User:Douglas M. Fox/Notebook/AU CHEM-571 F2011 Lab Support/2017/09/13

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Group Assignment

Group 1: Casey, Pauline, & Sanuja

  1. Bradford Analysis (UV/VIS)
  2. Dialysis
  3. Fluorescence
  4. Electrophoresis

Group 2: Anh, Dominique, Mackenzie, & Michael

  1. Dialysis
  2. Fluorescence
  3. Electrophoresis
  4. Bradford Analysis (UV/VIS)

Group 3: J.I., Kaiyu, Marjan, & Rachel

  1. Fluorescence
  2. Electrophoresis
  3. Bradford Analysis (UV/VIS)
  4. Dialysis

Group 4: Ashish, Maryam, Massie, & Noy

  1. Electrophoresis
  2. Bradford Analysis (UV/VIS)
  3. Dialysis
  4. Fluorescence

Bradford Analysis

The most direct method for measuring the protein concentration is the use of the Beer - Lambert Law, using published extinction coefficients (molar absorptivities) for the proteins at λ = 280 nm in a UV-VIS spectrum. For low concentrations of proteins, UV-VIS of just the protein is often not sensitive enough to accurately measure concentration. (The limit of detection is about 2 - 3 μM for most proteins.) During the semester, we will likely need to measure protein concentrations that are lower than this. In addition, molar masses and/or extinction coefficients of some proteins are not well quantified. One tool we have can use to measure protein concentrations on the μg/mL level is called the Bradford Assay. The Bradford Assay makes use of the Coomassie Blue dye, which binds to proteins. Upon binding to a protein, this dye undergoes a change in its absorption features. (No protein: peak at 460. Protein: peak at around 600). The dye binds to the basic and aromatic residues of the protein. We will be making calibration curves (using the Bradford Assay) for the different proteins we'll be using throughout the semester. Since this method depends on the number of peptide bonds, concentrations are reported by mass and the method is fairly independent of the particular protein being measured. There are a few interferences, such as co-factors that absorb near λ = 600 nm (e.g. hemes) or basic pH buffers.

The basic protocol can be found here (*Note: use section 2.3, page 5) or here.

  1. Prepare 50 mL of a stock solution of BSA that is roughly 5 mg in 5 mL of saline.
  2. Calculate your actual solution concentration.
  3. Using a quartz cuvette, record UV-VIS spectra between 200 nm and 800 nm.
    • remember to record UV-VIS spectrum for saline.
  4. Make 6 - 8 standard solutions (1 mL each) between 1 μg/mL and 20 μg/mL. It may be appropriate to use a serial dilution.
    • Determine the appropriate volume of stock solution to use and add it to a 1.5 mL centrifuge tube.
    • Add 200 μL of the Bio-Rad Protein Assay reagent. Use 1:4 concentrate diluted with water.
    • Add the correct amount of Tris/NaCl buffer such that the final volume is 1 mL.
    • Close the tubes and vortex them for 5 - 10 sec.
    • Let them sit for 5 min.
  5. Obtain a UV-VIS spectrum.
    • solutions must be measured within 1 hr of their preparation.
    • use PS cuvettes.
    • record between 400 nm and 800 nm
  6. Make duplicate blanks (4 solutions total) as well
    • 1 mL Tris/NaCl buffer
    • 200 μL Bradford reagent + 800 μL buffer
    • record their UV-VIS spectra between 400 nm and 800 nm
  7. Discard solutions in waste bottle and PS cuvettes in the tub or beaker, both in the fume hood.

Coordination will be helpful here. Record one spectrum at a time and allow other groups to measure a spectrum in between.

Data Analysis
First, you will need to use a difference spectra to construct a calibration curve. Align the baseline of all your spectra. There should be relatively little change in absorbance at 800 nm. Shift all spectra vertically so that they have the same absorbance at 800 nm. Then, find the average absorbance spectra for your Bradford Reagent control. Subtract these average values from each of your calibration and sample spectra. You should have negative values centered around 468 nm and positive values centered around 593 nm. You will use the absorbance differences at 593 nm to plot a calibration curve using typical Beer's Law methods.

Second, you will want to find the purity of your protein solution. Using the UV-VIS spectra of your stock solutions, calculate the concentrations of your solutions in both molarity (M or μM) and g/L. The extinction coefficient for BSA is 38,940 M-1cm-1 (λ = 279 nm). The purity of the lyophilized powder you used is [UV measured]/[mass measured].

The Bradford reagent has peaks at 460 nm and 630 nm. The Bradford-protein complex has a peak around 600 nm. There will be significant overlap, which will need to be resolved. The best method is to plot a difference spectrum. See an example in my 09/08/2014 notebook entry. Using the peaks of the difference spectra at 600 nm (594 nm in my plot) and the final protein concentration corrected for protein impurities, construct a calibration curve for your two sets of standards. You should find that these curves overlay each other.

Determine the minimum concentration (in μM) and maximum concentration (in μM) that you can measure using this analysis for each protein. The minimum occurs at a difference absorbance of 0.05 and the maximum occurs at a difference absorbance of 1.0. You should find that the concentration limits are different despite the same calibration curves.

Determine Concentration of your solutions

  1. Dilute your BSA and original AA stock solutions to the appropriate range.
  2. Use Bradford Analysis to determine the concentration.
    • Does the dye bind to free amino acids?
  3. Heat your BSA solution using a block heater to denature (unfold) the protein. Use Bradford Analysis.
    • Place 1 - 2 mL of solution in microcentrifuge and place in 95°C heating block for 5 min.
    • Does thermal denaturing change the concentration measurement?
  4. Try adding some SDS or guanidinium chloride
    • Prepare 10 mL of 2% SDS and 10 mL 2% GuanCl
    • Add equal mixtures of SDS and protein. Use Bradford analysis.
    • Add equal mixtures of GuanCl and Protein. Use Bradford Analysis.
    • Do these salts interfere with the analysis?
  5. If you have time, try a sample of poly(l-lysine)
    • 0.1 wt% solution is in refridgerator
    • Dilute to 10 μg/mL and measure Bradford Analysis

Many thanks go to Prof. Hartings, who wrote the original protocol for this class.


Dialysis is used to purify large molecules by using a membrane with small pores, allowing small molecules to freely move through while preventing large molecules from transferring. We will use dialysis to determine (a) how big the AuNP complexes are, (b) how strongly the stabilizer is bound to the AuNP, (c) if we can change the stabilizer, and (d) how quickly the molecules migrate through the tubing.

  1. Stabilizer exchange
    • Cut 2 8cm – 10 cm lengths of 3500 MWCO dialysis tubing & soak in DI water for 10 min.
    • Remove 1, clamp one end, open other end and transfer exactly 3 mL DI water into tubing. Clamp other end.
    • Place in 125 mL wide mouth jar, add exactly 100 mL 25 mM HdMe2ImCl and screw on lid. Record time.
    • Remove 2nd tubing, clamp end, transfer exactly 3 mL AuNP stabilized with basic AA. Clamp other end.
    • When 15 minutes after recorded time has past, place in 2nd 125 mL jar, add exactly 100 mL 25 mM HdMe2ImCl and screw on lid. Record time.
    • Every 30 minutes, remove exactly 1 mL of solution from jar and measure the absorbance at 211 nm using the spectrophotometer (single wavelength spectrometer). Note that you will be recording a value every 15 minutes.
    • At some point, you will need to measure the absorbance of the HdMe2ImCl stock solution and your AuNP solution.
    • At end of day, cap jar and leave solution until the next session.
  2. Other exchanges – perform in between UV measurements
    • Using same procedure to prepare dialysis tubes, prepare a total of 4 dialysis tubes.
    • Dialysis 3 mL AuNP stabilized with basic AA against Glycine buffer
    • Dialysis 3 mL AuNP stabilized with basic AA against water
    • Dialysis 3 mL AuNP stabilized with BSA against Glycine buffer
    • Dialysis 3 mL AuNP stabilized with BSA against water
    • Dialyze until your next session
  3. During the next session, you will record the UV (using spectrometer) of solutions inside and outside of each jar. If the UV spectrometer is too busy, you can store portions of the solutions in a centrifuge tube.


The stabilized AuNP colloids exhibit a plasmon resonance, which manifests itself with an absorbance in the range of 515 nm to 530 nm, depending on the size of the metal nanoparticle. This results in the observed absorbance peak and red color. Literature suggests that excitation of AuNPs at 308 nm will result in a photoluminescence band between 420 nm and 450 nm, caused by excitation of this same surface plasmon. We want to explore the nature of this phenomenon and determine if it can be used in electron transfer applications, such as FRET.

For fluorescence measurements, use excitation and emission windows of 8 nm and a scan peed of 50 nm / min. Start your emission scan at λex + 10 nm and scan a range of 200 nm.

  1. Prepare dilute (1 – 10 μM) solutions of acridine orange and fluorescein.
  2. Measure fluorescence of your stock solutions using excitation wavelengths of 308 nm, 440 nm, and 490 nm. You may need to dilute your stock solutions to record peaks below 1000 intensity. Make sure you note the dilutions required.
  3. Measure fluorescence of your AuNP using each AA and BSA. Use AuNP formed from the same concentration of HAuCl4 at all three excitation wavelengths. Then, measure the fluorescence of the more concentrated AuNP using the Au:AA = 10:1
  4. Choose any one of the AuNP solutions that had a photoluminescence band around 440 nm
    • Combine with acridine orange (at a concentration equal to one used in step 2) and excite at 308 nm. Measure the fluorescence up to 600 nm. Is the emission band associated with acridine orange visible?
    • Combine with fluorescein (at a concentration equal to one used in step 2) and excite at 490 nm. Measure the fluorescence up to 700 nm. Is the emission band associated with fluorescein quenched?
  5. If you have time, try with another AuNP solution that had slightly different fluorescence spectra


We will be using a Bio-Rad Mini Protean system with pre-cast Mini Protean TGX gels. The manual for this system can be found here We will be running SDS-PAGE followed by gel development with Coomassie Blue staining.

Below is a short description of how we will proceed. Please refer to the manual for more detailed instructions

  1. Prepare the Gel and Assemble the Electrophoresis Cell
    1. Remove comb and tape from the gels
    2. Rinse the wells with running buffer
    3. Assemble the electrophoresis cell (note diagrams in manual)
    4. Fill the inner and outer buffer chambers with running buffer
  2. Prepare and Load Samples
    1. use microcentrifuge tubes to prepare solutions below
    2. Take 10uL of your BSA stock and dilute to 1mL with the Glycine-HCl buffer
    3. Take 10uL of this diluted sample and mix with 10uL of the SDS-PAGE running buffer
    4. Repeat for a single AA solution (you choose), your AuNP from BSA and your AuNP from your chosen AA
    5. Take 100 μL of 0.1 wt% poly(l-lysine) from fridge and dilute to 1mL with the Glycine-HCl buffer
    6. Heat your samples for 5 minutes at 95C (in the thermocycler)
    7. Load 20uL of protein ladder into column 1 of your gel
    8. Load 20uL of your samples into the appropriate lane of your gel
  3. Perform electrophoresis
    1. Run 30 min at 200 V, 0.05 A, and 10 W
    2. While experiment is running, prepare solutions below.
  4. Develop/Stain your gel
    1. Prepare 100 mL of each solution
    2. Place gel in Fixative Solution (40% methanol, 10% acetic acid, 50% water) for 30 minutes
    3. Place gel in Stain Solution (0.025% (w/v) Coomassie Blue, 10% acetic acid, 90% water) for 1 hour
    4. Place gel in Destain Solution (10% acetic acid, 90% water) for 15 minutes
      • Repeat this step with fresh destain solution 2 more times