370 Labs 1 4 - 2013v1
370 Labs 1 4 - 2013v1
370 Labs 1 4 - 2013v1
This laboratory begins an experimental stream to construct and analyze a recombinant DNA molecule.
The starting reactants are the plasmids pAMP and pKAN, each of which carries a single antibiotic
resistance gene: ampicillin in pAMP and kanamycin in pKAN. The goal is to construct a recombinant
plasmid that contains both ampicillin and kanamycin resistance genes.
In Part A, Restriction Digest of Plasmids pAMP and pKAN, samples of both plasmids are digested in
separate restriction reactions with BamHl and Hindlll. Following incubation at 37°C, samples of digested
pAMP and pKAN are electrophoresed on an agarose gel to confirm proper cutting. Gel electrophoresis is
carried out in the following manner: the digested DNA samples are first loaded into wells of a 0.8%
agarose gel. An electric field applied across the gel causes the DNA fragments to move from their origin
(the sample well) through the gel matrix toward the positive electrode. The gel matrix acts as a sieve
through which smaller DNA molecules migrate faster than larger ones; restriction fragments of differing
sizes separate into distinct bands during electrophoresis. The characteristic pattern of bands produced
by each restriction enzyme is made visible by staining with a dye (ethidium bromide) that binds to the
DNA molecule. Each plasmid contains a single recognition site for each enzyme, yielding only two
restriction fragments. Cleavage of pAMP yields fragments of 784 bp and 3755 bp, and cleavage of pKAN
yields fragments of 1875 bp and 2332 bp.
In Part B, Ligation of pAMP and pKAN Restriction Fragments, the restriction digests of pAMP and
pKAN are heated to destroy BamHl and HindlIl activity. A sample from each reaction is mixed with DNA
ligase plus ATP and incubated at room temperature. Complementary BamHl and Hindlll "sticky ends”
hydrogen-bond to align restriction fragments. Ligase catalyzes the formation of phosphodiester bonds
that covalently link the DNA fragments to form stable recombinant DNA molecules.
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MATERIALS:
I. Set Up Restriction Digest (40-60 minutes, including incubation through Section III)
1. Use the table below as a checklist while adding reagents to each reaction. Read down each column,
adding the same reagent to all appropriate tubes. Use a fresh tip for each reagent. Refer to detailed
directions that follow.
pKAN
Reagent pAMP 2X BamHI /
0.2
Tube 0.2 µg/µl Buffer HindIII
µg/µl
Digested pAMP 5.5 µl 0 7.5 µl 2 µl
Digested pKAN 0 5.5 µl 7.5 µl 2 µl
2. Collect 2x buffer and BamHI/HindIII enzyme mix, and place in test tube rack on lab bench.
3. Add 7.5 µL of 2x restriction buffer to each of the two 1.5 mL tubes containing pAMP or pKAN.
4. Use fresh tip to add 2 µL of BamHI/HindIII to each tube.
5. Close tube tops. Pool and mix reagents by sharply tapping the tube bottom on lab bench or flicking
with your finger. Collect sample at bottom of tube by briefly (<10 seconds) microfuging.
6. Place reaction tubes in 37°C water bath, and incubate for a minimum of 60 minutes. Reactions can
be incubated for a longer period of time.
OPTIONAL STOP POINT: After a full 45-minute incubation (or longer), freeze reactions at -20°C until ready
to continue. Thaw reactions before proceeding to Section III, Step 1. and Part B
YOU WILL NOT BE CASTING THE AGAROSE GELS, HOWEVER YOU ARE RESPONSIBLE FOR
UNDERSTANDING HOW THE GELS ARE CAST AS WELL AS THE SAFETY PRECAUTIONS
REQUIRED WHEN CASTING AGAROSE GELS.
NOTE: Too much buffer will channel current over the top rather than through the
gel, increasing the time required to separate DNA. TBE buffer can be used
several times; do not discard. If using buffer remaining in electrophoresis box
from a previous experiment, rock chamber back and forth to remix ions that have
accumulated at either end.
OPTIONAL STOP POINT Cover electrophoresis tank and save gel until ready to continue. Gel will remain in
good condition for at least several days if it is completely submerged in buffer.
BIO370Y5 2013 LABORATORY #1 Page 5 of 9
NOTE: Only a fraction of the BamHII/HindIII digests of pAMP and pKAN are
electrophoresed to check whether plasmids are completely cut. These restriction
samples are electrophoresed along with uncut pAMP and pKAN as controls.
8. Electrophorese at 150 volts for 25 minutes. Adequate separation will have occurred when the
bromophenol blue band has moved 3-4 cm from wells.
NOTE: The 784 bp BamHI/HindIII fragment of pAMP migrates just behind the
bromophenol blue marker. Stop electrophroesis before the bromophenol blue band
runs off the end of the gel or this fragment may be lost.
9. Turn off power supply and disconnect leads from the inputs. Remove top of electrophoresis box.
BIO370Y5 2013 LABORATORY #1 Page 6 of 9
10. Carefully remove casting tray from electrophoresis chamber, and slide gel into disposable weigh boat
or other shallow tray. Label staining tray with your name.
Examine the photograph of your stained gel (or view on a light box). Compare your gel with the ideal gel
on following page, and check whether both plasmids have been completely digested by BamHI and
HindIII.
1. The Digested pAMP lane should show two distinct fragments: 784 bp and 3755 bp.
2. The Digested pKAN lane should show two distinct fragments: 1875 bp and 2332 bp.
3. Additional bands that comigrate with bands in the uncut Control pAMP and Control pKAN should be
faint or absent, indicating that most or all of the pAMP and pKAN plasmid has been completely
digested by both enzymes.
BIO370Y5 2013 LABORATORY #1 Page 7 of 9
4. If both digests look complete, or nearly so, continue on to Part B, Ligation of pAMP and pKAN
Restriction Fragments. The reaction will have gone to completion with the additional incubation during
electrophoresis.
5. If either or both digests look very incomplete, add another 1 µl of BamHI/HindIII solution and incubate
for an additional 20 minutes. Then continue on to Part B, Ligation of pAMP and pKAN Restriction
Fragments.
OPTIONAL STOP POINT: Freeze BamHI/HindIII reactions at -20°C until ready to continue. Thaw reactions
before proceeding to Part B, Ligation of pAMP and pKAN Restriction Reactions.
MATERIALS
PROCEDURE
1. Incubate tubes labeled “Digested pAMP” and “Digested pKAN” (from Part A) in 85°C block heater for
15 minutes.
NOTE: Step 1 is critical. Heat denatures protein, thus inactivating the
restriction enzymes.
2. Label one clean 1.5 ml tube Ligation (L) the other one Control Ligation (CL).
3. Use table below as a checklist while adding reagents to the Ligation tube. Use a fresh tip for each
reagent. Refer to detailed directions that follow.
BIO370Y5 2013 LABORATORY #1 Page 8 of 9
2 X Ligation
Reagent Digested Digested
Buffer / Water Ligase
Tube pAMP pKAN
ATP
Ligation (L) 3 µL 3 µL 10 µL 3 µL 1 µL
Control ligation (CL) 3 µL 3 µL 10 µL 3 µL 1 µL
4. Collect reagents (except ligase), and place in test tube rack on lab bench. Your digested pAMP and
pKAN and control digested pAMP, pKAN from the TA
5. Use fresh tip to add 3 µL of distilled water to two tubes L and CL
6. Use fresh tip to add 10 µL of 2 x ligation buffer with ATP to two tubes L and CL.
7. Use fresh tip to add 3 µL of dpAMP to L tube
8. Use fresh tip to add 3 µL of cdpAMP to CL tube
9. Use fresh tip to add 3 µL of dpKAN to L tube
10. Use fresh tip to add 3 µL of cdpKAN to CL tube
11. Use fresh tip to add 1 µL of DNA ligase to the L tube and 1 µL of DNA ligase to the CL tube.
Carefully check that droplet ligase is not on the inside wall of tubes.
12. Close tube top. Pool and mix reagents by pulsing in a microfuge or by sharply tapping tube bottom on
lab bench. Be sure to balance microfuge rotor by placing tubes of equal weight across from
each other.
13. Give your reaction to your TA for incubation at 16°C for overnight. Also, provide your TA with your
remaining digested plasmid for storage.
NOTE: For brief ligations of 2-4 hours, it is essential to use a high-concentration
T4 DNA ligase with at least 5 Weiss Units/µL or 100-500 Cohesive-end Units/µL
14. If time permits, ligation may be confirmed by electrophoresing 5 µL of the ligation reaction, along with
BamHI/HindlII digests of pAMP and pKAN. None of the parent BamHI/HindIII fragments should be
observed in the lane of ligated DNA, which should show multiple bands of high-molecular-weight
DNA high up on the gel.
OPTIONAL STOP POINT: Freeze reaction at -20°C until ready to continue. Thaw reaction before proceeding
to Laboratory 2.
Ligation of the four BamHI/HindIII restriction fragments of pAMP and pKAN produces many types of
hybrid molecules, including plasmids composed of more than two fragments. However, only those
constructs possessing an origin of replication will be maintained and expressed. Three different
replicating plasmids, with selectable antibiotic resistance, are created by ligating combinations of two
BamHI/HindIII fragments:
• Ligation of the 1875-bp fragment to the 2332-bp fragment regenerates pKAN.
• Ligation of the 784-bp fragment to the 3755-bp fragment regenerates pAMP.
BIO370Y5 2013 LABORATORY #1 Page 9 of 9
• Ligation of the 1875-bp fragment to the 3755-bp fragment produces the "simple recombinant"
plasmid pAMP/KAN, in which the kanamycin resistance gene has been fused into the pAMP
backbone.
1. Make a scale drawing of the simple recombinant molecule pAMP/KAN described above. Include
fragment sizes, locations of BamHI and HindIII restriction sites, location of origin(s), and location of
antibiotic resistance gene(s).
2. Make scale drawings of other two-fragment recombinant plasmids having the following properties.
Whenever possible, include fragment sizes, locations of BamHI and HindIII restriction sites, location
of origin(s), and location of antibiotic resistance gene(s):
a. Three kinds of plasmids having two origins.
b. Three kinds of plasmids having no origin.
3. Ligation of the 784-bp fragment, 3755-bp fragment, 1875-bp fragment, and 2332-bp fragment
produces a "double plasmid" pAMP/pKAN. Make a scale drawing of the double plasmid pAMP/pKAN.
4. Make scale drawings of several recombinant plasmids composed of any three of the four
BamHI/HindIII fragments of pAMP and pKAN. Include fragment sizes, locations of BamHI and HindIII
restriction sites, location of origin(s), and location of antibiotic resistance gene(s). What rule governs
the construction of plasmids from three kinds of restriction fragments?
5. What kind of antibiotic selection would identify E. coli cells that have been transformed with each of
the plasmids drawn in Questions 1 through 4?
6. Explain what is meant by "sticky ends." Why are they so useful in creating recombinant DNA
molecules?
7. Why is ATP essential for the ligation reaction?
BIO370Y5 2013 LABORATORY #2 Page 1 of 12
In Part A, Classic Procedure for Preparing Competent Cells, E. coli cells are rendered competent to
uptake plasmid DNA using a method essentially unchanged since its publication in 1970 by M. Mandel
and A. Higa (J. Mol. Biol., 53:159). The procedure begins with vigorous E. coli cells grown in suspension
culture. Cells are harvested in mid-log phase by centrifugation and incubated at O°C with two successive
changes of calcium chloride solution.
The classic procedure typically achieves transformation efficiencies ranging from 5x104 to 106 colonies
per microgram of plasmid. The enhanced efficiency over other methods is important when transforming
ligated DNA (composed primarily of relaxed circular plasmids and linear DNA), which produces 5-100
times fewer transformants than plasmid DNA purified from E. coli (containing a high proportion of the
super-coiled form).
In Part B, Transformation of E. coli with Recombinant DNA, competent E. coli cells are transformed
with the ligation products from Laboratory 1, Recombination of Antibiotic Resistance Genes. Ligated
plasmid DNA is added to one sample of competent cells and purified pAMP and pKAN plasmids are
added as controls to two other samples. The cell suspensions are incubated with the plasmid DNAs for
20 minutes at 0°C. Following a brief heat shock at 42°C, the cells recover in LB broth for 40-60 minutes
at 37°C. The recovery step is essential for the kanamycin selection in this lab. Samples of transformed
cells are plated onto three types of LB agar: with ampicillin (LB/amp), with kanamycin (LB/kan), and with
both ampicillin and kanamycin (LB/amp+kan).
The ligation reaction produces many kinds of recombinant molecules composed of BamHI/Hindlll
fragments including the religated parental plasmids pAMP and pKAN. The object is to select for
transformed cells with dual antibiotic resistance which must contain a 3755-bp fragment from pAMP
containing the ampicillin resistance gene (plus the origin of replication) and a 1875-bp fragment from
pKAN containing the kanamycin resistance gene. Bacteria transformed with a single plasmid containing
these sequences, or those doubly transformed with both pAMP and pKAN plasmids, form colonies on the
LB/amp&kan plate.
MATERIALS
NOTE: A tight pellet of cells should be easily seen at the bottom of the
tube. If the pellet does not appear to be consolidated, recentrifuge for
an additional 5 minutes.
NOTE: To ensure step 4 and step 5 are done entirely under sterile
conditions, plan out your manipulations. Organize lab bench, and work
quickly. Locate Bunsen burner in a central position on lab bench to avoid
reaching over flame. If working as a team, one person handles pipets, and
the other removes and replaces tube caps.
BIO370Y5 2013 LABORATORY #2 Page 3 of 12
4. Sterilely pour off supernatant from each tube into waste beaker for later disinfection. Be careful not
to disturb cell pellet.
a. Remove cap from culture tube, and briefly flame mouth. DO NOT
PLACE CAP ON LAB BENCH.
b. Carefully pour off supernatant. Invert culture tube, and tap gently on
surface of clean paper towel to drain thoroughly.
c. Reflame mouth of culture tube, and replace cap.
5. Use a 1 ml pipetman to sterilely add 1 mL of ice-cold CaCl2 solution to each culture tube:
a. Briefly flame pipet cylinder.
b. Remove cap from CaCl2, tube, and flame mouth. DO NOT PLACE CAP ON LAB BENCH.
c. Withdraw 1 mL of CaCl2. Reflame mouth of tube, and replace cap.
d. Remove cap of culture tube, and flame mouth. Do not place cap on lab bench.
e. Expel CaCl2, into culture tube. Reflame mouth of culture tube, and replace cap.
NOTE: The cell pellet becomes increasingly difficult to suspend the longer it sits in
the CaCl2 solution.
d. Hold tube up to light to check that suspension is homogeneous. No visible clumps of cells should
remain.
e. Add 4 mL CaCl2 to each tube and gently vortex.
8. Following incubation, re-spin cells in clinical centrifuge for 5 minutes at 2000-4000 rpm.
BIO370Y5 2013 LABORATORY #2 Page 4 of 12
NOTE: CaCl2 treatment alters the adhering properties of the E. coli membranes. The
cell pellet is much more dispersed after the second centrifugation.
9. Sterilely pour off CaCl2 from each tube into waste beaker. Be careful not to disturb cell pellet.
a. Remove cap from culture tube, and briefly flame mouth. DO NOT PLACE CAP ON LAB BENCH.
b. Carefully pour off supernatant. Invert culture tube, and tap gently on surface of clean paper towel
to drain thoroughly.
c. Reflame mouth of culture tube, and replace cap.
10. Use a 100 or 1000 µL micropipettor (or 1 mL pipet) to sterilely add 1000 µL (1 mL) of fresh, ice-cold
CaCl2 to each tube.
a. Remove cap from CaCl2 tube, and flame mouth. DO NOT PLACE CAP ON LAB BENCH.
b. Withdraw 1000 µL (1 mL) of CaCl2. Reflame mouth of tube, and replace cap.
c. Remove cap of culture tube, and flame mouth. Do not place cap on lab bench.
d. Expel CaCl2 into culture tube. Reflame mouth of culture tube, and replace cap.
11. Close caps tightly, and immediately finger vortex to resuspend pelleted cells in each tube. Hold tube
up to light to check that suspension is homogeneous. No visible clumps of cells should remain.
NOTE: Cell pellet may appear more diffuse than at the beginning of the procedure
and will resuspend more easily. Double check both tubes for complete resuspension
of cells.
OPTIONAL STOP POINT: Store cells in beaker of ice in refrigerator (approximately 0°C) until ready for use.
"Seasoning" at 0°C for up to 24 hours increases the competency of cells five- to tenfold.
pCLIG 100 µL -- 10 µL -- --
pAMP 30 µL -- -- 10 µL --
pKAN 30 µL -- -- -- 10 µL
BIO370Y5 2013 LABORATORY #2 Page 6 of 12
NOTE: Return remaining ligated and ligated control DNA to your TA for storage at 4°C for
use in Lab 4b.
4. Tap tubes with finger to mix. Avoid making bubbles in suspension or splashing suspension up sides
of tubes.
• Set for Ligation: Mark “L” on one LB/amp, one LB/kan, and one LB/amp&kan plate
• Set for Ligation: Mark “CL” on one LB/amp, one LB/kan, and one LB/amp&kan plate
• Set for pAMP: Mark “A” on one LB/amp, one LB/kan, and one LB/amp&kan plate
• Set for pKAN: Mark “K” on one LB/amp, one LB/kan, and one LB/amp&kan plate
7. Following 30 minute incubation on ice, heat shock the cells in all three tubes as follows:
a. Carry ice bucket containing your samples to the 42°C water bath. Remove tubes from ice,
and immediately immerse in 42°C water bath for 60 seconds.
b. Immediately return all three tubes to ice for at least 1 minute.
8. Use a 100-1000 µl micropipettor with a sterile tip to add 400 µl of LB broth to each tube. Gently tap
tubes with finger to mix.
9. Allow cells to recover by incubating all three tubes at 37°C in a shaking water bath (with moderate
agitation) for 60 minutes.
NOTE: If shaking water bath is not available, warm cells for several
minutes in 37°C water bath, then transfer to dry shaker inside 37°C
incubator. Alternately, occasionally swirl tubes by hand in non-shaking
37°C water bath.
OPTIONAL STOP POINT: Cells may be allowed to recover for up to several hours. A longer recovery period
assures the growth of as many kanamycin resistant recombinants as possible and can help to
compensate for a poor ligation or cells of low competence.
BIO370Y5 2013 LABORATORY #2 Page 7 of 12
10. Use table on the next page as a checklist for spreading the +pLIG, +pAMP, and +pKAN cells on each
type of antibiotic plate. The objective is to evenly distribute and separate cells on agar so that each
cell gives rise to a distinct colony of clones.
Read through steps 11-18 carefully before proceeding
Ligated control
Ligated DNA pAMP control pKAN control
DNA
(+pLIG) (+pAMP) (+pKAN)
(+pCLIG)
L A K
L
LB/amp 100 µL 100 µL 100 µL 100 µL
11. Lift the lid of first “L” plate only enough to fit the tip of the micropipettor; do not place lid on lab
bench. Use micropipettor with sterile tip to add 100 µL of cell suspension from the “pLIG” tube onto
each of the three plates marked L.
Do not allow suspensions to sit on plates too long before proceeding to Step 12.
NOTE: If too much liquid is absorbed by agar, cells will not be evenly
distributed.
12. Carefully and sterilely remove a cell spreader from the sterile package, and spread cells over surface
of each “L” plate. Use a new spreader for each plate. To spread cells, touch spreader to cell
suspension, and gently drag it back and forth several times across surface of agar. Rotate plate one
quarter turn, and repeat spreading motion. Be careful not to gouge agar.
BIO370Y5 2013 LABORATORY #2 Page 8 of 12
NOTE: If working as a team, one person handles pipets, one person lifts and
replaces plate lids, and one person handles cell spreader.
13. Use fresh sterile tip to add 100 µL of cell suspension from tube labeled “+pCLIG” onto each of the
three plates marked “CL”.
14. Repeat Step 12 to spread cells over the surface of each “CL” plate in succession.
15. Use fresh sterile tip to add 100 µL of cell suspension from tube labeled “+pAMP” onto each of the
three plates marked “A”.
16. Repeat Step 12 to spread cells over the surface of each “A” plate in succession.
17. Use fresh sterile tip to add 100 µL of cell suspension from tube labeled “+pKAN” onto each of the
three plates marked “K”.
18. Repeat Step 12 to sterilize spread cells over the surface of each “K” plate in succession.
19. Let plates set for several minutes to allow suspension to become absorbed into agar. Then wrap
plates together with tape.
20. Place plates upside down in 37°C incubator, and incubate for 12-24 hours.
21. After initial incubation, store plates at 4°C to arrest E. coli growth and to slow the growth of any
contaminating microbes.
22. Take time for responsible cleanup:
a. Segregate for proper disposal: culture plates and tubes, pipets, and micropipettor tips that
have come in contact with E. coli.
b. Disinfect overnight cell suspensions, tubes, and tips with 70% ethanol solution.
c. Wipe down lab bench with soapy water, 70% ethanol solution.
d. Wash hands before leaving lab.
BIO370Y5 2013 LABORATORY #2 Page 9 of 12
1. Record your observation of each plate in the matrix below. If cell growth is too dense to count
individual colonies, record "lawn."
LB/amp
LB/kan
LB/amp&kan
2. Compare and contrast the growth on each of the following pairs of plates. What does each pair of
results tell you about transformation and/or antibiotic selection?
“L” LB/amp and “CL” LB/amp
“L” LB/amp and “A” LB/amp
“L” LB/kan and “A” LB/kan
“A” LB/amp and “K” LB/kan
“L” LB/amp and “L” LB/kan
“L” LB/amp and “L” LB/amp&kan
BIO370Y5 2013 LABORATORY #2 Page 10 of 12
3. Calculate transformation efficiencies of “A” LB/amp and “K” LB/kan positive controls.
Remember that transformation efficiency is expressed as the number of antibiotic resistant
colonies per microgram of intact plasmid DNA.
The objective is to determine the mass of pAMP or pKAN that was spread on each plate and was
therefore responsible for the transformants observed.
a. Determine total mass (in micrograms) of pAMP and of pKAN used in Step 3 of Lab 2B:
mass = concentration x volume
b. Determine fraction of cell suspension spread onto “A” LB/amp plate (Step 15 of Lab 2B) and “K”
LB/kan plate (Step 17 of Lab 2B):
volume of suspension spread (Step 15 or 17 of Lab 2B)
fraction spread =
total volume of suspension (Steps 3 and 8 of Lab 2B)
c. Determine mass of plasmid pAMP and pKAN in cell suspension spread onto “A” LB/amp plate
and “K” LB/kan plate:
mass of plasmid total mass of plasmid fraction spread
= x
spread (answer from question 3 a) (answer from question 3 b)
d. Determine number of colonies per microgram of pAMP and pKAN. Express answer in scientific
notation:
colonies observed (answer from question 1)
transformation efficiency =
mass of plasmid spread (answer from question 3 c)
4. Calculate transformation efficiencies of the “L” LB/amp, “L” LB/kan, and the “L” LB/amp + kan
plates.
a. Calculate the mass of pAMP and pKAN used in the restriction reactions in Step 1 of Lab 1A.
b. Calculate the concentration of plasmid in each restriction reaction in Step 1 of Lab 1A.
c. Calculate mass of pAMP and pKAN used in the ligation reaction in Step 3 of Lab 1B.
d. Calculate the total concentration of plasmid in the ligation mixture in Step 3 of Lab 1B.
e. Use this concentration in calculations following Steps a-d of Question 3 above.
5. Compare the transformation efficiencies you calculated for control pAMP and pKAN versus the
ligated pAMP and pKAN. How can you account for the differences in efficiency? Take into account
the formal definition of transformation efficiency.
BIO370Y5 2013 LABORATORY #2 Page 11 of 12
1. Design and execute an experiment to compare the transformation efficiencies of linear versus
circular plasmid DNAs. Keep molecular weight constant.
2. Design and execute a series of .experiments to test the relative importance of each of the four major
steps of most transformation protocols: (1) preincubation, (2) incubation, (3) heat shock, and (4)
recovery. Which steps are absolutely necessary?
3. Design and execute a series of experiments to compare the transforming effectiveness of CaCl2
versus salts of other monovalent (+), divalent (+ +), and trivalent (+ + + ) cations.
a. Make up 50 mL solutions of each salt.
b. Check pH of each solution, and buffer to pH 7 when necessary.
c. Is CaCl2, unique in its ability to facilitate transformation?
d. Is there any consistent difference in the transforming ability of monovalent versus divalent versus
trivalent cations?
5. Repeat Experiment 4 above, but transform with a 1:1 mixture of pAMP and pKAN at each
concentration. Plate transformants on LB/amp, LB/kan, and LB/amp+kan plates. Be sure to include
a 40-60-minute recovery, with shaking.
a. Calculate percentage of double transformations at each mass: (colonies amp + kan plate) ÷
(colonies amp plate + colonies kan plate)
b. Plot a graph of DNA mass versus colonies per plate.
c. Plot a graph of DNA mass versus percentage double transformations. Under saturating
conditions, what percentage of bacteria are doubly transformed?
6. Plot a recovery curve for E. coli transformed with pKAN. Allow cells to recover for 0-120 minutes at
20-minute intervals.
a. Plot a graph of recovery time versus colonies per plate.
b. At what time point is antibiotic expression maximized?
c. Can you discern a point at which the cells began to replicate?
7. Attempt to isolate pAMP/KAN recombinants using the colony transformation protocol in Laboratory 2.
What trick would increase the likelihood of retrieving ampicillin/kanamycin-resistant colonies?
BIO370Y5 2013 LABORATORY # 3 Page 1 of 5
MATERIALS
CULTURES AND MEDIA SUPPLIES AND EQUIPMENT
“L” LB/amp plate w/colonies (from Lab 2) 2 replica-plating grids (see end of lab 3)
“L” LB/kan plate w/colonies (from Lab 2) sterile toothpicks or inoculating loop
“CL” LB/amp plate w/colonies (from Lab 2) beaker for waste
“CL” LB/kan plate w/colonies (from Lab 2) "bio-bag" or heavy-duty trash bag
1 LB/amp plate 10% bleach or disinfectant
1 LB/kan plate 37°C incubator
PROCEDURE
1. Attach a replica-plating grid to the bottom of an LB/amp plate and to the bottom of an LB/kan plate.
Use permanent marker to label the bottom of each plate with your name and the date.
NOTE: as an alternative to the replica plating grid provided, a 24 square grid may be drawn
on plate bottom with permanent marker.
BIO370Y5 2013 LABORATORY # 3 Page 2 of 5
2. Replica plate a sample of cells from one colony on the “L” LB/amp plate onto the fresh LB/amp and
LB/kan plates:
a. Use a sterile toothpick to scrape up a cell mass from a well-defined colony on the L LB/amp plate.
b. Immediately drag the same toothpick gently across agar surface to make a short diagonal (/)
streak within Square 1 of the LB/amp plate.
c. Immediately use the same toothpick to make a diagonal (/) streak within Square 1 of the LB/kan
plate.
NOTE: Lift plate lids only enough to select colony and streak. Do not place lids on lab
bench.
3. Repeat Step 2a-d using fresh toothpicks to streak cells from 5 different “L” LB/amp colonies onto
Squares 2-6 of both LB/amp and LB/kan plates.
NOTE: If you have fewer than 6 colonies on either plate, obtain a plate from another
experimenter.
4. Repeat Step 2a-d using fresh toothpicks to streak cells from 6 different “CL” LB/amp colonies onto
Squares 7-12 of both LB/amp and LB/kan plates.
5. Repeat Step 2a-d using fresh toothpicks to streak cells from 6 different “L” LB/kan colonies onto
Squares 13-18 of both LB/amp and LB/kan plates.
6. Repeat Step 2a-d using fresh toothpicks to streak cells from 6 different “CL” LB/kan colonies onto
Squares 119-24 of both LB/amp and LB/kan plates.
7. Place plates upside down in 37°C incubator, and incubate for 12-24 hours.
8. After initial incubation, store plates at 4°C to arrest E. coli growth and to slow the growth of any
contaminating microbes.
b. Disinfect overnight culture, tips, and supernatant from Step 4 with 10% bleach or disinfectant.
c. Wipe down lab bench with soapy water, 70% ethanol solution.
In general, the results of replica plating indicate the success of the ligation in Laboratory 1 and parallel
the results observed on the L LB/amp + kan plate from Laboratory 2. Thus, if there were a large number
of colonies on the LB/amp + kan plate, it is likely that there will be a high percentage of dual-resistant
colonies that grow on both the LB/amp and LB/kan replica plates. Experience indicates that 30-70% of
transformants selected with only ampicillin or kanamycin actually have dual resistance. Roughly equal
numbers of dual-resistant colonies are identified from L LB/amp and L LB/kan plates.
1. Observe the LB/amp and LB/kan plates. Use the matrix below to record as + the squares in which
new bacterial growth has expanded the width of the initial streak. Record as - the squares in which
no new growth has expanded the initial streak. Remember that nonresistant cells may survive,
separated from the antibiotic on top of a heavy initial streak; however, no new growth will be
observed.
1 13
2 14
3 15
4 16
5 17
6 18
7 19
8 20
9 21
10 22
11 23
12 24
a. Calculate the percentage of dual-resistant colonies taken from the L LB/amp plate and CL LB/amp
plate (Squares 1-12).
b.
c. Calculate the percentage of dual-resistant colonies taken from the L LB/kan plate and CL LB/amp
BIO370Y5 2013 LABORATORY # 3 Page 4 of 5
plate (Squares 13-24).
d. Give an explanation for the similarity or difference in the percentages of dual-resistant colonies taken
from the two source plates.
3. Draw restriction maps for different plasmid molecules that could be responsible for the dual
resistance phenotype.
2. The ampr protein, β-lactamase, is not actively secreted into the medium but is believed to "leak"
through the cell envelope of E. coli. Satellite colonies do not form on kanamycin plates because the
antibiotic kills all nonresistant cells outright. The following experiment tests whether resistance
protein escapes from ampicillin- and kanamycin-resistant cells.
a. Grow separate overnight cultures of an ampr colony from the A LB/amp plate and a kanr colony
from the K LB/kan plate (from Lab 2, or use other ampr and kanr strains). Inoculate 5 mL of plain LB
broth.
b. Pass each overnight culture through a 0.22 or 0.45 µm filter, and collect filtrate in a clean, sterile
15-mL tube. Filtering removes all E. coli cells.
c. Use permanent marker to mark one LB/amp plate and one LB/kan plate. Draw a line on plate
bottom to divide each plate into two equal parts; mark one half +.
d. Sterilely spread 100 µL of the A filtrate onto the + half only of the LB/amp plate. Sterilely spread
100 µL of the K filtrate onto the + half only of the LB/kan plate. Allow filtrates to soak into plates for
10-15 minutes.
e. Sterilely streak wild-type (nontransformed) E. coli cells on each filtrate-treated plate, taking care to
streak back and forth across the dividing line.
f. Incubate plates at 37°C for 12-24 hours. Compare growth on the treated versus untreated sides of
each plate.
3. The ampr protein is believed to leak primarily from stationary-stage cells. The following experiment
tests the hypothesis that leakage of β-lactamase is growth-phase-dependent.
a. Grow an overnight culture of an ampicillin-resistant colony from the A LB/amp plate in Lab 2 (or
other ampr strain). Inoculate 1 mL of plain LB broth.
b. Use overnight culture to inoculate 100 mL of fresh LB broth, and grow to make a Mid-log
Suspension Culture.
c. Sterilely withdraw 10-mL aliquots from the culture after 1, 2, and 4 hours, holding aliquots on ice.
d. Take the OD550 of each aliquot.
BIO370Y5 2013 LABORATORY # 3 Page 5 of 5
NOTE: The objective is to test resistance-protein "leakage" as a function of culture age, not
as a function of cell number. Because cell number increases over time, the cell number must
be equalized by diluting the 2- and 4-hour samples with sterile LB to the E. coli concentration
of the 1-hour sample. Since the OD550 values are proportional to the cell number, they can be
used to compute the dilution factor.
e. Pass each of the three samples through a 0.22 or 0.45 µm filter to remove the bacteria.
f. Prepare a 10- and 100-fold dilution for each filtrate, using sterile LB broth.
g. Use a permanent marker to draw a line dividing each of six LB/amp plates into equal parts.
h. Sterilely spread 100 µL of undiluted filtrate from the 1 hour sample over half of the first plate. Label
the plate with time point and dilution factor. Allow filtrate to soak into plate for 10-15 minutes.
i. Spread 100 µL of undiluted 2- and 4-hour filtrates over separate halves of the second plate, as
described above.
j. Repeat spreading procedure for the 10- and 100-fold dilutions, as described in Steps h and I
above.
k. After filtrates have soaked into the plates, sterilely streak wild type (non-transformed) E. coli cells
on each half of each plate.
Incubate plates at 37°C for 12-24 hours. Compare growth for each time point across each dilution.
Replica-plating grids:
__________
MATERIA
BIO370Y5 2013 LABORATORY # 4 Page 1 of 13
Growth of E. coli colonies on the L LB/amp + kan plate in Laboratory 2 confirms that they have been
transformed to a dual resistance (ampr/kanr) phenotype. This resistance is expressed by one or more
replicating plasmids, which were assembled in Laboratory 1 by ligating four BamHI/HindllI restriction
fragments of the parental plasmids pAMP and pKAN:
The goal of this laboratory is to determine the genotype responsible for dual resistance; that is, the
number and probable arrangement of any two or more pAMP and pKAN fragments.
In Part A, Plasmid Minipreparation of pAMP/KAN Recombinants, plasmid DNA is isolated from overnight
cultures of two different colonies from an L LB/amp+kan plate (Laboratory 2) or from replica plates
(Laboratory 3).
In Part B, Restriction Analysis of Purified Recombinant DNA, (which will be done next week), samples
of the plasmids isolated in Part A and a control sample of pAMP + pKAN are incubated with BamHl and
Hindlll. The three digested samples, uncut control and miniprep DNA and λ BstE II size markers as well
as the remainder of the ligation stored form lab 2 will be electrophoresed on an agarose gel. The
comigration of BamHI/HindIII fragments in the lanes of miniprep DNA and pAMP/pKAN controls, along
with an evaluation of the relative sizes of uncut supercoiled DNAs, gives evidence of the structure, size,
and number of plasmids present in each of the transformed strains.
PROCEDURE
NOTE: The cell pellet will appear as a small off-white smear on the bottom-side
of the tube. Although the cell pellets are readily seen, the DNA pellets in Step 14
are very difficult to observe. Getting into the habit of aligning tube with cap hinges
facing up in the microfuge rotor will allow you to always locate the pellet at tube
bottom beneath hinge.
6. Add 200 µL of SDS/NaOH solution to each tube. Close caps, and mix solutions by rapidly inverting
tubes five times.
7. Stand tubes on ice for 3-5 minutes [MAXIMUM 5 minutes!]. Suspension will become relatively clear.
8. Add 150 µL of ice-cold KOAc solution to each tube. Close caps and mix solutions by rapidly inverting
tubes five times. A white flocculent precipitate will immediately appear.
9. Stand tubes on ice for 5 minutes (you can leave the tubes longer on ice if you are waiting for tubes to
balance centrifuge).
10. Place tubes in a balanced configuration in microfuge rotor, and spin at maximum speed for 5
minutes to pellet precipitate along the side of the tube. This may need to be repeated if precipitate is
not pelleted well after first spin. If the supernatant still contains debris, remove the supernatant to a
clean tube and repeat this step to pellet the remaining precipitate.
11. Transfer 400 µL of supernatant from M1 into a clean 1.5 mL tube labeled M1. Transfer 400 µL of
supernatant from M2 into a clean1.5 mL tube labeled M2. AVOID PIPETTING PRECIPITATE, and wipe off
any precipitate clinging to the outside of the tip prior to expelling supernatant. Discard old tubes
containing precipitate.
12. Add 400 µL of isopropanol to each tube of supernatant. Close caps, and mix vigorously by rapidly
inverting tubes five times. Stand at room temperature for only 2 minutes. (Isopropanol
preferentially precipitates nucleic acids rapidly; however, proteins remaining in solution also begin to
precipitate with time.)
13. Place tubes in a balanced configuration in microfuge rotor, and spin at maximum speed for 5 minutes
to pellet the nucleic acids. Align tubes in rotor so that cap hinges point outward. The nucleic acid
residue, visible or not, will collect under hinge during centrifugation.
14. Pour off supernatant from both tubes. Be careful not to disturb nucleic acid pellets. Invert tubes,
and tap gently on surface of clean paper towel to drain thoroughly.
15. Add 200 µL of 70% ethanol to each tube, and close caps. Flick tubes several times to wash pellets.
OPTIONAL STOP POINT Store ethanol solution at -20°C until ready to continue.
16. Place tubes in a balanced configuration in microfuge rotor, and spin for 2-3 minutes.
17. Pour off supernatant from both tubes. Be careful not to disturb nucleic acid pellets. Invert tubes,
and tap gently on surface of clean paper towel to drain thoroughly.
BIO370Y5 2013 LABORATORY # 4 Page 4 of 13
NOTE: The pellet may appear as a tiny smear or small particles on the bottom-side of each
tube. Do not be concerned if pellet is not visible; pellet size is not a predictor of plasmid
yield. A large pellet is composed primarily of RNA and cellular debris carried over from the
original precipitate. A smaller pellet often means a cleaner preparation. Nucleic acid pellets
are not soluble in ethanol and will not re-suspend during washing.
18. Dry nucleic acid pellets by closing caps, and pulse tubes in microfuge to pool remaining ethanol.
Carefully draw off drops of ethanol using a 1-10 µl micropipettor. Place pellets at 37°C until dry.
19. All ethanol must be evaporated before proceeding to Step 20. Hold each tube up to light to check that
no ethanol droplets remain. If ethanol is still evaporating, an alcohol odor can be detected by sniffing
mouth of tube.
20. Add 15 µL of TE to each tube (2 x 7.5 µL using p10 micropipettor). Resuspend pellets by smashing
with the pipet tip and pipetting in and out vigorously. Rinse down the side of tube several times,
concentrating on area where the pellet should have formed during centrifugation (beneath cap
hinge). Check that all DNA is dissolved and that no particles remain in tip or on side of tube. May
help to resuspend by warming it in 37°C incubator for 1 min.
21. Keep the two DNA/TE solutions separate. DO NOT pool into one tube.
OPTIONAL STOP POINT Freeze DNA/TE solution at – 20°C until ready to continue. Thaw before using.
MATERIALS
REAGENTS PROCEDURE
miniprep DNA in TE (M1, M2, M3) SUPPLIES AND EQUIPMENTS
0.1 µg/µL pAMP/pKAN
0.5-10 µL micropipettor + tips
0.1 µg/µL pAMP
1.5 mL tubes
0.1 µg/µL pKAN
aluminum foil
BamHI/HindlIl enzyme mix
beaker for agarose
5 x restriction buffer with RNase
beaker for waste/used tips
distilled water
10% bleach
loading dye
electrophoresis box
0.8% agarose gel
microfuge
1 x Gel Running buffer
power supply
latex gloves
test tube rack
transilluminator (optional)
37°C water bath
M3 - = undigested miniprep 3
pA - = undigested pAMP
pK - = undigested pKAN
λ BstE II marker
1. Use permanent marker to label seven 1.5 mL tubes, in which restriction reactions will be performed:
M1 + = miniprep 1, BamHI/HindIII
M2 + = miniprep 2, BamHI/HindIII
M3 + = miniprep 3, BamHI/HindIII
AK + = pAMP/pKAN, BamHI/HindIII
M1 - = miniprep 1, no enzyme
M2 - = miniprep 2, no enzyme
1. Use table below as a checklist for setting up the restriction digests. Make sure the correct reagents
are added for each reaction and mark off once that reagent has been added to the tube. Read down
each column, adding the same reagent to all appropriate tubes.
BIO370Y5 2012 LABORATORY # 4 Page 6 of 13
2. ALWAYS use a fresh tip for each reagent. Refer to detailed directions that follow for the order in
which reagents should be added to the tube. Always add buffer prior to addition of restriction
enzyme.
4. Use one 10 uL tip to add proper volume of distilled water to each tube.
5. Use fresh tip to add 3.5 µL of M1 DNA to tubes labeled M1 - and M1 +.
6. Use fresh tip to add 3.5 µL of M2 DNA to tubes labeled M2 - and M2 +.
7. Use fresh tip to add 3.5 µL of M3 DNA to tubes labeled M3 + only.
8. Use fresh tip to add 5 µL of pAMP/pKAN to tube labeled AK +.
9. Use fresh tip to add 2 µL of restriction buffer/RNase to each reaction tube with plasmids which will be
digested.
10. Use fresh tip to add 2 µL of BamHI/HindIII to tubes labeled MI +, M2+, M3+ and AK+.
11. Close tube tops. Collect reagents at bottom of each tube and mix well. Spin down prior to incubation.
12. Place reaction tubes in 37°C water bath, and incubate for 45 minutes.
NOTE: Do not over incubate. During longer incubation, DNases in miniprep may
degrade plasmid DNA.
OPTIONAL STOP POINT Following incubation, freeze reactions at – 20°C until ready to continue. Thaw
reactions before continuing to Section III, Step 1.
1. Add 2 µL of loading dye to each reaction tube. Close tube tops, and mix by tapping tube bottom on
lab bench, pipetting in and out, or pulsing in a microfuge.
2. Using a p10 piptettor, load 10 µL of each reaction tube into a separate well in the gel, as show in
diagram below. Use fresh tip for each reaction. Expel any air in tip before loading, and be careful
not to punch tip of pipet through bottom of the gel.
3. Add loading dye to ligated DNA saved from Laboratory 1 (Recombination of Antibiotic Resistance
Genes). Load entire contents of “L” tube (5-10 µL) into the last well.
4. Electrophorese at 120 volts for 70 minutes, or longer. Good separation will have occurred when the
bromophenol blue band has moved 4-6 cm from wells.
• If time allows, electrophorese until the bromophenol blue band nears the end of the gel. This will
allow maximum separation of uncut DNA, which is important in differentiating a large
"superplasmid" from a double transformation of two smaller plasmids.
• Stop electrophoresis before bromophenol blue band runs off end of gel or the 784-bp
BamHI/HindIII fragment of pAMP, which migrates just behind the bromophenol blue marker, may
be lost.
5. Turn off power supply, disconnect leads from-the inputs, and remove top of electrophoresis box.
6. Carefully remove casting tray from electrophoresis box, and slide gel into disposable weigh boat or
other shallow tray. Label the staining tray with your name.
OPTIONAL STOP POINT Cover electrophoresis tank and save gel until ready to continue. Gel can be stored
in a zip-lock plastic bag and refrigerated overnight for viewing / photographing the next day. However,
over longer periods of time, the DNA will diffuse through the gel and the bands will become indistinct or
disappear entirely.
7. Use the gel documentation station to obtain an image of the gel. These images will be posted on the
course web page.
BIO370Y5 2012 LABORATORY # 4 Page 8 of 13
Observe your gel and determine which lanes contain control pAMP/ pKAN and which lanes contain
minipreps M1 and M2. Even if you have confused the prescribed loading order, the miniprep lanes can
be distinguished by the following characteristics:
• a background "smear" of degraded and partially digested chromosomal DNA, plasmid DNA, and
RNA
• undissolved material and high-molecular-weight DNA "trapped" at the front edge of the well
• a "cloud" of low-molecular-weight RNA at a position corresponding to 100-200 bp
• presence of high-molecular-weight bands of uncut plasmid in lanes of digested miniprep DNA
Refer to Results and Discussion section of Laboratory 1 “Restriction Analysis of Purified pAMP”, for more
details about interpreting miniprep gels and plasmid conformations.
Remember these three facts when considering possible constructions of the plasmids M1 and M2
resulting from the ligation performed in Lab 1:
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: i.e. a BamHI to a BamHI or
a HindIII to a HindIII. Thus, an intact plasmid must be constructed of an even number of fragments
(2, 4, 6, 8, etc.).
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
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.
The following questions and sample gels are provided to help you interpret the results of your mini prep
results. Once you have interpreted the sample gel, compare your gel with the ideal gels and Follow
Questions 1 through 8 to interpret each pair of miniprep results (digested and undigested).
1. Examine the ideal gel on shown on following page. The first task is to label the size of the fragments
in each lane of the gel starting with the molecular weight marker.
2. Label fragment sizes of, the four bands in AK+ lane located second from the right (AK is the digested
pAMP and pKAN mix) from top of gel to bottom: 3755 bp, 2332 bp, 1875 bp, and 784 bp.
NOTE: Every miniprep must contain the 3755-bp fragment containing the ampr gene and the
1875-bp fragment containing the kanr gene.
3. Locate these bands in the digested miniprep by comparing the M+ lanes (cut miniprep) with the AK+
lane (cut control).
BIO370Y5 2012 LABORATORY # 4 Page 9 of 13
4. Now look for evidence of any other bands in the M+ lanes. Compare the M+ lane with the AK+ lane.
The 2332-bp fragment and/or the 784-bp fragment may be present. If neither of these two additional
bands is present, the molecule is termed a "simple recombinant."
BIO370Y5 2012 LABORATORY # 4 Page 10 of 13
8. To gauge the size of the miniprep plasmid, and therefore which of the options in 5, 6 and 7 above
could be the recombinant plasmid, compare the M- lane (uncut miniprep) with the A- lane (uncut pAMP)
and the L/H lane (A markers). Remember that uncut plasmid can assume several conformations but that
the fastest moving form is supercoiled.
a. Locate the band that has migrated furthest in the A- lane; this is the supercoiled form of pAMP.
b. Now examine the band(s) furthest down the M - lane. If this band and the pAMP band have co-
migrated similar distances, your miniprep is likely a double transformation. The possible molecules
present in a double transformation range in size from 3106 bp (6c above) to 6077 bp (5c,d above),
and thus may appear noticeably lower or higher on the gel than supercoiled pAMP.
c. If the fastest moving band of the uncut miniprep is very high on the gel, your molecule is likely a
superplasmid. Compute the possible sizes of superplasmids composed of three or four fragments.
9. When bacteria are transformed with two different plasmids having related origins of replication, one
of the two plasmids is preferentially replicated within the host cell. Over generations, one of the two
plasmids is eventually lost. Thus, in double transformations with four different fragments, one pair of
fragments should be fainter than the other pair.
10. Based on your evaluation above, make scale restriction maps of your M1 and M2 plasmids.
BIO370Y5 2012 LABORATORY # 4 Page 11 of 13
• If all the restriction fragments are contained in a single superplasmid, all transformants will have dual
resistance.
• If three or four restriction fragments are distributed among separate plasmids, the transformants will
have mixed antibiotic resistance. Matching the observed pattern of antibiotic resistance with alternate
two-gene recombinants can often reveal the structure of the two plasmids involved.
Digesting miniprep DNA with the restriction enzyme XhoI can elucidate some of the structures of
superplasmids and plasmids in double transformations. This enzyme has a single recognition site within
the 1875-bp pKAN fragment and no sites within any of the other three BamHI/HindIII fragments.
Electrophorese XhoI digests of miniprep DNA with samples of uncut pAMP and λ/HindIII size markers.
2. If your miniprep DNA shows three fragments, including the 784-bp fragment (Question 4 in Results
and Discussion), then you may have either a superplasmid with one repeated fragment or a double
transformation.
a. The results of a XhoI digest of a superplasmid will differ according to which fragment is repeated:
• If the 1875-bp fragment containing the XhoI site is repeated, two fragments of 2659 bp and
5630 bp are produced.
b. A double transformation of the simple recombinant and relegated pAMP produces a linear 5630-
bp plasmid plus an uncut pAMP plasmid.
3. If your miniprep DNA shows three bands, including the 2332-bp fragment (Question 5 in Results and
Discussion), then you may have a superplasmid with one repeated fragment or a double
transformation.
BIO370Y5 2012 LABORATORY # 4 Page 12 of 13
a. The results of a XhoI digest of a superplasmid will differ according to which fragment is repeated:
b. A double transformation of the simple recombinant and relegated pKAN produces two fragments
of 5630 bp and 4197 bp.
c. A double transformation of the simple recombinant and ligated 3755-bp + 2332-bp fragments
produces a linear 5630-bp plasmid plus an uncut 6077-bp plasmid.
d. A double transformation of religated pKAN and ligated 3755-bp + 2332-bp fragments produces a
linear 4197-bp plasmid plus an uncut 6077-bp plasmid.
4. If your miniprep DNA contains all four fragments (Question 6 in Results and Discussion), the XhoI
digest will discriminate between a superplasmid and double transformations.
b. A double transformation of religated pAMP and religated pKAN produces a linear 4197-bp pKAN
plasmid plus an uncut pAMP plasmid.
c. A double transformation of the simple recombinant and ligated 2332-bp + 784-bp fragments
produces a linear 5630-bp plasmid plus an uncut 3106-bp plasmid.
5. Make a restriction map of the simple recombinant plasmid using BamHI, HindIII, and PvuI. Prior
experiments showed that the BamHI and HindIII sites are separated by 1875 bp. PvuI cuts the
recombinant plasmid at two positions.
a. Do double minipreps to obtain additional plasmid from a master colony known to contain the
simple recombinant.
PvuI
PvuI + BamHI
PvuI + HindIII
BamHI + HindIII
BamHI + HindIII + PvuI
c. Electrophorese the digested samples on a 1.2% agarose gel, stain, and photograph.
d. The expected number of fragments and their sizes are shown in the diagram on the next page.
BIO370Y5 2012 LABORATORY # 4 Page 13 of 13
6. Using the data from Experiment 5 above and applying a little logic, the relative positions of the
restriction sites can be positioned around a circle to produce a restriction map of the simple
recombinant plasmid.
a. The BamHI/HindIII digest reveals that the BamHI and HindIII sites are separated by 1875 bp.
b. The PvuI digest reveals that the two PvuI sites are separated by 900 bp.
c. The BamHI/HindIII/PvuI digest shows both 1875-bp and 900-bp fragments. This means that the
1670-bp and 1236-bp fragments separate the 900-bp PvuI fragment from the 1875-bp
BamHI/lHindIII fragment.
d. The PvuI/BamHI digest shows a 3545-bp fragment that must be composed of the 1875-bp
fragment plus the 1670-bp fragment.
e. The PvuIIHindIII digest shows a 3111-bp fragment that must be composed of the 1875-bp
fragment plus the 1236-bp fragment.
f. Results d and e indicate that the 1236-bp fragment is adjacent to the BamHI site and the 1670-bp
fragment is adjacent to the HindIII site.
g. Complete the restriction map showing all restriction sites and the distances between them.