Laboratory 1: Cloning and Construction of Recombinant DNA
I. Background:
Molecular biology received a huge surge in 1951 with the publication of Watson
and Crick’s seminal paper elucidating the structure of DNA. The biotech revolution,
however, may have been launched in 1972 by Herbert Boyer and Stanley Cohen, who
developed a procedure for making “recombinant DNA”, i.e., DNA consisting of
fragments from different organisms. Their technique was the basis of what is now a
multibillion dollar industry. Its promise and perils have probably not yet been fully
appreciated.
Cohen’s lab had developed a method of introducing plasmids containing
antibiotic genes into certain bacteria, while Boyer’s group had isolated an enzyme that
cut DNA in a precise, predicable, reproducible manner. These enzymes, of which
hundreds have been now identified, are called “restriction enzymes”. In the molecular
biology laboratory, they are used like molecular scalpels. They allow us to combine DNA
from different sources (“recombinant DNA”). The focus of the conference at which they
met was bacterial plasmids. Plasmids are circular, replicating pieces of DNA that contain
specialized genes such as those that confer antibiotic resistance to those bacteria that
contain them. They are usually small (< 10 kbp), and are thus easily transferred between
bacteria and relatively easily isolated from the much larger, more complex chromosomal
DNA that contains the majority of the bacterial genome. Because of their ease of
handling, plasmids are the most commonly used vectors for the manipulation and transfer
of recombinant DNA.
Their combined skills allowed for the development of a technique in which a
piece of DNA can be cut from, say, a wombat and placed in a plasmid that can be moved
into a bacterium, eg. E. coli. Termed “subcloning”, this type of manipulation is
ubiquitous and fundamental to modern molecular biology. Subcloning involves cutting
DNA with a restriction enzyme or enzymes and ligation of the fragment into a compatible
vector. The enzyme used to catalyze the joining of the two pieces is suitably named
ligase, discovered in 1967. The recombinant molecule is then moved into an appropriate
host, a process known as transformation. While plasmids are the most oft used vector,
they are by no means the only type of vector, and bacteria are not the only hosts.
After subcloning and transformation, the host is subjected to selection. This
involves growing the recipient of the plasmid on a medium containing a selective agent
(usually an antibiotic) to which the plasmid confers resistance. For example, the Kanr
gene encodes an enzyme that inactivates kanamycin. Kanamycin is an aminoglycoside
that interferes with prokaryotic translation by binding to the 70S ribosomal subunit. The
resistance enzyme, a 3-aminoglycoside phosphotransferase, inactivates the drug by
covalent modification, eliminating the antibiotic’s affinity for the ribosomal subunit.
The fragment containing the Kanr gene is 1300 bp long, and is excised from a
larger fragment using the restriction enzyme, Eco RI. Like all restriction enzymes, Eco RI
cuts at a specific recognition site, usually characterized by a palindromic sequence. Eco
RI cuts at the following:
A map of the resulting fragment is below:
Note that the fragments resulting from Eco RI digestion contain unpaired bases on
the 5’ end of each strand. Draw it out if you’re not convinced; understanding this is
ESSENTIAL!! Thus, these overhangs (“sticky ends”) can base pair with complementary
sticky ends from other DNA fragments. Different restriction enzymes leave different
ends. There are 3’ overhangs, 5’overhangs, and blunt cutter. Each has its use in the
molecular biologist’s toolkit. Below are some examples. For practice, you draw the
products of a 6-cutter restriction enzyme that leaves 3’ dinucleotide overhangs. Can you
see the advantages of overhangs for routine subcloning?
The vector into which we will ligate the Kanr gene fragment is a 3000 bp plasmid,
pSP64, derived from the pUC series (plasmid University of California). Like many
popular plasmid vectors, pUC vectors have been engineered to contain several unique
restriction sites, including a single Eco RI site. The multiple unique sites are clustered in
a region of the plasmid called a Multiple Cloning Site (MCS) or polylinker. The
polylinker in pUC18 is 44 bp long, containing 13 unique restriction sites. Below is a map
of pUC18, showing the restriction sites and other regions of interest. Other noteworthy
features of pUC are the lac Z gene, within which the MCS resides, an ampicillin
resistance gene (encodes the enzyme -lactamase), and an origin of replication (ori),
necessary for replication of the plasmid. What is the plasmid’s shape after cutting with
Eco RI? How would the insertion of a 1300 bp fragment affect Lac Z function?
After cutting, the fragments need to be ligated together to achieve a recombinant
plasmid. Ligase, derived from a bacteriophage, catalyzes the formation of a
phosphodiester bond between the 5’ phosphate of one fragment and the 3’hydroxyl of
another (below A, B). Complementary base pairing between the sticky overhangs
stabilizes the duplex to facilitate the reaction. The reaction requires energy
(simplecomplex) in the form of ATP. Because the enzyme is isolated from infected
E. coli, the Tm for T4 Ligase is 37˚C, yet ligation reactions are routinely performed at 4˚C
to 22˚C. Can you offer an explanation for this apparent inconsistency? Predict the
product(s) of the ligation reaction. What are concatamers? How can we increase the
percentage of recombinants among the molecular pool? To which end of the polylinker
will the Kanr promoter be closest (the 5’ end or the 3’ end)?
A.
B. Mechanism of Ligase Reaction (E=Enzyme) (from the Biochem 201 site at Stanford
University.)
NAD
ATP
When a fragment and vector are cut with a single restriction enzyme, further
analysis is required to determine the orientation of the insert. Why? In this laboratory
investigation, we will take advantage of a unique Cla I site in the Kanr fragment and a
unique Pvu I site that sits 3’ to the MCS in pSP64 (below). By cutting with these two
restriction enzymes (a double digest) we can determine the orientation of our insert, an
issue that can be of great practical importance.
In the diagram above, notice that the ClaI site is ~ 120 bp downstream from an
initiator codon (ATG) in the kanr fragment, i.e., it is very close to one end of the insert.
The PvuII site is located ~180 bp to the right of the EcoRI site into which the kanr
fragment was inserted. By cutting with ClaI and PvuII, we can determine the orientation
simply by looking at the size of the fragment released. Do you see why?
After the ligation reaction has been performed and verified (how?), the
recombinant plasmid is introduced into a suitable host by a process called transformation.
There are different ways to introduce foreign DNA into bacteria. One of the simplest,
though least efficient, is called chemical transformation. In this procedure, the bacteria
are made competent by treatment with CaCl2 or other salts. The mechanism by which
cations facilitate uptake is not precisely understood. It may be that they cause the DNA to
adhere to the cell wall, increasing the probability of casual uptake. The mechanism
whereby heat shock improves uptake is also not completely understood.
The bacteria used in this experiment are E.coli strain HB101. This strain has no
antibiotic resistance, no plasmids, and lacks restriction enzymes. It also lacks an enzyme,
RecA, which is responsible for intracellular recombination. Thus, any plasmid taken up
will not be incorporated into the bacterial chromosome. These features are typical of
strains used in cloning and subcloning experiments.
Once transformed, recombinant cells will make copies of the plasmid, making it
easier to generate large amounts of DNA for subsequent manipulations, eg. sequencing.
The amount of plasmid required for transformation is extremely small. Typically, 10 ng
or less are sufficient. In fact, too much plasmid (>100 ng) can actually reduce
transformation efficiency. Several other factors also affect the efficiency of uptake.
Linear vectors and large concatamers are not taken up well. Generally, the smaller the
plasmid, the more efficiently it is taken up.
The uptake event itself is rare; ~ 1 in 10,000 cells will take up exogenous DNA.
The number of cells obtained per µg DNA used is called the transformation efficiency.
Transformation efficiencies of 105-106/µg are sufficient for most subcloning, but much
higher efficiencies, up to 1010/µg, can be obtained using another technique,
electroporation. Can you think of a circumstance under which these high efficiencies will
be required?
The presence of an antibiotic resistance gene on the plasmid allows for easy
selection of transformants. Our cells will be plated on agar containing kanamycin. The
vast majority of bacteria will die; only those containing a functioning plasmid will
survive. Each transformed bacteria will multiply, forming colonies that are easily scored.
After transformation, the selective pressure needs to be maintained; otherwise the
transformant will eventually be cured of the plasmid. Can you offer an explanation?
Single colonies will be transferred to liquid broth containing antibiotic. This will be
cultured overnight until the broth is saturated with bacteria. We will extract DNA from
bacteria grown in these overnight cultures.
Alkaline-lysis plasmid extraction is an old, reliable method for obtaining large
amounts of fairly clean, intact plasmid. For many procedures, this DNA can be used as is.
For others, eg. sequencing, the plasmid must be further purified. It is important to
understand the roles of the various components of the procedure. SDS is a detergent, and
will thus dissolve the cell membrane. The procedure begins at a high pH to degrade
RNA and aids in protein denaturation. High pH denatures large chromosomal DNA and
any linearized or nicked DNA fragments. Plasmid, on the other hand, is a covalently
linked circle, so is relatively unaffected by this treatment. The high ionic strength
facilitates precipitation of the larger DNA. The supercoiled plasmids remain in solution.
Acidified potassium acetate neutralizes the pH, and causes the precipitation of free SDS
and the associated SDS/protein/membrane complexes. Chromosomal DNA precipitates
with this agglomerate, while the most of the plasmid remains suspended. RNAse is often
included to further facilitate the degradation of any contaminating RNA.
The supernatant containing the plasmid is usually treated further to remove any
contaminating salts, small molecules or proteins. Typically, the super is treated with
phenol followed by chloroform. This denatures any proteins, and they are discarded with
the organic phase. Treatment with ethanol in the presence of salts (Eg. NaCl, ammonium
acetate) causes the plasmid to precipitate while leaving small molecular contaminants in
solution. Alternatively, matrices have been developed which selectively remove protein
and RNA or selectively bind DNA. These resins have speeded up the purification
process.
Once you have obtained putative transformants, you will need to definitively
confirm transformation by isolating the plasmid and performing restriction analysis. The
results will be visualized by agarose gel electrophoresis. You will perform this procedure
after the initial ligation and again after transformation. Although size is a major
consideration in determining the mobility of DNA in an agarose gel, one must also
consider shape and density. Thus, supercoiled plasmid will migrate faster than linearized
DNA of the same size. Obviously, nicked concatamers will travel according to their size
(i.e., dimers will be lighter, and will thus travel further, than tetramers), By digesting with
the appropriate restriction enzymes you will obtain a series of size-separated bands that
can be evaluated to determine both the presence of the desired insert and its orientation.
How many fragments should you obtain from a digest with EcoRI? PvuI? ClaI? PvuI and
ClaI? How can you use these digests to determine the orientation of the insert?
Gel electrophoresis is a technique used for the separation of nucleic acids and
proteins. Separation of large (macro) molecules depends upon two forces: charge and
mass. When a biological sample, such as proteins or DNA, is mixed in a buffer solution
and applied to a gel, these two forces act together. The electrical current from one
electrode repels the molecules while the other electrode simultaneously attracts the
molecules. The frictional force of the gel material acts as a "molecular sieve," separating
the molecules by size. During electrophoresis, macromolecules are forced to move
through the pores when the electrical current is applied. Their rate of migration through
the electric field depends on the strength of the field, size and shape of the molecules,
relative hydrophobicity of the samples, and on the ionic strength and temperature of the
buffer in which the molecules are moving. After staining, the separated macromolecules
in each lane can be seen in a series of bands spread from one end of the gel to the other.