilutions may be necessary to make accurate readings in the spectrophotometer. Remember that a conservative estimate of the linear range of a spectrophotometer (at least, our spectrophotometers) is 0.1 - 1.0; readings outside this range generally are not used or are used with caution. If the reading is well above 1.0, dilutions need to be made until the reading is within the linear range. Ideally, you should repeat the process two more times and report the average value of the extinction coefficient (or molar absorption coefficient) along with the standard deviation; however, because of time limitations you will only use the one sample. Remember to clean the cuvettes thoroughly after you are done. Consider the λmax found in the visible region. Does this result make sense, given the color of the riboflavin solution? Comment on this question in the Discussion part of your report.

Figure 1.1 Chemical structure of riboflavin

Part II. Fluorescein. Your TA will give you a solution of fluorescein of unknown concentration and 0.3 mL of 1.0 M Tris⋅Cl, pH 8 buffer. Use a small test tube to make 2 mL of 0.1 M Tris⋅Cl. pH 8 buffer. How are you going to do this? (Hint: remember your old friend: M1V1=M2V2.)

Blank the spectrophotometer with 900μL of the 0.1 M Tris⋅Cl, pH 8 buffer, then add 100 μL of the fluorescein to the cuvette that contains the 900 μL and mix thoroughly by tightly covering the top of the cuvette with Parafilm® and inverting several times. (What is the dilution produced when you do this? Take the Spectrophotometry quiz to check your understanding.) Read this sample, and if the reading is in the appropriate, linear range you can use this reading. If not, adjust your dilution accordingly to get the sample to read. If not, adjust your dilution accordingly to get the sample to read within the linear range. Ideally, you should repeat the process two more times so that you would have three readings, from which you calculate the average and standard deviation; however, due to time constraints, you will use only one sample that reads within the linear range. Considering the molecular weight of fluorescein (332) and its extinction coefficient ( ϵ1mg/mL490=229), determine the concentration of the original, undiluted solution. Remember to take into account the "dilution factor" (What is a dilution factor and how is it determined? Take the Spectrophotometry quiz to check your understanding.) Determine also the ratio of its absorption maxima at 490 nm and 240 nm. From these data, determine the molar absorption coefficient M−1cm−1 of fluorescein at 240 nm. It is recommended that you again scan from 200 - 600nm and use the cursors to move the relevant peaks so that you can see what the absorbance values are at these peaks.

Figure 1.2 Chemical structure of fluorescein

Part III. Bradford Assay. Your TA will give you a solution of bovine serum albumin (BSA; 1 mg/mL). You will make a series of appropriate dilutions and plot a standard curve using the following protocol:

1. Prepare a range of standards (1.00mL volumes in 1.5 mL tubes) containing: 0.2 mg/mL, 0.4 mg/mL, 0.6 mg/mL, and 0.8 mg/mL of BSA. The fifth sample will be undiluted (1.0 mg/mL) BSA. You will use these same samples to construct you standard curve, which is a part of the Bradford assay.
2. Obtain six microcentrifuge tubes (1.5 mL size) and add 990 μL of Bradford reagent to each of these tubes. (Note: Your TA has already diluted the Bradford reagent 1:4.) Next, add 10 μL of dH2O to one of the tubes. This tube will be your blank.
3. Add 10μL of the 0.2 mg/mL BSA sample to a second tube, 10 μL of the 0.4 mg/mL BSA sample to a third tube and so on until you have used all of the BSA samples.
4. Cap each tube and be sure to label these tubes.
5. Mix the samples by gentle inversion of the tubes (about 5 times), and allow the samples to stand for 5 minutes.
6. After 5 minutes, add the blank mixture to one of the cuvettes and zero (or blank) the spectrophotometer at 595 nm. Dispose of the blank solution in the appropriate waste container and then add the lowest-concentration sample to the same cuvette and read the A595. This sample can be disposed as described above, and then add the next sample (in order of increasing concentration) to the same cuvette that held the blank and the first sample. The new sample will be read, disposed of, and the process repeated until the last, highest-concentration sample is read.
7. When you are done, clean the cuvette you used. You will need to use an acetone rinse to remove the residual Bradford reagent from this cuvette. Although this method is not ideal (in a perfect world we would use a clean cuvette for each sample), it should work for our purposes.

Figure 1.3 Coomassie Brilliant Blue G-250, whose absorbance shift is observed in a Bradford assay

Part IV. Standard Curve. Using the data from the solution of BSA (1 mg/mL) that was provided, each student in the group will plot a standard curve of the absorbance readings at 595 nm (A595) for the Bradford reaction of BSA versus protein mass (in μg). Does your standard curve appear linear? How good is the fit? What statistic allows one to estimate the "goodness of fit"? Make sure to save this standard curve as you will use it again later in the semester.

Interactive 1.1 Crystal Structure of Bovine Serum Albumin

Media will be placed here

Shown with bound calcium (green spheres) and acetate ligands. PDB accession number: 3v03

Part V. Protein Samples of Unknown Concentration. Your TA will give you different solutions containing different concentrations of different "unknown" proteins. (We, of course, know the identities and concentrations of these proteins.) Prepare each of these samples for Bradford analysis as described above and read the A595 of each of these unknowns after the appropriate incubation period. Did any of your unknown samples absorb outside the linear range? If so, what should you do to obtain a more accurate estimate of the concentration of this/these particular sample(s)? Make the appropriate adjustment and proceed until all of your readings are within the linear range of the instrument.