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Myxococcus xanthus possesses a form of surface motility powered by the retraction of the type IV pilus (T4P). Additionally, exopolysaccharide (EPS), the major constituent of bacterial biofilms, is required for this T4P-mediated motility in M. xanthus as the putative trigger of T4P retraction. The results here demonstrate that the T4P assembly ATPase PilB functions as an intermediary in the EPS regulatory pathway composed of the T4P upstream of the Dif signaling proteins in M. xanthus. A suppressor screen isolated a pilB mutation that restored EPS production to a T4P− mutant. An additional PilB mutant variant, which is deficient in ATP hydrolysis and T4P assembly, supports EPS production without the T4P, indicating PilB can regulate EPS production independently of its function in T4P assembly. Further analysis confirms that PilB functions downstream of the T4P filament but upstream of the Dif proteins. In vitro studies suggest that the nucleotide-free form of PilB assumes the active signaling conformation in EPS regulation. Since M. xanthus PilB possesses conserved motifs with high affinity for c-di-GMP binding, the findings here suggest that c-di-GMP can regulate both motility and biofilm formation through a single effector in this surface-motile bacterium.
Myxococcus xanthus is a motile bacterium well adapted to life on a solid surface. While a unicellular organism, it possesses a social or multicellular life style during both its vegetative and developmental phases1, 2. During vegetative growth, groups of M. xanthus cells swarm over solid surfaces to digest other bacteria and organic macromolecules as nutrients3. Its developmental program, which is triggered by nutrient limitation, involves the aggregation of ~105 cells on surfaces by directed cell movement4, 5; these multicellular aggregates eventually mature into fruiting bodies when vegetative cells within cease to move and differentiate into dormant spores embedded in an exopolysaccharide (EPS) matrix6, 7. When environmental conditions become conducive for growth, spores can germinate to resume proliferation and swarming. Motility is critical to this bacterium because both vegetative predation and developmental aggregation require cells to move on solid surfaces.
The surface motility of M. xanthus entails social (S) and adventurous (A) gliding systems4, 8. Mutations inactivating one system rarely affect the other, indicating the existence of two sets of motility machineries9–11. Phenotypically, A motility is functional even when cells are well isolated, whereas S motility requires cells to be in close proximity or in groups9. There is evidence that traveling motor complexes within cells drive A motility12–15. The motor for S motility is the type IV pilus (T4P)16–19. The T4P protein filament, which mostly localizes to one cell pole in M. xanthus at a given time20, 21, is also known to power the related twitching motility in other bacteria22, 23. The distal end of a T4P first attaches to an anchor and its ensuing retraction at the proximal end pulls the cell forward24, 25. In M. xanthus, the preferred anchor for T4P retraction is EPS either associated with the cell surface or deposited on a solid substratum by this bacterium26, 27. This preference explains the “social” aspect of T4P-mediated motility in M. xanthus because a cell requires its EPS-bearing or EPS-depositing neighbors to anchor and activate the retraction of its T4P motor. EPS, which is a key constituent of the matrix for biofilms and for the organization of M. xanthus fruiting bodies6, 7, has been shown recently to affect M. xanthus motility behavior independently of its role as anchors for T4P retraction in S motility28.
The level of EPS in M. xanthus is regulated by a signal transduction pathway consisting of the Dif chemosensory proteins as well as the T4P machinery (T4PM)24, 29–31. DifA, DifC and DifE resemble the methyl-accepting chemoreceptor proteins (MCPs), the scaffold CheW, and the histidine kinase CheA, respectively27. These three proteins form a transmembrane (TM) signaling complex that positively regulates EPS production through the kinase DifE32, 33. DifD, a CheY-like substrate of DifE phosphorylation, functions as a phosphate sink to negatively regulate EPS production29, 32. DifG, a homologue of the CheC phosphatase, is a negative regulator that can dephosphorylate DifD-phosphate29, 32. The occurrence of T4P correlates closely with that of EPS31. T4P− mutants are EPS- whereas the hyperpiliated pilT mutant is EPS+. Since mutations in dif are epistatic to those in T4P or pil genes, T4P has been proposed to function as a sensory apparatus that perceives and transmits signals to the Dif proteins downstream. The communication between T4P and Dif is mediated in part by the negative regulator StkA, a DnaK-like protein34, 35 that acts downstream of T4P but upstream of Dif36. Many questions remain concerning the mechanism of EPS regulation by this pathway although the transcription of eps genes does not appear to be the target of this regulation37.
We demonstrate here that the M. xanthus T4P assembly ATPase PilB21 functions in a regulatory capacity in signaling EPS production independently of T4P assembly. Genetic studies uncovered that mutations in pilB can suppress the EPS defects resulting from the deletion of the pilin gene pilA. A mutation known to eliminate the ATPase activity of PilB and its ability to support T4P assembly21 was found to strongly suppress the EPS defect of the pilA deletion strain. This observation indicates that the role of PilB in EPS regulation can be independent of its role as the T4P assembly ATPase. Analysis in vitro suggests that it is the nucleotide-free or the apo form of PilB that actively signals EPS production. Our results support the conclusion that PilB functions in a signaling capacity with dual roles in the regulation of motility and biofilm formation in M. xanthus.
To understand how the T4P filament functions to regulate EPS in M. xanthus, the pil genes encoding the T4P structural proteins were targeted in a genetic screen for suppressors of a pilA deletion (ΔpilA)17, 38. For this suppressor screen, 11 genes in three clusters at the M. xanthus pil locus (Fig. S1) were mutagenized. These are the pilB, pilT and pilC (pilBTC) genes in one cluster, pilG, pilH and pilI (pilGHI) as well as pilM, pilN, pilO, pilP and pilQ (pilMNOPQ) in two other clusters. We first deleted pilBTC, pilGHI and pilMNOPQ as individual clusters in the WT and the ΔpilA backgrounds. These mutants were confirmed to be defective in both S motility and EPS production (Fig. S2)31. Next, three complementation plasmids containing pilBTC, pilGHI and pilMNOPQ were constructed. These plasmids, which can integrate into the M. xanthus chromosome at a phage attachment site39, were transformed into and demonstrated to complement its corresponding deletions in S motility and EPS production in the WT background (Fig. S2). Transforming these plasmids into their respective deletion mutants in the ΔpilA background resulted in strains without S motility and EPS (Fig. S2) as expected for a ΔpilA strain31.
The three plasmids were mutagenized in an Escherichia coli mutator strain40, and pools of mutagenized plasmids were isolated and transformed into their respective deletion strains in a ΔpilA background for the suppressor screen. Approximately 20,000 transformants for each plasmid were screened on plates with Congo red where EPS+ colonies appear red but EPS− ones are unstained29, 35. Transformants of the mutated pilBTC plasmid yielded five red colonies but no transformant of the other two plasmids appeared EPS+. The five putative ΔpilA suppressor strains were confirmed to be EPS+ by an EPS binding assay using the fluorescent dye Calcofluor White. Genetic mapping by transformation using genomic DNA39, 41 determined that two out of the five isolates likely had suppressor mutations in the mutagenized pilBTC genes as they are linked to the kanamycin resistant (KanR) marker carried by the integrative plasmid. The other three, which must have occurred elsewhere29, 36, 39, 41, 42, were not pursued further in this study.
The mutations in the pilBTC gene cluster from the two EPS+ strains were identified by cloning and DNA sequencing as described in Methods. The same G to A transition mutation in pilB was found in both suppressor strains. This G to A mutation, referred to as pilB* hereafter, occurred at the third position in codon 388 of pilB, resulting in a methionine (M) to isoleucine (I) substitution (M388I) (Fig. S3). PilB is an ATPase with the conserved Walker A (WA) and Walker B (WB) boxes21, 43 and the M388I substitution resides in its Walker B box (Fig. S3A). Since no other mutation in pilBTC was found in the two suppressor strains, they probably originated from the same mutated plasmids.
To verify that the pilB* mutation was solely responsible for suppression of ΔpilA in the suppressor strains, the pilB M388I mutation was reconstructed by targeted mutagenesis in the plasmid containing the WT pilBTC gene cluster. When this plasmid with the M388I mutation was transformed into the pilABTC quadruple deletion (ΔpilABTC) mutant, the resulting strain had its EPS production restored (Fig. 1A). In addition, the M388I mutation was constructed on a plasmid containing pilB only. When transformed into a ΔpilA ΔpilB double mutant, this plasmid restored EPS production to this double deletion strain (Fig. 1B). These results indicate that the pilB* single mutation alone is sufficient to suppress the EPS defect resulting from ΔpilA.
We examined the relationship between PilB and other known EPS regulators (Fig. 2A) by genetic epistasis (Fig. 2B). stkA encodes a DnaK homologue that negatively regulates EPS production34–36. A ΔpilB ΔstkA double mutant was constructed and it was found to be EPS+. Likewise, a ΔdifD ΔdifG double mutation restored EPS production to a ΔpilB mutant as it did to ΔpilA 31. In contrast, a difE − pilB* double mutant was found to be EPS− similar as the single difE − mutant. Since the suppression of ΔpilA by pilB* showed that PilB functions downstream of the T4P filament in EPS regulation, these observations support a model wherein PilB acts as an EPS regulator downstream of the T4PM but upstream of StkA and the Dif pathway44.
A strain with either the ΔpilB mutation or the WT pilB (pilB WT) in a ΔpilA background is EPS− (Fig. 1), indicating that neither the pilB WT allele nor the null or loss-of-function (LOF) ΔpilB mutation can support EPS production in a ΔpilA background. The pilB* mutation is therefore likely gain-of-function (GOF) in EPS regulation instead of WT or LOF. If so, it should be dominant over pilB WT with regard to the EPS phenotype. To test this, a pilB WT/pilB* merodiploid was constructed in ΔpilA background by transforming the pilB*-containing plasmid into a ΔpilA mutant. As shown in Fig. 1B, the resulting strain with these two pilB alleles was EPS+, indicating pilB* is dominant over pilB WT and confirming that pilB* is indeed a GOF mutation with regard to EPS regulation.
Since PilB is the T4P assembly motor ATPase, the effect of the pilB* mutation on M. xanthus S motility in an otherwise WT background was examined as well. The pilB* and pilB WT alleles were introduced into a ΔpilB single mutant. The resulting strains were examined for S motility (Fig. 3A) and EPS production (Fig. 3B). As indicated by the clear binding of Calcofluor White in EPS assays, both the pilB WT and pilB* restored EPS production to the pilB mutant as expected. Interestingly, pilB* also complemented ΔpilB in S motility as analyzed on a soft agar plate. These results demonstrate that while pilB* is a GOF mutation in EPS regulation, it is WT with regard to T4P assembly. PilB* therefore likely retains sufficient ATPase activity to support T4P assembly and S motility in vivo despite the significant alteration in its function in EPS regulation. These observations suggest that the ability of PilB to signal EPS production and its role as the T4P assembly ATPase may be genetically separable and functionally distinct.
The WA and WB boxes of PilB, while separated by 56 amino acids in the primary sequence (Fig. S3A), are packed against each other in the tertiary structure of the protein43, 45. In the model of M. xanthus PilB (MxPilB) (Fig. 4), based on the crystal structure of Thermus thermophilus PilB/PilF (TtPilB)43, the WA box forms the C-terminal part of an α helix and a loop that connects the helix to a β strand. This strand is part of an antiparallel β sheet that lines one side of this α helix. M388 in the WB box is positioned in a strand in the center of this β sheet. The side chain of this methionine packs tightly against the α helix encompassing WA. Substituting this methionine with a bulkier isoleucine in MxPilB or TtPilB is expected to create clashes that would need to be resolved by altering the relative position between WA and WB. Despite its location in the WB box, the M388I substitution therefore may disturb the structure, orientation or conformation of both WA and WB.
It is known that residues in both WA and WB are crucial for the activity of many ATPases because they interact with the bound ATP molecule43, 46–48. Although the M388I mutation apparently does not eliminate the ATPase activity of MxPilB, it may still affect its ATP binding and/or ATPase activity. To test whether the ATPase activity of PilB has an effect on EPS regulation, we constructed mutations known to have more drastic effect on the ATPase activity of PilB. The strictly conserved residues K327 in WA and E391 in WB (Fig. 4) are known to be required for the ATPase activity of MxPilB in vitro and its ability to support T4P assembly and S motility in vivo 21. We constructed K327A (pilB WA) and E391A (pilB WB) substitution mutations, respectively, and confirmed that neither mutation supported M. xanthus S motility. As shown in Fig. 5A, pilB WA but not pilB WB restored EPS production to a ΔpilA mutant. pilB WA is in fact a more robust suppressor than pilB* because its presence in the ΔpilAB background results in a higher EPS level when compared to pilB* (Fig. S4). In addition, a pilB WA /pilB WT merodiploid strain in a ΔpilA background is EPS+ (Fig. 5B), and the dominance of pilB WA indicates that it is a GOF mutation in EPS regulation as pilB*. Because PilBWA fails to support S motility in vivo and showed no ATPase activity in vitro 21, the results here (Fig. 5) demonstrate that the role of PilB in EPS production is distinct from its function as the T4P assembly ATPase. That is, in the absence of pilA, the catalytically inactive PilBWA not only supports EPS production, but it is even more potent in doing so than its enzymatically active counterparts PilBWT and PilB* (Figs 1, ,55 and S4). The stronger EPS phenotype of the pilB WA mutant further strengthens the conclusion that the activation of EPS production by PilB does not require or can bypass its function in T4P assembly. These observations support PilB as a signaling or regulatory protein with a more direct role in EPS regulation and that the PilBWA variant may assume a more active signaling conformation without the pilus filament.
We explored the possibility that the EPS+ phenotype of pilB* and pilB WA mutations may have been due to altered the protein level of PilB by performing immunoblotting using anti-PilB antibodies21. The levels of PilB in the WT and the ΔpilA backgrounds were virtually indistinguishable (Fig. 6A), suggesting that the ΔpilA mutation itself does not alter PilB expression or stability. However, the levels of both PilB* and PilBWA were lower than PilBWT in an isogenic ΔpilA background (Fig. 6B). Since the experiments were conducted with strains where all pilB variants were expressed from the same promoter49, it is unlikely that transcription or translation are affected by these mutations. Instead, it is more likely that the stability of the protein in vivo was changed by these mutations. Regardless, since a pilB null mutation does not suppress the EPS− phenotype of ΔpilA, only an increase in PilB protein level could explain the suppression of ΔpilA by pilB* and pilB WA, but not a decrease. We propose that PilB* and PilBWA mutations changed the structure of the protein to favor its active signaling conformation and that this conformational change may have reduced the stability of PilB in vivo coincidentally.
To gain insights into the mechanisms of PilB in EPS regulation, we attempted to purify and examine the properties of PilBWT, PilBWA and PilBWB proteins in vitro. These three PilB variants were chosen because they reflect the three distinct phenotypes with regard to T4P assembly and EPS production in vivo. PilB* was not included in the in vitro studies because it resulted in an intermediate EPS phenotype. Purified MxPilB exhibits measurable but very low ATPase activity21 and it tends to form protein aggregates (unpublished). We henceforth expressed and purified the T. thermophilus PilBWT, PilBWA and PilBWB equivalents (Fig. S3A)50 for in vitro studies.
Differential scanning fluorimetry (DSF), which measures the thermal denauration of a protein over a temperature gradient51, 52, was used to first examine the structural differences among PilBWT, PilBWA and PilBWB. As shown in Fig. 7A, PilBWT and PilBWB behaved relatively similar in this assay: both show biphasic unfolding profiles with two melting temperatures or transition midpoints (T m) around 64–65°C and 79–81°C, respectively. In contrast, PilBWA unfolded over a broader temperature range with an apparent T m around 76°C. The obvious differences in thermal stability indicate that the structure of PilBWA, but not that of PilBWB, is significantly altered in comparison with PilBWT. We also examined the effect of AMP-PNP, a non-hydrolyzable ATP analogue53, 54, on PilB thermal stability (Fig. 7A). This nucleotide stabilized both PilBWT and PilBWB, resulting in the disappearance of the first unfolding transition. In contrast, the stability of PilBWA remained virtually unchanged by the addition of AMP-PNP. These results demonstrate that PilBWA no longer responds to its ligands and that its structure is distinct from those of PilBWT and PilBWB.
The circular dichroism (CD) spectra of these protein variants were collect to further explore their structural differences. In the far ultraviolet (UV) range (200–240nm), all three proteins showed similar spectra with or without ATP or ADP (Fig. S7). As the signals in this range are sensitive to changes in protein secondary structure55, the results indicate that the secondary structure content is unaffected by the mutations in PilBWA and PilBWB or by ligand binding. In the near UV range (260–320nm), changes in the CD spectra are related to protein conformation55. The signals in the 260–285nm range are contributed primarily by phenylalanine and tyrosine residues55, which the TtPilB contains 14 each (Fig. S3A). There is little signal beyond 285nm because the protein lacks tryptophan (Figs 7B and S3A). All three protein variants, PilBWT, PilBWA and PilBWB, showed similar CD spectrum without ligand. While the addition of ATP shifted the spectra of all three proteins downward in the 260–280nm range (Fig. 7B, left panel), the shift for PilBWA is less pronounced compared to those seen for the other two. The addition of ADP also shifted the spectra of PilBWT and PilBWB, but not that of PilBWA (Fig. 7B, right panel). These observations further underscored that PilBWA is diminished in its response to nucleotides when compared to PilBWT and PilBWB. In particular, PilBWA more closely resembles the ligand-free rather than the ligand-bound form of PilB in the presence of a nucleotide.
The binding affinity of PilB for ATP was examined using the ATP analogue MANT-ATP56. The fluorescence of this nucleotide is enhanced by a more hydrophobic environment frequently associated with its binding to a protein. Figure 8 shows the difference in fluorescence (ΔF) in the presence and absence of PilB variants over varying concentrations of MANT-ATP. A binding isotherm was fit to the data to estimate the binding affinity of the three PilB protein variants to this ATP analogue. The dissociation constants (Kd) for PilBWT and PilBWB were similar with values of 0.17μM and 0.19μM, respectively. The Kd for PilBWA is 1.23μM and this increase in Kd represents a significant reduction in its nucleotide binding affinity. These results suggest that the diminished response of PilBWA to nucleotides is attributed to its reduced affinity for its ligand. The in vitro studies (Figs 7 and and8)8) suggest that the nucleotide-free conformation of PilB may actively signal EPS production in M. xanthus.
This study identified the T4P assembly ATPase PilB as a regulator of EPS downstream of the T4P filament and upstream of the Dif signaling proteins in M. xanthus. A genetic screening was devised to first isolate pil mutations that could restore EPS production to a ΔpilA strain, leading to the discovery of pilB* (Fig. 1). Further genetic analysis indicated that PilB functions downstream of T4PM but upstream of the StkA and Dif proteins in EPS regulation (Fig. 2)44. Targeted mutagenesis of conserved residues revealed that the involvement of PilB in EPS regulation can be separated from and is independent of its function as the T4P assembly ATPase (Fig. 5). That is, the PilBWA mutant variant, which does not hydrolyze ATP or support S motility21, may be locked in a conformation that actively signals EPS production in a ΔpilA or a WT background (Fig. 5). Both pilB* and pilB WA are dominant over pilB WT (Figs 1 and and5),5), indicating that they are GOF mutations. In vitro studies using TtPilB indicated that the PilBWA variant showed diminished conformational response to the addition of nucleotides because of reduced binding affinity (Figs 7 and and8).8). These results lead to a model wherein the M. xanthus PilB ATPase functions as a signaling protein in EPS regulation and its nucleotide-free state may correlate with its actively signaling conformation.
As far as we are aware, XB2457 and the Na/K ATPase (NKA)58, 59 are the two other cases where ATPases have been proposed as signaling proteins. XB24 is a small cytoplasmic ATPase in rice with a role in immunity and defense against bacterial pathogens. It physically associates with and modulates the activity of the kinase XA21, which is a receptor for recognition of pathogen-associated molecular patterns (PAMPs). The ATPase activity of XB24 is required for its function in vivo and it is proposed that the signaling activity of XA21 is regulated by XB24. NKA is ubiquitous in the plasma membrane of all animal cells. It establishes and maintains the ion homeostasis essential for cell viability. All NKAs contain a binding site specific for steroid inhibitors such as ouabain and digoxin. There is evidence that NKAs can associate with and affect the activity of signaling proteins such as the Src kinase60, 61. It has been proposed that NKAs are receptors and signal transducers to regulate downstream targets in a signal transduction pathway.
Drawing analogy between G-proteins62, 63 and signaling ATPases57, 58, we hypothesized initially that it was the PilB in the ATP bound form that actively signaled EPS production. This was tested by mutating E391 in WB, a key catalytic residue for ATP hydrolysis47, 48 (Fig. 4). Since PilBWB has the same binding affinity for ATP as the PilBWT (Fig. 8) but is inactive as an ATPase21, it likely exists in its ATP-bound form in the cell. The pilB WB mutation, however, failed to suppress ΔpilA (Fig. 5), arguing against the ATP-bound PilB as the actively signaling conformation. On the other hand, the highly conserved K327 in WA is known to be critical for ATP binding47, 48 (Fig. 4). When mutated to an alanine, the resulting pilB WA turned out to be a robust suppressor of ΔpilA in EPS regulation (Figs 5 and S4). Binding assays indicated that PilBWA binds to ATP with much reduced affinity as expected (Fig. 8). Biophysical studies also suggested that the structure of PilBWA resembles the apo-form of PilBWT with or without its nucleotide ligands (Fig. 7B). We therefore propose that it is the apo form of PilB that actively signals EPS production in M. xanthus. It remains to be seen how prevalent signaling ATPases are in different biological systems and whether they share a common signaling conformation with M. xanthus PilB.
The regulation of EPS is the key aspect of bacterial biofilm formation64–66. The paradigm that emerged from the studies of biofilm in flagellated motile bacteria is the mutual exclusivity or inverse regulation of the motile state vs the biofilm state64–66. That is, the regulation is such that the motility of cells in a biofilm is inhibited whereas motile cells either exit from or exist mostly outside of biofilms in bacteria with flagellated motility. The small signaling molecule c-di-GMP is well known as the “master” regulator of the transition between the motile and the biofilm states. The overarching conclusion from studies of this signaling molecule is that it simultaneously enhances biofilm formation and inhibits flagellated motility. The known effectors or targets of c-di-GMP in biofilm regulation are diverse, ranging from biosynthetic enzymes and riboswitches to transcriptional activators and posttranslational regulators65. Nevertheless, EPS, the major constituent of bacterial biofilms, is the ultimate target of this regulation in most cases. For the regulation of flagellated motility, c-di-GMP generally targets the expression or activity of flagellar proteins. In V. cholerae, for example, such regulations are achieved by the c-di-GMP receptors and transcriptional regulators VpsR and FlrA/FliQ among others67. VpsR, which directly activates the expression of EPS or VPS (V ibrio polysaccharides) genes, binds c-di-GMP with high affinity. FlrA/FliQ is the master regulator of flagellar genes and the binding of c-di-GMP impairs its ability to activate the transcription of the flagellar operon.
More recently, a new c-di-GMP binding motif was discovered by the studies of V. cholerae MshE, the PilB equivalent in the Msh pilus system68–70. Sequence alignment with MshE shows convincingly that PilB from M. xanthus and many other T4P systems contain this c-di-GMP binding motif at their N-termini (Fig. S3B)68. This motif by itself and a few proteins with it have been verified or demonstrated to bind c-di-GMP with high affinity68. These include a PilB from Clostridium perfringens 71. It is therefore reasonable to assume that M. xanthus PilB is a functional c-di-GMP effector. With this in mind, the finding of M. xanthus PilB as an EPS regulator here suggests that c-di-GMP manages biofilm formation and T4P-mediated S motility through PilB as a direct target in M. xanthus. Drawing analogies with flagellated bacteria, we envision a similar working model for the regulation of EPS production and S motility by c-di-GMP in M. xanthus. We propose that c-di-GMP promotes EPS production and biofilm formation at high concentrations whereas the T4P-dependent S motility is favored at low or basal levels of c-di-GMP. In this model, when c-di-GMP is present at low or basal levels, PilB is active as the T4P assembly ATPase while EPS is produced only at a basal level to allow M. xanthus S motility to function26, 72, 73. When the cellular concentration of c-di-GMP is high, it binds to PilB to signal EPS production and to inhibit its activity as the T4P assembly motor simultaneously. We suggest that the binding c-di-GMP results in conformational changes in PilB mimicked or represented by the PilBWA mutant protein, and such conformational changes lead to the inhibition of ATPase activity and stimulation of EPS signaling.
Myxococcus xanthus strains used in this study are listed in Table 1. They were grown and maintained at 32°C on Casitone-yeast extract (CYE) agar plates or in CYE liquid medium74. XL1-Blue (Stratagene) and Rosetta (Novagen), the Escherichia coli strains used for plasmid construction and protein expression, were grown and maintained at 37°C on Luria-Bertani (LB) agar plates or in LB liquid medium75. Unless noted otherwise, plates contained 1.5% agar. Kanamycin and ampicillin at 100μg/ml and oxytetracycline at 15μg/ml were added to media for selection when appropriate.
Three sets of plasmids were generated to construct M. xanthus strains. The first set were constructs to delete single or multiple pil genes; these include pWB525 (ΔpilA), pWB581 (ΔpilB), pWB555 (ΔpilBTC), pWB556 (ΔpilAGHI), pWB605 (ΔpilGHI) and pWB557 (ΔpilMNOPQ). The second were for the expression of pil gene clusters and pilB alleles in M. xanthus; these constructs, which integrate at Mx8 att site76, include pWB559 (pilGHI), pWB565 (pilMNOPQ) and pWB566 (pilBTC) as well as pWB571 (pilB), pWB572 (pilB*), pGD5 (pilBWA) and pGD6 (pilBWB). The third includes the plasmid pWB606, which was used for the replacement of wild-type pilB with pilB*. The first and third sets were derivatives of pBJ11377. The second set used pWB425 as the cloning and M. xanthus expression vector39. pWB425 contains a BspHI/ApoI fragment from pZero-2 (Invitrogen) with the kanamycin resistance (KanR) gene and its promoter. The expression of all pil genes cloned into pWB425 is driven by this promoter because the multicloning site is immediately downstream of the KanR gene in this plasmid.
The details for the construction of plasmids used for M. xanthus strain construction are described here. Fragments with in-frame deletion alleles of pilA, pilBTC, pilGHI, pilAGHI and pilMNOPQ were generated by a two-step overlap PCR as described previously29. These fragments were cloned into pBJ11377 to construct pWB525 (ΔpilA), pWB555 (ΔpilBTC), pWB556 (ΔpilAGHI), pWB605 (ΔpilGHI) and pWB557 (ΔpilMNOPQ). In these plasmids, the ΔpilA allele deleted the codons from 7 to 218 of pilA, ΔpilBTC from 8 of pilB to 409 of pilC, ΔpilAGHI from 7 of pilA to 250 of pilI, ΔpilGHI from 5 of pilG to 250 of pilI and ΔpilMNOPQ from 7 of pilM to 896 of pilQ, respectively. The pilB deletion allele from DK1041618 was PCR amplified and cloned into pBJ113 to produce pWB581. For the allelic exchange of wild-type pilB with pilB*, the fragment from pWB572 (see below) was cloned into pBJ113 to create pWB606.
Plasmids that can integrate into M. xanthus chromosome at Mx8 phage attachment site (att)78 for ectopic expression were constructed using pWB42539 as the vector. pWB559, pWB565 and pWB566, which contain the respective pilGHI, pilMNOPQ and pilBTC gene clusters from pDW7979 or pSWU25717 in pWB425, were also used for mutagenesis in E. coli. With reference to the coding regions (Fig. S1), pWB559 (pilGHI) contains DNA from 73bp upstream of pilG to 328bp downstream of pilI, pWB565 (pilMNOPQ) from 120bp upstream of pilM to 19bp downstream of pilQ, and pWB566 (pilBTC) from 22bp upstream of pilB to 5bp downstream of pilC. In addition, pWB571 (pilB), which contains from 22bp upstream to 142bp downstream of pilB, was derived from pWB566. This was achieved by digestion with SacI (site in pilT) and BamHI (site on vector), treatment with T4 DNA polymerase and religation; this removed pilC and the bulk of pilT. pWB572 (pilB*), pGD5 (pilB WA) and pGD6 (pilB WB) were constructed using pWB571 as a template by a two-step overlap PCR using primers containing the M388I (pilB*), K327A (pilB WA) and E391A (pilB WB) mutations.
YZ690 through YZ1888 in Table 1 are the M. xanthus strains constructed in this study. For the construction of the pil deletions and the pilB* chromosomal replacement mutant, the plasmids with these mutant alleles were electroporated into DK1622 and used for allelic exchange as previously described29, 80, 81. These strains are YZ690 (ΔpilA), YZ1636 (ΔpilBTC), YZ1637 (ΔpilMNOPQ), YZ1639 (ΔpilAGHI), YZ1865 (ΔpilGHI) and YZ1875 (pilB*). The deletion strains YZ1638 (ΔpilBTC ΔpilA), YZ1640 (ΔpilMNOPQ ΔpilA) and YZ1682 (ΔpilB ΔpilA) were constructed similarly using YZ690 (ΔpilA) as the parent.
The plasmids constructed with pWB425 as the vector were electroporated into the appropriate deletion strains to construct YZ1644 (ΔpilBTC att::pilBTC), YZ1645 (ΔpilMNOPQ att::pilMNOPQ) and YZ1870 (ΔpilGHI att::pilGHI). Other strains similarly constructed are YZ1504 (ΔpilB ΔpilA att::pilB WA), YZ1505 (ΔpilB ΔpilA att::pilB WB), YZ1849 (ΔpilB ΔpilA att::pilB) and YZ1850 (ΔpilB ΔpilA att::pilB*) as well as YZ1512 (ΔpilB att::pilB WA), YZ1522 (ΔpilB att::pilB*), YZ1548 (ΔpilB att::pilB WB) and YZ1674 (ΔpilB att::pilB WB). The pilB merodiploid strain YZ1521 (ΔpilA att::pilB*) was constructed by transforming pWB572 into YZ690 (ΔpilA).
YZ1879 (ΔpilB stkA::tet R) and YZ1887 (pilB* difE::kan R) were constructed by transferring the stkA::tet R allele in LS1102 to DK10416 (ΔpilB) and the difE::kan R allele in SW501 to YZ1875 (pilB*) by genomic transformation41, respectively. pWB581 (ΔpilB) was used to replace pilB WT with ΔpilB in YZ641 (ΔdifD ΔdifG) to construct YZ1888 (ΔpilB ΔdifD ΔdifG).
pWB559, pWB565 and pWB566 were mutagenized by propagation in NR9458, an E. coli mutD5 mutator strain40. Cells were initially grown on plates with 1×Volgel-Bonner salts minimal media containing 0.4% glucose, 50μg/ml proline and 5μg/ml thiamine to minimize the mutation rate40. Plates containing approximately 100 colonies for each plasmid to be mutagenized were pooled and inoculated into LB broth to increase the mutation rate. Mutagenized pools of plasmid DNA were prepared from overnight cultures and used to transform the appropriate pil deletion strains. Potential EPS producing suppressor mutants were identified as reddish-orange colonies on CYE plates containing Congo red (30μg/ml)35. Genomic DNA from potential suppressor mutants was electroporated into the original parental strain and selected on plates with kanamycin and Congo red to examine the link of the EPS phenotype with the integrated plasmid39, 41. Genomic DNA from suppressor mutants was cut with PstI, religation and transformation into E. coli XL1-Blue to recover the plasmid and pil mutations were identified by DNA sequencing.
Log phase cells grown in CYE were harvested and resuspended in MOPS buffer (10mM morpholinopropanesulfonic acid [pH 7.6], 2mM MgSO4) at 5×109cells/ml. 5µl of this cell suspension was spotted onto CYE plates with 0.4% agar and regular CYE plates with Calcofluor white (50µg/ml) for S-motility and EPS analysis, respectively. Plates were incubated at 32°C for 5 days before documentation under white light for S motility and 365nm UV light for EPS production.
Plasmids pWB750, pWB751 and pWB752 (see SI Materials and Methods) were constructed and maintained in the E. coli strain XL1-Blue initially. For protein purification, they were transformed into the E. coli Rosetta strain containing pREP4 (Qiagen). The appropriate expression strains were grown at 35°C to an OD600 of 0.5–0.6 in 1 liter of LB plus ampicillin (100μg/ml) and kanamycin (25μg/ml). Protein expression was induced by addition of IPTG (Isopropyl β-D-1-thiogalactopyranoside) to a final concentration of 0.1mM, followed by incubation at 30°C for 4–5hrs. Please see SI Materials and Methods for more details for the purification.
Proteins were diluted to a final concentration of 8μM in 1× stock buffer (10mM HEPES [pH 8], 50mM KCl, 10mM MgCl2, 0.5mM EDTA, 1mM β-ME and 10% glycerol) using a 5× stock buffer. Sypro Orange (Ex. 490nm, Em. 530nm) from a 5,000× stock (Invitrogen) was diluted to 50× in deionized water and used at a 5× concentration in DSF assays. For examining the effects of ligand, adenosine 5′-(β,γ-imido)triphosphate (AMP-PNP) (Sigma-Aldrich) was added to a final concentration of 0.4mM. The experiment was carried out using a Bio-Rad CSX96 Real-Time System (Bio-Rad), starting at 25°C with temperature increments of 0.5°C to a final temperature of 99°C. Samples were held at each temperature for 30sec prior to measurement of fluorescence. Data were analyzed and Tm’s were obtained using Bio-Rad’s CFX Manager software. The data points starting at 45°C were presented in this paper.
Protein samples were adjusted to a final concentration of about 6µM for far UV (200–260nm) by diluting in storage buffer. For near UV (260–320nm), proteins were concentrating using Amicon stirred cells to 230µM and 170µM for the experiment with ATP and ADP, respectively. CD spectra were generated on a Jasco J-815 Spectropolarimeter equipped with the Jasco PFD-425S temperature-control unit in a 1-mm path-length quartz cell at 25°C. Each spectrum was from three accumulated scans with a 1second response time at a scan speed of 100nm per minute. Scans were performed with a bandwidth of 1nm in far UV and at 0.5nm increments in near UV. Data was recorded using Spectra Manager software (Jasco). For examining the effects of ligand, ATP and ADP (Sigma-Aldrich) was added at a final concentration of 0.1mM. Data was plotted after background subtraction of identical scans of controls without the protein. ATP and ADP were used in this experiment because these T. thermophilus PilB variants has no ATPase activity at 25°C (data not shown).
Binding of MANT-ATP [2′-/3′-O-(N-methylanthraniloyl) adenosine-5′-triphosphate] (Invitrogen) was monitored by measurement of fluorescent intensity using an Infinite M200 microplate reader (Tecan). The excitation was set to 356nm and fluorescence emission was recorded at 448nm. Samples with MANT-ATP at specified concentrations (0.0012 to 10μM) were incubated with or without the protein at 0.2μM at room temperature for 2hours in binding buffer [10mM Tris (pH 9), 50mM KOAc, 5mM MgOAc, and 1% glycerol] prior to fluorescence measurements. The difference in fluorescence (ΔF) with and without the protein at different MANT-ATP concentrations ([MANT-ATP]) was fitted to the binding isotherm ΔF=ΔFmax * [MANT-ATP]/(Kd+[MANT-ATP]) where ΔFmax is the maximum ΔF and Kd is the dissociation constant. Curve fitting and determination of Kd values were performed using XLfit (ID Business Solutions).
This work was partially supported by the National Institute of Health (GM071601 to Z.Y. and 1R21AI101774-01 to F.D.S), the American Heart Association (09SDG2260401 to F.D.S) and National Science Foundation (MCB-1417726 to Z.Y. and F.D.S. and MCB-1253234 to B.E.S.). L.W. is a recipient of a scholarship (File No. 201408440309) under the State Scholarship Fund from the China Scholarship Council. We are grateful to Drs. Webster L. Santos and Tijana Grove for their help with Jasco J-815 and we thank Gaurav Dogra for the constructions of pGD5 and pGD6. We are indebted to Dr. Sogaard-Andersen for PilB antibodies and Dr. Barkay for T. thermophilus HB27.
W.P.B. and Z.Y. designed research; W.P.B., L.W., J.X., R.C.S. and F.L. performed research; W.P.B., J.X., F.D.S., B.E.S. and Z.Y. analyzed data; W.P.B. and Z.Y. wrote the paper.
The authors declare that they have no competing interests.
Electronic supplementary material
Supplementary information accompanies this paper at doi:10.1038/s41598-017-07594-x
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