article: Although quorum-sensing (QS) systems are important regulators of virulence gene expression in the opportunistic human pathogen Pseudomonas aeruginosa, their detailed regulatory mechanisms have not been fully characterized. Here, we show that deletion of PA2588 resulted in increased production of pyocyanin and biofilm, as well as enhanced pathogenicity in a mouse model. To gain insights into the function of PA2588, we performed a ChIP-seq assay and identified 28 targets of PA2588, including the intergenic region between PA2588 and pqsH, which encodes the key synthase of Pseudomonas quinolone signal (PQS). Though the C-terminal domain was similar to DNA-binding regions of other AraC family members, structural studies revealed that PA2588 has a novel fold at the N-terminal region (NTR), and its C-terminal HTH (helix-turn-helix) domain is also unique in DNA recognition. We also demonstrated that the adaptor protein ClpS, an essential regulator of ATP-dependent protease ClpAP, directly interacted with PA2588 before delivering CdpR to ClpAP for degradation. We named PA2588 as CdpR (ClpAP-degradation and pathogenicity Regulator). Moreover, deletion of clpP or clpS/clpA promotes bacterial survival in a mouse model of acute pneumonia infection. Taken together, this study uncovered that CdpR is an important QS regulator, which can interact with the ClpAS-P system to regulate the expression of virulence factors and pathogenicity.
  P. aeruginosa is one of the most common nosocomial pathogens associated with fatal lung disease in cystic fibrosis patients [1]. This bacterium synthesizes a group of virulence factors consisting of pyocyanin, rhamnolipids, proteases, and biofilms that are regulated by a cell density-dependent quorum-sensing (QS) system [2,3] and second messenger c-di-GMP [4]. QS systems enable bacteria to communicate and regulate a large number of genes. P. aeruginosa has one of the most sophisticated QS systems of all bacterial species, which includes two systems (las and rhl) based on N-acyl-homoserine lactones (AHLs) and one based on 2-alkyl-4-quinolone (AQ) [2]. The LasI synthase produces the signal of the las system, N-3-oxo-dodecanoyl homoserine lactone (3-oxo-C12-HSL), which is recognized by the transcriptional regulator LasR [5]. The RhlI synthase catalyzes the synthesis of N-butanoyl homoserine lactone (C4-HSL), which is sensed by the transcriptional regulator RhlR [6]. These two systems control 10% of the P. aeruginosa genome [7,8]. Besides the AHLs systems, P. aeruginosa also produces the AQ signal molecule [9]. Pseudomonas quinolone signal (PQS) is synthesized from the precursor molecule 2-heptyo-4 (1H) -quinolone (HHQ) and finally converted into PQS by PqsH [10]. Moreover, these three systems and a group of transcriptional regulators (such as VqsR, QscR, VqsM, Vfr, and RpoN) form a complex regulatory network [11]. Intracellular protein degradation plays essential roles in many physiological processes such as cell growth and division. In Escherichia coli, proteolysis is mainly mediated by ATP-dependent proteolytic machineries (termed AAA+ proteases) [12], which consist of a proteolytic barrel (e. g. , heptameric ClpP) and a hexameric ATPase (e. g. , ClpA, ClpX). The ATP-binding proteins ClpX and ClpA recognize a specific peptide sequence of unfolded proteins [13,14], and translocate the tags into the peptidase barrel of ClpP, thus degrading the unfolded protein into small peptides [15]. In addition, adaptor proteins (i. e. , ClpS) are responsible for binding and transporting their target proteins to the protease complexes for degradation [16]. In P. aeruginosa, the Lon and ClpP proteases share 84% and 86% identity with their E. coli counterparts, which suggest that these proteases also play important roles in the degradation of unstable or misfolded proteins in P. aeruginosa. Moreover, the ATP-dependent ClpP protease has been shown to control virulence, antibiotic resistance, and a group of metabolic pathways [17]. However, the substrates and molecular mechanism of ClpP in regulating bacterial pathogenesis are still unclear. In this study, we showed that the new QS regulator CdpR (we named PA2588 in this study) directly activated PqsH, the PQS synthase. CdpR regulated the expression of virulence factors (i. e. , pyocyanin and motility) and bacterial pathogenicity of P. aeruginosa. Unlike other AraC-family proteins, the C-terminal HTH (helix-turn-helix) motif of CdpR has a unique conformation, which may play important roles in DNA recognition. Structural comparison also revealed that the N-terminal region (NTR) of CdpR has a novel fold whose function remains elusive. We also found that CdpR interacts with adaptor protein ClpS and is degraded by the ClpAP protease in vivo. Moreover, the deletion of clpP or clpS/clpA-promoted bacterial survival in a mouse model of acute pneumonia. To the best of our knowledge, this is the first report showing the degradation of a regulator of the alginate system by ClpAS-P proteases in P. aeruginosa. Overall, these findings establish a link between proteases ClpAS-P and virulence, which provides us with new insights into investigate the roles of proteases in the future.
  Previously, we showed that the QS regulator VqsM directly binds to the promoter region of the PA2588 (cdpR) gene [18]. Its N-terminus (amino acids 59–198) is homologous to the arabinose-binding domain of the AraC transcription regulator (N-term; pfam12625), suggesting that it may be an AraC-family transcriptional regulator. Given that lasI and cdpR are directly controlled by VqsM, we hypothesized that CdpR also regulates P. aeruginosa QS-associated virulence factors, such as pyocyanin production. To this end, we made a cdpR null mutant in the PAO1 strain by following procedures from our previous study [18]. The resulting ΔcdpR mutant exhibited higher pyocyanin production than wild-type PAO1 after 24 h of growth in Luria-Bertani (LB) medium. Overexpression of cdpR in a pAK1900 plasmid (T7 promoter) decreased the pyocyanin production in the cdpR mutant (Fig 1A). Two homologous operons are involved in the synthesis of phenazine compounds (i. e, pyocyanin) in P. aeruginosa, phzA1B1C1D1G1 (phzA1) and phzA2B2C2D2G2 (phzA2) [19]. The increased pyocyanin production in the ΔcdpR strain led us to test if cdpR regulates the promoter activity of phzA1. To this end, a phzA1-lux reporter was introduced into the wild-type, the ΔcdpR mutant, and its complemented strain. The expression of phzA1-lux in the ΔcdpR mutant was 3-fold higher than in the wild-type PAO1 or its complemented strain (Fig 1B). These results suggest that CdpR is a negative regulator of pyocyanin synthesis. The enhanced pyocyanin production in the ΔcdpR mutant led us to test if cdpR regulates other QS-related pathways, such as biofilm formation and bacterial motility. We performed microtiter dish and borosilicate tube binding assays to evaluate the biofilm formation of these strains. The ΔcdpR mutant exhibited better adherence than either the wild-type or the complemented strain (Fig 1C). In addition, deletion of cdpR resulted in reduced swarming motility (Fig 1D). Collectively, these results demonstrated that CdpR is a new and important regulator of P. aeruginosa virulence factors. In order to dissect the regulatory mechanism of CdpR, we performed a ChIP-seq assay to identify its direct targets in the Pseudomonas genome. Like untagged CdpR, CdpR-VSV (overexpressed in the plasmid pAK1900) was also able to decrease the expression of phzA1 (Fig 1B). Using MACS software for identifying conserved DNA motif bound by CdpR [20], 28 enriched loci (p-value = e−5) were identified to carry CdpR-binding peaks (S1 Table), which were enriched > 1. 5 folds compared to the control samples using the wild-type without VSV tags. Eighty-nine percent of these 28 loci are located within coding regions (Fig 2A). For example, CdpR binds to the coding region of PA0440 gene in the immunoprecipitated sample, while the enrichment in PA0440 was not present in the input sample (the wild-type PAO1 strain with empty pAK1900 vector) (Fig 2B). We further categorized the biological processes of CdpR targets based on gene ontology [21], including metabolism (28%), secretion (7%), regulation (4%), nucleic acid metabolism (3%), transportation (11%), virulence (11%), and unknown functions (36%) (Fig 2C). The ChIP-seq data also allowed us to define a 15-bp consensus-binding motif of CdpR (Fig 2D) using multiple EM for motif elicitation (MEME) analysis. We next sought to perform electrophoretic mobility shift assays (EMSA) to verify the CdpR targets in vitro. We first expressed and purified the CdpR protein (S1A Fig). CdpR bound to two promoter regions of cerN and sphR, but not to the lasR promoter as a negative control (S1B–S1F Fig). We further tested 18 binding sites, and 13 showed specific binding (S2A Fig). Overall, these EMSA results are consistent with the ChIP-seq data and confirm the binding motif of CdpR in the P. aeruginosa genome. The highest enriched region by CdpR was the shared promoter region between pqsH and cdpR (Fig 3A). EMSA showed that CdpR bound to the pqsH-cdpR intergenic region (pqsH-p) but not to the negative control (pqsR-p) (Fig 3B). Using a dye-based DNaseI footprinting assay, we further found a specific CdpR-bound sequence containing a 15-bp motif (5' -CTGCGCCTGGATGAT-3' ) (Fig 3C). The protected region extended from nucleotides -377 to -361 ahead of the pqsH translational start codon, which is consistent with the identified sequences by MEME analysis (Fig 2D). We have blasted the intergenic sequences between cdpR and pqsH in other Pseudomonas species. This region is largely conserved in P. aeruginosa, but not in other Pseudomonas species, suggesting that the shared promoter between cdpR and pqsH is universal in P. aeruginosa. We also noted that the motif (CTGCGCCTGGATGAT) is also present in the promoter regions of cerN (5' -CTTCGCCTGGTTGCC-3' , -87 to -72 from the cerN translational start codon) and sphR (5' -CTGCGCCTGGCCGCC-3' , -266 to -252 from the sphR translational start codon) (S2B Fig). In order to determine if the motif (CTGCGCCTGGATGAT) was important for binding to CdpR, we performed EMSAs again using a pqsH-p1 DNA fragment (−365 to +88 relative to the start codon of pqsH, without the binding motif) and a pqsH-M-p DNA fragment (the motif CTGCGCCTGGATGAT was mutated to TGACTTCTGGATGAT). Neither the pqsH-p1 nor pqsH-M-p fragments were able to bind to CdpR (S3A Fig). As expected, CdpR could not bind to the sphR-p1 or cerN-p1 fragment (without the binding motif) (S1D and S1E Fig). We next tested if CdpR controls the expression of pqsH or itself in vivo. As shown in Fig 3D, the promoter activity of pqsH (pqsH-lux, −500 to +88 relative to the start codon) in the ΔcdpR mutant strain was drastically weaker than that in the parental strain. Overexpression of cdpR gene in the ΔcdpR strain restored the expression of pqsH to the wild-type level. This result was further echoed by western blotting of the strains carrying an integrated mini-CTX-pqsH-flag plasmid (Fig 3E). We further measured QS molecules including PQS and C4-HSL, which showed that PQS production was lower in the cdpR mutant than in the wild-type PAO1 strain (Fig 3F); but no difference in C4-HSL production was observed between them (S3B Fig). Moreover, mutation in the consensus motif abolished the promoter activity of pqsH-M-lux in the wild-type strain (S3C Fig), suggesting that the binding sequence is required for full activity of the pqsH promoter. In addition, the expression of cdpR in the ΔcdpR mutant was 4-fold higher than that in the wild-type strain (S3D Fig), suggesting that the cdpR is negatively autoregulated. Taken together, these results indicate that CdpR regulates both pqsH and itself. As aforementioned, the deletion of cdpR increased pyocyanin production and compromised the swarming motility and improved biofilm formation (Fig 1). We next sought to test whether the deletion of cdpR changes P. aeruginosa virulence in a mouse model of acute pneumonia. Therefore, 6-wk-old C57BL/6 mice were intranasally inoculated with 1×107 wild-type, ΔcdpR mutant, or complemented strain (single copy integration of the cdpR gene). Kaplan-Meier survival analysis showed that loss of cdpR significantly decreased mouse survival compared to the wild-type PAO1. The ΔcdpR strain caused 70% and 100% of infected mice to die by 12 h and 48 h, respectively. In contrast, 60% of mice infected with wild-type PAO1 or the complemented strain were alive at 48 h (Fig 4A). We also observed more lung injury and inflammation in the ΔcdpR strain-infected mice than the wild-type PAO1 or complemented strain-infected mice by histological analysis with H&E staining (Fig 4B). The colony-forming units (CFUs) in alveolar macrophages (AM) infected with ΔcdpR were higher than those with the wild-type or the complemented strain (Fig 4C). In addition, the ΔcdpR-infected mice produced more superoxide and more polymorphonuclear neutrophil (PMN) cells than wild-type-infected mice (Fig 4D and 4E). In summary, these results indicate that CdpR is a negative regulator of P. aeruginosa pathogenicity in a mouse model. To gain more insights into the structural basis underlying the functions of PaCdpR, we carried out structural studies. The structure belongs to P41212 space group, and it contains one PaCdpR protein molecule in the asymmetric unit. As depicted in Fig 5A and 5B, the overall structure of PaCdpR can be divided into three regions, i. e. , NTR (aa 5–203), connector region (CR, aa 204–228), and C-terminal HTH domain (aa 229–329). The NTR (Fig 5C) is of PaCdpR that is of α/β fold in nature; it is composed of ten α-helices (α1 to α10) and five β-strands (β1 to β5). The β strands form a single-layer β-sheet. The last strand (β5) sits in the center, whereas β2 and β1 locate on one side, and β3 and β4 locate on the other side. Except β4, which is parallel to β3, all the other strands are antiparallel to each other. The α-helices can be divided in two groups: group A (α1 to α5) and group B (α6 to α10). Group B α-helices were clamped between the A group helices and the β-sheet. Four helices, α9, α10, α6, and α7 were packed against the β-sheet at the edge in an anti-clockwise direction, while α8 (the longest helices) crossed the β-sheet in the middle and formed extensive hydrophobic interactions with the three central strands (β2, β5, and β3). E. coli AraC (EcAraC) is composed of two domains, i. e. , N-terminal domain (NTD) and C-terminal HTH domain, and is the most well studied member of AraC family. Upon the binding of arabinose, EcAraC NTD can change its dimerization mode, which will in turn alter the HTH–DNA interaction and affect its gene regulation activity [22]. Though both CdpR NTR and EcAraC NTD are of α/β fold in nature, their overall folds are different from each other. EcAraC NTD has a jelly-roll fold β-barrel, which holds the arabinose in between the double-layer beta-sheet (S4A Fig). Moreover, no sequence similarity was observed between CdpR NTR and EcAraC NTD. These differences suggested that CdpR NTR and EcAraC NTD may have different functions. To this end, we carried out an in vitro DNA binding assay. As shown in S4B Fig, the CdpR–DNA interaction was almost identical in the presence or absence of arabinose, suggesting that arabinose is not a cofactor of CdpR. Structural comparison using DALI server [23] did not reveal significant similarity between CdpR NTR and other known AraC family structures. Altogether, these data indicate that CdpR NTR has a novel fold among AraC family, and its functions need to be further investigated. HTH domains are conserved in AraC family. Besides CdpR, the HTH domain structures of EcAraC (PDB ID: 2K9S) [24], EcMarA (multiple antibiotic resistant regulon A, PDB ID: 1BL0) [25], and Streptomyces griseus AdpA (PDB ID: 3W6V) [26] have also been reported. Although the overall folds are similar, structural comparison revealed that the detailed conformation of CdpR HTH is different from other AraC proteins (Fig 6A). CdpR HTH contains seven α helices (α13–α19), which can be further divided into two subdomains (S5A Fig), i. e. , HTH1 (aa 229–271) and HTH2 (aa 291–329). The largest difference occurs at the HTH1 motif, especially the helix α13. Consistently, the sequence similarity at this region is also very low (S5B Fig). The interaction modes between HTH1 and HTH2 motifs are also different within these structures. In EcMarA and sgAdpA structures, loop A and loop B interact with each other via backbone van der Waals contacts; however, as depicted in Fig 6B, such interaction was not observed in CdpR, indicated by the shortest distance (6. 7 Å) between the loop backbones. Instead, CdpR HTH1 and HTH2 mainly interact with each other through the backbone of loop B (aa 303–304) and the side chain of Tyr245 from loop A. This interaction was further enhanced by Arg281 and Arg284 of helice α16, which connects HTH1 and HTH2 and Tyr305 of loop B, via H-bonding and stacking. Both Tyr245 and Arg284 of CdpR are not conserved in other AraC-family proteins. HTH domains are responsible for target DNA recognition, and the complex structures have been solved for EcMarA and sgAdpA. To better understand the mechanism of CdpR-DNA recognition, we also carried out molecular modeling study. CdpR HTH may recognize the DNA base pairs using two positive charged residues (Arg260 and Arg264), corresponding to Arg262 and Arg266 of sgAdpA. In sgAdpA HTH–DNA structure, DNA backbones form several H-bonds with the side chains of positive charged residues (Fig 6C and 6D). Arg312 and Arg317 of CdpR HTH are corresponding to Arg315 and Arg320 of sgAdpA. The other four residues (Arg261, Arg269, Arg309, and His317) of sgAdpA are not conserved, but Arg263 of CdpR may replace Arg261 of sgAdpA during DNA recognition, supported by the similar orientations of their guanine groups. Structural analysis did not reveal any residues of CdpR, which could form H-bonds with DNA similar to Arg269, Arg309, and His317 of sgAdpA; lack of extensive interactions with DNA backbones may be the main cause of the weak CdpR-DNA binding affinity (Fig 6D). In order to experimentally confirm the proposed structural model for CdpR-DNA complex, conserved residues predicted to contact DNA (Arg263, Arg274, Arg312, and Arg317) were mutated to Ala. We next performed EMSAs for comparing the ability of purified wild-type CdpR to bind a DNA probe encompassing the pqsH-cdpR intergenic region with those of its mutated derivatives. As shown in Fig 6E, CdpRwt was able to shift about 50% of the probe at a concentration of 0. 5 μM, while CdpRR263A, CdpRR274A, and CdpRR312A/R317A were unable to shift the same DNA probe even at a concentration of 2 μM. To future confirm these residues are important for CdpR activity, we tested the expression of pqsH-lux in the cdpR mutant containing the expressing plasmid p-cdpRR263A, p-cdpRR274A, or p-cdpRR312A/317A. As shown in Fig 6F, mutation of these residues did not restore the pqsH activity of cdpR mutant to wild-type levels. These results clearly suggest that these residues are important for CdpR activity and confirm the CdpR-DNA model is accurate. In order to identify regulators of cdpR, its promoter was used as a reporter (CTX-cdpR-lux) and subjected to transposon mutagenesis. After three rounds of screening of 8,000 mutants, 4 positive and 10 negative mutants were selected. Genes inserted by transposon were determined via arbitrary primed PCR and subsequent DNA sequencing (S2 Table). Interestingly, mutation of lasI compromised the expression of cdpR compared to wild-type PAO1, suggesting that LasI is a positive regulator of cdpR. The expression of cdpR was increased in the M6 (cdpR: : Tn) strain compared to the wild-type strain, which is consistent with the observation in the cdpR deletion mutant (S6A Fig). As expected, pyocyanin production was also elevated in the M6 strain (S6B Fig). We noted that vqsM was missing in our screen, which might result from limited number (8,000) of mutants. Of these genes, we are most interested in the ATP-dependent protease ClpP, which associates with ClpS (ATP-dependent Clp protease adaptor protein) and ClpA (ATP-binding protease component). These mutants showed very strong phenotypes, revealing that they are involved in CdpR-controlled pathway. Moreover, previous study has shown that ClpP is related to bacterial virulence [27]. clpS and clpA share the same operon [28]. To further verify the effects of the clpS/clpP transposon insertion, we made a clpS-clpA double mutant (ΔclpSΔclpA) and a clpP single mutant (ΔclpP). As expected, the activity of cdpR-lux in these two deletion strains was higher than that in the wild-type strain (S7A Fig). We next wanted to verify whether the protein level of CdpR was affected by these proteases. To this end, we constructed the CTX-cdpR-flag fusion and the activity of CdpR-flag was shown the same as the wild-type CdpR (S7B Fig). Next, we tested the expression of CdpR-Flag in the wild-type PAO1, the ΔclpSΔclpA mutant, the ΔclpP mutant, and their complemented strains (ΔclpSΔclpA/p-clpSclpA, and ΔclpP/p-clpP) using western-blot assay. Higher levels of CdpR-Flag were detected in the ΔclpSΔclpA or ΔclpP mutant than in the wild-type strain. The complemented strains restored CdpR-Flag to the wild-type levels (S7C Fig). As expected, the expression of pqsH in ΔclpSΔclpA or ΔclpP mutants was also higher than the wild-type PAO1 (S8 Fig). Taken together, these results indicate that ClpS/ClpA and ClpP negatively regulate the expression of CdpR. Given that ClpS is a bacterial adaptor that recognizes and delivers N-degron (N-terminal amino acids) substrates to the ClpAP or ClpCP AAA+ proteases [29,30], we hypothesized that ClpS interacts with CdpR by a bacterial two-hybrid system. Like strain 1 (the positive control), strain 4 (bearing pBT-CdpR and pTRG-ClpS) also grew on dual-selective medium (Fig 7A), indicating a direct interaction between CdpR and ClpS. The direct interaction between ClpS and CdpR led us to examine if the CdpR protein can be degraded by the ATP-dependent ClpAP protease in vivo. We transformed mini-CTX-cdpR-flag into the wild-type PAO1, the ΔclpP, the ΔclpSΔclpA, and their complemented strains. Western-blotting analyses of cell lysates were performed over the course of 90 min. Degradation of CdpR-Flag was observed in the wild-type PAO1 and the complemented strains with a half-life of less than 30 min (Fig 7B). However, CdpR levels remained steady in the ΔclpP and the ΔclpSΔclpA strains (the second and fourth panels in Fig 7B), demonstrating that the ClpSA-ClpP protease system is responsible for CdpR degradation. A group of proteases has been shown to modulate P. aeruginosa virulence. For example, the deletion of Lon protease exhibits a defect in cell division and virulence-related properties, such as swarming, twitching, and biofilm formation [31]. It was also demonstrated that the lon mutant was less virulent in a mouse acute lung infection model as well as in a rat model of chronic infection [32]. The protease ClpP is also involved in antibiotic resistance and virulence-related phenotypes, such as motility and biofilm formation [17]. In the present study, ClpS/ClpA and ClpP controlled the expression of CdpR, which negatively regulated pyocyanin production and bacterial virulence (Fig 1). We showed that the ΔclpSΔclpA and ΔclpP mutants produced more pigments than the wild-type PAO1 (Fig 8A and 8B). The deletion of cdpR in the ΔclpSΔclpA or ΔclpP mutants restored pigment production and phzA1-lux activity to the wild-type level (Fig 8C). We also observed that the swarming motility of the ΔclpAΔclpS and the ΔclpP strains could be partially reversed by CdpR (S9 Fig). Taken together, these results indicate that ClpSA-ClpP plays important roles in regulating pigment and motility. CdpR interacts with these intracellular proteases with important functions. These altered virulence-associated phenotypes in the ΔclpP and ΔclpSΔclpA mutants led us to evaluate the pathogenicity of these mutants in a mouse model. We found that deletion of clpS/clpA or clpP improved mouse survival compared to the wild-type PAO1. The ΔclpSΔclpA and ΔclpP strains exhibited 100% and 85% of mice survival after 96 h, whereas the wild-type bacteria caused 70% death at 80 h (Fig 8D). These results demonstrated that ClpS/ClpA and ClpP proteases play important roles in P. aeruginosa pathogenicity in a mouse model.
  Here, we present the first characterization of CdpR as an important regulator of QS system that directly binds to the promoter region of pqsH. We also solved the crystal structure of CdpR and revealed that CdpR NTD has a unique fold that is different from other AraC-family proteins. In addition, we found that CdpR interacts with ClpS before degradation by ClpAP proteases, thus regulates the P. aeruginosa virulence. Although the QS systems in P. aeruginosa have been widely characterized in the last several decades, the complete signaling network and interactions with other systems still remain elusive. Previously, we showed that the QS regulator VqsM interacts with the promoter of cdpR [18]. Like VqsM, CdpR belongs to the AraC-family transcriptional regulators, which control a variety of cellular processes in bacteria, including carbon metabolism, stress responses, and virulence [21]. We performed a ChIP-seq assay that revealed 28 CdpR targets in the P. aeruginosa genome. More importantly, CdpR directly regulates the PQS system by binding to the promoter of pqsH (Fig 3 and S3 Fig). The ΔcdpR mutant exhibited higher pyocyanin and biofilm production as well as reduced bacterial motility (Fig 1), which are controlled by the PQS system [9,33]. Moreover, CdpR plays important roles in regulating bacterial pathogenicity in a mouse experiment. Mice inoculated with the ΔcdpR mutant showed more bacterial loads than mice infected with the wild-type strain (Fig 4A). The increased pathogenicity corresponds to the higher production of pyocyanin in the ΔcdpR mutant (Fig 1A). Like the other AraC-family proteins, CdpR has an NTD whose function is deemed as binding to small molecules and a HTH region, which binds to a special substrate. From our ChIP-seq results, CdpR can target many genes, but its DNA binding affinity is relatively lower than other identified AraC proteins. This is due to the lack of positively charged residues in HTH domain that can interact with the DNA backbones (Fig 6 and S5 Fig). Unlike other AraC-family proteins that can bind arabinose using NTD domain and alter the gene regulation activity, CdpR has a novel NTR region, and its interaction with target DNAs is not affected by arabinose (S4 Fig). CdpR NTR may also play a role in the gene regulation via the interaction with other proteins or small molecules. Besides, analysis of the CdpR structure revealed that the conformation of CdpR HTH is different from other AraC proteins (Fig 6A). The proposed structural model for CdpR-DNA complex demonstrated that Arg263, Arg274, Arg312, and Arg317 are important for DNA-binding ability of CdpR (Fig 6D), which has been verified by in vitro EMSA and in vivo complemented assays where these residues were mutated (Fig 6E and 6F). Overall, our results provide insights into the function of AraC-family protein in different bacteria. To further understand the regulatory mechanism upstream of CdpR, we performed a transposon mutagenesis assay that revealed several mutants showing altered expression of cdpR (S2 Table). The expression of cdpR was highly compromised in a mutant containing a transposon insertion in clpAP, which encodes ATP-dependent proteases [34]. Proteases ClpAP and Lon are involved in bacterial virulence expression, antibiotic resistance, and metabolism [27]. Lon protease negatively modulates QS by degrading autoinducer synthase LasI [35]. Our present study showed that CdpR interacted with the adaptor ClpS, and CdpR protein was degraded by the ClpAP protease in vivo (Fig 7). To the best of our knowledge, this is the first evidence showing the substrates of ClpS-ClpAP in P. aeruginosa. Further work needs to focus on the precise binding sites between ClpS and CdpR. In conclusion, we identified an AraC-family regulator, CdpR, which negatively modulates bacterial virulence (Fig 9). CdpR is a substrate of ATP-dependent ClpAP protease. CdpR also regulates the expression of pqsH by directly binding to its promoter regions. CdpR is positively modulated by LasI (S2 Table) and VqsM [18]. CdpR may work with other negative QS regulators, such as QscR and RsaL, to negatively regulate QS. In addition, ClpS interacts with CdpR and then associates with ClpAP to degrade CdpR. CdpR might indirectly regulate the expression of virulence factors, thus negatively controls P. aeruginosa pathogenicity. This work provides insight into roles of proteases in QS and bacterial virulence, which provides more cues to design effective antimicrobial drugs for the prevention of P. aeruginosa infections in the future.
  All the experiments involving animals were approved by the University of North Dakota Institutional Animal Care and Use Committee and performed in accordance with the animal care and institutional guidelines (IACUC approval #: 1204–5) (Assurance Number: A3917-01). Animal experimental procedures, including treatment, care, and end point choice, followed Animal Research: Reporting in Vivo Experiment guidelines. When moribund animals manifested distress and/or mortal signs, such as lethargy, weight loss (15%–25%), lack of eating or drinking, tissue rupture, or edema, we considered these as endpoints and humanely euthanized them immediately by CO2. Recovery of the animals was achieved without complications, but we monitored hourly until they were awake and active after procedures and then every 6 h. Animals were monitored daily throughout the experimental process. Once sedation was diminished, the mice acted normally and appeared playful. There generally was no apparent pain from the intranasal instilling procedure. If any pain sign occurred, a 1% solution of lidocaine was applied to lessen pain. The bacterial strains and plasmids used in this study are listed in S3 Table. P. aeruginosa PAO1 and derivatives were grown at 37°C on LB agar dishes or in broth with shaking at 220 rpm. Antibiotics were used at the following concentrations: for E. coli, gentamicin (Gm) at 15 μg/ml, ampicillin at 100 μg/ml, and tetracycline 10 μg/ml; for P. aeruginosa, gentamicin (Gm) at 50 μg/ml in LB or 150 μg/ml in PIA (Pseudomonas Isolate Agar); Tetracycline at 150 μg/ml in LB or 300 μg/ml in PIA and Carbenicillin at 500 μg/ml in LB. The procedures of chromatin immunoprecipitation (ChIP) were modified from previously described studies [18,36]. Wild-type PAO1 strain carrying pAK1900 or pAK1900-CdpR-VSV was cultured in LB medium until OD = 0. 6 and then crosslinked with 1% formaldehyde (final concentration) for 10 min at 37°C with shaking. 125 mM glycine (final concentration) was added into the culture to stop the crosslinking. Bacteria were centrifuged and washed three times with a Tris buffer (150 mM NaCl, 20 mM Tris-HCl, pH 7. 5). The pellets were resuspended in 500 μl IP buffer (150 mM NaCl, 50 mM HEPES-KOH pH 7. 5,1 mM EDTA, 0. 1% SDS, 1% Triton X-100,0. 1% sodium deoxycholate and mini-protease inhibitor cocktail (Roche), and the DNA was sonicated to 100–300 bp. The chromatins were centrifuged (12,000 rpm, 4°C), and the supernatant was saved. All samples were treated by protein A beads (General Electric) before adding 50 μl agarose-conjugated anti-VSV antibodies. Washing, crosslink reversal, and purification of the ChIP DNA were performed by following previous procedures [36]. We used agarose gel to cut DNA fragments between 150 and 250 bp, which were then used for library construction with NEXTflex ChIP-Seq Kit (Bioo Scientific). After sequencing the libraries on HiSeq 2000 system (Illumina), the reads were mapped to the genome of P. aeruginosa PAO1 with TopHat (Version 2. 0. 0) [37]. MACS software (version 2. 0. 0) were then used to call peaks [20], which was subjected to MEME analyses to calculate the specific CdpR-binding motif [38]. Plasmids p-cdpR, p-clpS/clpA, and p-clpP were constructed respectively by amplifying fragments with the corresponding primer pairs (S4 Table) pAK-cdpR-A/pAK-cdpR-S, pAK-clpS-A/pAK-clpA-S, and pAK-clpP-A/pAK-clpP-S by PCR. The PCR products were digested with the indicated enzymes and cloned into PAK1900 [39]. For construction of CTX-clpP and CTX-clpA/clpS, the sequence including its promoter and coding region was PCR-amplified with primers: CTX-clpP-F/CTX-clpP-R and CTX-clpA/clpS-F/CTX-clpA/clpS-R. These fragments were respectively ligated with Mini-CTX-lux [40]. The mini-cdpR-flag-A/mini-cdpR-flag-S primers were used to amplify the cdpR gene that intended to fuse with a C-terminal Flag-tag. The indicated enzyme-digested PCR products were cloned into the corresponding enzyme sites of Mini-CTX-lacZ to generate either Mini-CTX-cdpR-flag. The plasmid pMS402 carrying a promoterless luxCDABE reporter gene cluster was used to construct promoter-luxCDABE reporter fusions of the cdpR as previously described [18]. The cdpR promoter region was amplified by PCR using the primers cdpR-lux-F (with XhoI site) and cdpR-lux-R (with BamHI site) (S4 Table). The PCR products were cloned into the pMS402, yielding p-cdpR-lux. Besides the plasmid-based reporter system, an integration plasmid CTX6. 1 originating from plasmid mini-CTX-lux was used to construct chromosomal fusion reporter. The pMS402 fragment containing the kanamycin-resistance marker, the MCS, and the promoter-luxCDABE reporter cassette was then isolated and ligated to CTX6. 1, yielding CTX-cdpR-lux. The plasmid generated was first transferred into E. coli SM10-λ pir, and the P. aeruginosa reporter integration strain was obtained using biparental mating as previously reported [41]. All constructs were sequenced to verify that no mutations occurred for these constructs. For construction of gene knockout mutants, a SacB-based strategy was employed as described in previous and our studies [42]. To construct the cdpR null mutant (ΔcdpR), PCRs were performed to amplify sequences upstream (1,975 bp) and downstream (1,242 bp) of the intended deletion. The upstream fragment was amplified from PAO1 genomic DNA using primer pair, pEX-cdpR-up-S, and pEX-cdpR-up-A, while the downstream fragment was amplified with primer pair, pEX-cdpR-down-S, and pEX-cdpR-down-A (S4 Table). The two PCR products were digested and then cloned into BamHI/HindIII-digested gene replacement vector pEX18Ap, yielding pEX18Ap-cdpR. A 0. 9 kb gentamicin resistance cassette cut from pPS858 with XbaI was cloned into pEX18Ap-cdpR, yielding pEX18Ap-cdpR-Gm. The resultant plasmids were electroporated into PAO1 with selection for gentamicin resistance. Colonies showing both gentamicin resistance and loss of sucrose (5%) susceptibility were selected on LB agar plates containing 50 μg/ml of gentamicin and 5% sucrose, which typically indicates a double-crossover event and thus of gene replacement occurring. The pEX18Ap-cdpR-Tc was constructed by a similar strategy as described above. A 2. 3 kb tetracycline resistance cassette was amplified from integration vector mini-CTX-lacZ with primer pair Tc-S/Tc-A (with XbaI site) (S4 Table) for replacing the cdpR gene in PAO1. The ΔcdpR mutant was further confirmed by PCR. The pqsH, clpS/clpA, and clpP mutant was generated by a similar strategy with deletion of cdpR gene in PAO1. For generating cdpR/pqsH, cdpR/clpSclpA, and cdpR/clpP deletion strains (ΔcdpRΔpqsH, ΔcdpRΔclpSΔclpA, ΔcdpRΔclpP), the cdpR gene in ΔpqsH, ΔclpSΔclpA, and ΔclpP mutant was deleted by a similar strategy with plasmid pEX18Ap-cdpR-Tc. These resultant mutants were verified by PCR. The plasmid encoding the His-Sumo-PaCdpR was constructed, through PCR reaction and DNA ligation, and transferred into E. coli strain BL21 DE3. The recombinant strains were cultured in 1L LB medium supplemented with 50 ug/ml kanamycin at 37°C. Protein expression was induced at OD600 ≈ 0. 6 by addition of isopropyl β-D-1-thiogalacto-pyranoside (IPTG) with a final concentration of 0. 2 mM. The induced cultures were then grown at 18°C for additional 18 h. The cells were harvested by centrifugation and resuspended in lysis buffer (20 mM Mes pH 6. 5,200 mM (NH4) 2SO4,200 mM NaCl, 25 mM Imidazole pH8. 0) and then were lysed twice under high pressure via JNBIO homogenizer. The homogenate was clarified by centrifugation, and the supernatant was loaded onto Ni-NTA column (GE healthcare). The protein was eluted by elution buffer (20 mM Mes pH 6. 5,300 mM (NH4) 2SO4,150 mM NaCl, 500 mM Imidazole pH 8. 0) using a stage-wise gradient. The fractions containing the recombinant His-Sumo-PaCdpR protein were pooled and dialyzed against Buffer S (20 mM Mes pH6. 5,200 mM (NH4) 2SO4,200 mM NaCl, 1‰ β-Me) for three hours at room temperature with Ulp1 protease added. The cleaved sample was not stable during dialysis and additional 100 mM (NH4) 2SO4,100 mM NaCl, and 2‰ β-Me were added to redissolve the protein precipitation. The sample was loaded onto Ni-NTA column again to remove the cleaved His-SUMO tag. Target PaCdPR protein was contained in the flow-through, and its purity was analyzed by SDS-PAGE. Protein was concentrated via Amicon-Ultra centrifugal device from Millipore and stored at -80°C freezer until use. Selenomethionine-substituted PaCdPR were expressed in M9 medium supplemented with 60 mg/L Se-Met (J&K), its purification procedure is similar as the full-length native protein. The purity was verified by SDS-PAGE gel (S1A Fig). Crystallization was performed at 16°C using the Gryphon robot system from Arts Robbin Instrument Company, which identified the initial crystallization condition for full-length PaCdpR. The condition is composed of 1. 0 M (NH4) 2HPO4 and 0. 1 M Acetate (pH4. 5), which gives very tiny rod-shaped crystals. Relative larger crystals of Se-PaCdpR (0. 1×0. 1× 0. 2 mm) were obtained after several runs of optimization. The protein concentration is 3 mg/ml, and the well solution is similar as that of native proteins. 0. 001 M TCEP was added to the crystallization condition as additive. Crystals of Se-PaCdpR were cryoprotected by soaking in the mother liquid supplemented with 20% glycerol for 30 sec and then were flash-frozen by liquid nitrogen. The X-ray diffraction data was collected on beamline BL17U at Shanghai Synchrotron Radiation Facility (SSRF) at cryogenic temperature, maintained with Cryogenic system. One single crystal was used, and data processing was carried out with HKL2000 [43]. The data collection and processing statistics were summarized in S5 Table. The structure of Se-PaCdpR was solved using the SAD method with the Autosol program [44] embedded in the Phenix suite, which built an initial model that covered about 75% of the residues. The refinement was done using the Refmac5 program of ccp4 [45]; and during refinement, 5% of the data was randomly selected and set aside for free R-factor cross validation calculations. The 2Fobs-Fcalc and Fobs-Fcalc electron density maps were regularly calculated and used as a guide for the building of the missing amino acids and solvent molecules using Coot [46]. The Rwork and Rfree of the final model are 18. 8% and 23. 4%, respectively. The rmsd of bond and angle is 0. 005 Å and 0. 963°, respectively. Other refinement parameters were also summarized in S5 Table. The structure factors and atomic coordinates can be found in the Protein Data Bank with the access code 5CHH. The complex structures have been solved for two AraC family proteins, EcMarA (PDB code: 1BL0) and sgAdpA (PDB code: 3W6V). The overall structure of PaCdpR HTH is more similar to sgAdpA HTH than EcMarA HTH, with rmsds of 1. 7 Å and 2. 1 Å, respectively. In addition, the target DNAs of sgAdpA and PaCdpR are GC rich, whereas it is AT rich for EcMarA targeting DNA; therefore, the sgAdpA HTH–DNA structure was used as reference during the model building. The PaCdpR HTH–DNA complex was modeled in three steps. In the sgAdpA HTH–DNA complex structure, two symmetry-related DNA duplexes interact with the two DNA recognition helices (which are α15and α18 in PaCdpR) from one protein molecule; and the two DNA duplexes are not broken, due to the lacking of 5' -phosphate groups. Therefore, the first step was the generation of 5' -phosphate groups and bridging it with the 3' -OH of its neighboring nucleotide, resulting continuous DNA duplex. Second, the modified DNA was docked on the PaCdpR HTH structure, via the superimposing of the modified sgAdpA HTH–DNA structure with PaCdpR HTH. The rmsd between the overall structure of PaCdpR HTH and sgAdpA HTH is 1. 7 Å, based on 111 pairs of Catoms. Finally, the side chains of some residues, including Arg260, Arg263, Arg264, Arg265, Arg274, and Arg317, were adjusted to avoid the clashes between protein and DNA. No further conformational change or energy minimization was involved in the modeling. CdpR proteins were mixed with DNA probes (S4 Table) in 20 μl of the gel shift-loading buffer (20 mM HEPES, pH 8. 0,100 mM NaCl, 0. 5 mM dithiothreitol, 10% Glycerol, and 3 μg/ml sheared salmon sperm DNA). After incubation at room temperature for 20 min, the samples were analyzed by 6% polyacrylamide gel electrophoresis in 0. 5×TBE (Tris/Boric Acid/EDTA) buffer at 90 V for 90 min. The gels were stained by SYBR GOLD dye and subjected to screen on a phosphor screen (Tanon 5500). The DNA footprint assay was performed by following previous procedures [18,47]. A 588-bp DNA containing the pqsH promoter region (–500 to +88) was amplified with primers pqsH-gf (with a 6-FAM modification at the 5’) and pqsH-gr (S4 Table). Forty nM of 6-FAM-labeled pqsH promoter probe was incubated with 2 μM of CdpR in gel-shift loading buffer. The protein-DNA mixtures were then partially digested with 0. 05 units of DNase I (NEB) for 5 min at 25°C. The reaction was quenched by 0. 25 M EDTA and purified with phenol-chloroform-isoamylalcohol (25: 24: 1) and then QIAquick PCR Purification kit (Qiagen). Control samples were done without CdpR protein. The genotype samples were run with the 3730 DNA Analyzer, and viewed with Peak Scanner (Applied Biosystems). The PAO1 containing the CTX-cdpR-lux reporter fusion was subjected to transposon mutagenesis using the mariner transposon vector pBT20 [48]. Briefly, the donor strain (E. coli SM10-λpir) containing pBT20 and recipient PAO1-containing CTX-cdpR-lux were scraped from overnight plates and resuspended in 1 ml of M9 minimal medium. The bacterial suspensions were adjusted to an OD600 of 40 for the donor and an OD600 of 20 for the recipient. Next, 25 μl of each donor and recipient were mixed together and spotted on a dry LB agar plate and incubated at 37°C for overnight. Mating mixtures were scraped and resuspended in 1 ml of M9 minimal medium. Transposon-mutagenized bacteria were selected by plating on PIA plates containing gentamicin at 150 μg/ml. A transposon mutant library was constructed by picking 20,000 colonies grown on these selective plates. The mutants with altered expression of CTX-cdpR-lux were selected. The transposon insertion sites were determined by arbitrary primed PCR and subsequent sequencing of the PCR product [49]. Bacterial two-hybrid experiments were accomplished using the BacterioMatch II Two-Hybrid System Vector Kit (Agilent Technologies). The fragments of CdpR were cloned into the bait vector pBT in order to create a fusion protein with λ repressor protein (λcI). The DNA fragment of ClpS was inserted into the vector pTRG in frame with the α-subunit of RNA polymerase. The pBT-derived plasmids and pTRG-derived plasmid were then cotransformed into the validation reporter strain. Then, the resulting strains, grown at 37°C in SOC medium for 90 min, were harvested by centrifugation (5,000 g, 2 min) and washed twice with M9 + His-dropout broth. The cells were diluted to different gradients, spotted on the nonselective screening medium and selective screening medium, and finally incubated at 30°C for 2–4 d. The colonies grown on the selective screening medium were selected and streaked on the dual screening medium for further verification. The cotransformant containing pBT-LGF2 and pTRG-Gal 11P plasmids was used as the positive control. Expression of lux-based reporters from cells grown in liquid culture was measured as counts per second (cps) of light production in a Synergy 2 (Biotek) as previously described [18]. Overnight cultures were diluted to OD600 = 0. 2 with fresh LB medium, and then cultured for another 2 h. The cultures were added into a black 96-well plate with a transparent bottom. Sixty μl of sterilized mineral oil was added to culture. Promoter activities bacterial growth (OD = 595) were continuously measured every 30 min for 24 h in a Synergy 2 Plate Reader (BioTek). The in vivo degradation assay was carried out to assess the stability of CdpR-Flag protein in P. aeruginosa as previously described [50,51]. Briefly, overnight LB cultures of tested strains were diluted 100-fold in fresh 20 ml LB medium in an Erlenmeyer flask with a flask volume-to-medium volume ratio of 5: 1, and were aerated by shaking at 220 rpm. When OD600 value of the culture reached to approximately 0. 4, chloramphenicol (a final concentration at 75 μg/ml) was added to the culture in order to block the translation, and samples for western blot analysis were removed at the indicated times. For the detection of CdpR-Flag, overnight LB cultures of the indicated strains were 1: 100 diluted with LB medium recultured until A600 = 0. 6. One hundred μl cultures were washed and resuspended in 15 μl PBS buffer. The samples were solubilized in the SDS-PAGE loading buffer (50 mM Tris-HCl, pH 6. 8; 2% SDS; 0. 1% bromophenol blue; 1% mercaptoethanol; 10% glycerol) and then heated at 100°C for 15 min. SDS polyacrylamide gel electrophoresis was carried out according to the method of Laemmli [52] using a 10% slab gel with a 5% stacking gel and transferred onto PVDF (Bio-Rad) membranes. And then incubated with a mouse anti-Flag antibody (AOGMA, AGM12165) or anti-RNAP (Neoclone, WP003), followed by a sheep anti-mouse IgG antibody conjugated to horseradish peroxidase (HRP) (Code#: NA931, GE Healthcare), respectively. The relative abundance was determined by densitometric analysis using ImageQuant software. Biofilm formation was measured in a static system as previously described [53], with minor modifications. Visualization of biofilm formation was carried out in 15-mL borosilicate tubes. Briefly, cells from overnight cultures were inoculated at 1: 100 dilutions into LB medium supplemented with appropriate antibiotics and grown at 30°C for 10 h. Biofilms were stained with 0. 1% crystal violet (CV) and tubes were washed with water to remove unbound dye. Quantification of biofim formation was performed in 24-well polystyrene microtiter plates. LB and appropriate antibiotics was inoculated to a final OD600nm of 0. 01. The plates were incubated for 8 h or 14 h at 30°C. Crystal violet was added to each tube and stained for 15 min prior to removal by aspiration. Wells were rinsed three times by submerging the tubes in distilled water, and the remaining crystal violet was dissolved in 1 ml of 95% ethanol. A 1 ml portion of this solution was transferred to a new polystyrene tube, and the absorbance was measured at 600 nm. Pyocyanin was extracted from culture supernatants and measured using previously reported methods [54]. Briefly, 3 ml chloroform was added to 5 ml culture supernatant. After extraction, the chloroform layer was transferred to a fresh tube and mixed with 1 ml 0. 2 M HCI. After centrifugation, the top layer was removed and its A520 was measured. The amount of pyocyanin, in μg/ml, was calculated using the following formula: A520/A600 × 17. 072 = μg of pyocyanin per ml. Quantification of PQS production as described previously [55]. Briefly, singles colonies were inoculated in 10 ml LB for 16 h at 37°C. Cultures were diluted 1: 100 into fresh media and grown for 24 h as above. A 500 μl aliquot of each culture was mixed with 1 ml of acidified ethyl acetate, vortexed vigorously for 2 min, and then centrifuged for 10 min at 16,000 × g. The organic phase was transferred to a fresh tube and dried to completion. The solute was dissolved in 50 μl of a 1: 1 mix of acidified ethyl acetate: acetonitrile for analysis. The autoinducer of the rhl system, C4-HSL, was measured using an rhlA promoter-based P. aeruginosa, pDO100 (pKD-rhlA) as previously described [56]. Two microlitres of test bacterial culture (OD600 = 1. 0) were inoculated onto the seeded bioassay plates, and the plates were incubated at 37°C for 18 h. The dark halo zone around bacterial colonies indicates AHL activity. Overnight culture of bacteria were 1: 100 diluted with fresh LB medium and recultured until OD600 = 0. 6. Six-week-old female C57BL6 mice were bought from the Harlan Laboratory (Indianapolis, IN). The animal procedures have been approved by the University of North Dakota institutional animal care and use committee (UND IACUC). Mice were anesthetized with 40 mg/kg ketamine plus 5 mg/kg diazepam. 1 × 107 CFUs of P. aeruginosa were intranasally instilled into mice and survival rates were calculated for every bacterial strain. AM cells were cultured in 96-well plates at 37°C with 5% CO2 overnight. NBT dye (Sigma) was added to AM cells following the manufacturer’s procedures. The yellow-colored NBT changes to blue when oxidized by superoxide from AM cells [57]. The plate was kept at room temperature overnight for complete formation of dye product, which is monitored by a plate reader at 560 nm. Each experiment was conducted in three repeats. After BAL procedures and serum collection, lung tissues were fixed in 10% formalin using a routine histologic procedure. Ten μl of BAL and serum were applied on microscope slide. The PMN numbers were counted under a light microscope using a Hema staining kit (Thermofisher). Homogenizations of lung tissues were done using liquid nitrogen then dissolved in RIPA buffer and sonicated (10 s sonication with 10 s intervals, 3 times) for next analysis. Tissue damage was determined by H&E staining in formalin-fixed tissues [57].
summary: Although many transcriptional regulators tune P. aeruginosa virulence factor expression and secretion, the molecular mechanisms of the underlying regulatory network are still elusive. Quorum sensing, the ability of bacteria to communicate and detect cell density to determine the most advantageous time to orchestrate collective events, is known to govern P. aeruginosa virulence. In this study, we present a novel AraC-family transcription factor, CdpR (PA2588), that controls numerous virulence factors via directly regulating the quorum sensing Pseudomonas quinolone signal (PQS) system. We solved the crystal structure of CdpR, which showed that its N-terminal domain contains a unique fold that is different from other AraC-family proteins. In addition, we found that CdpR is regulated by the ClpAS-ClpP protease. CdpR interacts with the adaptor protein ClpS and then is degraded by the ATP-dependent ClpAP protease. This is the first example of a quorum-sensing regulator as a substrate of the ClpAS-ClpP protease in P. aeruginosa. These findings significantly extend our understandings of quorum sensing and virulence regulation and provide insights into the function of proteases in pathogenic bacteria.
keywords: sequencing techniques
  medicine and health sciences
  pathology and laboratory medicine
  enzymes
  pathogens
  gene regulation
  microbiology
  enzymology
  pseudomonas aeruginosa
  animal models
  regulator genes
  model organisms
  gene types
  sequence motif analysis
  molecular biology techniques
  dna
  bacteria
  bacterial pathogens
  promoter regions
  research and analysis methods
  pseudomonas
  sequence analysis
  proteins
  medical microbiology
  gene expression
  microbial pathogens
  mouse models
  pathogenesis
  molecular biology
  biochemistry
  nucleic acids
  virulence factors
  genetics
  biology and life sciences
  proteases
  organisms
title: Structural and Molecular Mechanism of CdpR Involved in Quorum-Sensing and Bacterial Virulence in Pseudomonas aeruginosa