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• W2017161880 abstract "ParG is the prototype of a group of small (<10 kDa) proteins involved in accurate plasmid segregation. The protein is a dimeric DNA binding factor, which consists of symmetric paired C-terminal domains that interleave into a ribbon-helix-helix fold that is crucial for the interaction with DNA, and unstructured N-terminal domains of previously unknown function. Here the ParG protein is shown to be a transcriptional repressor of the parFG genes. The protein assembles on its operator site initially as a tetramer (dimer of dimers) and, at elevated protein concentrations, as a pair of tetramers. Progressive deletion of the mobile N-terminal tails concomitantly decreased transcriptional repression by ParG and perturbed the DNA binding kinetics of the protein. The flexible tails are not necessary for ParG dimerization but instead modulate the organization of a higher order nucleoprotein complex that is crucial for proper transcriptional repression. This is achieved by transient associations between the flexible and folded domains in complex with the target DNA. Numerous ParG homologs encoded by plasmids of Gram-negative bacteria similarly are predicted to possess N-terminal disordered tails, suggesting that this is a common feature of partition operon autoregulation. The results provide new insights into the role of natively unfolded domains in protein function, the molecular mechanisms of transcription regulation, and the control of plasmid segregation. ParG is the prototype of a group of small (<10 kDa) proteins involved in accurate plasmid segregation. The protein is a dimeric DNA binding factor, which consists of symmetric paired C-terminal domains that interleave into a ribbon-helix-helix fold that is crucial for the interaction with DNA, and unstructured N-terminal domains of previously unknown function. Here the ParG protein is shown to be a transcriptional repressor of the parFG genes. The protein assembles on its operator site initially as a tetramer (dimer of dimers) and, at elevated protein concentrations, as a pair of tetramers. Progressive deletion of the mobile N-terminal tails concomitantly decreased transcriptional repression by ParG and perturbed the DNA binding kinetics of the protein. The flexible tails are not necessary for ParG dimerization but instead modulate the organization of a higher order nucleoprotein complex that is crucial for proper transcriptional repression. This is achieved by transient associations between the flexible and folded domains in complex with the target DNA. Numerous ParG homologs encoded by plasmids of Gram-negative bacteria similarly are predicted to possess N-terminal disordered tails, suggesting that this is a common feature of partition operon autoregulation. The results provide new insights into the role of natively unfolded domains in protein function, the molecular mechanisms of transcription regulation, and the control of plasmid segregation. Plasmids are of inherent interest because of their contribution to bacterial genome plasticity, as model systems for investigating a variety of biological processes, and for their utility in gene cloning technology. The role of plasmids in the dissemination of antibiotic resistance and other properties is also particularly significant (1.Hayes F. Methods Mol. Biol. 2003; 235: 1-17PubMed Google Scholar). The events that contribute to stable plasmid inheritance can be unraveled by analyzing the molecular mechanisms that underlie plasmid segregation. The active segregation of low copy number plasmids typically requires two plasmid-encoded proteins, one of which is most commonly a member of the ParA superfamily of Walker-type ATPases. The second protein is a DNA binding factor that interacts directly with a cis-acting partition site (2.Surtees J.A. Funnell B.E. Curr. Top. Dev. Biol. 2003; 56: 145-180Crossref PubMed Scopus (45) Google Scholar). The assembly of the ParA- and DNA-binding proteins on the partition site engenders a nucleoprotein complex that is required for the directional movement of paired plasmids away from the cell median (3.Li Y. Dabrazhynetskaya A. Youngren B. Austin S. Mol. Microbiol. 2004; 53: 93-102Crossref PubMed Scopus (35) Google Scholar). Plasmid movement is most likely mediated by ATP-dependent polymerization of the ParA protein or by its depolymerization (4.Barillà D. Rosenberg M.F. Nobbmann U. Hayes F. EMBO J. 2005; 24: 1453-1464Crossref PubMed Scopus (118) Google Scholar). Unknown host factors are also probably required during segregation, for example as tethers for plasmid pairing at the mid-cell position. The partition locus of multidrug-resistance plasmid TP228 consists of the tandem parF and parG genes and additional essential sequences located upstream of these genes (5.Hayes F. Mol. Microbiol. 2000; 37: 528-541Crossref PubMed Scopus (75) Google Scholar). The ParF protein (22.0 kDa), a member of a distinctive subgroup of the ParA family, and ParG protein (8.6 kDa) assemble on the upstream region; ParG binds to this locus, and ParF is recruited through interactions with ParG, thereby forming a nucleoprotein complex whose precise architecture is likely to be crucial for correct partitioning (6.Barillà D. Hayes F. Mol. Microbiol. 2003; 49: 487-499Crossref PubMed Scopus (36) Google Scholar). ParG is dimeric (6.Barillà D. Hayes F. Mol. Microbiol. 2003; 49: 487-499Crossref PubMed Scopus (36) Google Scholar), consisting of a pair of unstructured N-terminal tails and intertwined, symmetric C-terminal domains that adopt a ribbon-helix-helix (RHH) 1The abbreviations used are: RHH, ribbon-helix-helix; CDO, catechol 2,3-dioxygenase; EMSA, electrophoretic mobility shift assay; IR, inverted repeat; HSQC, heteronuclear single quantum correlation. fold similar to that of the Arc/MetJ family of DNA binding transcriptional repressors (7.Golovanov A.P. Barillà D. Golovanova M. Hayes F. Lian L.Y. Mol. Microbiol. 2003; 50: 1141-1153Crossref PubMed Scopus (72) Google Scholar). The interaction of ParG with its binding sites in the region 5′ of parFG is presumed to occur via the double-stranded antiparallel β-structure of the RHH domain that is inserted into the DNA major groove in a manner analogous to Arc/MetJ-type transcriptional repressors (8.Somers W.S. Phillips S.E. Nature. 1992; 359: 387-393Crossref PubMed Scopus (284) Google Scholar, 9.Gomis-Ruth F.X. Sola M. Acebo P. Parraga A. Guasch A. Eritja R. Gonzalez A. Espinosa M. del Solar G. Coll M. EMBO J. 1998; 17: 7404-7415Crossref PubMed Scopus (135) Google Scholar, 10.Raumann B.E. Rould M.A. Pabo C.O. Sauer R.T. Nature. 1994; 367: 754-757Crossref PubMed Scopus (257) Google Scholar). Plasmid partition genes are organized in operons whose expression is precisely regulated. The control of partition gene expression is mediated, at least in part, by transcriptional autoregulation exerted by one or both of the partition proteins (11.Kusukawa N. Mori H. Kondo A. Hiraga S. Mol. Gen. Genet. 1987; 208: 365-372Crossref PubMed Scopus (38) Google Scholar, 12.Friedman S.A. Austin S.J. Plasmid. 1988; 19: 103-112Crossref PubMed Scopus (98) Google Scholar, 13.Hayes F. Radnedge L. Davis M.A. Austin S.J. Mol. Microbiol. 1994; 11: 249-260Crossref PubMed Scopus (55) Google Scholar, 14.Kwong S.M. Yeo C.C. Poh C.L. Mol. Microbiol. 2001; 40: 621-633Crossref PubMed Scopus (29) Google Scholar). The importance of this regulation is revealed by the perturbation that takes place in segregation when partition genes are overexpressed, especially when the partition site is coresident (11.Kusukawa N. Mori H. Kondo A. Hiraga S. Mol. Gen. Genet. 1987; 208: 365-372Crossref PubMed Scopus (38) Google Scholar, 13.Hayes F. Radnedge L. Davis M.A. Austin S.J. Mol. Microbiol. 1994; 11: 249-260Crossref PubMed Scopus (55) Google Scholar, 15.Funnell B.E. J. Bacteriol. 1988; 170: 954-960Crossref PubMed Google Scholar, 16.Davis M.A. Radnedge L. Martin K.A. Hayes F. Youngren B. Austin S.J. Mol. Microbiol. 1996; 21: 1029-1036Crossref PubMed Scopus (74) Google Scholar). Inappropriate production of partition proteins might result in their permanent occupation of the partition site, which could interfere with the migration of replication and transcription complexes (17.Youngren B. Austin S. Mol. Microbiol. 1997; 25: 1023-1030Crossref PubMed Scopus (31) Google Scholar, 18.Sawitzke J.A. Li Y. Sergueev K. Youngren B. Brendler T. Jones K. Austin S. J. Bacteriol. 2002; 184: 2447-2454Crossref PubMed Scopus (8) Google Scholar). The experiments presented here establish that the flexible N termini of dimeric ParG are essential for the organization of a stable quaternary protein complex on the parFG operator site and thus play a crucial regulatory role. This is achieved by transient associations between the flexible and folded domains in complex with the target DNA. Strains and Media—Escherichia coli was grown at 37 °C in Luria-Bertani medium with appropriate antibiotics when needed. MacConkey agar plates with 1% maltose were used for two-hybrid assays. Strain DH5α was used for cloning, BL21(DE3) (Novagen) for protein overproduction and reporter assays, and SP850 (19.Shah S. Peterkofsky A. J. Bacteriol. 1991; 173: 3238-3242Crossref PubMed Google Scholar) was employed for two-hybrid analysis. Plasmids and Proteins—The parG gene previously was amplified by PCR and cloned in pET22b(+) for ParG overproduction and in pT18 and pT25 vectors for two-hybrid analysis (6.Barillà D. Hayes F. Mol. Microbiol. 2003; 49: 487-499Crossref PubMed Scopus (36) Google Scholar). Deletions in which 27, 57, and 90 bp were removed from the 5′-end of parG were constructed by PCR amplification and also cloned in pET22b(+), pT18, and pT25 vectors. Deletion proteins were purified as described for full-length ParG (6.Barillà D. Hayes F. Mol. Microbiol. 2003; 49: 487-499Crossref PubMed Scopus (36) Google Scholar). A 117-bp DNA fragment covering the parFG promoter region and the first 30 bp downstream of the parF translational start was amplified using oligonucleotides 5′-CGTCGAATTCCATATTAACCTTTACTC-3′ and 5′-GATAGGATCCTTTCGGATT-3′. This fragment was cloned in the BamHI site of plasmid pDM3.0 (20.Macartney D.P. Williams D.R. Stafford T. Thomas C.M. Microbiology. 1997; 143: 2167-2177Crossref PubMed Scopus (42) Google Scholar) to generate a transcriptional fusion to the xylE reporter gene (pDM-Oper). Two-hybrid Analysis—ParG-mediated association and functional complementation of the T18 and T25 fragments of the catalytic domain of Bordetella pertussis adenylate cyclase leads to cAMP synthesis, which triggers transcriptional activation of the maltose operon (6.Barillà D. Hayes F. Mol. Microbiol. 2003; 49: 487-499Crossref PubMed Scopus (36) Google Scholar). This results in red colonies on MacConkey plates with maltose at 30 °C or white colonies when this operon is not activated (21.Karimova G. Pidoux J. Ullmann A. Ladant D. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 5752-5756Crossref PubMed Scopus (1177) Google Scholar). Pentapeptide scanning mutagenesis of ParG produced by the pT25 vector was performed as described previously (22.Hallet B. Sherratt D.J. Hayes F. Nucleic Acids Res. 1997; 25: 1866-1867Crossref PubMed Scopus (77) Google Scholar, 23.Hayes F. Hallet B. Trends Microbiol. 2000; 8: 571-577Abstract Full Text Full Text PDF PubMed Scopus (39) Google Scholar, 24.Hayes F. Annu. Rev. Genet. 2003; 37: 3-29Crossref PubMed Scopus (131) Google Scholar) with the aim of assaying the effect of the insertions on protein dimerization using two-hybrid analysis. Chemical Cross-linking—Chemical cross-linking was performed using dimethyl pimelimidate (10 mm) (Sigma) as cross-linking agent and 20 μm protein according to the protocol detailed elsewhere (6.Barillà D. Hayes F. Mol. Microbiol. 2003; 49: 487-499Crossref PubMed Scopus (36) Google Scholar). Sedimentation Velocity Analysis—Proteins (∼1 mg/ml) were centrifuged in a 1.2-cm path length two-sector aluminum centerpiece cell in an An-60 Ti analytical rotor at 50,000 rpm in an Optima XL-I ultracentrifuge (Beckman) at 20 °C. Solute concentration changes were detected by Rayleigh interference and by monitoring A275. Results were subjected to g(s*) analysis (25.Stafford III, W.F. Anal. Biochem. 1992; 203: 295-301Crossref PubMed Scopus (521) Google Scholar) using DCDT+ Version 1.13 (26.Philo J.S. Anal. Biochem. 2000; 279: 151-163Crossref PubMed Scopus (237) Google Scholar) and to whole boundary analysis using Sedfit Version 8.0 (27.Schuck P. Biophys. J. 2000; 78: 1606-1619Abstract Full Text Full Text PDF PubMed Scopus (3065) Google Scholar). CDO Reporter Assays—E. coli BL21(DE3) was transformed with pDM3.0 or pDM-Oper or with pDM-Oper and pET22b derivatives carrying full-length parG or 5′-parG deletions. A colony was inoculated in Luria-Bertani broth under selective conditions and incubated at 37 °C until A600 ∼0.5. Cells were incubated for an additional 1 h before pelleting and resuspending in 100 mm potassium phosphate buffer, pH 7.4, with 10% acetone. After sonication, the cell lysate was cleared by centrifugation, and protein concentrations were determined using the Bio-Rad protein assay. CDO activity was measured by monitoring the A375 change for 1 min at 24 °C with 0.2 mm catechol. Measurements were performed at least in triplicate, each from an independent culture containing the plasmids. One CDO unit is the amount of enzyme that oxidizes 1 μmol catechol/min at 24 °C (28.Sala-Trepat J.M. Evans W.C. Eur. J. Biochem. 1971; 20: 400-413Crossref PubMed Scopus (203) Google Scholar). Electrophoretic Mobility Shift Assays (EMSA)—Biotinylated DNA substrates were a 48-bp oligonucleotide corresponding to the parFG operator (FS-48), a replica of this oligonucleotide in which one half of the palindrome was randomized (HS-48), and a shorter fragment (HS-23) covering one half-site (see Fig. 3E). Oligonucleotides (1 nm) were incubated at 25 °C for 20 min in binding buffer (10 mm Tris-HCl, pH 7.5, 50 mm KCl, 1 mm dithiothreitol, 5 mm MgCl2, 2.5% glycerol, 0.05 μg/ml poly(dI-dC) with different amounts of ParG proteins. Reactions were electrophoresed on 6% polyacrylamide native gels in TBE (Tris borate-EDTA) buffer and blotted onto positively charged nylon membranes (Roche Applied Science). DNA was immobilized by UV cross-linking and detected using the LightShift chemiluminescent EMSA kit (Pierce). Surface Plasmon Resonance—Surface plasmon resonance measurements used a Biacore 3000 instrument (Biacore AB). The surface of the CM5 sensor chip (Biacore) was sensitized using the Amine Coupling kit (Biacore AB) and coated with extravidin (Sigma). The same oligonucleotides used in EMSA were bound to the chip surface with the same resonance units to facilitate comparison of results between different substrates. An oligonucleotide with an identical base composition to FS-48 but with an unrelated sequence was bound to one flow cell as a reference. Proteins were injected in running buffer (10 mm HEPES-KOH, pH 7.5, 150 mm KCl, 5 mm MgCl2, 1 mm dithiothreitol, 0.005% Tween 20) at a flow rate of 5 μl/min. Data were reference-subtracted using the oligonucleotide of unrelated sequence and analyzed using Biaevaluation 3.1 software (Biacore AB). Ferguson Analysis—Δ9ParG-FS-48 and Δ9ParG-HS-48 complexes and accurate protein standards were electrophoresed in polyacrylamide gels ranging from 7 to 12% and stained with Coomassie Blue. Logarithms of the relative migrations of each species were plotted against acrylamide concentration. The log10 of the negative slope of each line (retardation coefficient) was plotted against the log10 of the molecular weight, and molecular weights of protein-DNA complexes were determined from the resulting linear plot (29.Orchard K. May G.E. Nucleic Acids Res. 1993; 21: 3335-3336Crossref PubMed Scopus (68) Google Scholar). Circular Dichroism (CD) Measurements—CD spectra in the near UV region (250-320 nm) were measured in a JASCO J-810 spectropolarimeter using a path length of 0.1 cm at 20 °C. ParG (10 μm) and FS-48 (1.25-20 μm) were prepared in CD buffer (20 mm potassium phosphate, pH 7.4, 50 mm KCl) and incubated at 20 °C for 20 min before recording the CD spectra. The stoichiometry of the ParG-FS-48 complex was determined by normalizing the change of ellipticity at 273 nm of the DNA in solution and in complex with ParG. NMR Spectroscopy—NMR measurements were performed on a Bruker DRX600 spectrometer with a triple resonance CryoProbe. Sequence-specific resonance assignment of ParG used three-dimensional 15N-edited TOCSY-HSQC and NOESY-HSQC spectra with uniformly 15N-labeled protein (1 mm). Spectra were acquired at T = 293 K. Proton chemical shifts were referenced to the methyl resonance of sodium 2,2-dimethyl-2-silapentane-5-sulfonate at 0 ppm. Chemical shift mapping analysis used proteins (50-100 μm) in buffer containing 90% H2O, 10% 2H2O, 100 mm NaCl, 50 mm NaH2PO4/Na2HPO4, 1 mm dithiothreitol, 40 mm arginine, 40 mm glutamate, pH 5.5. Arginine and glutamate were added to reduce protein aggregation and to increase the concentration of soluble protein (30.Golovanov A.P. Hautbergue G.M. Wilson S.A. Lian L.Y. J. Amer. Chem. Soc. 2004; 126: 8933-8939Crossref PubMed Scopus (318) Google Scholar). Time domain spectral data were processed by NMRPipe (31.Delaglio F. Grzesiek S. Vuister G.W. Zhu G. Pfeifer J. Bax A. J. Biomol. NMR. 1995; 6: 277-293Crossref PubMed Scopus (11570) Google Scholar), and spectra were visualized and assigned with NmrView (32.Johnson B.A. Blevins R.A. J. Biomol. NMR. 1994; 4: 603-614Crossref PubMed Scopus (2678) Google Scholar). Weighted amide chemical shift differences δ in spectra 1 and 2 resulting from deletion mutations or DNA binding were measured as shown in Equation 1 δ1−2=(δ1H−δ2H)+((δ1N−δ2N)/10)2(Eq. 1) where δH and δN are proton and nitrogen chemical shifts, respectively. If signals disappeared from the spectra, arbitrary values of δ >0.1ppm were assigned solely to represent this. NMR Analysis of Protein Complexes with FS-48—FS-48 oligonucleotide (80 μm) in 2 ml of NMR buffer was split in four equal aliquots. Concentrated protein solutions (50 μl) were added to 0.5 ml of DNA solution so that the concentration of dimeric protein in each case was <75 μm, allowing only one binding site to be occupied on each oligonucleotide. ParG Dimerization Does Not Require the Unstructured N-terminal Tail—ParG self-association can be monitored in vivo using a two-hybrid system based on reconstitution of adenylate cyclase activity in E. coli (6.Barillà D. Hayes F. Mol. Microbiol. 2003; 49: 487-499Crossref PubMed Scopus (36) Google Scholar, 21.Karimova G. Pidoux J. Ullmann A. Ladant D. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 5752-5756Crossref PubMed Scopus (1177) Google Scholar). A set of ParG derivatives with insertions at 13 different positions was constructed by pentapeptide scanning mutagenesis (22.Hallet B. Sherratt D.J. Hayes F. Nucleic Acids Res. 1997; 25: 1866-1867Crossref PubMed Scopus (77) Google Scholar, 23.Hayes F. Hallet B. Trends Microbiol. 2000; 8: 571-577Abstract Full Text Full Text PDF PubMed Scopus (39) Google Scholar, 24.Hayes F. Annu. Rev. Genet. 2003; 37: 3-29Crossref PubMed Scopus (131) Google Scholar). None of nine insertions in the ParG N-terminal tail detectably affected the association with wild-type ParG in two-hybrid analysis (Fig. 1A). In contrast, all four pentapeptide insertions in the folded domain abolished heterodimerization with wild-type ParG (Fig. 1A). The preceding pentapeptide insertions influence the sequence and composition of the ParG flexible tail but alter tail length less dramatically. More radical modifications of the N-terminal tail involved deleting 9, 19, and 30 residues (Δ9ParG, Δ19ParG, and Δ30ParG, respectively) (Fig. 1A). These deletions were chosen on the basis of NMR data that showed variations in the dynamic characteristics of the N-terminal domain, suggesting that this domain is not wholly a random coil (7.Golovanov A.P. Barillà D. Golovanova M. Hayes F. Lian L.Y. Mol. Microbiol. 2003; 50: 1141-1153Crossref PubMed Scopus (72) Google Scholar). The deletion variants both interacted with wild-type ParG (Fig. 1B) and self-associated (Fig. 1C) in the two-hybrid system as readily as full-length ParG. In chemical cross-linking, ParG principally forms dimeric species (6.Barillà D. Hayes F. Mol. Microbiol. 2003; 49: 487-499Crossref PubMed Scopus (36) Google Scholar). Δ9ParG, Δ19ParG, and Δ30ParG displayed similar patterns over a range of incubation temperatures (Fig. 1D). Furthermore, sedimentation velocity measurements revealed the predominant presence of a single dimeric species for each of the three proteins (Fig. 1E), similar to results obtained from analytical ultracentrifugation studies of full-length ParG (6.Barillà D. Hayes F. Mol. Microbiol. 2003; 49: 487-499Crossref PubMed Scopus (36) Google Scholar). Taken collectively, the two-hybrid, cross-linking, and sedimentation velocity data conclusively demonstrate that the N-terminal mobile tail of ParG plays no detectable role in dimerization. ParG Is the Autorepressor of the parFG Genes and Interacts with an Operator Site That Overlaps the Putative parFG Promoter—A 259-bp fragment upstream of the parF translational start codon is necessary for plasmid partitioning (5.Hayes F. Mol. Microbiol. 2000; 37: 528-541Crossref PubMed Scopus (75) Google Scholar) and is bound by ParG (6.Barillà D. Hayes F. Mol. Microbiol. 2003; 49: 487-499Crossref PubMed Scopus (36) Google Scholar). The 80-bp region immediately upstream of the parF initiation codon includes an inverted repeat (IR) that consists of imperfect 16-bp half-sites separated by a 4-bp spacer (see Fig. 3E). In silico analysis pinpointed the probable parFG promoter to the 5′-end of the 80-bp region. The candidate -10 and -35 elements for this promoter each contain 4/6 matches to the consensus promoter for σ70 of E. coli, with optimal separation of the boxes. The -10 element overlaps the left half-site of the 36-bp IR (Fig. 2, inset). This organization suggests that the IR is an operator site implicated in regulation of parFG expression. To assess the role of the upstream region in parFG expression, a xylE transcriptional fusion was constructed by cloning a 117-bp fragment covering the promoter, IR, and 5′-end of parF in a promoterless xylE cassette in plasmid pDM3.0 (Fig. 2). This fusion produced 301 ± 15 catechol 2,3-dioxygenase (CDO) units, compared with the low activity of the promoterless plasmid (∼0.5 CDO units), demonstrating that the parFG promoter is indeed located in the short region immediately upstream of the parF translational start. The ParG protein was provided from a pET22b(+) expression vector, and its effect in CDO assays was examined to assess whether it was a transcriptional repressor of the parFG promoter. Expression of the parF-xylE fusion was reduced 20-fold from 360 ± 25 CDO units in the presence of pET22b(+) to 18 ± 7 units in the presence of ParG (Fig. 2). To test the role of the IR in this repression, the rightward half of the palindrome was scrambled to a similar sequence to that present in the in vitro substrate HS-48 (Fig. 3). CDO production from a construct containing the mutated IR was similar to that from the unmutated plasmid, demonstrating that promoter activity is unaltered by the mutations. However, ParG repressed expression from this promoter only 3-fold. These results demonstrate that ParG is a transcriptional repressor of parFG expression and that the IR is an operator site required for this repression. The interaction between ParG and the IR region was explored further by EMSA with a biotinylated 48-bp oligonucleotide covering the IR (FS-48) (Fig. 3E). ParG formed two well defined complexes (I and II) with this substrate that eventually resolved into the more slowly migrating species (II) at high protein concentration (Fig. 3A). ParG did not retard a half-site of the IR located on either a 23-bp oligonucleotide (HS-23) (Fig. 3A) or on a 28-bp substrate bearing 6-bp extensions on both ends of the half-site (data not shown). In contrast, a substrate (HS-48) in which the right half-site of FS-48 was randomized formed a single complex with a migration similar to complex I (Fig. 3A). Thus, although ParG does not associate with an isolated half-site (HS-23), addition of the unrelated sequence in HS-48 stabilizes the interaction, perhaps by the association of ParG dimers bound to opposite half-sites. In this case, ParG bound to the intact half-site might be positioning additional ParG protomers non-specifically on the randomized sequences. The stoichiometry of ParG bound to FS-48 was assessed by a variation of the Ferguson method (33.Ferguson K.A. Metabolism. 1964; 13: 985-1002Abstract Full Text PDF PubMed Scopus (781) Google Scholar) adapted for protein-DNA complexes (29.Orchard K. May G.E. Nucleic Acids Res. 1993; 21: 3335-3336Crossref PubMed Scopus (68) Google Scholar). Gel filtration, analytical ultracentrifugation, and dynamic light scattering showed that the mobile tail of ParG greatly increases its hydrodynamic radius. 2F. Hayes, D. Barillà, and E. Carmelo, unpublished data. To reduce this effect, Δ9ParG was selected for Ferguson analysis as it retains the activities of full-length ParG in EMSA (see following section) but lacks approximately one-third of the tail. A set of protein standards and complexes I and II formed between Δ9ParG and FS-48 or HS-48 were analyzed in gels containing 7-12% acrylamide. The relative migrations of the main bands were plotted against the polyacrylamide concentrations of the gels (Fig. 4A). The retardation coefficients were represented against the molecular masses, and the sizes of the nucleoprotein complexes were estimated from the resulting plot (Fig. 4B). This yielded molecular masses of 72 and 101 kDa for complexes I and II, respectively. These figures are in close agreement with the 64.5 kDa calculated for complexes formed by FS-48 and two Δ9ParG dimers (29.5 kDa DNA + 2 × 17.5 kDa protein) and 99.5 kDa for FS-48 and four Δ9ParG dimers (29.5 kDa DNA + 4 × 17.5 kDa protein). The ParG:FS-48 binding stoichiometry was also analyzed by CD measurements, showing an increase of ellipticity at 273 nm caused by the titration of ParG with FS-48 compared with the free DNA. Fig. 4C shows the quantitative monitoring of that change, plotted as a percentage of the maximum change. The plot indicates that the ellipticity variations attain a plateau at a 4:1 molar ratio of ParG dimers:FS-48, establishing a complex similar to the previously described complex II. The N-terminal Tail Is Required for Repression and Modifies the Interaction of ParG with Its Operator Site—The Δ9ParG, Δ19ParG, and Δ30ParG derivatives were examined for repression of the parF-xylE reporter to assess whether the unstructured tail influences transcriptional repression by ParG (Fig. 2). Expression of the fusion was strongly repressed by Δ9ParG (13 ± 6.6 CDO units), producing a repression ratio very similar to that determined for full-length ParG (Fig. 2). In contrast, repression of the parFG promoter was dramatically reduced for Δ19ParG and Δ30ParG, which had repression ratios of only 2.3- and 1.5-fold, respectively. Thus the unstructured tail of ParG plays a crucial role in transcriptional repression of the parFG promoter, and increasingly longer deletions of the tail are accompanied by progressively weaker repression. The involvement of the N-terminal tail in the interaction of ParG with the operator was investigated further by EMSA (Fig. 3, B-D). Δ9ParG and Δ19ParG displayed patterns of binding to FS-48 that were very similar to those observed for ParG: complex I was most evident at low protein concentrations, with the appearance of complex II at higher protein:DNA ratios. For Δ9ParG and Δ19ParG, a large proportion of the substrate was bound into complex II at the highest protein concentration (Fig. 3, B and C). In contrast, interaction of Δ30ParG with FS-48 produced a smeared band with a different migration from either complexes I or II, suggesting that the association of Δ30ParG with the substrate was particularly perturbed (Fig. 3D). The singular behavior of Δ30ParG was emphasized by experiments with HS-48, with which the protein failed to form a defined complex, only smearing it from the unbound position. In contrast, Δ9ParG and Δ19ParG both associated with HS-48 with similar patterns as full-length ParG, producing complex I in each case. Like ParG, none of the deletion proteins interacted detectably with HS-23. Surface plasmon resonance experiments were performed to quantitate the ParG-operator interactions. Using a Biacore 3000 instrument, a CM Sensor Chip (Biacore) was extravidin coated and derivatized with FS-48 in Flow-Cell (FC) 1, HS-48 in FC2, and 48 bp of unrelated DNA as a reference in FC3. Proteins were passed over the chip surfaces at 0.05-5 μm (monomer equivalents) for 60 s (association) and allowed to wash off subsequently for 450 s (dissociation). ParG and the N-terminal deletion variants bound strongly to FS-48 at a concentration of 0.5 μm (Fig. 5A), and a similar effect was observed at every protein concentration tested (data not shown). Nevertheless, the interaction with this target was different between the proteins. Δ9ParG reproducibly gave the strongest response with FS-48, as opposed to Δ19ParG whose response was much weaker. Sensorgrams of ParG, Δ9ParG, and Δ19ParG had noticeably curved association and dissociation phases, strikingly different from those of Δ30ParG, which presented a much flatter pattern particularly in the dissociation phase when the slope was almost horizontal. The observation in EMSA analysis that ParG established two different complexes with FS-48 suggested that ParG and the N-terminal deletion derivatives follow a complex model in operator site binding. None of the kinetic models proposed by BIAevaluation 3.1 software gave a satisfactory close curve fitting for thi" @default.
• W2017161880 created "2016-06-24" @default.
• W2017161880 creator A5034687372 @default.
• W2017161880 creator A5040228041 @default.
• W2017161880 creator A5052378786 @default.
• W2017161880 creator A5056299775 @default.
• W2017161880 creator A5057928429 @default.
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• W2017161880 date "2005-08-01" @default.
• W2017161880 modified "2023-10-03" @default.
• W2017161880 title "The Unstructured N-terminal Tail of ParG Modulates Assembly of a Quaternary Nucleoprotein Complex in Transcription Repression" @default.
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