Isolation of Plasmid DNA
In addition to their complement of genomic DNA, bacteria often contain self-replicating
elements called plasmids: double stranded, covalently closed circular DNA structures. Usually,
bacteria can grow and reproduce in the absence of plasmids, but their presence may confer
additional useful characteristics such as antibiotic resistance upon the bacteria.
In the plasmid, the DNA is covalently closed, that is, each strand forms a continuous covalently
bonded loop. Since the structure is helical, each strand is also wound about the other and noncovalently bound to it through A:T and G:C base pairs. If the number of turns of helix is the
same as that for linear DNA (about 1 turn per 10 base pairs), the plasmid will appear as a circle.
If the number of turns is greater or less than this number, it can twist into a left- or right-handed
supercoil. Typical plasmids will exist in the cell as super coils. If one chain of the DNA duplex
in a super coiled plasmids is "nicked" (i.e., a phosphate-ribose link is broken), it will relax to form
an untwisted circle. Plasmids differing in the degree of superhelical twisting will have different
frictional properties and can be separated by gel electrophoresis.
Separation of plasmids from genomic DNA during the isolation procedure also depends on
special characteristics of the plasmid structure. If the small plasmid is denatured, as by the
addition of sodium hydroxide in the procedure given below, the two strands cannot separate, but
remain twisted about each other. Upon being returned to acidic conditions they return quickly to
their original structure, a process called renaturation. The genomic DNA, on the other hand, is
large and has been somewhat fragmented during the procedure. Strand segments which were
base-paired in the original structure become widely separated during denaturation. When acid
conditions are restored, they cannot find each other quickly, mismatches occur and the result is
a tangled precipitate which can be centrifuged out.
Another useful characteristic of DNA is its ability to bind the fluorescent dye, ethidium bromide.
A new reagent, SyberSafe® is a non-toxic alternative to ethidium bromide. We will be using this
safer reagent in class. The intensity of the fluorescence of the dye is strongly enhanced when it
binds by intercalation into DNA. Intercalation means that dye molecules are inserted between
some adjacent base pairs in the DNA molecules. Since intercalation causes an unwinding of
the double helix, this also affects the degree of supercoiling of covalently closed plasmids.
Electrophoresis on an agarose gel will separate DNA’ss varying in size and structure. The
separated bands may be located by illumination of the gel with ultra- violet light.
A. Isolation Procedure:
Safety: Wear safety glasses and nitrile gloves.
Caution: Plasmids are isolated from E.Coli cultures and residual culture must
be contained. BIOHAZARD
Disposal: Any item coming in contact E. Coli must be placed in a biohazard
bag for disposal.
Materials:
Qiagen® kit reagents
Sterile tubes and tips
Microcentrifuge
Ice bucket
Overnight culture—E.coli DH5α transformed in LB with amp
1
Qiagen Kit Protocol
Reprinted with permission from QIAGEN® Inc.
1.
Concentrate the E.coli cells (which have been grown overnight in Luria Broth containing
ampicillin) by pipetting 1.5 ml of culture into a microfuge tube and spin at max speed for
1 min. Decant the supernatant into a waste container without disturbing cells.
2.
Repeat #1 two more times, adding the aliquots to the same microfuge tube. Remove the
supernatant completely without disturbing the cell pellet. You now have concentrated
4.5 ml of E.coli culture into a small pellet of cells.
3.
Re-suspend and combine all the cells in 250 μl of Buffer P1 (insure that RNase A has
been added to Buffer P1) and transfer to clean microcentrifuge tubes. No cell clumps
should be visible after re-suspension of the pellet.
4.
Add 250 μl of Buffer P2 and gently invert tubes 4-6 times to mix. Do not vortex, as this
will result in shearing of genomic DNA. If necessary, continue inverting the tube until
the solution becomes viscous and slightly clear. Do not allow the lysis reaction to
proceed for more than 5 min.
5.
Add 350 ml of Buffer N3 and invert the tube immediately but gently 4-6 times. To
avoid localized precipitation, mix the solution gently but thoroughly, immediately after
addition of Buffer N3. The solution should become cloudy.
6.
Centrifuge for 10 min at 13,000 rpm (~17,000 x g) in a tabletop centrifuge. A compact
white pellet will form.
7.
Apply the supernatants from step 4 to the QIAprep Spin Column by decanting or
pipetting. Centrifuge for 60 sec at 13,000 rpm.
8.
Wash QIAprep Spin Column by adding 0.75 ml Buffer PE and centrifuging for 30-60
sec.
9.
Discard the flow-through, and centrifuge for an additional 1 min to remove residual
wash buffer. IMPORTANT! Residual wash buffer will not be completely removed
unless the flow-through is discarded before this additional centrifugation. Residual
ethanol from Buffer PE may inhibit subsequent enzymatic reactions.
10.
Place the QIAprep column in a clean 1.5 ml microcentrifuge tube and cut off the tube
lid. To elute DNA, add 50 l of sterile water to the center of each QIAprep Spin
Column, let stand for 1 min, and centrifuge for 1 min. Transfer to a clean microfuge
tube.
11.
The plasmid DNA is now ready to quantify.
B. Quantification of DNA
DNA in solution can be easily and rapidly quantified using spectrophotometric methods.
The absorbance of DNA measured at multiple wavelengths 260 nm, and 280 nm to determine
a variety of features about the DNA sample. Nucleotides absorb in the ultraviolet spectrum at
260 nm. So the absorbance at 260 nm gives an indication of the concentration of nucleic
acids in the solution. Different types of nucleic acids have different extinction coefficients so
they absorb at different levels. For example double stranded DNA (dsDNA) has a lower
extinction coefficient, therefore it absorbs less strongly than other forms of nucleic acids.
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Because it absorbs less strongly, it takes more dsDNA to equal the absorbance of other forms
of nucleic acids (see Table 1). Note the unusual extinction coefficient units for these nucleic
acids. Thus, if you know the type of nucleic acid you are measuring, it is very simple to
quantify the amount of DNA present in any sample simply by measuring the A260.
Table 1. Spectrophotometric Conversions
nucleic acid
extinction
concentration coefficient
type of Nucleic Acid
A260
(μg/ml)
(ml.cm.μg -1)
ds DNA
1 A260
50 μg/ml
0.020
RNA
1 A260
40 μg/ml
0.025
ss DNA
1 A260
33 μg/ml
0.030
oligonucleotides
1 A260
20 μg/ml
0.050
All measurements require a 1 cm path length
Because DNA is usually purified from cells, it is often contaminated with cellular protein. By
measuring the absorbance at 280 nm (A280), we can determine the amount of protein
contamination. A280 measures the presence of tyrosine, tryptophan and phenylalanine amino
acids in protein. However, nucleic acids also absorb at 280 nm. So some guidelines are
needed to determine when a nucleic acid sample is pure. A sample of pure DNA has a ratio of
A260/A280 of about 1.8. A sample of pure RNA has a ratio of A260/A280 of about 2.0. If your
sample has an A260/A280 ratio that differs significantly from these values, you likely have
contamination by cellular protein.
Quantification Procedure:
1. Generally you will need to make a dilution of your DNA sample to determine the amount of
DNA in any solution. A dilution of 1/25 (10μl in 250 μl water) is generally sufficient to
make your measurements. Note that DNA quantification is accurate only in the absence
of RNA.
2. Add 240 μl of sterile ultra pure water to 10 μl DNA. Using clean quartz micro-cuvettes,
measure absorbance at 260, 280 and 260./280 ratio. The Shimadzu UV 160
spectrophotometer is the best instrument for this purpose.
A260 x 50=ng/μl
3. DNA samples of good yield will be combined by partners.
4. For double stranded DNA, 0.05 g/ l of DNA gives an A260 of 1.0. Using this
information, determine the concentration of your DNA. (Remember that you diluted your
sample! 10L:240L= 25X dilution factor). TA will go over this calculation.
5. If your sample concentration is more than 1 µg/µL, prepare 25 µl of 1 µg/µL in a new sterile
microfuge tube. Typical recovery is 0.3-0.5 μg/μL.
C. Restriction Enzyme Digestion
To verify the quality of the plasmid DNA you have isolated it will be digested with different
restriction endonucleases, individually and in combinations, to generate restriction fragments.
The digestion products will be separated by agarose gel electrophoresis. The nucleotide
fragments will be visualized by staining with SyberSafe®, a non-toxic dye that intercalates into the
DNA and fluoresces under UV light. By comparison with markers of known sizes, you will be able
to determine the size of the fragments resulting from the digestion. By determining the sizes of
the bands, you can generate a plasmid map that graphically shows the structure of the plasmid.
This procedure will also be used at the end of the protocol to verify the sizes of the cloned DNA
fragments.
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Restriction endonucleases will be used to analyze the primary structure of the plasmids you
have isolated. Each of these enzymes is highly specific for a unique palindromic sequence in
double-stranded DNA. For example, BamH1 recognizes the following hexanucleotide sequence,
cleaving at the arrows,
v
5’-XYXG GATCCXXY-3’
3’-YXYCCTAG GYYX-5’
^
yielding fragments of DNA with protruding “sticky” ends. Other restriction endonucleases cleave
DNA to create blunt ends. The recognition sites of restriction enzymes are provided in the
catalogues of the companies that supply these enzymes.
A diagram showing the sites where restriction enzymes cleave a plasmid, and the length of
the resulting fragments, is known as a restriction map. A restriction map can be constructed
either from a knowledge of the sequence of the entire plasmid along with the known cleavage
sites of the restriction enzymes, or by analysis of restriction enzyme digests. Once the map is
known, a specific plasmid can be identified by comparison of the sizes of fragments resulting
from restriction digests with the expected sizes of the fragments from the map. Usually,
combinations of two or more restriction enzymes, used both separately and together in the
same digest, are used to map plasmids. By the appropriate choice of enzymes, this technique
can be used to identify a plasmid.
To determine a restriction map, you will use several restriction enzymes to cut the plasmid
into smaller pieces and determine the size of the smaller pieces. By comparison of the
restriction patterns, you should be able to align the fragments so that you can draw the structure
of the plasmid.
Restriction enzymes are enzymes; and just as other enzymes are sensitive to the salt
concentration and the pH of the buffer, so are restriction endonucleases. They will only function
within a defined range of pH and salt concentration. The temperature at which the reaction is run
is also important for optimal performance. At too low a temperature the rate of the reaction is too
slow. At too high a temperature the enzyme is denatured and thus inactivated.
The isolated plasmid will be digested with the two restriction endonucleases XbaI and XhoI,
separately and both together, generating restriction fragments. Each plasmid will have different
size fragments depending on where it is cut by the endonuclease.
Each of these enzymes is highly specific for a unique palindromic sequence in double-stranded
DNA. They recognize specific hexanucleotide sequences and cleave the DNA at these sites
yielding fragments of DNA with protruding ends. The activity of the enzymes is sensitive to the
salt concentration of the buffer and the temperature.
Materials:
Sterile water
P10 pipetman and sterile tips
Restriction Enzymes (BamHI and ScaI)
Enzyme buffer solution-Multi-core
Microfuge tubes
Heat block at 37oC
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Procedure:
1. Set-up restriction enzyme digestions.
Check your calculations with the Instructor prior to setting up the digestions below.
Using your plasmid DNA, prepare each of the digests following the table.
volumes may need to be adjusted according to your final yield.
Remember,
Restriction Digestion Table
Final volume for all digests is 10 μl.
Tube #
*DNA
(0.5 μg)
L
Rest.Enzyme I
BamHI
L
Rest. Enzyme II
ScaI
L
Buffer
*dH2O
L
L
1
2*
1
1
1
5*
2
2*
1
-
1
6*
3
2*
-
1
1
6*
4
2*
-
-
1
7*
*The use of listed volumes of water and DNA assumes that the DNA concentration is 0.25 μg/μl. If
your concentration is less, you will need to add more DNA and less water accordingly.
Allow all digests to incubate at 37o C in a heat block 1 hr or overnight. Make sure the tubes are
clearly labeled with your initials.
2. Electrophoresis
Agarose gels are the standard media used to detect, separate, and characterize small DNA
molecules by electrophoresis. The technique is simple, rapid, and relatively inexpensive.
Fragments that differ in molecular weight by as little as 1% can be resolved on agarose gels and
as little as 1 ng of DNA can be detected on a gel. For routine analyses of DNA fragments,
agarose concentrations in the range of 0.7% to 1% are most appropriate for separation by
electrophoresis.
The mobility of nucleic acids in agarose gels is influenced by the agarose concentration, the
molecular size of the nucleic acid, and the molecular shape of the nucleic acid. As do proteins,
nucleic acids migrate at a rate that is inversely proportional to their molecular weights.
Therefore, molecular weights can be estimated from electrophoresis results by comparison to
the migration of nucleic acids standard of known molecular size.
Using agarose gel electrophoresis in a mini-gel system, we will analyze the plasmid DNA you
isolated. After electrophoresis, the nucleic acids can be visualized in one of two ways: by
staining with SyberSafe®, a dye that fluoresces under ultraviolet light or by using ethidium
bromide (a carcinogen) which intercalates within the DNA structure. We will use Syber Safe®
for visualizing DNA.
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Materials:
Loading buffer-0.25% bromphenol blue, 0.25% xylene cyanol, 15% Ficoll 400
50X Tris acetate(TAE) buffer- diluted to 1X
1.0% Agarose Gel
Invitrogen1Kb Trackit ladder
SyberSafe®
Balance
Microwave and insulated gloves
Agarose gel electrophesis apparatus and power supply-CAUTION!
Glass tray for containing the gel casting apparatus
Biohazard bags
Safety:
Wear safety glasses and nitrile gloves. The melted agarose is very hot,
wear insulated gloves while handling the glassware.
Procedure:
1. Weigh 0.25 g of agarose into a 125 ml flask containing 25mL of 1X TAE buffer.
Microwave approximately 1 min. to dissolve the agarose. Watch carefully so as not
to boil over. Do not heat longer than necessary to melt the agarose.
2. Wearing insulated gloves, remove from microwave, and swirl gently to insure complete
melting of the agarose. While the agarose is cooling, prepare the gel tray (see #3
below). Cool to 50°C, then add 5 ml of SyberSafe® swirling thoroughly to mix, but
careful to avoid creating bubbles.
3. While the agarose is cooling, prepare the gel tray. The tray must be clean and
completely dry. Using lab tape seal the open edges of the gel tray. The instructor
will demonstrate this technique. It is important to get a good seal. A significant leak
will ruin the gel and it will have to be recast. Use 3 strips of lab tape, making sure
the first strip one rolls under the gel tray edge. Press each overlapping strip securely
in place.
4. Casting the gel: After SyberSafe® has been added and the solution mixed
thoroughly, pour the melted agarose into the prepared gel tray. If bubbles form,
quickly use a pasteur pipet and displace them before the gel sets. COVER
IMMEDIATELY WITH ALUMINUM FOIL TO PREVENT UV LIGHT PENETRATION. It
will take 20 to 30 minutes for the gel to set. The gel becomes opaque and firm to the
touch. Once set, remove the tape and place the gel and tray into the electrophoresis
apparatus.
5. The gel tray must be place in the chamber with the sample wells positioned to run from
negative to positive (BLACK TO RED).
6. Ready the electrophoresis apparatus by pouring 1X TAE buffer into the chamber
until the gel is submerged and air bubbles have been removed. A very gentle shake
of the gel box will usually displace all air bubbles from the wells.
7. Add 2 L tracking dye to the digested DNA samples. Centrifuge briefly to insure
proper mixing.
6
8. Working with one digest at a time, using a P10 and clean tip, load the entire digest
volume into one well of the prepared gel. Make sure to write down which sample you
put in each well. Using a clean tip for each digest, proceed to load all samples into the
wells. The last lane should contain 10 μl of the λ/HindIII standard. (The standard has
been heated at 37oC for 5 min to increase the intensity of the 4.3 kb band.)
9. Secure the lid on the gel box so that the wells are oriented to run from the black
electrode to the red electrode. Plug in the electrodes and turn the power supplies
on. Set the output to run at 90V and monitor progress until the marker dye migrates
to 1cm from the lower edge of the gel which takes approximately 45 minutes to 1
hour.
10. To stop the run, turn the power supply off and remove the lid from the apparatus.
Please, do not pull on the electrical leads.
With gloved hands, remove the gel tray containing the gel and obtain a photo
using the gel-documentation system. The instructor will assist you in taking a
picture of your gel.
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