Reorganization of terminator DNA upon binding replication terminator ...

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 1997 Oxford University Press

Nucleic Acids Research, 1997, Vol. 25, No. 3

Reorganization of terminator DNA upon binding replication terminator protein: implications for the functional replication fork arrest complex Andrew V. Kralicek, Paul K. Wilson, Greg B. Ralston, R. Gerry Wake and Glenn F. King* Department of Biochemistry, University of Sydney, Sydney, NSW 2006, Australia Received October 1, 1996; Revised and Accepted November 25, 1996

ABSTRACT Termination of DNA replication in Bacillus subtilis involves the polar arrest of replication forks by a specific complex formed between the replication terminator protein (RTP) and DNA terminator sites. While determination of the crystal structure of RTP has facilitated our understanding of how a single RTP dimer interacts with terminator DNA, additional information is required in order to understand the assembly of a functional fork arrest complex, which requires an interaction between two RTP dimers and the terminator site. In this study, we show that the conformation of the major B.subtilis DNA terminator, TerI, becomes considerably distorted upon binding RTP. Binding of the first dimer of RTP to the B site of TerI causes the DNA to become slightly unwound and bent by ∼40. Binding of a second dimer of RTP to the A site causes the bend angle to increase to ∼60. We have used this new data to construct two plausible models that might explain how the ternary terminator complex can block DNA replication in a polar manner. In the first model, polarity of action is a consequence of the two RTP–DNA half-sites having different conformations. These different conformations result from different RTP–DNA contacts at each half-site (due to the intrinsic asymmetry of the terminator DNA), as well as interactions (direct or indirect) between the RTP dimers on the DNA. In the second model, polar fork arrest activity is a consequence of the different affinities of RTP for the A and B sites of the terminator DNA, modulated significantly by direct or indirect interactions between the RTP dimers. INTRODUCTION Completion of a round of replication of the circular chromosomes of Bacillus subtilis and Escherichia coli involves arrest of replication forks at specific DNA sequences of 30 bp in length that are located in the terminus region approximately opposite the origin. Arrest at these sites requires the binding of a terminator protein to form a DNA–protein complex that blocks fork movement in a polar manner (i.e. it blocks forks approaching from one

direction only) (1,2). This complex has been proposed to cause replication fork arrest by inhibiting the DNA unwinding activity of the helicase present at the apex of the replication fork (3,4). A set of at least six DNA terminators has been identified in the terminus regions of both bacteria (Ter I–VI in B.subtilis and Ter A–F in E.coli) (5,6); at least three of these terminators are orientated to stop the anticlockwise replication fork, whilst the other three are positioned to stop the clockwise replication fork. In both organisms, the relative arrangement of the two groups of terminators is such that the terminus region acts as a replication fork trap, allowing replication forks to enter the region but not exit (see 7). Surprisingly, there is no significant sequence or structural homology between the terminator proteins of the two organisms (RTP in B.subtilis and Tus in E.coli) nor between the sequences of their DNA terminator sequences (8,9). Thus, the two organisms may use quite different molecular mechanisms to achieve helicase inhibition. RTP is a 29 kDa symmetrical dimer (10–12) which interacts specifically with an ∼30 bp region of the major B.subtilis terminator TerI (originally described as a 47 bp segment of DNA called IRI; see 13). This binding region (see Fig. 1) contains two overlapping RTP dimer binding sites, A and B, the latter having a higher affinity for RTP. The B site is located proximal to the approaching fork and is pseudosymmetrical in sequence (11). When RTP is bound to this site alone, replication fork arrest does not occur; a second RTP dimer must bind cooperatively to the low affinity A site to produce an active terminator (14,15). The crystal structure of RTP was recently solved at 2.6 Å resolution (12). As predicted on the basis of heteronuclear NMR experiments (11), RTP is a symmetrical dimer which contains both α-helix and β-sheet secondary structure; helices 1–3 are arranged in a similar fashion to the winged-helix DNA binding domain (16) and are believed to represent the primary DNA recognition motif (12,17). While the crystal structure has provided a foundation for constructing models that might explain how a single RTP dimer interacts with terminator DNA, much more information is required in order to understand how a functional fork arrest complex is assembled. In particular, the molecular details of the interaction of each RTP dimer with the A and B sites of the terminator have still not been resolved and many questions remain regarding the nature of the interaction between the two docked RTP dimers and the effect that RTP binding might have on the structure of the terminator DNA.

*To whom correspondence should be addressed. Tel: +61 2 9351 3902; Fax: +61 2 9351 4726; Email: [email protected]

591 Nucleic Acids Acids Research, Research,1994, 1997,Vol. Vol.22, 25,No. No.13 Nucleic

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Circular dichroic spectroscopy

Figure 1. Sequence features of the minimal functional region (29 bp) of the B.subtilis chromosomal terminator TerI. The boxed trinucleotide (TAT) occupies the central portion of the terminator such that 13 bp lie to either side of it. The overlapping lines above the sequence represent the two RTP binding sites, A and B, while the arrowheads below the sequence represent the limited symmetry between the A and B sites. The asymmetry in protein–DNA contacts between the two sites is readily seen from the filled circles, which show the prominent DNA–protein contacts as mapped using missing nucleoside interference experiments (11).

In this study, we have addressed the latter question by using circular dichroic (CD) spectroscopy and circular permutation band retardation assays to examine changes in the structure of TerI DNA upon binding of RTP. We show that binding of RTP to TerI causes considerable distortion of the terminator DNA; the DNA becomes noticeably bent and slightly unwound. We have used this information in combination with the RTP crystal structure (12), mutagenesis studies (18), missing nucleoside interference data (13) and comparative studies of various terminator DNA sequences (19) to critically examine various models of the mechanism by which the RTP2–DNA ternary complex causes replication fork arrest. MATERIALS AND METHODS Materials All chemicals were of analytical or molecular biology grade. Unlabelled RTP was prepared as described previously (11). The plasmid pBend2 was a generous gift from S.Adhya (Bethesda, MD). The plasmid pPW2 was produced in this laboratory (see below). Restriction endonucleases (EC 3.1.21.4) and mung bean nuclease (EC 3.1.30.1) were obtained from Boehringer Mannheim (Mannheim, Germany). T4 DNA ligase (EC 6.5.1.1) was obtained from New England Biolabs (Beverly, MA). T4 polynucleotide kinase (EC 2.7.1.78) was obtained from Amersham International (Amersham, UK). T7 DNA polymerase (EC 2.7.7.7) was obtained from Pharmacia LKB (Uppsala, Sweden).

All CD spectra were acquired using a Jasco J-720 spectropolarimeter under constant nitrogen flush at 20C. The spectropolarimeter was calibrated with an aqueous solution of ammonium D-camphor10-sulfonic acid (20). The temperature of the cell was maintained and adjusted with a Neslab RTE-111 circulating water bath. All spectra were acquired using the following parameters: four scans, resolution 0.2 nm, bandwidth 1.0 nm, sensitivity 50 mdeg, response time 1 s and scan speed 20 nm/min. Except for the titration study of B site DNA, all CD studies were performed using a 0.1 cm cell containing 300 µl samples with a buffer of 5 mM NaH2PO4, pH 7.8, 1 mM DTT. In all cases, prior to spectral acquisition, the sample was prepared at room temperature and incubated for at least 5 min within the spectropolarimeter to ensure temperature and solvent equilibration. Spectral signal-tonoise was optimized by multiplying the frequency power spectrum by a trapezoidal apodization function prior to inverse Fourier transformation to give the noise-reduced frequency domain spectrum. CD spectra of B site DNA alone (10 µM) were acquired over the wavelength range 180–320 nm. The stoichiometry of the RTP–B site interaction was monitored by the stepwise addition of 4 µl 0.1625 µM RTP to 2600 µl 0.5 µM B site DNA in 0.5 mM NaH2PO4, 0.1 mM DTT, pH 7.8, in a 1 cm cell. Previous attempts using a 5 µM B site sample in standard buffer in a 0.1 cm cell were subject to mixing problems, large dilutions and often transient precipitation of the added protein: these problems did not occur with the 1 cm cell. Spectra were acquired between 260 and 290 nm at the following ratios of RTP to B site: 0, 0.25, 0.5, 0.75, 1.0, 1.25, 1.5, 1.75, 2.0, 2.25, 2.5, 3.0, 3.5 and 4.0. Prior to spectral acquisition, the sample was allowed to incubate for 10 min at 20C to ensure that the binding reaction had reached equilibrium. Band retardation assay with B site DNA These were performed essentially as described previously (21). Binding reactions using RTP dimer:B site molar ratios of 0, 0.25, 0.5, 1.0 and 2.5:1 were performed under identical conditions to the CD experiments (i.e. 5 µM B site DNA in 5 mM NaH2PO4, 1 mM DTT, pH 7.8, 20C). After allowing 10 min for DNA– protein complexation, reaction mixtures were electophoresed on a 6% polyacrylamide gel and stained with ethidium bromide. DNA bending experiments with the TerI region

Preparation of a site B DNA fragment Two 20 nt oligonucleotides spanning the 16 nt complementary strands of the B site from TerI (see Fig. 1) were synthesized by Biotech International (Perth, WA): fragment 1, 5′-CTATGTACCAAATGTTCAGT-3′; fragment 2, 3′-GATACATGGTTTACAAGTCA-5′. Equimolar quantities of each DNA strand were mixed at room temperature in TE buffer (10 mM Tris, pH 7.5, 1 mM EDTA) and placed in a heating block at 95C for 5 min. The fragments were then allowed to anneal by slow cooling over 40 min to 37C, followed by incubation at this temperature for a further 1 h. The solution was then cooled to room temperature overnight. The duplex DNA was ethanol precipitated, aerated and resuspended in phosphate buffer (5 mM NaH2PO4, pH 7.8, 1 mM DTT) for CD and band retardation analysis. The concentration of each duplex was determined from the absorbance at 260 nm; it was assumed that an A260 of 1.0 is equivalent to a double-stranded DNA concentration of 50 µg/ml.

A double-stranded DNA oligonucleotide containing the 29 bp minimal region of TerI plus an additional 3 bp beyond the B site, as well as protruding single-strand XhoI ends, was phosphorylated (5′) and cloned into the SalI site of pBend2 (22) to give the plasmid pPW2. This plasmid was digested with each of the restriction enzymes MluI, BglII, NheI, XhoI, PvuII, StuI and BamHI to yield DNA fragments of approximately the same length (154 bp). The 5′-termini of the resulting fragments were labelled with [γ-32P]ATP (7000 Ci/mmol, 5 mCi) using T4 polynucleotide kinase. The labelled fragment was purified from the unincorporated radionucleotide using a Nick Column (Pharmacia). Band retardation experiments were performed using a DNA concentration of 6.8 × 10–13 mol/µl in a total volume of 6 µl TGMK (21). An RTP (monomer):DNA ratio of 1:1 was used to enable both of the DNA–RTP complexes, I and II, and the free DNA species to be visualized. After incubation at 25C for 30 min,

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Figure 2. CD spectra of B site DNA (5 µM) dissolved in phosphate buffer (5 mM NaH2PO4, 1 mM DTT, pH 7.8) either with (—) or without (•••) 80% TFE (v/v). θ, mean nucleotide ellipticity.

bromophenol blue (0.07%) was added and samples were loaded onto a 10% non-denaturing polyacrylamide gel (in 36 mM Tris, 30 mM NaH2PO4, pH 7.5) and run at 8.3 V/cm for 3–7 h. The gel was fixed in acetic acid (10%), exposed to a Storage Phosphor Screen and analysed using a PhosphorImager (Molecular Dynamics, Sunnyvale, CA). RESULTS The structure of uncomplexed B site DNA Figure 2 (solid line) shows the CD spectrum of a double-stranded oligonucleotide corresponding to the B site of B.subtilis TerI DNA. It has a number of features that identify it as being B-form in solution (23,24): it has a major longwave positive peak centred at 275 nm and the intensity of this positive peak is similar in magnitude to that of the negative peak centred at ∼240 nm. Addition of 80% trifluoroethanol (TFE) to B-form DNA has been shown to induce a B to A transition, resulting in a CD spectrum with characteristics totally different from those of B-form DNA; the longwave positive peak is larger with a maximum at ∼270 nm and a very large shortwave peak becomes apparent below 200 nm (24–26). The changes in the CD spectrum of B site DNA upon addition of 80% TFE (Fig. 2, dotted line) are consistent with a B to A transition, providing further evidence that ‘free’ B site DNA is B-form in aqueous solution. The structure of B site DNA when complexed with RTP As demonstrated above, the longwave positive CD band of nucleic acids is very sensitive to the local structure of DNA. This band has been used as an indicator of conformational changes in DNA upon binding various proteins, as well as a means of investigating the stoichiometry of the binding reaction; examples of such studies include those on Tet repressor (27), trp repressor (28) and CAP (29). As shown in Figure 3A, addition of RTP to B site DNA results in a marked enhancement in ellipticity and a slight blueshift (to 273.5 nm) of the band at 275 nm. These spectral changes indicate that binding of RTP causes a conformational change in B site DNA (see below); furthermore, the changes in ellipticity can be

Figure 3. (A) CD spectra of B site DNA (0.5 µM in 5 mM NaH2PO4, 1 mM DTT, pH 7.8) in the absence (—) or presence of RTP at RTP dimer:DNA molar ratios of 0.25:1 (•••), 0.375:1 (— - —), 0.75:1 (—•••—), 0.875:1 (- - -) and 1:1 (— — —). θ, mean residue ellipticity. (B) The change in ellipticity at 275 nm (∆θ) of B site DNA as a function of the RTP dimer:DNA molar ratio.

used to measure the stoichiometry of the protein–DNA complex. The titration curve shown in Figure 3B shows that there was an increase in ellipticity as more RTP was added, until a maximum increase of 66% was obtained at a ratio of one RTP dimer to one B site duplex. The 1:1 stoichiometry of the specific complex argues that a free RTP dimer cannot bind and produce an optical change in a region of DNA already in complex with RTP. This stoichiometry was confirmed by a band retardation experiment with the B site under the conditions of the CD experiment (data not shown), which showed binding of a single dimer to the B site oligonucleotide. Conformational changes in RTP upon complexation with B site DNA The only contribution of a protein to the CD signal in the wavelength range 260–290 nm comes from its aromatic residues. For most proteins, this contribution is small relative to the CD signal from DNA (30); indeed, the CD signal of concentrations of free RTP encompassing those used in the B site titration was found to be within the noise level of the solvent baseline for the


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Figure 4. A phosphorimage showing results of gel electrophoresis of 32P-labelled fragments of DNA (all ∼154 bp, with the RTP binding region at various locations) complexed with one dimer (complex I) or two dimers (complex II) of RTP. Free DNA is labelled F. The fragments analysed in lanes 1–7 were obtained by cutting pPW2 with MluI, BglII, NheI, XbaI, PvuII, StuI and BamHI respectively. The mid-points of the A+B binding region, from the end proximal to the A site, within the respective fragments were at positions 26, 32, 38, 62, 80, 92 and 129 bp respectively. Note that the BglII-cut fragment (lane 2) migrates anomalously.

260–290 nm region. If the substantial increase in ellipticity at 275 nm observed upon RTP binding to B site DNA was due to RTP alone, a radical reorganization of the protein would be required. While this seems unlikely, it is not impossible; for example, the DNA binding domains of bZIP transcription factors are disordered in the uncomplexed protein but become α-helical upon complexation with DNA (31). Thus, in order to investigate whether RTP undergoes a conformational change upon binding to B site DNA, a CD spectrum was acquired of RTP in the presence of an excess of the B site (RTP dimer:B site molar ratio of 1:2) to ensure complexation of all RTP molecules (data not shown). Because of a small contribution to the ellipticity of the complex from the B site DNA (especially in the region below 200 nm; see Fig. 2), this spectrum could not be deconvoluted into secondary structure components (20) and compared with the deconvoluted spectrum of free RTP. Nevertheless, the spectrum of the binary complex was very similar to that of free RTP over the region 200–260 nm, thus enabling us to conclude that there are no gross conformational changes in RTP upon binding to B site DNA and that the large changes in ellipticity at 275 nm are reporting exclusively on conformational changes in B site DNA. Bending of TerI DNA upon complexation with RTP Bent DNA migrates anomalously in non-denaturing polyacrylamide gels because of its reduced ability to reptate through the pores of the gel matrix (31–34). The mobility of the complex depends on the location of the bend within the DNA fragment; minimal migration occurs when the bend is located near the middle of the fragment, as this gives the largest effective Stokes radius (34). The circular permutation assay of Kim et al. (22), which takes advantage of the anomalous gel behaviour of bent DNA, was used to investigate whether the binding of RTP to TerI causes bending of the DNA. The RTP binding segment of TerI was cloned into the vector pBend2 to give pPW2, which was digested with a number of restriction endonucleases to produce a set of DNA fragments of similar length (∼154 bp) in which the binding region (A+B sites) was circularly permutated (i.e. the RTP binding sequence was

Figure 5. A graph showing the mobilities of complexes I and II relative to the free DNA fragment as a function of the mid-point of the relevant RTP binding region (B and A+B for complexes I and II respectively) within the ∼154 bp fragment. The relative mobilities are shown as the mean ± SEM (three experiments), with the SEM being less than the size of the data point in each case. Appropriate phosphorimages giving much narrower bands (particularly for complex II) were used for distance measurements.

placed at various positions relative to the fragment ends). These fragments displayed no systematic variation in their electrophoretic mobility in the absence of RTP (data not shown), implying that the binding region is linear when not complexed with the terminator protein. Figure 4 shows the result of an experiment analysing the behaviour of complexes I and II formed between RTP and the ∼154 bp fragments in which the binding region is located at various positions within each fragment. Complex I contains one dimer of RTP bound to the B site; complex II contains two bound dimers, with the second dimer binding to the A site. The RTP:DNA ratio was such as to leave some of the fragment unbound to RTP (species F). Figure 5 summarizes the relative mobilities (ratio of distance migrated by the complex to that of the free fragment) from several experiments. The mobilities are plotted against the mid-point of the B site in the case of complex I and the mid-point of the A+B sites for complex II. Suitable restriction sites were unavailable for generating fragments in which the binding sites were located between the 92 and 121 bp positions. The curves for both complexes show a systematic dependence of mobility on binding site position, with a minimum in each case when the site is located centrally in the ∼154 bp fragment. Significantly, the curvature is more pronounced in the case of complex II. Assuming that the site-dependent mobility reflects a simple bend in the DNA, bend angles of ∼40 and ∼60 were calculated for complexes I and II using the empirical equation of Thompson and Landy (35). While these calculated bend angles are only very approximate, the data show convincingly that the binding of a single dimer to the B site alone (complex I) causes a measurable conformational change (bend) in the DNA and that an additional conformational change (bend) accompanies binding of a second dimer to the A site (complex II).

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DISCUSSION Reorganization of terminator DNA upon RTP binding A theoretical study of the relationship between nucleic acid conformation and the resulting circular dichroism found a strong linear correlation between the magnitude of the CD signal at 275 nm and the helix winding angle and twist (36), i.e. as the winding angle increases the twist decreases, resulting in an increase in ellipticity. We know of only three atomic resolution structures that have been obtained for protein–DNA complexes in which binding of the protein to the DNA causes a substantial increase in intensity of the longwave CD signal: GCN4-ATF–CREB (31,37), Lac repressor headpiece–half operator (38–40) and trp repressor– trpO operator (28,41–43); in all cases, as expected on the basis of the CD data, the structures show that the DNA is underwound in the protein–DNA complex. Interestingly, the increases in ellipticity observed for these complexes (41–60%) and the DNA bending angle (15–20) are similar to those observed in the current work for RTP binding to a single B site (66% and 40 respectively). Thus, the increased ellipticity observed at 275 nm when RTP binds to the B site can be interpreted as an underwinding of B site DNA due to a decrease in twist angle. This can be crudely interpreted as B site DNA becoming more ‘A-like’ upon binding RTP, since B- and A-form DNA have average twists of 36 (10 bp/turn) and 33 (11 bp turn) respectively (44). Interpretation of the circular permutation assays described herein as simply reflecting DNA bending shows that TerI DNA is bent by ∼40 in the RTP–TerI binary complex and by ∼60 in the RTP2–TerI ternary complex. While in theory such bending could be achieved by either a roll or tilt between sequential base pairs, analysis of atomic resolution protein–DNA structures shows that such bends are always produced by alterations in roll rather than tilt (45). An increase in roll angle can, like the decrease in twist discussed above, be crudely interpreted as TerI DNA becoming more ‘A-like’ upon binding RTP (A- and B-form DNA have average roll angles of +20 and 0 respectively; see 44). Low twist and positive roll are typical of base pair steps with ‘low twist profile’ (46). Thus it would appear that binding of a single RTP dimer to the B site of TerI causes underwinding and bending of the DNA, which will serve to widen the minor groove and compress and deepen the major groove. It is possible that these conformational changes could affect the A site and be responsible for the cooperative binding of a second dimer at this site (10). It is significant that binding of the second dimer to the A site causes additional bending of TerI DNA. It has recently been suggested, on the basis of studies with symmetrical terminators (19) and experimentally determined RTP–DNA contacts (13), that in forming the ternary complex, the centres of the RTP dimers in the A and B sites are positioned on almost opposite faces of B-form DNA (19). If the binding of an RTP dimer to the B site induces a simple bend and if this applied also when a second dimer filled the A site, one might have expected to see a reduction in the overall bend angle in the absence of other effects. The apparent increase in bend angle may argue for a significant rearrangement of the ternary terminator complex which places the two dimers on closer faces of the DNA, thereby promoting protein–protein interactions. Alternatively, it may argue for something other than a simple bend in the DNA when both dimers are bound. It is possible that circular permutation assays of RTP

binding to ‘terminator’ constructs with altered spacing between two strong binding sites (so-called ‘phasing’ experiments; see 34) might provide more definitive information on the orientation of the two RTP dimers in the ternary complex. Models of replication fork arrest by the RTP–terminator ternary complex To date, the favoured model for generation of an RTP–terminator complex with polar fork arrest activity has evoked asymmetry in the ternary complex as a result of the different contacts between RTP and DNA at the A and B sites (which have substantially different sequences), modulated crucially by interactions between the RTP dimers bound at the two overlapping sites (19). We shall refer to this as the ‘induced conformational change’ (ICC) model, which is illustrated schematically in Figure 6A. In this model, RTP binds to the high affinity B site and, while there may be induced rearrangements in the DNA and (to a lesser extent) the protein (as indicated in the current work), the final conformation of the binary complex is unsuitable for impeding progression of the replicative helicase. However, it is proposed that subsequent binding of a second dimer to the A site causes further rearrangements (as evidenced by the increase in the DNA bending angle seen in the current study), possibly via protein–protein contacts between the two bound RTP dimers, to yield a unique asymmetrical ternary complex. The asymmetry is such that only the B site complex is appropriately configured for impeding the helicase. It should be pointed out that the ICC model does not necessitate a specific molecular interaction between RTP in the functional terminator complex and helicase. It intends to imply an inhibition of helicase unwinding by an, as yet, unexplained mechanism. The fact that the RTP–TerI system functions in E.coli (47,48), despite the apparent lack of homology between RTP and the E.coli terminator protein (Tus), argues against specific RTP–helicase interactions. However, Bastia and co-workers (49,50) have presented evidence in favour of a specific interaction between the replicative helicase and a particular surface of RTP. Figure 6B introduces a new model for polar fork arrest in B.subtilis which we will refer to as the ‘differential binding affinity’ (DBA) model. In this model, polar fork arrest is a consequence of the different affinities of RTP for the A and B sites of the terminator DNA. RTP is known to bind tightly to the high affinity B site with a macroscopic Kd of ∼10–11 M (10). However, this binding is no tighter than that between various repressor proteins and their operator sequences and these proteins are readily displaced by helicase during DNA replication; hence, binding of a single RTP dimer to the B site is insufficient to cause replication fork arrest. However, binding of RTP to the B site causes rearrangements in the terminator DNA (as seen in the current study), which presumably facilitates the cooperative binding of a second dimer at the A site. In the DBA model, binding of the second RTP dimer to the A site causes a further rearrangement of the terminator complex such that the B site dimer becomes more strongly bound. We suggest two possibilities that might explain this. The first and most obvious possibility is that the two bound dimers make protein–protein contacts in the ternary complex. The affinity of each dimer for the ternary complex is now enhanced as a result of their affinity for each other. The other possibility is that binding of a second RTP dimer to the A site causes an additional rearrangement of the DNA that enhances the affinity of the B site


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Figure 6. Highly schematic models for the generation of RTP–terminator complexes with polar fork arrest activity. The DNA terminator sequence is shown as either horizontal or bent double lines. The extent of the A and B sites are indicated by thin lines at the top of each model. The central trinucleotide (TAT) of the 29 bp minimal TerI is shown as a hatched box and the filled circles represent the centres of the flanking 13 bp A and B site regions; RTP is represented as shaded boxes of various shapes (conformations). (A) In the induced conformational change (ICC) model, the first RTP dimer binds at the B site and remodels the DNA (shown schematically as a slight bend) in preparation for cooperative binding of a second dimer at the A site. There may be minor conformational changes in the protein upon binding DNA. Binding of the second dimer causes additional remodelling of the DNA, possibly mediated by protein–protein interactions, which in turn alters the conformation of the RTP dimer–B site complex (shown simplistically as a shape change in RTP, although the DNA conformation could also be altered) such that it is now appropriately configured for impeding the replicative helicase. The RTP dimer bound at the A site has a different conformation as a consequence of different RTP–DNA contacts (see Fig. 1); this conformation cannot impede replication forks approaching from the A site end of the terminator complex. (B) In the differential binding affinity model, the first dimer binds with relatively high affinity to the B site, but the affinity (the extent of which is schematically indicated by arrows, with thick arrows representing lower affinities and hence a greater likelihood of displacement) is still not sufficiently high to impede the replicative helicase. As in the ICC model, binding of the first dimer to the B site remodels the DNA for cooperative binding of the second dimer at the A site. This results in the formation of a ternary complex, with resulting conformational changes in the DNA and possibly the protein. The affinity of the B site dimer for the ternary complex is now markedly increased (indicated by the thin arrow), possibly, though not necessarily, as a result of protein–protein interactions. A replication fork approaching from the B site end is unable to dissociate the B site dimer, as to do so would require complete unravelling of the RTP–terminator complex. On the other hand, the affinity for the A site dimer is markedly lower so that it can be peeled off the DNA by a replication fork approaching from that end. This markedly decreases the DNA binding affinity of the RTP dimer bound at the B site, thus enabling helicase to dissociate it from the DNA once the A site dimer has been displaced.

dimer for terminator DNA. Regardless of which possibility is preferred, the net result is an enhanced affinity of the RTP dimer for the B site and a substantially weaker affinity for the RTP dimer bound at the A site. Consequently, helicase approaching from the B site end of TerI cannot move through the terminator, as it would have to unravel the entire ternary complex, which is not possible because of the now significantly increased affinity of the B site dimer. On the other hand, helicase approaching from the other direction is able to displace the more weakly bound A site dimer, which in turn removes the binding energy necessary to enhance the binding affinity of the B site dimer; thus, the Kd of the B site dimer returns to its ‘normal’ value, enabling helicase to displace it. Hence, in the DBA model, the second A site dimer acts as an ‘affinity lock’ for the dimer that is already bound at the B site. The effect of DNA terminator complexes on transcription It is worth noting that the DBA model might explain why the RTP–terminator complex blocks RNA chain elongation by T7, SP6 and E.coli RNA polymerases with the same polarity as its contrahelicase activity (49). The DBA model predicts that the RTP–terminator complex will have similar effects on any protein

or protein complex (such as helicase or RNA polymerase) that requires DNA unwinding to precede its translocation along the DNA. Of course, this raises the question of why RTP can block the activity of replicative helicases but appears unable to inhibit those involved in DNA repair and conjugal DNA transfer, including Rep helicase, helicase I and helicase II of E.coli (48,51). However, one must be cautious in extending the results of such in vitro studies to the in vivo situation. For example, it has recently been demonstrated that an A173V mutant of Tus was only a very weak inhibitor of the DnaB replicative helicase in in vitro assays but it halted DNA replication in vivo at 75% of the efficiency of wild-type Tus (52). Of course, it is possible that the blocks to replication and transcription by the RTP–terminator complex occur via quite different molecular mechanisms. For example, the report that a specific point mutant of RTP has been constructed which fails to block replication forks but still imposes a polar block on RNA chain elongation (49) argues in favour of specific but different protein–protein or protein–DNA interactions in the inhibition of DNA replication and transcription. In this respect, it is worth noting that there is a statistically significant homology between RTP and the β′ subunit of the family of DNA-directed RNA

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polymerases, including that of E.coli. Residues 12–77 of RTP (GenBank accession no. S01271) align with residues 217–281 of the β′ subunit of E.coli RNA polymerase (GenBank accession no. RNECC) with 12 identities and 18 conservative substitutions, giving an overall homology of 45%; there is a single residue gap in the E.coli sequence corresponding to residue 34 of RTP. This may indicate that RTP is capable of displacing the β′ subunit from the core RNA polymerase complex or that it can bind some critical factor that is normally attached to the β′ subunit. Alternatively, this homology, which includes most of the winged-helix DNA binding motif of RTP (only the final β3 strand is missing: see 12,17), may simply indicate that RTP and the β′ subunit of the family of prokaryotic DNA-directed RNA polymerases bind DNA in a similar fashion. It should be stressed that while the DBA model can adequately explain the polar fork arrest activity of the RTP–terminator ternary complex without invoking specific RTP–helicase interactions, it does not rule out such a possibility. Furthermore, the model is not applicable to the E.coli terminator complex, where a single monomer binds to a smaller DNA terminator sequence; the DBA model requires two terminator proteins to bind with differential affinities in order to generate polarity of action. In the case of the Tus–terminator binary complex, evidence against the ‘molecular clamp’ hypothesis has already been described in favour of specific interactions between the terminator complex and DnaB helicase (52,53). However, given the lack of sequence homology between Tus and RTP and between their DNA terminator binding sites, as well as the formation of binary terminator complex in the case of Tus versus a ternary complex in the case of RTP, it is possible that the molecular mechanism of action of Tus and RTP are quite different. Indeed, the distinct lack of structural homology between Tus and RTP (8,9,12) argues in favour of significant differences between their mechanisms of achieving polar fork arrest. ACKNOWLEDGEMENTS This work was supported by research grants to G.F.K. and R.G.W. from the Australian Research Council and by the award of a University of Sydney Postgraduate Scholarship, an Australian Postgraduate Research Award and a University of Auckland William Georgetti Scholarship to A.V.K. We would like to thank Dr Tony Day (Department of Biochemistry, University of Oxford, UK) for alerting us to the homology between RTP and the family of DNA-directed RNA polymerases. REFERENCES 1 Hill,T.M. (1992) Annu. Rev. Microbiol., 46, 603–633. 2 Yoshikawa,H. and Wake,R.G. (1993) In Sonenshein,A.L., Hoch,J.A. and Losick,R. (eds), Bacillus subtilis and Other Gram-positive Bacteria: Biochemistry, Physiology and Molecular Genetics. American Society for Microbiology, Washington, DC, pp. 507–528. 3 Lee,E.H., Kornberg,A., Hidaka,M., Kobayashi,T. and Horiuchi,T. (1989) Proc. Natl. Acad. Sci. USA, 86, 9104–9108. 4 Khatri,G.S., MacAllistair,T., Sista,P.R. and Bastia,D. (1989) Cell, 59, 667–674. 5 Hill,T.M. (1996) In Neidhardt,F.C. (ed.), Escherichai coli and Salmonella—Cellular and Molecular Biology. American Society for Microbiology, Washington, DC, pp. 1602–1614. 6 Franks,A.H., Griffths,A.A. and Wake,R.G. (1995) Mol. Microbiol., 17, 13–23. 7 Baker,T.A. (1995) Cell, 80, 521–524. 8 Kamada,K., Horiuchi,T., Ohsumi,K., Shimamoto,N. and Morikawa,K. (1996) Nature, 383, 598–603.

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