Figures
Abstract
Background
The trinucleotide repeats AAT•ATT are simple DNA sequences that potentially form different types of non-B DNA secondary structures and cause genomic instabilities in vivo.
Methodology and Principal Findings
The molecular mechanism underlying the maintenance of a 24-triplet AAT•ATT repeat was examined in E.coli by cloning the repeats into the EcoRI site in plasmid pUC18 and into the attB site on the E.coli genome. Either the AAT or the ATT strand acted as lagging strand template in a replication fork. Propagations of the repeats in either orientation on plasmids did not affect colony morphology when triplet repeat transcription using the lacZ promoter was repressed either by supplementing LacIQ in trans or by adding glucose into the medium. In contrast, transparent colonies were formed by inducing transcription of the repeats, suggesting that transcription of AAT•ATT repeats was toxic to cell growth. Meanwhile, significant IS1E transposition events were observed both into the triplet repeats region proximal to the promoter side, the promoter region of the lacZ gene, and into the AAT•ATT region itself. Transposition reversed the transparent colony phenotype back into healthy, convex colonies. In contrast, transcription of an 8-triplet AAT•ATT repeat in either orientation on plasmids did not produce significant changes in cell morphology and did not promote IS1E transposition events. We further found that a role of IS1E transposition into plasmids was to inhibit transcription through the repeats, which was influenced by the presence of the H-NS protein, but not of its paralogue StpA.
Citation: Pan X, Liao Y, Liu Y, Chang P, Liao L, Yang L, et al. (2010) Transcription of AAT•ATT Triplet Repeats in Escherichia coli Is Silenced by H-NS and IS1E Transposition. PLoS ONE 5(12): e14271. https://doi.org/10.1371/journal.pone.0014271
Editor: David T. Kirkpatrick, University of Minnesota, United States of America
Received: June 23, 2010; Accepted: November 15, 2010; Published: December 9, 2010
Copyright: © 2010 Pan et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by grants from the Natural Science Foundation of the Beijing Institute of Technology (1060050320804), China, and the Medical Science foundation of Hebei University(2007Z02), China. And it has also been supported by a grant from Medical Research Council of the United Kingdom. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
AAT•TTA triplet repeats are types of trinucleotide repeats that are highly cumulative in human genome [1], [2], in contrast, they are much less abundant in many prokaryotic genomes, such as in E.coli, showing a biased distribution towards eukaryotic genomes [3], [4], [5], [6], [7]. Further, AAT•TTA repeats are also found to be over-represented in intronic and intergenic regions, but underrepresented in exons and UTRs, showing a nonrandom nature of distribution in an individual genome [1], [2].
AAT•TTA repeats have been characterized in vitro to form various types of non-B DNA secondary structures, including hairpin, triplex, non-H DNA [8], [9], [10], [11], which potentiallychallenge the stable maintenance of the repeats in genomes [8].
Consisting with this, it has been found in prokaryotes that certain proteins known as chromosome structural proteins bind preferentially to AT-rich DNA that is normally seen in promoter region, and which may make the DNA segments inactive to avoid forming non-B secondary structure during DNA transcription [12]. H-NS and its paralogue StpA are two of the main chromosome structural proteins in E.coli, which bind AT-rich DNA sequences with overlapped specificity; H-NS binds to more than 1000 genes when repressing the transcription of the target DNA region [12].
The non-B structures formed by AAT•TTA triplet repeats were found to be similar to those of non-B secondary structures formed by disease causing trinucleotide repeats such as CAG·CTG, CGG·CCG and GAA·TTC, which are associated with more than 40 human genetic diseases, including Huntington's disease or fragile X syndrome [8], [13]. Consistent with this, a typical expansion feature of AAT•TTA triplet repeats was found in an in vitro amplification assay [9], [10], [13], [14], [15], [16], [17], [18]. Moreover, in human populations similar instabilities of AAT•TTA triplet repeats were recently found to be associated with the high IgE blood syndrome in Chinese children [19], with schizophrenia [20], [21], cocaine addiction [22], and with the high prevalence of depression in adult Parkinson's disease patients [23]. More strikingly, a similar triplet repeat expansion was also found to be associated with the propagation of a 33-bp AT-rich repeat, which displayed chromosomal fragilities in humans, such as FRA3B, FRA16B, and FRA10B [24], [25].
However, the maintenance and biased distribution of AAT•TTA repeats in different chromosomal regions in a genome or in different genomes are still poorly understood. To gain some understandings of the maintenance of AAT•TTA triplet repeats in vivo and its biased distribution within a genome or in genomes, we performed experiments using a 24- triplet AAT•TTA repeats in the E.coli model system. We found that propagation of the repeats on plasmids was overall normal, but that transcription of the repeats in either orientation with either AAT or ATT serving as transcribing template induced significant changes in colony morphology, leading to formation of convex colonies from normal colonies, which then progressively transformed into transparent colonies. Interestingly, we found that IS1E transposition from the chromosome to either the proximal site of the promoter of lacZ in the plasmids or into the AAT•TTA triplet repeats themselves were capable of reversing colony changes by repressing transcription of the repeats. And we further found that the AT-rich repeats binding protein H-NS, a chromosomal structuring protein, but not its paralogoue StpA, played dual roles in triplet repeats binding and in facilitating IS1E transposition into the repeats containing plasmid, which ultimately prevented transcription of the triplet repeats.
Results
Transcription of a 24-triplet AAT•TTA repeat on a plasmid induces morphological changes of bacterial colonies from convex to transparent
Plasmids carrying a 24-triplet AAT•TTA repeat with either the AAT or the ATT strand as lagging strand template for DNA replication were initially constructed as described in the materials and methods. They were named as pAAT24 and pTTA24, respectively. Plasmid pAAT8 was a deletion product occasionally obtained when propagating pAAT24 in a recA mutant, and the pTTA8 plasmid was obtained by reversing the AAT8 into the opposite orientation.
When the pAAT24 and pTTA24 plasmids were propagated in the E.coli JM83 wildtype strain, normal convex colonies on LB plates containing ampicillin were formed, but the colonies started to become flat when plates were left at 4°C overnight (Figure 1). Almost all flat colonies became concave, and then progressively turned into transparent colonies. In contrast, a similar phenomenon was not observed with JM83 cells when they propagated pAAT8, pTTA8, and pUC18 plasmids growing under the same conditions (data not shown). This indicated that only colonies carrying longer AAT•TTA triplet repeats suffered morphological changes when grown overnight.
A) Organization of the AAT•ATT repeats in the lacZ gene in pUC18 plasmid and in the attB site of the genome; B) Papillation assays when the AAT•ATT repeats were subcloned in plasmids. (1) JM83 (pAAT24); (2) JM83Δhns (pAAT24); (3) JM83 (pATT24); (4) JM83Δhns (pATT24), which behaves similarly when a lacIQ plasmid was coexpressed, or 5% glucose was added in the LB plate (For clarity the data were not shown); (5) JM83 ΔstpA::cat (pAAT24); (6) JM83ΔstpA::cat(pAAT24) with 5% glucose; (7) JM83ΔstpA::cat(pATT24); (8) JM83ΔstpA::cat (pATT24) with 5% glucose; (9) W3110ΔstpA::cat (pAAT24); (10) W3110ΔstpA::cat(pAAT24) with 5% glucose; (11) W3110ΔstpA::cat(pATT24); (12) W3110ΔstpA::cat(pATT24) with 5% glucose; (13) AB1157 recF (pAAT24); (14) AB1157 recF (pAAT24) with 5% glucose; (15) AB1157 recF (pATT24); and (16) AB1157 recF (pATT24) with 5% glucose.
Several processes may be responsible for the observed alterations in colony morphology, including triplet repeats transcription from the lacZ promoter (Figure 1A) [26], DNA double strand breaks caused by formation of non-B secondary structures, such as non-H structure, which may be targeted by certain DNA structure specific nucleases [27], [28], or activation of the cryptic phageΦ 80 in the JM83 chromosome, leading to cell lysis.
To test these possibilities, plasmids pAAT24 and pTTA24 were also transformed into JM83 mutants defective in homologous recombination, such as ΔrecA, ΔrecB, recF::Tn10 KanR, and ΔruvABC. We wanted to test if the colony morphology changes may be hinted at AAT•TTA triplet repeats breaks in plasmids, which requires homologous recombination repair, and which relies on the rec gene functions. It was observed that DNA double strand breaks in E.coli genome or plasmid affected cell viability and also the colony morphology, leading to plasmid loss or cell death.
However, we found that compared to wild type cells, propagations of pAAT24 and pTTA24 plasmids in all rec mutants tested did not significantly enhance morphological changes (data not shown). Therefore, we excluded double strand break formation as the dominant cause for the observed changes in colony morphology.
To test whether the genetic background of the E. coli host made a difference, the growth and colony morphology of AB1157recF::Tn10 and W3110ΔstpA::cat cells containing plasmids pAAT24 and pTTA24 were compared to that of JM83. Although their genetic backgrounds were significantly different from JM83 and no similar cryptic phages were found, AB1157recF::Tn10 and W3110ΔstpA::cat colonies suffered the same morphological changes as JM83 cells (Figure1 B). These suggested that the observed morphological changes as associated with AAT•TTA triplet repeats only depended on the presence of the plasmids, but not on the genetic background or cryptic phage of the host. Therefore, all subsequent work was done using the JM83 wild type and its derivative strains.
As shown in Figure 1A, AAT•ATT triplet repeats were subcloned at the EcoRI site of pUC18, which is located downstream from the lacZ promoter (Figure 1A). Therefore, we wondered whether transcription of the AAT•ATT triplet repeats from the lacZ promoter occurred in the stationary phase, thus causing directly or indirectly colony morphology changes.
To test whether transcription was responsible for the observed changes in colony morphology, we constructed JM83Δhns and JM83ΔstpA::cat mutants defective in chromosomal structuring proteins H-NS and StpA, respectively. We therefore correlated the possible effects of transcription of AAT•ATT triplet repeats in pAAT24 and pTTA24 with morphological changes in the two strains by supplementing with IPTG, or by co-expressing LacIQ from a plasmid. Interestingly, compared to JM83 and its rec derivatives, propagation of pAAT24 and pTTA24 in the JM83Δhns mutant did not provoke changes in colony morphologies, but propagation in JM83ΔstpA::cat did, indicating that H-NS, but not StpA, was required for this process (Figure 1B).
H-NS binds to AT-rich DNA, which may have an effect on AAT•TTA triplet repeats transcription and somehow stimulate morphological changes of the colonies. To test additional effects, pAAT24 and pTTA24 were transformed into the ΔhnsrecB double mutant. No significant differences in morphological changes between the single Δhns and the ΔhnsrecB double mutant were observed (data not shown). Therefore, transcription of AAT•TTA triplet repeats did not cause double strand breaks in the presence or absence of the H-NS protein.
Transcription of AAT•ATT triplet repeats stimulates transposition of IS1E from the chromosome to plasmids
During prolonged incubation of morphologically changed colonies, secondary colonies can be slowly regenerated from the transparent cells containing pAAT24 and pTTA24 plasmids that eventually form healthy colonies in situ (Figure 1). The plasmid DNA was isolated from these regenerated colonies, and analyzed for variations on an agarose gel (Figure 2A). Plasmids recovered from the regenerated colonies were bigger in size, running between the monomeric and the dimeric plasmids (Figure 2A). Restriction enzyme digestions showed that extra PstI cutting sites were introduced to the repeats, because pUC18 has only one PstI restriction site. We sequenced the recovered plasmids and performed blast searches. This analysis indicated that the additional PstI cutting sites were the result of an IS1E insertion into the plasmid.
A) a. Transposition of IS1E elements into plasmids, plasmid isolated from the healthy colonies, and b. Restriction digestion of the plasmid DNA with transposons. Lanes 1, molecular weight (pBR322/BstN1); Lane 2 and 3, pAAT24 and pATT24 without IS1E transposition, digested by PstI; Lane 4, pAAT24, with IS1E transposition, digested by PstI; Lane 5, pAAT 24 with IS1E transposition, digested by EcoR1;Lane 6 and 7, pAAT24 with IS1E transposition, digested by PstI and EcoRI respectively; Lane 8 and 9 pAAT24 with IS1E transposition, digested by using PvuII. B) Mapping the IS1E inserting elements in the promoter and the AAT repeats region by DNA sequencing, the positions of the mostly recognized sites were shown.
The IS1E insertions were often on the 5′ side of the flanking sequence of the AAT•ATT triplet repeats, that is, within the promoter region of the lacZ gene (Figure 2B). For example, IS1E insertions occurred with a higher frequency into the “GGAATTGTG” site, where the first adenine base was the starting site for LacZ mRNA transcription. Another high frequency site was “GATTACGAA”, containing the “GAA” of the EcoRI restriction site 5′ to the repeats (Figure 2B). These findings suggested that transcription of the AAT•ATT triplet repeats was responsible for the negative effects on cell growth.
AAT•ATT triplet repeats serve as targets for IS1E transposition in repeats transcription
Besides recovering IS1E insertions upstream of the AAT•ATT repeats, we also isolated plasmids containing IS1E insertions in the AAT•ATT repeating array itself (Figure 2B and Figure 3). EcoRI fragments of ∼1 kb in size containing at least one IS1E element within the AAT•ATT repeats were recovered from both pAAT24 and pTTA24 plasmids (Figure 3A). These findings are of particular interest, as this is the first demonstration of AAT•ATT repeats acting as potential hotspots for transposition in vivo. In this case, we have further found that the insertion of IS1E into AAT•ATT repeats rely on the repeats orientations in transcription. IS1E appeared to favour inserting itself into the AAT•ATT repeats with its unique PstI restriction site distal to the PstI restriction site in the vector when AAT orientation is transcribed, generating larger DNA fragment after PstI digestion (Figure 3A lane 1 and 3, and 3B), while insertion of IS1E in the repeats array will use the opposite direction if the ATT orientation is transcribed, generating small DNA fragment by PstI digestion (Figure 3A lane 5 and 7, and 3B). These suggested that ISIE transposition into AAT•ATT repeats occurred in repeats transcription. Transcription of AAT repeats and ATT repeats may produce distinct repeats DNA conformations that may differently be recognized by H-NS, and therefore differently affect the IS1E transpositions. Based upon the fact that IS1E transposition into AAT•ATT repeats expands the repeats by 3 triplets of either AAT or ATT repeats (9 nts) at both the 5′ and 3′ ends flanking the IS1E insertion, which may imply a potential way of making repeats expansion by certain transposons in trinucleotide repeats [13], [29], [30], [31], [32].
A) Plasmids of AAT orientation propagated in LB medium were recovered and digested by Pst I (lane 1), EcoR I (lane 2); and propagated in LB medium in the presence of IPTG, and were digested by PstI (lane 3), and EcoRI (lane 4); Plasmids of ATT orientation propagated in LB medium with or without IPTG, digested by Pst I (lane 5), EcoR I (lane 6); Pst I (lane 7 with IPTG induction) and EcoR I (lane 8 with IPTG induction). M1 and M2 are DNA molecular weights; B) Schematic illustration of the directions of IS1E transposition into the AAT•ATT repeats in light of the transcription of either AAT or ATT orientation.
Transcription of AAT•ATT triplet repeats is inactivated by IS1E transposition
To understand the biological significance of the IS1E transposition during the transcription of AAT•ATT triplet repeats, we have further analyzed the effect of IS1E transposition on lacZ transcription in the plasmids. To this end, we eliminated the AAT•ATT triplet repeats in pAAT24IS1E-A using EcoRI digestion (as marked in Figure 2B), and subsequently re-ligated into the repeats-free pAAT24IS1E-A vector at the EcoRI restriction site.
As determined by DNA sequencing, insertion of an IS1E into pAAT24IS1E-A did not alter the lacZ gene promoter significantly (Figure 2), however, the IS1E element could potentially use its promoter to transcribe both the transposase gene (encoded by it) and the AAT•ATT triplet repeats downstream the IS1E. We conducted a conventional α-complementation analysis by using AAT•ATT triplet repeats free pAAT24IS1E-A and a positive control plasmid, pUC18 by transforming these plasmids into JM83 cells. The transformants were grown on LB plates containing ampicillin, IPTG, and X-gal. JM83 cells carrying the repeats free pAAT24IS1E-A plasmid formed white colonies while the pUC18 plasmid formed blue colonies (Figure 4). Therefore, transposition of IS1E into the upstream region of the AAT•ATT triplet repeats in the lacZ promoter repressed transcription of the repeats. IS1E elements have two partially overlapping open reading frames, InsA and InsB', which have a relative reading frame of 0 and -1, respectively [29], [30], [31], [32]. InsA protein regulates transcription of the gene producing the IS1E transposease, while InsAB' binding to both the left and right IS1E terminals represses IS1E transposase transcription from a promoter found partly in IRL and simultaneously inhibits transposition [29], [30], [31], [32]. As shown here, such a regulatory mechanism may contribute to the repression of AAT•ATT triplet repeats transcription.
a) α-complementation by an AAT•ATT repeats free pAAT24IS1E-A plasmid, and b) α-complementation by pUC18 plasmid (For detail see text).
This has been further confirmed that repression of AAT•ATT triplet repeats transcription by IS1E insertions helped plasmid stability; we co-propagated a LacIQ producing plasmid with either the pAAT24 or the pTTA24 plasmids. Co-propagation of the LacIQ plasmid and pAAT24 or pTTA24 improved plasmid recovery, which was also consistent with the result of adding 5% glucose to the medium to repress lacZ gene transcription (data not shown). These results further indicated that transcription of AAT•ATT triplet repeats was responsible for the morphological changes of the E.coli cells, and that inactivation of transcription by IS1E insertions into the lacZ promoter or the AAT•ATT triplet repeats proximal to the promoter helped cells recover from the transcriptional stress.
H-NS silences transcription by binding to AAT•ATT triplet repeats and promotes IS1E transposition
H-NS selectively silences bacterial genes associated with pathogenicity, and the gene's responses to environmental insults [33], [34], [35], [36]. H-NS binds preferentially to the AT-rich motifs displaying planar curvature, which is likely to appear in bacterial promoters [33], [36], [37]. In addition, H-NS also influences transposition and recombination [38], [39], [40]. In some transposition processes, strains with hns mutations show a low-level of transposase production. In IS1 transposition, reduction of InsAB' production to barely detectable levels has been reported [38], [39], [40]. Because we observed here that H-NS was required for IS1E transposition (Figure 1), while it was also implicated in the colony morphological changes in conjunction with the transcription of the 24 AAT•ATT triplet repeats, we determined whether the hns gene product itself may have a role in triplet repeats transcription by measuring the effects of H-NS on plasmid copy number variations in the wild type and the Δhns mutant of JM83. The results indicated that transcription of the two orientations of AAT•ATT triplet repeats was differently affected by H-NS. However, the plasmid copy number of pAAT24 was not significantly affected by the presence or absence of H-NS, or by the presence or absence of IPTG induction (Table 1). Only 86% of the plasmid copy number of JM83 was detected in the JM83Δhns mutant when induced by IPTG; 69% of the plasmid copy number of JM83 with pATT24 in the JM83Δhns was detected without IPTG induction; and 61.7% of the plasmid copy number of JM83 was detected in JM83Δhns with IPTG induction (Table 1). To rule out that multiple copies of pAAT24 and pTTA24 mitigated H-NS effects in the JM83 wild type, we integrated a single copy of a 24-triplet AAT•ATT repeat in two orientations into the attB site located on the E.coli JM83 chromosome (see Materials and Methods and Figure 1). Propagations of the repeats carrying strains were monitored by analyzing changes of bacterial cell morphology using confocal microscopy. Filamentous growth of the cells was noted when propagating the AAT•ATT repeats in the chromosome of the JM83Δhns mutant, but not in the JM83 wildtype (Figure 5), nor in JM83Δhns and JM83 strains carrying either the pAAT24 or pATT24 plasmid (data not shown). These suggested that H-NS affected the AAT•ATT triplet repeats on the chromosome in a dosage dependent manner.
Significant filamentous cells were observed in JM83Δhns -AAT and JM83Δhns -ATT,but cannot be seen with JM83 -AAT and JM83 -ATT, nor be seen with JM83(pAAT24), JM83(pATT24), JM83Δhns (pAAT24), JM83Δhns(pATT24) and JM83(pUC18), JM83Δhns (pUC18) (data not shown).
Discussion
In human beings, the expansion and contraction of trinucleotide repeats CAG•CTG, CGG•CCG and GAA•TTC are associated with more than 40 human genetic diseases and cancers, including Fragile X, Huntington's disease, SCA1-12, MD, and Fredericha 's ataxia [8], [13]. Similarly, as one of the most abundant and most polymorphic trinucleotide repeats in the human genome, the AAT•ATT triplet repeats also show instabilities, which have recently implicated in some human health symptoms, for example, high IgE blood symptom in Chinese children [19], schizophrenia [20], [21], cocaine addiction [22], and prevalence of depression in Parkinson's disease patients [23]. However, understanding the transactions of AAT•ATT triplet repeats in vivo has so far been largely elusive. In this work, we found that a 24-triplet AAT•TTA repeat can be propagated stably in E.coli when it was not transcribed. Transcription of the triplet repeats invoked cell toxicity, and therefore had to be silenced by using H-NS or by H-NS facilitated IS1E transposition. In contrast, propagation and transcription of a short AAT•TTA repeat of 8 triplets under otherwise similar conditions did not show cell toxicity, nor H-NS and H-NS facilitated IS1E transposition, suggesting that transcription mediated cell toxicity is dependent on the length of the AAT•TTA triplet repeats, and that similar sized AAT•TTA repeat may also possibly be intrinsically vulnerable for transcription in all eukaryotes and prokaryotes. Consisting with this, we found that AAT•TTA repeats longer than 16 triplets do not exist in the databases of human genomic plus transcript, the mouse genomic plus transcript and the others as set by NCBI. Although we realized that very long AAT•TTA repeats have been found in many species, including human being and Drosophila etc., this may further implicate that transcription acted as selective pressure against long AAT•TTA repeats (>17 triplets) to be distributed in the coding region of genes.
The mechanism underlying the induction of cell toxicity and the cellular morphological changes by transcription of AAT•ATT triplet repeats is complex. In this work, we have ruled out the effects of generation of DNA double strand breaks and the cryptic phage induced cell lysis on colony morphological alterations. While our work directly implicated an effect of transcription of longer AAT•ATT triplet repeats on cellular morphological changes, we reasoned that transcription of longer AAT•ATT triplet repeats may facilitate the repeats to form certain types of non-B DNA secondary structures, such as non-H structure, which recruits binding of histone-like protein such as H-NS etc (Figure 6) [9], causing depletion of nuclear structure associated proteins in the chromosome in cells. The cellular morphological changes could be induced due to the depletion of the histone-like proteins, which may also includes HU and IHF etc in some situations [35], [40], [41], [42]. In support of this idea, it was found that propagation of AAT•TTA triplet repeats containing different triplets formed non-H structure in vivo [9], and also simultaneous depletions of H-NS, HU and IHF in Escherichia coli K-12 are lethal [41], [42]. Cells under the situation of depletion of histone-like proteins decrease the negative superhelicity of their chromosomal DNA, show increased lethality [41], [42]. Similarly, we think that might also happen during the transcription of AAT•ATT triplet repeats in plasmids that may facilitate the repeats to form non-B secondary structures, and that depletes the H-NS and affects the cell growth.
RNA transcription by using lacZ promoter opened the double stranded AAT•ATT repeats that could promote the repeats to form DNA secondary structures that may recruit histone-like proteins, H-NS, to the repeats DNA, causing depletion of this protein in the chromosome, which further resulting in DNA instability and/or cell death. Repression by H-NS deactivated the transcription while also facilitating IS1E transposion, the repeats transcription is therefore further silenced by the transposition, over time, the repeats might be able to be converted into mixed repeats by IS1E and eventually form “Junk DNA”.
This fits with the observation of the involvement of H-NS, but not its paralogue StpA, in causing the transcription mediated problems. The H-NS was found to affect DNA compaction and transcription regulation through physical interactions with AT-rich DNA motifs with curvature [33], [34], [36], [37]. However, H-NS has also been implicated with helping targeting DNA molecules in transposition or cleaving the ends of the transposing elements (Figure 6)[30], [31], [32], [38], [39], [40]. Related to all these processes, our work suggested that H-NS affected IS1E transposition in a DNA sequence dependent manner, promoting distinct repression of AAT and ATT triplet repeats transcription and different transposition rates.
AAT•ATT triplet repeats may behave like some disease causing trinucleotide repeats, showing expansion and contraction instabilities. It was demonstrated in vitro assay that AAT•ATT triplet repeats produced expansion and contraction instabilities during replication [16], [17], [18]. However, similar triplet repeats instabilities were not detected in our in vivo assay. We have examined a large number of isogenic mutants defective in homologous recombination, methyl-directed mismatch repair, and nucleotide excision repair to search for the triplet repeats instabilities. However, under the condition of triplet repeats transcription, we failed to see any significant expansion and contraction instabilities, except for repeats instability caused by IS1E insertions (data not shown). Our work has therefore shown that longer AAT•ATT triplet repeats are vulnerable after transcription in E.coli cells that have therefore developed a set of mechanisms to inhibit its transcription by using both H-NS silencing and H-NS facilitated transposition inactivation, which somehow stabilize the repeats from causing cell morphological alteration and probably also repeats expansion and contraction. While bearing the findings of IS1E transposition into both the promoter region and the repeats array, we are unable to distinguish if the AAT•ATT triplet repeats serve as usual hotspots of transpositions, or does it do so only in the repeats transcription? Our findings of the IS1E transposition depended on the repeats orientation in transcription could suggest that the transposition is linked to the transcription of AAT•ATT triplet repeats. Interestingly, a similar transposition of Tn5 into GAA•TTC repeats during the repeats forming triplex has also recently been demonstrated in vitro [43].
Conclusions
We found that transcriptions of longer AAT•ATT triplet repeats located on plasmid and in the E.coli genome were responsible for the formation of abnormal cell colonies. The bacterial colonies were sick in morphology, which was accompanied by significant IS1E transposition and filamentous cell growth. H-NS was found to be a key protein for these processes; presumably inactivating triplet repeats transcription by differently binding to the repeat arrays and then promoting IS1E transposition (Figure 6). These findings suggest that RNA transcription of AAT•ATT triplet repeats may serve as a selective pressure for biased distribution of AAT•ATT repeats in different chromosomal regions.
Materials and Methods
Bacterial strains
Bacterial strains used in this work were AB1157recF:: Tn10KanR, W3110ΔstpA::cat, JM83 wildtype [44], and JM83 derivatives of ΔrecA, recB, Δhns, and ΔhnsrecB, ΔstpA::cat, respectively. Strains JM83-AAT24 and JM83-ATT24, were constructed by integrating the (AAT•ATT) 24 repeats at the attB site of the chromosome with two orientations, of which either AAT or ATT strand of the (AAT•ATT) 24 served as the template for transcription. Strains JM83Δhns –AAT24 and JM83Δhns –ATT24 were constructed by P1 transduction of the Δhns gene into the chromosome of JM83-AAT24, and JM83-ATT24, respectively. P1 transduction was performed as described in [44].
Plasmids
The plasmids used in this study were pKOV, pUC18 and its AAT•ATT repeats carrying derivatives, pAAT24 and pTTA24 [45], [46]. Plasmid pAAT24 was constructed by cloning the (AAT)24 repeats in the EcoRI site of pUC18 plasmid, which was on the lagging-strand template of the replication fork (a gift from C. Abbott, University of Edinburgh). Inversion of the trinucleotide repeat array of (AAT•ATT)24 to generate plasmid pTTA24 was performed by using EcoRI cleavage, and followed by religation using T4 DNA ligase. The plasmids pAAT24 and pTTA24 were all confirmed by DNA sequencing. Plasmids pAAT24IS1E-A, pAAT24IS1E-B, pAAT24IS1E-C, were plasmids of pAAT24 carrying IS1E elements in the promoter region of the lacZ (pAAT24IS1E-A, pAAT24IS1E-B) or in the AAT repeat array (pAAT24IS1E-C), respectively. Plasmids pTTA8 and pTTA8 were the deletion products of plasmid pAAT24 when propagated in a JM83recA::catR mutant. DNA sequencing primer used in this work is 5′-ATCCACATTGCCCTCCATC-3′, which was synthesized by Huada Co. Ltd (Beijing).
Enzymes, antibiotics and biochemicals
Restriction enzymes EcoRI, PstI, NotI used in this work were products of Promega (Beijing); T4 DNA ligase was purchased from New England Biolabs; Ampicillin was from Boehringer Mannheim. Isopropylthio-β-D-galactoside (IPTG) and 5-bromo-4-chloro-3-indolyl-β-D-galactopyronoside (X-gal) were bought from Sigma Chemical Company, respectively.
Media and bacterial cultivation
Luria-Bertani broth (L-broth) was utilized for the cultivation of bacteria at 30°C and 43°C when constructing the strains of JM83-AAT24, JM83-ATT24 by using pKOV integration [44], [47]; while all other types of cultivations were carried out at 37°C. Ampicillin was applied by a concentration of 100 µg/ml when it was required. Transformation was performed by using a CaCl2 method [47].
Plasmid DNA isolation and agarose gel electrophoresis
Plasmid DNA was prepared using a kit purchased from Qiagen after the propagations of the plasmid carrying strains for a period of time, normally overnight cultivation was applied. Agarose gel electrophoresis was conducted according the reference [47] on 0.8% gels (Flowgen).
Examination for the repeat tract instability
Plasmids population was examined following the method [28]: Briefly, monomeric plasmid DNA was used to transform E. coli strains of interest and plasmid DNA prepared from a population of transformants. Cells from roughly 4 primary transformants were harvested in 5 ml L-broth and 50 µl of this suspension was diluted into 5 ml L-broth and grown for 24 hours. This corresponds to 30 generations of cell growth. Plasmid DNA was isolated and cleaved with EcoRI. The fragments were end-labelled with S35-dATP using DNA polymerase I Klenow fragment and resolved on 8% native polyacrylamide gels. Bands were visualised either using X-ray film or on a Molecular Dynamics phosphor Imager [28].
DNA sequencing and determination of the transposon
DNA cycle-sequencing was performed using a kit purchased from PE Applied Biosystems. And DNA sequence was extracted from DNA sequencing has been used for searching the homology of the inserted DNA against NCBI database by using the Blast search engine on the NCBI website, IS1E transposition was therefore determined based on the Blast search.
Determination of the plasmid copy number
Plasmid copy numbers in the strains of JM83 wildtype and JM83Δhns mutant carrying pAAT24 and pATT24 were determined as the following: strains carrying pAAT24 and pATT24 were initially propagated overnight in LB medium. And such overnight cultures were then made into aliquots by 1 mL into two test tubes of 1.5 mL, respectively. Induction of the repeats transcription was applied to only one sample by using IPTG by a final concentration of 0.4 mmol/L, and the two test tubes derived from the overnight culture were further cultivated for 4 hours at 37°C. Total DNA including both chromosomal DNA and plasmid DNA were prepared by following the method described in the reference [48]: Briefly, the cultures were centrifuged at 10,000 rpm at 4°C for 5 minutes. And the pellets were resuspended by adding 50 uL 10 mmol/L EDTA and 50 uL freshly prepared solution containing 0.2 moL/L, NaOH, and 0.5% sucrose, and were further incubated at 70°C for 5 min. 1.5 uL of 4 Mol/L KCl and 0.5 uL of 0.4% bromophenol blue were added when the mixture was cooled down to the room temperature, and 20 uL was analyzed by running a 0.5% agarose gel. The ratio of the total plasmid DNA and the chromosomal DNA (mainly one bulky band) was obtained by measuring their band area appeared on the agarose gel by using software (Quantity One V.4.6.2); three independent measurements were performed.
Integration of the AAT•TTA repeats into chromosome attB site
Plasmid pKOV was used as an integrative tool for the construction of JM83-AAT24 and JM83-ATT24 [45]. Two DNA primers were designed and utilized as follows: upper strand primer: 5′ GTGTTCAGCGGCCGCTCCGGGCTATGAAATAGAAAAATGAATCCGTTGCCTGCGTTATC3′, and lower strand primer: 5′CAGGATGGCGGCCGCCCATCTGGTATCACTTAAAGGTATTAAAAACCCCACAGATGCG3′, which were synthesized by Shanghai Sangon Co. Ltd. They bear a NotI restriction site (underlined sequence), respectively, and also contain part of attB sequence, and the flanking sequence of lacZ open reading frame of plasmids pAAT24 and pTTA24. Integrative plasmids pKOV-AAT and pKOV-ATT were constructed by inserting the PCR products at the NotI sites.
Strains of JM83-AAT24 and JM83-ATT24 were selected against the following criteria [45]. For using antibiotic, 20 mg/ml of chromamphenicol was utilized; for selection against sacB, sucrose was added into the LB medium to a final concentration of 5%(w/v). PCR amplification using the same DNA primers as abovementioned was further performed for the confirmation of the JM83-AAT24, JM83-ATT24 .
Observation of the cell morphological changes during cultivation
Morphological alterations of the E.coli vegetative cells grown in LB broth were monitored by using a light microscope with oil immersion objective. Cells were stained by safranin before observation [49].
Acknowledgments
We are grateful to Drs D Leach and C. Abbott at the University of Edinburgh for the AAT•ATT repeats and some strains; Prof. B. Michel in CNRS, France, for the AB1157recF strain; Dr. Yasuhiko Sekine at Rikkyo University, Japan, for the Δhns mutant; Prof. Naotake Ogasawara at Nara Institute of Science and Technology, Japan, for the W3110ΔstpA::cat mutant; Prof. D Lilly at Dundee University for the lacIQ expression plasmid; Prof. George M. Church at Harvard Medical School, USA, for the pKOV plasmid; Prof. M Schlappi at Marquette University for manuscript editing and his constructive comments on the work, And all members of the Pan group for their help and discussion. YHL, YML and PC are research students in the laboratory and contributed equally to this work.
Author Contributions
Conceived and designed the experiments: XP. Performed the experiments: XP YL PC YL LL LY. Analyzed the data: XP HL. Contributed reagents/materials/analysis tools: XP HL. Wrote the paper: XP.
References
- 1. Astolfi P, Bellizzi D, Sgaramella V (2003) Frequency and coverage of trinucleotide repeats in eukaryotes. Gene 317: 117–125.
- 2. Subramanian S, Mishra RK, Singh L (2003) Genome-wide analysis of microsatellite repeats in humans: their abundance and density in specific genomic regions. Genome Bio.4: R13.
- 3. Rocha EPC, Matic I, Taddei F (2002) Over-representation of repeats in stress response genes: a strategy to increase versatility under stressful conditions? Nucleic Acid Res 30: 1886–1894.
- 4. Gur-Arie R, Cohen CJ, EitanY , ShelefL , Hallerman EM, et al. (2000) Simple sequence repeats in Escherichia coli: abundance, distribution, composition, and polymorphism. Genome Res 10: 62–71.
- 5. Toth G, Gaspari Z, Jurka J (2000) Microsatellites in different eukaryotic genomes: survey and analysis. Genome Res 10: 967–981.
- 6. Cox R, Mirkin SM (1997) Characteristic enrichment of DNA repeats in different genomes. Proc Natl Acad Sci USA 94: 5237–5242.
- 7. Todd J, Treangen TJ, Abraham A-L, Touchon M, Rocha EPC (2009) Genesis, effects and fates of repeats in prokaryotic genomes. FEMS Microbiol Rev 1–33:
- 8. Everett CM, Wood NW (2004) Trinucleotide repeats and neurodegenerative disease. Brain 127: 2385–2405.
- 9. Ohshima K, Kang S, Larson JE, Wells RD (1996) TTA•TAA triplet repeats in plasmids form a non-H bonded structure. J Biol Chem 271: 16784–16791.
- 10. Trotta E, DelGrosso N, Ebra M, Paci M (2000) the ATT strand of AAT/ATT trinucleotide repeats adopts stable hairpin structures induced by minor groove binding ligands. Biochemistry 39: 6799–6808.
- 11. Dayn A, Samadashwily GM, Mirkin SM (1992) Intramolecular DNA triplexes: Unusual sequence requirements and influence on DNA polymerization. Proc Natl Acad Sci USA 89: 11406–11410.
- 12. Oshima T, Ishikawa S, Kurakawa K, Aiba H, Ogasawara N (2006) Escherichia coli histone-Like protein H-NS preferentially binds to horizontally acquired DNA in association with RNA polymerase. DNA Res 13: 141–153.
- 13. Pan X (2006) Mechanism of trinucleotide repeats instabilities: the necessities of repeat non-B secondary structure formation and the roles of cellular trans-acting factors. J Genet Genomics 33: 1–11.
- 14. Otta E, Del Grosso N, Erba M, Melino S, Cicero D, et al. (2003) Interaction of DAPI with individual strands of trinucleotide repeats: effects of replication in vitro of the AAT · ATT triplet. Eur J Biochem 270: 4755–4761.
- 15. Hubacek JA, Pistulkova H, Valenta Z, Poledne R (1999) (TTA) n repeat polymorphism in the HMG-CoA reductase gene and cholesterolaemia. Vasa 28: 169–171.
- 16. Lyons-Darden T, Topal MD (1999) Effects of temperature, Mg2+ concentration and mismatches on triplet-repeat expansion during DNA replication in vitro. Nucleic Acids Res 27: 2235–2240.
- 17. Kappen LS, Xi Z, Jones GB, Goldberg IH (2003) Stimulation of DNA strand slippage synthesis by a bulge binding synthetic agent. Biochemistry 42: 2166–2173.
- 18. Lyons-Darden T, Topal MD (1999) Abasic sites induce triplet-repeat expansion during DNA replication in vitro. J Biol Chem 274: 25975–25978.
- 19. Leung TF, Liu EK, Tang NL, Ko FW, Li CY, et al. (2005) Nitric oxide synthase polymorphisms and asthma phenotypes in Chinese children. Clin Exp Allergy 35: 1288–1294.
- 20. Ujike H, Takaki M, Nakata K, Tanaka Y, Takeda T, et al. (2005) CNR1, central cannabinoid receptor gene, associated with susceptibility to hebephrenic schizophrenia. Mol Psychiatry 7: 515–518.
- 21. Martinez-Gras I, Hoenicka J, Ponce G, Rodriguez-Jimenez R, Jimenez-Arriero MA, et al. (2006) (AAT)n repeat in the cannabinoid receptor gene, CNR1: association with schizophrenia in a Spanish population. Eur Arch Psychiatry Clin Neurosci 256: 437–441.
- 22. Allon N, Leroy S, Roy C, Bourdel MC, Charles-Nicolas A, et al. (2006) (AAT)n repeat in the cannabinoid receptor gene (CNR1): association with cocaine addiction in an African-Caribbean population. Pharmacogenomics J 6: 126–130.
- 23. Barrero FJ, Ampuero I, Morales B, Vives F, de Dios Luna Del Castillo J, et al. (2005) Depression in Parkinson's disease is related to a genetic polymorphism of the cannabinoid receptor gene (CNR1). Pharmacogenomics J 5: 135–141.
- 24. Durkin SG, Glover TW (2007) Chromosome Fragile Sites. Annu Rev Genet 41: 169–192.
- 25. Schwartz M, Zlotorynski E, Kerem B (2006) The molecular basis of common and rare fragile sites. Cancer Lett 232: 13–26.
- 26. Pan X, Ding YF, Shi LF (2009) The roles of SbcCD and RNaseE in the transcription of GAA·TTC repeats in Escherichia coli. DNA Repair (Amst) 8: 1321–1327.
- 27. Kosmider B, Wells RD (2006) Double-strand breaks in the myotonic dystrophy type 1 and the fragile X syndrome triplet repeat sequences induce different types of mutations in DNA flanking sequences in Escherichia coli. Nucleic Acids Res 34: 5369–5382.
- 28. Pan X, Leach D (2000) The roles of mutS, sbcCD and recA in the propagation of TGG repeat in Escherichia coli. Nucleic Acids Res 28: 3178–3184.
- 29. Escoubas JM, Lane D, Chandler M (1994) Is the IS1 transposase, InsAB', the only IS1-encoded protein required for efficient transposition? J Bacterol 176: 5864–5867.
- 30. Lane D, Cavaille J, Chandler M (1994) Induction of the SOS response by IS1 transposase. J Mol Biol 242: 339–352.
- 31. Matsutani S (1997) Genetic Analyses of the Interactions of the IS1-encoded proteins with the left end of IS1 and its insertion hotspot. J Mol Biol 267: 548–560.
- 32. Biel SW, Berg DE (1984) mechanism of IS1 transposition in E.coli: Choice between simple insertion and cointegration. Genetics 108: 319–330.
- 33. Fang FC, Rimsky S (2008) New insights into transcriptional regulation by H-NS. Curr Opin Microbiol 11: 113–120.
- 34. Damea RT, Wymanb C, Goosena N (2001) Structural basis for preferential binding of H-NS to curved DNA. Biochimie 83: 231–234.
- 35. Amit R, Oppenheim AB, Stavans J (2003) Increased Bending Rigidity of Single DNA Molecules by H-NS, a Temperature and Osmolarity Sensor. Biophy J 84: 2467–2473.
- 36. Lang B, Blot N, Bouffartigues E, Buckle M, Geertz M, et al. (2007) High-affinity DNA binding sites for H-NS provide a molecular basis for selective silencing within proteobacterial genomes. Nucleic Acids Res 35: 6330–6337.
- 37. Bouffartigues E, Buckle M, Badaut C, Travers A, Rimsky S (2007) H-NS cooperative binding to high-affinity sites in a regulatory element results in transcriptional silencing. Nat Struct Mol Biol 14: 441–448.
- 38. Swingle B, O'Carroll M, Haniford D, Derbyshire KM (2004) The effect of host-encoded nucleoid proteins on transposition: H-NS influences targeting of both IS903 and Tn10. Mol Microbiol 52: 1055–1067.
- 39. Shiga YY, Sekine YK, Ohtsubo E (2001) Involvement of H-NS in transpositional recombination mediated by IS1. J Bacteriol 183: 2476–2484.
- 40. Rouquette C, Serre M-C, Lane D (2004) Protective Role for H-NS protein in IS1 transposition. J Bacteriol 186: 2091–2098.
- 41. Yasuzawa K, Hayashi N, Goshima N, Kohno K, Imamoto F, et al. (1992) Histone-like proteins are required for cell growth and constraint of supercoils in DNA. Gene 122: 9–15.
- 42. Dersch P, Schmidt K, Bremer E (1993) Synthesis of the Escherichia coli K-12 nucleoid-associated DNA-binding protein H-NS is subjected to growth-phase control and autoregulation. Mol Microbiol 8: 875–889.
- 43. Mancuso M, Sammarco MC, Grabczyk E (2010) Transposon Tn7 Preferentially Inserts into GAA•TTC Triplet Repeats under Conditions Conducive to Y•R•Y Triplex Formation. PLoS ONE 5: e11121.
- 44.
Miller JH (1992) A short course in bacterial genetics. N.Y: Cold Spring Harbor Laboratory, Cold Spring Harbor.
- 45. Phillips AJ, Church GM (1997) Methods for generating precise deletions and insertions in the genome of wild-type Escherichia coli: Application to open reading frame characterization. J Bacteriol 179: 6228–6237.
- 46. Yanisch-Perron C, Viera J, Messing J (1985) Improved M13 phage cloning vectors and host strains: nucleotide sequences of the M13mp18 and pUC19 vectors. Gene 33: 103–119.
- 47.
Sambrook J, Fitsch EF, Maniatis T (1989) Molecular Cloning: A Laboratory Manual. Cold Spring Harbor, Cold Spring Harbor Press.
- 48. Chen J, Zhang GC, Xu Y, Lu T, Wang CF (2001) Stability and copy number in host bacteria of a new plasmid cloning vector. Biotechnology 11: 6–8.
- 49. Pan X (2006) Effects of degU32(Hy), degQa and degR Pleiotropic Regulatory Genes on the Growth and Protease Fermentation of Bacillus subtilis Ki-2-132. J Genet Genomics 33: 373–380.