DNA CLONING
INTRODUCTION
Recombinant DNA technology is at the heart of the biotechnology industry. In this lab, we will be
performing restriction enzyme cloning to create a new (recombinant) plasmid. It is this same method that
Herbert Boyer and Stanley Cohen used in 1973 to herald in the field of genetic engineering. The only
difference is that we will be using updated techniques and materials, ones that you will find in any
molecular biology laboratory today! We be performing the steps a molecular biologist would carry out in
engineering a biotechnological product.
We will be taking a gene of interest (GOI) and moving it from one plasmid to another. GOI will be
coming out of a plasmid containing the kanR gene which supplies the bacteria with resistance to
kanamycin. We will call this plasmid pGOI. The GOI will be moved into an ampicillin resistant plasmid
called pGEM. Our final product should be pGEM-GOI. pGEM not only has an ampR gene, but the lacZ
gene as well. lacZ encodes for the enzyme -galactosidase. This enzyme is normally used by bacteria to
cleave disaccharides, such as lactose, for metabolism. In this lab, we are going use -galactosidase
activity to track whether we have cloned GOI into pGEM successfully. Since we will clone GOI into
restriction sites in lacZ, a successful cloning event should disrupt its function. So we will be engineering a
plasmid that would allow cells to be ampicillin resistant and have disrupted -galactosidase activity.
-galactosidase activity will be tracked by enzymatic cleavage of an artificial disaccharide, X-Gal. X-Gal,
when cleaved, will release a blue color. X-Gal and IPTG (lacZ is connected to the lac operon and must be
induced) will be plated with transformed cells, and successful pGEM-GOI transformants will not be blue,
while transformants with intact pGEM plasmids will turn blue. This typical selection procedure is called
blue/white selection and is a commonly used marker for successful recombination.
The reason for such an elaborate tracking system is because so many things can go wrong! First,
fragments produced by restriction digestion can recombine randomly, as long as sticky ends are
compatible. Therefore, various combinations of fragments can come together and become ligated into a
new plasmid creating a large number of possible clones. Second, restriction digests do not always cut
every piece of DNA. Plasmids that happen to be cut by only one of the enzymes will probably re-ligate
onto itself. Plasmids that are not cut at all will not undergo any changes, but still be transformed. By
understanding what the different parts of the plasmids are doing, we can predict what the final
recombined plasmids will look like based on the phenotype of the transformed colonies.
Maps of the two original plasmids are shown on the next page. Restriction sites for a few select enzymes
are shown. Total sizes are in the center of the maps and cut sites are displayed in parentheses next to the
restriction enzyme.
DNA CLONING
Ori
BamHI (1112)
Ori
pGOI
(7269)
pGEM
(3197)
GOI
lacZ
EcoRI (5)
BamHI (26)
HindIII (56)
KanR
AmpR
HindIII (2868)
The restriction enzymes we will use to cut out GOI are BamHI and HinDIII. pGEM will be cut with the
same enzymes. This will produce fragments with a BamHI sticky end on one side and a HinDIII sticky
end on the other side. When the fragments recombine, any combination will occur as long as the sticky
ends match. After ligation and transformation into E. coli, only circularized plasmids with an Ori and
antibiotic resistance (if selection is there) will be selected for. All other forms will not be maintained or
allow the bacteria to survive. Once colonies of transformed bacteria are grown, overnight cultures will be
made and restriction analysis will be used to check the correctness of the clones. This will involve the
same kind of analysis performed in the GFP lab. A table of the restriction cut sites of enzymes we will use
in this lab is shown below.
Table 1. Restriction enzyme recognition (↓ indicates cut site)
BamHI
5’ G↓GATCC 3’
EcoRI
5’ G↓AATTC 3’
HinDIII
5’ A↓AGCTT 3’
This entire lab will take 5 lab periods. A summary of the lab activities is shown in the grid below.
Day 1
Restriction digest and ligation
Day 2
Transformation and plating
Extra Day
Preparing overnight cultures
Day 3
Analysis of plates and miniprep
Day 4
Restriction analysis
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DNA CLONING
On Day 1, a double digest using restriction enzymes that will cut out GOI and open up pGEM will be
performed. This will create four different fragments, each having sticky ends that can recombine with
complementary sequences. The fragments will be allowed to randomly recombine and their
recombination made permanent by ligation.
On Day 2, the recombinant plasmids will then be used to transform E.coli bacterial cells. Transformed
cells will be screened for resistance to each ampicillin and kanamycin and both together. X-Gal and IPTG
will also be added to appropriate plates.
On Extra Day, plates will be removed from the incubator and briefly analyzed. A white colony will be
picked from the ligation LB amp and LB amp/kan plate. Overnights will be prepared for each colony for
minipreps.
On Day 3, plates will be fully analyzed. Colonies will be counted on each one. Minipreps of the two
overnights will be performed.
On Day 4, restriction analysis of the minipreps will be performed. You will cut them with the same
enzymes used to do the cloning. Gel electrophoresis and staining will be done. You will then be able to
determine if you successfully created recombinant plasmids.
DAY 1: RESTRICTION DIGEST
MATERIALS
0.075 µg/µl pGOI
0.075 µg/µl pGEM
5X restriction buffer/RNase
Enzyme mix (BamHI and HinDIII)
Micropipets
PROCEDURE
1. Use Table 2 as a guide to prepare your digest.
Table 2. Preparing restriction digest
pGOI
pGEM
5X Buffer
Enzyme mix
5 µl
5 µl
3 µl
2 µl
2. Obtain a centrifuge tube and label your group name on the tube.
3. Add the reagents into the tube in the order of Table 2.
4. Close caps, mix, and pulse in centrifuge.
5. Incubate at 37 °C for 30 minutes.
LIGATION OF RESTRICTION FRAGMENTS
MATERIALS
Digested pGOI and pGEM
6X ligation buffer/ATP
DNA ligase
Sterile microfuge tube
3
Total
15 µl
DNA CLONING
PROCEDURE
1. Incubate the digest tube from the previous step at 65°C for 10 minutes. This step is necessary
denature restriction enzymes.
2. Add 3 µl of 6X ligation buffer/ATP to the digest tube.
3. Add 2 µl of DNA ligase to the tube.
4. Mix and pulse tube in centrifuge. Incubate 24 hours at room temperature. (Refrigerate until next lab
period)
DAY 2: TRANSFORMATION
Ligated DNA from the previous day will be used to transform JM109 E. coli cells. Purified pGOI and
pGEM plasmids will also be used as controls. Transformed cells will be plated onto 4 types of LB agar:
with ampicillin (LB/amp); with kanamycin (LB/kan); with both ampicillin and kanamycin
(LB/amp+kan); and with plain LB (LB).
PREPARING COMPETENT CELLS
All of the steps in this lab must be prepared under sterile conditions.
MATERIALS
2 microfuge tubes of mid-log E. coli (JM109) cells
Sterile CaCl2 solution
Micropipets and sterile tips
Beaker of ice
PROCEDURE
Disinfect workspace.
1. Place sterile CaCl2 solution on ice.
2. Obtain two microfuge tubes of mid-log JM109 cells. There should be about 1.5 ml in each tube.
3. Make sure caps are securely closed and place tubes in microfuge and spin at top speed for 2 minutes.
Ensure proper balance in rotor head before starting.
4. Carefully and sterilely pour off supernatant from each tube into your biohazard waste container. Make
sure tube is well drained by gently tapping on a small piece of clean paper towel (dispose of in
hazardous waste), but don't lose the pellet.
5. Sterilely add 1,000 µl of cold CaCl2 to each tube with a sterile pipette.
6. Tightly close caps.
7. Use a micropipette set to 300 µl and pipette pellet in and out to thoroughly mix the cells into the
CaCl2. Hold tube up to the light to make sure all the cells are mixed into solution.
8. Return both tubes to ice, and incubate for 20 minutes.
9. After the 0°C incubation, spin down cells in the microfuge for 1 minute.
10. Carefully and sterilely pour off supernatant from each tube into your biohazard waste container. Make
sure tube is well drained by gently tapping on a clean paper towel (dispose of in hazardous waste), but
don't pour off pellet.
11. Sterilely add 500 µl of fresh, ice cold CaCl2 to each tube.
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DNA CLONING
12. Pipette the pellet in and out to thoroughly mix the cells into the CaCl2. Hold tube up to the light to
make sure all the cells are mixed into solution.
13. Return to ice for 20 minutes and prepare for transformation step.
14. After this incubation, the cells are "competent" to take up plasmid DNA.
15. Mix the contents of one tube and transfer all of its contents into the other tube (pool the competent
cells into one tube).
TRANSFORMATION
MATERIALS
pGOI (0.075 µg/µl)
pGEM (0.075 µg/µl)
X-Gal (25 µg/µl)
IPTG (12.5 µg/µl)
Sterile microfuge tubes
Beaker of ice
Micropipets and tips
3 LB/amp+kan, 3 LB/amp, 3 LB/kan, 3 LB agar plates
Spreader, dish, and 95% ethanol
PROCEDURE
1. Disinfect workspace.
2. Label 3 sterile microfuge tubes as follows:
+pLIG = this tube will contain your ligated recombinant plasmids from the previous lab.
+pGOI = this will be a control tube containing pGOI.
+pGEM = this will be a control tube containing pGEM.
3. Sterilely add 200 µl of competent cells to each tube.
4. Place all 3 tubes on ice.
5. Sterilely add 10 µl of your ligated recombinant plasmids to the "+pLIG" tube.
6. Use a fresh tip and sterilely add 10 µl of pGOI (0.075 µg/µl) to the " pGOI " tube.
7. Use a fresh tip and sterilely add 10 µl of pGEM (0.075 µg/µl) to the " pGEM " tube.
8. Close caps and gently mix. Avoid bubbles and getting the solution on the sides of the tube.
9. Return tubes to ice for 20 minutes.
10. During the incubation, obtain the 12 plates (3 each). Mark them as indicated in Table 3.
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DNA CLONING
Table 3. Marking and loading the spread plates
Volume from tube
pLIG
pGOI
100 µl cells
100 µl cells
pGEM
100 µl cells
100 µl cells
LB/kan
100 µl cells
25 µl X-Gal
10 µl IPTG
100 µl cells
100 µl cells
100 µl cells
25 µl X-Gal
10 µl IPTG
100 µl cells
LB
100 µl cells
100 µl cells
100 µl cells
LB/amp+kan
LB/amp
11. Carry the ice beaker to the 42°C water bath.
12. Heat shock the cells by immersing the tubes in the 42°C water bath for exactly 90 seconds.
13. Immediately return the tubes to the ice bath for at least 1 minute.
14. Sterilely add 800 µl of LB broth to each tube. Gently mix.
15. Allow cells to recover by incubating all three tubes at 37°C in a water bath for 30 min.
16. Review technique for making spread plates.
17. Following recovery, make spread plates for each culture dish by adding 100 µl of the appropriate
cells with a fresh sterile tip, and immediately spread with disinfected spreader. On the two plates that
require X-Gal and IPTG, add 25 µl and 10 µl each into your 100 µl puddle of cells and spread
normally.
18. Be sure to re-flame spreader before returning it to lab bench.
19. Place in wire basket, and place in 37°C incubator. Incubate for 2 days.
20. Recover the plates the next period.
EXTRA DAY AND DAY 3: PREPARATION OF OVERNIGHT CULTURES, PLATE
ANALYSIS, AND MINIPREP
Count the number of colonies on each plate and record in the Table 4 below. If the colonies are too
numerous to count, record "lawn".
1. On your pLIG – LB/amp plate, identify a white colony. Also identify a colony on your pLIG –
LB/amp+kan plate.
2. Obtain 2 test tubes with 5 ml of sterile LB/amp broth. Label each with your team name and color of
colony.
3. With the inoculating loop, transfer each colony to the appropriate tubes (don't forget to flame the loop
for a sterile transfer). You do not have to scrape up the entire colony, a mere touch with your loop
will transfer thousands to millions of cells!
4. Make sure the tubes are appropriately labeled and place in the rack in the 37 degree C incubator.
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DNA CLONING
Table 4. Colony counts in each plate
Color
pLIG
pGOI
pGEM
LB/amp+kan
White
LB/amp
Blue
LB/kan
LB
MINIPREP
MATERIALS
2 overnight cultures made from colonies growing on the pLIG - LB/amp plate
GTE
SDS/NaOH
KOAc
Isopropanol
100% Ethanol
TE
Micropipets and tips
PROCEDURE
1. Disinfect work area.
2. Label two microfuge tubes to indicate your team name and colony used.
3. Obtain your two overnight cultures and shake the tubes to resuspend cells.
4. Transfer 1.5 ml of culture into the correct microfuge tubes.
5. Centrifuge tubes for 1 minute.
6. Pour off supernatant into biohazard waste. Make sure the pellet remains in the tube.
7. Thoroughly drain the tubes by gently tapping on clean paper towel (be sure to discard towel in
biohazard container).
8. Add 100 µl of GTE solution to each tube. Resuspend pellets by pipetting solution in and out several
times. Hold tubes up to light to check that suspension is homogeneous and that no visible clumps of
cells remain.
9. Add 200 µl of SDS/NaOH solution to each tube. Close caps and mix by rapidly inverting tubes five
times.
10. Stand tubes on ice for 5 minutes. Suspensions should become relatively clear as the cells lyse.
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DNA CLONING
11. Add 150 µl of ice-cold KOAc solution to each tube. Close caps, and mix solutions by rapidly
inverting tubes 5 times. A white precipitate will immediately appear.
12. Stand tubes on ice for 5 minutes.
13. Centrifuge tubes for 5 minutes to pellet the precipitate.
14. Transfer 400 µl of supernatant from each tube into a clean microfuge tube, labeled appropriately.
(Save the supernatants)
15. Add 400 µl of isopropanol to each tube of supernatant.
16. Close caps and mix vigorously by rapid inversion five times.
17. Stand at room temperature for exactly 2 minutes. Isopropanol preferentially precipitates nucleic
acids, but with longer exposure proteins also begin to precipitate.
18. Centrifuge tubes for 5 minutes to pellet the precipitate. Hinge of microfuge tube up so you can tell
where to expect the nucleic acid residue.
19. Pour off supernatant from both tubes. Be careful not to disturb nucleic acid pellets. (Save the pellets)
20. Invert tubes over a paper towel and tap gently to drain thoroughly.
21. Rinse the pellets by adding 200 µl of 100% ethanol to each tube. Flick the tubes several times.
22. Centrifuge tubes for 2 minutes.
23. Pour off supernatant from both tubes. Save the pellets. Invert tubes over paper towel and drain
thoroughly.
24. Close caps and pulse tubes in centrifuge. Carefully remove any remaining ethanol with 2-20 µl
pipette.
25. Leave caps open and allow pellets to dry at room temperature for 10 minutes. All ethanol must be
evaporated before proceeding.
26. Add 30 µl of TE buffer to each tube. Resuspend pellets by smashing with the pipette tip and pipetting
in and out. Focus your efforts on the area where the pellet should have formed during centrifugation
(beneath the cap hinge). Check that all DNA is dissolved and that no particles remain in tip or on the
side of tube.
27. Keep the 2 DNA solutions separate. Do not pool into one tube.
28. Freeze the DNA/TE solutions at -20°C until you are ready to proceed with the restriction analysis of
the plasmid DNA.
DAY 4
RESTRICTION ANALYSIS AND GEL ELECTROPHORESIS
MATERIALS
Miniprep DNA from previous lab
0.025 µg/µl  marker (precut with HindIII)
0.075 µg/µl pGOI
0.075 µg/µl pGEM
BamHI/HindIII
5X restriction buffer/RNAase
DI water
Loading dye
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DNA CLONING
0.8% agarose
1X Tris/Borate/EDTA (TBE) buffer
Micropipets and tips
Microfuge tubes
PROCEDURE
Tube
Table 5. Preparing tubes for restriction digest
DNA
Buffer/RNAase
pLIG Amp
pLIG Amp/Kan
pGOI
pGEM
 marker
5 µl
5 µl
5 µl
5 µl
BamHI/
HindIII
3 µl
2 µl
3 µl
2 µl
3 µl
2 µl
3 µl
2 µl
10 µl supplied  DNA cut with HindIII
1. Disinfect workspace.
2. Label 6 tubes according to the first column in the table below.
3. Use Table 5 as a guide to add the proper reagents to each tube. Read down each column, adding the
reagent to all appropriate tubes. Use a fresh tip for each reagent. Your lambda marker is already
prepared for you. All you need to do is add loading dye.
4. When all tubes have their proper reagents, mix the tubes and pulse them in the microfuge.
5. Incubate at 37°C for 20 minutes.
6. Cast your agarose gel and prepare the gel box during the incubation
7. After removing tubes from the incubator, add 2 µl of loading dye to each tube.
8. Pulse in centrifuge to mix.
9. Load 12 µl of each tube into appropriate wells in gel in the same order they are in column 1 of the
table above.
10. Close gel box and connect to power supply. Electrophorese at about 100 volts until the smallest dye
marker has migrated about 3/4 of the way across gel.
11. Turn off power and remove gel from box. Take the gel to the lab technician for staining with
Ethidium Bromide.
12. The technician will photograph your gel.
Observe your data in table 4 and the photograph of your gel. The following are rules that are
required for a functional plasmid.
1. Every replicating plasmid must have an origin of replication. Recombinant plasmids with more than
one origin also replicate normally (however, only one origin is active).
2. Each adjacent restriction fragment can only ligate at a like restriction site, BamHI to BamHI and
HindIII to HindIII.
3. Repeated copies of a restriction fragment cannot exist adjacent to one another; that is they must
alternate with other fragments. Adjacent duplicate fragments form "inverted repeats" in which the
sequences, one on either side of the restriction site, are complementary along the entire length of the
duplicated fragment. Molecules with such inverted repeats cannot replicate properly. As the plasmid
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DNA CLONING
opens up to allow access to DNA polymerase, the single strand regions on either side of the
restriction site base pair to one another to form a large "hairpin loop," which fouls replication.
QUESTIONS FOR YOUR REPORT
1. Present a ½ page summary of purpose of this lab, what methods you used, and your
results/conclusions.
2. Calculate the expected sizes of fragments generated from each of the restriction digests from
Day 1. These are your pGOI and pGEM plasmids cut with BamHI and HinDIII
3. Explain what each of the following plasmid parts does: Ori, AmpR, KanR, GOI, LacZ.
4. Present your colony count data from table 4.
5. Explain what you expected to see on each of the LB plates in table 4 and what the purpose of
each plate was.
6. Explain what you expected to see on each of the LB/amp plates in table 4 and what the
purpose of each plate was. What is the significance of white vs. blue colonies?
7. Explain what you expected to see on each of the LB/kan plates in table 4 and what the
purpose of each plate was.
8. Explain what you expected to see on each of the LB/amp+kan plates in table 4 and what the
purpose of each plate was. What is the significance of white vs. blue colonies?
9. Did you see any differences with what you expected in questions 5-8? If so, explain these
differences.
10. What would be an explanation for finding only white colonies on the pGEM-LB/amp plate?
11. Present your gel picture with all lanes properly labeled. Explain any irregularities or
unexpected results.
12. Present your standard curve generated using the  marker.
13. Present your calculated estimations of fragment sizes in each lane on your gel. Include a table
with measured distances and calculated fragment sizes.
14. Based on your results, construct a map of each of the plasmids you created (from LB amp
and LB amp/kan). Use actual fragment sizes/cut sites, not estimated.
15. In the way we created our recombinant plasmids, explain why it is impossible to have created
one with an odd number of fragments?
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