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Research Article
Transparent Process
Open Access

Structural and functional characterization of TraI from pKM101 reveals basis for DNA processing

Annika Breidenstein, Josy ter Beek  Correspondence email, Ronnie P-A Berntsson  Correspondence email
Published 20 January 2023. DOI: 10.26508/lsa.202201775

Abstract

Type 4 secretion systems are large and versatile protein machineries that facilitate the spread of antibiotic resistance and other virulence factors via horizontal gene transfer. Conjugative type 4 secretion systems depend on relaxases to process the DNA in preparation for transport. TraI from the well-studied conjugative plasmid pKM101 is one such relaxase. Here, we report the crystal structure of the trans-esterase domain of TraI in complex with its substrate oriT DNA, highlighting the conserved DNA-binding mechanism of conjugative relaxases. In addition, we present an apo structure of the trans-esterase domain of TraI that includes most of the flexible thumb region. This allows us for the first time to visualize the large conformational change of the thumb subdomain upon DNA binding. We also characterize the DNA binding, nicking, and religation activity of the trans-esterase domain, helicase domain, and full-length TraI. Unlike previous indications in the literature, our results reveal that the TraI trans-esterase domain from pKM101 behaves in a conserved manner with its homologs from the R388 and F plasmids.

Introduction

Antibiotic resistance is one of the most pressing health challenges in today’s world. One of the main drivers of this worldwide problem is the ability of pathogens to spread resistance genes using horizontal gene transfer. This is a process that is often facilitated by conjugative type 4 secretion systems (T4SSs) (Juhas, 2015; Gonzalez-Rivera et al, 2016). These are megadalton-sized, multicomponent systems that transport DNA and proteins from a bacterial donor cell into a recipient cell (Grohmann et al, 2018; Waksman, 2019). In Gram-negative bacteria, T4SS consist of a channel that is made up of an inner membrane complex that is connected with an outer-membrane core complex by a stalk (Low et al, 2014; Chandran Darbari & Waksman, 2015; Costa et al, 2021). Other important features are the pilus, which extends out from the cell and is involved in mediating attachment to recipient cells, and the type IV coupling protein, an ATPase that helps to recruit the single-stranded DNA substrate which is coupled to a relaxase (Arutyunov & Frost, 2013; Grohmann et al, 2018). In recent years, researchers have generated structural and functional insight into these systems, especially from Gram-negative (G−) bacteria, among others the R388 plasmid and the pKM101 system (Khara et al, 2021; Macé et al, 2022). For a recent review of the structure and function of T4SSs, please see Costa et al (2021). The focus on this work is the T4SS of pKM101, which belongs to the class of minimal T4SS that consist of 12 proteins homologous to the paradigmatic VirB/VirD4 T4SS from Agrobacterium tumefaciens (Christie, 2004, 2016).

For all known T4SSs that transfer conjugative plasmids, the plasmid DNA must be processed before it can be transported. This is done via the relaxosome complex, which consists of a relaxase, accessory factor protein(s), and the DNA (Grohmann et al, 2018). To form this complex, one or several accessory proteins bind to the origin of transfer (oriT) on the DNA and locally melt the double-stranded DNA to promote relaxase binding to a defined sequence, often forming a hairpin, close to the nicking site (Zechner et al, 2017). The relaxase binds this single-stranded oriT DNA via its N-terminal trans-esterase domain (Garcillán-Barcia et al, 2009). This domain reacts with the DNA via a transesterification reaction at the specific nic-site, which generates the transfer intermediate consisting of the relaxase covalently bound to the 5′ end of the cleaved transfer strand (T-strand) (Byrd & Matson, 1997). Many relaxases have a second functional domain in the more variable C-terminal part of the enzyme. This is often a helicase domain, which unwinds the DNA to allow for transport of the single-stranded transfer-strand DNA (Garcillán-Barcia et al, 2009). The relaxase–transfer-strand complex (T-complex) is recruited to the T4SS by the type IV coupling protein and is transported through the T4SS channel into the recipient cell (Alvarez-Martinez & Christie, 2009). Once present in the recipient cell, the trans-esterase domain religates the DNA to regenerate the circularized plasmid (Waksman, 2019).

Relaxases have been phylogenetically classified into eight MOB families: MOBF, MOBH, MOBQ, MOBC, MOBP, MOBV, MOBT, and MOBB. Of these, pKM101-encoded TraI (TraIpKM101) belongs to the MOBF11 subclade together with its closest relative TrwC from plasmid R388 (TrwCR388) and the identical TraI from the sister plasmid pCU1 (TraIpCU1) (Paterson et al, 1999; Garcillán-Barcia et al, 2009; Guglielmini et al, 2011). Although structural information is available for the trans-esterase domains of several MOBF relaxases (De La Cruz et al, 2010; Zechner et al, 2017), structural data for the substrate DNA-bound state is only available for TrwCR388 and TraI from the F-plasmid (TraIF), which belongs to subclade MOBF12 (Larkin et al, 2005; Boer et al, 2006). Furthermore, there is only very limited data on full-length relaxases, again mostly from TraIF. TraIF consists of one trans-esterase domain, a vestigial helicase domain, and an active helicase domain (Fig 1A) and binds oriT DNA as a heterogenous dimer. One TraIF monomer adapts an open conformation and binds oriT at a hairpin positioned 5′ of the nic-site with the trans-esterase domain, whereas a second TraIF binds ssDNA with the helicase domains in a closed conformation. These two states are incompatible with each other in a single monomer (Ilangovan et al, 2017). A cryo-EM structure of the helicase-bound state is available and shows how the single-stranded DNA is almost entirely surrounded by the helicase domains. This structure is of full-length TraIF and visualizes all of the protein except the very flexible C-terminal domain (Ilangovan et al, 2017). It is not clear to what extent this structure and mode of action apply to TraIpKM101 as it is much shorter and completely lacks the vestigial helicase domain (Fig 1A).

Figure 1.
Figure 1. Schematic overviews of domain organization of relaxases and the cognate oriT of pKM101.

(A) Domain structure of three MOBF family relaxases TraIpKM101 (identical to TraIpCU1), TrwCR388, and TraIF (Ilangovan et al, 2017; Zechner et al, 2017). Trans-esterase domains are shown in green, vestigial helicase domain in light blue, active helicase domains in dark blue, and the C-terminal domains and linker regions are shown in gray. (B) oriT sequence of pKM101 and an overview of the different oriT nucleotides used in this study. * indicates the nic-site, underlined regions of the sequence indicate the bases participating in hairpin formation.

In this study, we present the crystal structure of the trans-esterase domain of TraIpKM101 bound to its substrate oriT DNA. This structure confirms the highly conserved DNA-binding mode of the trans-esterase domain observed in other MOBF family relaxases. We also present the apo structure with the flexible thumb-subdomain defined, which shows a large conformational change between the apo and substrate-bound structures. TraI is further characterized via electrophoretic mobility shift assays (EMSAs) and nicking and religation assays.

Results

Purification of TraI and its functional domains

TraI and its functional domains as predicted by PsiPred (Buchan & Jones, 2019), the trans-esterase domain (TraI-TE) and the helicase domain (TraI-H), were expressed in E. coli with a cleavable His-tag and purified to homogeneity in a three-step process. TraI, TraI-TE, and TraI-H, all with their His-tags cleaved off, were shown to be monomeric in solution by size-exclusion chromatography coupled to multi-angle light scattering (SEC-MALS) (Fig S1). It is interesting to note that full-length TraI was initially purified without removing the His-tag (TraIHis) and was then shown to be in an oligomer–monomer equilibrium, with the molecular weight of the oligomer determined to be 485±10 kD, which corresponds well to the expected molecular weight of a tetramer (492 kD) (Fig S2). Because we only observed oligomerization of TraIHis, but not for TraI without the His-tag or TraI-TE and TraI-H, we conclude that TraI is a monomer and that the initially observed oligomerization of the full-length protein was an artifact induced by the His-tag.

Figure S1.
Figure S1. Results from size-exclusion chromatography coupled to multi-angle light scattering experiments of TraI constructs without His-tag (left panels) and purified fractions of these protein constructs on SDS–PAGE (right panels).

In the left panels, the black chromatograms show the Rayleigh ratio which is directly proportional to the intensity of the scattered light in excess to the solvent (left axis). The red lines indicate the calculated molecular weight of the protein throughout the peak, plotted on the right axis. (A, B, C) Average molecular weights throughout the peaks were calculated to be: 116 ± 1 kD for TraI (A), 33 ± 1 kD for TraI-TE (B), and 49 ± 2 kD for TraI-H (C). These corresponded to the theoretical molecular weight of the monomers, which is indicated next to the SDS–PAGE fractions in each of the panels.

Figure S2.
Figure S2. His-tag induced tetramerization of TraIHis.

The chromatogram shows the elution profile and calculated molecular weight from size-exclusion chromatography coupled to multi-angle light scattering of TraI without removal of the His-tag (TraIHis). The Rayleigh ratio which is directly proportional to the intensity of the scattered light in excess to the solvent is indicated with a black line plotted on the left axis, and the calculated molecular weight of the protein throughout the peak is indicated with a red line and plotted on the right axis. The average molecular weight throughout the first elution peak is 485 ± 10 kD, corresponding to a tetramer, and throughout the second elution peak, this was calculated to be 123 ± 5 kD, corresponding to a monomer.

TraI is a functional relaxase capable of binding, nicking, and religating oriT-DNA

To biochemically characterize TraI and its functional domains, we performed several activity assays. To estimate the differences in binding affinity and specificity of the different TraI domains, we used EMSAs. We investigated DNA binding to a fluorescent 22-mer with the post-nic DNA sequence from the oriT of pKM101 (oriT22-F), a fluorescent 35-mer of the pre-nic DNA containing the hairpin structure (F-oriT35), and a fluorescent full-length 57-mer with both the pre- and post-nic sequence (F-oriT57), as well as randomly sequenced 57- and 35-mers (Fig 1B and Table S1).

We found that the TraI trans-esterase domain binds both F-oriT57 and F-oriT35 DNA with high affinity (at 200 nM over half of the DNA is bound, Figs 2A and S3A), whereas it only interacts with post-nic or random DNA at protein concentrations above 800 or 1,600 nM (Figs 2A and S3A). This indicates that the trans-esterase domain binds sequence-specific to the pre-nic region. In contrast, the TraI helicase domain shows only low affinity binding to both oriT and random DNA and seems to have a stronger interaction with the longer 57-mer than with the 35- and 22-mer DNA constructs (Figs 2B and S3B). This domain thus appears to have no sequence specificity and to prefer DNA fragments longer than 35 nucleotides.

Figure 2.
Figure 2. TraI binding to oriT or random DNA.

(A, B, C) Electromobility shift assay using 50 nM fluorescent 57-mer oriT (F-oriT57) (upper panel) or random DNA (lower panel) of the trans-esterase domain (TraI-TE) (A), helicase domain (TraI-H) (B), or full-length TraI (C).

Figure S3.
Figure S3. DNA binding by TraI.

(A, B, C) Electromobility shift assay using 50 nM F-oriT35 (upper panel), F-random35 (middle panel), or oriT22-F (lower panel) of the TraI trans-esterase domain (TraI-TE) (A), the TraI helicase domain (TraI-H) (B), or full-length TraI (C).

As full-length TraI includes both the helicase and the trans-esterase domains, we expected to find a combination of their DNA-binding capabilities. TraI was indeed found to bind both oriT and random DNA. Interestingly, the random DNA shows a significant mobility shift with TraI protein concentrations of 200 nM and above (Fig 2C), whereas 400 nM is needed when the TraI helicase domain is used in isolation (Fig 2B). Both F-oriT57 and F-oriT35 show a complete mobility shift with TraI concentrations of 200 nM and above (Figs 2C and S3C), compared with 400 nM for the trans-esterase domain (Figs 2A and S3A). Both observations therefore indicate a significant increase in the apparent affinity of the full-length TraI as compared with its individual domains. At higher TraI concentrations, additional supershifts are observed (Fig 2C). An additional observation was an initial very high shift that was observed for F-oriT57 and F-oriT35 already with 50 nM TraI (Figs 2C and S3C) (similar to the high shift seen at the highest concentration of TraI-H [Figs 2B and S3B]). However, this high–molecular-weight band was only observed up to 400 nM TraI and disappears as the protein concentration is increased further. This result has been consistently observed in mobility shift assays with oriT DNA (both oriT35 and oriT57) but not with random DNA (Figs 2C and S3C).

We also examined the capacity of TraI to nick and religate its substrate oriT DNA. TraI was incubated with the fluorescent full-length F-oriT57, containing the recognition hairpin, the nic-site, and a post-nic stretch of 22 bases, or a random DNA sequence of the same length. DNA products were consequently separated on a denaturing polyacrylamide gel. Our results (Fig 3A and C) demonstrate that both TraI and TraI-TE can nick about 50% of the tested oriT DNA substrate, whereas neither nicked the random DNA control under the same conditions. We were unable to detect a higher degree of cleaved F-oriT57 DNA and predicted that this was because of the expected religation ability of TraI. To test this, we added a shorter version of the fluorescent pre-nic DNA (F-oriT20) to the incubation mixture of TraI. In this experiment, F-oriT57 is expected to be (partly) cleaved by TraI into a fluorescent 35-mer and a non-fluorescent 22-mer that stays covalently bound to TraI. If religation occurs, the 22-mer could either be religated to the fluorescent 35-mer to re-form the original F-oriT57 or to the F-oriT20 to form a fluorescent 42-mer. This 42-mer was indeed seen on the denaturing gel (Fig 3B and D, left-side). This nicking and religation activity of TraI was further investigated by incubating the protein with a fluorescent F-oriT35 and a non-fluorescent oriT57. The appearance of F-oriT57 indicates that non-fluorescent oriT57 was nicked and that the resulting 22-mer was ligated to F-oriT35 (Fig 3B and D, right-side).

Figure 3.
Figure 3. DNA nicking and religation by TraI.

(A, B, C, D) Activity assays of TraI (A, B) and TraI-TE (C, D). (A, C) Nicking assays. F-oriT57 (left) and F-Random57 (right) were incubated with increasing protein concentrations, and the DNA was separated on a denaturing gel, showing the emergence of a 35-mer nicking product for oriT DNA only. (B, D) Religation assays. F-oriT57 and F-oriT20 were incubated with increasing protein concentrations (left-side). The appearance of a 35-mer product indicates nicking of F-oriT57, and the appearance of a 42-mer product indicates religation of the covalently bound post-nick DNA with the shorter F-oriT20 pre-nick DNA. F-oriT35 and oriT57 were incubated with increasing protein concentrations (right-side). The appearance of a 57-mer product shows nicking of the non-fluorescent oriT57 and religation with the fluorescent F-oriT35.

Crystal structure of TraI trans-esterase domain in complex with oriT DNA

To obtain a mechanistic understanding of the sequence-specific DNA-binding ability of TraI, purified TraI-TE was co-crystallized with single-stranded 11-mer oriT DNA (oriT11) (Fig 1B), and the obtained crystal structure was refined at a resolution of 2.1 Å. Virtually all residues were visible in the electron density, apart from five residues in flexible loop regions and at the C-terminus. We were also able to model 10 of the 11 bases into the electron density of the bound DNA (Fig 4).

Figure 4.
Figure 4. Crystal structure of DNA-bound trans-esterase domain of TraIpKM101.

In all panels, TraI is colored green, the DNA bound by TraIpKM101 is shown in purple and orange, and the bound divalent cation (here modeled as Mn2+) in gray, unless otherwise indicated. (A) Cartoon representation of the trans-esterase domains of TraIpKM101 colored from blue at the N-terminal to red at the C-terminal, with α-helices labeled from 1 to 10. (B) Superimposition of the trans-esterase domains of TraIpKM101 and TrwCR388 (protein in beige, DNA in gray, PDB: 2CDM). (C) Active site of TraI-TEpKM101 consisting of the histidine triad coordinating a Mn2+ (H149, H160, and H162), conserved aspartate (D84), and catalytic tyrosine (Y18). (D) Bound oriT DNA and its interactions with TraI-TE. Residues that form hydrogen bonds with DNA and the DNA itself are represented as sticks.

The overall architecture of TraI-TE can be described as a hand, with the bound DNA located between the palm and the C-terminal thumb subdomains (Figs 4A and S4A). The determined structure is similar to the previously described homologs TrwCR388 and TraIF (Fig 4B) (Guasch et al, 2003; Larkin et al, 2005; Boer et al, 2006). A conserved histidine triad coordinates a divalent metal that is required for the trans-esterase reaction in the homologs (Byrd & Matson, 1997; Datta et al, 2003; Boer et al, 2006). In TraI, this triad is formed by H149, H160, and H162 (Fig 4C). Electron density was observed in the middle of this triad, and refinement against the possible metal ions yielded the best fit for manganese (Fig S4B and C), albeit this does not rule out that other divalent cations are bound. The conserved aspartate, D84 in TraIpKM101, is situated within hydrogen-bonding distance of both H162 and Y18 (2.8 and 2.7 Å, respectively, Fig 4C) as earlier described for TrwCR388 and TraIF (Larkin et al, 2005; Boer et al, 2006). It is therefore in position to activate the catalytic tyrosine Y18, the first tyrosine of the conserved YY-X(5-6)–YY motif (Garcillán-Barcia et al, 2009; Zechner et al, 2017), which is predicted to occur during the trans-esterification reaction with the scissile phosphate at the oriT nic-site.

Figure S4.
Figure S4. Crystal structure of the trans-esterase domain of TraIpKM101.

(A) Cartoon representation of the DNA-bound trans-esterase domain of TraIpKM101, where the palm subdomain is colored green, the thumb subdomain is colored pink, the DNA is shown in orange and purple, and the divalent cation in gray. (B, C) Coordination of the divalent cation, here modeled as a Mn2+, by the histidine triad (H149, H160, and H162) of the apo structure (B) and DNA-bound structure (C). The electron density is shown as a mesh (SigmaA-weighted 2Fo−Fc maps, contoured at 2.5 σ).

The DNA substrate used for co-crystallization was oriT11, representing the 11 bases directly 5′ of the nic-site. Of these, the first base at the 5′ end T1 did not yield any defined electron density, likely because it was not properly stabilized (Fig 4D). Although the first visible base C2 does not show any specific contacts with TraI-TE, the following guanine (G3) forms four hydrogen bonds with R80 and D182 using both the Hoogsteen and Watson–Crick edge, and the following thymine (T4) forms two hydrogen bonds with N181. The following two bases, T5 and A6, are stacked on top of each other perpendicular to the orientation of the previous three bases and form hydrogen bonds with K188 (T5) and Q256 (A6). These initial bases are oriented in a position that is pointing away from the active site. The following bases place the DNA in a conformation resembling a U-turn that locates the scissile phosphate of T11 in close proximity to the catalytic tyrosine. This bent DNA conformation is stabilized with multiple hydrogen bonds, combined with hydrophobic and pi–pi interactions between the stacked bases G8 and G10. G7 forms two hydrogen bonds with L2 at its Watson–Crick edge and an additional one with K196 from its Hoogsteen edge. G8 forms two Watson–Crick hydrogen bonds with D3, and G10 forms two Hoogsten hydrogen bonds with R250. The phosphates of the three most 3’ bases are also stabilized with several hydrogen bonds (S236 for T9, S236 and K91 for G10, and R235 and R153 for T11) (Fig 4D). Taken together, these DNA–DNA and DNA–protein interactions illustrate a binding mechanism that relies both on the secondary structure and the specific sequence of the DNA.

Apo structure reveals rearrangement of the thumb subdomain upon DNA binding

An important feature of the DNA binding is the TraI-TE C-terminal thumb subdomain, consisting of two α-helices connected by a loop region (Figs 4A and S4A). This subdomain lies on top of the 3′ part of the DNA and forms hydrogen bonds with the oriT DNA via residues R235, R250, and Q256 (Fig 4D). The thumb subdomain is highly flexible and has previously only been observed in relaxase structures with DNA bound. Unexpectedly, our co-crystallization experiment resulted in an additional crystal with the space group P212121 that resolved to 1.7 Å. This structure had a different unit cell, which had the fortuitous outcome of supporting an apo state of TraI-TE, in which we were able to build most of the thumb subdomain in an open conformation (Fig 5A). The electron density corresponding to residues 261–271 was of insufficient quality to build these residues in the model, but inspection of the density suggests that the α-helix continues toward the base of the thumb. Between the apo- and the DNA-bound state, the thumb moves up to 40 Å and undergoes important structural changes. The observed conformational differences include a reorganization of α-helix 8 and the extension of α-helix 9 (numbered based on the DNA-bound structure as one helix disappears in the apo structure) to transition from the apo to the DNA-bound structure, as illustrated in Fig 5A and Video 1. These rearrangements reveal the structural changes that are necessary to first allow DNA to enter the binding site and to subsequently keep the DNA in place, highlighting the importance of the thumb subdomain in this process. The C-terminal helix at the base of the thumb subdomain (α-helix 10) interacts with the loop at the end of α-helix 1 via a hydrogen bond between R274 and D25 in both the DNA-bound and apo structures (Fig S5A). An additional interaction is made between H278 and Y26, the third tyrosine of the YY-X(5-6)–YY motif, but only in the DNA-bound structure.

Figure 5.
Figure 5. Overview of the conformational changes between the apo and the DNA-bound structures and the differences with the previously published structure of the identical TraI-TEpCU1.

(A) DNA-bound structure of TraIpKM101 (protein in green, DNA in purple and orange, Mn2+ as a gray sphere), superimposed with the apo structure (light gray). To provide a visual aid in understanding the conformational changes that occur in the thumb subdomain upon DNA binding, residue K241 (red), R251 (blue), and Q256 (cyan) are highlighted in both structures, and α-helix 8 and 9 are indicated in the DNA-bound structure. (A, B) DNA-bound structure of TraIpKM101 (colored as in panel (A)) with the thumb subdomain highlighted in pink, superimposed with the previously published structure of TraI-TEpCU1 (cyan, PDB: 3L6T). TraI-TEpCU1 lacks the thumb subdomain, and differences with TraI-TEpKM101 are also observed close to the active site, indicated by the dashed circle.

Video 1

Visual representation of the thumb subdomain movement during the transition from the apo to the DNA-bound state of TraIpKM101. The apo structure is represented in light gray, the DNA-bound structure is shown as a cartoon with protein in green and DNA in orange and purple, and the Mn2+ ion as a dark gray sphere. In both structures, the following residues are highlighted by color to make it easier to follow the conformational change: K241 (red), R251 (blue), and Q256 (cyan).Download video

Figure S5.
Figure S5. Overview of the interactions between the base of the thumb subdomain (α-helix 10) and the loop at the end of α-helix 1.

In all panels, TraI is shown in green, with important residues shown as sticks. (A) DNA-bound structure superimposed with the apo structure (in gray) of TraI. Hydrogen bonds are formed between R274 and D25 in both structures, whereas an additional bond between H278 and Y26 is only present in the DNA-bound structure. (B) DNA-bound structure of TraIpKM101 superimposed with TrwCR388 (PDB: 1S6M, purple). In TrwCR388, hydrogen bonds are formed between R274 and D24 or D25. (C) DNA-bound structure of TraIpKM101 superimposed with TraIF (PDB: 2Q7T, turquoise). In TraIF, hydrogen bonds are found between E271 and Y24.

Both the apo and the DNA-bound structure of TraI-TE presented here show differences when compared to the available structure from TraIpCU1, even though the domains have an identical primary sequence (Fig 5B, PDB: 3L6T) (Nash et al, 2010). In TraIpCU1, the C-terminal part of the protein containing the thumb subdomain and the base of the thumb are missing, and the loop at the end of α-helix 1 is pointing away from the active site, possibly because it is not stabilized by the missing α-helix 10 (Fig 5B).

Discussion

In this study, we structurally and biochemically characterized the TraI relaxase from the MOBF family plasmid pKM101 to better understand the diversity of the relaxases in this protein family as they are of key importance to type IV secretion.

We performed mobility shift assays (Figs 2 and S3) to estimate the DNA-binding affinity of full-length TraI and to compare this to the affinity of the trans-esterase (TraI-TE) and helicase (TraI-H) domains individually. Both full-length TraI and the helicase domain bound DNA in a sequence-unspecific manner as both bound to a similar degree to oriT and random DNA (Figs 2 and S3). This is as expected because these helicases are predicted to unwind the entire conjugative plasmid and therefore should not have any sequence specificity. In contrast, the trans-esterase domain of TraI (TraI-TE) showed a higher affinity for oriT DNA as compared with random DNA (Figs 2 and S3). Although the mobility shift assays presented here are not suitable for precise KD determinations, the apparent KD of the trans-esterase domain of TraIpKM101 for oriT DNA is <200 nM as more than half of the DNA was shifted upward at 200 nM (Fig 2A). This binding is sequence-specific as interactions with random DNA were only observed at protein concentrations above 800 nM (Fig 2A). Nicking and religation assays show that full-length TraI and trans-esterase domain (TraI-TE) are both equally capable of nicking and religating the oriT DNA but not random DNA. This shows that the trans-esterase domain is active on its own and confirms its oriT specificity (Fig 3).

The observed affinity and specificity of the trans-esterase domain for its cognate oriT DNA is a feature that has been described in various TraIpKM101 homologs, such as TrwCR388 and TraIF (Stern & Schildbach, 2001; Guasch et al, 2003; Carballeira et al, 2014). However, it is contradictory to a previous report on the trans-esterase domain of TraIpCU1 (Nash et al, 2010). pCU1 is an antibiotic resistance plasmid that shares a common ancestor with pKM101 and R46 (Paterson et al, 1999). The pKM101 and pCU1 conjugative plasmids have an identical oriT region and encode identical relaxases: TraIpKM101/TraIpCU1. In spite of this, the trans-esterase domain of TraIpCU1 was previously described to bind DNA in a weak and sequence-independent manner, with KD values for binding to oriT and non-oriT oligomers ranging from 0.7 to 1.2 μM (Nash et al, 2010). The DNA binding of TraIpKM101 and the relaxase domain of TraIpCU1 was measured using different methods and using slightly different protein constructs. The differences in the protein constructs are mainly the tags used and could explain the reduced affinity observed for the N-terminally tagged TraIpCU1 because adding additional residues at the N-terminus would likely disturb the active site. However, the differences are consistent also with the C-terminally tagged TraIpCU1, so we have no obvious explanation for the observed discrepancies between our experimental results and those of TraIpCU1. We do note that our observations for TraIpKM101 align well with published data for homologous proteins. We therefore conclude that TraIpKM101 binds and processes DNA in a similar fashion as its homologs TrwCR388 and TraIF.

Although His-tags sometimes cause dimerization (Amor-Mahjoub et al, 2006; Singh et al, 2020), it is very uncommon that they promote any defined higher oligomerization. This led us to initially think that TraI formed tetramers in solution (Fig S2), but this turned out to be an artifact of the His-tag (Fig S1). However, an interesting feature in the EMSAs with full-length TraI, and to some extent its helicase domain, is the appearance of supershifts at higher protein concentrations (Figs 2 and S3). The exact same data were observed both with His-tagged TraI and TraI with the His-tag removed. This suggests that multiple copies of the protein can bind to the same DNA molecule. Support for this can be found in the literature as TraIpKM101 was previously reported to self-interact in vivo (Li & Christie, 2020). In addition, TraIF was shown to bind oriT as a dimer where one subunit takes on a closed conformation, in which the helicase surrounds the DNA, and the other subunit takes on an open conformation, with the trans-esterase domain bound to oriT. These two conformations are proposed to be mutually exclusive as the directionality of the ssDNA was proposed to be inconsistent with the same protein binding with both the helicase and trans-esterase domain simultaneously (Ilangovan et al, 2017). Although there are significant differences between TraIF and TraIpKM101 (Fig 1A), it is possible that TraIpKM101 exhibits similar oligomerization when binding to DNA.

To understand the interaction between TraI and its substrate DNA on a mechanistic level, we solved the crystal structure of the trans-esterase domain in complex with its cognate oriT DNA. The DNA-bound structure of TraI-TEpKM101 (Fig 4A) shows a protein fold that can be described as a palm with the active site at the center and a thumb subdomain that is involved in holding the substrate DNA in place (Fig S4A). This structure is similar to the previously solved trans-esterase domains of its homologs (Guasch et al, 2003; Larkin et al, 2005; Boer et al, 2006) (Fig 4B). The highest structural homology is found with another member of the MOBF11 subclade: TrwCR388 (PDB: 2CDM, 50% identity, root mean square deviation (RMSD) = 1.7 Å over 276 residues). The structure is also similar to that of TraIF, a MOBF12 relaxase (PDB: 2A0I, 37% identity, RMSD = 2.3 Å over 269 residues) (Holm, 2020). The DNA-binding site is highly conserved between these proteins with the bound ssDNA making a characteristic U-turn (Carballeira et al, 2014). We modeled the divalent cation coordinated by the histidine triad as a Mn2+ because it clearly gave the best result in the refinements (Fig S4B and C). However, we do not have any experimental data to conclude with certainty that it is indeed Mn2+. The divalent cation found in the DNA-bound structure was likely retained during purification because no metals were added during purification or crystallization, whereas the apo structure was crystallized in the presence of both Mg2+ and Ca2+. Our activity assays were successfully performed in the presence of Mg2+. Because TraI homologs, containing the same active sites, have been shown to be able to function with various divalent metals, we did not investigate the identity of the metal further (Boer et al, 2006; Nash et al, 2010).

Although the DNA-bound structures of the MOBF family relaxases are very similar, there is more variability between the structures in the absence of DNA. This probably reflects the greater flexibility of the structures in the absence of the ligand. The apo structure of TraI-TEpKM101 presented here (Fig 5A) shows the thumb subdomain in an open conformation. Previously, this thumb subdomain could only be modeled in crystal structures with bound DNA (Guasch et al, 2003; Larkin et al, 2005; Boer et al, 2006). This thumb is also absent in the cryo-EM structure of TraIF, which has DNA bound to the helicase domain (Ilangovan et al, 2017). The RMSD between our ligand-free and DNA-bound structures is 3.3 Å over 277 residues, which is indicative of the large molecular movement of up to 40 Å (Fig 5A and Video 1). It is likely that the open conformation is important for oriT DNA to access the active site. The exact degree of opening observed in our structure is dependent on crystal contacts that stabilize the thumb subdomain. In solution, the protein is very likely in an open-closed equilibrium, which is shifted strongly toward probably many different open states in the absence of cognate DNA.

We were interested whether the thumb movement would be possible in the context of the full-length protein. To gain insight, we compared our TraI-TE structure to TraIF (PDB: 5N8O) and the alphafold model of full-length TraI (Jumper et al, 2021; Terwilliger et al, 2022). Superimposing TraI-TE on to the full-length TraI alphafold model shows that the thumb subdomain is facing the helicase domain in the model, but no direct contacts are predicted between the thumb and the helicase domains (Fig S6A). Furthermore, the cryo-EM structure of TraIF, which has DNA bound to the helicase domain, is missing the thumb subdomain (Fig S6B) (Ilangovan et al, 2017). This suggests that the thumb subdomain is flexible and exhibits an open-closed equilibrium also in TraIF. Taken together, this makes it likely that the conformational change of the thumb observed for the trans-esterase domain in isolation is also fully possible in context of the full-length protein.

Figure S6.
Figure S6. Structural comparison visualizing the thumb subdomain and its ability to open and close in the full-length protein.

Cartoon representation of the DNA-bound structure of the trans-esterase domain of TraIpKM101 with the palm subdomain in green, the thumb subdomain in pink, the DNA is shown in orange and purple, and the divalent ion in gray. (A, B) The structure is superimposed with the full-length TraIpKM101 AlphaFold model colored in gray (trans-esterase domain) and black (remaining protein) (panel (A)) or TraIF (PDB: 5N8O) colored in teal (trans-esterase domain), cyan (remaining protein), and yellow (bound DNA) (panel (B)). In both panels, the arrow represents the movement of the thumb subdomain upon opening.

There are important differences between the structures of TraI-TEpKM101 and the apo structure of TraI-TEpCU1 (PDB: 3L6T) despite their 100% sequence identity (Fig 5B). The overall fold is similar, with a RMSD of 1.4 Å over 208 residues, but the entire C-terminal region of TraIpCU1 is missing. This includes the thumb subdomain and the helix at the base of the thumb (α-helices 7–10 of the TraIpKM101 DNA-bound structure). More importantly, α-helix 1, which contains the beginning of the conserved YY-X(5-6)–YY motif that provides the catalytic tyrosine, mostly appears as a loop in the previously published TraIpCU1 structure. As a consequence, it points far away from the active site, resulting in a >10 Å displacement compared with the positions in the homologs (Nash et al, 2010). Possibly, the displacement of α-helix 1 is related to the destabilization of the thumb subdomain of the protein. In contrast, both structures presented here, as well as other relaxase structures (TrwCR388 PDB: 1S6M, TraIF PDB: 2Q7T), show interactions between the base of the thumb subdomain (α-helix 10) and the loop region following α-helix 1 (R274 to D25 in TraIpKM101) (Fig S5B and C). In the DNA-bound structure of TraIpKM101, an additional interaction was found between H278 (α-helix 10) and Y26 (the third Y of the YY-X(5-6)–YY motif), which might further contribute to this stabilization.

To conclude, we have shown that TraIpKM101 has a high affinity and specificity to its cognate oriT and can both nick and religate oriT in a similar manner as other MOBF family relaxases. The crystal structures of its trans-esterase domain (TraI-TEpKM101) visualize the large conformational change the protein undergoes upon DNA binding. Both the biochemical function and structures of TraIpKM101 reported here resemble those of the homologs TrwCR388 and TraIF more closely than the previously published work on the identical TraI-TEpCU1 (Nash et al, 2010). Our findings thus highlight the conserved mechanism of relaxases and the role they play in conjugation.

Materials and Methods

Plasmids

The full-length traI gene and shorter constructs encoding either the trans-esterase domain (TraI-TE) (residues 1–299) or the helicase domain (TraI-H) (residues 429–910) were PCR-amplified and cloned into p7XC3H and p7XNH3 via the FX cloning system (Geertsma & Dutzler, 2011) under the T7 promotor, with an N- or C-terminal 10-His-tag and a 3C protease sequence (see Table S1).

Protein expression and purification

All TraI variants were produced in E. coli BL21 (DE3). The cells were grown in 1.5 liters TB medium at 37°C using a LEX (large-scale expression) bioreactor (Epiphyte3). Once the cultures reached an OD600 between 1.0 and 1.5, the temperature was reduced to 18°C and expression was induced by adding 0.4 mM IPTG for ∼16 h. Cells were centrifuged at 6,000g, and pellets were resuspended in resuspension buffer (50 mM Hepes-NaOH, pH 7.0, 300 mM NaCl, 15 mM Imidazole, 0.2 mM AEBSF, and ca 0.02 mg/ml DNase I). Resuspended cells were lysed in a Cell Disruptor (Constant Systems) at 25 kPsi at 4°C, followed by centrifugation at 30,000g for 30 min at 4°C. The cell lysate was then incubated for 1 h at 4°C with Ni-NTA resin (∼2 ml/liters culture for TraI and TraI-H or ∼4 ml/liters culture for TraI-TE) before transfer to a gravity flow column. The resin was subsequently washed with 10 column volumes (CV) of wash buffer (50 mM Hepes-NaOH, pH 7.0, 300 mM NaCl, 50 mM Imidazole, 0.2 mM AEBSF), followed by 10 CV wash buffer with 2 M LiCl and another 10 CV wash buffer. For experiments without tag-cleavage, TraIHis was eluted with 5 CV elution buffer (50 mM Hepes-NaOH, pH 7.0, 300 mM NaCl, 500 mM Imidazole).

For experiments in which the tag was cleaved off, 3C PreScission Protease was added, while the protein was still bound to the IMAC resin, in an estimated ratio of 1:50–1:100 and incubated ∼16 h at 4°C. The cleaved protein was then recovered by collecting the flow through.

All samples were diluted 3× with dilution buffer (25 mM Hepes-NaOH, pH 7.0, 50 mM NaCl) after Ni-NTA purification and loaded on a HiTraP Heparin HP (5 ml) column, equilibrated with Buffer A (25 mM Hepes-NaOH, pH 7.0, 150 mM NaCl) with a peristaltic pump. After loading the protein, the column was connected to an ÄKTA Pure (Cytiva) and washed with Buffer A for ∼2 CV until absorbance at 280 nm was stable. All TraI variants were subsequently eluted via a salt gradient to 100% Buffer B (25 mM Hepes-NaOH, pH 7.0, 1.0 M NaCl) over 80 ml. Peak fractions with pure protein, as judged by a 260 nm/280 nm ratio of <0.8, were concentrated using Amicon Ultra Centrifugal filters with molecular weight cutoffs at 50 kD (TraI or TraIHis), 30 kD (TraI-H), or 10 kD (TraI-TE). Size-exclusion chromatography was performed in 25 mM Hepes-NaOH, pH 7.0, 300 mM NaCl on a Superdex 200 10/300 Gl Increase column, and fractions were analyzed on 12% SDS–PAGE stained with Coomassie blue.

SEC-MALS

SEC-MALS was performed with 250 μl of protein sample at a minimum concentration of 1 mg/ml in 25 mM Hepes-NaOH, pH 7.0, 300 mM NaCl. Experiments were done on a Superdex 200 10/300 Gl Increase column or a Superose 6 10/300 Gl Increase column for TraIHis, on ÄKTA Pure (Cytiva) coupled to a light scattering (Wyatt Treas II) and refractive index (Wyatt Optilab T-Rex) detector. Data were collected and analyzed using Astra software (version 7.2.2; Wyatt Technology) as described by Some et al (2019). The molecular weight of the protein samples was calculated as an average from a minimum of 3 measurements and is reported with the standard deviation.

DNA oligomers used for binding assays and crystallization

Single-stranded DNA oligomers were purchased from Eurofins Genomics, with and without the fluorescent label fluorescein isothiocyanate (FITC). The position of the label at the 5′ or 3′ end is indicated in Table S1, and a schematic of the different oriT DNA used in this study can be found in Fig 1B. Stock solutions of 100 mM were dissolved in MilliQ water, but all further dilutions were done in 25 mM Hepes-NaOH, pH 7.0, 300 mM NaCl. Oligomers were heated at 95°C for 5 min and cooled down to room temperature over at least 20 min before use.

EMSA

Fluorescently labeled DNA (50 nM, Table S1) was incubated for 15 min with 0–1,600 nM of protein (see the Results section) in 25 mM Hepes-NaOH, pH 7.0, 300 mM NaCl at room temperature before adding 6× native loading dye (3× TBE, 30% glycerol, 0.125% bromophenol blue). The samples were resolved on a native gel (5% polyacrylamide, 0.75× TBE) in 0.75× TBE running buffer at 50 V for 90 min at 4°C. Gels were imaged on an Amersham Typhoon 5 scanner with excitation at 488 nm, using the Cy2 emission filter (515–535 nm).

Nicking and religation assays

Nicking and religation assays were performed in 25 mM Hepes-NaOH, pH 7.0, 300 mM NaCl, 5 mM MgCl2. Nicking assays contained 100 nM F-oriT57, religation assays (version 1) contained 100 nM F-oriT57 and 100 nM F-oriT20, and religation assays (version 2) contained 100 nM F-oriT35 and 100 nM non-fluorescent oriT57 (Fig 1B and Table S1). DNA oligomers were incubated with increasing concentrations of TraI in a shaking heatblock at 37°C and 300 rpm, for 1 h. The reactions were stopped by adding 2× stop solution (96% formamide, 20 mM EDTA, 0.1% bromophenol blue). The samples were boiled for 5 min at 95°C before loading on a denaturing gel (16% polyacrylamide, 7 M urea, 1× TBE). The gel was run at 50 V for 15 min followed by 100 V for 1 h at room temperature and then imaged on an Amersham Typhoon 5 scanner with excitation at 488 nm, using the Cy2 emission filter (515–535 nm).

Crystallization and structure determination

Protein crystals were obtained with the sitting drop vapor diffusion method at 20°C. Substrate DNA, oriT11 (Fig 1B and Table S1), was added to TraI-TE, to yield a final concentration of 9 mg/ml TraI-TE and an equimolar amount of the DNA. This mixture was used for co-crystallization experiments, which yielded both the apo structure of TraI-TE and the DNA-bound structure. The apo structure crystallized in a 1:1 ratio with 10% wt/vol PEG 4000, 20% v/v glycerol, 0.03 M MgCl2, 0.03 M CaCl2, and 0.1 M MOPS/Hepes-Na, pH 7.5. The DNA-bound structure crystallized in a 1:1 ratio with 10% wt/vol PEG 20000, 20% vol/vol PEG MME 550, 0.03 M sodium nitrate, 0.03 M disodium hydrogen phosphate, 0.03 M ammonium sulfate, and 0.1 M MES/imidazole, pH 6.5. Crystals were flash-frozen in liquid nitrogen, without the addition of extra cryo-protectant. X-ray diffraction data were collected on beamline ID30B, at the ESRF, France. The data were processed using XDS (Kabsch, 2010; Monaco et al, 2013). Both crystals had the space group P212121 and contained a single copy of the protein in the asymmetric unit. The phase problem was solved using molecular replacement with the trans-esterase domain of TraIpCU1 (PDB: 3L6T) in PHENIX phaser (McCoy et al, 2007; Nash et al, 2010). The structures were built in Coot (Emsley & Cowtan, 2004) and refined at 2.1 Å (DNA-bound structure) and 1.7 Å (apo structure) in PHENIX refine (Afonine et al, 2012), to Rwork/Rfree values of 22.5/26.9% and 17.1/19.1%, respectively. Further refinement statistics can be found in Table S2.

Data Availability

Atomic coordinates and structure factors of both the apo and DNA-bound structures of TraI have been deposited with the Protein Data Bank (PDB: 8A1B and 8A1C).

Acknowledgements

A pET15b plasmid containing the pKM101 TraI gene that we used for further cloning in E. coli was generously donated by Prof. Peter J Christie, whom we also thank for fruitful discussions regarding the project. We acknowledge the MAX IV Laboratory for time on Beamline BioMax under Proposal 20180236. Research conducted at MAX IV, a Swedish national user facility, is supported by the Swedish Research council under contract 2018-07152, the Swedish Governmental Agency for Innovation Systems under contract 2018-04969, and Formas under contract 2019-02496. We also acknowledge the synchrotrons Swiss Light Source (Paul Scherrer Institute, Switzerland) for time at beamline PX1 and the ESRF (France) for time at beamlines ID23 and ID30. This work was supported by grants from the Swedish Research Council (2016-03599), Knut and Alice Wallenberg Foundation, and Kempestiftelserna (SMK-1762 and SMK-1869) to RP-A Berntsson.

Author Contributions

  • A Breidenstein: conceptualization, formal analysis, validation, investigation, visualization, methodology, and writing—original draft, review, and editing.

  • J ter Beek: conceptualization, supervision, project administration, and writing—original draft, review, and editing.

  • RP-A Berntsson: conceptualization, data curation, formal analysis, supervision, funding acquisition, investigation, visualization, project administration, and writing—original draft, review, and editing.

Conflict of Interest Statement

The authors declare that they have no conflict of interest.

  • Received October 19, 2022.
  • Revision received January 10, 2023.
  • Accepted January 10, 2023.
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This article is available under a Creative Commons License (Attribution 4.0 International, as described at https://creativecommons.org/licenses/by/4.0/).

References

  1. ↵
    1. Afonine PV,
    2. Grosse-Kunstleve RW,
    3. Echols N,
    4. Headd JJ,
    5. Moriarty NW,
    6. Mustyakimov M,
    7. Terwilliger TC,
    8. Urzhumtsev A,
    9. Zwart PH,
    10. Adams PD
    (2012) Towards automated crystallographic structure refinement with phenix.refine. Acta Crystallogr D Biol Crystallogr 68: 352–367. doi:10.1107/s0907444912001308
    OpenUrlCrossRefPubMed
  2. ↵
    1. Alvarez-Martinez CE,
    2. Christie PJ
    (2009) Biological diversity of prokaryotic type IV secretion systems. Microbiol Mol Biol Rev 73: 775–808. doi:10.1128/mmbr.00023-09
    OpenUrlAbstract/FREE Full Text
  3. ↵
    1. Amor-Mahjoub M,
    2. Suppini J-P,
    3. Gomez-Vrielyunck N,
    4. Ladjimi M
    (2006) The effect of the hexahistidine-tag in the oligomerization of HSC70 constructs. J Chromatogr B Analyt Technol Biomed Life Sci 844: 328–334. doi:10.1016/j.jchromb.2006.07.031
    OpenUrlCrossRefPubMed
  4. ↵
    1. Arutyunov D,
    2. Frost LS
    (2013) F conjugation: Back to the beginning. Plasmid 70: 18–32. doi:10.1016/j.plasmid.2013.03.010
    OpenUrlCrossRefPubMed
  5. ↵
    1. Boer R,
    2. Russi S,
    3. Guasch A,
    4. Lucas M,
    5. Blanco AG,
    6. Pérez-Luque R,
    7. Coll M,
    8. de la Cruz F
    (2006) Unveiling the molecular mechanism of a conjugative relaxase: The structure of TrwC complexed with a 27-mer DNA comprising the recognition hairpin and the cleavage site. J Mol Biol 358: 857–869. doi:10.1016/j.jmb.2006.02.018
    OpenUrlCrossRefPubMed
  6. ↵
    1. Buchan DWA,
    2. Jones DT
    (2019) The PSIPRED protein analysis workbench: 20 years on. Nucleic Acids Res 47: W402–W407. doi:10.1093/nar/gkz297
    OpenUrlCrossRefPubMed
  7. ↵
    1. Byrd DR,
    2. Matson SW
    (1997) Nicking by transesterification: The reaction catalysed by a relaxase. Mol Microbiol 25: 1011–1022. doi:10.1046/j.1365-2958.1997.5241885.x
    OpenUrlCrossRefPubMed
  8. ↵
    1. Carballeira JD,
    2. González-Pérez B,
    3. Moncalián G,
    4. la Cruz F
    (2014) A high security double lock and key mechanism in HUH relaxases controls oriT-processing for plasmid conjugation. Nucleic Acids Res 42: 10632–10643. doi:10.1093/nar/gku741
    OpenUrlCrossRefPubMed
  9. ↵
    1. Chandran Darbari V,
    2. Waksman G
    (2015) Structural biology of bacterial type IV secretion systems. Annu Rev Biochem 84: 603–629. doi:10.1146/annurev-biochem-062911-102821
    OpenUrlCrossRef
  10. ↵
    1. Christie PJ
    (2004) Type IV secretion: The Agrobacterium VirB/D4 and related conjugation systems. Biochim Biophys Acta 1694: 219–234. doi:10.1016/j.bbamcr.2004.02.013
    OpenUrlCrossRefPubMed
  11. ↵
    1. Christie PJ
    (2016) The mosaic type IV secretion systems. EcoSal Plus 7. doi:10.1128/ecosalplus.esp-0020-2015
    OpenUrlCrossRef
  12. ↵
    1. Costa TRD,
    2. Harb L,
    3. Khara P,
    4. Zeng L,
    5. Hu B,
    6. Christie PJ
    (2021) Type IV secretion systems: Advances in structure, function, and activation. Mol Microbiol 115: 436–452. doi:10.1111/mmi.14670
    OpenUrlCrossRefPubMed
  13. ↵
    1. Datta S,
    2. Larkin C,
    3. Schildbach JF
    (2003) Structural insights into single-stranded DNA binding and cleavage by F factor TraI. Structure 11: 1369–1379. doi:10.1016/j.str.2003.10.001
    OpenUrlCrossRefPubMed
  14. ↵
    1. De La Cruz F,
    2. Frost LS,
    3. Meyer RJ,
    4. Zechner EL
    (2010) Conjugative DNA metabolism in gram-negative bacteria. FEMS Microbiol Rev 34: 18–40. doi:10.1111/j.1574-6976.2009.00195.x
    OpenUrlCrossRefPubMed
  15. ↵
    1. Emsley P,
    2. Cowtan K
    (2004) Coot: Model-building tools for molecular graphics. Acta Crystallogr D Biol Crystallogr 60: 2126–2132. doi:10.1107/s0907444904019158
    OpenUrlCrossRefPubMed
  16. ↵
    1. Garcillán-Barcia MP,
    2. Francia MV,
    3. de La Cruz F
    (2009) The diversity of conjugative relaxases and its application in plasmid classification. FEMS Microbiol Rev 33: 657–687. doi:10.1111/j.1574-6976.2009.00168.x
    OpenUrlCrossRefPubMed
  17. ↵
    1. Geertsma ER,
    2. Dutzler R
    (2011) A versatile and efficient high-throughput cloning tool for structural biology. Biochemistry 50: 3272–3278. doi:10.1021/bi200178z
    OpenUrlCrossRefPubMed
  18. ↵
    1. Gonzalez-Rivera C,
    2. Bhatty M,
    3. Christie PJ
    (2016) Mechanism and function of type IV secretion during infection of the human host. Microbiol Spectr 4. doi:10.1128/microbiolspec.vmbf-0024-2015
    OpenUrlCrossRef
  19. ↵
    1. Grohmann E,
    2. Christie PJ,
    3. Waksman G,
    4. Backert S
    (2018) Type IV secretion in gram-negative and gram-positive bacteria: Type IV secretion. Mol Microbiol 107: 455–471. doi:10.1111/mmi.13896
    OpenUrlCrossRefPubMed
  20. ↵
    1. Guasch A,
    2. Lucas M,
    3. Moncalián G,
    4. Cabezas M,
    5. Pérez-Luque R,
    6. Gomis-Rüth FX,
    7. de la Cruz F,
    8. Coll M
    (2003) Recognition and processing of the origin of transfer DNA by conjugative relaxase TrwC. Nat Struct Mol Biol 10: 1002–1010. doi:10.1038/nsb1017
    OpenUrlCrossRefPubMed
  21. ↵
    1. Guglielmini J,
    2. Quintais L,
    3. Garcillán-Barcia MP,
    4. de la Cruz F,
    5. Rocha EPC
    (2011) The repertoire of ICE in prokaryotes underscores the unity, diversity, and ubiquity of conjugation. PLoS Genet 7: e1002222. doi:10.1371/journal.pgen.1002222
    OpenUrlCrossRefPubMed
  22. ↵
    1. Holm L
    (2020) Using dali for protein structure comparison. Methods Mol Biol 2112: 29–42. doi:10.1007/978-1-0716-0270-6_3
    OpenUrlCrossRefPubMed
  23. ↵
    1. Ilangovan A,
    2. Kay CWM,
    3. Roier S,
    4. El Mkami H,
    5. Salvadori E,
    6. Zechner EL,
    7. Zanetti G,
    8. Waksman G
    (2017) Cryo-EM structure of a relaxase reveals the molecular basis of DNA unwinding during bacterial conjugation. Cell 169: 708–721.e12. doi:10.1016/j.cell.2017.04.010
    OpenUrlCrossRef
  24. ↵
    1. Juhas M
    (2015) Horizontal gene transfer in human pathogens. Crit Rev Microbiol 41: 101–108. doi:10.3109/1040841x.2013.804031
    OpenUrlCrossRefPubMed
  25. ↵
    1. Jumper J,
    2. Evans R,
    3. Pritzel A,
    4. Green T,
    5. Figurnov M,
    6. Ronneberger O,
    7. Tunyasuvunakool K,
    8. Bates R,
    9. Žídek A,
    10. Potapenko A, et al.
    (2021) Highly accurate protein structure prediction with AlphaFold. Nature 596: 583–589. doi:10.1038/s41586-021-03819-2
    OpenUrlCrossRefPubMed
  26. ↵
    1. Kabsch W
    (2010) XDS. Acta Crystallogr D Biol Crystallogr 66: 125–132. doi:10.1107/s0907444909047337
    OpenUrlCrossRefPubMed
  27. ↵
    1. Khara P,
    2. Song L,
    3. Christie PJ,
    4. Hu B
    (2021). In situ visualization of the pKM101-encoded type IV secretion system reveals a highly symmetric ATPase energy center. mBio 12: e0246521. doi:10.1128/mBio.02465-21
    OpenUrlCrossRef
  28. ↵
    1. Larkin C,
    2. Datta S,
    3. Harley MJ,
    4. Anderson BJ,
    5. Ebie A,
    6. Hargreaves V,
    7. Schildbach JF
    (2005) Inter- and intramolecular determinants of the specificity of single-stranded DNA binding and cleavage by the F factor relaxase. Structure 13: 1533–1544. doi:10.1016/j.str.2005.06.013
    OpenUrlCrossRefPubMed
  29. ↵
    1. Li YG,
    2. Christie PJ
    (2020) The TraK accessory factor activates substrate transfer through the pKM101 type IV secretion system independently of its role in relaxosome assembly. Mol Microbiol 114: 214–229. doi:10.1111/mmi.14507
    OpenUrlCrossRef
  30. ↵
    1. Low HH,
    2. Gubellini F,
    3. Rivera-Calzada A,
    4. Braun N,
    5. Connery S,
    6. Dujeancourt A,
    7. Lu F,
    8. Redzej A,
    9. Fronzes R,
    10. Orlova EV, et al.
    (2014) Structure of a type IV secretion system. Nature 508: 550–553. doi:10.1038/nature13081
    OpenUrlCrossRefPubMed
  31. ↵
    1. Macé K,
    2. Vadakkepat AK,
    3. Redzej A,
    4. Lukoyanova N,
    5. Oomen C,
    6. Braun N,
    7. Ukleja M,
    8. Lu F,
    9. Costa TRD,
    10. Orlova EV, et al.
    (2022) Cryo-EM structure of a type IV secretion system. Nature 607: 191–196. doi:10.1038/s41586-022-04859-y
    OpenUrlCrossRef
  32. ↵
    1. McCoy AJ,
    2. Grosse-Kunstleve RW,
    3. Adams PD,
    4. Winn MD,
    5. Storoni LC,
    6. Read RJ
    (2007) Phaser crystallographic software. J Appl Crystallogr 40: 658–674. doi:10.1107/s0021889807021206
    OpenUrlCrossRefPubMed
  33. ↵
    1. Monaco S,
    2. Gordon E,
    3. Bowler MW,
    4. Delagenière S,
    5. Guijarro M,
    6. Spruce D,
    7. Svensson O,
    8. McSweeney SM,
    9. McCarthy AA,
    10. Leonard G, et al.
    (2013) Automatic processing of macromolecular crystallography X-ray diffraction data at the ESRF. J Appl Crystallogr 46: 804–810. doi:10.1107/s0021889813006195
    OpenUrlCrossRefPubMed
  34. ↵
    1. Nash RP,
    2. Habibi S,
    3. Cheng Y,
    4. Lujan SA,
    5. Redinbo MR
    (2010) The mechanism and control of DNA transfer by the conjugative relaxase of resistance plasmid pCU1. Nucleic Acids Res 38: 5929–5943. doi:10.1093/nar/gkq303
    OpenUrlCrossRefPubMed
  35. ↵
    1. Paterson ES,
    2. Moré MI,
    3. Pillay G,
    4. Cellini C,
    5. Woodgate R,
    6. Walker GC,
    7. Iyer VN,
    8. Winans SC
    (1999) Genetic analysis of the mobilization and leading regions of the IncN plasmids pKM101 and pCU1. J Bacteriol 181: 2572–2583. doi:10.1128/jb.181.8.2572-2583.1999
    OpenUrlAbstract/FREE Full Text
  36. ↵
    1. Singh M,
    2. Sori H,
    3. Ahuja R,
    4. Meena J,
    5. Sehgal D,
    6. Panda AK
    (2020) Effect of N-terminal poly histidine-tag on immunogenicity of Streptococcus pneumoniae surface protein SP0845. Int J Biol Macromol 163: 1240–1248. doi:10.1016/j.ijbiomac.2020.07.056
    OpenUrlCrossRef
  37. ↵
    1. Some D,
    2. Amartely H,
    3. Tsadok A,
    4. Lebendiker M
    (2019) Characterization of proteins by size-exclusion chromatography coupled to multi-angle light scattering (SEC-MALS). J Vis Exp. e59615. doi:10.3791/59615
    OpenUrlCrossRef
  38. ↵
    1. Stern JC,
    2. Schildbach JF
    (2001) DNA recognition by F factor TraI36: Highly sequence-specific binding of single-stranded DNA. Biochemistry 40: 11586–11595. doi:10.1021/bi010877q
    OpenUrlCrossRefPubMed
  39. ↵
    1. Terwilliger TC,
    2. Poon BK,
    3. Afonine PV,
    4. Schlicksup CJ,
    5. Croll TI,
    6. Millán C,
    7. Richardson JaneS,
    8. Read RJ,
    9. Adams PD
    (2022) Improved AlphaFold modeling with implicit experimental information. Nat Methods 19: 1376–1382. doi:10.1038/s41592-022-01645-6
    OpenUrlCrossRef
  40. ↵
    1. Waksman G
    (2019) From conjugation to T4S systems in gram-negative bacteria: A mechanistic biology perspective. EMBO Rep 20: e47012. doi:10.15252/embr.201847012
    OpenUrlAbstract/FREE Full Text
  41. ↵
    1. Backert S,
    2. Grohmann E
    1. Zechner EL,
    2. Moncalián G,
    3. de la Cruz F
    (2017) Relaxases and plasmid transfer in gram-negative bacteria. In Type IV Secretion in Gram-Negative and Gram-Positive Bacteria, Backert S, Grohmann E (eds), pp 93–113. Cham: Springer International Publishing.
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Characterization of pKM101 relaxase TraI
Annika Breidenstein, Josy ter Beek, Ronnie P-A Berntsson
Life Science Alliance Jan 2023, 6 (4) e202201775; DOI: 10.26508/lsa.202201775
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