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The vector product obtained by ligation between a vector previously digested with antarctic phosphatase lacks 5' phosphate groups. T4 ligase can ligate it to an insert with complementary sticky ends which provides 2 phosphate groups. But there are still 2 nicks in the vector product! How are these normally repaired ?
The nicks are not repaired during preparation. Instead, the vector (which is circular, despite the nicks) is transformed into the host where it is repaired by endogenous mechanisms, likely through base excision.
Molecular Biotechnology, Chapter 3
They are restriction endonucleases that can cut DNA molecules @ specific sites called recognition sites. The recognition sites contain PALINDROMIC DNA SEQUENCES and are between 4 to 8 bps long.
The most important basis of recombinant DNA technology is the presence of restriction endonucleases. Restriction enzymes cleave the large DNA molecules into smaller and manageable fragments. This is required to manipulate the DNA fragment and in gene cloning experiments. Also, the cleaving of DNA by restriction enzymes is HIGHLY SPECIFIC. THUS, restriction endonucleases help in cutting the DNA sample in a reproducible fashion.
The plasmid pCEL1 is a circular double-stranded plasmid and is treated with restriction enzymes one at a time and in combinations. The bands obtained in based pairs:
SO the size of the plasmid pCEL1 is 6 kilobase pairs bc it yields a single fragment of length 6.0 kb ON DIGESTION w/ EcoRI, BamHI, and HindIII. It also has single recognition site for these 3 enzymes. It has three sites for HaeII
size of plasmid --> 4.361 bp
has two genes that give resistance to ampicillin and tetracycline. they are referred as Amp^r and Tet^r genes.
Tet^r gene has unique BamHI, SaII, and HindII recognition sites. The amp^r gene has a unique PstI site, There is a unique EcoRI gene outside of the coding DNA
it has an origin of replication for the replication of DNA that works in E.coli
it is maintained as a high copy number plasmid include the E.coli cell.
The target DNA is now combined with the plasmid DNA and ligated using the enzyme T4 DNA ligase and ATP. This yields a mixture of recombinant plasmids having the target DNA and non recombinant plasmids that arise due to self-ligation of the plasmid DNA. These plasmids are then transformed into the host E.coli cells.
The cells containing the cloned DNA are IDed by screening the TRANSFORMED cells for the two antibiotic resistance markers that are present on the plasmid.
Immediately after transformation, the cells are incubated in an abx free medium to allow the cells to grow and express the abx resistance genes. The cells are then grown in the abx whose resistance gene in the plasmid is intact.
so like for example, if target DNA is inserted in BamHI site of the Tet^r gene, then the cells are 1st grown in ampillcin-containing medium (resistance gene in the plasmid is intact) The cells that have not transformed do not survive in this medium. The transformed cells with and without the target DNA grow on this medium
The insertion of the target DNA into the BamHI site disrupts the Tet^r gene and the cell containing this plasmid is no longer resistant to tetracycline. It is only resistant to ampicillin. If the plasmid does not contain the DNA insert, then it has an intact Tet^r gene and is resistant to both the antibiotics.
therefore in the next step, the cells have grown on ampicillin medium are plated onto a medium with tetracycline by marking the relative positions of the colonies.
Only cells that contain the intact Tet^r gene that is the self-ligated plasmids, grow on this medium.
size --> 2,686 bp
it has a gene that gives resistance to the abx ampicillin. referred to as amp^r gene.
it also contains a region of B-galactosidase gene (lacZ) that is part of the lactose operon in E.coli. this is under the control of the lac promoter that can be regulated. the lacI gene is also present and produces the repressor protein that helps in regulating the lacZ gene
it has an origin of replication from the plasmid pBR322 for the replication of DNA that works in E.coli.
contains a short DNA segment called the multiple cloning site or polylinker. this has a # of unique cloning sites that are clustered together. for example, EcoRI, BamHI, HincII, PstI, SacI etc are some of the sites present in the multiple cloning site. This is present in the lacZ gene and does not affect the production of the enzyme
the target DNA is now combined w/ the plasmid DNA and ligated using the enzyme T4 DNA ligase and ATP
this yields a mixture of recombinant plasmids having the target DNA and non-recombinant plasmids that arise due to self-ligation of the plasmid DNA. these plasmids are then transformed into the host E.coli cells.
the host cells should be able to produce the other part of B-galactosidase called LacZa. this combines with the portion of the enzyme produced by lacZ gene and forms an an active enzyme.
a medium containing ampicillin, IPTG and X-Gal is used to grow the treated host cells. IPTG acts as an inducer of the lac operon and X-Gal acts a substrate for B-galactosidase. upon hydrolysis, X-Gal produces a blue colored compound. the presence of ampicillin ensures that the non-transformed cells do not grow on the medium
cells that have transformed with self-ligated plasmids produce the active B-galactosidase enzyme and hydrolyze X-Gal to produce blue colored colonies. Cells that have been transformed with the recombinant plasmid will not produce the active enzyme and produce white colored colonies. this is because the target DNA that is introduced into the multiple cloning site disrupts the lacZ gene and so THE ACTIVE ENZYME IS NOT PRODUCED
- T4 DNA Ligase Buffer (NEB #B0202) should be thawed on the bench or in the palm of your hand, and not at 37°C (to prevent breakdown of ATP).
- Once thawed, T4 DNA Ligase Buffer should be placed on ice.
- Ligations can be performed in any of the four standard restriction endonuclease NEBuffers or in T4 Polynucleotide Kinase Buffer (NEB #B0201) if they are supplemented with 1 mM ATP.
- When supplementing with ATP, use ribo ATP (NEB #P0756). Deoxyribo ATP will not work.
- Before ligation, completely inactivate restriction enzyme by heat inactivation, spin column (NEB #T1030), or Phenol/EtOH purification.
- Heat inactivate (Antarctic Phosphatase, Quick CiP, rSAP) before ligation.
- Keep total DNA concentration between 1-10 µg/ml.
- Vector: Insert molar ratios between 1:1 and 1:10 are optimal for single insertions (up to 1:20 for short adaptors). Insert: vector molar ratio should be 6:1 to promote multiple inserts. Use NEBioCalculator to calculate molar ratios.
- For cloning more than one insert, we recommend NEBuilder ® HiFi DNA Assembly Products.
- If you are unsure of your DNA concentration, perform multiple ligations with varying ratios.
Subcloning is the process of moving insert DNA from one vector to another by means of preparing the DNA insert and new vector separately, ligating them together, and screening clones of the resulting mixture to identify the desired subclone.
Subcloning is done because the new vector or new arrangement of insert DNA(s) in the planned subclone offers some advantage over the old vector or arrangement.
Though there are as many potential advantages as there are vectors and there are hundreds if not thousands of different vectors. However, there are some common themes. One is that the optimal cloning vector, lambda phage, is not optimal for producing large amounts of DNA easily for use in other purposes, such as fragment isolation for subcloning. To solve this, DNA newly cloned from a lambda library usually is immediately subcloned into a plasmid vector, which offers much greater yields of DNA much more easily than lambda phage.
In a typical case, a cDNA (complementary DNA) for a specific gene is isolated by cloning a single recombinant phage (bacterial virus) from a recombinant phage cDNA library. The phage has a very large DNA genome and the cloned cDNA represents just 2 to 5% of it. After a day of work, one can isolate perhaps 25 micrograms of phage DNA, which contains just 0.5 to 1 micrograms of the cDNA. Subcloning it into another general purpose subcloning vector first, simply to allow more of it to be easily made. From such a subcloning vector construct, it is easy to make multiple milligram amounts of the cDNA in a short time.
Another reason to subclone is to put the DNA of interest into a new context for expression as RNA and/or protein. Typically, expression requires a promoter and terminator to start and stop mRNA synthesis and a components needed for translation: a ribosome binding site and open reading frame starting with a start (ATG) codon and ending with a stop codon. The vectors that are designed for gene expression usually provide everything except the open reading frame. Typical cloning vectors (usually phage) and typical general purpose subcloning vectors (plasmids) do not contain all of these elements. Expression elements differ greatly and are not compatible between types of organisms where the expression is desired: bacteria, yeast, insect cells, vertebrate animal cells, plant cells, so a different vector must be used for each.
Finally, to obtain specific changed in DNA, it is manipulated enzymatically (completely in vitro - in the test tube). This can be done to produce fusions of two pieces of DNA, deletions of some of the DNA, or mutations (changes in the DNA sequence). Typically, these enzymatic manipulations produce the desired DNA in very small and often unknown and undetectable quantities. To be useful, the resulting (desired) DNA must be subcloned and amplified in bacteria.
PCR and its use for Subcloning
PCR can be used to prepare DNA of interest for ligation into a subclone or for other in vitro manipulations such as DNA sequencing or restriction mapping. However, when used preparatively like this, PCR can introduce problems. One problem is that PCR produces a fairly small amount of DNA, with decreasing yield as the DNA gets longer. PCR does not work very well at all with DNA over 1000 base pairs, and most cDNAs are larger than that. Also, PCR amplification introduces mutations randomly at a rate of one per 2 or 3 kilobase-pairs, so any resulting subclones (or DNA sequence or mapping) may be done starting with a complex mixture of amplified DNAs. Also, PCR amplified DNA is heavily contaminated by the short single stranded DNA primers that interfere with the processing needed to subclone DNA made by PCR. Such primers must be removed prior to any processing, and this is an extra step that is not trivial to accomplish without losing a significant amount of the desired PCR product. Also, DNA made by PCR amplification, which is linear, is best subcloned if its ends are cut with one or two restriction enzymes, yet many restriction enzymes don't cut PCR amplified DNA well near its ends unless conditions are specially optimized. To make matters worse, unlike inserts cut from plasmids, one cannot tell by agarose gel analysis if the ends of the PCR amplified DNA have been cut by restriction enzymes, so a failure to cut shows up as a subcloning failure only after several days of additional effort.
Advanced PCR methods allow for PCR-mediated ligation of the amplified DNA into the vector by using PCR primers that anneal both to the sequence of interest and to the vector DNA. Though this requires longer primers, the cost for synthesis of primers has dropped dramatically in recent years such that longer primers are quite affordable. The problem of low yield is not important with these methods since the PCR-mediated insertion does not require the use of DNA ligase and bacteria can amplify even the vanishingly small quantities of circular DNA product from this method. The major remaining problem is that of the high error rate of high-temperature DNA polymerases used in PCR.
Traditional (Excision-Ligation) Subcloning Methodology
Preparing the Insert DNA
The "insert" DNA, which is the DNA to be put into a new vector via subcloning, can be obtained from another plasmid, from phage DNA, from genomic DNA, from cDNA made by reverse transcriptase, or from any of these DNAs that has been amplified by PCR (polymerase chain reaction). Also, synthetic DNA can be used as an insert. However, see the section above describing the disadvantages of preparing DNA to use for subcloning by PCR amplification. In the most traditional form of subcloning, the DNA to be subcloned is excised from another DNA clone or subclone using restriction endonuclease digestion.
Traditionally, the insert DNA cut out of its source DNA (usually phage or plasmid) by restriction digestion, must be isolated from the other pieces of DNA resulting from the digest reaction. This separation of the DNA fragments is done by agarose gel electrophoresis. The DNA band of interest is then literally cut from the gel with a razor blade and the DNA in the piece of agarose is extracted by one of a variety of methods. These methods remove the fragments of agarose that interfere with the subsequent ligation step.
For subcloning, the choice of how to cut the insert DNA out of its previous (initial) DNA context depends on what restriction enzyme sites are available for that purpose and where else those enzymes cut the insert DNA and its intial vector. It is desirable to have the enzymes that cut out the insert DNA not cut inside the insert DNA and to leave a short single stranded overhang that will be compatible with the overhang made when opening the subcloning vector DNA (see below). If possible, two different enzymes should be used to cut out the insert DNA (one on each end), as that allows for "directional subcloning" (see below). In some cases it may not be possible to avoid cutting with an enzyme that also cuts inside the insert DNA. In such cases a partial restriction digest is performed to produce the desired DNA fragment. That is done by cutting the starting vector with insert in a reaction that does not go to completion so only some of the DNA sites are cut. Since the enzyme cuts the sites in each molecule randomly, all possible partial digestion products are produced, including some DNA fragments contining internal, uncut restriction sites. In some cases, a "blunt-end" restriction enzyme must be used for cutting one or both ends of the insert DNA. In those cases, subcloning is more difficult because there are no "sticky ends" (overhangs) to assist the DNA joining by DNA ligase. Such ligations, while possible, are somewhat less efficient (fewer positives out of the resulting subclones) than those with fragments with overhangs on both ends. Since the ends of the prepared insert DNA must be able to be ligated with the prepared ends of the vector DNA, it may be that although overhangs can be produced on each, they may not be able to be generated with the same enzymes due to the lack of available or/or useful recognition sequences. In some cases enzymes with different recognition sequences can produce compatible overhangs (such as BamHI and BglII or NcoI and SalI). Otherwise, overhangs can be converted to blunt ends via trimming or filling in the recessed end with DNA polymerase.
The Vector DNA
Since the goal of subcloning is to create a new DNA construct (vector + insert) containing the insert DNA, the insert DNA fragment will be ligated into a site on the desired vector DNA (typically a plasmid). To prepare the vector DNA for this ligation, it must be cut (opened) at the site where the insert is desired to be ligated. Unlike insert DNA, partial digests typically are not feasible with vectors and the insertion site must consist of one or two unique restriction enzyme sites at the appropriate place on the vector relative to sequences needed for expression or other biological function (if any) and relative to each other. The presence or absence of the appropriate restriction sites in the insert DNA and vector strongly constrain the subcloning process and when those constraints are too constraining, considerable effort often must be given to working around them. Sometimes, when a specific site is not present in the insert DNA, the insert DNA can be subcloned first into another plasmid's multiple cloning site, then it can be cut out with the enzymes needed for subcloning into the real target vector. Alternatively, restriction enzyme sites in a vector can be changed by opening the vector and ligating in to that opening synthetic DNA "adapters" or "linkers" that introduce a new restriction endonuclease site. Finally, a restriction site can be added to an insert DNA where there was none before by using site-specific mutagenesis, a mostly distinct methodology.
In practice, for preparation of the vector DNA, often it is sufficient to simply to cut it with the one or two enzymes needed to open the subcloning site then to denature or otherwise remove the restriction enzymes as they will interfere with the subsequent ligation. However, for several reasons, it is often desirable to gel purify the linearized (cut) vector DNA by separation on an agarose gel and isolation of the DNA from the gel by cutting out piece of agarose as was described above for the insert DNA. For linear DNA vectors such as lambda and derivatives, each "arm" of the vector DNA is isolated as distinct DNA fragments. One reason is for gel purifying the vector DNA fragment(s) to be used in the subcloning is that this eliminates the need to use highly purified vector DNA (i.e., over CsCl density gradient). Secondly, when removed by gel purification, there is no chance of the "stuffer fragment" (the piece of DNA between the two restriction sites on the vector) to be re-ligated into the vector during the subcloning. Third, the vector may not be cut open completely, and any remaining uncut vector tends to interfere strongly with the subsequent screening of subclones. Uncut original vector usually can be removed easily by gel purification both of linear vectors (because of the size differences) and purification of plasmid vectors, since circular and supercoiled plasmid DNA migrate differently in a gel than the linear DNA fragment. Generally, it is always best to gel purify the vector DNA since it provides more robustness (less probability of failure). Since the insert DNA usually is gel purified, the vector can be done at the same time for virtually no extra labor cost.
In most cases, it is important that the insert DNA be subcloned such that it is in one correct orientation in the vector site, typically for expression as RNA and/or protein. If the vector is cut open with a single restriction enzyme and the insert DNA has the same overhangs on both ends, it is possible for the insert DNA to be ligated into the vector site in either of two orientations. If this is done, the subclones must be screened not only for the presence in the insert DNA but also for the orientation of the insert relative to the vector. Such "like-ended" ligations also are subject to the vector re-ligating in the absence of any insert DNA, which results in a high "background" of insert-negative candidate subclones. Both of these problems can be eliminated if two different restriction enzymes are used to open the vector and to prepare the insert DNA. For this to work properly the two enzymes must leave incompatible DNA "overhangs" so they will not ligate with the other end. This limits ligation of the insert into the vector to cases where the insert is oriented in the desired way. This is called "directional subcloning." Like all good things, it introduces at least one new problem. While cutting the insert out of its previous vector using two different restriction enzymes, it is easy to tell from the fragment prep gel if the digest went to the desired level of completion by the presence of the desired insert DNA fragment, the same is not usually true for double cutting the new vector. While cutting the new vector DNA with two enzymes, the sites are usually so close to each other (in the "multiple cloning sequence" or MCS) that once cannot tell from a gel separation if both enzymes or only one cut the vector. If one of the two enzymes cut inefficiently, the ligation will have a high background and a relatively low yield of the desired subclone. To make matters worse, many restriction enzymes do not cut well near the end of a DNA fragment, so often cutting with one will inhibit the reaction by the other. Also, if one of the two enzymes used is a blunt-end cutter, the reduction in efficiency of the ligation is so large that it is usually better to stick with a "like-ended" (single enzyme site) ligation with an enzyme that leaves an overhang, and simply screen the subclones for orientation later.
Reducing the background of like-ended ligations by removing the 5' phosphate at the ends of the vector DNA
The vector DNA in a "like-ended" ligation type of subcloning was opened by cutting with a single restriction enzyme. This means that the ends of the vector DNA can re-anneal and be ligated without the insert DNA during the ligation reaction. This is because the ends of a DNA cut with a restriction enzyme can always re-anneal and be ligated together, essentially reversing the action of the restriction enzyme. Not only is it possible for the vector to re-ligate with itself, this unimolecular reaction is actually strongly favored over the bimolecular reaction involving a molecule of vector with a molecule of insert DNA. This re-ligation of the vector without insert, however, does not produce the desired subclone, and so is an unwanted side reaction product. This side reaction negatively affects the process of subcloning in two ways. First, it consumes the vector so that less of it is available to anneal with and be ligated to the insert DNA. Secondly, it causes a larger background of insert-negative colonies in the subcloning, decreasing the fraction of positive subclones (efficiency) and potentially increasing the number of subclones that will need to be screened to get the desired construct.
There is a method that reduces the possibility of re-ligation of the vector without any insert DNA. This method involves treating the cut vector DNA with a phosphatase that removes the phosphate left on each 5' end of the cut DNA. Without a phosphate on the 5' end, the 5' and 3' ends of the vector strands cannot be ligated by DNA ligase (which requires the 5' end to have a phosphate on it). Since neither strand of the vector DNA can be re-ligated with their other end, the vector cannot be recircularized without insert. If insert is present and anneals with the ends of the vector, the DNA ligase can ligate one of the two strands (the one with the 5' end of the insert DNA and the 3' end of the vector DNA), but not the other one. Since the insert DNA is usually very long, there is a lot of DNA base-pairing between the two single strand breaks ("nicks") left after the ligation of insert with phosphatased vector is complete. So even though the DNA is not fully ligated (both strands have a break in them) the DNA appears to the bacteria as if it is fully ligated for purposes of it being taken up by the bacteria. Once inside the cells, the single strand breaks are repaired by normal DNA repair processes.
Ligation, transformation, and plating for colonies
Ligations only require an enzyme (T4 DNA ligase), buffer (including the magnesium and ATP needed) and the vector and insert DNA fragments. After incubating these together the entire mixture is put into bacteria, either by "transforming" bacteria made competent by chemical treatment or by electroporation (making holes in the bacteria with a short electrical pulse). These bacteria are then grown on agar plates containing an antibiotic (typically ampicillin) that prevents growth of bacteria that did not receive any vector DNA. After overnight incubation some of the numerous bacterial colonies on the plates ("subclones") are transferred to individual liquid cultures and small amounts (typically 3 ml of culture) are grown up as suspensions for isolation of plasmid DNA, which is then screened. Usually only 5 to 10 subclones need to be screened for any ligation. Subclones also can be screened immediately without growing them up as small suspension cultures by using analytical PCR. However, PCR screening usually only detects the presence of some of the insert DNA. (usually it is to long to detect all of it), and does not check uif the insert is intact. It can be used to screen orientation of the insert (one primer in the insert and one in the vector), but this does not detect unusual (and admittedly uncommon) rearrangements that sometimes occur during ligations, particularly during subclonings that occur at low efficiency.
Because the plasmid isolation and screening usually take an additional day or two, it is extremely helpful to avoid doing this if the ligation did not work well. Such early warning requires some useful observation to be made at the time of the appearance of bacterial colonies on the plates. Unfortunately, two simple methods are neither reliable nor widely applicable. One simple method is to use a vector with a LacZ (beta-galactosidase) A-chain which complements a short version of LacZ in the cells and catalyzes a reaction that turns the colony blue if a chromogenic substrate (X-Gal) is included in the plate agar. If the insert DNA is present, the LacZ A chain sequence in interrupted and a functional version of it does not get made and the colony does not turn blue. This method of prescreening colonies cannot be used in expression vectors because the LacZ sequences interfere with the expression of the vector cDNA. Also, it often fails, since the bacteria often will re-ligate (close) an opened vector without including an insert DNA yet often with a small mutation at the subcloning site that prevents the A chain of Lac Z from working (hence white colonies without inserts). This is particularly common with blunt-ended inserts and directional subcloning, but occurs frequently even with like-ended subcloning (particularly if the vector DNA was treated with phosphatase).
The second simple method to get early warning about a failed ligation is to just count the colonies of subclones. However, the number of colonies varies a lot, in part due to variation of the background and positive counts. However, simple colony counts are informative when they are compared to the proper controls. To do this, at the same time as a regular ligation reaction (involving vector and insert DNA) is done, a second, control ligation is done that contains vector DNA but not insert DNA. The colony counts from the control ligation can serve as an indication of the background. If the ligation reaction containing insert DNA results in a significant increase in colony counts compart to the insertless control reaction, then the ligation probably was successful. If there is no increase in colony counts resulting from inclusion of the insert DNA in the ligation reaction, then the chances are that the ligation failed. Note that this interpretation makes the logical assumption that the vector cannot ligate without insert. This is a valid assumption for directional subcloning and for like-ended ligations in which the vector has been treated with phosphatase, but it is not valid with like ended ligations that did not employ phosphatase treatment of vector DNA.
Screening of subclones
The isolated subclone plasmid DNA usually is screened by restriction enzyme digestion and agarose gel electrophoresis. Typically (though not always), this is done with the enzyme or enzymes used to open the vector and prepare the insert. For like-ended ligations, the screening process usually involves two stepss. First the screening must be done to find which subclones are positive for the presence of the insert DNA using the enzyme that was used to prepare the insert DNA and vector DNA for the ligation. The second step then analyzes those positive for insert DNA to determine the insert orientation in the vector. This "orientation screening" is usually done with one or two enzymes that cut the vector and the insert DNA asymmetrically to distinguish the two orientations. The two orientation can be predicted to yield different framgent patterns from the digest, and these are used to assign the orientation of the insert in each candidate subclone. In some subclonings, the first screening and orientation screening can be accomplished with a single screening digest, though such a digest cannot be done with the same enzyme that was used to open the vector for the ligation.
In the present study, we have demonstrated that integration of a long AT-DRS into a non-fragile region can drive fragile site expression. This provides a clear evidence for the direct role of AT-DRSs in driving chromosomal fragility. We further found that DNA fragments derived from the integrated AT-DRS have an increased tendency to form stable secondary structures under single stranded state which explains their ability to perturb DNA replication leading to genomic instability ( 20). The observation that a major proportion (∼45%) of long AT-DRSs along the human genome reside within cytogenetically defined unstable chromosomal regions further highlighting that AT-dinucleotide repeats are a key factor predisposing chromosomal regions to fragility.
The breakage frequency of the newly generated FS was significantly lower than the frequency at the endogenous FRA16C, from which the AT-DRS was derived. This indicated that other factors are contributing to the FRA16C fragility. Interestingly, in FRA16C, in addition to the long AT-DRS studied here, there are three additional long (>400 bp) AT-DRSs, which were shown to stall the replication fork progression under replication stress conditions ( 20) and are probably contributing to the higher fragility in the endogenous site. Moreover, replication timing data (http://www.replicationdomain.com) reveals that the endogenous FRA16C region is replicating later in S-phase relative to the HPRT locus, thus may further increase the difficulty to complete the replication along this region before entry into mitosis. It will be interesting to further characterize the novel generated fragile site, by studying the effect of the AT-DRS on the ability of the region to complete the replication during S-phase. Recent finding showed that fragile regions fail to complete their replication during S-phase and the under-replicated sequences are subjected to repair synthesis during mitosis termed MiDAS ( 34).
The question of whether integration of CFS sequences are sufficient to recapitulate FS-like instability was previously addressed using whole BACs containing FRA3B sequences ( 39). Integration of two adjacent FRA3B BACs, 150 kb each, into a non-fragile ectopic site in human cells was able to confer FS-like instability, implying an inherent instability of these sequences. However, a specific sequence motif responsible for the observed fragility in that study could not be identified. In the present work, however, we showed that a long AT-DRS is able and sufficient to drive the formation of an FS in a non-fragile region in the studied cellular system.
A number of fragile sites are enriched with AT-DRSs that have the potential to form alternative non-B DNA secondary structures ( 10, 21, 29, 40). AT-DRSs >200 bp in length are predicted to readily form secondary structures following unwinding of the DNA double helix during replication ( 10). Under aphidicolin treatment, which leads to uncoupling between the DNA polymerase and the helicase ( 41) longer stretches of ssDNA are exposed, which may enhance the formation of stable secondary structures of the AT-DRSs. Three in vitro studies tested the replication dynamics within plasmids containing different AT-DRSs derived from FRA16D ( 21, 22) and an AT-DRS derived from FRA16B ( 40). In one study ( 21), an ∼500 bp sequence which spans a FRA16D region highly enriched with perfect AT repeats, was shown to stall replication fork progression in yeast in a manner depending on the repeat length. Mfold predictions and gel electrophoresis migration analysis of these fragments suggested that the observed replication perturbation involves secondary structure formation ( 21). This suggestion was recently reinforced by showing that this AT-DRS is targeted by the MUS81 structure-specific endonuclease which cleaves the replication forks stalled at DNA secondary structures along this region ( 42). In another study, cell-free assays showed alleviated polymerase stalling within FRA16D-derived AT-DRSs by the addition of the Werner helicase, implicated in processing of secondary structures arising during replication ( 22). In the third study, a FRA16B-derived AT-rich fragment, cloned into a SV40 replication plasmid, was able to fold into branched secondary structures, promoting polymerase stalling ( 40). The probability to fold into secondary structures was increased when the structure-prone strand served as the lagging strand template ( 40). Recently, using HR reporters it was shown that the Bloom helicase activity and the FANCM fork reversal activity are required for preventing double strand breaks (DSBs) formation along AT-DRSs derived from FRA3B, FRA16C and FRA16D ( 14, 15). In an additional recent study, genome-wide analysis in human cells identified structure forming repeats, including quasi palindromic AT-rich repeats, as principal sites of fork collapse upon ATR inhibition ( 43). All these studies demonstrate that AT-DRSs can perturb the replication progression and generate DSBs. In the current study however, we have directly demonstrated that an AT-DRS that was shown to stall the replication fork progression ( 20) and form highly stable secondary structures (Figure 3B) has the ability to create recurrent chromosomal fragility in a non-fragile region. This provides for the first time a direct evidence for the role of the difficult to replicate AT-DRSs in the mechanism underlying fragile site formation.
DNA DSBs induced by hydroxyurea treatment were recently detected within poly(dA:dT) tracts in the CFSs FRA3B and FRA16D, suggesting that in addition to AT-DRSs, other sequences with potential to form non-B DNA sequences may also contribute to the fragility of CFSs under various stress conditions ( 44).
The landscape of CFS expression differs across cell types. This has been attributed to differences in the availability of replication origins and differences in replication timing programs among cell types ( 6). However, for example, FRA16D and FRA3B that are highly expressed in lymphocytes in which they exhibit a paucity of replication initiation events ( 45), are also expressed in fibroblasts (although to a lesser extent) in which replication initiation events are distributed along the fragility core ( 32, 45, 46). This intrinsic instability suggests that features as AT-DRSs which reside along FRA16D and FRA3B ( 18, 47) predispose these regions to breakage, independently of the tissue specific replication program and together with other key features underlying the fragility along CFSs.
CFSs are preferentially unstable in precancerous lesions and during cancer development [reviewed in ( 9, 48)]. A comprehensive study of the gene pairs involved in all recurrent chromosomal translocations in tumor cells found that over half of breakpoints are mapped to human fragile sites ( 49). Sequences within and flanking three randomly selected pairs of translocations prone genes showed enrichment of AT-DRSs, that were shown to form highly stable secondary structures ( 49). Another study analyzed ∼20 000 translocation breakpoints in cancer genomes and found significant association with potential non-B DNA forming sequences, including AT-DRSs ( 50). AT-DRSs were also found as hot spots for translocations causing genetic syndromes, as in the case of the translocation between chromosomes 11 and 22, t(1122), which underlies the Emanuel syndrome or the supernumerary-der(22)t(1122) syndrome. In most patients the translocation breakpoint is found at the center of the AT-DRS ( 51, 52). All these studies highlight the role of AT-DRSs in genomic instability driving cancer and genetic diseases. In summary, the results presented in the current work highlight the deleterious effect of intrinsic DNA features such as the AT-DRSs in driving recurrent genome instability.
1.4: DNA Modifying Enzymes
- Contributed by Michael Blaber
- Professor (Biomedical Sciences) at Florida State University
Just as the study of the bacterial restriction-modification system has provided a variety of specific endonucleases, there are also available a variety of specific DNA methylases.
- The recognition sequences of the methylases are the same as the associated endonucleases (e.g. EcoR1 methylase recognizes and methylates at the sequence "GAATTC").
- All methylases transfer the methyl group from S-adenosylmethionine (SAM) to a specific base in the recognition sequence, and SAM is a required component in the methylation reaction.
- Methylation of DNA usually has the effect of protecting the DNA from the related restriction endonuclease. However, there are methylases with minimal specificity. For example, Sss I methylase will methylate cytosine residues in the sequence 5' &hellip CG &hellip 3'. In this case, the methylated DNA will be protected from a wide variety of restriction endonucleases.
- Some restriction endonucleases will only cut DNA at their recognition sites if the DNA is methylated (e.g. Dpn I).
- Still other restriction endonucleases will cut both methylated and non-methylated DNA at their recognition sequences (e.g. BamH I).
Dam and dcm Methylation
- The methylase encoded by the dam gene (dam methylase) transfers a methyl group from SAM to the N6 position of the adenine base in the sequence 5' &hellip GATC &hellip 3'.
- The methylase encoded by the dcm gene (dcm methylase) methylates the internal cytosine base, at the C5 position, in the sequences 5' &hellip CCAGG &hellip 3' and 5' &hellip CCTGG &hellip 3'.
- Almost all strains of E. coli commonly used in cloning have a dam+dcm+ genotype. The point here is not that we particularly want our DNA to be methylated, but that to make a dam-dcm- host someone has to mutate the bacteria and isolate the correct mutant. That apparently has not been done for a lot of bacterial strains. Probably because the dam and dcm methylation affects only a small subset of potential restriction endonucleases
DNA isolated from dam+dcm+ strains will not actually be cut by a modest subset of available restriction endonucleases:
|Recognition sequence||Restriction enzyme||GATC||G me ATC|
DNA may have to be prepared from E. coli strains which are dam-dcm- in order to be cut by these enzymes.
A wide variety of polymerases have been characterized and are commercially available. All DNA polymerases share two general characteristics:
- They add nucleotides to the 3'-OH end of a primer
- The order of the nucleotides in the nascent polynucleotide is template directed
Figure 1.4.1:DNA Replication
In addition to the 5'->3' polymerase activity, polymerases can contain exonuclease activity. This exonuclease activity can proceed either in the 5'->3'direction, or in the 3'->5' direction.
- Exonuclease activity in the 3'->5' direction allows the polymerase to correct a mistake if it incorporates an incorrect nucleotide (so called "error correction activity"). It can also slowly degrade the 3' end of the primer.
- Exonuclease activity in the 5'->3' direction will allow it to degrade any other hybridized primer it may encounter. Without 5'->3' exonuclease activity, obstructing primers may or may not be physically deplaced, depending on the polymerase being used.
Different polymerases have differing error rates of misincorporation, and different rates of polymerization.
|E. coli DNA polymerase I||E. coli DNA polymerase I - Klenow Fragment||T4 DNA polymerase||T7 DNA polymerase||Taq DNA polymerase||M-MuLV Reverse Transcriptase|
|5'->3' exonuclease activity||*||*|
|3'->5' exonuclease activity||*||*||*||*|
|Error Rate (x10 -6 )||9||40||<1||15||285|
Uses of polymerases
The various activities of the different polymerases lend them to a variety of applications. For example, restriction endonucleases can yield fragments of DNA with either 3' or 5' nucleotide "overhangs".
- In the case of 5' overhangs, the 5'->3' polymerase activity can fill these in to make blunt ends.
- In the case of 3' overhangs, the 3'->5' exonuclease activity present in some polymerases (especially T4 DNA polymerase) can "chew back" these ends to also make blunt-ended DNA fragments.
Figure 1.4.2:Polymerase activity
This method is used to obtain highly radiolabeled single strand DNA fragments, which makes use of 5'->3' exonuclease activity present in some polymerases (E. coli DNA polymerase I, for example).
- In this method a DNA duplex of interest is "nicked" (i.e. one of the strands is cut see DNAse I).
- Then DNA pol I is added along with radiolabeled nucleotides. The 5'->3' exonuclease activity chews away the 5' end at the "nick" site and the polymerase activity incorporates the radiolabeled nucleotides. The resulting polynucleotide will be highly radiolabeled and will hybridize to the DNA sequence of interest.
- Thermostable polymerases have the ability to remain functional at temperature ranges where the DNA duplex will actually "melt" and become separated. This has allowed the development of the "Polymerase Chain Reaction" technique (PCR), which has had a profound impact on modern Biotechnology. We will discuss this method at a later date.
- The incorporation of dideoxy bases (i.e. no hydroxyl groups on either the 2' or 3' carbon of the ribose sugar) leads to termination of the polymerase reaction. This will be discussed in greater detail later. However, this chain termination by incorporation of dideoxynucleotides is the basis of the Sanger method of DNA sequencing, as well as therapies to try to inhibit viral replication.
- This is an exonuclease (starts at the termini and works inward) which will degrade both 3' and 5' termini of double-stranded DNA. It will not make internal cleavages ("nicks"), however, it will degrade the ends of DNA at existing internal "nicks" (which create both 3' and 5' termini).
- The degradation of termini is not coordinated, meaning that the product is not 100% blunt-ended (even though the original duplex may have been blunt ended).
- Such "ragged" ends can be made blunt by filling in and chewing back by a suitable polymerase (e.g. T4 DNA polymerase). The unit definition of 1 unit is the amount of enzyme required to remove 200 base pairs from each end of duplex DNA in 10 minutes at 30 °C.
Figure 1.4.4:Nuclease BAL-31 activity
- Catalyzes the stepwise removal of nucleotides from the 3' hydroxyl termini of duplex DNA.
- The enzyme will attack the 3' hydroxyl at duplex DNA with blunt ends, with 5' overhangs, or with internal "nicks".
- Since duplex DNA is required, the enzyme will not digest the 3' end of duplex DNA where the termini are 3' overhangs.
Figure 1.4.5:Exonuclease III Activity
Mung Bean Nuclease (isolated from mung bean sprouts)
- A single strand specific DNA and RNA endonuclease which will degrade single strand extensions from the ends of DNA and RNA leaving blunt ends.
- The single strand extensions can be either 5' or 3' extensions - both are removed and a blunt duplex is left.
Figure 1.4.6:Mung Bean Nuclease activity
Deoxyribonuclease I (DNAse I) from Bovine pancrease
- This enzyme hydrolyzes duplex or single DNA strands preferentially at the phosphodiester bonds 5' to pyrimidine nucleotides
- In the presence of Mg 2+ ion, DNAse I attacks each strand independently and produces nicks in a random fashion (useful for nick-translation)
- In the presence of Mn2+ ion DNAse I cleaves both strands of DNA at approximately the same position (but leaving "ragged" ends)
- Ligases catalyze the formation of a phosphodiester bond between juxtaposed 5' phosphate and 3' hydroxyl termini of nucleotides (potentially RNA or DNA depending on the ligase).
- In a sense, they are the opposite of restriction endonucleases, but they do not appear to be influenced by the local sequence, per se.
- Ligases require either rATP or NAD+ as a cofactor, and this contrasts with restriction endonucleases.
The following are different types of ligases and their characteristics.
T4 DNA ligase
- Isolated from bacteriophage T4.
- Will ligate the ends of duplex DNA or RNA.
- This enzyme will join blunt-end termini as well as ends with cohesive (complementary) overhanging ends (either 3' or 5' complementary overhangs).
- This enzyme will also repair single stranded nicks in duplex DNA, RNA or DNA/RNA duplexes. Requires ATP as a cofactor.
Taq DNA ligase
- This ligase will catalyze a phosphodiester bond between two adjacent oligonucleotides which are hybridized to a complementary DNA strand:
Figure 1.4.7:Taq DNA ligase activity
- The ligation is efficient only if the oligonucleotides hybridize perfectly with the template strand.
- The enzyme is active at relatively high temperatures (45 - 65 °C). Requires NAD+ as a cofactor.
T4 RNA ligase
- Will catalyze formation of a phosphodiester bond between RNA/RNA oligonucleotides, RNA/DNA oligonucleotides, or DNA/DNA oligonucleotides.
- Requires ATP as a cofactor.
- This enzyme does not require a template strand.
T4 RNA ligase can be used for a variety of purposes including constructing RNA/DNA hybrid molecules.
Figure 1.4.8:T4 RNA ligase activity
DNA ligase (E. coli)
- Will catalyze a phosphodiester bond between duplex DNA containing cohesive ends.
- It will not efficiently ligate blund ended fragments.
- Requires NAD+ as a cofactor.
Figure 1.4.9:DNA ligase (E. Coli) activity
T4 polynucleotide kinase
- Catalyzes the transfer and exchange of a phosphate group from the g position of rATP (adenine ribose triphosphate nucleotide) to the 5' hydroxyl terminus of double stranded and single stranded DNA or RNA, and nucleoside 3' monophosphates.
- The enzyme will also remove 3' phosphoryl groups.
- Oligonucleotides which are obtained from automated synthesizers lack a 5' phosphate group, and thus, cannot be ligated to other polynucleotides.
T4 polynucleotide kinase can be used to phosphorylate the 5' end of such polynucleotides:
Figure 1.4.10:T4 polynucleotide kinase activity
The In-Fusion™ cloning method was originally developed to produce a common entry vector for a multi-vector cloning system based on the cre-lox recombination (Clontech). As shown here and by others ( 18) (and Andrew Farmer: Clontech, personal communication), the enzyme can be used in combination with any 15 bp homology region to enable ligation-independent cloning of PCR products. We have exploited this property, in combination with single and multiple promoter vectors, to produce a simple and versatile system for expression of recombinant proteins. We have incorporated the method into a semi-automated pipeline for both vector construction and expression screening using standard laboratory liquid handling instruments. The pilot experiments in 96-well format showed cloning efficiencies of 89% after two clones per target tested, with 94% achievable with four clones tested, which we would recommend to maximize cloning efficiency. The reason(s) for some PCR products not cloning is not clear although the quality and quantity of the input DNA appears to be important. Over the last 12 months in the OPPF, we have used In-Fusion™ to construct a total of 661 vectors from 703 PCR products, an overall cloning efficiency of 94%. These results compare favourably with data reported by other HTP structural genomics consortia using either the Gateway™ recombinatorial system (e.g. 79% PCR product to expression clone efficiency ( 19, 20)) or ‘classical’ restriction enzyme and ligation-dependent methods (e.g. 87% ( 1)).
Multi-promoter vectors have been used to express recombinant proteins in more than one host in parallel, typically baculovirus-infected insect cells and E. coli e.g. ( 21, 22). Similarly, the pOPINE, pOPINF, pOPINJ or pOPINM vectors based on pTriEx2 (Novagen) described in this study can be used to survey expression in E. coli, insect cells, and mammalian cells from a single vector, which can be easily constructed in HTP mode with the associated savings in time and resources. Transient transfection of mammalian cells, notably COS7 and HEK293, is widely used to investigate the expression of eukaryotic proteins and can be readily scaled up for production purposes ( 7, 12, 23, 24). We describe the derivation and use of a secretion vector based on the pOPINF/pTriEx2 vector backbone for the HTP expression of extra-cellular proteins and domains. In addition, the expression from single vector constructs in all three hosts (E. coli, HEK293T and Sf9 cells) has been demonstrated for five target proteins. This ability to screen in multiple hosts would enable rapid scale-up in the most appropriate host (determined by small-scale screening) without the necessity for additional rounds of sub-cloning. The transient transfection protocol (HEK293 cells) and co-transfection protocols (Sf9 cells), which make use of a commercial transfection reagents, have both been automated to enable consistent HTP operation for functional/structural proteomic applications. As far as we are aware, this is the first report of such implementations.
In summary, we have addressed the primary characteristics we set for an improved strategy for HTP cloning as follows:
The ability to clone genes-encoding proteins, or domains thereof, in a rapid and reliable parallel or high-throughput (HTP) fashion. Cloning efficiencies of >90% have been achieved in a parallel 96-well plate format using a generic vector in combination with the In-Fusion™ enzyme.
The ability to accurately determine the final constructs without the addition of extraneous/vector or restriction-site-derived amino acids to the expressed protein. A suite of vectors has been described which make use of In-Fusion™ cloning to precisely engineer the expressed sequence, for example, introducing an N-terminal His6 tag and 3C protease cleavage site to enable removal of the tag during purification. The utility of this format has been exemplified (Case study I).
The process must be versatile in terms of insert sequence independence. Three case studies have been described in which eighty-two expression vectors have been produced from a variety of target genes using the vector system (Case studies I, II, III).
The process would preferably be single-step, to give rapid and cost-effective vector construction. Vector construction involves a single-step reaction that enables the simple and rapid construction of vectors in parallel using a 96-well plate format. Further, certain PCR products are compatible with multiple fusion vector formats (e.g. N-His, N-His-GST and N-His-MBP—see examples in Case study III) thereby enabling cost-effective use of PCR primers.
The constructs should be capable of expressing proteins from multiple hosts, i.e. a single vector capable of expression in E. coli, mammalian cell lines (e.g. HEK293T cells) and insect cell lines (e.g. Sf9 cells). By adapting a multi-promoter vector a single cloning step has been used to produce a single vector suitable for expression in E. coli, mammalian and insect cells. The utility of this has been exemplified (Case study III).
The expressed proteins must be capable of purification in HTP mode, i.e. they must all be fused to a common affinity purification ‘tag’ which may be removed, if desired, by enzymatic digest prior to crystallization. All vectors, regardless of additional fusion protein expressed, add either an N-terminal or C-terminal His6 tag onto the expressed target protein to facilitate detection and purification. The purification of proteins expressed both intracellularly in E. coli and secreted from mammalian cells using a common purification strategy has been exemplified (Case studies I and II).
The process should be amenable to automation. Laboratory automation to carry out liquid handling tasks involved in the various stages of the expression screening experiments in all three hosts has been described.
Cernunnos Promotes Joining of Mismatched Ends and Noncohesive Ends.
We purified Ku, DNA-PKcs, XL, and Cernunnos to apparent homogeneity (Fig. 1 A) and tested their ability to join cohesive 5′ ends (EcoRI–EcoRI) or cohesive 3′ ends (KpnI–KpnI). To measure joining efficiency, we used quantitative PCR, which amplified junctions produced from one pair of ends. The PCR primers eliminated signals from competing junctions, such as those produced from intramolecular ligation into circular monomers or intermolecular ligation of other ends. Ku, DNA-PKcs, and XL joined EcoRI–EcoRI ends and KpnI–KpnI ends with efficiencies of 1.2–2.2% (Fig. 1 B, rows 1 and 2). However, the addition of Cernunnos had no effect on joining efficiency.
Cernunnos stimulates the joining of mismatched DNA ends. (A) Purification of NHEJ proteins. Coomassie staining after SDS/PAGE shows our purified preparations of Ku, DNA-PKcs (Pk), Cernunnos (C), and XL. (B) Cernunnos stimulates the joining of noncohesive ends by Ku, DNA-PKcs, and XL. Linear DNA substrates were incubated with no protein (dark gray bars), Ku, DNA-PKcs and XL (light gray bars), or Ku, DNA-PKcs, XL, and Cernunnos (black bars). The orientation of DNA is depicted in the upper left corner. We tested compatible ends (rows 1–3) with cohesive 5′ overhangs (EcoRI–EcoRI), cohesive 3′ overhangs (KpnI–KpnI), or blunt ends (EcoRV–EcoRV). We also tested eight combinations of mismatched ends (rows 4–11). Note that SacII creates a 2-nt 3′ overhang. All other 3′ and 5′ overhangs were 4 nt. We measured joining efficiency by quantitative PCR and expressed efficiency as the percentage of input DNA ends joined (18). Because of large differences among experiments, we plotted joining efficiency on a logarithmic scale. Concentrations of Ku, DNA-PKcs, XL, and Cernunnos were 5, 5, 0.5, and 2.5 nM, respectively, here and elsewhere unless otherwise noted. (Insets) The overhangs from the KpnI–SacI, Acc65I–EcoRI, and EcoRI–BamHI ends.
Next, we tested whether Cernunnos might affect the joining of noncohesive DNA ends. The addition of Cernunnos to Ku, DNA-PKcs, and XL stimulated the joining of two blunt ends (EcoRV–EcoRV) by 6-fold (Fig. 1 B, row 3). In the absence of Cernunnos, Ku, DNA-PKcs, and XL joined blunt ends paired with either 5′ or 3′ overhangs with low and approximately equal efficiencies (Fig. 1 B, rows 4–7). However, the addition of Cernunnos stimulated joining 30- and 55-fold for the blunt-3′ ends, EcoRV–KpnI and EcoRV–SacII, and 8- and 9-fold for the blunt-5′ ends, EcoRV–EcoRI and EcoRV–BamHI. Thus, Cernunnos had a greater effect on blunt-3′ ends than on blunt-5′ ends.
Cernunnos also stimulated joining of ends with mismatched overhangs by 40- to 150-fold (Fig. 1 B, rows 8–11). Ku, DNA-PKcs, XL, and Cernunnos joined mismatched 5′ overhangs (Acc65I–EcoRI, 0.0039%) with 23-fold lower efficiency than mismatched 3′ overhangs (KpnI–SacI, 0.091%). However, Cernunnos facilitated more efficient joining when the 5′ overhangs contained partially complementary sequences (BamHI–EcoRI, 0.25%).
In summary, Cernunnos exhibited a preference for 3′ overhangs over 5′ overhangs. This preference also occurred when we omitted DNA-PKcs from the joining reaction [supporting information (SI) Fig. 6]. The effect of Cernunnos appeared to be specific, because Cernunnos had no effect on ligation of blunt ends to 3′ overhangs by bacteriophage T4 DNA ligase (SI Fig. 7).
Cernunnos Requires Ku to Promote Mismatched End Joining by XL.
We were particularly interested in the joining of mismatched 3′ overhangs because they are produced during V(D)J recombination in vivo (15) and because Cernunnos stimulated their joining by 40-fold in vitro (Fig. 1 B, row 9). Interestingly, purified XL joined KpnI–SacI ends with low but detectable efficiency, although the addition of Cernunnos failed to stimulate joining by XL alone (Fig. 2 A). However, even a substoichiometric concentration of Cernunnos (0.05 nM) stimulated joining by XL (0.5 nM) in the presence of Ku (5 nM) and DNA-PKcs (5 nM). A stoichiometric excess of Cernunnos (15 nM) stimulated joining 200-fold, up to a level of nearly 1%.
Ku, DNA-PKcs, XL, and Cernunnos act in concert to join mismatched 3′ overhangs. (A) Cernunnos stimulates the joining of mismatched ends with Ku, DNA-PKcs, and XL but not with XL alone. DNA substrates with mismatched 3′–3′ overhangs (KpnI–SacI) were incubated with XL alone or XL, Ku, and DNA-PKcs (Pk). Concentrations of Ku, DNA-PKcs, and XL were 5, 5, and 0.5 nM, respectively. Cernunnos (C) was added at concentrations of 0.05, 0.1, 0.5, 2.5, 5, and 15 nM. The point in the lower left shows the background PCR signal in the absence of protein. Error bars represent the variation in duplicate measurements. (B) Cernunnos requires Ku to promote mismatched end joining by XL. DNA substrates with mismatched 3′ overhangs (KpnI–SacI) were incubated with different combinations of the core NHEJ proteins Ku, DNA-PKcs, XL, and Cernunnos (2.5 nM).
To determine which proteins were required for joining mismatched ends, we incubated the DNA with Ku, DNA-PKcs, XL, and Cernunnos in different combinations (Fig. 2 B). As single proteins, only XL produced detectable joining of the KpnI–SacI ends. The addition of Ku and DNA-PKcs to XL increased joining efficiency 12-fold. Starting from the reaction with all four protein preparations, we omitted each protein one at a time. The omission of DNA-PKcs reduced joining only 4-fold, but the omission of Ku or Cernunnos reduced joining 150-fold or 40-fold, respectively. Note that the omission of Cernunnos reduced joining 200-fold when the concentration of Cernunnos was 15 nM rather than 2.5 nM (Fig. 2 A). Significantly, the omission of XL eliminated joining activity completely. Thus, Ku, XL, and Cernunnos were required for efficient joining of mismatched ends. These data suggest that Cernunnos and Ku stimulated a latent joining activity for mismatched ends contained in Ligase IV.
Cernunnos Promotes the Preservation of 3′ Overhangs.
To further characterize the joining of mismatched KpnI–SacI ends, we sequenced the junctions created by Ku, DNA-PKcs, and XL with or without the addition of Cernunnos. Surprisingly, the sequence from one 3′ overhang was retained in every junction, whereas the sequence of the other 3′ overhang was lost (Fig. 3). In the absence of Cernunnos, 21 of the 22 sequenced junctions retained the 3′ KpnI overhang, whereas only 1 junction retained the SacI overhang. This preference occurred only in the absence of Cernunnos and remains unexplained. However, in the presence of Cernunnos, the 3′ overhangs were retained with equal probability.
Cernunnos promotes the retention of overhanging ends. We incubated DNA substrates with Ku, DNA-PKcs, and XL with or without the addition of Cernunnos. The DNA substrates had mismatched 3′–3′ ends (KpnI–SacI), blunt-3′ ends with 4-nt overhangs (EcoRV–KpnI), or blunt-3′ ends with 2-nt overhangs (EcoRV–SacII). We used quantitative PCR to measure the percentage of DNA ends joined and sequenced products from each reaction to characterize the junctions. The key shown at the top indicates how the data are presented for each reaction. The junctions recovered from PCR amplification are depicted to show nucleotide addition (black background) and nucleotide deletion (white background).
We also characterized the joining of blunt ends to 3′ overhangs of 4 nt (EcoRV–KpnI) or 2 nt (EcoRV–SacII). In the absence of Cernunnos, most junctions (20 of 29) lost the 3′ overhang, and only a minority (9 of 29) retained the 3′ overhang (Fig. 3). In the presence of Cernunnos, 62 of 62 sequenced junctions retained the 3′ overhang (Fig. 3). Cernunnos exhibited the same effect when added to Ku and XL in the absence of DNA-PKcs (SI Fig. 8). Thus, the addition of Cernunnos promoted retention of sequences from the 3′ overhangs.
Finally, we characterized the joining of blunt ends to 5′ overhangs (SI Fig. 9). Cernunnos exhibited only a moderate effect in promoting retention of the 5′ overhang for EcoRV–BamHI ends, and no effect on retention of the 5′ overhang for EcoRV–EcoRI ends.
Ku, XL, and Cernunnos Ligate One of the Two Strands from Mismatched DNA Ends.
In an effort to account for the retention or deletion of 3′ overhangs, we examined our purified protein preparations for polymerase or nuclease activities. Polymerase activity was absent because the joining reactions did not include dNTPs and because joining efficiency was unaffected by addition of dNTPs (data not shown). Nuclease activity also was absent because Cernunnos strongly stimulated joining of blunt-3′ ends with retention of 3′ overhanging sequences (Fig. 3 and SI Fig. 8). Furthermore, Cernunnos, Ku, XL, and DNA-PKcs failed to degrade radiolabeled single-strand DNA substrates (SI Fig. 10) and did not delete any nucleotides in 112 junctions from pairs of blunt ends (EcoRV–EcoRV) and cohesive ends (EcoRI–EcoRI and KpnI–KpnI) (SI Fig. 11).
Taken together, our data suggest that Cernunnos and Ku stimulated XL to ligate one of the two strands from mismatched DNA ends. The deletion or retention of 3′ overhangs would depend on which strand was ligated. If Ligase IV catalyzes ligation of only one strand from mismatched ends, the 5′ phosphate group of one DNA substrate should be dispensable for ligation. To test this hypothesis, we removed 5′ phosphates from the DNA substrates with 3′ overhanging KpnI ends or blunt EcoRV ends and incubated the DNA with Ku, XL, and Cernunnos with or without DNA-PKcs (Fig. 4). When we removed the recessed 5′ phosphate on the KpnI end, joining actually increased because of decreased competition from intramolecular circularization of the KpnI substrate. This result demonstrated that the 5′ phosphate on the KpnI end was dispensable for joining. By contrast, when we removed the 5′ phosphate from the blunt EcoRV end, joining decreased to the levels observed when both DNA ends lacked 5′ phosphates. Taken together with the junction sequences (Fig. 3), these data indicated that Cernunnos stimulated the ligation of one strand from each end, joining the overhanging 3′ hydroxyl on the KpnI end to the 5′ phosphate on the blunt EcoRV end.
Ku, XL, and Cernunnos join mismatched ends with the 5′ phosphate from only one end. To determine how Ku, XL, and Cernunnos joined mismatched ends, we treated the EcoRV and KpnI substrates with DNA phosphatase (Antarctic phosphatase New England Biolabs). EcoRV* and KpnI* denote dephosphorylated DNA molecules. We confirmed the successful removal of the 5′ phosphates by incubating each DNA preparation with T4 ligase and were able to demonstrate a loss of >90% of the ligation products by using agarose gel electrophoresis (data not shown). We incubated mismatched DNA substrates (EcoRV–KpnI, EcoRV–KpnI*, EcoRV*–KpnI, or EcoRV*–KpnI*) with Ku, XL (1 nM), and Cernunnos (2 nM), with or without DNA-PKcs amplified the junctions by quantitative PCR and plotted joining efficiency on a linear scale.
We wanted to obtain direct evidence for what we shall refer to as mismatched end (MEnd) DNA ligase activity. To avoid PCR amplification, we designed a gel-based assay with radiolabeled DNA. To unambiguously identify the joined products and improve joining efficiency, we assayed intramolecular joining of linear DNA. Each DNA molecule contained cohesive EcoRI–EcoRI ends, blunt EcoRV–EcoRV ends, or mismatched blunt-3′ EcoRV–KpnI ends (Fig. 5).
Ku, XL, and Cernunnos join mismatched ends by ligation of one strand. (A) Schematic of the gel-based assay for joining. We incubated Ku, XL, and Cernunnos (Ku/XL/C) or T4 ligase (T4) with a linear DNA substrate containing EcoRI–EcoRI, EcoRV–EcoRV, or EcoRV–KpnI ends digested the DNA products with XmnI and used T4 polynucleotide kinase to radiolabel the DNA. Concentrations of XL and Cernunnos were 1 nM and 5 nM, respectively. For EcoRV–EcoRV and EcoRV–KpnI ends, we performed a second digest with ApoI or MspA1I to remove the radiolabel from one DNA strand. To confirm that ligation created a new phosphodiester bond, we digested the products of the EcoRV–EcoRV joining reaction with EcoRV. Radiolabeled DNA strands are red with red asterisks at the 5′ end, and unlabeled DNA strands are black. DNA products were resolved with denaturing gel electrophoresis. (B) Ku, XL, and Cernunnos joined cohesive EcoRI–EcoRI ends. Ku, XL, and Cernunnos (lane 3) and T4 ligase (lane 4) produced a higher molecular weight band of 498 nt. The molecular weight markers consisted of a radiolabeled 50-bp ladder (lane 1). Joining efficiencies for Ku/XL/C and T4 (calculated from the ratio of intensities in the 498-nt band to the 339-nt band) were 39% and 89%, respectively. Sizes of the ligated higher molecular weight bands in B–D appear in blue typeface. (C) Ku, XL, and Cernunnos joined blunt EcoRV–EcoRV ends. Ku, XL, and Cernunnos (lane 3) and T4 ligase (lane 4) produced a higher molecular weight band of 470 nt, with joining efficiencies of 20% and 14%, respectively. ApoI digestion converted the 470-nt band to 413-nt and 57-nt bands and converted the 281-nt band to 224-nt and 57-nt bands (lane 6). MspA1I digestion converted the 470-nt band to 434-nt and 36-nt bands and the 189-nt band to 153-nt and 36-nt bands (lane 8). Intensities of the 413-nt and 434-nt bands were reduced to 50% of the 470-nt band because the second digestion removed the radiolabel from one strand. EcoRV digestion eliminated the 470-nt band (lane 10), demonstrating that the DNA junction contained new phosphodiester bonds. The 153-, 57-, and 36-nt bands are not shown. (D) Ku, XL, and Cernunnos joined only one strand from mismatched EcoRV–KpnI ends. Ku, XL, and Cernunnos (lane 3) produced a higher molecular weight band of 472 nt, with a joining efficiency of 5%. ApoI digestion converted the 472-nt band to 415-nt and 57-nt bands and the 281-nt band to 224-nt and 57-nt bands (lane 5). The intensities of the 472-nt and 415-nt bands were equivalent (lanes 3 and 5). MspA1I digestion converted the 472-nt band to a 36-nt band and converted the 191-nt band to a 36-nt band (lane 7), demonstrating the ligation of only one strand.
We incubated the DNA with Ku, XL, and Cernunnos. To detect the joined products by denaturing gel electrophoresis, we cleaved the DNA products with XmnI and labeled the ends with [γ- 32 P]ATP and T4 polynucleotide kinase (Fig. 5 A). This procedure labeled the 5′ ends of DNA strands terminating at XmnI, EcoRI, and EcoRV sites. We measured joining by appearance of a higher molecular weight DNA fragment of the appropriate size. To determine whether joining occurred for a specific strand, we removed the radiolabel on one strand or the other by cleaving the DNA with ApoI or MspA1I.
For the cohesive 5′ overhanging EcoRI–EcoRI ends (Fig. 5 B), Ku, XL, and Cernunnos joined 39% of the input DNA. By contrast, T4 ligase joined 89% of the input DNA. For blunt EcoRV–EcoRV ends (Fig. 5 C), Ku, XL, and Cernunnos joined 20%, and T4 ligase joined 14% of the input DNA. When we removed the radiolabel from one strand by cleaving the DNA with ApoI or MspA1I, 50% of the signal from the higher molecular weight DNA disappeared (Fig. 5 C). Thus, the assay was able to discriminate the joining of one strand vs. the other. The junction formed by joining of the blunt EcoRV ends was susceptible to cleavage by the phosphodiesterase activity of EcoRV, indicating that Cernunnos stimulated a ligation event that created a bona fide phosphodiester bond.
For blunt-3′ EcoRV–KpnI ends, Ku, XL, and Cernunnos joined 5% of the input DNA (Fig. 5 D), which was higher than the result from quantitative PCR of the same DNA products, which detected ligation of 1.6% of the input DNA (data not shown). When we removed the radiolabel from the strand containing the 3′ overhang by MspA1I cleavage, 100% of the signal from the higher molecular weight DNA disappeared. By contrast, removal of the radiolabel from the other strand by ApoI reduced the size of the higher molecular weight fragment by 57 nt without diminishing the signal. This result provided direct evidence for single-strand ligation of the hydroxyl group of the 3′ overhang to the 5′ phosphate of the blunt end. Remarkably, Ku, XL, and Cernunnos ligated blunt-3′ ends with an efficiency of 5%, even in the absence of DNA-PKcs. For comparison, T4 ligase ligated blunt ends with an efficiency of 14%.
Targeted nucleases are powerful tools for mediating genome alteration with high precision. The RNA-guided Cas9 nuclease from the microbial clustered regularly interspaced short palindromic repeats (CRISPR) adaptive immune system can be used to facilitate efficient genome engineering in eukaryotic cells by simply specifying a 20-nt targeting sequence within its guide RNA. Here we describe a set of tools for Cas9-mediated genome editing via nonhomologous end joining (NHEJ) or homology-directed repair (HDR) in mammalian cells, as well as generation of modified cell lines for downstream functional studies. to minimize off-target cleavage, we further describe a double-nicking strategy using the Cas9 nickase mutant with paired guide RNAs. This protocol provides experimentally derived guidelines for the selection of target sites, evaluation of cleavage efficiency and analysis of off-target activity. Beginning with target design, gene modifications can be achieved within as little as 1𠄲 weeks, and modified clonal cell lines can be derived within 2𠄳 weeks.
Funding for this work was provided by the National Institutes of Health (R01GM121698 to D.A.B, R21AG056706 to D.A.B, R01GM106081 to X.W.) and the Arizona Biomedical Research Commission (ADHS16-162401 to D.A.B). N.B. was supported by a fellowship from the International Foundation for Ethical Research.
Nicholas Brookhouser, Toan Nguyen, Stefan J. Tekel and Kylie Standage-Beier contributed equally to this work.
School of Biological and Health Systems Engineering, Arizona State University, 501 E. Tyler Mall, ECG 334A, Tempe, AZ, 85287, USA
Nicholas Brookhouser, Toan Nguyen, Stefan J. Tekel, Kylie Standage-Beier, Xiao Wang & David A. Brafman
Graduate Program in Clinical Translational Sciences, University of Arizona College of Medicine-Phoenix, Phoenix, AZ, 85004, USA
Molecular and Cellular Biology Graduate Program, Arizona State University, Tempe, AZ, 85287, USA