INTRODUCTION TO NUCLEIC ACIDS AND MOLECULAR CLONING
References:
 E.coli: Cellular and Molecular Biology: F.C. Neidhardt ed.
Volumes 1,2. (1987)- a detailed reference on things E.colial.
 www.slic2.wsu.edu:82/hurlbert/micro101/pages/101hmpg.html
an excellent review of most of the things we will do in the last
section of the lab. A MUST-READ-BEFORE-DOING-ANEXPERIMENT website.
 Voet, Biochemistry, pp916-918, 932-933, 897-902.
 Jacobs, “Life in the Balance- Cell Walls…” Science, 278, 1731
(1997)
The next series of experiments will be devoted to the nucleic acids. In them se
we are going to learn some of the basic techniques that are used by
biochemists who study nucleic acids. At one time, nucleic acid research was a
separate branch of the field of biochemistry practiced mainly by
microbiologists and a few hardy organic chemists and physicists. Researchers
could and did study enzymes without knowing much about how to work with
nucleic acids, except perhaps how to get rid of them in the early steps of
enzyme purification. This is no longer so. The advent of recombinant DNA
technology, a technology which includes the discovery of restriction
endonucleases i.e., the ability to cut at specific DNA sites, thus enabling us to
cut out defined chunks of DNA, the discovery of methods of inserting foreign
DNA into other organisms and also of having them expressed in quantity, has
given biochemists a useful and general way of producing large quantities of
almost any DNA, RNA and protein they want to study. It has also provided a
way to produce modified enzymes on demand. This makes it possible to test
models for catalysis, for example, by making an enzyme that lacks a particular
amino acid in the active site.
The experiments in the DNA section of the course cover some of the
basic methods used in recombinant DNA work. Your instructors have not been
able to reduce the content of some of recombinant DNA protocols so that they
can be completed in a single afternoon. Therefore, you may sometimes be
required to come in early to get an experiment started. Labs sometimes
stretch over several lab periods so you may be required to come in on an off
day to get things ready for the next session. We have, however, been able to
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adapt a good cross-section of useful techniques to a suitable time frame and
those of you who may be going on to do further work in biochemistry should
find this a useful exercise.
Plasmids
Chromosomal DNA contains the biological information we need. Chromosomal
DNA is difficult to obtain in pure form and, if allowed unrestricted
transcription and translation would produce a large mass of biochemical
products. Thus, if we want to use chromosomal DNA as a source from which we
produce our material of interest, we must painfully remove the very small
fraction of the components of interest from a larger mass of irrelevant
products with very similar chemical and physical properties. Historically, the
solution to this problem has been to study smaller pieces of DNA as a
substitute for chromosomal DNA. Ideally, small pieces need to be constructed
so that they not only contain the component we want to examine, but also DNA
components that will allow them to be replicated by cellular machinery. An
example of such a piece of DNA is called a plasmid. Plasmids are
extrachromosomal, semi-autonomously-replicating pieces of DNA that occur
naturally in bacterial cells. They allow genetic information to be transferred
both vertically and horizontally. Plasmids are circular, and contain genetic
elements that allow replication in a suitable host by using the enzymes and
energy provided by the host. In short, plasmids are molecular parasites. Like
viruses they use cells for propagation of the species. They usually provide a
benefit for the host by carrying genes that might confer selective advantages
to the host. Some of these selective advantages are providing genes for
antibiotics resistance, for trace element metabolism, or that enhances
disease-causing ability.
One of the most widely used plasmids in the laboratory is called
pBR322. It is a circular bit of DNA of about 4600 base pairs. The designation
"pBR" derived from the first letters of "plasmid Bolivar Rodriguez" the latter
two letters being the initials of the constructors of the plasmid-Bolivar and
Rodriguez. pBR322 is derived from a much larger natural plasmid, and has
been stripped down to the bare minimum of sequence required to carry out its
laboratory functions. This plasmid has two protein-coding genes, one that
confers resistance to ampicillin, a form of penicillin, while the other confers
resistance to tetracycline. The presence of these genes allow bacteria which
carry them, to grow on nutrients containing those antibiotics, whereas ordinary
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bacteria lacking that gene will not grow. This process of growing bacteria
under conditions where only a desired genotype is viable is called “selection”.
We will make extensive use of selection to isolate recombinant species of
interest. There is also an origin of replication, which governs plasmid
replication. Plasmids such as pBR322 are replicated by a "relaxed" or “low
stringency” mode. These plasmids can replicate without host protein synthesis
requiring only DNA polymerase I activity. “Relaxed replication” plasmids are
present in about 10-100 copies per cell and are passed to each daughter cell as
the parent cell divides.
Unlike most natural plasmids, pBR322 and its derivatives have no system
for spreading horizontally through the bacterial population; they cannot be
transferred by bacterial conjugation. The use of non-conjugative plasmids is a
sort of a safety feature, so those antibiotic-resistant traits cannot easily and
inadvertently, be transferred to non-laboratory strains of bacteria. The only
way to get the DNA into a new strain of bacteria is through deliberate
manipulation in the laboratory.
Another useful feature of pBR322 derivatives is that only a single type
of recombinant plasmid can stably exist in one cell at any one time. If we
introduce mixture of pBR322 and another plasmid, such as pIH1034 with the
same origin of replication, into a population of bacteria, and we pick a single
bacterial cell to grow up (easily done, with standard bacteriologic techniques),
we will find that the cell we choose will have only one of the two plasmids.
a.
The plasmids used this semester are both derivatives of pBR322.
They have been made by modifying pBR322 in different ways.
Commercial plasmids contain a sequence of DNA, which allow the
insertion of foreign DNA into the plasmid in such a way that the
replication of the plasmid is not affected. These sequences are called
“Linker” regions or “multiple cloning sites (MCS)”. These foreign
sequences are then carried along as silent passengers as the plasmid
replicates and can be translated by the host's translation systems.
Thus plasmids, with independent replication and translation, can
provide large amounts of almost any piece of DNA or its gene
products, which we have cloned into it. the isolation experiment. The
addition of the fusion protein gene raises the Mr of the pIH1034 to
about 11 kbp.
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Bacterial Hosts
The bacterial host we will use in our experiments is a laboratory strain of
Escherichia coli, E. coli. E.coli is a "gram-negative" bacterium that normally
grows in the human gut and is the best-studied of all micro-organisms. Some
strains are modestly pathogenic, although most laboratory strains are believed
to be completely safe, because they are significantly disadvantaged relative to
the wild bacteria already in your body. In spite of the fact that laboratory
strains are non-pathogenic, you must follow "good microbiological practice" in
handling live cells because you do not want to contaminate your experimental
set-up. All waste should be placed in containers for sterilization before
disposal, and you should wash your hands and the bench-top at the end of the
day. You must not mouth pipette when handling cultures or other material that
potentially contains living cells. We also take these precautions to protect
ourselves in case an unwanted contaminating strain of bacteria entered our
cultures from the environment.
Restriction Endonucleases and formation of Recombinant DNA.
Restriction endonucleases are enzymes derived from a variety of bacterial
species. Their normal biological function is to prevent foreign DNA from
subverting the cellular machinery. They do this by chopping foreign DNA to
pieces. Host DNA is protected because host-specific methylation enzymes
modify it, so that they are not substrates for the host's restriction system.
Thus this specific methylation prevents host endonucleases from destroying
the host's own DNA.
Restriction enzymes are enormously useful in the laboratory. In
general, DNA in the laboratory will lack the strategic, species-specific
methylations, and so will be cut to bits. In a given piece of DNA, the cuts
will always be in the same place, at fixed sites corresponding to recognition
sequences three to six base pairs in length. These recognition sequences are
the restriction sites for the enzyme. There are some restriction
endonucleases that don't cut at fixed places but we will not run across them
in this course.
Most restriction sites (all the ones in our experiments) are
palindromic. This means there is a center of symmetry. Because of this
palindromic symmetry, the bottom strand reads exactly the same as the top.
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The sites of actual cleavage are offset, by three base pairs in the example
below. The consequence of this is that the ends are cohesive or sticky i.e.
has complementarity with the other fragment produced by the hydrolysis.
Even after they have been cut, they have some tendency to be held together
by base pairing. This tendency is not enough to hold them together under at
room temperature, but, at lower temperature, this cohesion provides enough
net stabilization to greatly aid the joining of free ends by the DNA ligase.
Any piece of DNA cut by the same enzyme will be also be cohesive with the
cut plasmid. This is what allows us to readily create recombinant DNA by
being able to join two different DNA’s cut with the same restriction
endonuclease. The pIH1034 plasmid was created by ligating two ends of the
chromosomal lacZ gene to complementary ends i.e. cut with the same
restriction enzyme, in the vector pMAL-c2 in such a way that the LacZ DNA
was in the reading frame of the upstream malE DNA. This ensured that the
insert would code for a fusion protein containing an amino terminal malE
protein fragment covalently linked to an active -galactosidase.
5'-GAATTC-3'
3'-CTTAAG-5'
EcoRI
5'-G-OH
+
3'-CTTAA-O-P
POAATTC - 3'
HO-G-5'
The recognition site for the restriction endonuclease EcoRI. Note that the
sequence is palindromic; the last three nucleotides are complementary to the
first three. Cleavage sites are indicated by the arrows. All EcoRI-cut ends
have a three-base 5' overhang, and because of the symmetries, all ends from
all EcoRI sites are cohesive.
Fig: 30: EcoRI Hydrolysis of DNA
Restriction enzymes are named according to an agreed scheme. The
first letter corresponds to the genus, and the second two to the species of
bacteria from which the enzyme is derived. There may be additional letters
or numbers to indicate a particular strain. Some bacteria contain several
5
restriction enzyme activities with different specificities, so the final part of
the enzyme name is a Roman numeral indicating whether it is the first, second,
or third enzyme in the particular strain. As examples, the enzymes you use in
this course may include: Pst I derived from Providencia stuarti, Pvu II from
Proteus vulgaris and Bgl I from Bacillus globigii.
Many of your experiments will involve the enzyme -galactosidase. This
enzyme is encoded by the lacZ gene of E. coli. lacZ is part of the Lac operon
which encodes several genes required for the metabolism of lactose. Those of
you who have completed a course in molecular genetics will know that the
genes of the lac operon were among the first E. coli genes to be characterized
in detail. Many of our ideas about bacterial gene organization, transcription,
and repression were originally formed through study of the Lac system.
-Galactosidase is a tetramer of three identical 116,000 Da (Da is the
IUPAC symbol for g/mole) subunits. It catalyzes the hydrolysis of the
substrates with galactose at the non-reducing end of the molecule. Among the
most useful of these synthetic substrates is bromo-chloro-indolyl galactoside
(abbreviated, thankfully, as Xgal). Upon hydrolysis, the bromo-chloroindolyl
moiety (the "X") turns blue and precipitates from solution. Thus, a colony of
bacteria expressing -galactosidase will turn an attractive shade of blue when
grown on nutrient agar containing X gal. You will make use of this phenomenon
in the first experiment when you use this blue color to identify the clone
containing -galactosidase.
In the first set of experiments in the lab involve proteins; assaying,
characterizing and isolating them. We will learn to operate and understand the
instrumentation used in these processes and to generate some of the
important numbers connected with these processes.
In the next set, the DNA section we are going to learn some basic
bacteriological methods, including sterile technique, preparation of liquid and
solid media, isolation of pure cultures, and use of selective and indicator
media to isolate a given bacterial genotype. We will also isolate plasmid DNA
from a selected clone, a method called the miniprep method-a quick method
for isolating DNA from small volumes of cells. The miniprep method usually
yields microgram quantities from 5ml of cells. We should isolate enough
plasmid DNA in this small volume to provide for our plasmid DNA needs for
the entire semester.
In experiment eight we show you how to "transform" an appropriate
host, that is introduce your plasmid into a plasmidless (wild type) E. coli strain
and show that the transformed strain expresses the plasmid phenotype.
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Experiment nine, part A, sets the stage for your Southern blot
experiment. In this experiment you will label a lacZ DNA fragment by PCR.
This labeled DNA will be used later as a probe for part B, the Southern blot
proper. In part B you will use probe will be used as a reporter molecule for the
presence of the lacZ gene in a chromosomal DNA mixture.
Experiment ten, our last canned experiment before the independent
projects, is one in which we transcribe some DNA to produce a self-splicing
RNA. We will then use a polyacrylamide gel to look at the kinetics of the selfsplicing reaction.
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Experiment 6 - Selecting a Clone.
Reading in Handouts:
 Genotypes and Phenotypes - Brown "DNA Sequencing".
 E.coli growth Medium - Russel Hopper, Gen. Eng. News.
 WSU website mentioned before — see bacterial growth and
selection sections.
Overview - You will be given aliquots of pure cultures from three E.coli strains.
These had been grown overnight and diluted out so that single cells could be
formed on plating out. These single colonies, or clones, consist of billions of
cells grown from the single cell are easily visible on the agar surface. You are
going to check out the phenotype of the unknowns and select the pure culture
containing the plasmid you have been assigned. Clone selection will be based on
the fact that the phenotype is based on expression of all genetic information
contained in the cell. One of the strains will be a wild type containing only
chromosomal DNA while the two other strains will contain, in addition to
chromosomal DNA, a plasmid. The wild type can express only proteins coded for
by the chromosomal DNA, while the plasmid-containing strains will show a
phenotype determined by the genetic information expressed by both
chromosomal and plasmid DNA, and as the plasmids contain sequences which
allow them to survive in some antibiotic-containing environment, we can
distinguish one pure culture form the other by inspecting their viability in our
selection media. Thus we need to find out the genotype of the host strain and
that of your plasmid and, by adding these genotypes, predict the phenotype
expected of a host cell containing that plasmid. Information in the lab
handouts, manual and in lecture will describe common bacterial genotypes and
phenotypes. These sources of information should enable you to select your
particular clone.
What we will do Today
A. Prepare sterile agar plates for the transformation experiment.
B. Prepare sterile equipment for the transformation experiment.
C. Select single colonies for Plasmid isolation.
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A - Preparing Solid media and Sterile Techniques
Preparing Solid Selective Medium. We have provided you with all the plates
and supplies needed for today's experiments. Your job will be to make your
own plates for the transformation experiments to be done in a couple of weeks.
A group, the students on one side of each aisle, usually two or three students,
will cooperate in the making of a set of plates.
We are trying to grow a defined culture and so the only non-sterile
component is the bacteria we introduce into our medium. Everything else
needs to be sterilized. If things are not marked as sterile, don't use them.
Unlabeled equipment lying about are generally not sterile. Use sterile pipettes
or pipette tips to introduce the antibiotic into the autoclaved solutions.
Each group will make up 400ml of LB (Luria-Bertani) medium in a 600ml
Erlenmeyer flask following the recipe in the reagents section at the back of
the lab book. Add enough agar to give a final concentration of 1.5% (w/v). The
1.0% formulation of a solution means that 1.0 g of solute is dissolved in
enough solvent to give 100 ml of the final solution. Place a stirring-bar in
your Erlenmeyer and cover with aluminum foil cap. The stirring bar will be used
to disperse the antibiotic solution we will add later to the sterilized medium.
Agar is not soluble in water at room temperature so it needs to be autoclaved
for dissolution. The autoclave not only dissolves all solids, but also sterilizes
the resulting solution by raising its temperature to 120°C at a pressure of
about 2 atmospheres. A time setting of 30 minutes will sterilize most
solutions. Your instructors will show you how our autoclave works. It might be
necessary to coordinate with other groups, as autoclave space is limited.
After autoclaving is done, remove the Erlenmeyer using the thermal
gloves and eye protection and allow the solution to cool down in a water bath to
about 65°C before adding ampicillin. The ampicillin, at a final concentration of
50 g/ml allows for the selection for the clones of interest, i.e. those able to
grow in the presence of ampicillin. As too much antibiotic will kill all the cells,
we need to calculate how to add enough ampicillin to allow resistant cells to
grow while killing only sensitive cells. So while the solutions are cooling
calculate the volume of stock 50mg/ml ampicillin to be added to your 400 ml of
cooling LB-agar solution to give a final concentration of 50 g/ml. The
ampicillin is heat-sensitive so we have to cool the solution before adding the
agar. While waiting for the agar to cool, mark a set of Petrie dishes by running
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a marking pen down its side to make a single hash mark. In this lab no hash
mark means a plain LB plate, a single hash mark indicates an LB-amp plate, and
two hash marks means a tetracycline plate.
The formula for the ampicillin dilution calculation is one you last used in your
freshman year in General Chemistry
CiVi = CfVf
Ci = initial concentration
Vi = initial volume
Cf = final volume
Vf = final volume
Our stock (initial) concentration of ampicillin is 50 mg/ml.
Mix the amp-agar solution gently on the stirring plate and pour the plates
quickly. Pour each plate by removing the lid, retaining it in your hand, and
dispensing about 30 ml of the agar solution into the dish. Replace the cover
and allow the plates stand out overnight to set and dry. Do not throw the
sleeves away, they will be used to store your amp plates. Also cut off the top
of the sleeves neatly to remove the sterile Petrie dishes, don't just rip the
sleeve apart.
B. Preparing Sterile Equipment for the Transformation experiment
This will be done while we are doing other things. This sterile equipment
will be used in the transformation of a wild type E.coli in a few weeks time.
Each group of students - group is defined as two students, one assigned a
pUR290 plasmid and the other a pAtRNA-1 plasmid, - will combine to sterilize
enough equipment for their own use. Each group needs to sterilize:






1.7ml eppendorff tubes.
0.7ml Eppendorff tubes.
Spreaders.
Yellow pipette tips.
Blue pipette tips.
50 ml Erlenmeyer tubes.
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C. Selecting Clones.
This experiment will be done in groups of two.
Reagents:




Sterile Spreaders.
Bring gloves for this experiment.
Three sets each of solid medium selection plates containing
either, no antibiotic, just solid rich medium, (no hash marks); an
ampicillin-Xgal mix in solid rich medium, (one hash-mark) or
tetracycline in solid rich medium, (two hash-marks).
A set of unknown cultures labeled A, B and will be provided.
Attach group numbers to the unknown numbers in your lab book
as the unknowns will be scrambled so that each group has a
different allocation i.e. one groups A, B and C will be another
groups’ C, A and B. There are 6 groups and three scrambled
unknowns so be sure to record your group number in your lab
book. The unknown are all 106-fold dilutions of overnight pure
cultures.
Protocol
 Pure cultures stored at -70C are revived in LB-amp liquid medium and
grown overnight.
 They were then diluted out 106-fold in fresh LB-amp liquid medium and
distributed to the groups. A 106-fold dilution should be enough to
allow the formation of single colonies.
 Each culture is to be plated out on each of the selective solid media.
There is thus a set of selection plates for each unknown.
 Plate 50L of unknown A onto the surface of each member of the
selection plate set.
 Repeat for unknowns B and C using a fresh set of selection plates for
each unknown.
 After plating, allow the plates to stand, lid-side up for about 15
minutes.
 Tape the three plates together, about 5 inches of tape along one side
will do. This is just to keep them together during incubation and
storage.
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
Place the plates, lid-side down, in the incubator by the west door in
lab M227. This has been set to 37C. Tomorrow morning I will take
the plates out and store them in the refrigerator until the next lab
period, when you can score them.
What is happening in the Plates
The presence of the antibiotics tetracycline or ampicillin in a medium, allows
for selection of bacteria with plasmids containing the relevant antibiotic
resistance genes. Xgal is a substrate for -galactosidase and allows for the
detection of cells containing an active lacZ gene and IPTG is an inducer of the
LacZ gene. The induction of the lacZ gene makes for a more sensitive assay
for cells containing this gene. Some of these induced cells die and the
-galactosidase leaks through to the external medium where it hydrolyzes
Xgal. This hydrolysis produces an insoluble blue product around colonies
expressing the lacZ gene and allows us to identify these colonies. Even with
induction it might take more than an overnight incubation to produce enough
Xgal product for detection.
The hydrolytic reaction is as shown in the figure below. The final
product, the one allowing visualization, is the same insoluble indigo product we
saw in the Western blot.
Fig. 31: X-Gal - Another Substrate for -Galactosidase.
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Marking and Filling Plates for the Clone Selection
Use Sharpies, permanent markers, for all labeling. You should get your own
sharpie for the semester, the lab does not provide these.
Label each plate with your name. The general rule for Petrie dishes: all labeling
is done on the bottom of the plates, not on the lids, because lids are easily
mixed up. Place all contaminated spreaders in a container and autoclave them
along with the biological waste before you go.
g
III: Scoring Plates and Selection of Clones.
I will remove your plates for the incubator and store them for you in the
refrigerator so you can score them next time. “Scoring” means making a table
of clone number versus behavior on the different media (usually a “+” to
indicate growth, a “-“ to indicate no growth and a “?” to indicate uncertainty).
From this information, you will be able to deduce what plasmid, if any, each
clone carries. If in doubt, consult your instructor. But before consulting the
instructor, make some effort to understand the information on your own.
Use the following table to record the data from this experiment. The plasmids
do not correspond to the phenotypes shown. The data are entirely fictional, the
form for data entry is the point.
Unknowns.
A
B
C
LB
+
+
+
Amp/Xgal/IPTG
+
+
?
Tet
+
Plasmid
Wild type
Plasmid I
Plasmid II
Lab report


Count the number of colonies on each plate and record the results in a
table as above.
Calculate the number of colonies in the original over-night culture,
before dilution. See pages 108-109 of the lab book.
13

Identify, if possible, all the unknowns. Construct a second table with
unknown letter, your identification and the genotype expected for all
selective conditions. Be sure to explain (briefly, almost
monosyllabically), in a separate paragraph, your identifications
criteria, including why some unknowns could not be identified (if this,
in fact, turns out to be true). The information for your identifications
depends upon you applying the knowledge that the phenotype of each
strain is the sum of chromosomal and Plasmid DNA information
expressed in the selection media. The wild type, E.coli strain C90, has
no antibiotic resistance.

Articulate your choice of clones "Clone number “x” was selected for my
plasmid prep because….".
Highlight and identify the entry on the list that you would use for your
plasmid prep.
Note any unexpected phenotypes and rationalize their appearance.
Design a procedure for isolating and identifying separate strains from
a common mixed liquid culture.



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Experiment 7 :Miniprep Isolation of Plasmid DNA
Reading:
 P.Serwer, Agarose Gels, Electrophoresis, 1983, 4, 375-381.
 Voet, Biochemistry, 862-870.
It has been known since the 1920’s that living cells could be
“transformed” by other cell, even “dead” other cells. Griffith’s
demonstration that extracts of killed virulent pneumococcus strain could
“transform” a live avirulent strain into a live virulent strain paved the way
for a scientific examination and explanation of the phenomenon. Some
twenty years later the work of Avery, Macleod and McCarty capped
Griffith’s discovery by proving that “transforming” material was DNA. In
experiment 8 we are going to follow in Griffith’s footsteps and use
exogenous DNA to change the genotype of an E.coli strain. In this
experiment, experiment 7, we are going to isolate the DNA for that
transformation. The genetic information we are going to add, in one case, is
an expressible LacZ gene, and in the other case, information coding for selfsplicing RNA. The hard work of getting these sequences out of the
ancestral chromosomal DNA and into a conveniently expressible form has
already been done for us. Our source for these genes will be plasmids
contained in an E.coli strain called TG2. All we have to do is allow the strain
to divide and isolate the plasmid produced. We need only picograms of
plasmid to transform a significant number of bacteria, but we do need
relatively pure DNA, so that we can limit the information we add to the cells
genetic repertoire to only that contained in the plasmid. What we do today
will allow us to approach the transformation experiment with a
characterized DNA and thus be able to specify our transformation
conditions. Toady we are going to:
A. Isolate plasmid DNA from an overnight culture.
B. Characterize the plasmid by agarose gel electrophoresis.
C. Quantitate plasmid recovery by a fluorescent DNA assay.
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A. Plasmid Isolation
At first glance, plasmid isolation looks like a difficult task, as we need
to separate the plasmid DNA from all the other unwanted components of the
cell, some of which have similar properties to the plasmid to wit,
chromosomal DNA. Other cellular components are low molecular weight and
could be difficult to remove from a plasmid solution, and could easily
interfere with the transformation. However the situation is less complicated
than it looks at first glance. We are going to use a commercial kit called the
Qiaprep-spin plasmid purification kit that will allow us to isolate pure plasmid
DNA rapidly and efficiently from the bacterial mix. Plasmid isolation is a
standard technique in all biochemistry labs and our method is just one of the
many simple and effective commercial methods for plasmid purification.
Background for Qiaprep-spin Plasmid Isolation Protocol
This kit has been specially designed for the rapid isolation of plasmid DNA
from cell cultures and the secret of its success is a silica matrix to which
nucleic acids will bind specifically. Thus all we really need to do to isolate
plasmid DNA is to break open the cells to release the DNA, treat the
cytoplasmic solution, allow the DNA to bind to a silica filter, wash off all
non-binding components and then wash off the pure DNA. All in all, this is a
very simple procedure.
 The initial step in the isolation involves resuspending the cell pellet
in a resuspension buffer containing RNase. The RNase is added to
hydrolyze the RNA released when the cells are lysed. RNA has
similar physical properties to DNA and would tend to be coprecipitated with DNA if no precautions are taken. So we use a
chemical difference- susceptibility to RNase- to separate RNA
from DNA. RNase is a very stable enzyme and is not irreversibly
denatured even under the harsh condition used to break the cells
open. It is thus ready to remove RNA when conditions allow.
 The cells are then lysed under alkaline conditions, a process
naturally called alkaline lysis. Alkaline lysis is the most useful of
several methods used to release DNA from intact bacteria. It is
quick and gives good yields of relatively clean DNA. Alkaline lysis
involves treating a cell suspension with a lysis solution containing
SDS (Sodium Dodecyl Sulfate or lauryl sulfate), EDTA and sodium
16



hydroxide. The SDS along with EDTA, added in the resuspension
step, weaken and disrupt the E.coli cell walls to allow release of the
cytoplasmic contents. SDS also functions to denature released
proteins. At the same time the sodium hydroxide works to
denature DNA partly hydrolyze RNA and denature protein. These
reactions serve to convert some components to a form called an
insoluble precipitate that facilitates their removal from the
plasmid solution.
Next, we neutralize the alkaline solution by adding a neutralizing
solution containing guanidine hydrochloride (GuHCl), a chaotropic
salt. On neutralization the small circular plasmid DNA re-anneals
rapidly and remains in solution. The large, more complex,
chromosomal DNA cannot re-anneal rapidly enough and, along with
the denatured proteins and cell membrane components, form a
dense insoluble network which can be pelleted by centrifugation.
The supernatant thus contains the soluble, mainly plasmid, DNA
and the pellet contains the unwanted cellular debris.
The next step is a binding of the DNA to a silica membrane. It has
been shown that, under conditions of high concentration of
chaotropic salts, DNA will bind selectively to silica surfaces. The
mechanism of this binding is unknown. Small non-binding DNA
fragments and proteins do not bind to silica and are washed off
the membrane. So filtering the solution through the silica pad
allows the retention of the bound DNA.
The retention step is followed by a series of washing steps to
remove unbound or weakly bound material from the filter. Washes
containing buffer, chaotropic salts and ethanol remover everything
we want to remove and we are left at the end of the washing steps
with only DNA bound to the filter. The DNA is then washed with a
TE buffer. This serves to remove, or elute, the DNA. The plasmid
is now in a small volume of TE in a clean container and is ready for
analysis.
Care and Feeding of the Pipetman
Today you will be introduced to the use of an automatic pipettor for
handling small volumes (microliters) of reagents. These volumes are
probably much smaller than anything you’ve handled in previous courses. The
17
small volumes used in molecular biology reflect the very small amounts of
material that are actually needed for the experiment as well as the very
expensive cost of almost all the reagents. Most of you will use the
PipetmanTM brand of automatic pipettor. These are a basic tool of the
molecular biology laboratory. Your instructors will teach you how to use the
Pipetman. If any of these pipetmen are lost, everyone sharing in their use,
will be apportioned their cost for the replacement (about $200). If you
have not used a pipettor before, you should listen carefully to a description
of its use. These instruments are a vital part of the protocol and should be
cared for and used as instructed. They are quite expensive to buy and
maintain, as well as being fragile. When you had done with them for the day
be sure they are returned to where they were found before you leave the
lab.
To keep your sanity, remember that the amounts of DNA we are
isolating in today’s experiment are very small and cannot be seen by the
naked eye. These first few experiments will allow one to become familiar
with working with molecular biological amounts of reactant and should
impress one with the role of faith in science.
The second part of today’s lab is the characterization and
quantitation of your plasmid. Characterization means to determine the
purity, restrictability and molecular weight of your plasmid. You should read
the assigned reading s on restriction enzymes and gel electrophoresis
before coming to class. Electrophoresis will be done in a submarine gel
electrophoresis unit using agarose as the sieving material. After resolving
the DNA bands we will visualize them in the gel by a dye-binding method.
For this experiment we will provide the gels and all other necessary
materials. In later experiments, you will have a chance to make these
yourself. While you are electrophoresing your DNA you are going to learn to
use the spectrofluorimeter to determine DNA concentration.
Plasmid Miniprep Protocol.
Our isolation protocol is based on that suggested by the manufacturers of
the Qiaprep-spin columns.
1. Pellet about 3.4 ml of the overnight culture in an eppendorff tube by
filling the tube with 1.7 ml of culture, microfuging for 1 minute, decanting
and discarding the supernatant, and repeating the process with another
18
1.7 ml of the culture. Be careful to remove all the excess liquid from the
top of the pellet in both centrifugation steps.
2. Add 0.250 ml of resuspension buffer (P1) to the pellet and resuspend
using a pipetman. Resuspend thoroughly so that there are no clumps.
Clumps will reduce your yield of plasmid.
3. Add 0.250 ml of lysis buffer (P2) and mix by inversion. The suspension
should clear as the cells lyse. Allow 5 min at room temp for lysis.
4. Add 0.350 ml of neutralization buffer (N3) to neutralize the solution.
This contains guanidinium.HCl, GuHCl, an irritant, so wear gloves and eye
protection in this step.
5. Microfuge for 10 min.
6. Place a Qiaprep-spin column in a 2 ml microcentrifuge tube and add the
supernatant from the previous step to the column.
7. Microfuge for 60 seconds to remove unbound material.
8. Wash the column with 0.5 ml of wash buffer (PB). This also contains
GuHCl so wear gloves. The volumes of the various washes will exceed the
volume of the collection tubes so, after each wash, decant the spent
solution into the waste container. Use a fresh collecting tube.
9. Wash again with 0.75 ml buffer PE and microfuge for 60 seconds.
10. Microfuge for a further 2 minutes seconds in a clean collecting tube to
remove the traces of last (PE) wash solution.
11. Place the Qiaprep column in a clean collecting tube and elute DNA by
adding 100l of Elution buffer (EB), a TE buffer, and microfuging for one
minute. Transfer the plasmid solution to an eppendorff tube.
12. Label the eppendorff containing your plasmid and keep the tube in the
assigned container in the freezer when not in use.
19
B: Analysis of DNA by Agarose gel electrophoresis
The basis of the electrophoretic method is differential migration
based on differences in the mobility of the migrating species. Charges will
migrate in an electric field toward the electrode of opposite charge. Cations
will move toward the cathode (negatively charged electrode) and anions move
toward the anode (positively charged electrode). On the electrophoresis
unit, the anode is traditionally marked in red and the cathode in black. In
the field alone, in the absence of any frictional effects, the migration
distance is dependent on the charge/mass ratio of the molecule. When both
of these parameters are independent, electrophoresis can be a sensitive
discriminator of molecular mass and/or shape. However DNA has a constant
charge/mass ratio, each nucleotide being of roughly the same mass and
having one negative charge at the pH of electrophoresis. Thus in a medium
which offers no resistance to migration, no differentiation of differentsized DNA molecules should take place. However, in agarose, a cross-linked
galactose polymer with a continuum of varying pore sixes, the DNA has to be
pulled by the field through the various pores of the sponge toward the
cathode and thus encounters a resistance which depends on its molecular
volume or size. The smaller molecules move through the agarose matrix
meeting little resistance and migrate faster i.e. larger distances in a give
time. Larger molecules move with more difficulty and migrate smaller
distances. Thus migration is inversely proportional to mass. The basic unit
of the agarose is the disaccharide consisting of the 1,3-linked -D galactose
and the 1,4-linked 3,6 anhydro--L galactose as shown in the figure below.
Fig. 33: Agarose unit
20
Pouring the Gel
Agarose is a linear polymer of the above disaccharide. An average agarose
molecule is 400 units long and is consequently insoluble in aqueous solutions
at lower temperature. It, however, dissolves at 90°C. As it cools, it forms a
porous gel consisting of hydrogen-bonded networks of agarose helices and
unordered agarose chains, through which the DNA must move.
Gel Preparation

Set up the gel unit as shown in the figure below. Note the small gap
between the bottom of the comb and the cradle surface. This allows the
formation of a well in the solidified agarose.
Figure 34: Setting up Agarose Gel





Add 0.5g of agarose to 50 ml of 1X TAE buffer in a 100ml Wheaton
bottle. This makes a 1% agarose gel which will separate DNA fragments
ranging in size from about 8000 bp to about 1500bp.
Add 3L of ethidium bromide and heat in the microwave for 1min and 20
seconds.
Cool under the faucet for about 30 seconds and pour into the gel unit.
Allow cooling for 30 minutes before using.
Remove the container and place in the transfer unit. Remove the comb
only when electrode buffer just prior to use, covers the gel.
21
The figure below shows the arrangement of a gel during the run.
Fig. 35: Agarose gel electrophoresis.
Preparing Restriction Digests




Add 10 L of your plasmid prep to each of two 0.7ml eppendorff. To one
of these –one marked “cut” add 10 L of HindIII restriction
endonuclease solution (to be provided), mix, and incubate at 37 oC for 30
minutes. To the other marked, “uncut”, add 10L of the same buffer as
added to the cut sample and incubate in the same manner.
At the end of the incubation add 5L of the 10X loading buffer to both
samples, mix and bring them to the TA, who will show you how to load
them on the gel. Loading buffer contains some dyes to enable you to
track the progress of the electrophoresis. It also contains a high density
component, such as glycerol, to enable one to sink the DNA solution into
the well so that migration would take place through the sieving matrix
and not through the relatively non-selective electrode buffer. Loading
buffer is sometimes called ”anticonvective” or “sample buffer”. Load as
much of both solutions as the wells will hold. This is usually about a 20L
volume.
Electrophoresis is carried out at a constant current of 90 mA for 50-60
minutes. Sketch the set-up of the electrophoretic apparatus and jot
down the power supply settings needed for agarose gel electrophoresis of
DNA in you lab book. Next time you run an agarose gel you need to be
able to set up the run yourselves.
After the electrophoresis run is complete look at the gel in the
transilluminator. The gel contains ethidium bromide (about 3 L of
22
10 mg/ml ethidium bromide per 50 ml gel solution) so wear gloves when
you handle the gel. Ethidium bromide binds to duplex DNA by
intercalation. This binding allows interaction between the DNA and the
ethidium bromide so that the latter can absorb radiation in the UV region
and re-emit some of that radiation as visible orange-red light. The
intercalated ethidium-DNA complex is therefore visible on the gel and
can easily be seen and sketched or photographed for a permanent record
in you lab book.
Ethidium bromide is thought to be a mutagen, as might be expected for a
compound that intercalated double-stranded DNA. Wear gloves when
working with ethidium bromide, and avoid getting any of the solution on your
skin. Discard all waste solution into the carboy provided.
Migration of Various DNA Species in Our System
Your plasmid prep might contain some of the following nucleic acid species.
Any of these, if not removed by a particular step, can be carried through
the isolation and will show up on the gel. We need to be looking for these
when we interpret our gel.
Supercoiled DNA is the most compact form of DNA and will consequently
migrate more rapidly than other forms of DNA of the same or larger
molecular weight.
Nicked relaxed Plasmid DNA has an intermediate mobility under our
electrophoretic conditions. It has a more extended conformation and,
effectively, a larger volume, therefore migrating more slowly than supercoiled DNA. Other even more slowly migrating bands are supercoiled
plasmid dimers and protein-bound DNA forms.
Plasmid Multimers can be formed during replication. These will be migrate at
higher molecular weights and will only be seen in the uncut lane. In the
cut lane the restriction endonucleases will hydrolyze these to the plasmid
monomer molecular weight
Chromosomal DNA is large and will migrate the least distance. It is
usually sheared during the isolation so look along the line of migration to
see if there is a faint column of DNA all the way down the migration
path. Also look in the wells for very large chunks of DNA which cannot
enter the gel.
23
A schematic diagram of an agarose gel separation is shown below.
Fig. 36: Agarose Gel profile.
Your goal in this experiment is to isolate a pure plasmid. Analysis of the
electrophoretic profile of your plasmid prep will tell you whether you have
been successful or not. A successful experiment is one in which we get lots
of clean plasmid DNA of the correct molecular weight. So inspection of the
uncut lanes will tell you if there is extensive contamination of your miniprep
and the inspection and analysis of the linear lane will allow you to determine
the molecular weight of your plasmid. The molecular weights of the
components of the -Hind III standard DNA set are given in the reagents
and recipe’s section at the end of the manual.
C: Fluorimetric Assay to Determine DNA Concentration
Basis of the fluorescence method: When compounds absorb light the
energy of the photon is used to move one or more of their outer electrons
farther away from the nucleus. This is called the excited state. In most
cases the lifetime of the excited state is short (about 10-8 sec) and then the
electrons fall back to the stable ground state emitting a photon of identical
24
energy to the one absorbed. In some cases the compounds in the excited
state dissipates some of its energy internally (by converting it into thermal
energy). As a consequence the light emitted when the electron falls back to
the ground state is less energetic (it has a longer wavelength). If the time
frame for the photon emission is about 10-7 seconds the process is called
fluorescence. If the emission time is of the order of 10-3 seconds or longer
the process is called phosphorescence. Fluorescence is an extremely useful
method of analysis because compounds that fluoresce have two
characteristic and independent electromagnetic properties, a shorter
wavelength at which they absorb the light and a longer wavelength at which
they emit the less energetic light. Using the property of fluorescence we
can determine the concentration of the compound by measuring either
absorbance or emission spectra. This is because the intensity of light,
whether absorbed or emitted, is proportional to the number of molecules
doing the absorption or emission.
The fluorimeter differs from the spectrophotometer in that the
detector is at right angles to the light source and we measure the intensity
of the emitted light – the lower energy radiation – at right angles to the
light path. This has one important consequence, the instrument does not
have to read and interpret small differences in relatively large intensities as
it does in the spectrophotometer, but rather compares the emitted light
with darkness (no fluorescence). This makes for a much more sensitive
analytical method. In addition this fluorimeter is equipped with series of
quartz mirrors that reflect and concentrate the emitted radiation to
further increase the sensitivity of the assay.
In our particular assay Hoechst 33258 a bis-benzimidazole derivative
binds specifically to DNA. The structure is shown below.
Under the conditions of the assay the dye binds predominantly to A-T rich
regions and the electron excitation is facilitated. The signal due to RNA
binding is well below 1% that of the DNA and so the assay is relatively
specific for DNA. Single stranded DNA has about 50% of the fluorescent
signal of the duplex DNA and, therefore, the mode of fluorescent
enhancement does not appear to be primarily by intercalation.
25
Figure 37:
Hoescht 33258 DNA-binding Fluorophore
The dsDNA-dye complex is excited at 365nm and the Hoechst dye emits
light at about 460nm. The light is detected and its intensity measured by a
photon detector in the fluorimeter. The assay is sensitive to about 1-2 ng
of supercoiled DNA under our conditions. The spectral characteristics of
DNA-dye complex depends on the nature and shape of the DNA bound by
the dye. It is therefore best to use the same form of DNA in the standard
as is present in the unknown DNA being quantified. Hoescht like ethidium
bromide dye is intercalating and therefore a potential carcinogen.
Always use gloves when working with solutions which contain either of
these dyes.
Pictures of the mode of binding of the dye to DNA are shown below.
Figure 38: Hoescht-DNA Complex
26
Things to be Done when Measuring DNA Concentration
1. Allow the instrument to warm up for at least 30 min. With the scale
knob in the most sensitive position (fully clockwise) adjust the meter to
read “000” with only working dye solution in the cuvette. The instrument
is fairly stable and the zero point should not change much during the time
of the assays. Do not change the scale knob on the fluorimeter during
the assay. As we all will be using a common standard, we need therefore
to measure our unknowns under the same conditions used in measuring
the standard. We have to relate our unknown’s fluorescence intensity to
the concentration scale fixed by the standard. This means that the
instrument settings must be held constant during a run. The same scale
factor used in determining intensities for the standards must be used to
determine intensities for the unknowns.
2. Run a blank. We need to use a proper blank to zero the fluorimeter
before taking readings. In this case the blank consists of Hoescht dye at
the same concentration as it is in the unknown plasmid DNA solution. This
is because the dye itself fluoresces at the wavelength of measurement
and we need to be able to subtract the dye’s contribution from the
fluorescent intensity we observe in order to determine the DNA
concentration accurately.
3. Run a standard Curve . This method of analysis does not provide
consistent extinction coefficients and so we need to run a standard curve
in order to calculate concentration. We will run a standard curve for the
class so as to demonstrate the use of the fluorimeter. The DNA used in
the standard curve will be pBR322 a commercial plasmid. Be sure to
record the results in your lab book, as you will need them to determine
your concentration of plasmid DNA.
Protocol for Determining DNA Concentration


Turn the instrument on and allow it to warm up.
Make the blank by mixing 20 L water with 20 L of 2X assay solution.
There is some fluorescence in the absence of DNA. Draw up about 10 L
into the capillary. Only one blank is needed to set the instrument for the
afternoons readings.
27

Make your unknown DNA solution as follows:
1. Add 5 L of your miniprep DNA to 5L of water in a small
eppendorff.
2. Add 10L of 2X-assay solution (a solution of Hoescht dye and
buffer) to your DNA solution and mix with a pipetman.
3. Pull up your diluted DNA solution into a capillary tube and wait 10
minutes before doing the assay.
4. If your DNA signal is lower than the lowest standard signal you
need to assay a more concentrated solution of your unknown. Do
this by diluting 5L of your DNA solution in 5L of 2x assay
solution.
Measuring Fluorescence



Place the capillary with the blank Hoescht solution into the unit and
adjust the meter to zero with the “ZERO” adjust knob. Needs to be only
once per session.
Place the capillary containing the standard into the unit and record the
reading in fluorescent intensity units. Needs to be done only once per
session, therefore, already done for you.
Place the capillary containing the unknown into the unit and record its
intensity.
Lab Book information for Plasmid Prep





Look at the picture of your gel and get qualitative information about its
purity and restrictability
Determine the molecular weight of your plasmid using a plot of log kbp vs.
migration distance.
Calculate the concentration of your DNA solutions in ng/L. This unit is
particularly convenient for calculation the volumes of DNA needed in the
transformation experiments.
Have you achieved the goal of isolating pure plasmid DNA?
How do you know this?
28
Experiment 8 – Transformation of E.coli
Readings: Hengen, “Electrotransformation”, TIBS, 20, 248(1995)
Chung and Miller, 1993, Methods in Enzymology, 18, 621.
Transformation is an indispensable molecular biological too. It is impossible
to imagine the rapid progress in our knowledge of things biological in its
absence. The goal of transformation is to be able to insert foreign DNA into
a host in such a fashion that the host will express the foreign information as
if it were the host’s own. In some cases the foreign DNA consists of a
plasmid, a vector - a piece of replicable DNA with its own origin of
replication - and an insert containing the gene of interest. The hosts we use
are bacteria. They are easy to grow and grow rapidly; doubling about every
half-hour. It is easy to get transforming material inside bacteria, easy to
disrupt the cell and release its cytoplasmic contents. Bacteria are relatively
simple in composition so our isolation of our gene product is simplified.
Once inside the cell, the plasmid, with accompanying insert
information, is replicated, transcribed and translated by the host’s enzymes
as if it were host information. The plasmid is now a permanent part of the
genetic repertoire of the host and the host is said to be “transformed”. Any
time we allow the host to grow it will express the added information. We can
thus use the host as a source for the inserted DNA or as a source of the
protein coded for by the insert.
Our Transforming Material
Our transforming experiment follows the footsteps of Griffith in the
1920’s, and that of Avery and MacLeod in the late 1940’s. They established
that foreign DNA, isolated from all the other components of dead cells,
could be incorporated into, and expressed by, live cells of a different strain.
We will be using plasmids to “transform” bacterial cells. Each student will
use the plasmid isolated in the previous experiment to effect the
transformation. Both plasmids encode a gene for -lactamase, an enzyme
that hydrolyzes ampicillin. The E.coli we transform lack this enzyme, but
upon taking up the plasmid, they begin to express the plasmid -lactamase as
they grow. The enzyme diffuses out of the cell and inactivates local
ampicillin. Cells are thus able to grow in the ampicillin-free medium.
29
Barriers to transformation
E.coli does not willingly to take up foreign DNA. As in all life forms there is
an “immune” system which is designed to maintain the genetic status quo.
This means it is designed to prevent the incorporation of foreign DNA.
E.coli has developed such a defense system to and expresses enzymes that
can distinguish between self and non-self DNA and take action to destroy
the not-self DNA. The strains we use have been engineered to remove
these defense systems. So once foreign DNA gets in, it can be replicated
along with the host chromosomal DNA and express its particular set of
proteins. Even with the cytoplasmic “immune system” inactivated, the
foreign DNA still needs to get inside the cell to subvert the normal
workings of the cell, and the polysaccharide outer cell wall of E.coli
presents a formidable physical barrier to invasion. We will therefore abuse
the cells briefly to make them more “competent”. For a cell “competence”
means that their cell walls have become permeable enough to allow the
indiscriminate entry of large foreign macromolecules. The fraction of cells
that are made competent at any time is very small, sometimes only about
0.0001% but considering that we start with about 1010 cells/ml, even such a
small competent fraction will give us thousands of transformants.
Those cells, which do not take up our plasmids, are of no use to us. They
use nutrients and make components that interfere with our isolation of what
we want. We therefore need to get rid of them. We will do so by growing
our transformation mix on an ampicillin-containing plate. Only those cells
expressing -lactamase will grow and those without it will not. As the only
source of this enzyme is the plasmid, only plasmid-containing cells will grow.
Thus the ampicillin-containing plates are called selective media – selecting
for the presence of -lactamase. The strain of E.coli to be used in this
experiment is TG2. You should be familiar with the TG2 genotype and
phenotype from earlier experiments.
Overview of the Experiment
We are going to compare two different types of transformation
methodologies. One of these is an electrical method called
electrotransformation and the other is a chemical method called the onestep method. Each student will need to know something about the other
30
method. Each group will make their own competent cells. The competent
cells from one method will be pooled before using them in the
transformation. This will allow us to look at the effect of plasmid and
method on transformation efficiency. The lab instructor will assign a
transformation method to each student at the appropriate time.
Outline of the Experiment:
I.
II.
III.
IV.
V.
VI.
VII.
I.
Make Cells Electrically Competent.
Transform Electrically Competent Cells.
Making Cells Chemically Competent.
Transforming Chemically Competent Cells.
Plating Out Cells on a Selective Medium.
“Scoring” the Plates.
Analysis of the Results.
Making Cells Electroporetically Competent
Electrotransformation is the process of transforming cells by applying an
electric field across them. It is based on the observation that placing the
cells in an electric field causes a transient breakdown of their membranes,
just as one would blow holes in the dielectric of a capacitor if one put a high
electric field across it. In some cells these holes will be resealed as the
cells repair themselves. Thus, if we include some replicative form of foreign
DNA in a cell suspension and subject the suspension to a high electric field
(about 18kV/cm – for a short period of time – about 5 msec), the foreign
DNA can diffuse inside the cell during the time the membrane is
compromised. When the membrane reseals, the foreign DNA will be
trapped inside. A diagram of the electroporator is shown below.
31
Fig. 39: Schematic diagram of the Electroporator
Protocol

A small aliquot, 0.2ml, of a stock TG2 is diluted into 15ml LB broth,
and grown overnight at 37ºC.

Six ml of the overnight culture is added to 300ml of fresh LB and
allowed to shake at 370C until the OD650 (optical density at 650
nm) is about 0.4. We use the term optical density instead of
absorbance in the context of cell suspension. This is because the
decrease in intensity of the transmitted light measured by the
spectrophotometer is due to the scattering of the incident light by
the relatively large bacterial cells and not by electronic transitions
taking place in molecular chromophores in solution. At this optical
density the cells are doubling exponentially and are said to be in
the log phase of growth. At the log phase there being no lack of
nutrients or oxygen, all cells are growing rapidly and the cell
population doubles in about 30-40 min.

Each group will process 30 ml of log phase cells. We begin by
pouring the cells into a cold, sterile centrifuge tube and placing it
on ice for 5 min.

Centrifuge at 4000 rpm/5min/100C.

Decant and discard the supernatant and gently resuspend the cell
pellet in about 30 ml of cold, sterile water.
32
II.

Repeat the wash step three more times, using volumes of 30 ml,
30ml and 20ml respectively. Washing the cells in the water dilutes
out the salt from the growth medium and lowers the ionic strength
of the suspension. This, in turn, increases the resistance of the
solution and prevents current flow across the electrodes during
the voltage pulse. This minimizes heat and reduces cell mortality.
Washing is all that is needed to make cells competent for
electroporetic transformation.

After the last wash resuspend the cells gently in 0.5 ml of a 10%
glycerol/water solution. This solution should be made with sterile
water and glycerol and should not be autoclaved.

Combine all competent cell mixtures before using them in your
transformation. Competent cells stored in 10% glycerol/water
retain their competence for years at –700C.
Transforming the Electrocompetent Cells
Keep everything ice-cold until otherwise instructed. Each student
should have their own stock of plasmid from their miniprep and should use
this to transform some cells. We are going to use 100ng of DNA in all
transformations. You should calculate the volume of plasmid solution that
needs to be added to a volume of 100 L of competent cells to give the
specified mass. If dilutions are to be made they need to be made with
sterile water and not LB broth or buffer. A control transformation – cells
electroporated without added DNA – will be used to demonstrate the use of
the electroporator and serve as standard against which we will calculate the
efficiency of the transformation.
Protocol

Use a sterile pipetman tip to transfer 0.10ml of your cell
suspension to an electroporetic cuvettes, add a volume of your
DNA corresponding to 100ng, to the suspension and shake to mix
and to settle the suspension between the electrodes.
33

Put the cuvette in the safety chamber and push the slide in until
contacts at the sides of the cuvette touch the electrodes in the
base of the chamber.

Have 0.5 ml of LB broth held ready in a sterile pipetman tip to add
to the cuvette as soon as possible after the shocking. This
addition of LB will cool the cells down quickly. The sooner the cells
are cooled after their shocking treatment, the better their
chances of survival.

Press both PULSE buttons until the unit beeps. This indicates that
the pulse has decayed. Record the time constant displayed on the
unit. Ideally this should be about 4-5 msec.

Add the 0.5ml LB to the cuvette, mix and transfer to a sterile,
labeled 1.7ml eppendorff tube. Place the eppendorff in the 370C
incubator in the balance room.

Allow the cells to remain in the incubator for about 30 minutes.
This allows them time to recover and begin the expression of the
genes that allow them to survive the selection process.

During the dead time you are going to clean up your electroporation
cuvette for re-use. The method used is Paul Hengen’s “paranoid
method” from the Bionet “Methods and Reagents FAQ” with the
web address bionet.molbio.methds-reagnts, a good source for
information about molecular biology protocols.







Wash out with bleach.
Rinse six times in distilled water.
Fill with 0.25M HCl and allow to stand for 30 minutes.
Rinse in distilled water.
Boil for 10 minutes in distilled water.
Remove immediately and rinse in 95% ethanol.
Air-dry upside down and store in a clean container.
34
III. Preparing Chemically Competent Cells
The chemical method makes use of a mixture of PEG (polyethyleneglycol), magnesium ions and DMSO (dimethlysulfoxide) to permeabilize cell
membranes so that endogenous material can enter the cell. DMSO has been
used to carry dissolved compounds such as arthritis medicine, across the
skin to relieve arthritic joint pain. So one may imagine that it fulfills such a
role in the transport of DNA across the E.coli cell membrane. Each group
will make their own competent cells. The cells will be combined before being
transformed
Protocol

A small aliquot, 0.2ml, of a stock wild-type TG2 are grown
overnight at 37ºC in 15ml LB broth.

Add 6 ml of the overnight culture to 300ml of fresh LB and allow
the diluted culture to shake at 370C until the OD650 (optical
density at 650 nm) is about 0.4. We use the term optical density
instead of absorbance in the context of cell suspension. This is
because the decrease in intensity of the transmitted measured by
the spectrophotometer, is due to the scattering of the incident
light by the relatively large bacterial cells and not by electronic
transitions taking place in molecular chromophores in solution. At
this optical density the cells are doubling exponentially and are
said to be in the log phase of growth. At the log phase there being
no lack of nutrients or oxygen, all cells are growing rapidly and the
cell population doubles in about 30-40 min.

Each group will process 30 ml of log phase cells. We begin by
pouring the cells into a cold, sterile centrifuge tube and placing it
on ice for 5 min.

Pellet the cold culture at 1000g for 10 minutes and discard the
supernatant.

Gently resuspend the cells in 2 ml ice-cold TSS - Transformation
and Storage Solution and incubate on ice for 15 min.
35

IV.
The cells are now competent for transformation. Combine the
various preps into a common cell suspension for the transformation.
What we do not use we can store at –700C for up to a year.
Transforming Chemically Competent Cells
Keep everything ice-cold until otherwise instructed. Each student
should have their own stock of plasmid from their miniprep and should use
this to transform some cells. We are going to use 100 ng of DNA in all
transformations. You should calculate the volume of your plasmid solution
that needs to be added to a volume of 100 L of competent cells to give the
specified amount of transforming DNA. A control transformation – cells
treated as below without added DNA – will be used as a standard for
determining the efficiency of each transformation. Either the TA or
assigned students will do the standard for the section.
Protocol
V.

Pipette 100 L of competent cells into a 1.7ml sterile eppendorff.

Add 100 ng of your plasmid DNA to the competent cells, mix and
incubate on ice for 40 minutes.

Add 0.5ml TSS containing 20mM glucose and incubate the cells at
370C for 30 minutes.

Some of the cells are now transformed.
Selection of Transformed Cells
Remember we are going to select for transformants by plating out on LBampicillin solid medium. These are the plates you made as part of
experiment six. While the cells are recovering, prepare your plates for the
selection process. By this I mean label your plates on their bottoms with
indelible marker, with script which identifies the owner of the plate, the
plasmid plated and the dilution on each plate. Your plates will go into a big
36
incubator with everyone else’s, so be sure each of your plates can be
identified. When the cells are recovered we need to dilute them out suitable
before plating them. Even at low transformation probabilities there will be
far too many on a plate to count individually. At low dilutions the bacteria
would cover the plate in what is accurately described as “a lawn” and be
uncountable. Therefore we need to make dilutions of our recovered culture
before plating. A good volume to spread out on the plate is in the range
50L. A smaller volume is hard to spread evenly while a greater volume can
result in a watery plate where the colonies are smeared out. Therefore we
dilute out or cultures before plating and use 50L of the dilution for
spreading. Use LB not water as a diluent for your culture. During the
recovery time we can also set up some eppendorff tubes in a plastic rack for
diluting out our culture before plating. The plating protocol for controls and
experiments are given in below.
Not every group will need to plate out a control. Only ONE control is
needed for each transformation method, as each section will be
transforming the same competent cell population a TA or an assigned
student will prepare the control for each method for each section.
Serial Dilution and Plating of Control Cultures

Add 0.5ml of LB to each of three eppendorff tubes, and 0.49ml of LB to
a fourth. Label these, “102”, “104”, “106” and “107” respectively. The
exponents refer to the dilution factor, the higher the dilution factor the
more dilute the cell culture.

Add 5L of the recovered control culture to the tube labeled “102”
dilution and mix. This will give a 100-fold dilution of the culture.

Add 5L of the 102 dilution to the “104” tube and mix. This will give a one
hundred thousand-fold.

Add 5L of the 104 dilution to the “106” tube and mix. This will give a
million-fold dilution.

Add 50L of the 106 dilution to the “107” tube and mix. This gives a ten
million-fold dilution.
37

Spread 50L of the “106” and “107” dilutions on LB plates and spread 50L
of the “104” dilution on a LB-amp plate. The plating on LB allows us to
assess the viability of the cells following their treatment. The plating on
the LB-amp allows us to check for revertants, contaminants and whether
we made an error in making or marking, the plates.

It is not necessary to use a fresh tip or spreader every time you add
some culture to, or spread some culture on, a plate. As long as the set
contains the same type of culture – except for a dilution factor – it is
possible to pipette out or spread the entire set using one tip or spreader.
To do this just set out your plates in order of increasing concentration of
bacteria from left to right. Add 50L of “107” dilution to your “107”
plate. Use the same tip to add 50L aliquots of increasing concentration
culture to its corresponding plate. When done, you have all your dilutions
lying on the agar surface.
Spread the cultures in the same order as you plated them and you should
be able to use the same spreader for all dilutions.
Allow the plates to stand for 15 minutes on the bench and then place
them upside-down in the 37ºC incubator on the north wall of the lab.
They will be incubated overnight.
Plates should be removed from the incubator after an overnight
incubation because, if they are not removed, the colonies will grow
overgrow each other and counting becomes difficult. The TA will remove
your plates from the incubator and place them in the refrigerator. They
will keep in the refrigerator until the next lab session when they can be
scored.



Diluting and Plating Transformed Cells
Modify the control protocol above and make a ten-fold and a hundred-fold
serial dilution of your transformed culture. Spread 50 L of each of these
two dilutions on labeled LB-amp plates. In addition spread 50L of your
original transformed culture on a LB-amp plate
38
VI.
Scoring Plates
“Scoring plates” is the microbiological buzzword for counting transformed
colonies on each plate. You need to score your plates so that you can
calculate the efficiency of your transformation. This does not usually mean
“count every colony”! Remember that your plates are just dilutions of the
same original sample, and that “counts” do not have to be exact in bacterial
work. You might want to count only one quarter of a crowded plate and
multiply by four to get an estimate of the total colonies on the plate.
Record your results in you lab book in the form of a table as shown below.
Control
Control
Control
Transformed
Transformed
Transformed
Dilutions
104
106
107
100
101
102
Colonies/plate
Colonies/ml
VII. Analysis of the Experiment
There are two numbers usually used in transformation experiments to assess
the success of the transformation. The first is the probability of a cell
being transformed and the second is the transformation efficiency or XFE.
We are going to use our data to calculate both these numbers.


Transformation probability – for this you need to know how many cells
survived our treatment to make competent and how many of those live
cells were actually transformed. If we take the ratio of the
concentration (cell/ml) of amp-resistant (transformed) cells to that of
the control plates, we have the probability of a single cell being
transformed.
Transformation Efficiency or XFE – for this value find the total number
of transformed cells and divide this by the mass of DNA, in g, used to
transform them. XFE has units of transformed cells/g DNA.
39
Sample Calculation – any resemblance between these and actual
transformation results is coincidental.
Sample Data
We added 100 ng DNA to 100uL of competent cells and, after
transformation, added 2ml of LB broth to aid recovery. We then plated out
50 L of a ten-fold and a hundred-fold dilutions of our transformed cultures
on amp plates and got 25 and 3 colonies respectively. When 50L of the
control culture, diluted by a factor of 106, was spread, it showed 30 colonies,
when one-quarter of the plate was counted.
i.
Calculating colonies/ml

Ten-fold Dilution calculation
25 colonies Plate
colonies
X
X10  5000
plate
50uL
ml

Applying the calculation above to hundred-fold dilution should give 6000
colonies/ml. The number used to represent colonies/ml for the original
should be the average of the two above results.
 Control calculation
4 quarter - plates
30 colonies
cells
X
X106  2.4X109
quarter - plate - 0.050ml
plate
ml
ii.
Calculating probabilities of transformation
Take the ratio of colonies/ml for transformed cells to colonies/ml for
control cells.
5500
colonies/m l
 2.3X10-6
2.4X10 colonies/m l
9
40
iii.
Calculating XFE – XFE’s have units of colonies/ug. We therefore need
to get “colonies” and “g DNA” and take their ratio to give us XFE.
According to the protocol above we added 100 ng of DNA to 100L of our
recovered cells and subjected them to transformation. We then diluted our
transformed mixture with 2ml LB broth to cool them off, and plated 50L.
In our 100L recovery mixture there must have been:
5500 colonies 2.1ml
X
 11,550 colonies
ml
Assuming there has been no cell division during recovery, we can calculate
the XFE as follows:
XFE 
11550 colonies
colonies
 1.2 X 10 5
0.1g DNA
uL
Lab Book


Determine transformation probabilities and XFE’s for your
transformation.
As a separate exercise we are going to use the class data to do a
statistical analysis of the effect of method and plasmid on
transformation probabilities. You will get additional information about
this on the web site. This will not go into your lab book so don’t delay
writing up this experiment waiting for class data.
41
Experiment 9A – Preparing for the Southern blot
In our next experiment we are going to learn how to do a Southern blot, one
of the most important techniques in molecular biology. This is the
indispensable technique of the human genome project. The Southern blot
uses a short piece of DNA to probe for its - the short piece’s – target, a set
of complementary sequences in a complex DNA mixture. Our DNA target
will be the lacZ gene of E.coli. This is a single copy gene which comprises less
than 0.1% of the E.coli chromosomal DNA. Our probe will be made from two
short sequences of the lacZ gene. It might seem like circular reasoning to
use lacZ to find lacZ, but short sequences of DNA, which can easily be
obtained from partial amino acid sequences, allow us to detect larger
sequences. From these larger sequences we get new probes which we can
fish out other large pieces. This process continues until eventually, we have
all the pieces of the gene. By overlapping these pieces, we obtain the linear
sequence of the lacZ gene.
Today’s Work – Two Southern’s per aisle.
A. Preparing Chromosomal DNA.
 Determining DNA concentration by Spectrophotometry.
 Practice Calculation for Restriction Digests.
 Restrict Chromosomal DNA.
B. Synthesize a LacZ Probe by PCR.
A. Preparing Chromosomal DNA

Determining DNA Concentration by Spectrophotometry
J.A.Glasel, Biotechniques, 18,62(1995)
E.coli chromosomal DNA from Sigma, a commercial biotech company
has been dissolved in TE buffer. You will be given an aliquot of this
solution and will need to determine its concentration. You need some
chromosomal DNA for your Southern blot and we want to know what
volume of the DNA is needed for adequate detection. The bases of
DNA absorb strongly in the near UV region with a maximum
42
absorbance around 260nm. At this wavelength the extinction
coefficient used to determine concentration is usually taken to be
0.02/g/ml/cm. In solution as detailed in experiment I, in solution
we have absorption bands not spectral lines. The band for DNA
band falls away gradually and an A260/A290 of 1.9 is considered to be
pure DNA. We are going to read Glasel’s paper for some discussion
as to the validity of this assumption. The protocol for the
determination of the DNA concentration is as follows:







Use quartz cuvettes for the experiment.
Zero the cuvettes from 340nm to 240 nm using TE in both
reference and sample cuvettes.
Add 50L of your unknown DNA solution to 1.0ml TE buffer.
Run the unknown spectrum from 340nm to 240 nm.
Get the Shimadzu to print out the spectrum and a list of
absorbances, at 10nm intervals, which cover the range of the
spectrum. Make a copy of the information for each student.
Check the DNA concentration using the fluorimeter.
Restricting Chromosomal and Plasmid DNA.
We are going to double-digest our pIH1034 plasmid and single-digest our
chromosomal DNA. The restriction endonucleases we are going to use on the
plasmid are, DraI and NcoI. These cut on both sides of the lacZ gene in
pIH1034 and produce a fragment of about 3000 bp. This fragment
comprises almost the entire lacZ gene in pIH1034. The NotI will cut at a
specific 8 base-pair-long sequence. This will produce, in E.coli, fragments
which are, on the average, about 66000 base pairs long.

Sample Restriction Calculations: restriction type calculations that we are
going to do today are done routinely in molecular biology labs. Each
student will do the following sample restriction calculations before
restricting our own plasmid and the provided chromosomal DNA.
“We have been given a plasmid with a concentration of 100 g/ml,
some 10X restriction buffer. “10X” in the context we are using means
“the buffer is ten times more concentrated than is necessary for
running the digestion” This means the restriction digest is routinely
run in a 1X concentration buffer. We also have a 10,000 units/ml
43
solution of Ear I. We want to restrict a given mass of our plasmid
simultaneously with enough EarI in a reaction mix with a 30L volume,
to restrict all the plasmid. Devise a protocol to restrict 200 ng of
your plasmid using 3 units of each restriction endonuclease in a total
volume of 30L 1X buffer. You are also given chromosomal DNA with a
concentration of 2 mg/ml. Devise a protocol for double-digesting 5 g
of the DNA under the same conditions. These calculations need to be
handed in before the start of the class and are merely practice for
the similar calculations you need to do for today’s restriction of your
own plasmid and the group’s chromosomal DNA.
Fig. 41 - Restriction Map of pUR290

Restrict an aliquot of Chromosomal DNA for use as target DNA in
the Southern Blot. Try to get as much DNA in the restriction
digest as you can. Remember lacZ is a single copy gene. Try for 5g
DNA per well. The concentrations of the chromosomal DNA and of
the restriction endonuclease will, of course, be different from
that given in the calculations above. NotI is sold at a concentration
of 15U/uL.
44
Your restriction protocol for the practice calculation should be handed in, in
the matrix form below. The real restriction conditions should go into your
lab book in this form as well.
DNA
Chromosomal
PUR290
Volumes in L
10X
Ear I
NotI
-
H 2O
Fig 40. Form for Restriction Digest Information
B. Synthesizing a Probe by PCR
An Overview of the PCR Reaction - we are going to use the Polymerase
Chain Reaction (PCR) to replicate a small portion of the lacZ gene using a
cell-free system. PCR was developed as a working technique for the in
vitro amplification of DNA in 1987. The idea had been considered in 1971
by Khorana but was not pursued, as there were no hear-stable
polymerases around. In 1987 by Gary Mullis turned Khorana’s idea in to
reality and developed, what is now, one of the premier techniques of
biotechnology. PCR and its alter ego, the Southern blot, are the two
methodologies mainly responsible for the stunning explosion of
fundamental discoveries in molecular biology. The basis of PCR is the 5’ –
3’ extension of two small primers complementary to sequences
downstream from each other, but on different DNA strands, of a larger
target DNA molecule. If the target DNA is made single-stranded, the
primer will bind and a polymerase will extend the primer in the 3’
direction using the target strand as a template. The other target strand
will have a different primer site and will also be extended, but in the
opposite direction, being on the other strand. The net result of these
reactions produces a dsDNA copy of sequences between the two primer
sites. The way the PCR reaction is run allows this target region to be
amplified many-fold. The diagram of the PCR reaction is shown below Fig. 41.
45
The Components of the PCR Reaction

Taq Polymerase - the polymerase we are using is called Taq polymerase
and is a DNA polymerase that has been cloned from the gene in Thermus
Aquaticus, a thermophilic bacterium. Taq has an Mr of about 95 kDa, no
3’ -5’ exonuclease activity, a half-life of 40 minutes at 95°C and can add
thousands of nucleotides to the primer in the time, usually minutes, we
allow for extension in the PCR machine. Taq incorporates with an error
rate of about one substitution per 9000 bases and a frame-shift error of
about one in every 41,000 bases. It can therefore replicate target
strands quite rapidly and exactly. Taq polymerase can also incorporate
modified nucleotide containing bulky side-chains, albeit a little more
slowly than its regular substrates. Thus a digoxigenin-labeled DNA, DigdUTP, will compete with the regular dTTP substrate and will be
incorporated into the sequence being amplified.


Primers – our system includes two primers specific to the lacZ gene
that whose replication will result in a 300bp region of the gene being
amplified.
Label – in addition the system contains a modified nucleotide some
digoxigenin-11-dUTP - Dig-labeled dUTP. The label does not radically
interfere with the use of its dUTP as a substrate for the polymerase
during the replication of the target strand. It is incorporated fairly
46
efficiently into the growing daughter strand in the place of dTTP. In
addition, the Dig appendage, derived from a plant hormone, has no
bacterial equivalents. Thus a mammalian antibody directed against
this plant molecule would find it even in a complex bacterial protein
mixture. Thus on labeling DNA with digoxigenin we have synthesized a
bipolar probe, one end of which is DNA specific- it will base-pair to
the target DNA, and the other end is digoxigenin - it will bind only to
the anti-digoxigenin antibody. This arrangement forms the basis for a
sensitive, non-radioactive assay for the location of its complementary
target fragment.

Target DNA – this will be E.coli chromosomal DNA purchased from
the Sigma chemical company. It will have been dissolved in TE buffer.
The Detection System
The detection system contains an anti-digoxigenin-antibody covalently
linked to an alkaline phosphatase. This is the same detection system used for
the Western blot. The DNA-binding end of the probe will find the target
DNA, and the Anti-dig antibody will find the Dig label and the alkaline
phosphatase goes along for the ride. Once binding has taken place we can
use the NBT/BCIP combination to locate the region on the membrane
containing target sequences. See the figure below for a diagram of the
visualization sandwich.
Figure 42: Visualization Sandwich
47
Protocol for Labeling Probe
Reagents




Target DNA (Template): E. coli chromosomal DNA 2.5 g/ml to be
provided.
Primers: Both primers diluted to a concentration of 50 M in water.
Primer I: TGC CAA TGA ATC GTC TGA CC
Primer II: GGACCA TTT CGG CAC AGC
PCR Supermix: A complete PCR solution manufactured by Gibco. All
one needs to do is add primers and target DNA. The mix consists of
22M Tris HCl, pH 8.4, containing 55 mM KCl, 1.65 mM MgCl2, 220 M
of dATP, dTTP, dGTP, and dCTP and some proprietary stabilizers.
Label: 1mM digoxigenin-11-dUTP in water.
Protocol

DNA
5
Add the reagents in the volumes specified below to a 0.7ml sterile
eppendorff tube.
Supermix
45
All volume in L
Primer I
Primer II
2
2
Dig-11-dUTP
3

Mix and run in the PCR using the hot bonnet and the
3-step protocol program in the PCR instrument. The
3-step protocol is as follows:







1 minute at 940C.
40 seconds at 920C – the denaturation step.
40 seconds at 600C – the annealing step..
1 minute and 30 seconds at 750C – the extension step.
Repeat the 920C to 750C cycle 29 times.
5 minutes at 750C- a final extension step.
Indefinite hold at 40C.
48
I will remove these reactions from the machine and store them at
-200C until the next session when we will to check to see whether the probe
has been labeled.
Day II: Checking the labeled probe
We need to know if our probes were successfully labeled before using them
in our Southern blot. We will check the probes by covalently linking a small
aliquot to a nylon membrane. This is done by UV irradiation. The UV light
causes the formation of covalent bonds between thymidine bases and the
nylon membrane, a process very similar to the thymidine dimer formation
when UV light mutates DNA. We then wash the membrane with solutions to
remove contaminants and cover the membrane surface with a protecting
protein and DNA layer - a process called blocking. Finally we treat the
membrane with detection reagents to visualize the location of the target
DNA. In addition to our probes we are also going to run a set of dig-labeled
DNA standards through the same washing, blocking and visualizing
procedure. This standard set will allow us to obtain an estimate of the
concentration of our probes.
Protocol for Checking Probes





Prepare 10X and 100X dilutions of your probe using 1L aliquots.
Spot 1L of your concentrated probe and your two dilutions onto a
properly equilibrated air-dried nylon membrane. Try to keep the are of
the spot as small as possible. The TA will show you how to do this. The
membrane has to be fairly dry before small spots can be made. Make sure
you know which slots you used for your probes.
Allow your DNA spots to air-dry for 10 minutes.
Along with your probe each section should include a biotinylated DNA
standard strip purchased form Roche Biochemicals. These strips consist
of precise masses, 3, 10, 30 100 and 300pg, of biotin-labeled DNA crosslinked to the strip.
Covalently cross-link the DNA to the membrane using the UV
stratalinker. The stratalinker cross-links by breaking double bonds in
thymidine bases and allowing the unpaired electrons to form covalent
49
bonds with the carbonyl groups in the membrane. This reaction is
analogous to thymidine dimer formation in DNA cross-linking.
Equilibrate the membrane in 15 ml of 1X wash buffer (a 0.1M Tris-buffer
pH 7.5, containing 0.15M NaCl and 0.3% Tween-20, a non-ionic detergent)
for 3 minutes.
Block for 45 minutes in 70 ml blocking buffer, a proprietary buffer
containing detergent and both nonspecific DNA and non-specific protein.
“Nonspecific” means in this context that it will bind nonspecifically to
membrane “hot-spots” and coat the membrane, so that components of the
visualization solutions will not bind there, and produce false positives.
Discard the blocker and incubate the membrane in 20 ml of fresh blocker
containing anti-digoxigenin-antibody/alkaline phosphatase conjugate
solution for 10 minutes. The concentrations and volume will be given at
the appropriate time. We buy several different types of conjugate and
do not know beforehand what the active concentration will be.
Discard the conjugate solution and wash the membrane in 70 ml of 1X
wash buffer for 15 minutes.
Repeat the above washing process one more time.
Place you membrane in a weighing boat and cover it with substrate
solution. Substrate solution is BCIP (bromo-chloro-indolyl-phosphate) and
NBT (nitro-blue-tetrazolium in high pH Tris buffer). The TA will show
you the recipe for this reagent.
Wait for about 30 minutes. If no signal is seen at this time cover the
solution and leave it overnight in a drawer to develop.
Stop the reaction by washing copiously with water and store in 0.5M
EDTA in the seal-a-meal bag.








The diagram for the detection sandwich has been given earlier and the
reactions giving rise to the visualization are shown on page 50.
Lab Book

Compare the intensities of your probes’ signal with that of the standard
DNA set to derive a qualitative measure of the initial concentration of
your probe.
50
Experiment 9B – The Southern Blot
Purpose: To locate the lacZ gene in E.coli.
Introduction
We have seen in earlier experiments that moving fragmented DNA down an
electric field through a sieving matrix will resolve those fragments
according to size. If it were possible to discriminate between those
fragments by making use of some internal differences, such as differences
in DNA composition or sequence, one could take advantage of these
differences to explore individual or population DNA differences. Using
different fragmentation methods and detecting only parts of one particular
fragment would allow us to “fingerprint” the chromosomal DNA of an
individual. The figure below illustrates this idea.
Fig. 43:
“Fingerprinting” Chromosomal DNA.
In 1975 E.M. Southern devised a technique to do just that- to detect
individual differences in DNA. This technique, called the “Southern blot”,
has become an essential tool of anyone attempting to understand the
relationship between phenotype and genotype, or merely studying DNA at
the molecular level. To perform the Southern blot the gel is fragmented in
some fashion and resolved on a gel. After the resolution, the DNA is
51
denatured in the gel so that it becomes single-stranded. It is then reneutralized and becomes capable of H-bonding to complementary sequences.
The re-neutralized DNA is transferred to a membrane, making a mirror
image of gel DNA pattern on the membrane. The membrane is then exposed
to a single-stranded DNA probe that contains sequences complementary to
those on the target DNA on the membrane. Hybridization then takes place
on the membrane and, if the probe has been labeled with some reporter
group, it is possible to locate the complementary target fragments on the
membrane.
Uses of the Southern Blot




Determination of the chromosomal location of genes.
Forensic usage - comparing or “finger-printing” DNA from
different sources.
Ferreting out genetic linkages.
Diagnostics.
An Outline for a Southern Blot
This experiment is not particularly difficult, but does require organization
and must be done according to the schedule outlined in the protocol section.
The outline of the experiment is as follows:
I.
II.
III.
IV.
V.
VI.
Resolve fragments on an Agarose Gel.
In-gel treatment – denaturation and neutralization.
Transfer of DNA from the gel to a membrane.
On-membrane treatment – cross-linking, prehybridization and
hybridization.
Washing the membrane.
Visualization of the Complementary Fragments.
We will give some additional information about each of the above steps and
before giving the detailed protocol.
I.
Resolution of fragments – this will be done as before using a 1%
agarose gel cast in IX TAE. The gel is to be run at 90ma for about
one hour.
52
II.
In-Gel treatment


III.
Denaturation - raising the pH deprotonates the bases and the
hydroxyl groups of the sugars. The combination of these effects –
the loss of hydrogen bonding and the formation of a charged
functional group – destabilizes (or denatures) the dsDNA and the
chains move away from each other. Remember that the interstitial
spaces of the gel are much larger than DNA molecules, and the
DNA is essentially in solution in the interstitial fluid of the gel.
Neutralization – lowering the pH restores the H-bonding capacity
of the bases. Even though the molecules are now capable of basepairing, they do not form dsDNA. The rate-limiting step in DNA
renaturation is the nucleation event where complementary bases
find each other and begin to zip up the helical structure. The time
needed for this process depends on the complexity of the
molecule. The more complex the DNA the longer the “zipping up”
takes. As our target DNA is quite complex, it essentially remains a
combination of single-stranded and heteroduplex (mismatched)
forms in the gel. These are less soluble than the double-stranded
forms and tend to precipitate out of solution.
Transfer of the DNA to the Membrane –the DNA in the gel is
transferred from there to the membrane by capillary action. The
solvent flow from the solvent reservoir through the gel to the
solvent sink – a stack of dry absorbent paper - carries the DNA along
with it to the membrane. And, as the DNA binds quite strongly to the
membrane, it is stuck there as the solvent moves off. The capillary
set-up is shown on the next page. The membranes are sold as dry
strips and need to be equilibrated before use. You will be given the
equilibration details in class.
53
Fig.44: Capillary Transfer Apparatus
IV.
On-Membrane Treatment
 Cross-linking the DNA to the Membrane – the DNA is only weakly bound
to the nylon and, if not somehow permanently fixed there, would be
washed off during the washing steps preceding the visualization. The gel
is incubated under UV light for 30 seconds to covalently link the DNA to
the nylon membrane. As we said before the cross-linking process is
something akin to the formation of pyrimidine dimers in dsDNA with
electrons from the base forming covalent bonds with the nylon matrix.

Prehybridization - after transferring the DNA to the membrane we have
to treat the membrane so that the DNA probe and detection proteins will
bind only to their specific targets on the membrane and not nonspecifically to chemical hot spots on the membrane. To do this we wash
the membrane in detergent, non-homologous DNA and protein so that
these will occupy the non-specific membrane sites. This protection
process is called prehybridization. After the prehyb treatment we can
incubate the probe with the target DNA, hybridize them, and be
reasonably sure that areas of the membrane that light up after the
detection process will all be complementary to target DNA sequences and
not random.

Hybridization – The membrane is now ready for probing and is incubated
with single-stranded probe so that it can hybridize with its target single54
stranded DNA on the membrane surface. The components of the
hybridization solution and the conditions of the hybridization reaction,
under high temperature, are such that only a high complementarity
between probe and target will be result in binding. These conditions are
called high stringency conditions. The presence of formamide in the
hybridization solution lowers the melting point of the probe-target DNA
duplex by about 32ºC. Thus we are effectively conducting the
hybridization at 75º - a temperature that maximizes the requirements
for H-bonding between probe and target for duplex formation.
V. Washing the membrane – we wash the membrane extensively in buffer
and detergents to remove weakly bound species.
VI. Visualization – the washed membrane is incubated in a solution containing
an anti-Digoxigen antibody conjugated to alkaline phosphatase. The
antibody conjugate binds to the dig label and on adding BCIP and NBT as
we did in the Western blot, the reaction produces insoluble diformazan
and indigo. We thus “light up” the target fragment on the membrane
surface.
A schematic diagram of the on-membrane processes is shown on the next
page.
55
Fig. 45: Diagram of Membrane Treatment
Schedule for the Southern Blot
Day I




Day II



Separate DNA on the agarose gel.
Denature the DNA in the gel by incubation in base.
Neutralize by incubating in a neutralization buffer
Transfer the ssDNA from the gel to the membrane by capillary
action.
Come in before 10:00 a.m. to dismantle the transfer apparatus and
covalently link the ssDNA to the membrane using the Stratalinker.
Incubate the membrane in pre-hybridization buffer for 3-4 hours.
Do an overnight hybridization of your ss-probe to the
complementary ssDNA immobilized on the membrane.
56
Day III



Incubate the membrane in blocking buffer to saturate non-specific
protein binding sites on the membrane.
Incubate the membrane with an antidigoxigenin-alkaline
phosphatase conjugate. Wash off non-specifically bound conjugate.
Add an alkaline phosphatase substrate to visualize the target DNA
in situ.
Protocol
Preparing the Restriction Digests for Running
We restricted our DNA last session during the probe synthesis. You may
remember that sensitivity required that we digest five times more
chromosomal DNA than plasmid DNA. Unequal amounts are used because the
lacZ sequence is present in very low concentrations in the chromosomal DNA
and our plasmid DNA is, by contrast, present in relatively large
concentrations. Loading unequal volumes will help equalize the hybridization
signals between the two DNA targets, the plasmid and the chromosomal
DNA. Set up the following samples in 0.7ml eppendorff tubes:
Type of DNA
Chromosomal
Plasmid
Vol. DNA (uL)
30
20
Vol. Loading buffer (uL)
4
3
The loading buffer contains glycerol and some colored dyes. The glycerol
makes the solution denser than the electrode buffer and so it will sink to
the bottom of the well and the DNA will migrate through the agarose. The
dyes, bromophenol blue and xylene cyanol allow us to monitor the progress of
the electrophoresis. The bromophenol blue will co-migrate with DNA of size
1000bp while the xylene cyanol co-migrates with about 5000 bp DNA.
57
I. Agarose Gel Electrophoresis of DNA
The following controls will be provided and should be run once on each gel.



DNA Standard - Dig-labeled HindIII digest. This will allow us to
determine the molecular weights of bands on the gel.
House Chromosomal DNA – undigested E.coli chromosomal DNA.
Negative control - unlabelled HindIII digest.
Two groups will pour a 1% agarose gel in 1XTAE and use it in common. The
combs we have form, at most, so load in the following order:
Lane
Lane
Lane
Lane
Lane
Lane
1:
2:
3:
4:
5:
6:
…
Lane 12:
10 L negative control- unlabeled HindIII digest.
10 L labeled HindIII standard.
20 L House uncut E.coli.
20 L your chromosomal DNA.
2 L your plasmid digest.
5L your plasmid digest.
10 L labeled HindIII digest.
The above order is for one group. The second group on the gel will load
their samples according to the same order in the available wells. The
standard on both ends allows us to determine molecular weights where
migration is not uniform across the gel.

When all the lanes are loaded, run the gel at a constant current of
80-90 mA. During electrophoresis you will equilibrate your
membranes. After the gel has run we will look at the gel on the
transilluminator to inspect the pattern, before going further.
58
II. On-Membrane Treatment
These processes prepare the gel for probe binding by first making the DNA
single-stranded and then neutralizing it, so that it can hydrogen bond to the
probe.

Denature the DNA in the gel by shaking in 50 ml of 0.5M NaOH
containing 1.5M NaCl for 15 minutes at room temperature. Repeat
the denaturation in fresh base for an additional 15 minutes. Note
the change in color of the bromophenol blue dye.

Neutralize the DNA in the gel by shaking in 50 ml of 1M phosphate
buffer, pH6.5, for 10 minutes. Repeat the neutralization two more
times. The DNA is now ready for transfer and competent for base
pairing.
III. Transfer of DNA to the Membrane
Set up the transfer unit according to the instructions that come with the
unit. The solvent used for the transfer is 20X SSC. The transfer should
take only about 2-3 hours, but can safely be to take place overnight as we
shall do. After the transfer remove the membrane from the transfer
sandwich. Before separating the gel from the membrane mark the face of
the membrane where the membrane contacted the gel, this is the DNA side-up surface of the membrane i.e. the surface to which the DNA is
bound, and which should be exposed to the cross-linking radiation. Cutting
off one of the corners can do this. This also allows you to orient yourself
for the identification of the various bands on the membrane. Even though
the bands as loaded on the gel, are not symmetrical, it is sometimes useful
to have a clear-cut orientation marker on the gel.
IV. Membrane Treatment

Place the membrane, DNA side up on a piece of filter paper, in the
Stratalinker. Use the autolink mode to cross-link the DNA. This
program exposes the DNA to 1200J of UV light which covalently
links the DNA to the membrane.
59

Lay your membrane down on the nylon mesh about three
membranes per mesh. Roll the mesh package up into a cylinder and
place in the hybridization bottle. Add about 50 ml of prehyb
solution to the bottle and use the hybridization oven to
prehybridize the membranes at 42oC for at least 2 hours.

Boil your probe for 10 minutes in 10 ml of fresh hybridization
solution and store it in the freezer until you need it. Combine all
probes for the common hybridization.

Decant the prehybridization solution. It can be reused, so decant
it into the container provided. If none is provided find a clean
beaker and label it “prehyb” solution and decant into it.

Add the single-stranded probe to the hybridization bottle and
incubate overnight at 43oC.

Decant the used probe solution and save it. The probe can be
reused several times if stored at –20oC.

Wash the membranes twice for 5 minutes a piece in 40 ml of
2XSSC containing 0.1% SDS.

Wash the membranes twice, 15 minutes apiece, at 68oC in 50 ml of
0.1XSSC containing 0.1% SDS.
VI. Visualization of Complementary Fragments
The membranes are now ready for visualization. We use the almost the
same protocol for visualizing our Southern as we did for checking our probes
so consult experiment 9A. The only differences are that now, that we
equilibrate the membrane for about 10 minutes in the wash buffer instead
of three minutes, and we reduce the blocking time to twenty minutes.
60
Lab Report
Do one formal report for both parts of experiment 9. Arrange your
discussion around your data. Address your data interpretation to answering
the questions below.
a.
b.
c.
d.
e.
f.
What bands do you expect to see on the gel?
How many fragments of what sizes lighted up?
Did you see bands you expected?
Were there any unexpected bands on the gel?
What concentration of probe did your target DNA see?
Does your fragment size agree with that expected from the E.coli DNA
sequence
61
Making Buffers in a Biochemistry Lab
The first lab periods will be devoted to making buffers which to be
used during the coming semester. In order to save space, and make enough
of a given buffer for the entire semester we usually make the buffers at a
higher concentration than we use them at. In the lab terminology we
describe the situation above as, we make buffers at, say a, 10X
concentration (ten-fold higher than we need) and dilute them to 1X (the
concentration of use). To make buffers we need to brush up on some basic
calculations from genchem, to relearn the use of chemical balances, and to
understand and use pH meters.
Choosing a Buffer
As you recall from genchem, a buffer is a solution of a weak acid and its
conjugate base, the combination of which is able to minimize changes in pH
caused by the addition of external acid or base. You also will, no doubt, also
recall that the pH of the solution is determined by the pKa of the acid and
the ratio of the concentration of the conjugate base to the concentration of
the acid. The Henderson-Hasselbach expresses this relation:
pH = pKa + log[(conjugate base) / (acid)]
Biological systems, including enzyme reactions, often have an optimal pH so
that the most sensitive assays are usually be carried out near that
optimal pH. So, knowing the optimum pH, we need to choose an appropriate
buffer i.e., one which would buffer (or hold the pH relatively constant) at
that pH. Buffers are most effective when the pH of the solution is equal to
the pKa of the system, as, at this pH, the concentration of the acid form
equals the concentration of the conjugate base form, so we have maximal
conversion of one form to the other, or buffering. We would be lucky to
find both the above conditions fulfilled at the arbitrary pH of one out of
many enzymes. Therefore, the best choice of a buffer would be one whose
pKa is close to the required pH. Physiological pH is about 7.4 so usually
reactions can be run close to pH 7 and we consequently need a buffer with a
pKa near 7. Two commonly used buffers are tris-hydroxymethyl amino
methane, Tris base, with a pKa of 8.07, and one of the three ionization
forms of phosphoric acid, H 2PO 4-, with a pKa of 7.20.
62
Making the Buffer - Theory
After the appropriate pKa has been chosen, a buffer of the appropriate pH
can be made by adjusting the ratio of the base form to that of the acid
form, of that buffer, in one of three ways:
1. A salt of the acid form can be added to a salt of the base form in the
correct molar ratio. This will give the desired pH.
2. The acid form of the buffer can be dissolved and its conjugate base can
be
formed by the addition of an appropriate amount of strong base.
HA + OH - = A- + H 2O
3. The salt of the base form can be dissolved, and an appropriate amount
of the acid form can be formed in solution by the reaction with a strong
acid.
A- + H + = HA
The Concentration of the Buffer
In making solutions we need to make up a given concentration of the solute in
a given volume of solvent. In Biochemistry the concentration of the buffer is
defined as the concentration of both the acid and the base forms of that
buffer. Thus all we need to know when making a buffer is its Mr, its
concentration and its volume. When we adjust the pH the forms will
proportion themselves out spontaneously to give the desired pH.
We will use Tris as a general example of how a buffer is made. First, weigh
out the correct mass, in grams, of the base form. “Correct mass” means the
mass that will give the desired molar concentration when dissolved in the
given volume of solution. We allow the solute to dissolve in a somewhat
smaller volume than the final volume calls for, and then titrate the solution
to the correct pH with hydrochloric acid. After the correct pH is reached
we add more solvent until the volume of the solution is brought to the
desired total volume, mix and check the pH. So this is an example of the
third method of making a buffer.
63
Procedure:
The recipe in the appendices in the lab manuals, unless specified
otherwise, indicates concentrations corresponding to their working, or 1X,
concentration. As an example of a buffer preparation, let us make 5L of a
4X solution of Tris HCl buffer with a working concentration of
0.1M.concentration at pH 7.6. TrisHCl indicates that HCl is the acid
needed to titrate the base form of Tris. You could also titrate with
H2SO4 Or CH3COOH to make TrisSO4 or Tris Acetate, if needed.
1. Find all the components of your assigned buffer. In this case the
solid Tris base and the HCl solution. All solids are in the west
balance room or by the pH meters. The Acids are under the hood in
the east lab. Look for the formula weight of Tris, or any solid, on
its container. Record the Mr in your lab book.
2.
Calculate the number of grams of each component you will need to
weigh out. (Remember, grams = 4(V x C x M r), where V = volume,
C = molar concentration and M r = molecular weight. This calculation
gives the mass of Tris base to be weighed out.
(4x5X0.1X121)g Tris base = 242g of Tris base.
3. When you get to the balance, first put the weighing container on the
pan and tare the balance. Using a clean spatula, add solid to the
container until the correct weight is obtained. Add the solid to a
beaker and rinse the weighing container with water into the beaker.
Repeat this for each solid component. When you are finished with
the balance, clean out any spilled chemicals with the brush
provided, and turn off the balance. Clean the used spatulas and
return them to their containers. Sloppy lab practices are not
tolerated.
4. Dissolve solids into somewhat less water than the total volume needed.
For our example the initial volume would be about 4.7L. Use a
magnetic stirrer and stir bar.
5. With the standard buffers provided, calibrate the pH meter
according to the published instructions. For maximum accuracy the
calibrations standards should bracket the desired pH. Place the pH
electrode into the buffer solution.
64
6. Titrate the buffer with strong acid, HCl to pH 7.6, the desired pH.
Rinse the electrode and put the electrode back into the storage
buffer. Turn off the pH meter.
7. Bring the buffer to the desired final volume by adding water and recheck your pH. Adjust again if necessary.
8. Transfer the buffer to an appropriate storage container. Label the
container with (a) the buffer name, (b) buffer concentration, (c) date
made, and (d) your initials.
8. To use the buffer dilute it buffer to its working, or 1X concentration.
For example, if you want 200 ml of a 1X solution starting from a 10X
solution then, use the calculation from genchem and insert the “X”
designation for concentration instead if molarity:
V 1 x C 1 = V 2 x C 2, (200ml)(1X) = (xml)( 10X)
Xml = 200(1X/10X) = 20ml.
9. Add 20 ml 10X to 180ml H2O, mix. Decant into appropriate container
for storage.
10. Clean all glassware used to make solutions. Hang out to dry on the
racks.
65
Reagents
Agarose Gels
Ampicillin Solution (1000X)
Buffers
Bradford Reagent for Protein
Assay
Column Buffer
Coomassie Blue Stain for Gels
Destaining Solution (SDS)
DNA Denaturing Solution
DNA Loading Solution (10X)
DNA Neutralization Solution
Hoescht Assay Solution
IPTG Solution (100X)
LB (Luria Broth)
Liquid Medium
LB (Luria Broth)
Add 0.5g agarose to 50 ml 1X TAE in a
Wheaton bottle. Microwave for 1’ 20”. Cool hot
solution under running water, and pour. This
makes a 1% gel
A filter-sterilized solution of 50mg/ml in
distilled water.
50mM appropriate buffer – weigh out enough
salt to give a 50mM concentration in desired
volume, dissolve in water, pH and then take to
volume. Check pH.
100mg Serva Blue dissolved in 50ml 95%
ethanol. Add 100ml 85% H3PO4, mix, add water
to one liter. Mix.
20mM TrisHCl, pH 7.4, containing 200mM NaCl
and 1mM EDTA.
4X 100mg Coomassie blue tablets in 1L of
40:10:50/methanol: glacial Acetic Acid: water.
40:10:50/methanol: glacial Acetic Acid: water.
0.4M NaOH containing 1M NaCl.
0.25g Bromophenol Blue, 0.25g Xylene Cyanol in
50ml 1mMTrisHCl, pH 8.0. Add 50ml glycerol
and mix.
1M phosphate pH 6.5.
Add 2μL 33528 Hoescht dye to 1 ml 2X TNE in
an eppendorff.
Make 100mM in water and filter-sterilize.
10g bactotryptone, 5g yeast extract and 10g
NaCl in one liter of water. Autoclave after
dissolving.
Add Bacto-agar to 1.5% (w/v) to LB, liquid
medium. Autoclave and pour into Petrie dishes.
66
Solid Medium
PNPG Solutions
SDS Loading Buffer (10X)
If antibiotics are to be added, wait until the
agar solution is about
65 C before adding.
p-nitrophenyl--D-galactose, PNPG, in Z
buffer.
Add 15 ml glycerol to 35ml 0.5M Tris HCl, pH
6.8, containing 4g SDS and 15mg Bromophenol
blue.
67
SDS Gel Electrode Buffer
(10X)
Separating Gel Solution
SSC (20X)
Stacking Gel
Substrate Solution for Blots
TAE Buffer (10X)
TE Buffer
Tetracyclin Solution (1000X)
TNE Solution – (2X)
Towbin Buffer
TSS Solution
Western Blocking solution
Western Salt Solution
Western Wash Solution
250mM Tris base containing 1.92mM glycine
and 1g SDS, pH 8.3.
Add 120ml 30% Acrylamide/bis-acrylamide to
280 ml of 0.54M TrisHCl, pH 8.8, containing
300μL of TEMED, mix. This will give a 9% gel
88.2g sodium citrate and 175.3g of NaCl per
liter. Adjust pH to 7.0 with glacial acetic acid
and autoclave before use.
13.6 ml of 30% acrylamide/bis-acrylamide to
87ml of 0.145M Tris HCl, pH 6.5, containing
100μL TEMED. This will give a 4% gel.
0.1M TrisHCl, pH 9.5, containing 50mM Mg+2
and 100mM NaCl.
TAE is an acronym for Tris-acetate-EDTA. We
buy a 50X TAE solution and dilute it with water
to make a 1X running buffer. The 50X contains
2M Tris acetate and 50mM EDTA, pH 8.3,
10mM TrisHCl, pH 8.0, containing 0.15M NaCl
and 1mM EDTA.
15mg tetracycline in 70% ethanol-water.
20mM TrisHCl, pH 7.4, containing 2mM EDTA
and 0.2M NaCl.
0.192M glycine, 0.025M Tris base, 0.0013M
SDS, pH 8.3. Make to 15% in methanol.
Add 10g of PEG 3350 or PEG 8000 to 100 ml of
sterile LB. Add filter-sterilized Mg+2 to a final
concentration of 50mM; adjust the pH to 6.66.8 and add enough DMSO to give a final
concentration of 5%(w/v) and filter-sterilize.
1g Blocker in 200ml Western Salt solution
containing 0.2ml Tween 20.
48.6g Tris base, 47gNaCl in 4L of water. Take
the pH to 9.5 with HCl.
0.2ml Tween 20 per liter Western salt solution.
68
eXgal Stock Solution
Xgal Agar Plates
YT Medium
Z Buffer
25mg Xgal/ml in 50%mDMSO/water.
Dilute the stock Xgal by a factor of 1000, to
give a final concentration of 25μg/ml.
Dissolve 0.8g bactotryptone, 0.5g yeast extract
and 0.5g NaCl in 100 ml of water. Adjust pH to
7.4 and autoclave.
50mM TrisHCl, pH 7.5, containing 50mM Mg+2
and 2mM DTT, dithiotreitol.
69