Biotechnology
Homework 3 Fall 2010
Answers
Vectors for cloning large DNA segments
1. (i) The objective is to generate as many different end-points for clones as possible. XhoI sites
are not as frequent as Sau3A sites.
(ii) There are two significant considerations. The smaller point is that large gaps between clone
end-points is proportionately more significant for smaller inserts than for larger ones. More
significantly, BAC clones have flexibility in their sizes. Thus, for any XhoI site cutting at the left
end of a fragment there would likely be around 20 or more right-hand XhoI cuts which could
generate a fragment of suitable size for cloning. With lambda, however, where the suitable insert
size range may be about 15-25kb there may most commonly be only one or two suitable righthand XhoI cuts (if any). This will severely diminish the number of potential insert fragments that
can be cloned and lead to several regions with no representatives at all.
(iii) Three factors were explicitly mentioned for Sau3A libraries in lambda phage, so those issues
simply have to be adapted to this question.
(1) Size selection; purify (from a gel) fragments from the partial XhoI digest that are in the range
of roughly 15-25kb
(2) Treat the insert fragments with alkaline phosphatase so they cannot ligate to each other.
(3) Avoid self-ligation by using the inverse of the strategy shown in the slides for Sau3A
fragments and XhoI-cut vector. Here, fill the insert XhoI sites with T and C, and fill vector
BamHI-cut ends with G and A, then the modified vector & inserts can ligate only to each other.
This would, of course, be an alternative to phosphatase treatment.
2. (i) Purify a restriction fragment from close to each end of the 20kb insert (by using your map to
identify convenient fragments). Hybridize to the genomic library, pick cognate clones, make
DNA and map. The clones should overlap the original probe and extend in both directions,
probably occupying at least 50Kb (but repeat the process if necessary).
(ii) (a) Hybridize the labeled cDNA to a Southern blot of the cloned recombinant phage DNAs
(although you could do a genomic Southern that would be many times less sensitive and might
miss short regions of complementary sequence). From your map you can see which genomic
regions hybridize and hence a minimal span of the transcription unit.
(b) Since genes can give rise to many transcript structures and your cDNA might not be fully
representative or, indeed, even full-length you may be missing part of the transcribed region.
You may also be missing that region from the cloned recombinant genomic DNA you were
testing. Beyond those considerations, cDNAs cannot show you where regulatory, nontranscribed regions of a gene lie. Some such sequences are typically 5’ from the start of the
transcript but it is not unusual for enhancer sequences to be many kb distant and also to be (in
part) 3’ of the transcript.
(iii) The cosmid vector is cut at one site to ligate to an insert just like with a plasmid vector.
Additionally, it would typically be cut between the Cos sites so that the products of ligating to an
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insert are linear. After ligation the products are packaged in vitro into phage and used to infect
cells. Once inside cells the Cos sites anneal to circularize the DNA, nicks are repaired and it
behaves just like a ColE1 plasmid.
Two reasons were emphasized previously for why large DNAs are hard to clone in ColE1
plasmids- reduced efficiency of transformation and instability of the clone because slow growth
introduces strong selective pressure for rare variants to take over a population (in a nascent
colony or in liquid culture). The second (more serious) issue is not changed by using a cosmid.
A cosmid does, however, allow high efficiency introduction of DNA into a cell (because the
packaging extract is very efficient and because ligation can be maximized by using DNA at high
concentration since circles do not need to form). Hence, the (former) value of cosmids (it is true
that they hold more DNA than lambda vectors but the comparison here is with ColE1 plasmids).
After the introduction of electroporation, however, there was no serious constraint on the
efficiency of introducing 50kb molecules into E.coli, so I think there is currently no significant
virtue of cosmids. The ColE1 stability issue was overcome by using low-copy BACs.
(iv) The vector must be cloned in E.coli as part of the process of constructing it, and then to have
adequate quantities of starting material. Theoretically that could now be accomplished simply
through PCR reactions but it is much more efficient to clone.
(v) After ligation DNA is introduced into yeast by a chemical procedure or electroporation, and
yeast are plated on selective media so that only those with the YAC vector sequences can grow.
Only large enough YACs (with suitable inserts) are stable and hence all emerging colonies will
also have large inserts.
PLEASE START A NEW PAGE
3.
(i) The PCR product must be made with primers that add attB1 and attB2 sequences at either end.
Fortunately these are only 21bp long and can be added relatively conveniently at the 5’ end of
primers with around 20-25nt that hybridizes to the target DNA.
(ii) The Entry to Destination vector reaction is reversible. Thus, if you add the second product of
that reaction (a kanamycin resistant vector with ccdB flanked by attP1 and attP2) and BP clonase
(rather than LR clonase) you will make an entry clone from a destination clone (select for
kanamycin resistance in cells killed by the ccdB gene). You could also, of course just use the
destination vector as a template for PCR to make a product exactly analogous to the one you
inserted into the entry vector in the first place.
(iii) (a) You could directly infect cells in liquid culture and allow growth over several hours to
amplify the phage population. It is better, however, to spread infected cells on plates to produce
plaques because (i) it allows you to determine the size of the library (how many independent
clones) and (ii) amplification of phage is less competitive so different clones will grow more
evenly (but certainly not equally- significant distortions in representation are inevitable). You
then wash the plates to elute much of the phage into liquid (with Magnesium to stabilize the
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phage). These procedures will amplify at least a million-fold or so. Hence, there is plenty to go
around and you should be very generous with your friends (now you are so phage-rich).
(b) If it were not for the internal EcoRI sites you could simply purify the mixed phage from liquid
culture, isolate the mixed DNA, cut with EcoRI and ligate to EcoRI cut plasmid (using suitable
amounts to create a few million colonies). However, inserts with EcoRI sites would not be
cloned intact because the DNA would lose its methylation when amplified in lambda clones.
You might postulate some rare-cutters, like NotI, are immediately adjacent to the EcoRI inserts in
the lambda vector (true in many vectors) and think about excising the fragments en masse that
way. That would be an improvement but there are still going to be some clones with internal
NotI sites (or whatever other site you pick).
A better strategy, suggested by the subject matter of Q3, is to amplify the whole batch of
recombinant lambda DNAs with primers containing attB1 and attB2 sequences hybridizing to
vector sequence either side of the inserts. The whole set of PCR products can then be cloned into
Entry vector.
4.
(i) No, primer hybridizing to synthesized DNA will have a break in the template immediately
adjacent to the hybridization site. The original DNA strands serve as templates throughout the
procedure. Hence, production of newly synthesized DNA is not exponential and accumulate only
arithmetically.
(ii) Any double-stranded molecule containing an original DNA strand will have methylated
GATC on at least one strand and will be cut into fragments, preventing such strands from ever
creating a viable plasmid transformant.
(iii) The relevant molecules are those in which two complementary synthesized strands hybridize
to each other. Those strands may initially form a linear molecule with complementary singlestranded overhangs equal in length to the length of the primers (typically 30nt or more). Those
ends will anneal well under conditions of the annealing phase of PCR during each cycle (after the
first). That annealing prevents any action of DNA polymerase (linear molecules would have the
sticky ends filled in giving dead-end molecules) during the upcoming cycle. At each cycle those
molecules would be denatured but they can anneal again in the next cycle. Those annealing
during the last cycle are the actual molecules that will produce the desired clones (DNA
synthesized in the last round serves only to ensure DpnI digestion of template strands- DpnI acts
only on double-stranded DNA).
(iv) You simply design the primers so they lack the 30bp region and have adequate length of
sequence either side to ensure hybridization. To be conservative that length should be at least
15nt but I would prefer even longer.
5.
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(i) The PCR primers must include at the 5’ end some length of identical sequence for any two
fragment ends intended to be joined (if you use the vector as is without PCR you would add
vector sequences at a site opened up by a restriction enzyme as the sequence segment to be added
to the insert). To allow directional cloning you would use different sequences for the two
junctions being made. The longer the common sequence the more efficient will be the annealing
in vitro. However, you should also note that annealing will only be effective if the exonuclease
treatment makes the entire common sequence (and therefore likely more in most cases) singlestranded.
(ii) All of the approaches are fairly easy once you are used to them. To me that means they are
efficient and reliable, as well as being easy to apply (e.g. by designing appropriate PCR primers).
In some cases (Gateway) you need to purchase specific reagents.
The short discussion following assumes you are cloning a cDNA into a vector as a primary
example (just for convenience). Junctions with naturally occurring restriction sites are seamlessyou don’t have extra or altered sequences at the junction. However, you rarely find restriction
sites exactly where you want them so your design of junctions is almost always compromised.
Also, each design is case by case and sometimes there is no good solution-far from a universal
application of identical strategies that work in all cases. Gateway cloning involves addition of
fairly long segments of DNA (att sites of different types) between tags and a cDNA or between
promoter elements and a cDNA. It is not seamless. It is universal because the nature of the DNA
being cloned is of no consequence- the same strategy works every time. PCR cloning by adding
restriction sites overcomes the major shortcoming of just cloning with restriction enzymes. You
can place a new restriction site where you want it. Sometimes this changes the normal sequence
but generally minimally, so it is close to seamless. The technique can work in every case but the
exact sites used will depend on the DNAs involved (sites in the vector and sites that must be
absent from the body of a cDNA), so the design is not universal (one design fits all) in that sense.
PCR followed by exonuclease & then hybridization can be used seamlessly (if you add sequences
present from one molecule-here vector- to another rather than adding the same foreign sequence
to both DNAs to be joined). It can be applied to any junction. Also, if you were happy to add
extra DNA at the junctions you could always add the same sequence if you wished & hence apply
the technique universally. Perhaps the biggest limitation is that you do need fairly long PCR
primers to hybridize to templates and include at least 30nt of homology to the sequences you are
joining to.
(iii) Using the strategy of (i) you could design your PCR primers so that (a) two overlapping
segments of a circular plasmid are converted into linear DNA products, (b) the overlap entirely
corresponds to the primer sequences (just like in Quik-change mutagenesis), and (c) the primers
are designed to have the required alterations to the normal plasmid sequence (making the primers
long enough that they hybridize well despite some mismatches.
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