Structural basis of the 3′-end recognition of a leading strand in stalled replication forks by PriA
2007; Springer Nature; Volume: 26; Issue: 10 Linguagem: Inglês
10.1038/sj.emboj.7601697
ISSN1460-2075
AutoresKaori Sasaki, Toyoyuki Ose, Naoaki Okamoto, Katsumi Maenaka, Taku Tanaka, Hisao Masai, Mihoko Saito, Tsuyoshi Shirai, Daisuke Kohda,
Tópico(s)CRISPR and Genetic Engineering
ResumoArticle26 April 2007free access Structural basis of the 3′-end recognition of a leading strand in stalled replication forks by PriA Kaori Sasaki Kaori Sasaki Division of Structural Biology, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan Search for more papers by this author Toyoyuki Ose Toyoyuki Ose Division of Structural Biology, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan Search for more papers by this author Naoaki Okamoto Naoaki Okamoto Olympus Corp., Tokyo, Japan Search for more papers by this author Katsumi Maenaka Katsumi Maenaka Division of Structural Biology, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan Search for more papers by this author Taku Tanaka Taku Tanaka Genome Dynamics Project, Tokyo Metropolitan Institute of Medical Science, Tokyo, Japan Search for more papers by this author Hisao Masai Hisao Masai Genome Dynamics Project, Tokyo Metropolitan Institute of Medical Science, Tokyo, Japan Search for more papers by this author Mihoko Saito Mihoko Saito Department of Bioscience, Nagahama Institute of Bioscience and Technology, and JST-BIRD, Siga, Japan Search for more papers by this author Tsuyoshi Shirai Tsuyoshi Shirai Department of Bioscience, Nagahama Institute of Bioscience and Technology, and JST-BIRD, Siga, Japan Search for more papers by this author Daisuke Kohda Corresponding Author Daisuke Kohda Division of Structural Biology, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan Search for more papers by this author Kaori Sasaki Kaori Sasaki Division of Structural Biology, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan Search for more papers by this author Toyoyuki Ose Toyoyuki Ose Division of Structural Biology, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan Search for more papers by this author Naoaki Okamoto Naoaki Okamoto Olympus Corp., Tokyo, Japan Search for more papers by this author Katsumi Maenaka Katsumi Maenaka Division of Structural Biology, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan Search for more papers by this author Taku Tanaka Taku Tanaka Genome Dynamics Project, Tokyo Metropolitan Institute of Medical Science, Tokyo, Japan Search for more papers by this author Hisao Masai Hisao Masai Genome Dynamics Project, Tokyo Metropolitan Institute of Medical Science, Tokyo, Japan Search for more papers by this author Mihoko Saito Mihoko Saito Department of Bioscience, Nagahama Institute of Bioscience and Technology, and JST-BIRD, Siga, Japan Search for more papers by this author Tsuyoshi Shirai Tsuyoshi Shirai Department of Bioscience, Nagahama Institute of Bioscience and Technology, and JST-BIRD, Siga, Japan Search for more papers by this author Daisuke Kohda Corresponding Author Daisuke Kohda Division of Structural Biology, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan Search for more papers by this author Author Information Kaori Sasaki1, Toyoyuki Ose1, Naoaki Okamoto2, Katsumi Maenaka1, Taku Tanaka3, Hisao Masai3, Mihoko Saito4, Tsuyoshi Shirai4 and Daisuke Kohda 1 1Division of Structural Biology, Medical Institute of Bioregulation, Kyushu University, Fukuoka, Japan 2Olympus Corp., Tokyo, Japan 3Genome Dynamics Project, Tokyo Metropolitan Institute of Medical Science, Tokyo, Japan 4Department of Bioscience, Nagahama Institute of Bioscience and Technology, and JST-BIRD, Siga, Japan *Corresponding author. Division of Structural Biology, Medical Institute of Bioregulation, Kyushu University, Maidashi 3-1-1, Higashi-ku, Fukuoka 812-8582, Japan. Tel.:+81 92 642 6968; Fax: +81 92 642 6764; E-mail: [email protected] The EMBO Journal (2007)26:2584-2593https://doi.org/10.1038/sj.emboj.7601697 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info In eubacteria, PriA helicase detects the stalled DNA replication forks. This critical role of PriA is ascribed to its ability to bind to the 3′ end of a nascent leading DNA strand in the stalled replication forks. The crystal structures in complexes with oligonucleotides and the combination of fluorescence correlation spectroscopy and mutagenesis reveal that the N-terminal domain of PriA possesses a binding pocket for the 3′-terminal nucleotide residue of DNA. The interaction with the deoxyribose 3′-OH is essential for the 3′-terminal recognition. In contrast, the direct interaction with 3′-end nucleobase is unexpected, considering the same affinity for oligonucleotides carrying the four bases at the 3′ end. Thus, the N-terminal domain of PriA recognizes the 3′-end base in a base-non-selective manner, in addition to the deoxyribose and 5′-side phosphodiester group, of the 3′-terminal nucleotide to acquire both sufficient affinity and non-selectivity to find all of the stalled replication forks generated during DNA duplication. This unique feature is prerequisite for the proper positioning of the helicase domain of PriA on the unreplicated double-stranded DNA. Introduction When cells are stressed by ultraviolet irradiation or other damaging treatments, DNA replication forks often encounter template DNA lesions, which can stall the progression of the replication forks (Kowalczykowski, 2000). Processing of the stalled replication forks by recombination proteins, including RecA, RecG, and other Rec and Ruv proteins, is necessary in Escherichia coli cells to form intermediate DNA structures such as a chicken-foot fork structure and a D-loop (displacement loop) structure (Kowalczykowski, 2000). These branched DNA structures are the target of two proteins, PriA (McGlynn et al, 1997; Liu and Marians, 1999; Marians, 2000; Xu and Marians, 2003) and PriC (North and Nakai, 2005; Heller and Marians, 2005a). The two proteins ultimately load the DnaB replicative helicase and initiate the reassembly of a replisome to continue the duplication of genomic DNA. PriA was originally identified as a protein essential for the replication of the small, single-stranded DNA (ssDNA) of the phage ϕX174 (Shlomai and Kornberg, 1980). PriA functions as a scaffold that recruits other proteins, including PriB, PriC, DnaT, DnaC, DnaB, and DnaG. The protein cluster is collectively referred to as the ϕX174-type primosome. Previous studies clarified that the mechanism and the order of the primosome assembly (Ng and Marians, 1996; Jones and Nakai, 1997; Liu and Marians, 1999). In addition to the phage replication, PriA is necessary for the induction of RecA-dependent stable DNA replication (Masai et al, 1994), normal cell proliferation, and resistance to various genotoxic agents, including UV and mitomycin C (Lee and Kornberg, 1991; Kogoma et al, 1996). PriA contains a DEXH-type DNA 3′ → 5′ helicase domain in the C-terminal region (Ouzounis and Blencowe, 1991). Despite the conservation of the helicase motifs among eubacteria, the helicase activity of the C-terminal domain is unexpectedly dispensable for most of the PriA functions (Zavitz and Marians, 1992; Liu et al, 1999), with the exception of its requirement for full-level DNA synthesis in recombination-dependent modes of DNA replication (Tanaka et al, 2003). Recently, the PriA helicase activity was reported to prevent RecA from provoking unnecessary recombination during replication fork restarting (Mahdi et al, 2006), and to unwind the nascent lagging strand DNA to help PriC load the DnaB helicase onto stalled replication forks where there is a gap in the nascent leading strand (Heller and Marians, 2005b), although another helicase, Rep, can substitute for the function of the PriA helicase in both cases. Replication fork reassembly requires the recognition of a specific DNA structure by PriA. PriA reportedly binds to a D-loop-like structure by recognizing a bend at the three-strand junction, and to duplex DNA with a protruding 3′ single strand (McGlynn et al, 1997; Nurse et al, 1999; Chen et al, 2004), but the entity that PriA recognizes was unclear. We discovered that the N-terminal 181-residue domain of E. coli PriA specifically bound to a D-loop structure and an A-fork structure in gel shift assays (Tanaka et al, 2002; Mizukoshi et al, 2003), and to a ssDNA in a surface plasmon resonance (SPR) analysis (Mizukoshi et al, 2003). In both experiments, the phosphorylation of the 3′-hydroxyl group of the 3′-terminal nucleotide residue decreased the binding substantially, suggesting that the N-terminal domain of PriA recognizes an unmodified 3′ end of ssDNA. It is reasonable that PriA monitors the occurrence of an unusual 3′ end of DNA as a landmark to detect the stalling of chromosomal DNA replication. Recently, we showed that the presence of a free 3′ end at the branch point of an arrested fork structure and the 3′-end-binding ability of PriA were both required for the stabilization of the arrested fork structure (Tanaka and Masai, 2006). The binding of the N-terminal domain to the 3′ end of the leading DNA strand is required to place the helicase domain in an 'unwinding-deficient' orientation. Otherwise, the helicase destabilizes the fork structure and the repair of the arrested replication fork is aborted. In the present study, we analyzed the interaction between the N-terminal 105-residue domain of PriA and oligonucleotides carrying different bases at the 3′ end by fluorescence correlation spectroscopy (FCS) and crystallography. The results clearly indicated that PriA interacts with the base, deoxyribose, and 5′-side phosphodiester group of the 3′-terminal nucleotide residue. The deoxyribose moiety, in particular, the 3′-OH group is recognized by a network of polar interactions. By contrast, the direct recognition of the base moiety was really unexpected, considering the nearly equal affinity of PriA for ssDNA, irrespective of the base at the 3′ end. After a systematic mutagenesis study, we concluded that E. coli PriA utilizes a unique amino acid–nucleotide interaction mode to acquire sufficient affinity for the 3′ end of DNA, in a manner that neither discriminates the 3′-terminal base nor disturbs the Watson–Crick base pairing. The unique 3′-terminal nucleotide recognition is prerequisite for the proper sensing of the stalled replication forks by the PriA protein. Results The N-terminal 181-residue domain of PriA consists of two subdomains We prepared the N-terminal 181-residue fragment of E. coli PriA and measured the NMR spectrum (Supplementary Figure 1A). The good dispersion of the cross peaks in the [1H,15N]HSQC spectra indicates its stable tertiary structure, that is, the 181-residue fragment is a structural domain. The fragment 1–181 was cleaved after Arg105 and Arg108 by a limited proteolysis with trypsin (data not shown). We made two new constructs, 1–105 and 109–181, and found that both fragments had a stable fold by NMR (Supplementary Figure 1B and C). Thus, the fragment 1–181 consists of two subdomains, residues 1–105 and residues 109–181. The NMR spectrum of PriA[1–181] roughly equals the sum of the two spectra of PriA[1–105] and of PriA[109–181], suggesting the independency of the two domains. The N-terminal domain of PriA recognizes the 3′-terminal nucleotide residue The interaction of the N-terminal 105-residue domain of E. coli PriA (PriA[1–105]) with oligodeoxyribonucleotides was analyzed using FCS. FCS is a spectroscopic technique for studying molecular interactions in solution (Rigler, 1995). FCS monitors stochastic spontaneous fluctuations of fluorescently labeled particles, due to their entrance into and exit from a defined, small volume irradiated by a focused laser beam. The correlation function analysis of the fluorescent intensity provides the diffusion time, an average time for the particle to cross the small volume. The diffusion time is dependent on the mass of the particle, and thus the increase in the diffusion time of a fluorescently labeled molecule indicates an interaction with an added protein. First, we showed the specific recognition of the 3′-terminal nucleotide residue by PriA[1–105] (Figure 1A). As the PriA[1–105] protein was added to 5′TAMRA-d(A8) (octadeoxyadenylate with a terminal 5′ fluorophore and an unmodified 3′-hydroxyl group), a dose-dependent increase in the diffusion time was observed, due to complex formation. The curve fitting revealed a dissociation constant of about 25 μM. By contrast, neither 5′TAMRA-d(A8)p (terminal 3′ phosphate) nor d(A8)-3′TAMRA (terminal 3′ fluorophore) increased the diffusion time at all, indicating that the modification of the 3′-hydroxyl group by a bulky group completely blocks the interaction of PriA[1–105] and the oligonucleotide. The longer version of the N-terminal fragment, PriA[1–181], and the full-length PriA[1–732] bound to the 5′TAMRA-d(A8) with higher affinities of 7 and 3 μM, respectively, but the helicase domain of PriA[194–732] did not (Figure 1B). We further analyzed the role of the 3′-hydroxyl group in the recognition by changing the 3′-terminal deoxyribose to a dideoxyribose and prepared 5′TAMRA-d(A7)ddC. This oligonucleotide ends with a dideoxycytidine, and thus lacks hydroxyl groups at the 3′ end. The binding was abolished (Figure 1C), indicating that a hydrogen bond involving the 3′-hydroxyl group is essential. Figure 1.FCS analyses of PriA binding to TAMRA-labeled oligonucleotides. (A) Plots of FCS diffusion time against protein concentration. The points are connected by solid lines to aid in visualization. The N-terminal domain of PriA (PriA[1–105]) was mixed with 5′TAMRA-d(A8), 5′TAMRA-d(A8)p, and d(A8)-3′TAMRA. (B) Same as in (A) for the full-length PriA (PriA[1–732]), the longer version of the N-terminal domain (PriA[1–181]) and the helicase domain (PriA[194–732]) mixed with 5′TAMRA-d(A8). (C) Effect of the deletion of the 3′-OH group of the 3′-terminal deoxyribose moiety on binding. PriA[1–105] was mixed at a final concentration of 100 μM with 5′TAMRA-d(A7)ddC (dideoxycytidine at the 3′ end). 5′TAMRA-d(A7C) was used as a positive control and 5′TAMRA-d(A7C)p was a negative control. The corrected diffusion time (marked by *) was calculated by subtracting the value at zero protein concentration. (D) Effect of the oligonucleotide length on binding. PriA[1–105] was mixed at a final concentration of 250 μM with 5′TAMRA-d(An), where the number of residues, n, varied from 2 to 8. (E) Base preference at the 3′ end of oligonucleotides. PriA[1–105] was mixed at a final concentration of 250 μM with 5′TAMRA-d(A7N), where N is A, C, G, or T. 5′TAMRA-d(A8)p was used as a negative control. (F) Same as in (E) for the full-length PriA[1–732] at a final concentration of 100 μM. Download figure Download PowerPoint Then, we analyzed the interaction with oligonucleotides of differing lengths, ranging from 2 to 8 nt (Figure 1D). The optimal length was around 4 nt, but the shortest, 2 nt, can still bind to PriA[1–105] with comparable affinity. This result strengthens the notion that PriA[1–105] mainly recognizes the 3′-terminal nucleotide of DNA, and the other parts of the DNA may assist in the binding. Finally, we analyzed the interaction with oligonucleotides bearing different bases at the 3′ end (Figure 1E). We prepared 5′TAMRA-d(A7C), 5′TAMRA-d(A7G), and 5′TAMRA-d(A7T) in addition to 5′TAMRA-d(A8). PriA[1–105] bound to the four oligonucleotides with almost the same affinity. This intriguing result is not an artifact of domain excision, as the full-length PriA had the same base-non-selectivity (Figure 1F). Structure determination and overview The structure of PriA[1–105] in the absence of a ligand was determined using the multiwavelength anomalous diffraction (MAD) method to a resolution of 2.5 Å (Table I). The asymmetric unit in the crystal contains four PriA[1–105] molecules. Overall, the four PriA[1–105] structures in the crystal are very similar, with Cα r.m.s.d. values in the range of 0.8 to 1.3 Å. In the crystal, two PriA[1–105] molecules exchange their N-terminal seven amino-acid residues and form an intertwined dimer (Supplementary Figure 2), and thus there are two independent, intertwined dimers in the asymmetric unit. The PriA[1–105] protein also forms a stable dimer in solution, as demonstrated by gel filtration and analytical ultracentrifugation (data not shown). SPR and NMR analyses in the previous study (Mizukoshi et al, 2003), and FCS data (Figure 1) in the present study clearly showed that dimer formation by PriA[1–105] in solution does not interfere with the 3′-terminal nucleotide binding of the oligonucleotides, and that the binding properties of PriA[1–105] are similar to those of the full-length PriA. Table 1. Refinement statistics Ligand Free PriA[1–105] PriA[1–105] complex with d(AA) d(AC) d(AG) d(AT) d(CCC) Resolution range (Å) 20.0–2.52 20.0–3.30 20.0–3.20 20.0–3.35 20.0–3.15 20.0–3.0 No. reflections (work/test) 18 496/1418 8466/633 8987/685 8152/646 9894/763 11 132/859 Rwork/Rfree 24.9/29.5 26.0/30.9 24.7/30.2 28.1/33.3 27.1/31.8 23.8/29.3 Number of copies in AU Protein 4 4 4 4 4 4 Ligand 0 4 2 2 3 1 Number of atoms Protein 3296 3267 3296 3296 3296 3217 Ligand — 84 77 38 78 19 Water 56 — — — — — B-factors Protein 74.82 94.968 87.962 108.36 104.01 89.458 Ligand — 148.61 135.83 132.14 130.94 149.59 R.m.s. deviations Bond lengths (Å) 0.008 0.005 0.010 0.011 0.009 0.009 Bond angles (deg) 1.5 1.2 1.7 1.7 1.9 1.6 Ramachandran plot Most favored (%) 94.3 80.7 85.2 88.6 87.5 90.7 Additionally allowed (%) 4.5 19.2 13.6 11.4 12.5 7.0 Generously allowed (%) 1.1 0 1.1 0 0 2.3 Disallowed (%) 0 0 0 0 0 0 PDB entry 2D7E 2D7G 2DWL 2DWN 2DWM 2D7H Each data set was collected from a single crystal. Data collection statistics were reported previously (Sasaki et al, 2006). The functional form of the N-terminal 105-residue fragment should be monomeric because the full-length PriA is monomeric. We reconstituted the monomeric form of PriA[1–105] by swapping the coordinates of the N-terminal segments between the two intertwined chains and used it in the following discussion (Figure 2A). PriA[1–105] is composed of two antiparallel the β-sheets, a one-turn 310 helix, and two α-helices connected by a short loop. The secondary structure with the multiple alignment of the N-terminal domain of PriA from representative bacterial species is shown in Figure 2B. The architecture of the N-terminal domain of PriA appears to be novel, as no equivalent fold was found in a DALI search of the Protein Data Bank (Holm and Sander, 1998). Figure 2.Structure and multiple sequence alignment of the N-terminal domain of PriA. (A) The functional monomeric form of the N-terminal 105-residue domain of E. coli PriA was reconstituted from the intertwined dimeric form in crystal (Supplementary Figure 2). The swapped N-terminal seven-residue segment is drawn blue. The hinge region (Leu8–Arg14), for which the structure was not determined, is shown as an orange, broken curve. In a DALI search of the Protein Data Bank (Holm and Sander, 1998), the proteins showing the highest structural similarity had z scores in the range of 2.1 to 2.8 and r.m.s.d. values of 2.7–4.3 Å for 43–70 paired Cα atoms. (B) The secondary structure is shown above the alignment. β, α, T, and η represent a β-strand, an α-helix, a turn, and a 310 helix, respectively. Residues involved in the binding of the 3′-terminal nucleotide in E. coli PriA are labeled with blue triangles. Representative sequences were selected from each clade of the phylogeny of eubacteria (Supplementary Figure 6). The figure was produced with ESPript 2.2 (http://espript.ibcp.fr/ESPript/ESPript/index.php). Download figure Download PowerPoint 3′-Terminal nucleotide-binding site We carried out complex structure determinations of PriA[1–105] with four dinucleotides, d(AA), d(AC), d(AG), and d(AT) (Table I). The dinucleotides used were designed to have a constant adenine base at the 5′ end and a variable base at the 3′ end for analyzing the 3′-terminal base recognition by PriA[1–105]. In all of the complexes, the ligand binding did not change the overall protein conformation, with r.m.s.d. values in the range of 0.19 to 0.80 Å. Not all of the binding sites of PriA[1–105] were occupied by the ligands (Table I). In the crystal structure with d(AA), the electron density of the ligand existed in all of the four independent copies of the protein, but in the structures with the other dinucleotides, the densities of only two or three sites in the four copies were adequate for building nucleotide models, and the low occupancy and/or the high temperature factor prevented us from building a model at the remaining sites. The small variation in the affinity due to subtle differences in the crystallographic environments might explain the reason why not all binding sites are equally occupied. Even at the visible sites, the electron density corresponding to the constant 5′-terminal adenylate was poorly observed, and only that of the 3′-terminal nucleotide was positioned on the surface of the PriA[1–105] protein (Figure 3A). All of these observations were attributed to the fact that PriA[1–105] only recognizes the 3′-terminal nucleotide portion (base, deoxyribose, and 5′-side phosphodiester group), with a weak binding affinity. We also prepared a cocrystal of PriA[1–105] with a trinucleotide, d(CCC). As expected, just the density corresponding to the 3′-terminal cytidylate was visible. Comparisons of the structure with d(CCC) to those with d(AA) and d(AC) demonstrated that no serious biases arose in the binding mode from the different lengths of the ligands or the different base next to the 3′-terminal nucleotide. Figure 3.3′-Terminal nucleotide-binding site of PriA with bound ligands. (A) Stereoview of the electron density of the Fo–Fc omit map calculated from the final coordinates of the PriA[1–105]·d(AA) complex structure, minus the coordinates of the ligand, contoured at 3σ (green). The final refined structure is superimposed on the map. (B) Superimposed five complex structures with d(AA) (green), d(AC) (cyan), d(AG) (magenta), d(AT) (yellow), and d(CCC) (salmon). Only the 3′-terminal nucleotides were visible in the electron density maps and drawn. Amino-acid residues in contact with the 3′-terminal nucleotides are labeled. (C) Different view of the binding site, showing the interactions between the 3′-terminal base, Asp17, and the loop segment, Ser53–Glu54–Leu55. (D) Schematic representation of the interactions at the 3′-terminal nucleotide-binding site in the LIGPLOT style. The 5′-side phosphodiester group, the deoxyribose moiety, and the nucleobase of the 3′-terminal nucleotides are recognized by PriA. Amino-acid residues involved in hydrophobic contacts with the nucleotide are drawn schematically as brown spoked arcs. (E) Site of the 3′-nucleotide binding in the overall structure. Download figure Download PowerPoint The crystal structure of PriA[1–105] in complexes with oligonucleotides facilitated the identification of the residues that contact the 3′ end of the DNA. PriA[1–105] possesses a pocket, consisting of Phe16, Asp17, Tyr18, Gly37, Leu55, and Lys61, to accommodate the 3′-terminal nucleotide of an oligonucleotide (Figure 3B). In accordance, the backbone amide cross peaks of Asp17, Tyr18, and Gly37 were the most affected during the titration of an 8-nt oligonucleotide in the previous NMR study (Mizukoshi et al, 2003). Figure 3D summarizes the binding mode schematically. The oxygen atoms of the 5′-side phosphodiester group are liganded to the Gly37 backbone N and the Lys61 side-chain N. The deoxyribose moiety is anchored to the bottom of the binding pocket, by a stacking interaction on Tyr18, and in contact with the Lys61 backbone N and the Asp17 backbone O. Finally, the base is situated in a cleft formed by Phe16, Asp17, and Leu55 and adopts the anti conformation about the glycosyl bond. The base is sandwiched between Phe16 and Leu55, whereas the minor groove face of the base interacts with Asp17 (Figure 3C). Note that the major and minor groove faces of the bases are defined as the directions accessible from the outside in the duplex DNA structure (see Figure 1 of the literature; Luscombe et al, 2001). Effects of point mutations on binding affinity and specificity We used FCS analysis to monitor the binding of PriA[1–105] mutants to octadeoxyadenylate with a 5′-end fluorophore. We observed a pronounced reduction in the binding affinities by the alanine-scanning mutations of the amino-acid residues in contact with the 3′-terminal nucleotide: Phe16, Asp17, Gly37, Leu55, and Lys61 (Figure 4). To check the structural integrity of the mutants, we measured 1D 1H NMR spectra (Supplementary Figure 3). The chemical shifts of the upfield shifted methyl peaks, which are highly sensitive to the tertiary fold, are unchanged upon mutations. The mutation of Phe36, next to Gly37, also impaired the binding. As a control, the mutation of Lys38 had little effect on the binding affinity. Notably, the mutation of Tyr18 was not tested because of inclusion body formation, suggesting the important role of Tyr18 in the PriA structure. Figure 4.FCS analyses of the binding of alanine-scanning mutants of PriA[1–105] to octadeoxyadenylate. Amino-acid residues that contact the 3′-terminal nucleotide in the crystal structures were substituted to an alanine. Wild type and alanine point mutants of PriA [1–105] were mixed at a final concentration of 500 μM with 5′TAMRA-d(A8). As a negative control, the wild-type PriA[1–105] was mixed with 5′TAMRA-d(A8)p. Download figure Download PowerPoint The crystal structures suggested that the backbone N and side-chain O of Asp17 are involved in the interaction with the 3′-terminal base. We generated six mutants of Asp17 that differ in charge and size, and monitored the binding specificity for the 3′-terminal base. We used four 5′TAMRA-labeled octanucleotides, each with one of the four nucleotides at the 3′ end, in the FCS analysis (Figure 5A). The substitution of Asp17 to any other amino acid always negatively affected the affinity. At the same time, the substitution of Asp17 makes PriA[1–105] have distinctive specificity for the 3′-terminal base. A subtle change, such as the extension of the length of the side chain of Asp17 by a methylene group (Asp → Glu) or the amidation of the carboxyl group (Asp → Asn), causes PriA[1–105] to have base specificity. Figure 5.FCS analyses of the binding of mutant PriA[1–105] to oligonucleotides bearing different 3′-terminal bases. (A) Point mutants with the substitution of Asp17 with various amino acids were mixed at final concentrations of 50–100 μM with 5′TAMRA-d(A7N), which possesses different bases, A, C, G, or T, at the 3′ end. (B) Same as in (A), except for point mutants with the substitution of Leu55. Download figure Download PowerPoint We also prepared mutations of Leu55, which hydrophobically contacts the 3′-terminal base (Figure 5B). The substitution of Leu55 to Val reduced the binding affinity moderately, and that to Ala showed a more pronounced reduction of affinity. The substitution to Glu abolished the binding (data not shown). Interestingly, no obvious differences in the affinity for the 3′-terminal base were generated by the Leu55 mutations, in sharp contrast to the case of the Asp17 mutations. Discussion The crystal structure of PriA[1–105] in complexes with di- and trinucleotides facilitated the identification of residues that contact the 3′ end of DNA (Figure 3). The electron density of the 3′-terminal nucleotides is of moderate quality. We consider that this is due to the inherent nature of the loose binding mode of PriA. To allow judging the relevance of the models, the ligand-unbiased Fo–Fc maps, calculated from the protein coordinates before the nucleotide coordinates were added during refinement, are shown in Supplementary Figure 4. In addition, the 2Fo–Fc maps and the Fo–Fc omit map, both calculated from the final coordinates, are shown in and Figure 3A and Supplementary Figure 4, respectively. In parallel with the crystal screening of PriA[1–105], we attempted to crystallize the PriA[1–181], which has three times higher affinity for the oligonucleotides, but we have not obtained any crystals, probably due to the flexibility between the two subdomains. Mechanism for the 3′-terminal nucleotide recognition The modification of the 3′-OH group of the deoxyribose by a bulky group, such as phosphate or the TAMRA dye, completely blocked the interaction (Figure 1A). The removal of the 3′-OH group also abolished the binding (Figure 1C). Thus, the size and hydrogen bonding at the 3′ position of deoxyribose are crucial for the 3′-end recognition by the PriA protein. The stacking interaction between the deoxyribose ring and the tyrosine residue at position 18 also appears important because Tyr18 is one of the small number of invariably conserved amino-acid residues in the PriA protein family (Figure 2B). PriA interacts with the 5′-side phosphodiester group using the side-chain amino group of a lysine residue and the backbone amide of a glycine residue, but few interactions exist with the 5′ neighboring nucleotide residue in all of the crystal structures determined (Figure 3). The extensive interactions of Asp17 and Lys61 with the 3′-terminal nucleotides make the 3′-terminal nucleotide-binding site of PriA[1–105] narrow and compact. In summary, PriA has a small pocket that accommodates the 3′-terminal nucleotide only. In fact, the extension of oligonucleotides enhanced the affinit
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