CHE 341 Lab 1 Lesson

Procedures

PART 1: Pipetting

Purpose

Before attempting an experiment that requires precision pipetting (Part 2), every student will first assess their accuracy and precision of your pipetting technique.

Materials

• Pipettors and tips
• A weigh boat on the pan of a digital balance
• Two 50 mL beakers, one filled with water, next to the balance

Procedure (individual)

1. Weigh distilled water that you will dispense into a weigh boat on a balance. Between you and your partner, one will weigh either 70 μL of water with a 10-100 μL pipettor or 700 μL of water with a 100-1000 μL Pipetman.

2. Perform and record three replicate measurements of the mass of water delivered to the balance. Practice forward and reverse pipetting. Be sure to tare the balance between measurements.

3. Ask your instructor or TA to sign-off on the precision and accuracy of your technique before continuing to Part 2.

 

PART 2: Bradford Protein Assay

Purpose

Measure the total protein concentration of a sample of non-fat milk.

Materials (work with a partner)

• Spectrophotometer set to 595 nm
• Bradford reagent purchased from Sigma-Aldrich (be sure to report the manufacturer in the Experimental section of your reports)
• Cuvettes
• Eleven 13 x 100 mm test tubes and test tube rack
• 100 μl and 1000 μl pipettors and pipette tips
• BSA (bovine serum albumin) protein standard (1.0 μg/μL) in a microcentrifuge tube
• Non-fat milk in a microcentrifuge tube

Procedure

1. Warm up the spectrophotometer at 595 nm for least 30 min.

2. Set up 11 clean test tubes to use for the assay. You will get the best results if you mix the reaction in a large test tube and then pour a portion into the cuvette (~3 mL) for measurement.

3. Set up your protocol based on the following table. Copy the table below into your notebook.

 

4. Fill-in the "Calculated Amount of Protein (μg)" column for the eight BSA Standards based-on the concentration of the BSA standard listed above.

5. Choose the ideal sample volume and water volume for Milk Sample A. Each assay in the table should contain 100.0 μL of water plus protein solution (either BSA or milk). Select a milk sample volume that you expect to be within the range of your standard curve (0–100 μg of BSA). Since a Bradford assay standard curve may lose linearity at high concentrations, a low-to-middle protein mass of 1–60 μg should be ideal for your samples. Based on the manufacturer's nutrition label, the milk contains about 30 μg/μL protein. The class will discuss this important decision before lab begins. You may choose to use a different volume in the actual lab. But it is critical that you think about this decision before lab.

6. Using a micropipettor, transfer the appropriate amount of BSA standard into the tubes. For accurate pipetting, make sure the tip touches the bottom of the tube when dispensing. Adding it to the bottom also reduces the risk of contaminating the pipette you will use repeatedly to add the Bradford reagent.

7. Obtain a 1 mL sample of milk. Pipette your chosen volume of the milk sample into the last four tubes and add water so that the sample total is 100 μL.  

8. Add 3 ml of Bradford reagent to each tube. Only touch the tip to the upper part of the inside wall of the tube to avoid cross-contamination by protein as you reuse the same pipette tip. Cover the top of the tube with parafilm or a plastic cap and invert the tube a few times to mix immediately after adding the reagent to each tube. Do not wait until you have added it to all tubes.

9. Let the tubes sit for at least 10 minutes before reading the absorbance. Once the color develops, it is stable for over an hour.

10. Read absorbance at 595 nm. Zero the spectrophotometer on Standard 1. To avoid the need to wash the cuvette between readings: (1) read from lowest to highest protein concentration and (2) decant the solution back into its original tube and then remove as much residue as you can with a Pasteur pipette and discard it onto a paper towel or Kimwipe.

11. In your notebook, make a rough plot to determine which BSA concentrations cause the curve to stray from linearity. If you have a laptop available, you can alternatively plot your data in Excel.

 

PART 3: Make a 50 mM pH 6.5 Buffer

Purpose

Prepare 500 mL of 50 mM pH 6.5 phosphate buffer for future experiments. Observe the effects of ionic strength on the pH of a buffer.

Materials (work with a partner)

• Bottles of monobasic sodium phosphate, NaH₂PO₄ (MW = 119.98 g/mol) or NaH₂PO₄·H₂O (MW = 137.99 g/mol)
• Bottle of solid NaCl
• pH meter
• Clean 600 ml beaker
• 1.0 M NaOH
• 1.0 M HCl

 

Procedure

1. Weigh-out enough monobasic phosphate salt to give the target molarity and volume. Record the calculated mass here in your notebook before lab (calculate for both the anhydrous and the hydrated salt, since either one may be potentially available).

2. Add the accurately weighed mass of phosphate salt to a clean 600 mL beaker, and dissolve to ~80% of your final volume with deionized water (ask you instructor if you are unsure which water to use).

3. Calibrate the pH meter using the pH 4, 7, and 10 buffer solutions. Set up a beaker with your buffer solution on a stir plate (if available) so that you can stir the solution and read the pH continuously. If a stir plate is not available, just swirl the beaker frequently with the electrode while you add acid or base.

4. Use either 1 M NaOH or 1 M HCl to titrate to the desired pH by adding the strong acid or base a drop at a time. By doing this, you effectively change some of the buffer acid form to the conjugate base form, or vice versa, until you achieve the desired ratio of the two species (desired pH).

5. Recheck the pH to make sure it has not changed. If it has, correct it with NaOH or HCl.

6. Bring the volume to the final volume of 500 mL in a volumetric flask.

7. Record the pH of this solution.

8. Remove 10 ml of your 50 mM buffer and dilute it in a volumetric flask to a final volume of 100 ml of deionized water.

9. Record the new pH and the expected concentration of your buffer.

10. Add 416 mg of NaCl.

11. Measure the pH again.

12. Turn-in what remains of your 500 mL buffer to your instructor or TA. You will use this buffer in future labs this quarter.

• Dispose of all solutions in the chemical waste container. Only water should go down the sink.
• When you clean-up, you are responsible for filling the base tubes, acid tubes, and pH calibration vials for the next section.
• Do not discard any solution tubes that you find on your bench in this course. They are meant to be reused.

13. Answser the following questions clearly in your notebook. Your answers will be graded as part of the post-lab assignment grade for Lab 1.

1. Briefly explain why the pH of the buffer changed when you diluted it.
2. Briefly explain why adding NaCl salt restored the pH of the diluted buffer.