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Lab Day 7:
Part I – Blue/White Cloning of DNA Fragment and Assay of β–
galactosidase Module II cont.
Count the CFUs on both of your plates and determine the transformation efficiency of your plasmid
DNA. Assume that 25 ng of DNA was used in each transformation reaction.
Additional questions to answer in your analysis (include in your notebook):
1. Did your transformations work for both plates?
2. If there are no colonies on one or both of your transformation plates, what could be the reason
that the transformation did not work? Include this analysis in your lab notebook.
3. What additional controls could you have done alongside this experiment?
4. Assuming the control plate had 87 colonies and the Calculate the transformation efficiency.
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Part II – Blue/White Cloning of DNA Fragment and Assay of β–
galactosidase Module III:
In a previous lab session, you ligated a linearized pUC8 plasmid with a DNA fragment (the ‘insert”).
Following the ligation reaction, the vector was then inserted into an E. coli JM109 host strain by
transformation. As discussed in the previous handouts, a ligated vector without an insert (“self” ligated)
will have an uninterrupted lacZ’ gene and therefore a functional lacZα subunit will be transcribed. Thus,
colonies with a plasmid that has no insert in the multiple cloning site will have a functional β–
galactosidase enzyme. Cells with a functional β–galactosidase enzyme (Lac+ transformants) that are
grown on a plate with the X-gal substrate will be blue. However, if a fragment was inserted in the lacZ’
gene, then a functional lacZα subunit will not be made, resulting in a lack of β–galactosidase enzyme
activity, hence colonies will be white (Lac- transformants).
In today’s lab, you will grow Lac+ and Lac- E. coli cultures and assay for β–galactosidase enzyme
activity. β–galactosidase enzyme activity assays will be performed using 4-orthonitrophenalgalactopyranoside (ONPG) as a substrate. Recall that upon catalysis by β–galactosidase, ONPG will form a
yellow color.
Lac+ cells (blue colonies) will have a functional β-galactosidase enzyme and therefore will hydrolyze
ONPG, hence a yellow color will be observed. Lac- cells (white colonies) do not have a functional βgalactosidase enzyme and therefore will not hydrolyze ONPG, hence no yellow color will be observed.
pUC8 without insert  β-galactosidase is expressed  blue colonies  yellow observed
pUC8 with insert  β-galactosidase is not expressed  white colonies  no yellow observed
Follow Module III directions below for performing β-galactosidase enzyme assays.
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Record your results in your notebook as follows:
Additional Questions:
1. Do all the white and blue colonies in the LB/X-gal/Amp plates contain a plasmid?
2. Will lysozyme used to lyse cells denature β-galactosidase?
3. Which restriction enzyme is best suited for cloning pUC8? (Refer to previous handouts for pUC8
map and multiple cloning site).
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Part III – Making Electrocompetent Cells
In the previous lab sessions you made competent E. coli cells using two methods; 1) by harvesting cells
from an LB plate and treating them with cold CaCl2 and 2) using a modified CaCl2 method by growing
cells to an OD 0.4-0.6 and performing a series of washes with TFBI and TFBII solutions containing
RbCl2. These are called “chemically competent” cells. In today’s lab you will make “electrocompetent”
E. coli cells. Electrocompetent cells are made by a series of washes to remove any salts which can
increase the conductivity of the cell suspension (the success of electroporating cells depends on a cell
suspension with low conductivity).
1. Inoculate 25 ml of fresh LB with 1 ml of overnight culture.
2. Grow cells at 37 ºC on a shaker at 220 rpm until the Optical Density at λ600 (OD600) = 0.4 - 0.6
(Should take 2-3 h).
3. Centrifuge the cells at 3000 rpm for 5 min at 4 ºC and discard supernatant.
Note: It is important to keep the cells cold at all times for the remainder of the procedure. Any
bottles or solutions that the cells come into contact with must also be pre-chilled on ice.
4. Resuspend the cell pellet in 25 ml of ice cold H2O.
5. Centrifuge the cells at 3000 rpm for 5 min at 4 ºC and discard supernatant.
6. Resuspend the cell pellet in 25 ml of ice cold H2O.
7. Centrifuge the cells at 3000 rpm for 5 min at 4 ºC and discard supernatant.
8. Resuspend the cell pellet in 25 ml of ice cold 10% Glycerol.
9. Centrifuge the cells at 3000 rpm for 5 min at 4 ºC. Carefully decant the supernatant.
Note: Cells lose their adherence in 10% glycerol, so the pellet may not be as compact as after
previous centrifuge spin cycles.
10. Prepare 5 microcentrifuge tubes by placing them in ice. Make certain to properly label your tubes.
Remember, you’ll have two varieties of competent cell; the chemically competent cells that were
previously prepared, and the electrocompetent cells.
11. Resuspend the cell pellet in 1 ml of ice cold 10% Glycerol.
Question: How would you make 500 ml of 10% glycerol using a 100% glycerol stock and water?
12. Aliquot cells 200 μl/tube, flash freeze and store at -80 °C.
To flash freeze put tubes in a bath of dry ice and ethanol, followed by just dry ice.
Make sure you wipe the ethanol off the tubes before placing them in just dry ice
Or, flash freeze with liquid nitrogen.
13. These cells are now ready for transformation by electroporation and do not require the addition of
CaCl2 as in the first step of Module II.
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Part IV – Comparing Chemically Competent and Electrocompetent Cells
We will compare the transformation efficiency of the chemically competent cells to the electrocompetent
cells.
To transform the chemically competent cells:
1. Add 2 μl of purified undigested pET3a vector (from previous experiments) to one of the tubes of
chemically competent cells.
2. Incubate the tube for 15 minutes on ice.
3. Heat shock cells by placing the tube in a 42 °C water batch for 45 seconds.
4. Place the tube on ice for 2 minutes.
5. Add 800 μl of LB to the tube and incubate for one hour at 37 °C.
6. Spread 100 μl of cells onto an LB-amp agar plate.
7. Observe for colonies during the next lab period.
To transform the electrocompetent cells:
1. Gently thaw one of the tubes of electrocompetent cells on ice.
2. Remove sterile cuvettes from their pouches and place them on ice.
3. In a cold 1.5 ml microcentrifuge tube, mix 40 μl of cell suspension with 2 μl of pET3a vector
DNA. Mix well and let sit on ice for 1 min.
4. Set the electroporator apparatus to 2.5 kV when using the 0.2 cm cuvettes. Set it to 1.8 kV when
using 0.1 cm cuvettes.
5. Transfer the mixture of cells and DNA to a cold electroporation cuvette, and shake the suspension
to the bottom. Place the cuvette in a chilled safety chamber slide. Push the slide into the
chamber until the cuvette is seated between the contacts in the base of the chamber.
6. Pulse once.
7. Remove the cuvette from the chamber and immediately add 1 ml of LB or SOC medium to the
cuvette and quickly but gently resuspend the cells with a pipette.
8. Record pulse parameters. The time constant should be close to 5 msec and the field strength can
be calculated as volts (kV)/cuvette gap (cm).
9. Transfer the cell suspension to a 37 °C incubator and shake for at 220 rpm for 1 hr.
10. Plate 100 μl onto an LB-amp plate and incubate overnight at 37 °C.
During the next lab period record the number of CFUs and calculate the transformation efficiency for
each method.
Questions:
1. Which method resulted in more transformants?
2. How could the comparison procedure be improved to provide a more accurate comparison
between the two methods?
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