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2 The abbreviations used are:
RMrestriction modificationModmodificationResrestrictionMDRSmodification-dependent restriction system5mC5-methylcytosine5hmC5-hydroxymethylcytosine5ghmC5-glucosylhydroxymethylcytosineEcE. coliRMCmethylated binding site, where R is a purine and MC is a methylcytosineHpH. pyloriHpR1Δ136the N-terminal DNA-binding domain of H. pylori LlaJI.R1 proteinAtA. thalianaSeMetselenomethionineSADsingle-wavelength anomalous diffractionSECsize exclusion chromatographyMALSmulti-angle light scatteringEMSAelectrophoretic mobility shift assayNSP3nonstructural protein 3.
Restriction modification systems consist of an endonuclease that cleaves foreign DNA site-specifically and an associated methyltransferase that protects the corresponding target site in the host genome. Modification-dependent restriction systems, in contrast, specifically recognize and cleave methylated and/or glucosylated DNA. The LlaJI restriction system contains two 5-methylcytosine (5mC) methyltransferases (LlaJI.M1 and LlaJI.M2) and two restriction proteins (LlaJI.R1 and LlaJI.R2). LlaJI.R1 and LlaJI.R2 are homologs of McrB and McrC, respectively, which in Escherichia coli function together as a modification-dependent restriction complex specific for 5mC-containing DNA. Lactococcus lactis LlaJI.R1 binds DNA site-specifically, suggesting that the LlaJI system uses a different mode of substrate recognition. Here we present the structure of the N-terminal DNA-binding domain of Helicobacter pylori LlaJI.R1 at 1.97-Å resolution, which adopts a B3 domain fold. Structural comparison to B3 domains in plant transcription factors and other restriction enzymes identifies key recognition motifs responsible for site-specific DNA binding. Moreover, biochemistry and structural modeling provide a rationale for how H. pylori LlaJI.R1 may bind a target site that differs from the 5-bp sequence recognized by other LlaJI homologs and identify residues critical for this recognition activity. These findings underscore the inherent structural plasticity of B3 domains, allowing recognition of a variety of substrates using the same structural core.
). These systems consist of a restriction endonuclease and a methyltransferase, which provide the dual function of cleaving exogenous DNA site-specifically, whereas protecting the host genome via methylation of the corresponding recognition sequence (
). Three variants of RM systems, type I, type II, and type III, have been identified and differ in their structural composition and mechanism of restriction. Type I systems are multifunctional complexes containing separate restriction, methylation, and DNA-sequence recognition subunits. These machines require Mg2+ and ATP, catalyze both restriction and methylation, and cut DNA nonspecifically far from their recognition sites (
). Type II systems are the simplest, generally existing as dimers that carry out the recognition and restriction activities. These enzymes do not require ATP and have a separate, associated methyltransferase (
). Homodimeric type II restriction enzymes recognize DNA sequences that are symmetric, whereas those that are heterodimeric can bind asymmetric sequences (
). Type III systems contain separate modification (Mod) and restriction (Res) subunits that form homodimeric Mod2 and heterotetrameric Res2Mod2 complexes and catalyze both restriction and methylation in a Mg2+ and ATP-dependent manner (
). The McrB and McrC proteins together form a conserved, two-component restriction complex capable of long-range DNA translocation similar to type I and type III enzymes. Escherichia coli (Ec) McrB contains an N-terminal DNA-binding domain (
A mutational analysis of the PD.D/EXK motif suggests that McrC harbors the catalytic center for DNA cleavage by the GTP-dependent restriction enzyme McrBC from Escherichia coli.
). Biochemical studies suggest a model for DNA cleavage in which McrB and McrC assemble at two distant RMC sites (where R is a purine, and MC is a methylcytosine) and translocate in a manner that requires stimulated GTP hydrolysis (
The GTP-dependent restriction enzyme McrBC from Escherichia coli forms high-molecular mass complexes with DNA and produces a cleavage pattern with a characteristic 10-base pair repeat.
). Structural studies of McrB, MspJI, PvuRtsI, and AbaSI suggest type IV systems employ a generalized base-flipping mechanism for recognition of the modified DNA (
The LlaJI restriction cassette was first identified in Lactococcus lactis on the naturally occurring plasmid pNP40 and shown to confer resistance against common lactococcal phages (
). It consists of an operon encoding two 5mC-methyltransferases, LlaJI.M1 and LlaJI.M2, and two restriction proteins, LlaJI.R1 and LlaJI.R2, both of which are absolutely required for restriction activity in vivo (
). Although formally classified as a type II R/M system (REBASE enzyme number 10100, New England Biolabs), LlaJI.R1 and LlaJI.R2 share domain homology with McrB and McrC, respectively. R1 contains sequence motifs that identify its C-terminal portion as a GTP-specific AAA+ domain and R2 contains a conserved C-terminal PD-(D/E)XK endonuclease domain. These features suggest LlaJI enzymes function more like McrBC than other type II systems.
Unlike McrB, however, L. lactis LlaJI.R1 binds DNA site-specifically, recognizing the asymmetric 5′-GACGC-3′ sequence in one strand and 5′-GCGTC-3′ in the other strand (
). How LlaJI proteins recognize DNA site-specifically is unknown. Here we present the structure of the N-terminal DNA-binding domain of H. pylori LlaJI.R1 (HpR1Δ136) at 1.97-Å resolution, which adopts a B3 domain fold. Structural comparison to B3 domain-containing plant transcription factors and restriction endonucleases identifies the key recognition motifs responsible for site-specific DNA binding. Additional evidence from biochemistry and structural modeling argues that HpLlaJI.R1 binds a target site that differs from the 5-bp sequence recognized by the L. lactis and C. cellulovorans LlaJI homologs. Mutagenesis further identifies residues Arg-17 and Arg-60 as critical determinants of HpLlaJI.R1 DNA binding. Together, these findings underscore the inherent structural plasticity previously noted for B3 domains, which confers specificity to different sequences via the same structural core.
Results
Structure and topology of the HpLlaJI.R1 N-terminal domain
Although previous studies show LlaJI.R1 binds DNA site-specifically (
), the molecular means through which this is achieved remains unknown. Numerous attempts to purify either the full-length L. lactis LlaJI.R1 or its isolated N-terminal DNA-binding domain for structural studies were unsuccessful. Bioinformatics identifies various other species harboring the LlaJI operon, including H. pylori (Hp) (
). Computational analyses of these homologs by fold matching and structural prediction algorithms failed to identify a reliable template for modeling DNA interactions. To understand how the LlaJI.R1 binds DNA site-specifically, we therefore crystallized the N-terminal domain of HpLlaJI.R1 (HpR1Δ136) and determined its structure at 1.97 Å by selenium SAD phasing (
Figure 1Structure and topology of HpR1Δ136.A, crystal packing of HpR1Δ136. AB and CD dimers are labeled. B, cartoon representations of HpR1Δ136 in two orientations. Molecules A and B are colored green and blue, respectively. The asymmetric β7 strand is colored raspberry. C, topology diagram of HpR1Δ136. The core-fold of each monomer is shown in blue. The relative position and connectivity of the asymmetric β7 strand associated with molecules B and D is denoted by dashed outlines and colored in raspberry. The N- and C-arms are colored orange and cyan, respectively. D, 2Fo − Fc electron density (blue mesh) of the α4–β7 region in molecule B contoured to 1σ. E, superposition of HpR1Δ136 dimers AB and CD. AB dimer is colored green and blue, whereas the CD dimer is colored gray. β7 from molecules B and D are colored raspberry and gray, respectively.
HpR1Δ136 crystallizes in the space group P1 with four molecules (A–D) in the asymmetric unit organized as two homodimers packed end to end, with molecules A and B and molecules C and D pairing together (Fig. 1A). These dimers superimpose with an r.m.s. deviation of 0.589 Å. Each HpR1Δ136 monomer consists of a core six-stranded β sheet that folds into a pseudo-β barrel flanked on four separate edges by α helices (α1–α4) (Fig. 1, B and C). An additional β-strand (β7) inserts at the dimer interface and breaks the symmetry, adopting an antiparallel configuration with β1B/β1D and a parallel configuration with β1A/β1C (Fig. 1, B and C). Clear connectivity between β7 and α4 can be traced in molecule B (Fig. 1D). Structural superposition of the two asymmetric dimers suggests β7 is connected in the same manner in molecule D despite the lack of density for the α4–β7 loop (Fig. 1E). We observe no density for the corresponding β7 strands in either molecule A or molecule C.
β7 residues Leu-127 and Phe-129 interact with a hydrophobic cluster sandwiched between β1 and the amphipathic α2 helix in each monomer (Fig. 2A). Ile-24, His-27, and Phe-28 in α2 and Val-115, Leu-116, and Leu-119 in α4 provide additional stabilizing contacts across the dimer interface (Fig. 2, A and B). β7 insertion helps space these elements and prevent steric clashing that otherwise would occur. A total interaction surface of 800 Å2 is shared between the monomers. Size exclusion chromatography coupled to multi-angle light scattering (SEC-MALS) indicates HpR1Δ136 dimerizes in solution (Fig. 2C), suggesting the observed molecular organization in the crystal lattice is not simply a packing artifact. Deletion of β7 renders HpR1Δ136 insoluble. Point mutations at the dimer interface in α2 (I24N, H27E, F28E) and α4 (L116E and L119E) displayed similar insolubility phenotypes and could not be purified. Only the V115N mutant retained solubility and, like WT, forms stable dimers in solution when analyzed by SEC-MALS (Fig. 3A). These data support the notion that HpR1Δ136 dimerization is required for stability and biologically relevant.
Figure 2Critical structural contacts stabilizing the HpR1Δ136 dimer.A, zoomed view of β1, α2, and β7 interactions at the dimer interface. B, hydrophobic interactions between α4 helices across the dimer interface. C, SEC-MALS analysis indicates HpR1Δ136 exists as a dimer in solution. UV trace (black) and calculated molecular weight based on light scattering (blue) are shown. Dashed red lines denote the predicted molecular weight of an HpR1Δ136 monomer and dimer.
Figure 3SEC-MALS of HpR1Δ136 mutants. V115N is located at the dimer interface (see Fig. 2B), whereas H14A, R17A, P59A, and R60A are putative binding site mutations based on structural homology (see Fig. 6, D and E). UV trace (black) and calculated molecular weight based on light scattering (blue) are shown. Dashed red lines denote the predicted molecular weight of an HpR1Δ136 monomer and dimer.
) identifies several structural homologs of HpR1Δ136. These include the DNA-binding domain of the Arabidopsis thaliana (At) auxin-dependent transcription factor ARF1 (PDB code 4ldx, Z-score = 8.6, r.m.s. deviation = 3.0), the C-terminal fragment of the BfiI restriction endonuclease (PDB code 3zi5, Z-score = 9.7, r.m.s. deviation = 2.1), and the N-terminal fragment of the EcoRII restriction endonuclease (PDB code 3hqf, Z-score = 8.1, r.m.s. deviation = 2.7). Each is comprised of a B3 domain (
Structural insight into the specificity of the B3 DNA-binding domains provided by the co-crystal structure of the C-terminal fragment of BfiI restriction enzyme.
). Structural superposition confirms HpR1Δ136 similarly adopts a B3 domain-fold (Fig. 4A, Fig. S1). Importantly, this fold is structurally distinct from the analogous region in EcMcrB, which preferentially binds DNA containing methylated cytosines.
Figure 4HpR1Δ136 adopts a B3-fold for site-specific DNA recognition.A, superposition of HpR1Δ136 (monomer A, green) with the A. thaliana (At) ARF1 B3 domain (monomer A, pink; PDB code 4ldx) confirms similar structural fold. B, superposition of HpR1Δ136 (light blue) with the AtARF1A B3–DNA complex (gray; PDB code 4ldx) identifies the N-arm (orange and green) and C-arm (cyan and purple) regions that are necessary for target recognition. Structures are oriented looking down the helical axis of bound AtARF1 DNA. C, electrostatic surfaces of the individual HpR1Δ136 and AtARF1 B3 domains. Arrows indicate the N- and C-arms. Scale bar indicates electrostatic surface coloring from −3 KbT/ec to +3 KbT/ec. D, electrostatic surface of the HpR1Δ136 AB dimer in two orientations. N- and C-arms are labeled. The black circle indicates the location of Glu-131 in molecule B.
B3 domains contain two critical regions that confer DNA target site–specificity. These recognition motifs, termed the N-arm and C-arm, reside on opposite edges of the pseudo-barrel core and form a wrench-like structure that contacts the major groove (
Structural insight into the specificity of the B3 DNA-binding domains provided by the co-crystal structure of the C-terminal fragment of BfiI restriction enzyme.
). In this arrangement, the N-arm specifically associates with the 5′-half of the target site and the C-arm engages the 3′-half. Comparison to the DNA-bound AtARF1 structure identifies these key features within the HpR1Δ136 model: the N-arm encompasses the β1–β2 loop and the α1 helix and the C-arm localizes to the β3–β4 loop (Figs. 1C and 4B). Both proteins display a comparable electrostatic surface, with an extensive basic patch positioned between the N- and C-arms and coincident with the DNA-binding face of AtARF1 (Fig. 4C). This mode of substrate binding is conserved among DNA-bound B3 domain structures and is consistent with other HpR1Δ136 structural superpositions (Fig. S1). An exception is the B3 domain of NgoAVII, whose orientation on DNA is inverted such that the N-arm associates with the 3′-half of the target site and the C-arm with the 5′-half (
Structural insight into the specificity of the B3 DNA-binding domains provided by the co-crystal structure of the C-terminal fragment of BfiI restriction enzyme.
The Arabidopsis B3 domain protein VERNALIZATION1 (VRN1) is involved in processes essential for development, with structural and mutational studies revealing its DNA-binding surface.
). HpR1Δ136 similarly lacks these segments, suggesting it either evolved from a more simplified common ancestor or lost these segments over time due to a lack of selective pressure.
The putative DNA-binding surface of each HpR1Δ136 monomer faces away from the dimer interface, suggesting that HpR1Δ136 has the capacity to bind two DNA target sites simultaneously. The asymmetric orientation of the β7 strand, however, positions Glu-131 close to one of the binding sites and alters its surface charge potential in a manner that makes it less basic (Fig. 4D). This intrinsic difference would allow one monomer to bind more efficiently and could bias the arrangement of HpLlaJI on DNA.
All previously characterized B3 domains exist as monomers (
Structural insight into the specificity of the B3 DNA-binding domains provided by the co-crystal structure of the C-terminal fragment of BfiI restriction enzyme.
The Arabidopsis B3 domain protein VERNALIZATION1 (VRN1) is involved in processes essential for development, with structural and mutational studies revealing its DNA-binding surface.
). To understand what hinders dimerization in these contexts, we superimposed the coordinates of other B3 domains onto our HpR1Δ136 dimer and examined the orientation of secondary structure features relative to the dimer interface (Fig. 5). Although EcoRII and BfiI have structurally equivalent β-strands that partially align with β7, they also contain helical segments that sterically prevent two monomers from coming together (Fig. 5, A and B). The α1 helices of VRN1 and UbaLAI would similarly clash and block dimerization (Fig. 5, C and D). AtARF1, NgoAVII, RAV1, and At1g16640.1, in contrast, all lack a corresponding β7 strand (Fig. 5, E–H), suggesting monomers cannot be properly spaced to avoid collision. The stabilizing hydrophobic interactions provided by β7 would also be absent. These observations highlight the importance of secondary structure features in modulating the oligomeric state of B3 domains and will be useful for predicting interactions in other uncharacterized proteins that contain this conserved fold.
Figure 5Structural constraints of B3 domain dimerization. Molecule A (green) and molecule B (blue) of HpR1Δ136 are shown in each panel with the asymmetric β7 strand (raspberry) and flanking β1 strands emphasized to delineate the dimer interface. Coordinates from different B3 domains were superimposed with molecule A and the overlapping structural elements that spatially align with the dimer interface are labeled and highlighted. PDB codes are indicated for each structure. A, superposition with EcoRII (gray). B, superposition with BfiI (yellow). C, superposition with VRN1 (cyan). D, superposition with UbaLAI (light blue). E, superposition with AtARF1 (pink). F, superposition with NgoAVII (light orange). G, superposition with RAV1 (magenta). H, superposition with At1g16640.1 (orange).
). HpR1Δ136 shows weak affinity for DNA containing this sequence (Ll) when assessed by filter binding (Fig. 6A, blue). Scrambling the putative binding sequence (Llscr) has no effect on the affinity (Fig. 6A, green), suggesting this represents the basal level for nonspecific DNA binding by HpR1Δ136. EcMcrB, in contrast, preferentially binds 5mC-containing DNA (5mC) but does not bind a nonmethylated version of the same substrate (nm) under the same assay conditions (Fig. 6A, black versus red). EcMcrB similarly does not bind either the Ll or Llscr substrates (Fig. 6A, yellow and orange), underscoring how its binding depends on the presence of methylated cyotsines.
Figure 6Structural modeling of HpR1Δ136 substrate recognition.A, filter binding analysis of HpR1Δ136 (Hp) and full-length EcMcrB (Ec) interactions with different DNA substrates. Substrate abbreviations are as follows: 5mC, methylated EcMcrB-specific substrate; nm, nonmethylated EcMcrB-specific substrate; Ll, site-specific substrate containing the L. lactis LlaJI.R1 5′-GACGC-3′ target site sequence; Llscr, substrate with the L. lactis LlaJI.R1 target site sequence scrambled as a control. Sequences for each substrate can be found under “Experimental procedures.” Binding was performed at 30 °C for 10 min in a 30-μl reaction mixture containing 14.5 nm unlabeled DNA and 0.5 nm labeled DNA. Samples were filtered through KOH-treated nitrocellulose and binding was assessed by scintillation counting. B, relationship between C-arm length and target site length in previously determined B3 domain–DNA complexes. C, orientation of C-arm loops relative to DNA in various B3 domain homologs. DNA from the AtARF1 complex (PDB code 4ldx) is shown. C-arm coloring is labeled below along with corresponding PDB codes. D, key residues in AtARF1 DNA binding. E, residues predicted to be important for HpR1Δ136 DNA binding based on structural comparison. AtARF1 DNA modeled as in D. F, filter binding analysis of HpR1Δ136 mutants. Point mutations of predicted binding residues identified in D (H14A, red; R17A, orange; P59A, green; R60A, light blue; R17A/R60A; brown) were assessed for binding to the nm DNA substrate. Filter binding was carried out as described in A. The WT curve (purple) is the same as shown in A (Hp+nm).
Unexpectedly, HpR1Δ136 showed enhanced affinity for the E. coli-specific 5mC and nm substrates (Fig. 6A, light blue and purple) relative to Ll and Llscr substrates. We attribute this to subtle sequence differences as the binding is independent of methylation status. The 5mC and nm substrates likely contain sequence fragments that more closely mimic the preferred recognition site of HpR1.LlaJI, which is distinct from both EcMcrB and other LlaJI homologs.
Despite a common fold, B3 domains exhibit divergent sequence preferences. Previous structural and biochemical data show that the C-arm length can influence the length of the recognized target site (Fig. 6B). A longer C-arm is excluded from the major groove (Fig. 6C), decreasing the overall binding footprint and biasing recognition toward a 5-bp site (
Structural insight into the specificity of the B3 DNA-binding domains provided by the co-crystal structure of the C-terminal fragment of BfiI restriction enzyme.
). A shorter C-arm affords greater access to the DNA bases, which in some instances increases the number of specific contacts and extends the target site to six bases (
Structural insight into the specificity of the B3 DNA-binding domains provided by the co-crystal structure of the C-terminal fragment of BfiI restriction enzyme.
). The amino acid composition of the N- and C-arms ultimately dictates specificity, however, and thus some B3 domains with shorter C-arms still bind 5-bp sites (
). Structural superposition reveals a shorter C-arm in HpR1Δ136 (Fig. 6C).
In the absence of a DNA-bound complex and without explicit knowledge of the HpLlaJI target site, we used the AtARF1–DNA structure as a proxy to identify side chains that might contribute to specificity. The N-arm residue His-136 and C-arm residues Arg-181, Pro-184, and Arg-186 are critical for AtARF1 DNA binding (Fig. 6D). Structural modeling reveals similar residues in HpR1Δ136 (Fig. 6E), with His-14 and Arg-17 in the N-arm and Pro-59 and Arg-60 in the C-arm poised to provide base-specific contacts. Interestingly, Arg-17 is spatially oriented like Arg-181 in AtARF1, hinting that it would contact the 3′-half of the target site despite being localized in the N-arm.
To confirm the significance of our structural modeling, we mutated the predicted binding residues in HpR1Δ136 and assessed how each substitution affects interaction with the E. coli-specific nm DNA substrate via filter binding (Fig. 6F). H14A (red), R17A (orange), and R60A (light blue) mutations show a marked decrease in affinity for nm DNA versus wildtype (WT, purple), whereas P59A (green) shows less of an effect. The R17A/R60A double mutant (brown) completely abolishes binding. This finding was corroborated using electrophoretic mobility shift assays (EMSAs) to measure the association of HpR1Δ136 with digested, nonmethylated λ-phage DNA (Fig. 7). We observed a significant gel shift with WT HpR1Δ136. H14A and P59A show similar shifts in this assay, whereas the individual R17A and R60A substitutions produce a moderate reduction in binding. The R17A/R60A double mutant, however, significantly impairs binding (Fig. 7), similar to its effects on the nm DNA substrate in the filter-binding assay (Fig. 6F). All of these mutants form stable dimers in solution (Fig. 3B), arguing that their effects are not due to global structural perturbations. Together these data implicate Arg-17 and Arg-60 as critical determinants of HpR1Δ136 DNA binding.
Figure 7EMSA analysis of predicted HpR1Δ136-binding mutants. Binding was carried out at 25 °C for 30 min in a 16-μl reaction mixture containing 10 ng/μl of digested (BamHI/NdeI) nonmethylated λ-phage DNA, and increasing concentrations (0–100 μm) of each HpR1Δ136 construct. Gels were stained with SYBR® Green in 1× TAE overnight at 25 °C. Calculated sizes (bp) of the digested DNA products are noted on the left of each gel.
Structural insight into the specificity of the B3 DNA-binding domains provided by the co-crystal structure of the C-terminal fragment of BfiI restriction enzyme.
). Arg-177 and Lys-126 form these interactions in AtARF1 (Arg-81 and Lys-23 in EcoRII; Arg-272 and Lys-340 in BfiI; Lys-27 and Lys-82 in UbaLAI; Lys-212 and Lys-275 in R.NgoAVII). In HpR1Δ136, Arg-6 is poised to act on one strand, whereas Lys-50 could perform a similar function on the opposing strand. Lys-50 is positioned away from the modeled DNA backbone in the apo state and may be reoriented upon target recognition. Conformational rearrangements in the BfiI and R.NgoAVII B3 domains have previously been observed upon DNA binding (
Structural insight into the specificity of the B3 DNA-binding domains provided by the co-crystal structure of the C-terminal fragment of BfiI restriction enzyme.
Here we described the structure of the HpLlaJI.R1 DNA-binding domain and demonstrated that it adopts a B3 domain fold. B3 domains are prevalent among bacterial restriction endonucleases and plant transcription factors, where they function as site-specific DNA-binding modules (
Structural insight into the specificity of the B3 DNA-binding domains provided by the co-crystal structure of the C-terminal fragment of BfiI restriction enzyme.
). Previous crystallographic studies revealed that the N- and C-arms determine the specificity of each individual B3 domain and confer structural plasticity to the conserved core scaffold (
Structural insight into the specificity of the B3 DNA-binding domains provided by the co-crystal structure of the C-terminal fragment of BfiI restriction enzyme.
The Arabidopsis B3 domain protein VERNALIZATION1 (VRN1) is involved in processes essential for development, with structural and mutational studies revealing its DNA-binding surface.
). Our structural data and modeling identifies the N- and C-arms in HpR1Δ136 along with key residues that likely form direct contacts with the DNA backbone, clamp phosphates, and specific bases. HpR1Δ136 has weak affinity for DNA containing the asymmetric 5-bp site that other LlaJI homologs target (
) and a surprisingly stronger affinity for the EcMcrB-specific DNA substrates, regardless of their methylation status (Fig. 6A). We note that HpR1Δ136 contains a shorter C-arm and thus could potentially bind a 6-bp site. Although further studies are required to pinpoint the target site of HpLlaJI.R1, our findings offer a general model for site-specific binding and provide a structural explanation for why LlaJI homologs do not target modifications despite sharing a similar domain organization with McrBC.
Importantly, we identify the N-arm Arg-17 and C-arm Arg-60 residues as critical determinants of DNA binding and specificity in HpR1Δ136. Individual point mutations at these positions display moderate defects, whereas a combined double mutant completely abolishes DNA binding in all assays tested (Figs. 6F and 7). The H14A and P59A mutations show varying effects depending on the specific substrate used and the sensitivity of the assay. Although His-14 and Pro-59 may also impart specific binding interactions, their contribution is likely context dependent.
HpLlaJI.R1 is unique in that its isolated B3 domain dimerizes, both in solution (Fig. 2C) and in crystallo (Fig. 1). β7 is absolutely essential for HpR1Δ136 dimerization and structural stability, as a truncation of this motif renders the protein insoluble. Our structure shows that direct dimerization of other B3 domains is hindered by either (i) the intrinsic lack of a structurally equivalent β7 strand or (ii) the presence of additional helical motifs at the N or C terminus that sterically clash with β7 or α4 at the dimer interface (Fig. 5). Dimerization of other B3 domain-containing proteins instead occurs through additional structural elements. For instance, AtARF1 monomers associate through a separate dimerization domain, which facilitates cooperative binding of the B3 domains to two anti-parallel 5′-TGTCTC-3′ sites on opposing strands (
). These observations will help in classifying uncharacterized B3 domains and predicting their architectural organization.
Our structural data show that the β7 strand from one HpR1Δ136 monomer is asymmetrically stabilized at the dimer interface, whereas the corresponding region in the other monomer remains disordered. The orientation of this strand dictates the electrostatic landscape on the dimer surface, making the DNA-binding site in one monomer more basic than the other. Although we cannot completely rule out that this is an artifact of crystallization, an analogous phenomenon was noted in the rotavirus A nonstructural protein 3 (NSP3) homodimer (
). There the asymmetric stabilization of a helix from one monomer creates a single positively charged binding site that ultimately leads to a stoichiometry of 2:1 NSP3:viral mRNA (
). We speculate that HpLlaJI.R1 may bind DNA with a similar 2:1 stoichiometry, but that only one B3 domain will directly contact the target site.
Asymmetric binding could have important implications for the assembly of a cleavage-competent LlaJI restriction complex. Like McrB, LlaJI.R1 contains a conserved GTP-specific AAA+ domain at its C terminus (
A mutational analysis of the PD.D/EXK motif suggests that McrC harbors the catalytic center for DNA cleavage by the GTP-dependent restriction enzyme McrBC from Escherichia coli.
). Although the exact organization of McrBC on DNA has yet to be elucidated, biochemical and structural studies have shown the EcMcrB N-terminal domain binds a single methylated cytosine via base flipping (
). The intrinsic asymmetry of the McrBC complex therefore imposes constraints on how the individual subunits interact with the RMC site and suggests that some monomers are directly engaged whereas others are not. The asymmetric HpR1Δ136 dimer could reflect a similar structural constraint in the LlaJI system wherein the alternative positioning of the β7 strand dictates which monomers bind the target sequence. Further structural characterization of both systems will be necessary to parse out how substrate binding, GTP-dependent assembly, and nuclease recruitment are coordinated in each case. Although LlaJI and McrBC differ in their specificity and targeting, we predict the general molecular mechanisms governing the function of LlaJI and McrBC will be conserved.
Experimental procedures
Cloning, expression, and purification of HpLlaJI.R1 constructs
DNA encoding the H. pylori LlaJI.R1 protein (DOE IMG/M ID 637022177) was codon optimized for E. coli expression and synthesized commercially by Bio Basic Inc. DNA encoding the N-terminal domain (HpR1Δ136; residues 1–136) was amplified by PCR and cloned into pET21b, introducing a His6 tag at the C terminus. Selenomethionine-labeled (SeMet) HpR1Δ136 was expressed in minimal media using methionine auxotrophs (T7 Express Crystal Competent E. coli, New England Biolabs) according to manufacturer protocols. Native HpR1Δ136 was transformed into BL21(DE3) cells, grown at 37 °C in Terrific Broth to an A600 of 1.0, and then induced with 0.3 mm isopropyl 1-thio-β-d-galactopyranoside overnight at 19 °C. All cells were harvested, washed with nickel load buffer (20 mm HEPES, pH 7.5, 500 mm NaCl, 30 mm imidazole, 5% glycerol (v/v), and 5 mm β-mercaptoethanol), and pelleted a second time. Pellets were typically flash frozen in liquid nitrogen and stored at −80 °C.
Thawed pellets from 500-ml cultures were resuspended in 30 ml of nickel load buffer supplemented with 10 mm phenylmethylsulfonyl fluoride, 5 mg of DNase (Roche Applied Science), 5 mm MgCl2, and a Roche complete protease inhibitor mixture tablet (Roche). Lysozyme was added to 1 mg/ml and the mixture was incubated for 15 min rocking at 4 °C. Cells were disrupted by sonication and the lysate was cleared of debris by centrifugation at 13,000 rpm (19,685 × g) for 30 min at 4 °C. For native and SeMet HpR1Δ136, the supernatant was filtered, loaded onto a 5-ml HiTrap chelating column charged with NiSO4 and then washed with nickel load buffer. HpΔ136 was eluted with an imidazole gradient from 30 mm to 1 m. Pooled fractions were dialyzed overnight at 4 °C into SP loading buffer (20 mm HEPES, pH 7.5, 50 mm NaCl, 1 mm EDTA, 5% glycerol (v/v), and 5 mm DTT). The sample was applied to a 5-ml HiTrap SP HP column equilibrated with SP loading buffer and then washed with SP loading buffer. HpR1Δ136 was eluted with a NaCl gradient from 50 mm to 1 m. Pooled fractions were concentrated and further purified by SEC using a Superdex 200 10/300 column. All proteins were exchanged into a final buffer of 20 mm HEPES, pH 7.5, 150 mm KCl, 5 mm MgCl2, and 1 mm DTT (5 mm for SeMet labeled) during SEC and concentrated to 5–40 mg/ml. Concentrations of purified proteins were determined by SDS-PAGE and densitometry was compared against BSA standards. All amino acid substitutions were introduced into HpR1Δ136 in pET21b by QuikChange PCR and mutant proteins were purified as described for WT.
Cloning, expression, and purification of EcMcrB
DNA encoding the E.coli McrB protein (Uniprot P15005) was codon optimized for E. coli expression and synthesized commercially by GENEART. DNA encoding the full-length protein (EcMcrB FL) was amplified by PCR and cloned into c2xP, a modified pMAL c2x vector with an HRV3C protease site replacing the Factor Xa site directly upstream of the mcrB gene. Native EcMcrB FL was expressed as N-terminal maltose-binding protein fusion in BL21(DE3) cells. Transformed cells were grown at 37 °C in Terrific Broth to an A600 of 0.8–1.0, and then induced with 0.3 mm isopropyl 1-thio-β-d-galactopyranoside overnight at 19 °C. All cells were harvested, washed with amylose loading buffer (20 mm Tris-HCl, pH 8.0, 500 mm NaCl, 5% glycerol (v/v), 1 mm EDTA, and 1 mm DTT), and pelleted a second time. Pellets were typically flash frozen in liquid nitrogen and stored at −80 °C.
Thawed pellets from 500-ml cultures were resuspended in 30 ml of amylose loading buffer supplemented with 10 mm phenylmethylsulfonyl fluoride, 5 mg of DNase (Roche), 5 mm MgCl2, and a Roche complete protease inhibitor mixture tablet (Roche). Lysozyme was added to 1 mg/ml and the mixture was incubated for 15 min rocking at 4 °C. Cells were disrupted by sonication and the lysate was cleared of debris by centrifugation at 13,000 rpm (19,685 × g) for 30 min at 4 °C. The supernatant was filtered, loaded onto 40 ml of packed amylose resin (New England Biolabs), and then washed with amylose loading buffer. EcMcrB FL was eluted with amylose loading buffer supplemented with 10 mm d-maltose. Hrv3C protease was added to pooled fractions and dialyzed overnight at 4 °C into Q loading buffer (20 mm Tris-HCl, pH 8.0, 50 mm NaCl, 1 mm EDTA, 5% glycerol (v/v), and 1 mm DTT). The sample was applied to a 5-ml HiTrap Q HP column equilibrated with Q loading buffer and then washed with Q loading buffer. EcMcrB FL was eluted with a NaCl gradient from 50 mm to 1 m. Pooled fractions were concentrated and further purified by SEC using a Superdex 75 10/30pg column. All proteins were exchanged into a final buffer of 20 mm HEPES, pH 7.5, 150 mm KCl, 5 mm MgCl2, and 1 mm DTT during SEC and concentrated to 5–40 mg/ml.
Crystallization, X-ray data collection, and structure determination
SeMet HpR1Δ136 was crystallized by sitting drop vapor diffusion in 0.1 m 1:2:2 dl-malic acid:MES:Tris base (MMT), pH 6.5, 25% PEG 1,500 (v/v) by mixing 1 μl of protein with 1 μl of the condition with a final drop size of 2 μl and reservoir volume of 65 μl. Crystals appeared within 2–8 days at 20 °C. Samples were cryoprotected with Parabar 10312 and frozen in liquid nitrogen. Single-wavelength anomalous diffraction (SAD) data of two crystals were collected remotely on the tuneable NE-CAT 24-ID-C beamline at the Advanced Photon Source at the selenium edge energy at 12.663 kEv (Table 1). Crystal 1 was of the space group P1 with unit cell dimensions a = 37.47, b = 44.39, c = 84.78 Å and α = 98.04°, β = 94.37°, γ = 98.52° and showed strong anomalous signal. Crystal 2 was of the space group P1 with unit cell dimensions a = 37.53, b = 43.77, c = 85.09 Å and α = 97.87°, β = 93.86°, γ = 97.77°. Both crystals were prepared in the same condition but exhibited mosaicities of 0.20871° and 0.11033°, respectively. Data were integrated and scaled using XDS (
) via the NE-CAT RAPD pipeline. Se-SAD phasing with the data from crystal 1 yielded an initial model that was incomplete and contained a few regions of ambiguity. Heavy atom sites were located using SHELX (
), but failed to improve significantly the density and refinement statistics. A more complete model was obtained using the diffraction data from crystal 2. The structure was solved by molecular replacement with PHASER (
) using the SAD-derived structure from crystal 1 as the search model. This vastly improved the resulting maps and statistics following subsequent rounds of manual model building and refinement. The final model of crystal 2 was refined to 1.97-Å resolution with Rwork/Rfree = 0.1927/0.2233 (Table 1) and contained four molecules in the asymmetric unit: molecule A, residues 1–121; molecule B, residues 1–131; molecule C, residues 1–121; molecule D, residues 1–131. Threonine 57 exists as a Ramachandran outlier with relatively weak density in the β3–β4 loop of molecules B and D, respectively, and could not be refined further. All structural models were rendered with PyMOL (Schrödinger, Inc.) and surface electrostatics were calculated with APBS (
Purified HpR1Δ136 at 4 mg/ml was subjected to size-exclusion chromatography using a Superdex 200 10/300 column (GE Healthcare) equilibrated in SEC buffer (20 mm HEPES, pH 7.5, 150 mm KCl, 5 mm MgCl2, and 1 mm DTT). The column was coupled to a static 18-angle light scattering detector (DAWN HELEOS-II) and a refractive index detector (Optilab T-rEX) (Wyatt Technology). Data were collected continuously at a flow rate of 0.5 ml/min. Data analysis was carried out using the program Astra VI. Monomeric BSA at 4 mg/ml (Sigma) was used for normalization of the light scattering detectors and data quality control.
Preparation of oligonucleotide substrates
The following DNA oligonucleotides for filter binding were synthesized commercially by Integrated DNA Technologies (IDT): 5mC_us, 5′-CCGGGTAAGA(5mC)CGGTAGCGAGCCCGG; 5mC_ls, 5′-CCGGGCTCGCTA(5mC)CGGTCTTACCCGG; nm_us, 5′-CCGGGTAAGACCGGTAGCGAGCCCGG; nm_ls, 5′-CCGGGCTCGCTACGGTCTTACCCGG; Ll_us, 5′-CAGATCTGACGCTAGAGCTT; Ll_ls, 5′-AAGCTCTAGCGTCAGATCTG; Llscr_us, 5′-CAGATCTCGACGTAGAGCTT; Llscr_ls, 5′-AAGCTCTACGTCGAGATCTG.
Lyophilized single-stranded oligonucleotides were resuspended to 1 mm in 10 mm Tris-HCl and 1 mm EDTA and stored at −20 °C until needed. Single-stranded oligonucleotides were 5′ end-labeled with [γ-32P]ATP using polynucleotide kinase (New England Biolabs) and then purified on a P-30 spin column (Bio-Rad) to remove unincorporated label. Duplex substrates were prepared by heating equimolar concentrations of complementary strands (denoted with suffixes “us” and “ls” indicating upper and lower strands) to 95 °C for 15 min followed by cooling to room temperature overnight and then purification on an S-300 spin column (GE Healthcare) to remove ssDNA. Four duplex DNA substrates were prepared: a methylated EcMcrB-specific substrate, 5mC (5mC_us and 5mC_ls), a nonmethylated EcMcrB-specific substrate, nm (nm_us and nm_ls), a site-specific substrate containing the L. lactis LlaJI.R1-binding site Ll (Ll_us and Ll_ls), and a substrate with the L. lactis LlaJI.R1-binding site sequence scrambled as a control Llscr (Llscr_us and Llscr_ls).
Filter-binding assays
The standard buffer for the DNA-binding assays contained 25 mm MES, pH 6.5, 2.0 mm MgCl2, 0.1 mm DTT, 0.01 mm EDTA, and 40 μg/ml BSA. Binding was performed with purified HpR1Δ136 (WT or mutants) or EcMcrB FL at 30 °C for 10 min in a 30-μl reaction mixture containing 14.5 nm unlabeled DNA and 0.5 nm labeled DNA. Samples were filtered through KOH-treated nitrocellulose filters (Whatman Protran BA 85, 0.45 μm) using a Hoefer FH225V filtration device for ∼1 min. Filters were subsequently analyzed by scintillation counting on a 2910TR digital, liquid scintillation counter (PerkinElmer Life Sciences). All measured values represent the average of at least two independent experiments and were compared with a negative control to determine fraction bound.
EMSA
The standard buffer for the EMSAs contained 10 mm Tris-HCl, pH 8.0, 100 mm NaCl, 1 mm MgCl2, and 1 mm DTT. Binding was performed with purified HpR1Δ136 (WT or mutants) at 25 °C for 30 min in a 16-μl reaction mixture containing 10 ng/μl of N6-methyladenine-free λ-phage DNA (New England Biolabs) digested with BamHI and NdeI (New England Biolabs) and purified via a NucleoSpin® Gel and PCR Clean-Up kit (Machery-Nagel). Following incubation, samples were analyzed by 0.7% agarose gel in 1× TAE at 4 °C and 80 V for 90 min. All gels were stained with SYBR® Green in 1× TAE overnight at 25 °C (Thermo-Fisher Scientific) and visualized using a Bio-Rad Gel DocTM EZ imager system.
Author contributions
C. J. H. and J. S. C. conceptualization; C. J. H. and J. S. C. data curation; C. J. H. software; C. J. H. and J. S. C. formal analysis; C. J. H. and J. S. C. validation; C. J. H. and J. S. C. investigation; C. J. H. and J. S. C. visualization; C. J. H. and J. S. C. methodology; C. J. H. and J. S. C. writing-original draft; C. J. H. and J. S. C. writing-review and editing; J. S. C. resources; J. S. C. supervision; J. S. C. funding acquisition; J. S. C. project administration.
Acknowledgments
We thank April Lee for assistance with initial crystallization experiments, the Northeastern Collaborative Access Team (NE-CAT) beamline staff at the Advanced Photon Source (APS) for assistance with remote X-ray data collection, Dr. Holger Sondermann for critical reading of the manuscript and use of his SEC-MALS instrument, Dr. John O'Donnell for assistance with SEC-MALS data analysis, and Dr. Eric Alani and Dr. Carol Manhart for guidance and assistance with filter binding experiments. This work is based upon research conducted at the NE-CAT beamlines (24-ID-C and 24-ID-E) under the General User Proposals GUP-51113 and GUP-41829 (PI: J. S. C). NE-CAT is funded by National Institutes of Health program project Grant P41 GM103403. The Pilatus 6M detector on 24-ID-C beam line is funded by National Institutes of Health-ORIP HEI Grant S10 RR029205. This research used resources of the Advanced Photon Source, a U.S. Dept. of Energy (DOE) Office of Science User Facility operated for the DOE Office of Science by Argonne National Laboratory under Contract DE-AC02-06CH11357.
A mutational analysis of the PD.D/EXK motif suggests that McrC harbors the catalytic center for DNA cleavage by the GTP-dependent restriction enzyme McrBC from Escherichia coli.
The GTP-dependent restriction enzyme McrBC from Escherichia coli forms high-molecular mass complexes with DNA and produces a cleavage pattern with a characteristic 10-base pair repeat.
Structural insight into the specificity of the B3 DNA-binding domains provided by the co-crystal structure of the C-terminal fragment of BfiI restriction enzyme.
The Arabidopsis B3 domain protein VERNALIZATION1 (VRN1) is involved in processes essential for development, with structural and mutational studies revealing its DNA-binding surface.
This work was supported by Cornell University and National Institutes of Health Grant GM120242 (to J. S. C). The authors declare that they have no conflicts of interest with the contents of this article. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.
The atomic coordinates and structure factors (code6C5D) have been deposited in the Protein Data Bank (http://wwpdb.org/).
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