Restriction Digest of pAMP and pKAN
INTRODUCTION
There are several methods used to analyze DNA. The purpose of this laboratory is to introduce
one method commonly used to analyze a DNA plasmid (circular, double-stranded DNA). The protocol
uses a mixture of two restriction enzymes, BamH I and Hind III, to digest two plasmids and
electrophoresis to separate the restriction fragments. DNA that is cut with restriction enzymes will leave
a specific electrophoresis gel pattern. This restriction fragment pattern should be consistent for any given
piece of DNA. Because of the consistency of cutting, a plasmid can be identified by the pattern of
restriction fragments visible in the gel.
Plasmids are circular pieces of DNA that are naturally found in bacterial cells but have been
modified through genetic engineering to facilitate gene cloning and protein production (expression) in
bacteria. Antibiotic resistant genes have been engineered into these plasmids and function as selectable
markers – that is to say, these genes allow us to select between bacteria that harbor the plasmids from
those that do not. If a bacterium carries a plasmid with an antibiotic resistant gene, the bacterium will be
able to grow and reproduce in the presence of that antibiotic; those bacteria without the plasmid will not
be able to grow. Thus, antibiotics can be used to select bacteria that are resistant, and presumably carry a
plasmid with the resistant gene, from those bacteria that do not carry the plasmid.
Two plasmids will be used in this laboratory exercise. One contains a gene for ampicillin
resistance, ampr, and the other plasmid contains a gene for kanamycin resistance, kanr. The plasmids,
pAMP and pKAN, were engineered by the DNA Learning Center at Cold Spring Harbor Laboratory. The
pAMP plasmid was engineered from the pUC19 plasmid which contains ampr and a 1904 bp restriction
fragment cut from  DNA. The  restriction fragment contains both BamH I and Hind III restriction sites.
The ligation, or joining, of this fragment into the pUC19 plasmid gives the engineered plasmid both
BamH I and Hind III restriction sites. The
pKAN plasmid was engineered from the pUC19 plasmid and an 1861 bp restriction fragment cut from the
pKC7 plasmid. This pKC7 restriction fragment contains both a kanamycin resistance gene and BamH I
and Hind III restriction sites. Study the following restriction maps of pAMP and pKAN.
EcoR I
0/4539
Hind III
234
ori
pAMP
4539 bp
r
amp
BamH I
1120
pKAN
4194 bp
r
kan
ori
Hind III
1904
EcoR I
2116
BamH I
2095
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Because Hind III is a specific restriction endonuclease, it will consistently cut DNA wherever it
encounters the six-base recognition sequence indicated below. The precise location that is cut is called its
restriction site. The DNA molecule consists of two strands of nucleotide building blocks. These building
blocks are oriented in the opposite direction on each strand. Thus, the two strands that make-up a DNA
molecule are said to be “anti-parallel.” For convenience, we can say that one strand is oriented in a 5’
(“five prime”) to 3’ (“three prime”) direction while the other strand is oriented 3’ to 5’. Careful
examination of the restriction sequence will reveal that the sequence of nucleotides is a palindrome; that is
to say, it reads the same on both strands when read in a 5’  3’ direction.

5’…………….AAGCTT……………..3’
3’…………….TTCGAA……...……...5’

Therefore, whenever Hind III encounters this six-base sequence, it will cut the DNA helix between the
adjacent adenine bases. This leaves four unpaired bases forming a “sticky end.”
5’…………A 3’
3’…………TTCGA 5’
Sticky end  5’ AGCTT…………3’
 Sticky end
3’ A………….5’
The recognition sequence for BamH I is:

5’…………….GGATCC……………..3’
3’…………….CCTAGG……...……...5’

Can you figure out the 5’3 prime direction “sticky ends” for BamH I?
MATERIALS
Reagents
pAMP (60 ng/μL)
pKAN (60 ng/μL)
Distilled water dH20
Restriction enzyme mix
BamH I + Hind III
2.5x restriction buffer
Equipment and supplies
P-20 micropipette and tips
1.5 mL microfuge tubes
Minicentrifuge
37ºC water bath
Permanent marker
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METHODS
The plasmids pAMP and pKAN have been cloned in overnight cultures of E. coli (mm294) and
purified using a commercial plasmid purification kit. This purification produces a high yield of
supercoiled plasmid, which increases the efficiency of transformations. The restriction enzymes, BamH I
and Hind III, should always be kept in ice to reduce degradation.
Preparing the pAMP and pKAN restriction digest
1. Obtain pAMP, pKAN, 2x buffer and restriction mix from the instructor. Keep these tubes in ice.
2. Use a marker to label four 1.5 mL microfuge tubes as follows:
A + = pAMP + BamH I and Hind III
A- = uncut pAMP (pAMP without enzymes)
K + = pKAN + BamH I and Hind III
K - = uncut pKAN (pKAN without enzymes)
Include your period and group number on each tube so that you can locate them for the next
lab period.
3. The following reaction matrix summarizes the volumes used to set-up the restriction digest.
Tube
pKAN
pAMP
2.5x Buffer
dH2O
Enzyme
Mix
Total
Volume
K+
4 μL
-
4 μL
-
2 μL
10 μL
K-
4 μL
-
4 μL
2 μL
-
10 μL
A+
-
4 μL
4 μL
-
2 μL
10 μL
A-
-
4 μL
4 μL
2 μL
-
10 μL
4. Add 4 μL of pAMP to tubes labeled A+ and A-. Change the tip and add 4 μL of pKAN to tubes
labeled K+ and K-.
5. Use a fresh tip and add 4 μL of 2.5x restriction buffer to all tubes.
6. Use a fresh tip and add 2 μL of the enzyme mix, containing BamH I and Hind II, to the tubes
labeled A+ and K+ only.
7. Using a fresh tip, add 2 μL of dH2O to the A- and K- tubes. What is the purpose of these tubes?
8. Cap the tubes and use the minicentrifuge to spin down the reagents.
9. Place all four tubes into the 37ºC water bath, and incubate for at least 60 minutes.
10. Following the 60-minute incubation period, the digest can be frozen at -20º C until time is
available for electrophoresis
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Electrophoresis of pAMP and pKAN restriction fragments
INTRODUCTION
It is important at this stage of our experimental procedure that we need to onfirm that Hind III and
BamH I have digested the original plasmids and that we have the correct restriction fragments. Gel
electrophoresis is a procedure commonly used to separate fragments of DNA according to molecular size
or number of base pairs. DNA fragments will migrate through the agarose maze. DNA, because of the
phosphate groups, is negatively charged and will move towards the positive (red) electrode. Because it is
easier for small molecules to move through the agarose matrix, they will migrate faster than the larger
fragments. Picture a group of cross-country runners that are racing through a dense tropical rain forest.
All other factors being equal, the shorter runners will be able to navigate through the tangle of
overhanging vines and dense foliage faster than the taller runners. So, smaller DNA fragments will move
through the tangle of agarose molecules faster than the longer fragments.
We’ll take all of our plasmid samples, digested, and undigested, and use electrophoresis to
separate these pieces. You might have predicted that your uncut plasmids would produce only a single
DNA band; there’s no reason why you would think otherwise. However, it is likely that two or three
bands will appear in the undigested plasmid lanes. This is because plasmids isolated from cells exist in
several forms.
One form of plasmid is called “supercoiled.” You can visualize this form by thinking of a
circular piece of plastic tubing that is twisted. This twisting or supercoiling results in a very compact
molecule; one that will move through the gel very quickly for its size.
A second plasmid form is called a “nicked-circle” or an “open-circle.” Often a plasmid will
experience a break in one of the covalent bonds located in its sugar-phosphate backbone along one of the
two nucleotide strands. Repeated freezing and thawing of the plasmid or other rough treatment can cause
the break. When this break occurs, the tension stored in the supercoiled plasmid is released as the twisted
plasmid unwinds. This circular plasmid form will not move through the agarose gel as easily as the
supercoiled form; although it is the same size, in terms of base pairs, it will be located closer to the well
than the supercoiled form.
The last plasmid form we are likely to see is called the “multimer.” When bacteria replicate
plasmids, the plasmids are often replicated so fast that they end up linked together like links in a chain. If
two plasmids are linked, the multimer will be twice as large as a single plasmid and will migrate very
slowly through the gel. In fact, it will move slower than the nicked-circle. Your uncut pAMP and pKAN
samples then may each have three bands that appear in the gel. Starting closest to the well, you might
observe a multimer, followed by a nicked-circle band and finally, a fast traveling supercoiled band.
We will use a special staining technique that permits us to see the fragments embedded within the
gel, then make a photographic record of your gel to document this important step.
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MATERIALS
Reagents
pAMP (60 ng/μL)
pKAN (60 ng/μL)
DNA size marker (1 kb ladder)
Agarose (0.8%)
2.5x restriction buffer
Distilled water
1x SB buffer
5x loading dye
Ethidium Bromide
Equipment and supplies
P-20 micropipette and tips
1.5 mL microfuge tubes
Minicentrifuge
Electrophoresis apparatus
UV transilluminator
Poloroid camera
Permanent marker
METHODS
1. An 0.8% agarose solution has been prepared and is available in a screw-top culture tube. The
agarose has been pre-measured (about 25 mL) for your gel trays.
2. Be sure the sides of the well tray are in the up position before pouring in the agarose solution.
When the agarose has solidified, about 20 minutes, carefully remove the comb taking care not to
tear the wells. Set the tray into the gel box so that the well-end of the gel is closest to the negative
(black) electrode.
3. Fill the box with 1 x SB buffer to a level that just covers the entire surface of the gel to a depth
of 1-2 mm. Check to see that the gel is covered with buffer and that no “dimples” appear over the
wells. Add more buffer if needed.
4. Determine which samples your team will be loading into the gel: K+ and A+ and ONE of the
following: K-, A-, or DNA marker.
5. Add 2uL of loading dye to each of the tubes containing plasmids (2 or 3, depending upon your
group – be sure to change tips for each tube.
6. If your group is loading the DNA marker, pick it up from your instructor. This sample already
contains the loading dye.
7. Microfuge the tubes.
8. Using a fresh tip for each sample, load 10 uL of each sample into the cells based on the “map”
given in class.
 As you do this, slowly lower the pipette tip below the surface of the buffer directly over, but
not into, the well. Putting the tip into the well can damage the wall of the well or puncture the
bottom of the well. These are not good things to do!
 Use two hands to steady the pipettor. Slowly dispense the sample by pushing to the second
stop of the pipettor. Because of the loading dye, the sample will have a greater density than
the SB buffer. This will allow the sample to sink into the well.
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SB buffer

Important: While holding the button on the second stop, slowly remove the pipette tip from
the gel box. If you’ve loaded your sample correctly, the well will be filled with a blue colored
solution.
9. Secure the cover to the electrophoresis box and connect the electrical leads to the power
supply. Be certain that the anode is connected to the anode (red to red) and the cathode is
connected to the cathode (black to black).
10. Turn on the power and set to 130-135 volts.
11. When the loading dye has moved to within 1 cm from the far edge (+end) of the gel, turn off
the power supply. Unplug the electrical leads from the power supply by grasping the plugs, not
the cords. Remove the cover to the electrophoresis chamber.
12. The instructor will remove the gel and stain the gel,
13. Take a photograph of the gel on the UV light box, and include the photo in your lab write-up.
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