ING: Day 2
Decide with your partner how to distribute the effort required to complete the lab experiments. Our recommended distribution is to have one partner be responsible for setting up the electrophoresis experiment and preparing the electrophoresis samples, while the other can begin setting up the enzyme assays. Once the gel analysis is underway, both partners should work together to complete the remainder of the experimental work in the allotted time.
Enzyme Assay
Mix 0.5 mL of toluenized culture with 1.5 mL of p-nitrophenyl galactoside (PNPG) (1.0 mg/mL) in 50 mM sodium phosphate buffer, pH 7.0. Incubate the reaction mixture at room temperature for 10 minutes; then quench the reaction by adding 1.0 mL of 1 M Na2CO3. Read the absorbance at 400 nm versus a suitable substrate blank (see instructions for EZA lab). If the absorbance exceeds a value of 1.0, repeat the assay using an appropriate dilution of the sample. Record the absorbance values and dilution factors, if required, on DATA SHEET II.
Gel Electrophoresis
Click Here for Gel Standards
Materials
The formulation described below is for 12% polyacrylamide gels. In combination with a Tris/glycine buffer system, these gels can be used to provide rapid separation of polypeptides from 20 to 80 kDa. For separation of proteins from 1 to 100 kDa, a Tris/Tricine buffer system provides much better results, although the time required for electrophoresis is about 50% longer (Schäger, H. and von Jagow, G. (1987). “Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa.” Anal. Biochem., 166, 368-379).
AND IS HIGHLY TOXIC TO THE UNBROKEN SKIN. WEAR DISPOSABLE GLOVES WHILE HANDLING UNPOLYMERIZED ACRYLAMIDE SOLUTIONS. NEVER MOUTH-PIPETTE THESE SOLUTIONS! DO NOT HANDLE UNCONTAMINATED EQUIPMENT WHILE WEARING CONTAMINATED GLOVES. FOLLOW ALL INSTRUCTIONS REGARDING THE DISPOSAL OF CONTAMINATED PIPETTES, GLOVES, ETC.
- 10% SDS Solution: 10 g of SDS dissolved in a final volume of 100 mL of deionized water;
- 30% Acrylamide Solution: Dilute 75 mL of 40% acrylamide to a final volume of 100 mL with deionized water;
- 10% Ammonium Persulfate Solution: 0.1 g of ammonium persulfate diluted to a final volume of 1 mL with deionized water;
- TEMED: use as a neat solution;
- Resolving Gel Buffer: 24.2 g of Tris Base diluted to a final volume of 100 mL with deionized water, and carefully adjusted to pH 8.8. Add 0.5 g of SDS for denaturing gels;
- Sample buffer: 5 mL of Stacking buffer containing SDS, 50 μL of β-mercaptoethanol, a few specks of bromophenol blue diluted to a final volume of 10 mL with 85% glycerol;
- 10X Electrode buffer: 288 g of glycine free base, 60 g of Tris Base diluted to a final volume of 2000 mL with deionized water. This buffer should be pH 8.5, and normally should not be adjusted;
- Staining solution: 1000 mL of methanol and 200 mL of glacial acetic acid diluted to a final volume of 2000 mL with deionized water. Add 1.86 g of Coomassie Blue R;
- Destaining solution: 300 mL of methanol and 100 mL of glacial acetic acid diluted to a final volume of 1000 mL with deionized water.
Procedure: Day 2
One person from each pair should work to prepare the gel while the other partner performs the β-galactosidase assays. In order to complete this laboratory in the allotted time, you will have to work efficiently. Carefully review the following procedures before coming to lab, so that you will be familiar with the order of preparations required.
1. Preparation of the resolving gel
1.1. Put on your gloves.
1.2. Assemble a gel apparatus, as demonstrated by the Teaching Assistant in the pre-lab lecture.
1.3. Check for leaks by filling the apparatus with water, and have a Teaching Assistant check it.
1.4. Add 1.0 mL of the Resolving Gel Buffer, and 2.0 mL of deionized water to the 2.0 mL of acrylamide/TEMED solution provided in a scintillation vial labeled “R.”
1.5. Add 25 μL of the 10% Ammonium Persulfate Solution to the acrylamide vial.
1.6. Gently mix the solution by swirling the flask. Try to avoid foaming the SDS or mixing air bubbles into the solution.
1.7. Using a long-tipped Pasteur pipette, immediately transfer the solution into the gel apparatus. Fill the unit to ~2 mm below the notches in the front face of the gel apparatus.
1.8. Using the same Pasteur pipette, gently layer some deionized water on top of the acrylamide solution. This will help to form a smooth polyacrylamide surface, on which to pour the stacking gel. Do not move the gel until polymerization has occurred. Watch for the appearance of a visible separation between the water and the gel. While the polyacrylamide gel is polymerizing, prepare the bacterial cell lysates. You will have about 30 minutes to complete this task.
2. Preparation of the bacterial cell lysates
2.1. Place the labeled microfuge tubes from the first day of the experiment in a 95-97°C water bath for 20 min to lyse the bacterial cells.
2.2. Remove the microcentrifuge tubes from the water bath. Allow to cool and spin in a microcentrifuge for 2 minutes to pellet any remaining undissolved material.
3. Preparation of the stacking gel
3.1. Put on your gloves.
3.2. Carefully invert the gel apparatus to pour off the water layer. Remove any water remaining from the corners using a piece of paper towel.
3.3. Get a microfuge tube, labeled “S,” containing 0.15 mL of an acrylamide/TEMED solution, 0.3 mL of the Stacking Buffer, and 0.8 mL of deionized water.
3.4. Add 20 μL of the 10% Ammonium Persulfate Solution and mix carefully.
3.5. Close the cap and gently mix the solution by inversion.
3.6. Using a long-tipped Pasteur pipette, immediately transfer the solution into the gel apparatus. Add acrylamide nearly to the top of the glass plate.
3.7. Carefully insert a comb into the acrylamide in the gel apparatus. Try to avoid splashing the acrylamide when the comb is inserted. Center the comb as necessary between the edges of the gel.
3.8. Examine the stacking gel for the presence of bubbles, and remove them as necessary.
3.9. Wait for the stacking gel to polymerize. The stacking gel should polymerize within 30 min.
4. Loading the sample wells
4.1. Obtain 80 mL of 10X Electrode Buffer. Dilute it to 1X and use that in steps that follow.
4.2. With a grease pencil, draw on the glass plate around the teeth of the comb. Then remove the comb from the polymerized stacking gel. Fill the sample wells with the diluted Electrode Buffer to displace the bubbles.
4.3. Load molecular weight standards in two lanes of each gel. Load the standards asymmetrically (for example, in lanes 1 and 9) so that you can distinguish between the two sides of gel once it is removed from the plates. Also load the standards differently in the two gels (for example, lanes 1 and 9 for gel 1 and lanes 1 and 8 for gel 2) so that you can distinguish between the two gels. Load 10 μL for one of the standards and 5 μL for the second standard.
4.4. In the first gel, load 10 μL of the Control samples from 0, 50, 90, and 110 minutes into the 1st, 2nd, 3rd, and 4th open lanes. Load 10 μL of the Antibiotic samples from 0, 50, 90, and 110 minutes into the 5th, 6th, 7th, and 8th open lanes. In the second gel, load 10 μL of the IPTG samples from 0, 50, 90, and 110 minutes into the 1st, 2nd, 3rd, and 4th open lanes. Load 10 μL of the Lactose samples from 0, 50, 90, and 110 minutes into the 5th, 6th, 7th, and 8th open lanes.
4.5. Attach the gel assembly to the electrode assembly.
4.6. Place the electrode assembly into the electrode chamber. Fill the inner chamber.
4.7. Add enough Electrode buffer to fill the lower electrode chamber up at least above the electrode wires.
5. Applying the voltage
5.1. Place the cover on the electrophoresis unit. Be sure to connect each plug to the like-colored socket.
5.2. Turn on the power supply and run at a constant voltage of 200 V.
5.3. Continue the electrophoresis until the dye front is within 1 cm of the bottom of the gel. The electrophoresis should take about 45 min to complete.
5.4. Turn off the power and unplug the leads from the power supply. Remove the cover.
6. Staining and destaining the gel
6.1. Put on your gloves.
6.2. Unclamp the gel from the apparatus. Slide the Teflon spacers out of the gel sandwich. With a spatula or one of the Teflon spaces, carefully pry apart the two gel plates. Apply pressure only at the middle of the gel sandwich, NEVER near the “corners” as they will break off. The gel should adhere to one of the glass plates. Cut off the stacking gel. Nick one corner of one of the two Resolving Gels your group has run. Both gels from your group will share a staining box, so this nick will tell you which gel is which -- even if your standards don’t stain well enough to differentiate the gels, based on their different lanes of loading.
6.3. Transfer the gel from the glass plate into a staining box by inverting the plate with the gel over the box and squirting water along the edge of the gel until it comes loose and slides into the dish. Never try to move the gel unless it is immersed in liquid; it will tear. If the force of the water from the squeeze bottle doesn’t remove the gel from the plate, it may be necessary to run a flat spatula along the sides of the gel.
6.4. Holding the gel in place, pour off the water. With a spatula, cut away a 1 cm wide piece of the gel from the right bottom corner. You should place the top of this cut to mark the position of the dye front. This is essential in order to be able to determine Rf values!
6.5. Cover the gel with Coomassie Blue staining solution (about 125 mL).
6.6. After at least 15 minutes, rinse your gel several times with deionized water. Then cover the gel with destain. Crumple a Kimwipe, and saturate it with destain in your dish. Be sure the Kimwipe isn’t touching your gel. The Kimwipe helps pull the stain out of your gel by absorbing the dye out of the destaining liquid. Place on the shaker overnight.
7. Clean up your gel apparatus, glass plates, and work area.
8. Mount your gels in cellophane and dry (TA will demonstrate).
8.1. Follow the instruction sheets that are hanging in the lab for mounting and drying your gel.
Report
Check the lab manual and class website for detailed instructions and late-breaking tips on how to prepare the lab report. Prepare a cover sheet for your lab report containing your name, section, experiment title, and name of lab partner. Include the complete DATA SHEETS I and II. The dried gel can be photocopied. Prepare plots or answers as indicated below.
β-galactosidase Data Sheet
[PDF] or [Word doc]
Induction Experiments
1. From DATA SHEET I, make a single plot showing the growth curves for each of your three cultures. Clearly label the axes, scales, and units for each growth curve.
2. Prepare a plot of ln[A410/A0] = t. From this plot, compute the doubling time of your cultures and indicate the values in a Figure caption describing your plots. If you think the calculation of doubling time is inappropriate for any of your cultures, state your reasons for believing so in the Figure caption.
3. Assume the molar absorptivity of the p-nitrophenolate anion (MW 138) is 14,000 M-1cm-1 at 400 nm, that the pKa is 7.15 and that the absorbance of the uncleaved substrate is negligible at 400 nm. Calculate umol of p-nitrophenolate produced and enter these values in the appropriate place in DATA SHEET II. Convert these values to I.U. per mL of culture and I.U. per 108 cells, and enter these values in DATA SHEET II.
4. Using DATA SHEET II, prepare one plot showing the enzyme activity versus incubation time. Scale the left axis to show I.U. per mL of culture; scale the right axis to show I.U. per 108 cells. Hint: see the web site for a description of how to make overlay plots using Excel.
5. Provide a short, written comparison of the effect of chloramphenicol cell growth and β-galactosidase activity. Express your answer in quantitative terms, using the doubling times and the specific activity data from your experiments.
Electrophoresis Experiments
1. Attach your dried electrophoresis gel (or a photocopy) to a sheet of white, unlined paper. Indicate your partner, and which report contains the original gel. Indicate the following:
a. The identity of the sample in each lane and the loading volume;
b. The electrode polarity (positions and charge of anode and cathode);
c. The direction of migration of proteins, Na+, and Cl-;
d. The position of the top of the resolving gel; and
e. The position of the dye front.
2. For the molecular mass standards, prepare a Table containing log (molecular mass) and relative mobility, where
|
relative mobility = Rf = |
distance of the protein band from the resolving gel top |
|
distance of dye front from resolving gel top
|
3. Make a molecular mass calibration curve for the standards. Provide a linear least squares fit analysis of the data, and plot the best-fit line. On your graph, provide a 1-sentence answer describing the range of molecular masses that you believe have a linear relationship with the determined Rf-values.
From your comparative examination of the bacterial cell lysates, can you identify any induced protein bands? If so, clearly label these bands on the page containing your gel, calculate Rf-values for these bands, and enter these values in the table you have prepared for the molecular weight standards.
From the relative mobility of the induced polypeptide(s) you have identified on your gel, determine the molecular weight(s) of these protein(s). Enter these molecular weights into your table of standard values. How confident are you in the accuracy of the molecular weights obtained for the unknown proteins?
4. What polypeptide(s) do you think might correspond to the induced proteins you have identified? Give reasons that support your choice.
Would some of the samples have been better run on a different kind of gel? If so, what modifications would you make to the protocol to achieve better accuracy?
Is there a difference in the accuracy obtained for lanes adjacent to the spacer and lanes in the middle of the gel?
5. Can a polypeptide corresponding to β-galactosidase be identified in your denaturing gel? If so, mark this band on the gel. If not, why not?