β-Lactams are a class of antibiotics that target the synthesis of peptidoglycan, an essential component of the cell wall. β-Lactams inhibit the function of penicillin-binding proteins (PBPs), which form the cross-links between strands of peptidoglycan. Resistance to β-lactams complicates the treatment of bacterial infections. In recent years, the spread of β-lactam resistance has increased with growing intensity. Resistance is often conferred by β-lactamases, which inactivate β-lactams, or the expression of alternative β-lactam-resistant PBPs. σP is an extracytoplasmic function (ECF) σ factor that controls β-lactam resistance in the species Bacillus thuringiensis, Bacillus cereus, and Bacillus anthracis. σP is normally held inactive by the anti-σ factor RsiP. σP is activated by β-lactams that trigger the proteolytic destruction of RsiP. Here, we identify the penicillin-binding protein PbpP and demonstrate its essential role in the activation of σP. Our data show that PbpP is required for σP activation and RsiP degradation. Our data suggest that PbpP acts as a β-lactam sensor since the binding of a subset of β-lactams to PbpP is required for σP activation. We find that PbpP likely directly or indirectly controls site 1 cleavage of RsiP, which results in the degradation of RsiP and, thus, σP activation. σP activation results in increased expression of β-lactamases and, thus, increased β-lactam resistance. This work is the first report of a PBP acting as a sensor for β-lactams and controlling the activation of an ECF σ factor.
IMPORTANCE The bacterial cell envelope is the target for numerous antibiotics. Many antibiotics target the synthesis of peptidoglycan, which is a central metabolic pathway essential for bacterial survival. One of the most important classes of antibiotics has been β-lactams, which inhibit the transpeptidase activity of penicillin-binding proteins to decrease the cross-linking of peptidoglycan and the strength of the cell wall. While β-lactam antibiotics have historically proven to be effective, resistance to β-lactams is a growing problem. The ECF σ factor σP is required for β-lactam resistance in B. thuringiensis and close relatives, including B. anthracis. Here, we provide insight into the mechanism of activation of σP by β-lactams.
The bacterial cell wall is essential for cell viability under most environmental conditions. Peptidoglycan is the major component of the cell wall and is responsible for maintaining cell shape, preventing lysis under turgor pressure, and protecting the cell from extracellular stresses. Peptidoglycan is composed of chains of repeating N-acetylglucosamine (NAG) and N-acetylmuramic acid (NAM) subunits that are cross-linked by pentapeptide side chains extending from the NAM subunits (1, 2). In Gram-positive organisms, the peptidoglycan forms a multilayer matrix that encases the plasma membrane (3).
Penicillin-binding proteins (PBPs) are some of the enzymes responsible for peptidoglycan synthesis. In the cytosol, dimers of NAG-NAM with pentapeptide side chains are synthesized and then flipped outside the cell membrane. These dimers are added to the growing peptidoglycan polymer by transglycosylation, which results in strands of repeating NAG-NAM subunits. These strands are cross-linked by transpeptidation of their pentapeptide side chains in a reaction carried out by PBPs. There are two types of high-molecular-weight PBPs. Type a PBPs have both transglycosylase activity and transpeptidase activity. Type b PBPs have only transpeptidase activity but work in concert with monofunctional SEDS (shape, elongation, division, sporulation) family transglycosylases to synthesize peptidoglycan (2, 4). The activities of type a PBPs and type b PBPs are required for cell viability (5–7).
β-Lactam and cephalosporin antibiotics inhibit peptidoglycan synthesis by forming a covalent bond with the transpeptidase active-site serine of PBPs (5, 8, 9). This inhibition prevents cross-linking of the peptide side chains, which results in peptidoglycan instability and lysis during cell growth (10). Resistance to β-lactams and cephalosporins is a growing problem that complicates the treatment of bacterial infections. Resistance to β-lactams is usually due to the secretion of β-lactamases, which destroy the antibiotic by cleaving the β-lactam ring, or mutations that lead to modification of the transpeptidase active sites of PBPs and prevent β-lactam binding (11, 12).
In response to stresses like antimicrobial peptides or antibiotics, many bacteria utilize alternative σ factors to regulate subsets of genes required for the stress response. The extracytoplasmic function (ECF) σ factor family is the largest and most diverse group of alternative σ factors and represents the “third pillar” of bacterial signal transduction (13–15). ECF σ factors are part of the σ70 family but contain only region 2 and region 4.2 of σ70. These regions bind to the −10 and −35 regions of promoters, respectively (13, 16). Many ECF σ factors are held inactive by anti-σ factors (13, 17, 18). The activation of these ECF σ factors requires release from their cognate anti-σ factors to allow the transcription of specific stress response genes.
A recent study identified >150 different families of ECF σ factors (15). The roles of the vast majority of these σ factors remain poorly understood; however, of the studied ECF σ factors, the mechanisms of ECF σ factor activation are diverse (18–20). One common mechanism known to control ECF σ factor activation is the proteolytic destruction of the anti-σ factor (18, 21). Among those ECF σ factor systems that use proteolytic destruction of the anti-σ factor, the mechanisms controlling the initiation of this proteolytic cascade are diverse (21). In Escherichia coli, the activation of σE is controlled by the binding of misfolded outer membrane proteins to the site 1 protease DegS and lipopolysaccharide (LPS) binding to RseB (a negative regulator of σE activation) (22–25). These binding events lead to the cleavage of the anti-σ factor RseA at site 1 by DegS (26). In Bacillus subtilis, the activation of σV by lysozyme is controlled by the direct binding of the anti-σ factor RsiV to lysozyme and then cleavage of RsiV at site 1 by signal peptidase (27–32).
In Bacillus anthracis, Bacillus cereus, and Bacillus thuringiensis, resistance to penicillin and other β-lactam antibiotics is dependent upon σP, an ECF σ factor (33, 34). σP was originally classified as a member of the ECF01 group of ECF σ factors but was recently reclassified to the ECF265 group, the members of which are primarily found in Firmicutes (14, 15). Approximately 50% of ECF265 σ factors are associated with an anti-sigma factor that contains a single transmembrane helix (15). Little is known about how the activity of the ECF265 σ group is controlled, and σP could represent a model to understand the activation of this subclass of ECF σ factors.
σP activity is inhibited by the anti-σ factor RsiP, which contains a single transmembrane helix. The activation of σP results in the expression of at least two genes that encode β-lactamases and are involved in resistance to penicillin, ampicillin, and other β-lactam antibiotics. σP also activates the expression of its operon, thus controlling the expression of sigP and rsiP (33, 34). We previously demonstrated that σP is activated in the presence of a subset of β-lactams, ampicillin, methicillin, cefoxitin, cephalothin, and cefmetazole, but not other cell envelope stresses (34). We also identified a subset of β-lactams that do not activate σP: piperacillin, cefsulodin, and cefoperazone (34). In response to the activating β-lactams, RsiP is destroyed by a cascade of proteases, resulting in σP activation (34). An unidentified site 1 protease initiates the proteolytic cascade by cleaving RsiP at site 1, which is then followed by cleavage at site 2 by RasP, the highly conserved site 2 protease (34) (Fig. 1B). Here, we demonstrate that β-lactam activation of σP is dependent on the PBP HD73_3488 (also known as HD73_RS17405), which we have named PbpP. Our data indicate that PbpP is required for site 1 cleavage of RsiP in response to β-lactams, but PbpP is likely not the site 1 protease. Our data suggest that PbpP likely functions as a sensor of β-lactams by directly binding β-lactams and triggering σP activation by promoting site 1 cleavage of RsiP.
PbpP is required for σP activation.
Bacillus thuringiensis, B. cereus, and B. anthracis contain two open reading frames in the sigP region that encode predicted penicillin-binding proteins (PBPs). In Bacillus thuringiensis subsp. kurstaki HD73, these PBPs are called pbpP (HD73_3488) and bt3491 (HD73_3491). We also identified a third open reading frame that appears to be found only in Bacillus thuringiensis subsp. kurstaki HD73, called bt3487 (HD73_3487) (Fig. 1A). Although they are not located in the same operon as sigP and rsiP, we hypothesized that they may play a role in the response of σP to β-lactams because PBPs have been well characterized as targets of β-lactam antibiotics (9, 35). Additionally, genes involved in the same signaling system are often located in the neighboring regions. To determine if BT3487, PbpP, and BT3491 were required for the response of σP to β-lactams, we generated strains with in-frame deletions of each of the genes and measured the effect on ampicillin resistance. We found that the deletion of pbpP led to a dramatic decrease in the ampicillin MIC similar to that of a ΔsigP mutant (Table 1) (33, 34). In contrast, strains with deletions in bt3487 and bt3491 had no effect on ampicillin resistance (not shown). We also determined that a ΔpbpP mutant is more sensitive to cefoxitin and cefmetazole than the wild type (WT) (Table 1).
Experiments were performed in biological and technical triplicate.
We noted that a ΔsigP-rsiP mutant is more sensitive to β-lactams than a ΔpbpP mutant. We hypothesized that a ΔpbpP mutant may block σP activation in response to β-lactams but retains a basal level of σP activation that allows a low level of resistance to β-lactams. To monitor σP activity, we took advantage of the fact that σP is required for the transcription of its promoter (PsigP); thus, we inserted a PsigP-lacZ promoter fusion into the thrC locus (33, 34). To determine if PbpP played a role in σP activation, we tested the effect of a pbpP deletion on σP activity by monitoring PsigP-lacZ expression. Interestingly, we did not observe activation of σP in the ΔpbpP mutant in the presence of cefoxitin (see Fig. S1A in the supplemental material). We complemented the ΔpbpP mutant with pbpP+ on a plasmid under the control of its native promoter. We found that σP was activated in the presence of cefoxitin to an extent similar to that observed for the WT (Fig. S1A). To reinforce our finding that ΔpbpP results in the loss of σP activation, we conducted β-galactosidase assays to quantify the effect on σP activation. As previously reported, PsigP-lacZ expression is induced in a dose-dependent manner in response to increased cefoxitin concentrations in the WT (Fig. 1C) (34). Consistent with previous observations, we did not observe induction of PsigP-lacZ in the ΔsigP-rsiP mutant because σP is required for transcription from PsigP (34). We found that the deletion of pbpP resulted in the loss of PsigP-lacZ expression at every concentration tested (Fig. 1C). Taken together, our data suggest that PbpP is required for the activation of σP, thereby altering the transcription of the σP regulon and β-lactam resistance.
PbpP is required for site 1 cleavage of RsiP.
Because our data suggest that PbpP is required for σP activation, we hypothesized that PbpP is required for RsiP degradation. To test this, we compared the effects of cefoxitin on the degradation of green fluorescent protein (GFP)-RsiP in WT, ΔpbpP, and ΔrasP mutant strains. We previously showed that GFP-RsiP is functional and localized to the membrane (34). We found that the levels of full-length GFP-RsiP decreased in the WT in the presence of cefoxitin (Fig. 1D) (34). When a ΔrasP mutant, which lacks the site 2 protease, was incubated with cefoxitin, we observed a decrease in full-length GFP-RsiP and the buildup of an intermediate GFP-RsiP fragment, indicating the loss of site 2 cleavage (Fig. 1D) (34). This GFP-RsiP fragment is approximately the predicted size for a site 1 protease cleavage product. In contrast, we found that full-length GFP-RsiP levels did not decrease in the ΔpbpP mutant when grown in the presence of cefoxitin (Fig. 1D). This suggests that PbpP is required for site 1 cleavage of RsiP and, thus, σP activation.
PbpP is a penicillin-binding protein.
A defining feature of PBPs is the ability to covalently bind β-lactams (9, 36). We sought to determine if PbpP has the capacity to bind β-lactams. We tested if PbpP could bind Bocillin-FL (Boc-FL), a fluorescent β-lactam consisting of penicillin V and BODIPY FL dye (37). We found that Bocillin-FL was degraded when σP was activated (Fig. S3B). In a ΔsigP-rsiP mutant, we found that Bocillin-FL was not degraded, suggesting that σP-regulated β-lactamases are likely responsible for Bocillin-FL degradation (Fig. S3B). To perform Bocillin-FL labeling experiments, we expressed pbpP from an isopropyl-β-d-thiogalactopyranoside (IPTG)-inducible promoter in a ΔsigP-rsiP mutant. We labeled cells with Bocillin-FL and blotted them with anti-PbpP antisera (37). We observed a fluorescent band at approximately 66 kDa with both Bocillin-FL and anti-PbpP antisera. This band was the predicted size of PbpP; it increased in intensity with increasing IPTG concentrations and was not observed in the empty vector (EV) control (Fig. 2A and Fig. S3A). This demonstrates that PbpP binds β-lactams. We also noted that the lack of a fluorescent band corresponding to PbpP in the EV suggests that the levels of PbpP in wild-type cells are not high enough to be detected by Bocillin-FL labeling.
All PBPs have an active-site serine that is acylated by β-lactams (36). We identified serine 301 (S301) as the likely active-site residue required for transpeptidation based on homology to other PBPs. To determine if S301 is the active-site serine, we mutated it to an alanine by site-directed mutagenesis and expressed pbpPS301A under the control of an IPTG-inducible promoter. In the strain producing PbpPS301A, the 66-kDa band was lost when imaging for Bocillin-FL (Fig. 2A and Fig. S3A). However, immunoblotting using anti-PbpP antisera detected a 66-kDa band corresponding to PbpPS301A, which is produced at levels similar to those of the WT protein (Fig. 2A; Fig. S3A). Thus, PbpPS301A cannot covalently bind Bocillin-FL. This suggests that PbpP is a penicillin-binding protein, S301 is required for binding β-lactams, and S301 is likely the active-site serine.
β-Lactam binding by PbpP is required for β-lactam-dependent activation of σP.
We sought to determine if β-lactam binding to PbpP was required for σP activation using a PbpPS301A active-site mutant. We complemented the ΔpbpP mutant with pbpP+ and pbpS301A under the control of their native promoter in a single copy by integrating constructs at the B. subtilis integrative conjugative element (ICEBs1) site in the B. thuringiensis chromosome (38). We found that PbpP+ restored PsigP-lacZ expression in the presence of cefoxitin (Fig. 2B). In contrast, when we complemented the strain with pbpPS301A, we observed no increase in PsigP-lacZ expression in the presence of cefoxitin (Fig. 2B). These data suggest that binding of PbpP to β-lactams is required for β-lactams to activate σP.
Overexpression of pbpP and pbpPS301A leads to activation of σP.
We noted that the basal level of PsigP-lacZ expression was higher in the strains complemented with pbpP+ and pbpS301A integrated at ICEBs1 than in WT B. thuringiensis (Fig. 2B). We reasoned that this might be due to higher basal levels of expression of pbpP and pbpPS301A at the ICEBs1 site. Thus, we sought to determine the effect of the overexpression of pbpP+ and pbpPS301A on σP activation. We expressed pbpP+ or pbpPS301A from a tetracycline-inducible promoter on a multicopy plasmid (34, 39). We observed that increased expression of pbpP+ or pbpPS301A leads to a dose-dependent increase in the expression of PsigP-lacZ, in the absence of β-lactams (Fig. 3A). We also found that the addition of cefoxitin led to a further increase in PsigP-lacZ expression when pbpP+ was overexpressed (Fig. S4B). We noted increased basal levels of PsigP-lacZ expression in the absence of anhydrotetracycline (ATc) and concluded that this is likely due to leaky expression of Ptet-pbpP and Ptet-pbpPS301A (Fig. 3A and Fig. S4B). These data suggest that the overexpression of both the WT and the active-site mutant (S301A) can activate σP even in the absence of β-lactams. We interpret this to mean that the requirement for β-lactam binding to PbpP can be compensated for by increased levels of PbpP; however, β-lactam binding to PbpP further enhances σP activation (Fig. S4B). The activation of σP in WT cells is likely not due to β-lactam-induced pbpP transcription as the expression of pbpP is not induced by β-lactams (Fig. S1B and C). The pbpPS301A mutant also fails to induce σP activation when expressed under the control of its native promoter, further suggesting that pbpP is not induced by β-lactams (Fig. 2B).
Since the loss of PbpP results in little to no degradation of RsiP in the presence of β-lactams, we tested if the increased expression of pbpP leads to the degradation of RsiP in the absence of β-lactams. We introduced Ptet-pbpP+ or Ptet-pbpPS301A into a strain containing IPTG-inducible gfp-rsiP. We found that the overexpression of PbpP and PbpPS301A leads to decreases in full-length GFP-RsiP levels, suggesting that PbpP can induce RsiP degradation and, thus, σP activation (Fig. 3B). This suggests that PbpP controls σP activation by controlling RsiP degradation.
PbpP is likely not the site 1 protease for RsiP.
The site 1 protease required for initiating RsiP degradation has not yet been identified. Since PbpP is required for site 1 cleavage of RsiP, the possibility exists that PbpP is the site 1 protease. We sought to determine if basal-level site 1 cleavage occurred in the absence of pbpP, which would suggest that another protein can cleave RsiP. Since site 2 cleavage is rapid (34), we expressed gfp-rsiP in a ΔpbpP ΔrasP double mutant, which should allow the buildup of any GFP-RsiP site 1 cleavage product. We observed the accumulation of a band corresponding to a GFP-RsiP fragment in the ΔrasP mutant in the absence of cefoxitin, and the intensity of this band increased in the presence of cefoxitin (Fig. 4A). We observed the same band in the ΔpbpP ΔrasP mutant; however, the band did not increase in the presence of cefoxitin. We concluded that in a ΔpbpP ΔrasP mutant, there is a basal level of site 1 cleavage of RsiP occurring in the presence and absence of cefoxitin (Fig. 4A). This suggests that site 1 cleavage can occur in the absence of PbpP, but it is not β-lactam inducible. Presumably, in this strain, the unidentified site 1 protease still retains its basal level of activity but cannot be further activated in the presence of cefoxitin due to the absence of PbpP.
To test if PbpP is sufficient for site 1 cleavage of RsiP, we introduced IPTG-inducible gfp-rsiP into the Bacillus subtilis chromosome (which does not encode a homolog of sigP or rsiP) and expressed pbpP using a xylose-inducible promoter. We grew the cells in the presence of 0.01 mM IPTG and increasing concentrations of xylose. We asked if PbpP was expressed and presumably properly folded by labeling with the fluorescent β-lactam Bocillin-FL. We observed a fluorescent band corresponding to PbpP that increased in intensity with increasing concentrations of xylose (Fig. 4B). We also monitored GFP-RsiP levels by performing immunoblot analysis using anti-GFP antisera. We did not observe degradation or a decrease in RsiP levels even at the highest levels of PbpP, indicating that PbpP is not sufficient for RsiP degradation in B. subtilis (Fig. 4C). Taken together, these data lead us to conclude that PbpP is not the site 1 protease but is required for sensing of β-lactams in B. thuringiensis.
Affinities of β-lactams for PbpP do not correlate with their ability to activate σP.
Since PbpP is likely not acting as the site 1 protease, we hypothesized that PbpP functions as a sensor that binds β-lactams and subsequentially activates σP. Therefore, we hypothesized that the reason why some β-lactams do not activate σP is that they have a lower affinity for PbpP. To test this hypothesis, we determined the affinity of PbpP for eight different β-lactams by modifying a Bocillin-FL inhibition experiment previously described by Kocaoglu and colleagues (40). We calculated the 50% inhibitory concentration (IC50) (the concentration of β-lactam at which 50% of Bocillin-FL labeling of PbpP is inhibited) to determine the binding affinity of different β-lactams. We found that while the β-lactams had different IC50s for PbpP, the differences did not correlate with the ability of the β-lactams to activate σP (Fig. 5A and B). For example, we found that some of the nonactivating β-lactams (cefoperazone and cefsulodin) had IC50s similar to those of activating β-lactams (Fig. 5A and B). Thus, the disparity in the β-lactams’ ability to activate σP is not simply due to the inability of PbpP to bind different β-lactams. These data also suggest that simple binding of any β-lactam to PbpP is not sufficient for σP activation.
Cefsulodin inhibits activation of σP by cefoxitin.
We found that β-lactam binding to PbpP is not sufficient for σP activation because nonactivating β-lactams covalently bind PbpP with affinities similar to those of the activating β-lactams (i.e., cefsulodin and ampicillin have nearly identical binding affinities for PbpP). We hypothesize that the β-lactams that activate σP induce a conformational change in PbpP that permits a protein-protein interaction. If this hypothesis were true, the β-lactams that do not activate σP would be able to inhibit the activation of σP by occupying the PbpP active site. To test this, we pretreated cells with cefsulodin (a nonactivator of σP) and then added cefoxitin (an activator of σP). We found that cefsulodin inhibited the activation of σP by cefoxitin in a dose-dependent manner (Fig. 6). We also show that pretreatment with cefmetazole (an activator of σP) does not inhibit activation (Fig. 6). Therefore, nonactivating β-lactams inhibit σP activation presumably by occupying the active site of PbpP and preventing activating β-lactams from binding PbpP and activating σP (Fig. 7).
Our data argue that PbpP is a sensor for β-lactams that is required for σP activation by indirectly promoting the degradation of RsiP (Fig. 7). This is supported by our observation that the loss of the penicillin-binding protein PbpP blocks σP activation and RsiP degradation. Our data indicate that the binding of a subset of β-lactams to PbpP results in σP activation. However, inhibition of PbpP transpeptidase activity by β-lactams is not the signal that activates σP since the transpeptidase mutant PbpPS301A does not activate σP. Interestingly, the overexpression of PbpP and PbpPS301A activates σP even in the absence of β-lactams; however, PbpP is not the site 1 protease. Together, these results argue that PbpP is a sensor of β-lactams and controls σP activation.
PbpP is required for σP activation.
The principal finding of this work is the demonstration that PbpP is required for the activation of σP in response to some β-lactams. Based on our findings, we propose the following working model for how PbpP functions as a sensor for β-lactams. In WT cells in the absence of stress, RsiP binds σP and inhibits σP activation (Fig. 7). When activating β-lactams are present, they bind the active-site serine of PbpP. The binding of the activating β-lactams results in a conformational change in PbpP that allows it to interact with a component of the σP system. This interaction initiates regulated intramembrane proteolysis of RsiP and, thus, σP activation (Fig. 7). This model is supported by ample evidence: (i) deletion of pbpP blocks RsiP degradation and σP activation, (ii) mutants of PbpP unable to bind β-lactams fail to activate σP in response to β-lactams, and (iii) overexpression of PbpP or PbpPS301A leads to constitutive RsiP degradation and σP activation. Thus, PbpP plays an essential role in sensing the presence of inducing β-lactams and controlling σP activation.
PbpP is not the site 1 protease.
It is possible that PbpP is a site 1 protease that initiates RsiP degradation; however, we think that it is unlikely. While PbpP is required for site 1 cleavage of RsiP in response to β-lactams, the totality of our data does not support PbpP as the site 1 protease. First, the overexpression of PbpP in B. subtilis does not induce the degradation of RsiP as it does in B. thuringiensis. Our data indicate that PbpP is functional, folded, and localized properly when expressed in B. subtilis since it can be labeled on whole cells by Bocillin-FL. This argues that PbpP is not sufficient for site 1 cleavage of RsiP and suggests that an unidentified B. thuringiensis protease is required. Second, in B. thuringiensis, we observed low-level site 1 cleavage of RsiP in the absence of PbpP. This argues that PbpP is not absolutely required for site 1 cleavage. If PbpP were a site 1 protease, there must be a second protease in B. thuringiensis that has low basal activity and cleaves RsiP at site 1 in the absence of PbpP. Finally, PbpP lacks any predicted protease domains. Future work will be required to identify the protease(s) required for site 1 cleavage of RsiP and, thus, σP activation.
PbpP is the β-lactam sensor for the σP system.
We hypothesize that PbpP is the sensor of β-lactams for the σP system. In support of this, we found that σP is not activated in the ΔpbpP mutant or when pbpPS301A is expressed from the native PpbpP promoter. However, the overproduction of either PbpP or PbpPS301A results in the activation of σP in the absence of β-lactams. This suggests that the overproduction of PbpP can compensate for β-lactam binding to PbpP to activate σP. Importantly, activation of σP is not due to inhibition of PbpP transpeptidase activity by β-lactams because PbpPS301A is catalytically inactive yet does not result in σP activation. This loss of σP activity is not due to an instability of PbpPS301A as it is produced at levels similar to those of WT PbpP. Activation of σP by β-lactams is not simply due to increased expression of pbpP since β-lactams do not induce pbpP expression. In addition, if increased expression of pbpP in response to β-lactams was responsible for σP activation, then we would have expected the pbpPS301A allele to induce σP activation when expressed under the control of the native PpbpP promoter. Taken together, these data suggest that PbpP interacts with some component of the signal transduction system.
In support of this hypothesis, we found that a subset of activating β-lactams bind PbpP with affinities similar to those of nonactivating β-lactams. We found that cefsulodin, a nonactivating β-lactam, can inhibit the activation of σP by an activating β-lactam, cefoxitin, presumably by competing for the active-site serine of PbpP. We hypothesize that nonactivating β-lactams do not induce the appropriate conformational change in PbpP to render it active and able to interact with its target. One obvious target for PbpP interaction is the anti-σ itself. However, we did not observe an interaction between the extracellular domains of RsiP76–275 and PbpP35–586in vitro using a copurification assay (see Fig. S8 in the supplemental material). This raises the possibility that PbpP interacts with another protein like the as-yet-unidentified site 1 protease. Alternatively, it may interact indirectly with RsiP or the site 1 protease via an unknown protein. Future work will need to determine what PbpP interactions drive RsiP degradation and, thus, σP activation.
Comparison of the BlaRI response to β-lactams to σP activation.
While the identification of a PBP required for the activation of an ECF σ factor is novel, there is precedence for a PBP transpeptidase-like domain functioning as a sensor of β-lactams. Found in diverse organisms, including Staphylococcus aureus and Bacillus licheniformis, BlaR1 (MecR1) contains an extracellular transpeptidase-like domain that senses β-lactams and a cytoplasmic protease domain. BlaR1 is a β-lactam sensor that directly binds β-lactams in its extracellular transpeptidase-like domain (41). The covalent bond formed with the β-lactam ring causes a conformational change in BlaR1 that activates the cytoplasmic protease domain (42). The protease domain cleaves the repressor of the β-lactamase operon, BlaI, thus activating the transcription of β-lactamase and increasing resistance to β-lactams (42). While the BlaIR system is clearly not synonymous with σP, it is worth noting that there is precedence for PBP domains that function as sensors of β-lactams.
MATERIALS AND METHODS
Media and growth conditions.
All B. thuringiensis strains are isogenic derivatives of AW43, a derivative of B. thuringiensis subsp. kurstaki strain HD73 (43). All strains and genotypes can be found in Table 2. All B. thuringiensis strains were grown in or on LB media at 30°C unless otherwise specified. Liquid cultures of B. thuringiensis were grown with agitation in a roller drum. B. thuringiensis strains containing episomal plasmids were grown in LB medium containing chloramphenicol (Cam) (10 μg/ml; Ameresco) or erythromycin (Erm) plus lincomycin (Linc) (MLS) (1 μg/ml Erm [Ameresco] and 25 μg/ml Linc [Research Products International]). E. coli strains were grown at 37°C using LB-ampicillin (Amp) (100 μg/ml; Ameresco) or LB-Cam (10 μg/ml) medium. B. subtilis strains were grown on LB medium with antibiotics (Cam at 10 μg/ml, spectinomycin [Spec] at 100 μg/ml [Amresco], or Erm at 10 μg/ml). To screen for threonine auxotrophy, B. thuringiensis strains were patched onto minimal medium plates without or with threonine (50 μg/ml). The β-galactosidase chromogenic indicator 5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside (X-Gal; Research Products International) was used at a concentration of 100 μg/ml. Anhydrotetracycline (ATc; Sigma) was used at a concentration of 100 ng/ml unless otherwise indicated. IPTG (Research Products International) and xylose (Acros) were used at the concentrations indicated in the figure legends. Additional β-lactams used in β-lactam-binding experiments were used at the concentrations indicated in the figure legends and were acquired from the following sources: cefsulodin, piperacillin, cefmetazole, and cefoxitin from Sigma-Aldrich; cephalothin from Chem-impex International Inc.; methicillin from Alfa Aesar; and cefoperazone from Toronto Research Chemical Inc.
TABLE 2 Strains
Reference or source
B. thuringiensis subsp. kurstaki HD73 cured of both pAW63 and pHT73; Nalr
endA1 rpsL thr leu thi lacY galK galT ara tomA tsx dam dcm supE44 Δ(lac-proAB) [F′ traD36 proAB lacIqZΔM15]
F−ompT hsdSB(rB− mB−) gal dcm (DE3) pRARE (Camr)
pCE593 in Rosetta DE3
Strain and plasmid construction.
All plasmids are listed in Table 3 and Table S1 in the supplemental material, which includes information relevant to plasmid assembly. Plasmids were constructed by isothermal assembly (44). Regions of plasmids constructed using PCR were verified by DNA sequencing. The oligonucleotide primers used in this work were synthesized by Integrated DNA Technologies (Coralville, IA) and are listed in Table S2. All plasmids were propagated using OmniMax 2-T1R as the cloning host and passaged through the nonmethylating E. coli strain INV110 before being transformed into a B. thuringiensis recipient strain.
To construct deletion mutants, we cloned 1 kb of DNA upstream and 1 kb downstream of the site of the desired deletion using primers listed in Table S2 into the temperature-sensitive pMAD plasmid (erythromycin resistant) between the BglII and EcoRI sites (45). Mutants were constructed by shifting temperatures as previously described (45).
B. subtilis ICEBs1 conjugation strains were constructed by transforming JAB932 as previously described (38). The resulting transformants or donor strains were grown in LB medium with d-alanine (100 μg/ml) for 2 h, at which point 1% xylose was added and cells were grown for 1 h. Recipient strains of B. thuringiensis were grown to an optical density at 600 nm (OD600) of ∼0.8. The donor and recipient strains were mixed at equal concentrations, plated on LB medium containing d-alanine (100 μg/ml), and incubated for 6 h. Transconjugants were isolated by plating on LB plates containing chloramphenicol.
B. thuringiensis DNA transformation.
Plasmids were introduced into B. thuringiensis by electroporation (46, 47). Briefly, recipient cells were grown to late log phase at 37°C from a fresh plate. For each transformation, cells (1.5 ml) were pelleted by centrifugation (8,000 rpm) and washed twice in room-temperature (RT) sterile water. After careful removal of all residual water, 100 μl of filter-sterilized 40% polyethylene glycol 6000 (PEG 6000; Sigma) was used to gently resuspend cells. Approximately 2 to 10 μl of unmethylated DNA (>50 ng/μl) was added to cells and transferred to a 0.4-cm-gap electroporation cuvette (Bio-Rad). Cells were exposed to 2.5 kV for 4 to 6 ms. LB medium was immediately added, and cells were incubated at 30°C for 1 to 2 h prior to plating on selective media.
To quantify expression from the sigP promoter, we measured the β-galactosidase activity of cells containing a PsigP-lacZ promoter fusion. Cultures grown overnight were diluted 1:50 in fresh LB medium and incubated to mid-log phase (OD of 0.8 to 1.5) at 30°C with or without ATc or IPTG. One milliliter of each subculture was pelleted (8,000 rpm), washed (in LB broth), and resuspended in 1 ml LB broth lacking or including specified antibiotics. After 1 h of incubation at 37°C, 1 ml of each sample was pelleted and resuspended in 1 ml of Z-buffer (0.06 M Na2HPO4, 0.04 M NaH2PO4*H2O, 0.01 M KCl, 0.001 M MgSO4). Cells were permeabilized by mixing with 16 μl of chloroform and 16 μl of 2% Sarkosyl (27, 48). Permeabilized cells (50 μl) were mixed with 100 μl of Z-buffer and 50 μl of 2 mg/ml chlorophenol red-β-d-galactopyranoside (CPRG; Research Products International) (50 μl), which is considerably more sensitive than X-Gal (49). The OD578 was measured over time using an Infinite M200 Pro plate reader (Tecan). β-Galactosidase activity units [(micromoles of chlorophenol red formed per minute) × 103/(OD600 × milliliters of cell suspension)] were calculated as previously described (50). Experiments were performed in technical and biological triplicate, with the means and standard deviations shown.
To determine the MICs for various antibiotics, we diluted cultures of bacteria grown overnight (washed in LB medium) 1:1,000 in medium containing 2-fold dilutions of each antibiotic. All MIC experiments were performed in round-bottom 96-well plates. Each experiment was performed in triplicate, and the cultures were allowed to incubate for 24 h at 37°C before observing growth or no growth by centrifuging the plates at 1,000 rpm for 5 minutes and observing the presence or absence of pellets.
Samples were electrophoresed on a 15% SDS-polyacrylamide gel, and proteins were then blotted onto a nitrocellulose membrane (GE Healthcare, Amersham). Nitrocellulose was blocked with 5% bovine serum albumin (BSA), and proteins were detected with a 1:10,000 dilution anti-GFP antisera. Streptavidin IR680LT (1:10,000) was used to detect two biotin-containing proteins, PycA (HD73_4231) and AccB (HD73_4487), which served as loading controls (51, 52). To detect primary antibodies, the blots were incubated with a 1:10,000 dilution of goat anti-rabbit IR800CW (Li-Cor) and imaged on an Odyssey CLx scanner (Li-Cor) or Azure Sapphire (Azure Biosystems). All immunoblots were performed at room temperature a minimum of three times, with a representative example shown.
Bocillin-FL labeling assay.
Cultures grown overnight at 30°C were diluted 1:50 and grown to an OD of ∼1.0. The cultures were aliquoted in 1-ml aliquots and pelleted at 8,000 rpm. The cells were washed twice in 500 μl of 1× phosphate-buffered saline (PBS) and resuspended in either 50 μl of 50 μg/ml Bocillin-FL (Thermo Fisher) or 50 μl of 10-fold dilutions of β-lactams (0.0005 to 5,000 μg/ml). The samples resuspended in β-lactams were incubated for 30 min at room temperature and then pelleted and resuspended in 50 μl or 50 μg/ml Boc-FL for 15 min. After incubation in Boc-FL, all the samples were pelleted and resuspended in 200 μl sample buffer with 5% β-mercaptoethanol (βME). The samples were sonicated, heated, and electrophoresed on a 12% polyacrylamide gel. The gels were imaged on an Azure Sapphire system (AzureBiosystems) by excitation at 488 nm and detection at 518 nm. The Bocillin-FL labeling experiment was performed in biological triplicate for each antibiotic, and the Bocillin-FL intensity for the PbpP band was quantified on each gel. The average intensity was used to calculate the IC50 using GraphPad Prism, with means and standard errors or deviations shown.
This work was supported by National Institute of Allergy and Infectious Diseases, NIH, grant R21AI146769.
We thank Christopher A. Voigt for providing the B. subtilis ICEBs1 conjugation strains, plasmids, and protocols. We also thank members of the Ellermeier and David S. Weiss laboratories for helpful comments.
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