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Department of Biochemistry and Molecular Biology, Medical University of South Carolina, Charleston, South Carolina 29425
Received 19 June 2006/ Returned for modification 17 July 2006/ Accepted 21 August 2006
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ABSTRACT |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Termination of DNA replication has been well studied for prokaryotes (reviewed in references 6, 10, and 53). Thorough analysis of protein-DNA and protein-protein interactions, as well as cocrystal structures of terminator protein-DNA complexes, have generated a wealth of information leading towards an understanding of the mechanism of fork arrest in these systems (11, 27, 38, 51, 52, 58, 62, 67). The site-specific DNA binding terminator proteins Tus and RTP of Escherichia coli and Bacillus subtilis, respectively, form asymmetric protein-DNA complexes and arrest the oncoming replisome by preventing DNA unwinding by the replicative helicase. Fork arrest is polar or orientation specific, arresting only forks approaching the site from one direction, and this polarity is apparently generated by direct interaction of the functionally asymmetric terminator protein with the replicative helicase (46, 52). Site-specific fork arrest has also been described for various eukaryotic systems (reviewed in references 6 and 56). In S. cerevisiae, the protein Fob1 binds to two sites, Ter1 (RFB1) and Ter2, located within the rDNA intergenic spacer to cause polar fork arrest (30, 50). Similarly, the murine RNA polymerase I transcription terminator and the homologous Reb1 of S. pombe have been shown to cause fork arrest in the intergenic regions of mouse and fission yeast rDNA, respectively (22, 60). Although several recent studies have addressed the complex regulation of S. cerevisiae fork arrest in vivo (12, 20, 49, 65), biochemical analysis of eukaryotic fork-blocking complexes is lacking, and therefore the precise mechanism(s) of replisome blockage remains unknown. In fact, aside from crude DNase I footprinting of Fob1 (30) and Reb1 (68) with their respective binding sites, no studies have yet addressed the topology of eukaryotic terminator protein-Ter complexes.
In S. pombe, replication forks pause at four defined fork barriers, Ter1-3 (RFB1-3) and RFP4, within the rDNA intergenic spacer region (35, 60). The RNA polymerase I transcription terminator Reb1 arrests replication at Ter2 and Ter3 (60), whereas the switch-activating protein Sap1 binds to Ter1 to cause polar fork arrest at this site (34, 48). Sap1 is a multifunctional DNA binding protein, first identified by its binding to a site, SAS1, located centromere proximal to the mating-type switching locus mat1, which is required for mating-type switching (4). However, as contrasted with Ter1, Sap1 does not block replication at SAS1 (28, 34). Subsequent studies established that Sap1 is required for viability independently of its function in mating-type switching (2), being required also for maintenance of genome stability. These functions may require, at least in part, Sap1 binding to DNA without sequence specificity (18). In addition, a reiterative screening procedure has identified preferred synthetic Sap1 binding sites, consisting mostly of directly or indirectly repeated core TAA/GCG core motifs (23). Thus, Sap1 appears to be a rather promiscuous DNA binding protein involved in diverse biological functions.
Because analysis of terminator protein-DNA interactions has contributed to understanding fork arrest in prokaryotes, we wished to extend this type of analysis to the study of eukaryotic fork arrest in the rDNA intergenic spacer. We have used a variety of protein-DNA interaction techniques to probe the interaction of Sap1 with Ter1 as contrasted with SAS1. The results yield for the first time significant insight into the architecture of a eukaryotic fork-blocking complex and are compared to the strikingly different architecture of the Sap1-SAS1 imprinting complex. Upon DNA binding, Sap1 causes distinct local helical distortions, which mirror the architecture of the protein-DNA complex. In addition, we demonstrate by mutational analysis of the nonnatural direct repeat-type Sap1 binding site DR2 that modulating the interaction of the fork-proximal subunit of the Sap1 dimer converts this site from an inefficient to an efficient fork barrier in vivo without affecting the affinity of the dimer for the DNA. Local structural changes in the binding site accompany this conversion. The potential biological and mechanistic relevance of the findings is discussed below.
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MATERIALS AND METHODS |
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Preparation of substrates for DNA-protein interaction studies.
All binding sites were PCR amplified
with Vent polymerase (New England Biolabs) from pTer1(IRT2),
pDR2(IRT2), pSAS1(IRT2), and the indicated mutant sites
(34) to yield fragments
of 
160 bp in length. The PCR products were eluted from agarose
gels and 500 ng (5 pmol) was treated with
Optikinase (USB) and [
-32P]ATP (3,000 Ci/mmol;
Perkin-Elmer) according to the manufacturer's instructions. Excess
[-32P]ATP was removed with Sephadex G-25, and the
radiolabeled products were restricted with either AscI or EagI to yield
products labeled exclusively on either the top or the bottom strand,
respectively. The digested products were desalted using Sephadex G-25.
Sequences of all primers and plasmids used are available upon
request.
DNase I footprinting. For DNase I footprinting reactions, 30 fmol of wild-type or mutant Ter1- or SAS1-containing fragments (prepared as described above) was incubated with or without His6-Sap1 for 15 min at room temperature in Sap1 binding buffer (10 mM HEPES, pH 8.0, 100 mM NaCl, 7 mM ß-mercaptoethanol, 0.05% Triton X-100, 5% glycerol) containing 2.5 mM MgCl2 and 0.5 mM CaCl2. All reactions were performed in 10-µl volumes and contained 200 ng sheared salmon sperm carrier DNA. DNase I (0.015 U; New England Biolabs) was added for 2 min at room temperature. Reactions were stopped with 100 µl phenol and were immediately vortexed for 10 s and chilled on ice. The aqueous volume was adjusted to 100 µl with 20 mM EDTA and extracted, and the DNA was precipitated with ethanol and glycogen (Sigma), washed with 70% ethanol, and electrophoresed through 8% polyacrylamide-7 M urea sequencing gels containing 30% formamide (to avoid compression artifacts). Gels were dried and exposed to Biomax MS autoradiography film (Kodak).
Missing base contact interference. Missing base contact interference assays (9) were performed using either formic acid (depurination; Sigma) or hydrazine (depyrimidation; Fisher Scientific) as follows. For depurination reactions, 500 fmol (50 ng) of precut Ter1-, DR2-, or SAS1-containing fragments (prepared as described above) was incubated with 1 µl 4% formic acid in the presence of 1 µg sheared salmon sperm carrier DNA in a volume of 10 µl for 30 min at 37°C. The modified DNA was ethanol precipitated, washed with 70% ethanol, and resuspended in 10 µl distilled water. Three reaction mixtures were pooled, and binding reactions were performed using 6 µl modified DNA and 150 to 300 ng His6-Sap1 in Sap1 binding buffer (10 mM HEPES, pH 8.0, 100 mM NaCl, 7 mM ß-mercaptoethanol, 0.05% Triton X-100, 5% glycerol). The amount of protein was sufficient to bind the entire probe in the absence of any modifications. Binding was allowed to proceed for 15 min at room temperature. The reaction mixtures were electrophoresed through 5% native acrylamide gels containing 2.5% glycerol and 0.5x Tris-borate-EDTA (TBE). Bound and unbound bands were excised from the gel and purified using QIAGEN columns according to the manufacturer's instructions. Purified DNA was subsequently cleaved with 1 M piperidine for 30 min at 90°C and precipitated with n-butanol. Equal amounts of radioactivity were electrophoresed through 8% polyacrylamide-7 M urea sequencing gels containing 30% formamide. Gels were dried and exposed to Biomax MS autoradiography film (Kodak). Depyrimidation reactions were performed similarly, excepting that the DNA was modified with 30 µl hydrazine for 30 min at room temperature.
DMS protection. For dimethyl sulfate (DMS) protection assays, 30 fmol end-labeled substrates (prepared as described above) was incubated with indicated amounts of His6-Sap1 for 10 min at room temperature in Sap1 binding buffer (10 mM HEPES, pH 8.0, 100 mM NaCl, 7 mM ß-mercaptoethanol, 0.05% Triton X-100, 5% glycerol) containing 400 ng salmon sperm DNA. Binding volumes were 20 µl. Guanines were subsequently methylated by addition of 1 µl of 10% dimethyl sulfate (Sigma) for 2 min, and reactions were stopped by the addition of ammonium acetate to a 2 M final concentration and 2 volumes of ethanol. Modified DNA was precipitated, washed with 70% ethanol, and dried under vacuum. Cleavage at modified bases was performed by the addition of 100 µl 1.5 M piperidine and incubation at 90°C for 30 min, followed by precipitation with n-butanol. The dried DNA was resuspended in formamide loading buffer, boiled and chilled, and electrophoresed through 7 M urea-8% acrylamide sequencing gels containing 30% formamide. Gels were dried and exposed to Biomax MS autoradiography film (Kodak).
Hydroxyl radical protection. For hydroxyl radical protection assays, 15 fmol of end-labeled substrates (prepared as described above) was incubated with indicated amounts of His6-Sap1 for 15 min at room temperature in Sap1 binding buffer lacking glycerol and containing 400 ng salmon sperm DNA. Binding reaction volumes were 20 µl. To the binding reaction mixtures were added 1.5 µl of a 5 mM Fe-10 mM EDTA solution, 1.5 µl of 0.5 mM ascorbate, and 1 µl of 30% H2O2 for 4 min at room temperature, and the reactions were stopped with 75 µl of 100 mM thiourea-250 mM EDTA, followed by phenol extraction and ethanol precipitation. Precipitated DNA was washed once with 70% ethanol, dried thoroughly, and electrophoresed through 12% acrylamide-7 M urea sequencing gels containing 20% formamide. Gels were dried and exposed to Biomax MS autoradiography film (Kodak).
Ethylation interference. Ethylation interference assays were performed essentially as described previously (54). Briefly, 4.5 pmol of end-labeled substrates (prepared as described above) was brought to a 100-µl volume in 50 mM sodium cacodylate buffer. An equal volume of saturated ethyl nitrosourea (1 g/5 ml ethanol; Sigma) was added, and the mixture was incubated at room temperature for 30 min, followed by two rounds of ethanol precipitation in the presence of glycogen and subsequent washing with 70% ethanol. The precipitated nucleic acids were dissolved in 10 mM Tris, pH 8.0, and 1 mM EDTA and incubated with the indicated amounts of His6-Sap1 for 15 min at room temperature in Sap1 binding buffer (10 mM HEPES, pH 8.0, 100 mM NaCl, 7 mM ß-mercaptoethanol, 0.05% Triton X-100, 5% glycerol) containing 400 ng salmon sperm DNA. Binding volumes were 20 µl. The reaction mixtures were electrophoresed through 5% native acrylamide gels containing 2.5% glycerol and 0.5x TBE. Bound and unbound fractions were excised, and the DNA was cleaved at ethylated sites by the addition of sodium hydroxide to 150 mM for 30 min at 90°C. Cleavage reaction mixtures were neutralized with acetic acid (to 150 mM), ethanol precipitated, washed with 70% ethanol, and electrophoresed through 12% acrylamide-7 M urea sequencing gels containing 20% formamide. Gels were dried and exposed to Biomax MS autoradiography film (Kodak).
Potassium permanganate probing. Determination of sensitivities of His6-Sap1-bound binding sites to potassium permanganate (KMnO4) oxidation was performed essentially as described previously (13). Briefly, end-labeled substrates (prepared as described above) were incubated with indicated amounts of His6-Sap1 for 15 min at room temperature in Sap1 binding buffer lacking reducing agents and containing 400 ng salmon sperm DNA in 20-µl reaction volumes. Subsequently, 2 µl of 50 mM KMnO4 was added for 2 min at room temperature, after which the reaction mixtures were quenched by the addition of 2 µl ß-mercaptoethanol. The DNA was ethanol precipitated in the presence of glycogen, washed with 70% ethanol, and cleaved by the addition of 150 µl of 1 M piperidine for 30 min at 90°C. Cleaved DNA was precipitated with n-butanol and resolved through 8% acrylamide-7 M urea sequencing gels containing 30% formamide. Gels were dried and exposed to Biomax MS autoradiography film (Kodak).
Comparative gel mobility shift assays. Gel mobility shift assays, used to determine the relative affinities of His6-Sap1 for its various binding sites, were performed essentially as described previously (34), except for the following minor modifications. DNA probes containing the indicated binding sites were amplified from the respective plasmids, purified, and radiolabeled as described above and subsequently digested with KpnI followed by ethanol precipitation to yield substrates labeled only on the bottom strand. His6-Sap1 was diluted into buffer containing 10 mM HEPES, pH 8.0, 60 mM KCl, 1 mM dithiothreitol, 5 mM MgCl2, 100 µg/ml bovine serum albumin, and 50% glycerol. Binding reactions were performed in 20-µl reaction volumes containing 12 mM HEPES, pH 8.0, 100 mM NaCl, 50 µg/ml bovine serum albumin, and 7 mM ß-mercaptoethanol for 15 min at room temperature, followed by electrophoresis through 7.5% native acrylamide gels containing 2.5% glycerol and 0.5x TBE. Gels were directly exposed to phosphorimager screens at room temperature for 3 to 4 h for quantification of bound and unbound fractions.
2D agarose gel electrophoresis. Preparation and separation of replication intermediates by two-dimensional (2D) gel electrophoresis were performed as described previously (25, 35). In all cases, PvuII-digested plasmid intermediates were probed with a 2.2-kb fragment of the LEU2 gene.
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RESULTS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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25 and 30 bp on the top and bottom strands of Ter1,
respectively, from DNase I cleavage under the conditions used. As
expected, all three core motifs were protected (Fig.
1B). In contrast, Sap1
protected 30 and 35 bp on the top and bottom strands
of SAS1, respectively (Fig.
1B), providing initial
clues that the two complexes differ in mode of Sap1 binding.
Importantly, we found reactivity internal to SAS1 (Fig.
1A and B), which was
absent in Ter1, revealing that DNase I has access to the minor groove
in this region. These results suggested that the Sap1 dimer forms a
more compact complex on Ter1 than on SAS1. In order to further examine differences in the two complexes, we expressed and purified two His6-Sap1 truncations, Sap1(1-157) (with residues 1 to 157 deleted) and Sap1(22-157), for use in DNase I protection assays. These truncations were chosen for analysis as they retain the DNA binding domain as well as a region of the coiled C-terminal dimerization domain and have been shown to remain functional for SAS1 dimer binding as demonstrated by gel mobility shift assay (3, 23). The results of the footprinting experiments suggest that the Sap1(1-157) truncation and the wild-type protein protect similar regions of Ter1 and SAS1 from DNase I cleavage (Fig. 1C). In contrast, the Sap1(22-157) truncation, which lacks the ill-defined N-terminal domain presumed to mediate alternative protein dimerization in the absence of the C-terminal domain (23), reveals patterns of protection different from those of the wild-type and Sap1(1-157) proteins on both Ter1 and SAS1. Significantly, the patterns of protection differ also between Ter1 and SAS1 (Fig. 1C). Sap1(22-157) allowed for increased DNase I access to Ter1 in the region of the proximal core motif a on both the top and the bottom strand (Fig. 1C) but not in the distal region adjacent to motif c. In contrast, although both truncations bound SAS1 less tightly than Ter1, it was clear that Sap1(22-157) allowed increased DNase I access from both the proximal and the distal side of SAS1 on both strands (Fig. 1C). In summary, the DNase I footprinting results suggest that the Sap1-Ter1 and Sap1-SAS1 complexes exist as structurally distinct complexes. Experiments with the truncation mutants suggest that the N-terminal domain of at least one Sap1 subunit may reside in the vicinity of the proximal core motif a of Ter1. In contrast, the N-terminal domains appear to reside near both the proximal a and the distal c core motif of SAS1. The architectural models built by additional experiments in this paper further support this conclusion (see below).
Contrasting modes of sequence recognition at Ter1 versus SAS1. Identification of terminator protein-base pair contacts has resulted in significant insight into the mechanism(s) of fork arrest in prokaryotes (14, 38, 51, 58, 62). We wished to extend this analysis to the study of eukaryotic fork arrest at Ter1. Additionally, we reasoned that determination of base-specific contacts at Ter1 versus SAS1 should shed further light on the differences between the Sap1 fork-blocking and imprinting complexes, which may result in different biological functions of the complexes. Towards these goals, we performed missing base contact interference as well as DMS protection assays of Sap1 bound to either Ter1 (Fig. 2) or SAS1 (Fig. 3). Missing base contact interference involves the chemical removal of bases in order to determine which bases are required for stable protein binding (9). DMS methylates N7 of guanine residues and N3 of adenine residues within the major and minor grooves of the double helix, respectively, and complexes are subsequently assayed for residues that are protected from methylation by the bound protein. Thus, the techniques are complementary for determining base-specific contacts within protein-DNA complexes.
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The base-specific contacts of the Sap1-SAS1 complex differed markedly from those at Ter1 (Fig. 3A to C). As is the case for Ter1, Sap1 makes extensive contacts with motif a, as revealed by both missing base interference and DMS protection. However, few missing bases in the inverted motif b affected binding, and no significant methylation protection was observed in this region, suggesting that Sap1 makes few if any base contacts in this region. Instead, removal of bases from inverted motif c affected binding, and methylation protection confirmed that the protein made contact in this region (Fig. 3A to C). As was the case for Ter1, Sap1 appears to make only major-groove contacts with SAS1. Notably, as contrasted with the Sap1-Ter1 complex, we found that only a single base (an adenine on the bottom strand) (Fig. 3A and C) located between the core motifs affected Sap1 binding, perhaps suggesting that this region tolerated alterations in the backbone better than Ter1. This interpretation would be consistent with our DNase I footprinting and DNA bending results (34) suggesting that Sap1 is subject to considerably less space restraint when bound to SAS1 than when bound to Ter1.
In summary, base contacts of Sap1 with Ter1 and SAS1 reveal that the protein makes similar contacts with motif a of both sites. In contrast, directly repeated motif b contributes important base contacts to the Sap1-Ter1 complex, whereas Sap1 contacts primarily the inverted motif c in SAS1. Taken together with our results verifying that a single subunit of the dimer binds motif a of Ter1 (see below), the results suggest that the two complexes differ in sequence recognition and therefore binding configuration of the second but not the first subunit of the dimer and that the arrangement of motifs b and c may direct the mode of binding.
A single Sap1 subunit recognizes motif a of Ter1, and the Sap1 dimer subsequently nucleates at this site.
Both the monomeric Tus
and dimeric RTP replication terminator proteins of E. coli and
B. subtilis, respectively, have been shown to expose
asymmetric faces of the complex to the oncoming replication fork
(46,
52). Because Sap1 is a
dimer in solution and when bound to DNA
(5,
23), we wished to further
define the manner by which individual Sap1 subunits recognize and bind
Ter1, as asymmetric assembly of the dimer could, in principle,
contribute to polar fork arrest. Such asymmetry has been well
documented for the cooperative assembly of RTP dimers
(38,
41), although other
mechanisms must also contribute to polarity generation in this system
(19,
46). Towards this goal,
we utilized a truncated Sap1 protein, Sap1(1-136), which
retains the DNA binding domain but lacks the entire C-terminal
dimerization domain and is therefore unable to dimerize properly to
form a stable complex on either SAS1 or direct repeat-type binding
sites, as determined by gel mobility shift analysis
(3,
23). As expected, the
mutant is also unable to shift Ter1 in a gel mobility shift assay (data
not shown). However, Sap1(1-136) was able to recognize and bind
to Ter1, although with reduced affinity, as determined by DNase I
footprinting analysis. Significantly, only the proximal one-half of the
binding site was occupied by the mutant, even at high protein
concentrations (Fig. 4D and
F). Using a previously characterized triple mutant of motif
a (the TM1 mutant) (34),
we detected absolutely no DNase I protection, even at very high protein
concentrations (Fig. 4D,
right panel). Because the Sap1(1-136) mutant retains the entire
DNA binding domain and Sap1 is known to bind Ter1 as a dimer
(34), we postulated that
the Sap1(1-136) footprint represented the binding of a single
Sap1 subunit to Ter1. To gain further support for this notion, we
utilized a Ter1 
1 mutant, previously shown to be completely
defective in stable Sap1 binding as determined by gel mobility shift
assay (34). This mutant
contains a G deletion immediately preceding motif b
(34) (Fig.
4F). DNase I footprinting
analysis of wild-type Sap1 on the Ter1 1 mutant resulted in a
footprint identical to that of Sap1(1-136) on wild-type Ter1
(compare Fig. 4A and D),
suggesting that the dimer can fill only the proximal one-half of Ter1
in the presence of the 1 mutation, which presumably prevents
recognition of motif b. DMS protection experiments support these
results, as guanine residues within only motif a and not motif b are
protected by binding of Sap1 to the 1 mutant site (Fig.
4B).
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1 mutant and of
Sap1(1-136) on wild-type Ter1 reveal that the respective
proteins make backbone contacts with sugar-phosphates only in the
proximal half of Ter1 (Fig. 4C, E, and
F), as contrasted with contacts made by full-length Sap1 on
the wild-type Ter1 (Fig.
5A). Minor differences in protection from hydroxyl radical
cleavage were noted for the bottom strand. Specifically, additional
sugar-phosphates were protected in the Ter1 1 mutant-bound
Sap1 complex compared to the wild-type-Ter1-bound Sap1(1-136)
complex (Fig. 4C, E, and
F). In light of the data as a whole, we interpret these
minor differences to perhaps represent conformational differences
between the full-length Sap1 dimer and the Sap1(1-136)
truncation when bound to motif a, rather than differences in site
recognition per se.
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Sap1-induced helical distortion. We have demonstrated previously that Sap1 bends both Ter1 and SAS1 upon binding, although bending of Ter1 was reproducibly greater than that of SAS1 (34). These results suggested that the helices were distorted to different extents within the two complexes. As DNA bending and helical distortion are common among bacterial terminators (27, 33) and may contribute to the mechanism of fork arrest in these systems, we were interested to investigate further the Sap1-mediated structural changes at Ter1 and to compare these with the Sap1-SAS1 complex. KMnO4 efficiently oxidizes thymine (and, to a lesser extent, cytosine) bases only when the DNA helix is locally unwound or distorted, and the modified bases subsequently may be detected by piperidine cleavage (61). We therefore probed His6-Sap1-bound and unbound Ter1 and SAS1 with KMnO4 in order to detect protein-mediated helical distortions at these sites.
Sap1 binding resulted in strand-specific exposure of KMnO4-sensitive bases at Ter1, as only the bottom strand showed hypersensitivity to KMnO4 (Fig. 6A, left two panels). Specifically, the second-position bottom-strand thymine of motif a became reactive after Sap1 binding. In addition, the first-position thymine of the inverted motif c was reactive, although to a greatly reduced extent (Fig. 6A). In addition, a bottom-strand cytosine immediately preceding motif b was reactive (Fig. 6A). Reactivity at this site was severely reduced compared to that of the thymines, but this may be because KMnO4 oxidizes cytosines inefficiently compared to thymines (57). For comparison, we performed similar experiments with the His6-Sap1-SAS1 complex. As expected, the pattern of Sap1 binding-induced KMnO4 reactivity was strikingly different at this site compared to that at Ter1. Notably, both strands of SAS1 became hypersensitive to KMnO4 upon Sap1 binding (Fig. 6A, right two panels). Although reactivity of the second-position thymine of motif a was analogous to that at Ter1, the reactivities at motifs b and c differed. As with the a motifs of both Ter1 and SAS1, second-position thymines of SAS1 motifs b and c were modified. It should be noted that in each case (motif a of Ter1 and motif a of SAS1, as well as motifs b and c of SAS1), the Sap1-exposed thymines were on the strand opposite the defined TAA/GCG core sequences. This likely reflects the orientation of subunit binding to these motifs, as summarized below. It is interesting to note also that the helical distortion at these bases is exquisitely local, as demonstrated by the KMnO4-insensitive neighboring thymines in the third position of motif a (Fig. 6A and C). In addition to exposure of these motif-embedded thymines, a single top-strand thymine located between motifs b and c of SAS1 became KMnO4 sensitive upon Sap1 binding (Fig. 6A and C).
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Alternate symmetries of the Sap1 terminator and imprinting complexes. We have used a series of footprinting and chemical modification, protection, and interference techniques in order to identify the protein-base pair and protein-backbone contacts of Sap1 at the fork barrier site Ter1 and have contrasted the results with contacts at SAS1. Accordingly, we have been able to describe the molecular architectures of both complexes. The results of these experiments have been synthesized and modeled onto double-helical representations of the respective binding sites, as depicted in Fig. 7. Representation of the data in such a manner allows for visualization of the striking architectural differences between the two functionally distinct complexes.
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Helical projection of the Sap1-SAS1 complex reveals the stark architectural differences of this complex compared to Sap1-Ter1 (Fig. 7). Base pair and sugar-phosphate contacts at motif a resemble analogous contacts at Ter1, and backbone contacts are also made on the opposite lip of this major groove, suggesting that a single Sap1 subunit binds to this region in a manner similar to that of Ter1. However, motif c appears to contribute most of the remaining base-specific contacts. Furthermore, Sap1 makes extensive sugar-phosphate contacts with motifs b and c from the opposite face of the helix (Fig. 5 and 7). These contacts reveal mirror image symmetry with respect to contacts at motif a, thus providing the complex with rotational symmetry rather than the translational symmetry evident at Ter1 (Fig. 7). Thus, although the C-terminal dimerization domain of Sap1 appears to be required for stable binding of both SAS1 and Ter1 as determined by gel mobility shift assay (23; this study), the second subunit of the dimer appears to be oriented in opposite orientations in the two complexes (Fig. 7 and 8). In addition, the complex accommodates an extra 1.5 helical turns between subunits (Fig. 7). Sap1-induced Ter1 and SAS1 structural distortions mirrored the alternate symmetries of the two complexes (Fig. 6 and 7).
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Because the
sequences of Ter1 and DR2 (Fig.
8A) as well as the base
pair contacts of Sap1 with the two sites (Fig.
2) are very similar, we
were interested to determine which region of Ter1 conferred upon this
site the ability to arrest forks more efficiently than DR2. We
therefore mutated several regions of DR2 and analyzed the mutants for
extrachromosomal replication fork arrest (Fig.
8A). We chose not to
mutate motif a, as this region is crucial for recruitment of Sap1 to
Ter1 (Fig. 4) and
mutations in this region of Ter1 abolish stable Sap1 binding and fork
arrest (34,
48). Two-dimensional gel
analysis of the DR2 mutants revealed that only one of the mutants (the
DR2-D mutant), in which motif b (TAGCG
CAAGG) as well as the
region corresponding to motif c of Ter1
(AGCCCGAGCTAT) has
been altered to resemble the analogous bases of Ter1, was able to
arrest replication efficiently. Mutating either of these regions
individually had no effect on fork arrest (Fig.
8A and B). Interestingly,
although we were unable to detect any base-specific contacts at motif c
of Ter1 by the methods used (Fig.
2), this region appears to
cooperate with motif b to cause efficient fork arrest. Because Sap1
makes several sugar-phosphate contacts with motif c of Ter1 (Fig.
5), we hypothesize that
these contacts are critical for efficient fork arrest. In summary, the
results suggest that modulating the interaction of the fork-proximal
Sap1 subunit with its direct repeat-type binding site suffices to
convert an inefficient fork barrier into an efficient
one.
Relative affinities of various Sap1 binding sites.
Several
studies of prokaryotic systems have suggested that the mechanism of
fork arrest is probably not explained simply by strong binding of the
terminator proteins to their cognate sites
(19,
21,
63), nor can DNA binding
alone account for the polarity of the process
(46,
52). In our system, the
lack of fork arrest at SAS1 and the inefficient arrest at DR2 could
theoretically be explainable by the decreased affinity of Sap1 for
these sites compared to that for Ter1. Alternatively, striking or
subtle architectural differences between the various Sap1 complexes may
determine whether or not forks are blocked. In order to begin to
distinguish between these possibilities, we determined the relative
binding affinities of Sap1 bound to Ter1, DR2, DR2-D, or SAS1 by using
comparative gel mobility shift assays. As shown in Fig.
9A, the binding curves of Sap1 bound to Ter1, DR2, and DR2-D are nearly
identical. We approximated the relative Kd values
from these curves by determining which concentrations of Sap1 resulted
in half-maximal DNA binding. Accordingly, the mean half-maximal binding
values (±standard deviations) of Sap1 to Ter1, DR2,
and DR2-D were 5.5 (±1.4) nM, 2.6 (±0.5) nM, and 4.4
(±0.4) nM, respectively. As the results clearly show that the
binding affinities of Sap1 for the sites are comparable, the relatively
inefficient fork arrest at DR2 could not be explained by a lower
affinity of Sap1 for this site than for Ter1. Similarly, the DR2-D
mutation restored efficient fork arrest not simply by increasing the
affinity of Sap1 for this site. The results are consistent with the
aforementioned studies of prokaryotes and suggest that DNA binding
affinity alone cannot account for the mechanism of Sap1-mediated
replication fork arrest. The relative affinity of Sap1 for SAS1 was
reduced 7- to 8-fold compared to the affinity for the direct
repeat-type sites, with an approximate Kd value of
37.2 (±8.9) nM (Fig.
9A), precluding
conclusions about whether binding affinity and/or the architecture of
the Sap1-SAS1 complex accounts for the lack of fork arrest at this
site.
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DISCUSSION |
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Sap1 binding induces conformational changes in its binding sites (Fig. 6 and 9B). As expected, these helical distortions reflect the mode of dimer binding. However, we were surprised to find that, although mild helical distortion was evident internal to motifs a and b of both Ter1 and SAS1, the major distorted regions were located within the core recognition motifs in all sites analyzed. Thus, protein binding apparently led to helical distortion directly at the site of subunit contact. Furthermore, the distortion was extremely localized, as even immediate thymine neighbors of the core motifs were unaltered as probed by KMnO4 (Fig. 6A and C). The symmetry of distortion directly paralleled the symmetry of binding motifs, such that every second-position bottom-strand thymine of the sequence TAACG or TAGCG was modified regardless of which site was analyzed (Fig. 6 and 9B). These results suggest that KMnO4 reactivity within the recognition motifs may be due to the mechanism of binding of the Sap1 DNA binding domain itself rather than to passive untwisting of the area during dimer binding. This would differ from the distortion induced between the motifs in Ter1 and SAS1 (but lacking at DR2), which is probably due to dimer-induced bending. The potential significance of these structural changes for fork arrest is discussed below. It is interesting to note that such localized KMnO4 reactivity has been found to be due to a base-flipping mechanism in several methyltransferases and DNA repair enzymes (55, 61), suggesting that Sap1 may also flip out bases during binding. Alternatively, Sap1 may open the helix at the core motifs to insert contact elements.
We have demonstrated that nonnatural Sap1 binding site DR2, which is highly homologous in sequence to Ter1, binds Sap1 in a manner similar to Ter1 yet acts as only a very inefficient fork barrier in vivo (Fig. 8). Notably, the DNA binding affinities of Sap1 for these two sites are nearly identical, suggesting that binding affinity alone cannot explain the differences in fork arrest. These results are consistent with similar evidence from prokaryotic systems (19, 24, 52, 63). Modulating the interaction of the replication fork-proximal Sap1 subunit with DR2 can convert this site from an inefficient to a very efficient replication fork barrier (DR2-D) (Fig. 8A). Again, binding affinities of Sap1 for DR2 versus DR2-D are nearly identical, suggesting that the mutations affect fork arrest by means other than enhancement of binding affinities. It is therefore interesting to note that Sap1 causes distinctly different KMnO4 reactivities at Ter1 versus DR2 and that KMnO4 reactivity at DR2-D is similar to that at Ter1. We hypothesize that, although the Sap1 dimer appears to bind both Ter1 and DR2 in architecturally similar configurations, subtle changes in the nature of the protein-DNA complex may determine whether or not forks are arrested. These structural changes could, in principle, lead to an altered DNA-protein complex that forms a more stable impediment to the oncoming replication fork. It is perhaps notable, therefore, that the mutations which increased fork-blocking efficiency of DR2 are located at the replication fork-proximal side of the complex. Binding-dependent structural changes could thus expose the Sap1-DNA complex to the oncoming replisome in a manner suitable to arrest fork progression, either through direct interaction with components of the replisome or more indirectly. Such DNA binding-induced allosteric protein-DNA conformational changes have been suggested to modulate transcription factor function (40). Bacterial terminator proteins block fork progression by inhibiting replicative helicases (29, 39). This is accomplished through both protein-DNA and protein-protein interactions of the terminator proteins with the replicative helicase (46, 51, 52). It will be interesting to determine whether eukaryotic fork-blocking proteins, such as Sap1, also act as contrahelicases and whether this is dependent on the structural context.
Most prokaryotic and eukaryotic replication fork barriers, including Ter1, are polar in nature. In bacteria, polarity is generated by the assembly of a functionally asymmetric terminator-protein complex (reviewed in reference 6). For instance, both RTP and Tus, the terminator proteins of B. subtilis and E. coli, respectively, are situated on the DNA to make specific contacts with the oncoming replicative helicase of the replisome from only one orientation (46, 52, 62). In addition, the crystal structure of the Tus-Ter complex reveals asymmetric backbone contacts (27), and RTP, which requires cooperative binding of two dimers to its binding site in order to cause fork arrest (41, 58), makes asymmetric nucleoside contacts with the fork-proximal and fork-distal halves of the Ter site (38). Although we have not detected any notable asymmetry in the DNA-protein contacts of the Sap1-Ter1 complex, DNA ligand-induced asymmetry of the protein could still generate polarity. In this respect, it is interesting to note that the Sap1 subunits arrange themselves into a complex with translational symmetry (Fig. 7), resulting in a structurally polar complex. The replication fork approaches Ter1 in situ from the side containing motif c, so the fork would encounter drastically different faces of the fork-pausing complex depending on the direction from which it approached the site. Future experiments should address whether the blocking-competent complex achieves polarity by direct interactions of the fork-proximal subunit with a component of the oncoming replisome or whether the mechanism of polarity generation is more indirect.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Published ahead of print on 28 August 2006.
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