Appl Microbiol Biotechnol (2014) 98:9545–9560 DOI 10.1007/s00253-014-6151-3

MINI-REVIEW

Squaring up to DNA: pentapeptide repeat proteins and DNA mimicry Shama Shah & Jonathan G. Heddle

Received: 8 September 2014 / Revised: 8 October 2014 / Accepted: 9 October 2014 / Published online: 26 October 2014 # Springer-Verlag Berlin Heidelberg 2014

Abstract Pentapeptide repeats are a class of proteins characterized by the presence of multiple repeating sequences five amino acids in length. The sequences fold into a right-handed β-helix with a roughly square-shaped cross section. Pentapeptide repeat proteins include a number of examples which are thought to function as structural mimics of DNA and act to competitively bind to the type II topoisomerase DNA gyrase, an important antibacterial target. DNA gyrase-targeting pentapeptide repeat proteins can both inhibit DNA gyrase—a potentially useful therapeutic property—and contribute to resistance to quinolone antibacterials (by acting to prevent them forming a lethal complex with the DNA and enzyme). Pentapeptide repeat proteins are therefore of wide interest not only because of their unusual structure, function, and potential as an antibacterial target, but also because knowledge of their mechanism of action may lead to both a greater understanding of the details of DNA gyrase function as well as being a useful template for the design of new DNA gyrase inhibitors. However, many puzzling aspects as to how these DNA mimics function and indeed even their ability to act as DNA mimics itself remains open to question. This review summarizes the current state of knowledge regarding pentapeptide repeat proteins, focusing on those that are thought to mimic DNA, and speculates on potential structure-function relationships which may account for their differing specificities. Keywords DNA gyrase . Topoisomerase . Pentapeptide repeat proteins . MfpA . Qnr . DNA mimicry S. Shah : J. G. Heddle (*) Heddle Initiative Research Unit, RIKEN, 2-1, Hirosawa, Wako, Saitama 351-0198, Japan e-mail: [email protected] S. Shah Department of Life Science and Medical Bioscience, Waseda University, 2-2 Wakamatsu-cho, Shinjuku, Tokyo 162-8480, Japan

Introduction DNA has a central role in all cellular life, and cells expend significant resources in maintaining, repairing, modifying, and copying their genomes. Up to 2,600 of the genes in the human genome, for example, may contain a DNA-binding motif (Babu et al. 2004), and many enzymes have DNA as their substrate. It is therefore, perhaps no surprise that nature has also evolved proteins that mimic DNA as a means to compete with it for binding to DNA-binding proteins as possible protective or regulatory measures. There are a wide variety of proteins that bind DNA (and RNA, which is reviewed elsewhere (Nakamura and Ito 2011)), some are vital to cellular life, and some are also specific to particular taxonomic groups. DNA-mimicking proteins specific to certain species or classes maybe useful as lead molecules for the production of therapeutics targeting pathogenic organisms through inhibiting their unique DNA-binding proteins (Putnam and Tainer 2005). It has also been suggested that DNA mimics could also be used in place of natural DNA substrates in difficult to crystallize protein-DNA complexes (Dryden 2006). DNA gyrase targeting pentapeptide repeat proteins (PRPs, Table 1) may be useful for development as inhibitors of gyrase, which is already known as an excellent antibacterial target (Collin et al. 2011). Proteins can mimic DNA via one of two routes: They may be able to mimic the interactions of the DNA with its target (this does not necessarily require overall mimicry of DNA structure throughout the bulk of the molecule), or they may be structural mimics of DNA which are able to satisfy proteinDNA binding interactions through overall similarity to DNA structure. Antirestriction and pentapeptide repeat proteins are two examples of proteins showing overall similarity. A list of proteins mimicking DNA and their main features is given below. Further details can be found in recent reviews (Putnam et al. 1999; Wang et al. 2014a) including details of

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Table 1 PRP proteins known to affect gyrase PRP protein

Organism in which discovered

No. of amino acids

MW of monomer (kDa; according to ProtParam (Gasteiger et al. 2005))

Predicted PI (according to ProtParam (Gasteiger et al. 2005))

QnrA1 QnrB1 QnrC QnrD1 QnrS1 AhQnr MfpA EfsQnr McbG AlbG

Klebsiella pneumoniae Klebsiella pneumoniae Proteus mirabilis Salmonella enterica Shigella flexneri Aeromonas hydrophila Mycobacterium tuberculosis Enterococcus faecalis Escherichia. coli Xanthomonas. albilineans

218 226 221 214 218 216 183 211 187 200

24,774.7 25,169.4 25,226.3 23,934.8 24,739 24,236.1 20,019.5 24,252.4 21,848.2 22,749.5

4.82 5.29 5.08 4.93 5.33 4.31 5.33 4.66 5.69 4.63

For Qnr proteins, one representative of each Qnr type is shown

Ugi. Ugi (uracil-DNA glycosylase inhibitor protein; Fig. 1) was the first DNA-mimicking protein to be discovered (Wang and Mosbaugh 1989). It was detected as a

factor capable of inhibiting the N-glycosidase activity of Bacillus subtilis infected with bacteriophage PBS2 (Friedberg et al. 1975). The DNA of this phage contains deoxyuridine in place of thymidine (Takahashi and Marmur 1963). Host cell uracil-DNA glycosylase (UDG) can act as a defensive measure against PBS2 phage infection by catalyzing release of free uracil from DNA (Friedberg et al. 1975). Ugi was subsequently purified and characterized (Cone et al. 1980; Wang and Mosbaugh 1988; Wang et al. 1991), and the X-ray crystal structure of the protein bound to UDG was solved (Mol et al. 1995). This showed that Ugi acted as a structural and electrostatic mimic of dsDNA, which inserted the edge of a beta strand (β1) into the DNA-binding groove of UDG. The majority of the interaction between the β1

Fig. 1 Comparison of three types of DNA mimics. a Crystal structure of a 12 bp dsDNA (pdb 1bnA) (Drew et al. 1981). b Structure of a small DNA mimic, Ugi (pdb 1ugi) (Putnam et al. 1999). c Structure of MfpA (pdb 2bm5) (Hegde et al. 2005). d Structure of Arda (pdb 2w82) (McMahon et al. 2009). Each protein is shown as (i) cartoon format, (ii) surface charge format, and (iii) surface representation with S atom of cys colored yellow, N and H atoms of free amines in lysine colored blue, and

carboxylate groups of Asp and Glu colored red. Images were produced using PyMOL (Schrodinger 2010) and APBS (Baker et al. 2001). Surface charge formats are shown using solvent-accessible surface charge with negative charge shown in red and positive charge in blue. For ease of visualization, max and min settings for displaying charge differed for ArdA compared to Ugi and MfpA. The same molecules are shown in similar orientations and scale. All molecules are shown approximately to scale

proteins, only portions of which mimic DNA (e.g., TAFII230 (Tora 2002) and P53 (Kruse and Gu 2009)). Examples of different classes of DNA-mimicking proteins are shown in Fig. 1.

Short DNA mimicking proteins Short DNA mimics resemble double-stranded DNA (dsDNA) but only over a short length, typically a single helical turn or less. These include the following:

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edge and the DNA-binding groove is accounted for by hydrogen bonding (Putnam et al. 1999). Overall, Ugi mimics the kinked structure of approx. 10 bp of UDGbound dsDNA with the edges of the protein equivalent to the two phosphate backbones of dsDNA where appropriate negative charges are provided by numerous carboxylate-containing side chains (Putnam et al. 1999). A protein from Bacillus phage ϕ29 known as p56 (Asensio et al. 2011; Serrano-Heras et al. 2007) has also been shown to bind to UDG in competition with DNA (Serrano-Heras et al. 2007), and SAUGI from Staphylococcus is a further example of a DNAmimicking UDG inhibitor (Wang et al. 2014b). DinI. DinI (Kenyon and Walker 1980) is an 81-amino acid protein from E. coli known to inhibit RecA coprotease activity (Yasuda et al. 1998). RecA is an important protein in the SOS response and initiates it via binding to single-stranded DNA (ssDNA) (Cox 2007). DinI appears to achieve inhibition by DNA mimicry; the solution structure (Ramirez et al. 2000) showed that like Ugi, DinI possesses a ridge with negative charges provided by Glu and Asp residues. It was proposed that the ridge mimicked the phosphate backbone of DNA, competing with it for binding to RecA, something which was later confirmed by biochemical experiments (Voloshin et al. 2001) and was suggested to correspond to the “groove mode” of DinI binding from threedimensional reconstruction studies (Galkin et al. 2011). HI1450. HI1450 is a Haemophilus influenzae protein, whose solution structure shows some similarity to Ugi (Parsons et al. 2004). In particular, the distribution of negative charge, mimicking part of a phosphate backbone, is similar, and it has therefore been suggested that it also likely acts as a DNA mimic in an analogous way (Parsons et al. 2004). It binds to HU-α, a bacterial histone-like protein (Dorman and Deighan 2003; Grove 2011). The protein DMP12 found in Neisseria species (Wang et al. 2013) may also mimic DNA bound to HU although this appears to have only a relatively weak affinity. NuiA. This protein inhibits an endonuclease found in Anabaena spp. The nuclease NucA is able to nonspecifically digest both ssDNA and dsDNA as well as RNA, and NuiA likely represents a form of selfimmunity (Muro-Pastor et al. 1997). The protein adopts an alpha-beta-alpha sandwich structure (Kirby et al. 2002), and the crystal structure of the complex between NucA and NuiA shows that a large contribution to binding stems from numerous negatively charged residues which are presumed to mimic the phosphate backbone of substrate DNA (Ghosh et al. 2007). CarS. CarS is one half of the CarA-CarS repressorantirepressor in Myxococcus xanthus. When the cell is

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exposed to blue light, the carB operon is expressed and carotenoid synthesis occurs (Fontes et al. 1993). CarS is the antirepressor in this system, is expressed by the cell in the presence of blue light (Lopez-Rubio et al. 2002), and inhibits the inhibitory action of CarA on carB (Whitworth and Hodgson 2001). CarS contains a five-stranded beta sheet domain similar to SH3 domains (León et al. 2010). The DNA mimicry occurs in CarS due to the arrangement of negatively charged surface residues in a way which mimics the phosphate backbone of distorted, bound dsDNA. This mimicry region binds to the DNA recognition helix α2 in CarA (León et al. 2010). DMP19. DMP19 is a protein found in species of pathogenic Neisseria (Exley et al. 2009; Stabler et al. 2005). The crystal structure of the protein revealed surface negative charges arranged in a fashion mimicking that of dsDNA (Wang et al. 2012). A Neisseria transcriptional repressor which binds to DNA to control expression of a nitrogen-response protein was identified as a target for DMP19 and was shown to have a DNA-binding region consisting of positively charged residues which mirrored the negative charge of DMP19, suggesting a largely electrostatic binding interaction (Wang et al. 2012).

Long DNA mimicking proteins We arbitrarily classify long mimics as those that mimic more than approx. 10 bp of dsDNA. Long mimics include the following: Ocr. Ocr (overcome classical restriction) protein is a bacteriophage T7 homodimer protein encoded by gene 0.3. It mimics a 24-bp bent DNA molecule both in terms of overall structure and in distribution of negative charge. Impressively, Ocr is able to inhibit all known type I restriction enzymes (Walkinshaw et al. 2002). Gam. Gam protein from lambda phage is part of a viral homologous recombination system, and its role is to protect virally generated ssDNA ends from the host RecBCD nuclease (Murphy 1991). The crystal structure of the protein (Court et al. 2007) shows it to have a similar shape and size as Ocr and also has numerous negatively charged surface residues (Court et al. 2007). Gam is able to compete with DNA for binding to RecBCD, and the central part of the protein is proposed to act as a dsDNA mimic with a helix positioned at one end of the elongated structure (helix H1) proposed to mimic ssDNA (Court et al. 2007). ICP11. This is a protein produced by the white spot syndrome virus (Wang et al. 2007). Its role appears to be inhibition of binding to DNA of a number of histone

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proteins in the host cell (Wang et al. 2008). This seems to be achieved by DNA mimicry with the functional dimer displaying negatively charged side chains at positions roughly equivalent to phosphates in dsDNA (Wang et al. 2008). ArdA. Like the Ocr protein, ArdA is able to bind and inhibit type I restriction enzymes (Fig. 1) (Belogurov et al. 1992; Nekrasov et al. 2007). ArdA is a long DNA mimic protein, equivalent to a 42 bp dsDNA (McMahon et al. 2009), and is the only non-pentapeptide repeat DNA-mimicking protein which rivals them in length.

Pentapeptide repeat proteins The first pentapeptide repeats (PRs) were identified in the sequence of the hglK gene in Cyanobacterium anabaena species strain PCC 7120 in 1995 (Black et al. 1995). HglK was suggested to be a membrane protein with a role in transport and/or assembly of glycolipids. The protein, 727 amino acids in length, was found to contain 36 PRs of degenerate sequence, AXLXX preceded by four membranespanning regions. A wider survey of a number of species revealed that PR-containing proteins were numerous and constituted a protein family of both membrane and cytoplasmic proteins, and a PR repeat consensus sequence A(D/N)LXX was described (Bateman et al. 1998). The current consensus sequence is [S,T,A,V][D,N][L,F][S,T,R][G] (Vetting et al. 2006). PRs have since been found as motifs within numerous larger proteins (Vetting et al. 2006). “Pentapeptide repeat protein” (PRP) may refer to both proteins that contain a PR domain as a smaller part of a larger structure and those where the PR constitutes the majority. DNA-mimicking PRPs fall into the latter category. PR containing proteins are found in both prokaryotes and eukaryotes, and bioinformatics approaches have identified hundreds of such proteins (summarized in Vetting (Vetting et al. 2006)). A recent query (June 2014) of the Pfam database (http://pfam.xfam.org/) for members of the pentapeptide family (PF00805) returns 11,082 sequences from 1,513 species with protein structures of a number of proteins having been solved (these include PRPs from Nostoc sp. (Ni et al. 2009; Vetting et al. 2007), Cyanothece 51142 (Buchko et al. 2006; Buchko et al. 2008), Arabidopsis thaliana, Enterococcus faecalis (Vetting et al. 2009), Klebsiella pneumoniae (Vetting et al. 2011a), Xanthomonas albilineans (Vetting et al. 2011b), Aeromonas hydrophila (Xiong et al. 2011), and Mycobacterium tuberculosis (Hegde et al. 2005) (Fig. 2). PRPs are characterized by a right-handed quadrilateral beta helix (an Rfr-fold) giving them their square cross section (Fig. 3). Each face is composed of five amino acids. The central amino acid is termed i, with the preceding two amino

acids being i−2 and i−1 and the following two amino acids as i+ 1 and i+2. The side chains of residues i and i−2 typically form the hydrophobic core of the structure while the side chains of the remaining three residues are part of the exterior (Hegde et al. 2005; Vetting et al. 2006). Four pentapeptide faces complete one 360° turn and are called a coil. PRPs vary in their length and hence the number of coils. Turns in the beta helix are achieved by the beta turn motif (Richardson 1981) which may be either type II or type IV (see Buchko (Buchko et al. 2006) for an in-depth description of the beta turns in PRPs). The N- and C-termini of PRPs typically lie at opposite ends of the stacked coils. Those PRPs that exist as dimers in solution all have C-terminal “dimerization domains” (Fig. 3) which consist of an alpha helix sandwiched between two beta strands. Hydrophobic residues along this structure provide an interface with the corresponding dimerization domain in the partner PRP (Vetting et al. 2011b). PRPs that target topoisomerases and for which high-resolution structures have been reported are all dimers (Fig. 2) and show some conservation in the position of hydrophobic residues in the dimerization domain; this conservation extends to those topoisomerase targeting PRPs for which no structure is yet known, suggesting that all topoisomerase targeting PRPs will adopt the dimer structure (Vetting et al. 2011b). A dimer structure may be necessary to allow a more flexible rather than linear structure that is more able to mimic potential distortions experienced by the DNA as the target enzyme changes conformation (Vetting et al. 2011b).

PRPs and DNA gyrase DNA gyrase is protein nanomotor able to negatively supercoil dsDNA using energy from ATP hydrolysis (Higgins et al. 1978; Mizuuchi et al. 1978; Sugino and Cozzarelli 1980) (Fig. 4). The enzyme is a type II topoisomerase and consists of two proteins, GyrA and GyrB with the functional enzyme being an A2B2 tetramer. The supercoiling mechanism has been studied in detail (Nollmann et al. 2007; Papillon et al. 2013): Gyrase is thought to assemble onto dsDNA which binds to a “saddle” region spanning the GyrA dimer which includes the catalytic tyrosine residues (Horowitz and Wang 1987). Regions of GyrB also contribute to this binding pocket (Heddle and Maxwell 2002; Yoshida et al. 1991). The bound DNA is referred to as the gate or “G” segment. After Gsegment binding, DNA is positively wrapped by the Cterminal domains of GyrA, and a second strand of dsDNA is captured by dimerization of GyrB upon ATP binding (Brino et al. 2000; Wigley et al. 1991). This T-segment is passed through the gap in the cleaved DNA, which is resealed and passed out through the exit gate of the enzyme (Kampranis et al. 1999a; Williams et al. 2001; Williams and Maxwell 1999a; Williams and Maxwell 1999b). This reaction results

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Fig. 2 Crystal structures of nine PRP proteins. Structures are shown in a cartoon format, with each monomer colored either green or cyan and b surface view with S atom of cys colored yellow, N and H atoms of free amines in lysine colored blue, and carboxylate groups of Asp and Glu colored red. Molecules are shown in similar orientations with N-termini toward the top and at the same scale. Proteins are (i) pdb 3J8I, NP275 from Nostoc punctiforme (Vetting et al. 2007), (ii) pdb 2F3L Lumenal Rfr-domain protein from Cyanothece sp. (Buchko et al. 2006), (iii) pdb

3n90, AT2G44920 from Arabidopsis thaliana thylakoid lumen (Ni et al. 2011), (iv) pdb 3DU1, HetL protein from Nostoc sp. strain PCC 7120 (Ni et al. 2009), (v) pdb 2W7Z, Efsqnr from Enterococcus faecalis (Vetting et al. 2009), (vi) pdb2XTW QnrB1 from Klebsiella pneumoniae (Vetting et al. 2011a), (vii) pdb 2XT2, AlbG from Xanthomonas albilineans (Vetting et al. 2011b), (viii) pdb 3PSS, AhQnr from Aeromonas hydrophila (Xiong et al. 2011), and (ix) pdb 2bm5 MfpA protein from Mycobacterium tuberculosis (Hegde et al. 2005)

in a change in the linking number of the DNA of two (Bates and Maxwell 2005). Topoisomerase IV (topo IV) is another type II topoisomerase found in some prokaryotes (Champoux 2001). It consists of ParC and ParE proteins, analogous to GyrA and GyrB, respectively. Topo IV has a high structural similarity to gyrase (Laponogov et al. 2007; Wohlkonig et al. 2010) but differs in the DNA-wrapping domain meaning that it does not carry out negative supercoiling of DNA but is instead mainly responsible for DNA decatenation in the cell (Drlica and Zhao 1997; Levine et al. 1998; Zechiedrich and Cozzarelli 1995). Gyrase and topo IV are the targets of the important quinolone class of antibacterials (Emmerson and Jones 2003), perhaps the most well-known of which is the fluoroquinolone ciprofloxacin (CFX). The mechanism of action of quinolones has been well characterized (Collin et al. 2011); they bind to a pocket consisting of the G-segment and residues of both GyrA (or ParC) and GyrB (or ParE) (Bax et al. 2010b; Heddle and Maxwell 2002; Heddle et al. 2000; Laponogov et al. 2010; Laponogov et al. 2009; Laponogov et al. 2013; Wohlkonig et al. 2010). These structures show that quinolones intercalate via base stacking with one quinolone either side of the cleavage site. This likely distorts the DNA and disfavors the religation of the cleaved strands leading to stabilization of

the enzyme and DNA in a state in which the DNA is cleaved (the so-called “cleavage complex”). Once in this state, further supercoiling cannot occur. Furthermore, when DNA breaks are revealed, they lead to cell death. A recent report has suggested a second mode of binding for fluoroquinolone drugs which may reflect the initial binding conformation to a separate binding pocket also near to the active site, before the DNA is cleaved (Mustaev et al. 2014). A number of protein and peptide inhibitors are known to target gyrase; these include CcdB (Kampranis et al. 1999b; Loris et al. 1999; Ogura and Hiraga 1983), MccB17 (Heddle et al. 2001; Moreno et al. 1995), GyrI (Chatterji and Nagaraja 2002; Romanowski et al. 2002), and the topoisomerasetargeting pentapeptide repeat proteins (TTPRPs) (Vetting et al. 2006). The biggest class of PRPs which target gyrase is the Qnr proteins, the majority of which are plasmid-borne. Non-Qnr proteins include McbG, MfpA, and AlbG, all of which are considered below. Qnr proteins One of the earliest proteins consisting solely or predominantly of PRs was QnrA which was discovered in 1998 (MartínezMartínez et al. 1998) with Shewanella algae having been

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Fig. 3 Features of topoisomerase-targeting PRPs. The structure of QnrB1, a typical TTPRP, is shown (pdb 2XTW (Vetting et al. 2011a). One monomer is colored green, the other cyan. The green monomer is modified as follows: Red “C” and “N” represent the location of C- and Ntermini, respectively. Yellow: one face of the quadrilateral consisting of the sequence EKIDR as shown. Superscripts above the sequence show the nomenclature for naming the position of each residue within the pentapeptide. Blue: loop 1. Magenta: loop 2. Red: the C-terminal dimerization domain

identified as the putative natural reservoir (Poirel et al. 2005). QnrB (Jacoby et al. 2006), QnrC (Wang et al. 2009), QnrD (Cavaco et al. 2009), and QnrS (Hata et al. 2005) have subsequently been discovered and are covered in depth in recent reviews (Rodríguez-Martínez et al. 2011; Strahilevitz et al. 2009). QnrA is a 218-amino acid protein which has a protective effect on the activity of DNA gyrase against the inhibitory action of quinolone antibacterials (Tran and Jacoby 2002; Tran et al. 2005). A tagged QnrA (C-terminal His6 or Tag-100-QnrA-His6) was found to protect gyrase against the inhibitory effects of CFX on DNA supercoiling (Tran and Jacoby 2002; Tran et al. 2005). It was shown that this inhibition did not require the presence of DNA, ATP, or CFX and that QnrA was able to bind to isolated GyrB and GyrA subunits as well as to the complex. Filter binding assays showed that QnrA decreased gyrase binding to DNA and the presence of QnrA did not appear to affect the formation of the gyrase holoenzyme. Similar results were obtained against E. coli topo IV with QnrA-His6 showing binding to the enzyme independent of the presence of DNA or CFX. Binding was also shown to be able to occur to isolated ParE and ParC subunits (Tran et al. 2005). These results led to the suggestion that as QnrA binding does not require conformations

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associated with ATP binding and hydrolysis, DNA binding, and CFX binding, it binds to a conformation preceding the formation of the cleavage complex in the catalytic cycle. Possibly one in which the enzyme is in the early stages of interaction with DNA. Given the model for highly similar structures (such as MfpA) where binding is supposed to be across the active site for DNA cleavage, it is not clear how this could function given that the displacement of DNA from the site should abrogate supercoiling (see below for detailed discussion). QnrB is another PRP which is a plasmid-mediated quinolone resistance (PMQRP) protein, discovered in K. pneumoniae, which is capable of protecting purified gyrase against inhibition by CFX (Jacoby et al. 2006). Strains harboring qnrB show resistance to quinolones. The amino acid sequence of QnrB1 is 39.5 % identical to QnrA1 (Jacoby et al. 2006). qnrC (Wang et al. 2009), qnrD (Cavaco et al. 2009), and qnrS (Hata et al. 2005) are all examples of subsequently discovered genes encoding Qnrs found in Proteus mirabilis, Salmonella enterica, and Shigella flexneri, respectively, and having 64, 45, and 59 % amino acid sequence identity to QnrA, respectively. qnrA was discovered as an example of a PMQRP, and the majority of Qnr proteins fall into this category. However, genes for similar proteins have been found on bacterial chromosomes. These include efsqnr, which is found in E. faecalis and shows similarities to both typical plasmid-borne Qnr proteins and MfpA (Hegde et al. 2011) (see below). AhQnr is another chromosomally encoded Qnr. Like plasmid-borne Qnrs, its structure includes two external loops (Xiong et al. 2011); the N-terminal loop is referred to as loop 1, while the C-terminal loop is loop 2. Interaction of PRPs with DNA gyrase PRPs known to interact with gyrase include the following: McbG. McbG protein is a 187-amino acid protein that confers immunity to microcin B17 (MccB17) (Garrido et al. 1988). MccB17 is a member of the microcin family, a class of small peptide antibiotics synthesized by Enterobacteriaceae strains (Kolter and Moreno 1992). MccB17 is 3.1 kDa post-translationally modified peptide (Moreno et al. 1995) and targets DNA gyrase, acting, like the quinolones, as a gyrase poison (Heddle et al. 2001) although the details of mechanism and binding site await elucidation (Parks et al. 2007). McbG gives immunity to MccB17-producing cells (Garrido et al. 1988) presumably in a fashion analogous to the way in which Qnr proteins protect against the action of quinolones. Indeed, McbG is likely to give immunity to some quinolones (Lomovskaya et al. 1996).

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A

i

ii

iii

B

Fig. 4 Potential mechanisms of TTPRP action. a Crystal structure of the 59 kDa N-terminal region of GyrA using APBS (Baker et al. 2001) to show electrostatic potential from −5 (red) to +5 (blue) shown in mutually orthogonal views in (i) and (ii). (iii) Shows the approximate predicted position of MfpA and Qnrs represented as a green rectangle lying across the DNA-binding saddle. b Mechanism of supercoiling by DNA gyrase: (i) GyrB and GyrA dimers assemble on a piece of DNA. The G-segment (“G”) binds to the active site at the DNA gate (“D”). (ii) A protein-DNA complex is formed. (iii) GyrB N-terminal regions (shown in purple, “N”) dimerize and capture the T-segment (“T”) upon ATP binding. The Gsegment is cleaved across both strands. (iv) The DNA gate is opened and

the T-segment is transported into the lower cavity of the enzyme. (v) The exit gate (“E”) opens and the T-segment passes out. c Inhibition of supercoiling via DNA competition. A PRP (green rod) competes with DNA for binding to the G-segment binding site. The resulting complex is incapable of supercoiling. d Protection against quinolones. Fluoroquinolone binds to the gyrase-DNA complex at the point at which the Gsegment is cleaved, stabilizing it in this conformation. Here, the PRP is able to recognize this conformation (red arrow) and destabilize the complex, causing loss of quinolone binding, which allows the enzyme to continue its catalytic cycle

AlbG. AlbG is a protein found in the plant pathogen X. albilineans. It provides resistance to albicidin, a small molecule pathogenesis factor employed by the bacteria (Birch and Patil 1987a; Birch and Patil 1987b). Albicidin appears, like fluoroquinolones, to act as a gyrase poison (Hashimi et al. 2007). The structure of AlbG has been solved (Vetting et al. 2011b) and is a PRP, similar in structure to other gyrase targeting PRPs such as MfpA and EfsQnr. The AlbG structure (Vetting et al. 2011b) highlights the importance of the C-terminal regions of the AlbG monomer; this region that contains a dimerization

module consisting of a strand-helix-strand motif was found to be conserved in topoisomerase poisoning PRPs, and the dimer interface is small and imparts flexibility to the dimer, something which may be important for binding/inhibition of gyrase. MfpA. Mycobacterium fluoroquinolone resistance protein A (MfpA) is a TTPRP that was originally identified as a factor giving resistance to fluoroquinolones in Mycobacterium smegmatis (Montero AAC 45 3387 (2001). The M. smegmatis protein consists of 192 amino acids and 32 uninterrupted PRs (Vetting et al. 2006). MfpA from

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M. tuberculosis consists of 183-amino acid residues and is 67 % identical to M. smegmatis MfpA protein. Crystallographic studies reveal that MfpA has the familiar right-handed quadrilateral β-helix consisting of consecutive pentapeptides with a slight left-handed twist to the helical axis. MfpA forms a dimer in solution due to the Cterminal dimerization domain. The dimer has a rod-like shape with a length of 100 Å; diameter varies from 27 to 18 Å from the N-terminus to the dimer interface (Hegde et al. 2005). Due to the dimensions and distribution of negative charge, it was found that MfpA could be convincingly modeled into the G-segment binding pocket of GyrA (Hegde et al. 2005). This leads to a model wherein part of MfpA lies in close contact with electropositive groove of gyrA59 and blocks formation of the gyrase poisoning cleavage complex by inhibiting G-segment binding. Biochemical tests show that MfpA is able to inhibit negative supercoiling by DNA gyrase (Hegde et al. 2005). Unlike other topoisomerase targeting peptides, MfpA shows no significant protection of negative supercoiling by the enzyme from quinolones (Mérens et al. 2009). As M. tuberculosis contains only gyrase as its sole type II topoisomerase, it would be interesting to know if, like QnrA (Tran et al. 2005), MfpA is also able to inhibit topo IV; however, this is currently not known.

The conundrum of TTPRP action MfpA is similar in structure to other TTPRPs, but while MfpA consists almost exclusively of stacked coil structures, many other TTPRPs are interrupted by non-coil sequences, which form protruding loops. The majority of TTPRP structures solved to date have such loops with the exception of MfpA (Hegde et al. 2005) and EfsQnr (Vetting et al. 2009). MfpA differs from these loop-containing proteins in that it is, in vitro, able to inhibit the supercoiling function of gyrase at relatively low concentrations but does not protect the functioning of the enzyme from fluoroquinolones (Hegde et al. 2005; Mérens et al. 2009). The differences in effects between these different TTPRPs are intriguing given that the similarity in structure strongly suggests that they share a common binding site. When the structure of MfpA was solved, it was suggested that MfpA would compete with dsDNA for binding to the saddle region of the enzyme where DNA is bound and cleaved (Hegde et al. 2005). This is supported by experiments showing that mutation of D87 of M. tuberculosis GyrA gives resistance to MfpA. This residue is close to the active site of gyrase where the G-segment is bound and cleaved. It was proposed that as mutations at GyrA87 are known to increase

gyrase-DNA stability, this mutation makes the DNA less likely to be displaced by MfpA (Mérens et al. 2009). Binding of TTPRPs to this region would clearly inhibit DNA supercoiling, but if DNA is displaced from the binding site, it is unclear how this can result in resistance to the effects of fluoroquinolones while still allowing supercoiling to proceed as supercoiling would clearly be impossible without a bound gate strand (Fig. 4). How then do these other “quinolone rescue” TTPRPs achieve this? One step in solving this mystery has been taken; the loop structures on TTPRPs have been shown to be responsible for the protective effect against fluoroquinolones. This has been shown in the case of AhQnr (Xiong et al. 2011) and QnrB1 (Vetting et al. 2011a, b). Loop 2, in particular, seems to be responsible for the protective effect. In both cases, where loops were mutated and/or deleted, the protective effect was lost. However, the resulting loopfree proteins did not behave like MfpA; i.e., they did not gain the ability to inhibit the supercoiling reaction of the enzyme. EfsQnr which is structurally similar to MfpA, also lacking external loops, is more similar to it in in vitro behavior than other known TTPRPs. EfsQnr is able to inhibit supercoiling activity of gyrase at similar concentrations to MfpA while also acting like a typical Qnr protein in being able to protect the enzyme from the inhibitory effects of fluoroquinolones at submicromolar concentrations (Hegde et al. 2011). Alanine scanning of the residues in loop 2 (106–116) has been carried out in QnrS1 (Tavío et al. 2014) and shows that the protective effect against CFX is severely inhibited by a number of these mutations with Val108-Ala and Cys115-Ala causing a particularly large increase in the concentration of QnrS1 that rescues half of the supercoiling inhibition. It may be that the existing model for MfpA is only applicable to the loop-containing Qnrs when they are at high concentration and exhibit MfpA-like supercoiling inhibition. A different binding mode may obtain for the quinolone rescue effects of Qnrs. Such a model envisages that protection of gyrase from quinolones is not through competition for the Gsegment binding site. Instead it has been suggested that a Qnr may recognize and destabilize the cleavage complex, causing dissociation of drug (Vetting et al. 2011a). Presumably, this effect is mediated by external loops of Qnrs that are not present in MfpA. A second problem arises when comparing the in vitro and in vivo MfpA results. In vivo, the presence of mfpA does in fact show protection against the action of quinolones (Montero et al. 2001). However, it was recently discovered that this requires the presence of a second protein factor called MfpB (Tao et al. 2013) which essentially fulfills the functional role of the missing loops on MfpA. In vitro experiments have confirmed this result. MfpB is a GTPase and the GTP-bound form is necessary for interaction with MfpA (Tao et al. 2013). Care must be taken in testing a PRP from one species against the gyrase of another; for example, QnrB4 from

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enterobacteria is able to protect E. coli gyrase from low concentrations of fluoroquinolones (Mérens et al. 2009) but does not show this effect when tested against M. tuberculosis gyrase (Mérens et al. 2009). There may also be differences in the apparent effect of PRPs in vivo versus in vitro (as discussed above for MfpA). Another potential confounding factor lies with the question of whether the TTPRPs are tested as monomers or dimers. To the best of our knowledge, of the Qnrs, only AhQnr has been tested to determine its oligomeric state in solution, with results indicating that it is a dimer (Xiong et al. 2011). Of the three studies on MfpA, one (Hegde et al. 2005) produced N-terminally His-tagged protein, and the His-tag was cleaved before crystallization and biochemical testing, with the crystal structure showing it to be a dimer. In the other reports on TTPRPs where more in-depth biochemical assays have been carried out (Mérens et al. 2009; Tao et al. 2013; Tavío et al. 2014; Tran and Jacoby 2002; Tran et al. 2005; Xiong et al. 2011), C-terminal His-tagged proteins where the tags are not removed are used. It is possible that the presence of these tags could interfere with dimerization, but in no reports was the tagged protein tested to see if it existed as a dimer or monomer in solution. It is notable that in experiments where C-terminally tagged MfpA proteins were used against E. coli gyrase, MfpA activity against the enzyme appears to be associated with the production of nicked and/ or linear DNA (Mérens et al. 2009; Tao et al. 2013). One possible explanation then for the discrepancy between MfpA and Qnr action could lie with the effect of the C-terminal purification tag; i.e., one C-terminal tagged protein is able to form dimers while the other is prohibited from dimer formation. It is notable that EfsQnr is an exception in that it was assessed with the His-tag cleaved and showed somewhat different activity from other Qnrs, being more MfpA-like (i.e., showing some ability to inhibit gyrase supercoiling at low concentration while also demonstrating a quinolone rescue effect). It is interesting to note that of the monomer PRP structures solved to date, all except HetL have either no or only one loop. HetL, like the Qnr proteins, has two loops, although it appears to lack a C-terminal dimerization domain and is therefore expected to be a monomer. It would be interesting to assess HetL for interaction with gyrase. Recently, a somewhat different mechanism of action has been proposed for TTPRPs to try to explain how protection against the action of fluoroquinolone is achieved without affecting supercoiling. In this suggestion (Vetting et al. 2011a), it is proposed that rather than mimicking the initial bound state of DNA and competing for its binding site, the TTPRPs recognize the quinolone-stabilized cleaved complex and interact with it, somehow causing destabilization and loss of quinolone from the binding site thus allowing the supercoiling reaction to proceed (Vetting et al. 2011a).

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Whether this mechanism would require DNA mimicry at all is unclear.

Phylogenetic and structural relationship between TTPRPs Since the initial discovery of QnrA, Qnr proteins have been found in numerous species and various subtypes of each Qnr protein are known which differ in only a small number of amino acids: an updated list is maintained at http://www.lahey. org/qnrStudies (Jacoby et al. 2008). We have taken representatives of each Qnr type along with several other TTPRPs and used them to produce an updated phylogenetic tree (Fig. 5 using the software Mega 6 (Tamura et al. 2013) which confirms previous findings that EfsQnr is somewhat distant from the plasmid-borne Qnrs (Cavaco et al. 2009), as is MfpA (Wang et al. 2009). It also confirms that within the Qnrs, QnrA, QnrC, and QnrS form one family with QnrB and QnrD forming another (Wang et al. 2009). We have also included McbG, AhQnr, and AlbG in our analyses and find that AhQnr is most closely related to existing plasmid-borne Qnrs but forms a separate branch. McbG is most closely related to MfpA while AlbG is most closely related to EfsQnr. As the structure of a number of TTPRPs is known, we used I-TASSER (Roy et al. 2010; Zhang 2008) to carry out homology modeling of example molecules for which no structural data exists. Inputting examples of plasmid-borne Qnr proteins (QnrA1, QnrC, QnrD1, QnrS1 QnrVc1) all returned

Fig. 5 Maximum parsimony analysis of MfpA taxa: The evolutionary history was inferred using the maximum parsimony method. Sequences were aligned using ClustalW (Larkin et al. 2007). The most parsimonious tree with length=744 is shown. The consistency index is (0.833058), the retention index is (0.547085), and the composite index is 0.472817 (0.455754) for all sites and parsimony-informative sites (in parentheses). The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (500 replicates) are shown next to the branches (Felsenstein 1985). The MP tree was obtained using the subtree pruning-regrafting (SPR) algorithm (pg. 126 in (Nei and Kumar 2000)) with search level 1 in which the initial trees were obtained by the random addition of sequences (ten replicates). The analysis involved 11 amino acid sequences. All positions containing gaps and missing data were eliminated. There were a total of 162 positions in the final dataset. Evolutionary analyses were conducted in MEGA6 (Tamura et al. 2013)

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predictions of high structural similarity to QnrB1 (example shown in Fig. 6), suggesting that all Qnr proteins likely have similar structure-function relationships. Inputting MfpA into I-TASSER and disallowing a selfmatch return the HetL structure as the closest match (Fig. 6), but I-TASSER correctly does not try to match the MfpA structure to the loop regions of HetL giving confidence in the homology modeling process. Inputting the amino acid sequence of McbG returns the best homology prediction with MfpA and predicts only a single loop for McbG consisting of residues 82–87 (MFPCTF; Fig. 6). This is consistent with the phylogenetic analysis, which predicts that MfpA and McbG are closely related. Interestingly, AlbG also has a single loop at a similar position (residues 87–99, TSAQWPSVKMEGA). All of the structures that we aligned and whose homologybased structures were predicted maintained the PR pattern, i.e., where i−2 and i residues are internal and predominantly hydrophobic and i−1 and i+1 residues were external and predominantly polar (Fig. 7). The requirement of loops for quinolone resistance has recently been investigated further. One report into QnrS1 showed that the effect of various point mutations in the Cterminal loop (loop 2, residues 102–113) affected the minimal inhibitory concentration (MIC) for CFX in a BL21 (DE3) background. This could lead to either an increase or a decrease in the MIC depending on the position of the mutation (Tavío et al. 2014; Vetting et al. 2011a). Deletion of the N-terminal loop (loop 1, residues 46–51) caused a significant decrease in the ability of the protein to rescue gyrase from CFX while deletion of loop 2 resulted in complete loss of protection (Vetting et al. 2011a). One model of QnrC suggests that it is closer to MfpA in structure (i.e., lacking loops) (Guo et al. 2010); however, our own modeling and the known

Fig. 6 I-TASSER homology models. a An example of two Qnr proteins aligned (QnrVc1 (cyan (Fonseca et al. 2008)) and QnrB1 (green, pdb 2XTW (Vetting et al. 2011a)). b MfpA (green, pdb) aligned with HetL (cyan, pdb 3DU1 (Ni et al. 2009)). c McbG (cyan) aligns with MfpA (green). All images have N-terminus at the top

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characteristics of QnrC (i.e., protection against quinolones at low concentration) would tend to support the existence of loops. Combining these results with known structures, we can suggest that all known plasmid-borne Qnr proteins and the chromosomally encoded Qnr proteins AhQnr and EfsQnr have a PRP structure, which includes two loops per monomer. We can expect that all of these proteins will show a characteristic protection of gyrase against fluoroquinolones while inhibition of the enzyme (if it occurs at all) will be at higher concentrations. Furthermore, EfsQnr, MfpA, AlbG, and McbG have no (MfpA, EfsQnr) or one (AlbG, McbG) small loop per monomer, suggesting that they may function differently. Indeed, for MfpA, this is known as described above. EfsQnr is also known to have unusual properties which lie somewhere between those of the other Qnrs and MfpA; i.e., it does protect against quinolones at low concentrations but is also able to inhibit supercoiling at relatively low concentrations (Arsène and Leclercq 2007). AlbG is known not to protect gyrase from CFX inhibition (Hashimi et al. 2007), and so, we may predict that AlbG and McbG will similarly diverge from standard Qnrs and be closer to MfpA. This perhaps is not surprising given that they are resistance factors to gyrase poisons that are significantly different to quinolones in structure. This observation suggests that loops themselves directly interfere with gyrase poison binding pockets.

Alternative models for gyrase-targeting PRPs We suggest a model (Fig. 8), which takes into account the requirement of loop 2 for protection of gyrase against fluoroquinolone activity. In this model, the TTPRP again acts as a

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Fig. 7 I-TASSER homology model of QnrC aligned with template QnrB1. Models are shown looking along the longitudinal axis from the N-terminus and are represented in cartoon format. QnrC is shown in green and QnrB1 is shown in cyan. a Residues i, i+1, and i+2 are shown in blue, magenta, and orange, respectively, for QnrC and red, yellow, and

dark gray, respectively, for QnrB1. b Residues i−1 and i−2 are shown in magenta and blue, respectively, for QnrC and yellow and red, respectively, for QnrB1. Internal and external positioning of residues match for the template (QnrB1) and modeled (QnrC) proteins. Similar results are found for all other proteins modeled

DNA mimic, but in this case, it mimics the T-segment and is captured by a gyrase-DNA complex to which the G-segment is already bound. Upon capture and approaching the G-segment, loop 2 of the Qnr recognizes structural features unique to the quinolone complex (this could be the quinolone, the distorted DNA, features of the protein revealed by distortion of the DNA or a combination). The interaction of loop 2 destabilizes this complex. Destabilization could result in dissociation of the whole enzyme-drug DNA complex or simply dissociation of the drug whereby the supercoiling cycle is presumably either reset or continues and the Qnr is passed

out of the “top” or “bottom” gate of the enzyme. In the case of MfpA, the structural similarity to Qnrs would suggest that it too would mimic the T-segment and would be captured by the DNA clamp; however, lacking loop 2 it would not be able to recognize a quinolone-bound complex and so would not be able to rescue the enzyme from quinolone inhibition. In the absence of quinolones, MfpA, however, can inhibit gyrase. We can speculate that this may be due to the location and/or duration which the PRP dwells in gyrase. An example could be if the presence of loop 2 triggers a “resetting” of the enzyme in which the Qnr is passed out of the ATP clamp

Fig. 8 Possible model for the interaction of PRP proteins with gyrase for those proteins that have a protective effect against quinolones. The crystal structure of Staphylococcus aureus DNA gyrase (GyrB27–A56 (GKdel/ Tyr123Phe) bound to G-segment DNA and two CFX molecules (pdb 2XCT (Bax et al. 2010a)) is shown. The gyrase dimer is colored yellow, and the phosphate backbone of the DNA is shown in orange with the CFX molecules in red. The crystal structure of the PRP, AhQnr dimer (pdb 3PSS (Xiong et al. 2011)), is shown with one monomer colored

green, the other cyan. Structures are shown along the longitudinal axis of AhQnr (a) or rotated approximately 90° to it around the gyrase dimer axis (b). AhQnr is placed over the G-segment in a position thought to mimic that of the T segment as it approached the DNA gate. Loop 2 of the green AhQnr monomer (colored black) positions close to the bound CFX. c A zoom of the region in b which includes the proposed possible interaction region between loop 2 and the DNA/protein

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while, in contrast, MfpA, lacking the loop, would instead either bind more stably to the ATP clamp or, alternatively, pass into the exit gate and dwell there for a sufficient period of time to significantly inhibit supercoiling. The reverse model where Qnrs are passed and MfpA remains “above” the DNA gate is also possible. A weakness of this model is that the positioning of the TTPRP is such that only one subunit of the dimer appears to interact with gyrase (Fig. 8). It may be that dimer formation is required for a different stage of the inhibition cycle (e.g., initial capture) or indeed that such TTPRPs are in fact able to function as monomers. Alternative T-segment mimicking mechanisms can easily be envisaged where the TTPRP functions as a dimer by positioning centrally over the DNA gate such that the GyrA and PRP dimer interfaces align. This would place both loop 2s (one from each monomer) at positions where they may interact with the C-terminal region of GyrB and/or GyrA “horns.” This would require a more indirect mechanism of TTPRP quinolone protection than the model illustrated. A further alternative model would utilize the exit (“C-gate”) of gyrase. It is possible that Qnrs and/or MfpA enter through this gate and bind in the exit cavity of the enzyme (MfpA) or disrupt the cleavage complex (Qnrs) from “below.” Entrance of T-segment into the complex via the Cgate is thought to be possible, e.g. in the presence of ADPNP (Williams et al. 2001).

Conclusion The first PRP was identified 19 years ago, and while the structure and function of many TTPRPs are now known in detail, the mechanism of how effects such as gyrase inhibition and rescue of gyrase from poisons are achieved remains puzzling. In particular, the model where TTPRPs mimic the G-segment and bind to the G-segment binding and cleavage site across the GyrA saddle (Hegde et al. 2005) appears incompatible with the results of quinolone protection of supercoiling observed for a number of Qnrs. Indeed, an alternative model whereby the Qnrs achieve supercoiling protection by recognizing and destabilizing the gyrase-DNAquinolone complex has been proposed although a T-segment mimicry model was not suggested (Vetting et al. 2011a). From our own study of recent results, we are lead to suggest that TTPRPs function by T-segment mimicry. Future research is perhaps best addressed by considering the mechanism of quinolone inhibition separately from the mechanism of action of TTPRP inhibitors such as MfpA. For the former, the mode of action as originally envisaged for MfpA (Hegde et al. 2005) seems incompatible with in vitro results where quinolone protection without supercoiling inhibition occurs. Therefore, the priority should be to construct and test a new model for the mechanism of action of these

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proteins where, clearly, the function of the external loops will be key. If the G-segment-mimicry model for MfpA is incorrect and MfpA does not have quinolone rescue activity, then the interaction of MfpA with gyrase in toto can be questioned. The evidence accumulated in favor of MfpA inhibition of gyrase thus far is limited (Hegde et al. 2005; Mérens et al. 2009; Tao et al. 2013) but would argue against this, particularly in the case of MfpA inhibition of M. tuberculosis gyrase (Mérens et al. 2009). The T-segment mimicry model for TTPRPs in general, including MfpA, should be tested perhaps by experiments that are able to follow the passage of such a mimic though the various gates of gyrase (it is possible that the gate utilized to exit the complex would be different between quinolone rescue TTPRPs and supercoiling inhibitor TTPRPs). The question of whether TTPRPs function as monomers or dimers could be addressed firstly by assessing TTPRPs with Cterminal tags to ascertain their oligomeric state. Once the mechanism of action of either TTPRPs or MfpA is fully understood, the apparent incompatibilities in their functioning should be resolvable. However, for MfpA, one mystery will still remain: that of the role of MfpB. MfpB appears to be able to endow MfpA with gyrase rescue from quinolone functionality. How this approx. 20 kDa protein mimics the function of one or two small loops on other PRPs amounting to only a few amino acids in length is unclear. For the moment, the interaction of TTPRPs with gyrase remains puzzling. We envisage that elucidating it could give a deeper understanding of gyrase mechanism as well as opening up new avenues for production of novel antibacterials. Acknowledgments We thank Anthony Maxwell, James Berger, TingYu Lin, and Soshichiro Nagano for critical reading of the manuscript, Soshichiro Nagano for useful discussions and assistance with sequence alignments, and Ting-Yu Lin for elements of Fig. 4. SS and JGH were funded by RIKEN Initiative Research Funding awarded to JGH, and SS was funded as a RIKEN Junior Research Associate.

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