| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
J. Biol. Chem., Vol. 278, Issue 36, 33848-33856, September 5, 2003
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
¶ ||
From the
Department of Molecular & Medical
Genetics, University of Toronto, Ontario M5A 1X8, Canada and the
¶Program of Genetics & Genomic Biology, The
Hospital for Sick Children, Toronto, Ontario M5G 1X8, Canada
Received for publication, April 23, 2003 , and in revised form, June 9, 2003.
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
34 can
be genetically unstable, and expansions can be as large as 3000 repeats. In
affected families there is a common tendency for expansion of the disease
allele, although there have been a few DM1 families reported that
preferentially transmit repeat contractions to subsequent generations
(2–5).
Dramatic differences in CTG length variations have been observed in several
regions of the brain in DM1, HD, and DRPLA patients when compared with blood
of the same patient (1). While
many regions of the cerebral cortex show heterogeneous expansions larger than
those observed in blood of the same patient
(6), the cerebellar cortex
consistently shows remarkably shorter lengths in DM1
(7,
8), DRPLA
(9), HD, SBMA, SCA1, and SCA3
(1) patients, suggestive of
large somatic contractions in this tissue.
The precise mechanism(s) responsible for triplet repeat instability remains to be elucidated. Expansions are dynamic, in that longer repeats tracts are more likely to incur further expansion mutations (1). The process of instability is likely to involve DNA slipped/secondary structures (out-of-register interstrand mispairings at repeat sequences). One hypothesis predicts that expansion events arise from errors at DNA replication forks (10, 11). We have shown that cis-elements such as the location of replication initiation and fork direction, relative to a repeat tract, can lead to different types and frequencies of repeat instability (10). Depending upon the location of replication initiation, (CTG)n·(CAG)n repeat tracts incurred expansions or deletions, or were stably replicated. Repeat instability may be linked to the location of the repeat tract within the single-stranded Okazaki initiation zone of the replication fork. It is possible that the replication fork may become blocked as it passes through the triplet repeat tract (12–14). This blockage may be caused by slipped structures, formed either between the nascent and template strands or between the un-replicated template strands. Elimination of the blockage and subsequent resumption of fork progression may require a double-strand break-mediated recovery mechanism (15–17), which can lead to expansions and/or deletions (18). A second hypothesis is that recombination between repeat tracts containing slipped intermediates can yield repeat expansions or deletions as a result of unequal genetic exchange (19, 20).
While these and other theories remain plausible, it is apparent that
several models involve the presence of a double-strand break (DSB). Mammalian
cells are constantly threatened by the occurrence of endogenously or
exogenously created DSBs. In human cells, it has been estimated that DSBs
arising through the DNA replication process occur at a frequency of 
10
per cell cycle, and proper repair is essential for genomic stability and
cellular survival (15). Work
in bacteria has shown that gene conversion can influence the expansion of
CTG·CAG repeats (21).
Experiments using yeast have indicated that triplet repeat tracts are
"fragile," such that they are susceptible to DSBs, possibly
through endonucleolytic cleavage of secondary structures
(22,
23). The formation of DSBs
within or proximal to the repeats is length-dependent. It has been suggested
that subsequent repair of the break can lead to instability of the repeat
tract, in some cases yielding large deletions of the repeat and flanking
sequences
(22–24).
The use of a donor sequence containing (CAG)98 to repair a DSB lead
to integration and expansion of the triplet repeat tract
(25). However, in that study,
the initial DSB substrate did not harbor any (CTG)·(CAG) repeats. Thus,
what remains to be addressed is the influence and fidelity of DSB repair
within a repeat tract upon tract stability. It is also important to determine
the effects of both repeat tract length and the presence of alternative DNA
structures on the fidelity of DSB repair. Furthermore, it is of interest to
utilize primate cells, as yeast and mammalian cells are known to
preferentially use homologous recombination and non-homologous end joining
(NHEJ), respectively (26). The
differences between these two DSB repair mechanisms may therefore result in
distinct repair products.
In this study, we have used an established primate DSB repair system (27) to investigate the fidelity of repair of a DSB within a (CTG)·(CAG) repeat tract. Following repair of the DSB substrates in primate cells, products were characterized for sequence alterations within the repeat tract length and the flanking sequence. The presence of a DSB within the repeat tract lead to several forms of deletion events, removing either exclusively repeats, non-repetitive flanking sequences, or both. The repeat-containing end of the DSB was preferentially deleted, relative to the non-repetitive end. The presence of intrastrand slipped structures within the repeat tract enhanced this "polarized" deletion effect.
|
EXPERIMENTAL PROCEDURES |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
|
Large scale plasmid preparations were prepared from dam+ Escherichia coli cells as previously described (19). Briefly, cells were harvested and lysed with lysozyme (Invitrogen) and a detergent solution of 1% Brij 58 (Sigma) and 0.4% deoxycholate (Sigma). Plasmids were treated with RNase A and T1 (Sigma), phenol-extracted, and purified twice by cesium chloride/ethidium bromide centrifugation.
Template Linearization—The linearization of (CTG)n-containing plasmids was performed by BsmI restriction digestion (New England Biolabs). The completion of linearization was verified through visualization on a 1% agarose gel, where only a single band (linearized DNA) was observed. Linearized templates were also transformed into E. coli cells and yielded no colonies, confirming the poor ability of E. coli to rescue (re-circularize) linear plasmids (28).
Heteroduplexing/Reduplexing—The induction of slipped structures in the repeats was performed as described previously (19, 29–31). For reduplexing (S-DNA), only the pDM79 plasmid was treated (5 µg), and for heteroduplexing (SI-DNA), equal quantities of pDM47 and pDM79 were mixed (2.5 µg each). Briefly, the DNAs were denatured for 5 min in denaturing solution (0.5 M NaOH, 1.5 M NaCl, pH 13), diluted 50-fold in modified 5x TE (50 mM Tris-HCl (pH 8.0), 5 mM EDTA, 0.2755 M NaCl), and renatured at 68 °C for 3 h. The relative amounts of DNA containing slipped structures were determined by densitometric analysis of electrophoretic patterns of radiolabeled repeat-containing DNAs (Fig. 1C).
Cell Culture/Transfections—COS1 cells, which express SV40 T-antigen (T-Ag) were grown in DMEM supplemented with 10% (v/v) fetal bovine serum (FBS). Cells were plated to a confluency of 20% in 100-mm dishes, 1 day prior to transfection. The following day, cells at 40% confluency were incubated according to the manufacturers protocol (Stratagene) with 5 µg of linearized plasmid DNA and 50 µl of Lipotaxi transfection reagent in 4 ml of DMEM lacking FBS. The amount of DNA used per cell was such that any given cell was transfected with only one plasmid. After4hof incubation, 5 ml of DMEM supplemented with 20% FBS was added. The medium was replaced with fresh DMEM (10% FBS) following 24 h of incubation. Episomal DNAs were then extracted 48 h after the addition of DNA, as described (10), with 100 mM Tris-HCl (pH 7.5), 1 mM EDTA, and 0.6% SDS. DNA was purified with multiple phenol/chloroform extractions, ethanol-precipitated, and resuspended in TE (10 mM Tris-HCl (pH 7.6), 1 mM EDTA).
Measuring Efficiency of DSB Repair—Intersubstrate DSB repair efficiencies (Fig. 2) were determined as follows. The linear repeat-containing repair templates pDM79, pDM79-S-DNA, and pDM47x79-SI-DNA (linearized with BsmI) were each mixed with the control SV40-ori plasmid (pJM1), which is void of repeats, and these mixtures were co-transfected into COS1 cells. Following DSB repair and replication, purified products were linearized with XhoI, and unrepaired material eliminated by DpnI digestion. Products were electrophoresed on a 1% agarose gel to resolve the pDM and pJM1 repair products, and Southern blotted using a 32P-labeled SV40-ori probe. The relative amounts of pre-transfection starting mixtures and repaired material were determined by densitometric analysis. The efficiency of DSB repair was determined by the relative differences in the amounts of repeat-containing plasmids compared with control plasmid. The repeat-containing plasmids replicated with equal efficiency to the control pJM1 plasmid.
|
Analysis of Stability/STRIP Assay—Repeat stability
was assessed using the Stability of Trinucleotide
Repeats by Individual Product analysis (STRIP
assay). This assay has been described in detail
(10,
11). Briefly, DSB repair
products in the Hirts lysate were digested with 10 units of DpnI (New
England Biolabs) to eliminate unrepaired and unreplicated parental templates.
DpnI-digested material was transformed into DH5
-mcr E.
coli, and only the DpnI-resistant material gave rise to
colonies. Individual bacterial colonies, each representing an individual
product of primate DSB repair and replication, were picked and cultured.
Primate replication-mediated instability was minimized as the starting
templates are stably replicated
(10). Bacterial contribution
to repeat length alterations was minimized by: 1) having the
(CTG)·(CAG) repeat in the stable orientation relative to the bacterial
ColE1 replication origin, 2) the limited colony growth time (
6 h), and 3)
bacterial strain selection. Through the use of these strict specifications,
255 CTG repeats can be stably maintained in bacteria
(31).
Miniprep DNA was analyzed for repeat instability through analysis of the migration pattern of BamHI/HindIII fragments on high resolution 4% polyacrylamide gels (19). The magnitude of repeat length changes were ascertained by electrophoretic sizing of the repeat-containing fragments relative to starting length material and a size marker. All expansions and samples of each deletion type were confirmed through sequence analysis.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
DSBs were produced by linearizing the pDM plasmids with the class II-S restriction endonuclease BsmI. The unique cleavage site for this enzyme is located in the first CTG unit of the repeat tract (Fig. 1A) as shown in Sequence I, thereby causing the complete repeat tract to comprise one free end of the linearized plasmid (Fig. 1B). In order to be "rescued," there were limitations to the deletion sizes that could be tolerated by the DSB templates without jeopardizing their ability to replicate in either primate or bacterial cells (Fig. 1B).
|
Linearized pDM47 and pDM79 plasmids were also denatured and renatured under conditions that induce the formation of slipped structures within the repeat tract (19, 29). Slipped homoduplexes (S-DNA), containing the same number of repeats on each strand, and slipped intermediate heteroduplexes (SI-DNA), containing a different number of repeats on each strand, have been characterized (19, 29–31). The amount of slipped structures formed was determined by densitometric analysis of the electrophoretic pattern of radiolabeled repeat-containing DNAs (Fig. 1C). S-DNAs alone amounted to 70% of the repeat-containing fragments (middle lane). Together, the amount of sister SI-DNAs, containing an excess of CTG or CAG repeats, represented 50% of the repeat-containing fragments (right lanes). The visualization of each product within the heteroduplex mixture was facilitated by uniquely radiolabeling either the (CTG)47 or the (CTG)79 strands, indicated by an asterisk. These relative amounts are consistent with our previous studies (19, 29–31).
Double-strand Break Repair Efficiency—To determine potential differences in DSB repair efficiency, which may be caused by the presence of (CTG)·(CAG) repeats or slipped structures, the degree of repair of repeat-containing substrates was compared with that yielded by an identical linear plasmid lacking a repeat tract (pJM1). Repeat-containing substrates and the control plasmid were mixed at equimolar concentrations (Fig. 2, "starting material"), co-transfected and repaired/replicated material was isolated 48 h later. DNA from the starting plasmid mixtures was electrophoresed along with the repaired material, and relative concentrations determined through densitometric analysis (Fig. 2). It was found that the pDM79 substrate, with the repeats in their linear, Watson-Crick form, was repaired with 91% percent efficiency relative to the non-repetitive control plasmid (pJM1). The presence of homoduplex SDNA structures decreased the DSB repair efficiency, with the pDM79-S-DNA substrate being repaired at 76% efficiency relative to pJM1, and 83% relative to the linear form of the repeats. The pDM47x79-SI-DNA substrate was repaired at 71% efficiency relative to pJM1, and 78% efficiency relative to the linear form of the repeats. The differences observed are due to variable efficiencies of DSB repair, as the presence of repeats does not affect the efficiency of replication (data not shown). These values suggest that the presence of both the repeat sequence and slipped DNA structures reduced the efficiency of DSB repair.
The STRIP Assay—To address the effect of DSB repair on (CTG)·(CAG) repeat instability, we modified an assay we have developed to determine the role of primate DNA replication on the STRIP assay (10). Following enzyme-induced linearization, repeat-containing plasmids were transfected into COS1 cells, and the DSB-repaired (re-circularized) products were recovered as replicated DNAs using Hirts lysis and DpnI digestion. To reduce contributions to DSB repair by homologous recombination, the transfections were performed such that any cell contained a single plasmid ("Experimental Procedures"). The DpnI-resistant material was transformed into bacterial cells to generate individual colonies, each derived from a single product of DSB repair. Plasmids of each colony were analyzed for repeat length alterations by restriction digestion and visualization on polyacrylamide gels.
Classification of Repeat Instabilities—As a means of determining repeat stability, the plasmids isolated from individual colonies were digested with BamHI and HindIII to release both the repeat tract (BamHI-HindIII) and the SV40-ori (BamHI-BamHI) (Fig. 3A). An assessment of stability was then made through electrophoretic comparison of the BamHI-HindIII and BamHI-BamHI fragments to the starting material. As previously observed, several rescued plasmids contained insertions of either genomic or plasmid DNA (27) and yielded aberrant restriction digestion patterns. The occurrence of these recombinants did not correlate with either repeat tract length or the presence of slipped structures (data not shown). As in previous studies, these recombinants were not considered for our results.
|
In general, rescued recombinant plasmids could be broadly classified as having sustained deletions or expansions of repeats. Yet due to the location of the DSB at the extreme 5'-end of the repeat, deletion events could potentially remove flanking regions as well, and therefore a more elaborate classification system was established (Fig. 3A).
Repeat Expansions—Rescued plasmids that had experienced no sequence alterations were termed "stable" (Fig. 3, A–i) and displayed BamHI-HindIII and BamHI-BamHI restriction fragments, containing the CTG repeat and the SV40-ori, respectively, that co-migrated with the fragments of the starting plasmid (Fig. 3B, see lanes labeled "intact"). Plasmids that incurred repeat expansions (Fig. 3, A–ii) displayed a CTG repeat fragment of slower electrophoretic mobility, and an SV40 fragment identical to the control (Fig. 3B; see lanes labeled expansion). All repeat expansions were confirmed through sequence analysis.
Only low levels of repeat expansions following DSB repair were observed for the linear pDM47 and pDM79 substrates (Fig. 4A), and these levels were not significantly above the length heterogeneity present in the starting bacterially-produced DNA preparations (data not shown). Repair of substrates containing slipped-homoduplex S-DNAs (pDM79-SDNA) produced expansions at a similarly low level. Repair of an SI-DNA heteroduplex substrate (pDM47x79-SI-DNA) did not generate products with repeat tracts greater than 79 repeats (it is not possible to know if repair products containing 48–78 repeats were expansions of 47 or deletions of 79). Notably, the BsmI site was conserved in the majority of cases where an expansion event or no sequence alteration was present; as confirmed through sequencing and digestion (data not shown). We conclude that primate cell repair of DSBs within CTG repeats in either the Watson-Crick or slipped conformation did not yield significant levels of expansions.
|
The Spectrum of Deletions—Following DSB repair, deletion events represented the majority of the repaired products (Fig. 4A). Plasmids that underwent a deletion event within the repeat (termed Class I deletions) (Fig. 3, A–iii) displayed a BamHI-HindIII CTG-repeat fragment of greater electrophoretic mobility, and a BamHI-BamHI SV40 fragment identical to the control (Fig. 3B; see lanes labeled Class I). Also, several plasmids harbored deletions that encompassed more than just the repeat. The nature of these deletions was assessed as follows: rescued plasmids with Class II deletions displayed only the BamHI-BamHI SV40 fragment (Fig. 3B; see lanes labeled Class II), having sustained deletions of the entire repeat tract, as well as the HindIII site on their 3'-end (Fig. 3, A–iv). Another subset of rescued plasmids, termed Class III deletions, showed a single band that varied in size between plasmids, but was consistently larger than the standard BamHI-BamHI SV40 fragment (Fig. 3B; see lanes labeled Class III). These rescued plasmids had sustained deletions that removed the BamHI site nearest the BsmI-induced DSB, and possibly a portion of the repeat tract (Fig. 3, A–v). A sample of each deletion class was confirmed through sequences analysis.
DSB repair induced high levels of deletion products for each of the linear substrates. The pDM17 substrate incurred deletions at a frequency of 43%, of which 19, 9, and 15% were Class I, Class II, and Class III events, respectively (Fig. 4A). The pDM47 substrate incurred deletions at a frequency of 54%, of which 31, 8, and 15% were Class I, Class II, and Class III events, respectively (Fig. 4A). The pDM79 substrate incurred deletions at a frequency of 66%, of which 34, 8, and 24% were Class I, Class II, and Class III events, respectively (Fig. 4A). Class III deletions were increased only for the pDM79 substrate. It is noteworthy that while the frequency of Class II deletions was similar between the different repeat lengths, the amount of DNA lost was greater for the longer repeat tracts. Both Class II and Class III deletion products were specific to DSB repair, as neither of these has ever been observed following bacterial or primate DNA replication (10). However, Class I deletion products, having lost only repeat units, were observed following DSB repair (herein) and bacterial or primate replication (10). As stated above, the plasmids used as DSB repair substrates have been shown previously to be stably replicated in primate cells (10). This is summarized in Fig. 4B, where it can be seen that primate background deletion levels (stippled bars) are equal to bacterial background deletion levels (hatched bars), which means that the deletions observed following primate replication were originally present in the starting plasmid preparation. Likewise, a proportion of the Class I deletions observed in our DSB-repaired material can also be attributed to background length heterogeneity. Consequently, the levels of DSB repair-induced Class I deletions above the background were 19% (19–0%) for pDM17, 11% (31–20%) for pDM47, and were negligible (34–36%) for pDM79 (Fig. 4B). For pDM17, Class I deletions occurred only following DSB repair, but were not detected following either bacterial or primate replication (Fig. 4B), providing further support for the enhanced deletions induced by DSB repair. The apparently reduced levels of Class I events for the longer repeats may be due to enhanced deletions into the repeat tact and the adjacent flanking regions, a phenomenon that may be stimulated by an inherent ability of the longer repeats to "fray" into alternative DNA structures (19, 29).
The presence of homoduplex slipped-strand S-DNAs affected the type of DSB repair-induced deletion products, but not the overall deletion frequency. The pDM79-S-DNA substrates incurred overall deletion events at a frequency of 66%, which is similar to that of the fully paired linear substrate. However, the slipped substrates incurred enhanced deletions into the repeat tract, as is evidenced by the relative increase in Class II deletions and decrease in Class III deletions compared with the fully base-paired linear substrates (Fig. 4A). It therefore appears that homoduplex S-DNAs are more susceptible to deletions into the repeat tract, relative to duplex linear repeats.
DSB repair of substrates containing heteroduplex slippedintermediate SI-DNAs also affected the types of deletion products generated. Repair products of the pDM47x79-SI-DNA substrate also contained high levels of deletions, totaling 64%, and comprising of 10% that were Class II deletions and 17% that were Class III deletions (Fig. 4A). It was not possible to assess the frequency of Class I deletions for the heteroduplexed substrates; however, they occurred at a minimum frequency of 30% (represented by deletion products <47 repeats; see Fig. 4A).
Since the DSB was induced by BsmI digestion, it was of interest to determine if the restriction site was maintained. The status of the BsmI site in the three deletion classes varied (data not shown). For all repeat lengths, in both linear and slipped-structure conformations, Class I deletions showed a relatively even distribution of products that preserved the BsmI site and products that lost the site. The exception was for rescued pDM47x79-SI-DNA heteroduplex substrates that resulted in repeat fragments whose size was between 47 and 79 repeats, where the BsmI site was consistently preserved. Class II deletions showed a similar, seemingly random distribution of BsmI site-preservation frequencies. Finally, the BsmI site was abolished in all examples of Class III deletions, which is not surprising since Class III products have incurred loss of sequence in the non-repeating end that contains the BsmI recognition site.
Preferential Deletion of the Repeat-containing End—Remarkably, it was found that the repeat-containing end of the DSB was preferentially degraded during DSB repair. Moreover, the presence of slipped structures within the repeat tract enhanced this directional deletion effect. DNA ends at a double-strand break are susceptible to exonucleolytic degradation, and subsequent ligation of these ends can result in deletions. Assuming that the rate of exonucleolytic digestion of DNA ends is insensitive to DNA sequence or structure, one would expect both the repeat end and the non-repeat end to be degraded at similar rates. Intuitively, Class II deletion frequencies should decrease as repeat tract length increases, simply due to the greater number of nucleotides between the BsmI DSB and the HindIII site (Fig. 5A). However, Class III deletion frequencies should remain relatively constant, since the number of nucleotides between the BsmI DSB and the proximal BamHI site is the same for all repair substrates. In this manner, the proportion of Class III deletions should increase relative to Class II deletions for substrates with longer repeat tracts, and the Class III/Class II ratio for a given substrate would be directly related to the differential lengths from each end to their respective diagnostic sites.
|
Taking these assumptions into account, we have estimated the expected proportion of Class III to Class II frequencies based on the nucleotide lengths that would be lost for each event. More specifically, the length ratios are the distances from the DSB site to the Class II and Class III diagnostic restriction sites, namely from BsmI to HindIII and BsmI to BamHI, respectively (Fig. 5A, left column). Comparing these expected III/II ratios to the experimentally observed III/II ratios revealed considerably lower values in the latter (Fig. 5B), indicating a marked bias for the deletion of the repeat-containing end of the DSB.
The difference between observed and expected values for the slipped
structure-containing pDM79-S-DNA substrate was statistically significant
(
2-test, p = 0.0025). Additionally, as both the S-DNA
and SI-DNA mixes did not consist entirely of molecules containing slipped
structures (70 and 50%, respectively), the effect of slipped structures on the
observed deletion bias may have been underestimated due to the presence of
repeat tracts in their linear Watson-Crick form. Thus, the presence of slipped
structures within the repeat tract enhanced the preferential deletion of
repeats.
This method of analysis was verified by performing an identical DSB repair experiment using a plasmid (pJM1) similar to the pDM substrates, lacking only the repeat tract (Fig. 5A). Repair products of the linearized control plasmid produced an observed Class III/Class II ratio similar to that which was expected (Fig. 5B). This further illuminates the effect that both repeat sequence and slipped repeat structures have on the outcome of DBS repair.
To address the possibility that repeat tracts of different lengths were
deleted at different rates, we compared the expected over observed Class
III/Class II ratios between different substrates
(Fig. 5A, right
column). As predicted, there was little difference observed for the
control pJM1 plasmid, which lacked repeats (0.85), and a minimal difference
for the pDM17 substrate that contained only 17 repeats (1.31). For both the
pDM47 and pDM79 substrates, which contained longer repeat tracts, the
difference was considerable (2.06 and 2.16, respectively). The pDM79-S-DNA
substrate, which contained slipped structures, showed the greatest difference
(4.71). Thus, the rate of polarized deletion was 2-fold greater for
tracts that contained greater than 17 repeats, while the presence of slipped
structures led to a nearly 5-fold enhanced deletion rate. Thus, not only are
repeat-containing ends more susceptible to deletions than non-repeat ends, but
longer repeat tracts are deleted more frequently or rapidly than shorter
tracts. To our knowledge, this is the first observation of a polarized
deletion bias for any sequence or DNA structure.
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
The improper repair of a DSB located within a triplet repeat tract resulted most commonly in deletions. As tract length increased above the unstable threshold length of 34, the occurrence of deletions into the repeat end increased. This is the first time this phenomenon has been observed for a DSB located within a repeat tract in primate cells. Similar double-strand break repair assays that have not concerned repeating sequences have also found deletions to be the most common result of improper DSB repair (27, 32). It has been proposed that this bias toward deletions is likely a result of fraying/unwinding at the two free ends of the double-strand break, thereby becoming suitable substrates for repair/recombination enzymes (33). For the repeat-containing substrates used herein, the preferential deletion of the repeat end in duplex molecules may be due to the increased ability of this end to fray and form substrates for nucleases that would enhance deletions. Free ends with different base compositions are known to vary in their degree of end fraying (breathing) (34), which may affect their ability to form alternative structures, as in the terminal repeats of the linear adenovirus (35). It is possible that CTG·CAG repeats at free ends are inherently prone to breathing and structure formation, with subsequent enzymatic processing possibly leading to deletions (Fig. 6). This model is supported by the enhanced deletion of DSB substrates containing slipped structures.
|
Our results propose a role for sequence- or structure-specific protein(s) which preferentially remove triplet repeat-induced slipped structures located at a DSB as an early step in the repair process (Fig. 6). We have recently demonstrated that both S-DNA and SI-DNA structures can be specifically recognized and cleaved by both junction- and single strand-specific nucleases (36). Mammalian cells are believed to use non-homologous end joining (NHEJ) as a primary means of DSB repair (26), making proteins such as Ku70, Ku80, DNA-PKcs, DNA ligase IV, XRCC4, and Artemis likely candidates. Similarly, the human Mre11-Rad50-NBS1 complex, which is a part of the homologous recombination process, has been directly implicated in the scission and/or unwinding of DNA hairpins (37). Other junction- or structure-specific endonucleases (38) may act upon slipped structures. The structure-specific WRN DNA helicase may participate in the repeat instability observed (39). Many of these proteins are involved in the maintenance of telomeric repeats through telomere-specific DNA structures (40). Thus, triplet repeats situated at a DSB end may not be protected from end degradation, as are telomeric repeats at chromosome ends (41). While it is not known which specific proteins participate in the processing of slipped repeat structures, we have clearly shown that when present at a DSB, their processing results in deletions.
Repeat tract length changes may result from a combination of replication fork blockage and DSB repair (12–14, 16, 18). The re-establishment of stalled fork progression in mammalian cells may involve the induction of DSBs and subsequent break repair (15, 17, 18). Previously, we reported template-specific instability of (CTG)·(CAG) tracts following episomal replication in primate cells (10). Depending upon replication direction and the location of the repeat tract relative to replication initiation, we observed either a bias for deletions, a bias for expansions or stable replication. Replication fork blockage at the repeats and DSB-induced re-establishment of fork progression may contribute to the type of mutations incurred by specific template configurations. The results presented here suggest that DSBs within triplet repeats increase the frequency of repeat deletions. This would suggest that the circular replication templates pDM79AP, pDM79HD, and pDM79E that show a predominance of deletions (10) may be more commonly subject to fork blockage within the repeat tract, and that DSB induction and aberrant repair lead to preferential repeat contractions.
The DSB in our substrates may share some, but not all, structural features present in DSBs associated with stalled replication forks, and as such may not be subject to the same repair/recombination processes. Such differences may explain the paucity of expansion events observed in our DSB repair assay. Conversely, the DSB present in our substrates may more closely reflect breaks that occur spontaneously throughout the life of a cell (22). The deletions incurred by genome maintenance-associated repair may reflect the age-dependent length heterogeneity observed in non-proliferative cells of mice and patients harboring expanded trinucleotide repeats (42–46). Our results suggest that the natural or environmental induction of DSBs within a (CTG)·(CAG) tract might lead to repeat deletions.
Large repeat deletions have been detected in the cerebellar cortex of DM1 (7, 8), DRPLA (9), HD, SBMA, SCA1 and SCA3 (1) patients. Furthermore, a few DM1 families have been shown to preferentially pass on repeat contractions of the expanded repeat to subsequent generations (2–5). It is unknown how and when these deletions arise. Perhaps these somatic or intergenerational contractions are the result of faulty DSB repair specific to these families/tissues. Further studies establishing the natural causes of DSBs in human triplet repeat tracts, and their repair by human proteins, will further our understanding of the manifestation and treatment of trinucleotide repeat-associated diseases.
|
FOOTNOTES |
|---|
Supported by the Hospital for Sick Children Restracomp Award and the
Ontario Graduate Scholarship.
|| A CIHR Scholar and a Canadian Genetic Disease Network Scholar. To whom correspondence should be addressed. Tel.: 416-813-8256; Fax: 416-813-4931; E-mail: cepearson{at}genet.sickkids.on.ca.
1 The abbreviations used are: DM1, myotonic dystrophy type 1; DSB,
double-strand break; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal
bovine serum; nt, nucleotides; STRIP assay, Stability of Trinucleotide Repeats
by Individual Product analysis.
|
ACKNOWLEDGMENTS |
|---|
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati What's this?
This article has been cited by other articles:
|
|
 |
|
  L. M. Pollard, R. L. Bourn, and S. I. Bidichandani Repair of DNA double-strand breaks within the (GAA*TTC)n sequence results in frequent deletion of the triplet-repeat sequence Nucleic Acids Res., February 2, 2008; 36(2): 489 - 500. [Abstract] [Full Text] |
