CLONES
Basic Plasmid Cloning
Restriction enzymes are used to cleave the vector, and cleave a compatible fragments out
of the insert. DNA ligase is used to join insert to vector. The mixture is transformed into
E. coli, and then the bacteria are cloned. Each clone will have only one of the possible
ligation products, because of 'incompatibility' (meaning that if multiple plasmids of the
same kind enter a bacterium, all but one of them gets kicked out).
Modern cloning vectors have engineered multiple cloning sites. This is the map of
pUC19, from the New England BioLabs catalog:
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For a result with the minimal amount of trouble, one should gel purify both the cleaved
vector and the cleaved insert before ligating. This removes extraneous DNA species that
may interfere (the most troublesome of which is residual uncleaved vector). It also
allows verification that a fragment of the expected size has in fact been produced, and
allows crude quantitation of vector and insert. Vector and insert are usually ligated in ~
1:1 molar ratios. Excessive amounts of either or both can lead to concatemerization
instead of circle formation. Concatemers do not produce colonies on transformation.
Interfering enzyme activities can usually be removed by a heat treatment.
Type II restriction enzymes leave ends that may be 5' overhanging, 3' overhanging, or
blunt. In all cases each end is left with a 3' OH and a 5' phosphate. All blunt ends, and
any complementary overhanging ends may be religated with T4 DNA ligase, as long as at
least one 5' phosphate is present. A common cloning strategy is to dephosphorylate the
vector. This prevents ligation of vector ends to each other, but does not inhibit ligation of
vector to insert. Dephosphorylation has no impact on whether a restriction site is restored
after ligation.
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In the tables below G^AATTC means that the end will be:
-----G
-----CTTAA
3'
5'
GTGCA^C means that the end will be:
-----GTGCA
-----C
3'
5'
Compatible ends
Isoschizomers - Enzymes that recognize the same sequence. Isoschizomers may not
leave the same kind of end (and also may not be affected the same by methylation).
Type II enzymes cleaving different recognition sites may leave complementary
(compatible) ends. Clones that are made by joining compatible ends may not regenerate
either of the restriction sites, or they may generate some other site. For example an AccI
cut insert joined to a ClaI cut vector will create a junction recognized as a TaqI site.
Examples of compatibility groups:
AccI
ClaI
FspII
TaqI
GT^CGAC
AT^CGAT
TT^CGAA
T^CGA
BamHI
BclI
MboI
BglII
G^GATCC
T^GATCA
^GATC
A^GATCT
HgiAI
NsiI
PstI
GTGCA^C
ATGCA^T
CTGCA^G
All blunt ends are compatible:
HaeIII
AluI
SmaI
HincII
GG^CC
AG^CT
CCC^GGG
GTY^RAC; R is A or G; Y is C or T
Some other commonly used restriction enzymes that are incompatible with other ends are
listed below. Notice that for some cases ends produced by the same enzyme are not
necessarily compatible (eg. HinfI).
EcoRI
HindIII
SacI
HinfI
DdeI
BglI
G^AATTC
A^AGCTT
GAGCT^C
G^ANTC; N is A,T,G, or C
C^TNAG
GCCN4^NGGC
Verifying a construct by restriction endonuclease cleavage:
Most typically restriction digests are assayed by electrophoresis on a small gel of 0.7 to
1% agarose. This technique is fast and simple. Although polyacrylamide gels have
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higher resolution for fragments below 500 bp, most people try to get by with higher
percentage agarose, or certain brands of agarose engineered for resolution of smaller
fragments. Capillary gel electrophoresis is also possible, but the equipment available
within the department is configured for fluorescently labeled fragments, and is only
applicable to special applications.
Visualization: Although other staining agents are available, most commonly the DNA is
visualized by staining in 0.5 ug/ml ethidium bromide, which can either be soaked into the
gel or placed in the gel during its original formation. Visualization requires illumination
with a UV light. A wave length of 265 nm is most effective, but also does more damage
to the DNA and is a safety concern. Illumination at 300 nm is common, particularly if
the DNA is to be recovered from the gel for cloning. For imaging the gel may be either
illuminated from below or above. The investigator should never stare into a gel laying on
top of a UV light source, at least not without a UV face shield. There are a number of
high tech imaging devices ranging from specialized gel imagers to home-made CCD
cameras mounted on laboratory computers. The Biochemistry Dept. has a departmental
imager in 550C. Contact Sandy Mathis to arrange the required brief training if you wish
to use it. Sensitivity typically allows visualization down to 5-10 ng in a band. One
should always record an image of every gel.
The experiment depicted above is suboptimal for a number of reasons. Can you see why?
There should be more markers, and there should be markers extending below the lowest
experimental band of interest and markers better defining the onset of curvature. The gel
could have been run longer to make better use of its revolving power.

Understanding circular DNA
Plasmid DNA as isolated from cells is a mixture of forms: monomer covalently
closed circle (ccc) (supercoiled), dimer and higher forms of ccc, and nicked circles (alias
relaxed circles). The supercoils generally run as if they were about half the size of the
corresponding linear, although the relationship between ccc and linear migration is
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dependent on gel conditions. The sizing curve for circles remains linear higher up on the
gel.
On storage, ccc DNA converts slowly to the nicked circle (not linear) form. As soon as
one cleaves with a restriction enzyme, the pattern simplifies to the pattern of bands you
would expect from the map of the plasmid.
JOINING INCOMPATIBLE ENDS
Incompatible ends may be polished to blunt ends to permit ligation either by cleaving off
the overhang, or by filling in the overhang.
Alternatively, a synthetic linker may be used. The following linker is used to join blunt
ended fragments to an EcoRI site:
5
5'-pCCGGAATTCCGG-3'
Note that the linker is self complementary. When annealed to itself and ligated in excess
to blunt ended fragments, it creates an array of RI sites at each end:
RRRRRRR
RRRRRRR
RRR
||||||| frag.
||||||| frag.
|||
------------------------------------
Recleavage with RI leaves a fragment with RI ends. Note that you might have to first
methylate the fragment with EcoRI methylase to protect any internal EcoRI sites during
the recleavage. Alternatively, you could create the overhang directly with two separate
oligos, thus avoiding the recleavage:
5'-AATTCCCGGG-3'
GGGCCCp-5'
Note that by leaving the phosphate off of the upper strand, you preclude joining of linker
RI ends to each other. The phosphates for joining to the vector will have to be provided
by the vector ends.
Public repositories of cDNA clones and sequence information.
Large projects are underway to obtain and array cDNA clones from human, mouse, rat,
and zebrafish, etc; to add their sequences to the EST database; and to make the clones
physically available.
NIH's project is called Mammalian Gene Collection
(http://mgc.nci.nih.gov/). If you identify a clone you want by a database search, you can
obtain the cDNA clone from an associated supplier called the I.M.A.G.E. Consortium.
See: http://image.hudsonalpha.org/ (Bonaldo, Lennon and Soares, Genome Res., 1996,
791-806). An index of other projects attempting to compile full length cDNA clones for
a variety of genomes is found at http://www.ncbi.nlm.nih.gov/genome/flcdna. A number
of
commercial
enterprises
sell
clones
from
their
own
collection:
http://www.genecopoeia.com/,
http://www.origene.com/cdna/,
and
http://clones.invitrogen.com.
NCBI has a help page for finding cDNA clones through its database system:
http://www.ncbi.nlm.nih.gov/genome/clone/finding_cdna.shtml.
Map browsers for various genomes exist to enable correlating cDNA structure
with splicing patterns of various genomes:
 http://www.ncbi.nlm.nih.gov/Genomes/ - index of NCBI's map viewers.
 http://genome.ucsc.edu/cgi-bin/hgGateway?org=human - Gatway to UCSC's
genome browsers.
 http://www.ensembl.org/index.html - ensemble genomes
 http://img.jgi.doe.gov/cgi-bin/pub/main.cgi - D.O.E.'s Integrated Microbial
Genomes.
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PCR Amplification from mRNA to recover or extend a cDNA.
If one conducts first strand synthesis with reverse transcriptase, it is possible to
add PCR primers and complete the synthesis of a cDNA, as well as amplifying it, in the
PCR machine. This is sometimes called RT PCR (although the designation 'RT' is also
often used to mean 'real time' instead of 'reverse transcription'). Both primers could be
specific for a particular gene, or one specific primer could be used with oligo dT.
The following figure shows amplification between a gene specific primer (GS1)
and the poly(A) tail which has incorporated two improvements over the basic technique.
1) The reverse transcription is conducted with oligo dT with a 5' extension. The PCR
primer matches the 5' extension. This avoids trying to amplify with oligo dT itself, which
would force badly balanced TM's between the two primers. 2) The first population of
cDNAs produced are re-amplified with a second (nested) gene-specific primer. This
restores the specificity of a two primer amplification.
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A similar procedure can be used to extend towards the 5' end.
This procedure has been named RACE (Rapid Amplification of cDNA Ends).
Ref: Frohman et al., PNAS 85, 8998-9002 (1988) Current Protocols in Molecular
Biology (Ausubel et al.) has a more thorough accounting of this an most basic cloning
procedures.
Directly Cloning PCR Products
Remember that PCR primers as delivered from an oligonucleotide synthesis facility do
not have 5' phosphates. Hence a mixture of blunt ended PCR products should not be able
to ligate to one another. They can join to a blunt ended vector, if the vector has 5'
phosphates. However, the common practice of dephosphorylating the vector to prevent
circularization without an insert would also block ligation to the insert. In this case, one
would have to phosphorylate either the PCR primers before amplification, or the PCR
product after amplification. Since single stranded ends are more reliably phosphorylated,
it would be best to do it at the primer level. The classic way to measure efficiency of a
phosphorylation reaction is to use [32]P-ATP, and then measure the specific activity of
the product. Since PCR products typically have 3' overhangs (see below), direct cloning
is inefficient unless one first 'polishes' the ends by reaction with a DNA polymerase with
3' -> 5' exonuclease activity. The most commonly used enzymes are T4 DNA
polymerase or Pfu DNA polymerase (see below).
TA cloning
During PCR, the polymerase tends to add an extra untemplated base to the 3' ends
of each synthesized fragment. Most commonly, an A is added, essentially making a one
base sticky end. This arrangement can be used for highly efficient cloning into a vector
cleaved so as to leave a 3' T overhang. The most popular systems are vended by
InVitrogen, and are based on "Topo Cloning". In this procedure, the vector was opened
by a topoisomerase instead of a restriction enzyme. The topoisomerase cleaves to leave
the 3' T overhang, but also remains covalently attached to the vector. When provided a
PCR product with an A overhang, the topoisomerase will reverse its reaction and join the
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PCR product to the vector. No ligase in involved, and phosphorylation of the 5' ends of
the PCR product is unnecessary.
TOPO® TA Cloning® of Taq-amplified DNA (From the InVitrogen web site).
The topo cloning procedure is highly efficient. It does have the drawback that the
vector is expensive and loses efficiency during on storage. It also has the drawback that
one is limited as to the choice of vectors and vector sites into which to insert. One could
engineer any blunt end restriction site in a vector for TA cloning by cleaving the vector
and then adding the T overhang by exposing to Taq polymerase + dTTP. In this case the
joining would require ligase, and one would have to pay attention to 5' phosphates.
Different polymerases differ in their propensity to add the overhang, and in their
preference of which base to add [Hu, DNA Cell Biol. 12:763-70(1993)]. Furthermore,
the identity of the last templated base on the 3' end of the PCR product influences the
efficiency of adding the overhang, and the identity of the base to be added. For example,
Taq polymerase most efficiently adds an A if the last templated base is a C. Hence, I
recommend a practice of putting a G on the 5' end of PCR primers if possible. Even if
TA cloning is not part of the intended strategy, it helps to have it as a fallback option if
the intended strategy fails to produce clones.
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Pfu polymerase (best known for conducting PCR with higher fidelity than Taq
polymerase) has a 3' exonuclease that precludes leaving an overhang. Hence Pfu
polymerase would be a good choice if blunt end ligation was planned. However, Pfu
products could be given the overhang by exposure to Taq polymerase plus dATP after the
amplification, and Taq products can be polished to blunt ends by exposure to Pfu
polymerase after amplification. Finally, the addition of the overhang by Taq polymerase
is promoted by adding an extra 10 minute extension after the last cycle. If you use Taq
and don't want the overhang, You should remove this extension from the procedure and
also keep the other extension times as short as is practical.
Cloning PCR products with Restriction sites.
Alternatively, one can add a restriction site to the back of the PCR primers, and
then cleave the product to expose the cohesive ends. It is best to gel purify the PCR
product before attempting the cleavage, because the DNA polymerase will otherwise
modify the ends exposed by the cleavage. PCR primers may also interfere with some
restriction enzymes. After cleavage the fragments can then be ligated into a restriction
enzyme-cleaved vector in the ordinary way. It helps to gel purify the fragments again
after cleavage, since the little ends cleaved away will religate to both vector and insert
and inhibit production of the desired product.
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Six extra bases should be added behind each restriction site, so that the enzyme is not
trying to cut too close to the end of the fragment.
Troubleshooting:
A common problem with this strategy is that either not enough restriction enzyme
is used to cut off, or too much is used and the ends are damaged making them
unligatable.
 Not enough enzyme: Restriction enzymes generally come with activity
labeled in units similar to 10U/ul, meaning 1 ul of enzyme will cleave all
EcoRI sites in 10 ug of some test DNA in 1 hour under suitable reaction
conditions. Typically one uses a 10 fold excess to assure cleavage, given that
some sites are slower to cut than others, and the enzyme may have lost some
potency in storage. But the Unit = 1 ug/hour definition assumes some
particular density of EcoRI sites. If the test molecule was typical of ordinary
DNA sequence, it will have about 1 EcoRI site/2000 bp ( 1/4^6). For a 400
bp PCR product with an EcoRI site on each end, the concentration of EcoRI
sites is 10x higher than the test molecule the enzyme was assayed against. It
will be necessary to up the amount of enzyme used (or the length of the
reaction) by 10 fold to get comparable cleavage efficiency. Another problem
is that sites near the end of a DNA molecule tend to exhibit suboptimal
cleavage.
 Too much enzyme: Commercial restriction enzymes are not pure. They
contain activities such as phosphatases that will destroy the ligatability of the
ends produced. They are typically purified enough such that a 100 fold excess
of the enzyme leaves > 95% of ends in a ligatable condition. There should be
a specification sheet that came with the enzyme that states this. If a 1000 fold
excess of the enzyme is used (including amount of enzyme used and number
of hours reacted), then there may be damage to more than half of the ends and
the fragments produced will not clone. The onset of this problem also has to
be prorated by the number of ends/ug of cleaved product relative to the typical
test molecule producing 2 ends per 2000 bp. The usual way to get into this
problem is to react 1 ul of enzyme with some unknown tiny amount of DNA.
Problematically, a gel electrophoresis assay will usually not detect the loss of the
small piece of DNA cleaved away, and hence can not be used to diagnose the problem.
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Compounding this is that the problem might have nothing to do with ligation. The
problem could be in the transformation or electroporation procedure.
Ligatablilty after cleavage could be assayed by killing the restriction enzyme and
then just religating the mixture and running it on a gel. For example, if the above product
were cleaved with RI and then religated, you would expect 1/2 of it to dimerize, while the
remainder rejoined to the small piece cleaved away.
If this procedure is not working, an excellent option is to TA clone the whole
fragment, and then cleave the desired fragment out of the TA vector. In that case,
cleavage is unambiguous by gel assay.
Construction tricks with PCR
1) One can create a fragment with a blunt end at any position by placing the PCR primer
at that position. The blunt ended fragments can then be fused by ligation, or used
as described below as PCR primers in a subsequent construction step. The
overhanging A should be avoided for these purposes.
2) One can add any arbitrary sequence to the end of a PCR product by simply adding it to
the back of the PCR primer. This is illustrated above for adding a convenient
restriction site, but the concept can be extended to adding promoters, translation
initiators, or mutated sections of a coding region. Primers of up to 100 nucleotides
can now be synthesized relatively cheaply. Primers with large 5' extensions are
highly prone to false priming problems. Hence they should only be used on noncomplex templates (like a purified fragment or a plasmid clone). If you need to
both recover the fragment from a complex template (like the human genome), and
add 5' extensions, consider amplifying first with a version of the primers without
the 5' extensions and then using the purified product to reamplify with the larger
primers.
3) One can fuse any two overlapping fragments by coamplifying them with the right pair
of primers.
--->
============================ frag. A
========================== frag. B
<---resulting product:
=========================================
4) One can place any mutation (even a large deletion) into a fragment by placing the
mutation within a PCR primer.
5) One can rescue a band from a gel by pricking it with a pin and swirling the pin in a
PCR reaction mix with the appropriate primers. This also works even if the DNA
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is single-stranded, or if there isn't very much of it. Thus one can rescue a fragment
from an SSCP gel, or a hybridization filter.
Often an individual PCR product is purified from a complex PCR product by the
above method. The most common failure in this experiment is to add too much of
the template to the reamplification reaction. This results in a smear high up on the
gel and/or DNA stuck in the slot.
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Problems
1. A clone is described as having been made by cloning a SmaI fragment into an EcoRI
site.
Sequencing through the insert/vector junction, you find
GAATTCATCCCGGG. How did they do this?
2. You wish to add an elaborate promoter construct of 200 bp in length to the beginning
of a gene. The promoter design is theoretical; there is no piece of DNA to get it
from either by restriction enzyme cleavage or PCR amplification. You can locate
no oligonucleotide facility that is able to make a 220 bp PCR primer that might be
used to add this sequence in the same way a restriction site is appended by adding it
to the 5' end of the primer sequence. How might you build this sequence onto your
gene?
3. A colleague created PCR primers to add EcoRI sites to a segment of DNA during PCR
amplification. The PCR fragment would not clone after EcoRI cleavage.
Examination of the primer sequences reveal that the colleague neglected to add the
6 bases after the EcoRI site to the primer sequences. Can you think of a way to
salvage the PCR product without making another pair of primers?
4. A colleague obtains a cDNA clone. He runs some on a gel uncut and some cut with an
enzyme that is supposed to cleave once in the circle. He gets the patterns below,
and notes that relative to a marker lane on the gel, the cut material is the wrong
size. What's the problem with this experiment?
last revised 3/29/2011 - Steve Hardies
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