Real-time monitoring of peptidoglycan synthesis by membrane-reconstituted penicillin-binding proteins

Víctor M Hernández-Rocamora; Natalia Baranova; Katharina Peters; Eefjan Breukink; Martin Loose; Waldemar Vollmer

doi:10.7554/eLife.61525

. 2021 Feb 24;10:e61525. doi: 10.7554/eLife.61525

Real-time monitoring of peptidoglycan synthesis by membrane-reconstituted penicillin-binding proteins

Víctor M Hernández-Rocamora ^1,^†, Natalia Baranova ^2,^†,^‡, Katharina Peters ^1,^§, Eefjan Breukink ³, Martin Loose ^2,^✉, Waldemar Vollmer ^1,^✉

Editors: Jie Xiao⁴, Olga Boudker⁵

PMCID: PMC7943195 PMID: 33625355

Abstract

Peptidoglycan is an essential component of the bacterial cell envelope that surrounds the cytoplasmic membrane to protect the cell from osmotic lysis. Important antibiotics such as β-lactams and glycopeptides target peptidoglycan biosynthesis. Class A penicillin-binding proteins (PBPs) are bifunctional membrane-bound peptidoglycan synthases that polymerize glycan chains and connect adjacent stem peptides by transpeptidation. How these enzymes work in their physiological membrane environment is poorly understood. Here, we developed a novel Förster resonance energy transfer-based assay to follow in real time both reactions of class A PBPs reconstituted in liposomes or supported lipid bilayers and applied this assay with PBP1B homologues from Escherichia coli, Pseudomonas aeruginosa, and Acinetobacter baumannii in the presence or absence of their cognate lipoprotein activator. Our assay will allow unravelling the mechanisms of peptidoglycan synthesis in a lipid-bilayer environment and can be further developed to be used for high-throughput screening for new antimicrobials.

Research organism: E. coli

Introduction

Peptidoglycan (PG) is a major cell wall polymer in bacteria. It is composed of glycan strands of alternating N-actetylglucosamine (GlcNAc) and N-acetylmuramic acid (MurNAc) residues interconnected by short peptides. PG forms a continuous, mesh-like layer around the cell membrane to protect the cell from bursting due to the turgor and maintain cell shape (Vollmer et al., 2008). The essentiality and conservation of PG in bacteria make PG metabolism an ideal target of antibiotics.

Class A penicillin-binding proteins (PBPs) are bifunctional PG synthases which use the precursor lipid II to polymerize glycan chains (glycosyltransferase [GTase] reactions) and crosslink peptides from adjacent chains by DD-transpeptidation (Goffin and Ghuysen, 1998). Moenomycin inhibits the GTase and β-lactams the transpeptidase function of class A PBPs (Sauvage and Terrak, 2016; Macheboeuf et al., 2006). In Escherichia coli, PBP1A and PBP1B account for a substantial proportion of the total cellular PG synthesis activity (Cho et al., 2016) and are tightly regulated by interactions with multiple proteins (Egan et al., 2015; Typas et al., 2012; Egan et al., 2020; Egan et al., 2017), including the outer membrane-anchored activators LpoA and LpoB (Egan et al., 2018; Typas et al., 2010; Jean et al., 2014).

Historically, in vitro PG synthesis assays have been crucial to decipher the biochemical reactions involved in PG synthesis and determine the mode of action of antibiotics (Izaki et al., 1968). However, these studies were limited by the scarcity of lipid II substrate and the inability to purify a sufficient quantity of active enzymes. Lipid II can now be synthesized chemically (VanNieuwenhze et al., 2002; Schwartz et al., 2001; Ye et al., 2001) or semi-enzymatically (Breukink et al., 2003; Egan et al., 2015), or isolated from cells with inactivated MurJ (Qiao et al., 2017). Radioactive or fluorescent versions of lipid II are also available to study PG synthesis in a test tube. However, there are several drawbacks with currently available PG synthesis assays. First, most assays are end-point assays that rely on discrete sampling and therefore do not provide real-time information about the enzymatic reaction. Second, some assays involve measuring the consumption of lipid II or analysing the reaction products by SDS-PAGE (Egan et al., 2015; Barrett et al., 2007; Qiao et al., 2014; Sjodt et al., 2018) or high-pressure liquid chromatography (HPLC) after digestion with a muramidase (Bertsche et al., 2005; Born et al., 2006). These laborious techniques make assays incompatible with high-throughput screening and hinder the determination of kinetic parameters. A simple, real-time assay with dansyl-labelled lipid II substrate overcomes these problems but is limited to assay GTase reactions (Schwartz et al., 2001; Offant et al., 2010; Egan et al., 2015).

Recently, two types of real-time TPase assays have been described. The first uses non-natural mimics of TPase substrates such as the rotor-fluorogenic 470 D-lysine probe Rf470DL, which increases its fluorescence emission upon incorporation into PG (Hsu et al., 2019). The second assay monitors the release of D-Ala during transpeptidation in coupled enzymatic reactions with D-amino acid oxidase, peroxidases, and chromogenic or fluorogenic compounds (Frére et al., 1976; Gutheil et al., 2000; Catherwood et al., 2020). Coupled assays are often limited in the choice of the reaction conditions, which in this case must be compatible with D-amino acid oxidase activity. Hence, each of the current assays has its limitations and most assays exclusively report on either the GTase or TPase activity, but not both activities at the same time.

Another major drawback of many of the current assays is that they include detergents and/or high concentration (up to 30%) of the organic solvent dimethyl sulfoxide (DMSO) to maintain the PG synthases in solution (Offant et al., 2010; Biboy et al., 2013; Huang et al., 2013; Lebar et al., 2013; Qiao et al., 2014; Egan et al., 2015; Catherwood et al., 2020). However, both detergents and DMSO have been shown to affect the activity and interactions of E. coli PBP1B (Egan and Vollmer, 2016). Importantly, a freely diffusing, detergent-solubilized membrane enzyme has a very different environment compared to the situation in the cell membrane where it contacts phospholipids and is confined in two dimensions (Gavutis et al., 2006; Zhdanov and Höök, 2015). Here, we sought to overcome the main limitations of current PG synthesis assays and establish a system with more physiological experimental conditions. We used sensitive Förster resonance energy transfer (FRET) detection for simultaneous monitoring of GTase and TPase reactions. The real-time assay reports on PG synthesis in phospholipid vesicles or planar lipid bilayers. We successfully applied this assay to several class A PBPs from pathogenic Gram-negative bacteria, demonstrating its robustness and potential use in screening assays to identify PBP inhibitors.

Results

Real-time assay for detergent-solubilized E. coli PBP1B

To develop a FRET-based real-time assay for PG synthesis using fluorescently labelled lipid II, we prepared lysine-type lipid II versions with high quantum yield probes, Atto550 (as FRET donor) and Atto647n (as FRET acceptor), linked to position 3 (Figure 1—figure supplement 1A, B; Mohammadi et al., 2014; Egan et al., 2015). For assay development, we used E. coli PBP1B (PBP1B^Ec) (Egan et al., 2015; Bertsche et al., 2005; Biboy et al., 2013) solubilized with Triton X-100 and a lipid-free version of its cognate outer membrane-anchored lipoprotein activator LpoB (Typas et al., 2010; Egan et al., 2014; Egan et al., 2018; Lupoli et al., 2014; Catherwood et al., 2020).

PBP1B^Ec can utilize fluorescently labelled lipid II to polymerize long glycan chains only when unlabelled lipid II is also present in the reaction (Van't Veer et al., 2016). We therefore included unlabelled meso-diaminopimelic acid (mDAP)-type lipid II into reactions of PBP1B^Ec with lipid II-Atto550 and lipid II-Atto647n (Figure 1A). Both probes were incorporated into the produced PG or glycan chains as indicated by SDS-PAGE analysis (Figure 1B, I). After the reaction, fluorescence spectra taken at the excitation wavelength of the donor fluorophore (Atto550, λ_abs=552 nm) showed a reduced donor emission intensity (λ_fl=580 nm) and an increased emission of the acceptor fluorophore (Atto647n, λ_fl=665 nm) (Figure 1C, I) indicative of FRET between the two fluorophores. Analysis of the fluorescence spectra allowed to calculate FRET efficiencies which we found to be 29 ± 6% (Figure 1D, Figure 1—figure supplement 2). Ampicillin, which inhibits the TPase, blocked the formation of crosslinked PG (Figure 1B, II) and reduced the FRET efficiency to one third (Figure 1C, II, D). Moenomycin, which blocks the GTase, and, indirectly, TPase activities completely abolished the incorporation of fluorescent lipid II and the associated signal (Figure 1B, III, D; Bertsche et al., 2005). These results demonstrate that incorporation of the labelled probes into PG by PBP1B^Ec results in fluorescence energy transfer that depends on the GTase and TPase activity, with the latter being the major contributor.

Figure 1. — (A) Scheme of the reactions of a class A penicillin-binding protein (PBP) (GTase-TPase) with unlabelled lipid II and the two versions of labelled lipid II, yielding a peptidoglycan (PG) product that shows FRET. (B) SDS-PAGE analysis of PG products by PBP1B^Ec (0.5 µM) reactions with unlabelled lipid II, Atto550-labelled lipid II, and Atto647n-labelled lipid II at a 1:1:1 molar ratio (each 5 µM), in the absence of antibiotics (I, red) or in the presence of 1 mM ampicillin (II, blue) or 50 µM moenomycin (III, yellow). Samples were incubated for 1 hr at 37°C and boiled for 5 min. (C) Representative fluorescence emission spectra taken after reactions performed as described in B and following the same labelling pattern. (D) FRET efficiency for PBP1B^Ec reactions carried out as indicated in B, calculated using the (ratio)_A method (see Materials and methods). Values are mean ± SD of at least three independent samples. (E) Representative reaction curves from FRET assays of detergent-solubilized PBP1B^Ec. The same components as indicated in B were incubated in the presence or absence of 2 µM LpoB(sol). Reactions were performed in the absence of antibiotic (left), with 1 mM ampicillin (Amp) or 50 µM moenomycin (Moe) (middle), or by omitting unlabelled lipid II (right). The numbers indicate the corresponding lane of the gel in Figure 1—figure supplement 2D. Samples were incubated for 1 hr at 25°C. (F) Averaged initial slopes from reaction curves obtained by the FRET assay for detergent-solubilized *E. coli* PBP1B in the presence (blue) or absence (red) of LpoB, and in the presence or absence of ampicillin. Values are normalized relative to the slope in the absence of activator for each condition and are mean ± SD of 2–3 independent experiments.

Figure 1—source data 1. Numerical data to support graphs in Figure 1 and original gel images for Figure 1B.

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Figure 1—figure supplement 1. — (A) Scheme of the reactions of a class A penicillin-binding protein (PBP) (GTase-TPase) with unlabelled lipid II and the two versions of labelled lipid II, yielding a peptidoglycan (PG) product that shows FRET. (B) SDS-PAGE analysis of PG products by PBP1B^Ec (0.5 µM) reactions with unlabelled lipid II, Atto550-labelled lipid II, and Atto647n-labelled lipid II at a 1:1:1 molar ratio (each 5 µM), in the absence of antibiotics (I, red) or in the presence of 1 mM ampicillin (II, blue) or 50 µM moenomycin (III, yellow). Samples were incubated for 1 hr at 37°C and boiled for 5 min. (C) Representative fluorescence emission spectra taken after reactions performed as described in B and following the same labelling pattern. (D) FRET efficiency for PBP1B^Ec reactions carried out as indicated in B, calculated using the (ratio)_A method (see Materials and methods). Values are mean ± SD of at least three independent samples. (E) Representative reaction curves from FRET assays of detergent-solubilized PBP1B^Ec. The same components as indicated in B were incubated in the presence or absence of 2 µM LpoB(sol). Reactions were performed in the absence of antibiotic (left), with 1 mM ampicillin (Amp) or 50 µM moenomycin (Moe) (middle), or by omitting unlabelled lipid II (right). The numbers indicate the corresponding lane of the gel in Figure 1—figure supplement 2D. Samples were incubated for 1 hr at 25°C. (F) Averaged initial slopes from reaction curves obtained by the FRET assay for detergent-solubilized *E. coli* PBP1B in the presence (blue) or absence (red) of LpoB, and in the presence or absence of ampicillin. Values are normalized relative to the slope in the absence of activator for each condition and are mean ± SD of 2–3 independent experiments.

Figure 1—source data 1. Numerical data to support graphs in Figure 1 and original gel images for Figure 1B.

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Next, we monitored reactions in real time by measuring fluorescence emission of the donor and acceptor fluorophores (FI_donor and FI_acceptor, respectively) after excitation of the donor (540 nm) in a microplate reader for 60 min (Figure 1E, Figure 1—figure supplement 3A). As controls, we also performed reactions containing unlabelled lipid II plus only one of the labelled lipid II versions (lipid II-Atto550 or lipid II-Atto647n) in parallel (Figure 1—figure supplement 4). Changes in FI_donor and FI_acceptor were much higher when both fluorescent lipid II versions were present, in agreement with energy transfer. Thus, we used the ratio between both signals (FI_acceptor/FI_donor) as a real-time readout for FRET and PG synthesis. Without LpoB, FRET appeared after ~5 min and slowly increased until it plateaued after 50–60 min (Figure 1E, left). By contrast, reactions with LpoB(sol) showed an immediate and rapid increase in FRET which reached the plateau after 10–20 min, consistent with faster PG synthesis (Figure 1E, left). In agreement with the end-point analysis described above, we found no FRET in samples containing moenomycin (Figure 1E, middle), and ampicillin generally reduced the final fluorescence ratio level by approximately threefold (Figure 1E, middle). Analysis of reaction products by SDS-PAGE also confirmed that crosslinked PG was only produced in the absence of antibiotics, while the presence of ampicillin still allowed the formation of glycan chains (Figure 1—figure supplement 3B).

The GTase reaction began after a lag phase, consistent with previously published data (Schwartz et al., 2002; Egan et al., 2014), which is likely caused by a slower initiation of glycan chain synthesis compared to the rate of polymerization. We measured the slope of FRET reaction curves during the linear raise in signal after the lag phase (when present) and compared the slopes with or without activator. Slopes with LpoB were approximately ten- or twentyfold higher than without the activator, in the absence or presence of ampicillin, respectively (Figure 1F). Although this result is comparable to the approximately tenfold activation of the GTase rate by LpoB measured with dansyl-lipid II (Egan et al., 2014; Egan et al., 2018), a quantification of the individual GTase and TPase reaction rates would require a more exact knowledge of how these two activities contribute to the final FRET signal, which is currently not available (see Discussion).

Intra-chain versus inter-chain FRET

Because ampicillin substantially reduced the FRET signal, we hypothesized that FRET arises mainly between fluorophores on different glycan chains of a crosslinked PG product (Figure 1A). To determine the relative contribution of intra-chain versus inter-chain FRET, we digested PG produced in the presence of labelled lipid II with either the DD-endopeptidase MepM, which cleaves crosslinks between glycan chains (Singh et al., 2015; Singh et al., 2012), or the muramidase cellosyl, which cleaves the β-(1,4)-glycosidic bond between MurNAc and GlcNAc-producing muropeptides (structures 1–3 in Figure 2C; Rau et al., 2001; Figure 2A, B). As a control, glycan chains produced by PBP1B^Ec in the presence of ampicillin were also digested with both hydrolases. SDS-PAGE analysis confirmed that MepM substantially reduced the amount of crosslinked PG in the samples while cellosyl digested the PG into muropeptides (Figure 2A). Next, we measured the FRET efficiency after digestion. MepM digestion had a negligible effect on the FRET efficiency of glycan chains produced in the presence of ampicillin but reduced the FRET efficiency by approximately twofold for crosslinked-PG samples (Figure 2B). This confirms that inter-chain FRET is a major contributor to the final FRET signal. MepM did not reduce FRET efficiency to the same value as ampicillin, presumably because of incomplete digestion of the labelled PG. Finally, cellosyl completely abolished FRET for both glycan chains and crosslinked PG (Figure 2B).

Figure 2. — (A) Peptidoglycan (PG) synthesized in reactions of PBP1B^Ec in the presence or absence of 1 mM ampicillin was incubated with no PG hydrolase (U), DD-endopeptidase MepM (M), or muramidase cellosyl (C), and aliquots were analysed by SDS-PAGE. Reaction conditions were the same as indicated in Figure 1B–D. (B) FRET efficiency for samples prepared as indicated in A, calculated using the (ratio)_A method (see Materials and methods). Values are mean ± SD of at least three independent experiments. (C) PBP1B^Ec (0.5 µM) was incubated with 5 µM each of lipid II-Atto647n, lipid II-Atto550, and ¹⁴C-labelled lipid II. At indicated time points, aliquots were taken and reactions were stopped by addition of moenomycin. After measuring fluorescence (see D), the PG was digested with the muramidase cellosyl, and the resulting muropeptides were reduced with sodium borohydride and separated by HPLC. The structures of muropeptides corresponding to peaks 1–3 are shown below the chromatograms. (D) Fluorescence spectra taken with excitation at 522 nm for the samples described in C. (E) Quantification of peak 2 (GTase product, blue), peak 3 (GTase+TPase, black), or the sum of both 2 and 3 (yellow) from chromatograms in C, along with the FRET signal (red) calculated as the ratio of acceptor emission over donor emission from data in D.

Figure 2—source data 1. Numerical data to support graphs in Figure 2 and original gel images for Figure 2B.

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Figure 2—figure supplement 1. — (A) Peptidoglycan (PG) synthesized in reactions of PBP1B^Ec in the presence or absence of 1 mM ampicillin was incubated with no PG hydrolase (U), DD-endopeptidase MepM (M), or muramidase cellosyl (C), and aliquots were analysed by SDS-PAGE. Reaction conditions were the same as indicated in Figure 1B–D. (B) FRET efficiency for samples prepared as indicated in A, calculated using the (ratio)_A method (see Materials and methods). Values are mean ± SD of at least three independent experiments. (C) PBP1B^Ec (0.5 µM) was incubated with 5 µM each of lipid II-Atto647n, lipid II-Atto550, and ¹⁴C-labelled lipid II. At indicated time points, aliquots were taken and reactions were stopped by addition of moenomycin. After measuring fluorescence (see D), the PG was digested with the muramidase cellosyl, and the resulting muropeptides were reduced with sodium borohydride and separated by HPLC. The structures of muropeptides corresponding to peaks 1–3 are shown below the chromatograms. (D) Fluorescence spectra taken with excitation at 522 nm for the samples described in C. (E) Quantification of peak 2 (GTase product, blue), peak 3 (GTase+TPase, black), or the sum of both 2 and 3 (yellow) from chromatograms in C, along with the FRET signal (red) calculated as the ratio of acceptor emission over donor emission from data in D.

Figure 2—source data 1. Numerical data to support graphs in Figure 2 and original gel images for Figure 2B.

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To confirm that the formation of peptide crosslinks is required to produce substantial FRET in the absence of LpoB, we analysed the PG synthesized by PBP1B^Ec from radioactively labelled mDAP-type lipid II and the two fluorescent lipid II analogues (Figure 2C–E). We monitored the reaction at different time points by fluorescence spectroscopy (FRET measurements) and digested aliquots with cellosyl before separating the resulting muropeptides by HPLC. The monomers and crosslinked muropeptide dimers were quantified by scintillation counting using an in-line radiation detector attached to the HPLC column (Figure 2C). FRET increased over time and correlated well with the formation of crosslinked muropeptide dimers, but not the rate of lipid II consumption (peak 2) (Figure 2D, E). Overall, we conclude that, in the absence of LpoB, FRET can arise from GTase activity alone (intra-chain FRET), but the overall contribution from the TPase activity (inter-chain FRET) is dominant.

To study in more detail the contribution of intra-chain FRET, we varied the molar fraction of fluorescent lipid II and measured the activity of PBP1B^Ec in the presence or absence of activator. Confirming a previous study (Van't Veer et al., 2016), PBP1B^Ec alone was unable to use lipid II-Atto550 and lipid II-Atto647n for polymerization when unlabelled lipid II was not present (Figure 1E, right). Surprisingly, addition of LpoB allowed PBP1B^Ec to produce short, non-crosslinked individual PG chains (Figure 1E, Figure 1—figure supplement 3B) that gave rise to a slow but large increase in FRET (Figure 1E, right, Figure 1—figure supplement 3A), indicating that polymerization of labelled lipid II occurred in the absence of unlabelled lipid II. To investigate this effect further, we varied the proportion of fluorescent lipid II over non-fluorescent (but radioactive) lipid II and measured the reaction slopes and, at the end of the reaction time, the final FI_acceptor/FI_donor ratio and amounts of unused lipid II versions, in the presence or absence of activator (Figure 2—figure supplement 1). LpoB slightly increased consumption of all versions of lipid II by PBP1B^Ec (Figure 2—figure supplement 1A) but did not affect the proportion of fluorescent material that was incorporated into PG, which reflected the initial percentage of fluorescent lipid II (Figure 2—figure supplement 1B). In the absence of LpoB, the final FI_acceptor/FI_donor ratio increased with increasing proportions of labelled lipid II, but this increase was steeper in the presence of LpoB (Figure 2—figure supplement 1C). As similar proportions of fluorescent material were incorporated into PG with or without activator, the difference in the final FRET must arise by fluorophores located closer together within the PG produced when LpoB is present. Finally, the reaction slopes did not change significantly with increasing proportions of labelled lipid II in the absence of LpoB but increased in its presence up to 50% of fluorescent lipid II, and then plateaued (Figure 2—figure supplement 1A). Overall, these results suggest that LpoB stimulates the incorporation of fluorophores in consecutive positions along the glycan chain and thus increases the contribution of intra-chain FRET. Thus, the increase in slopes observed with activator (Figure 1F) reflects not only a higher PG synthesis rate but also a higher contribution of intra-chain FRET.

FRET assay to monitor PG synthesis in liposomes

To establish the FRET assay for membrane-embedded PG synthases, we reconstituted a version of PBP1B^Ec with a single cysteine at the cytoplasmic N-terminus into liposomes prepared from E. coli polar lipids (EcPL) (Figure 3—figure supplement 1A). The liposome-reconstituted PBP1B^Ec became accessible to a sulfhydryl-reactive fluorescent probe only after disrupting the liposomes with detergent (Figure 3A), showing that virtually all PBP1B molecules were oriented with the N-terminus inside the liposomes. This suggests that the large, extracellular portion of PBP1B^Ec is not transferred through the membrane during the reconstitution into liposomes (Rigaud and Lévy, 2003). Next, we reconstituted unmodified PBP1B^Ec and tested its activity by adding radioactive lipid II. In contrast to the detergent-solubilized enzyme, the liposome-reconstituted PBP1B^Ec required the absence of NaCl from the reaction buffer for improved activity (Figure 3—figure supplement 1B–E), suggesting that ionic strength affects either the structure of PBP1B^Ec in the membrane, the properties of EcPL liposomes, or the delivery of lipid II into the liposomes.

Figure 3—figure supplement 1. — (A) Class A penicillin-binding proteins (PBPs) were reconstituted in *E. coli* polar lipid (EcPL) liposomes. To assess the orientation of the liposome-reconstituted PBPs, MGC-⁶⁴PBP1B-his C777S C795S containing a single cysteine in the N-terminal region was reconstituted as in A. The accessibility of the cysteine was determined by staining with sulfhydryl-reactive fluorescent probe, Alexa Fluor555-maleimide, in the presence or absence of Triton X-100 (TX). Samples were analysed by SDS-PAGE with fluorescence scanning to detect labelled protein followed by Coomassie staining. (B) To perform activity assays in liposomes, class A PBPs were reconstituted along a 1:1 molar ratio mixture of Atto550-labelled lipid II and Atto647n-labelled lipid II in liposomes as in A. Reactions were started by addition of unlabelled lipid II in the presence or absence of lipoprotein activators (lpo). Using this methodology, we monitored the activity of PBP1B^Ec (**C, D**), PBP1B^Ab (**E, F**), and PBP1B^Pa (**G, H**). Representative reaction curves are shown. Reactions were carried out in the presence (blue lines) or absence (red lines) of the lipoprotein activators (LpoB(sol) for PBP1B^Ec, LpoP^Ab(sol) for PBP1B^Ab, and LpoP^Pa(sol) for PBP1B^Pa), and either in the absence of antibiotic (left) or presence of 1 mM ampicillin (Amp) or 50 µM moenomycin (Moe, black and yellow lines) (middle). For PBP1B^Ec, control reactions in the absence of unlabelled lipid II (right) are also shown. Products were analysed by SDS-PAGE followed by fluorescence scanning at the end of reactions (right side). Curves are numbered according to the corresponding lane on the SDS-PAGE gels. PBP1B^Ec, PBP1B^Ab, and PBP1B^Pa were reconstituted in EcPL liposomes containing labelled lipid II (0.5 mol% of lipids, 1:1 molar ratio mixture of Atto550-labelled lipid II and Atto647n-labelled lipid II), at protein-to-lipid molar ratios of 1:3000, 1:2000, and 1:3000, respectively. Reactions were started by adding unlabelled lipid II (final concentration 12 µM) and incubated at 37°C for 60 min (PBP1B^Ec and PBP1B^Ab) or 90 min (PBP1B^Pa) while monitoring fluorescence at 590 and 680 nm with excitation at 522 nm. (D), (F), and (H) show averaged initial slopes from reaction curves obtained by the FRET assay for liposome-reconstituted PBP1B^Ec, PBP1B^Ab, and PBP1B^Pa, respectively, in the presence (blue) or absence (red) of lipoprotein activators and in the presence or absence of ampicillin. Values are normalized relative to the slope in the absence of activator and are mean ± variation of two independent experiments.

Figure 3—source data 1. Numerical data to support graphs in Figure 3 and original gel images for Figure 3C, E and G.

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We next aimed to adapt the FRET assay to study PG synthesis on liposomes to mimic the situation in the cell (Figure 3, Figure 3—figure supplement 2). As PBP1B^Ec did not accept Atto550- or Atto647-derivatized lipid II for GTase reactions in the absence of unlabelled lipid II (Figure 1E), we reconstituted PBP1B^Ec in liposomes along both Atto-labelled substrates and initiated the reaction by adding unlabelled lipid II (Figure 3B). PBP1B^Ec reaction rates in liposomes were slower than in the presence of Triton X-100 for all conditions tested (compare curves in Figure 3C, measured at 37°C, with the ones in Figure 1E, measured at 25°C), and there was a longer lag time before FRET started to increase (Figure 3C, left). Moenomycin blocked the increase in FRET, while ampicillin reduced the final FRET levels (Figure 3C, middle). For unknown reasons, the FRET signal with moenomycin was initially higher than without moenomycin and then decreased to initial values without moenomycin (Figure 3C, middle), independent of the class A PBP used (see below) but not in empty liposomes (Figure 3—figure supplement 3). LpoB(sol) produced an approximately tenfold increase in the initial slope, measured as explained above (Figure 3D), and the resulting final FRET was much higher (Figure 3C, left). In some experiments with PBP1B^Ec liposomes in the presence of ampicillin and LpoB(sol), we noticed a slow decrease in FRET after a fast initial increase, and the production of short glycan chains instead of the long chains produced normally (Figure 3—figure supplement 2A). As in detergents, without unlabelled lipid II membrane-bound PBP1B produced a FRET signal only in the presence of LpoB(sol) (Figure 3C, right). The analysis of the final products by SDS-PAGE confirmed that both Atto550 and Atto647n were incorporated into glycan chains or crosslinked PG during the reaction in liposomes (Figure 3C, right, Figure 3—figure supplement 2B). As expected, controls with PBP1B^Ec liposomes reconstituted with only lipid II-Atto550 or only lipid II-Atto647n showed significantly lower changes in FI_donor and FI_acceptor than when both fluorescent versions were present together (Figure 3—figure supplement 4).

In summary, using our FRET-based assay we demonstrated real-time monitoring PG synthesis in membrane by PBP1B^Ec and showed that the FRET signal was sensitive to the presence of PG synthesis inhibitors (moenomycin and ampicillin).

Activities of other membrane-bound class A PBPs

To demonstrate the usefulness of the FRET assay to study class A PBPs of potential therapeutic interest, we next tested two PBP1B homologues from Gram-negative pathogens, Acinetobacter baumannii (PBP1B^Ab) and Pseudomonas aeruginosa (PBP1B^Pa). We set up reactions in the presence or absence of a soluble version of the lipoprotein activator LpoP^Pa(sol) for PBP1B^Pa (Greene et al., 2018). There is currently no reported activator of PBP1B^Ab, but next to the gene encoding PBP1B^Ab we identified a hypothetical gene encoding a lipoprotein containing two tetratricopeptide repeats (Uniprot code D0C5L6) (Figure 3—figure supplement 5) which we subsequently found to activate PBP1B^Ab (see below, Figure 3—figure supplement 6). We named this protein LpoP^Ab and purified a version without its lipid anchor, called LpoP^Ab(sol). We were able to monitor PG synthesis activity by FRET for both PBPs in the presence or absence of their (hypothetical) activators using the Triton X-100-solubilized (Figure 3—figure supplements 7 and 8) or liposome-reconstituted proteins (Figure 3E–H, Figure 3—figure supplement 2D–E). Our experiments revealed the differences in the activities and the effect of activators between both PBP1B-homolgoues which we discuss in the following paragraphs.

PBP1B^Ab showed GTase activity in the presence of Triton X-100 (Figure 3—figure supplement 6A) and was stimulated ~3.3-fold by LpoP^Ab(sol) (Figure 3—figure supplement 6B); LpoP^Ab(sol) also accelerated the consumption of lipid II-Atto550 and glycan chain polymerization (Figure 3—figure supplement 6C). We measured a low FRET signal for PG produced by the detergent-solubilized enzyme in the FRET assay (Figure 3—figure supplement 7A) and poor production of crosslinked PG (Figure 3—figure supplement 7C), unlike in the case of the other PBPs. However, the liposome-reconstituted PBP1B^Ab displayed a higher TPase activity than the detergent-solubilized enzyme (compare gels in Figure 3E, right, and Figure 3—figure supplement 7C). In addition, the final FRET signal was substantially higher in liposomes than in detergents (Figure 3E, Figure 3—figure supplement 7A). Moenomycin completely blocked FRET development, while ampicillin had a negligible effect on the final FRET levels in detergents and only a small effect in liposomes (~1.2-fold reduction), indicating that intra-chain FRET is the major contributor to FRET (Figure 3E, Figure 3—figure supplement 7A). LpoP^Ab(sol) stimulated PBP1B^Ab, with a higher effect in detergents (12.3-fold increase) than liposomes (~2.5-fold increase) (Figure 3E, F, Figure 3—figure supplement 7A, B).

PBP1B^Pa displayed robust TPase activity in detergents and liposomes (Figure 3G, right, Figure 3—figure supplement 8C), and ampicillin reduced the final FRET signal by ~1.8-fold in Triton X-100 and ~1.5-fold in liposomes, indicating a substantial contribution of inter-chain FRET to the FRET signal (Figure 3G, Figure 3—figure supplement 8A). The addition of LpoP^Pa(sol) resulted in an increase in the final FRET by ~2.2-fold in the membrane and ~2.1-fold in detergents (Figure 3G, Figure 3—figure supplement 8A), and accelerated initial slopes by ~4.2-fold in the membrane and ~11.5-fold in detergents (Figure 3H, Figure 3—figure supplement 8B); lipid II consumption was increased under both conditions (Figure 3G, right, Figure 3—figure supplement 8C). Overall, these results indicate that LpoP^Pa(sol) stimulates both GTase and TPase activities in agreement with a recent report (Caveney et al., 2020).

PG synthesis on supported lipid bilayers

As we were able to successfully reconstitute active class A PBPs in membranes and monitor their activity in real time, we next aimed to characterize the behaviour of these enzymes in the membrane in more detail by reconstituting them on supported lipid bilayers (SLBs). SLBs are phospholipid bilayers formed on top of a solid support, usually a glass surface, and they allow for studying the spatial organization of transmembrane proteins and their diffusion along the membrane by fluorescence microscopy at high spatiotemporal resolution.

We optimized the reconstitution of PBP1B^Ec in SLBs formed with EcPL and used the optimized buffer conditions for activity assays on liposomes. To support lateral diffusion and also improve stability of the proteins incorporated into SLBs, we employed glass surfaces coated with polyethylene glycol (PEG) end-functionalized with a short fatty acid (Roder et al., 2011) to anchor the EcPL bilayer (Figure 4A). We noticed a decrease in membrane diffusivity and homogeneity at a high surface density of PBP1B^Ec (Figure 4—figure supplement 1). To maintain the integrity of the SLB, we reduced the density of PBP1B^Ec on SLBs from ~10⁻³ mol protein/mol lipid in liposomes to a range of 10⁻⁶ to 10⁻⁵ mol protein/mol lipid. Using a fluorescently labelled version of PBP1B^Ec reconstituted in SLBs, we were able to track the diffusion of single PBP1B molecules in the plane of lipid membrane in the presence or absence of substrate lipid II by total internal reflection fluorescence (TIRF) microscopy (Figure 4B, D, Video 1). PBP1B^Ec diffused on these supported bilayers with an average D_coef of 0.23 ± 0.06 µm²/s. Addition of lipid II slowed down PBP1B^Ec diffusion (Figure 4C), resulting in a lower average D_coef of 0.10 ± 0.06 µm²/s. Upon addition of lipid II, we could not detect a prolonged confined motion within particle tracks (Figure 4D); however, the average length of displacements between two sequential frames was reduced (Figure 4E). Thus, we successfully reconstituted diffusing PBP1B^Ec in SLBs and observed that lipid II binding slowed down the diffusion of the synthase.

(A) Schematic illustration of the approach (not to scale). A single-cysteine version of PBP1B^Ec (MGC-⁶⁴PBP1B-his C777S C795S) labelled with fluorescent probe Dy647 in its single Cys residue (PBP1B^Ec-Dy647) was reconstituted into a polymer-supported lipid membrane formed with *E. coli* polar lipids, and its diffusion was monitored using TIRF microscopy in the presence or absence of substrate lipid II. (B) Single-molecule TIRF micrograph of PBP1B^Ec-Dy647 diffusing in the lipid membrane in the presence of 1.5 µM lipid II (corresponding to Video 1). Calculated particle tracks are overlaid. (C) Histograms of diffusion coefficients (D_coef) of PBP1B^Ec-Dy647 particles in the presence (red) or absence (black) of lipid II. The average D_coef decreased from 0.23 ± 0.06 µm²/s to 0.1 ± 0.04 µm²/s upon addition of lipid II. Values are mean ± SD of tracks from three independent experiments. (D) Representative tracks for diffusing PBP1B^Ec-Dy647 particles in the absence (black, top) or presence of lipid II (red, bottom), showing the absence of confined motion in the presence of lipid II. (E) Displacement distributions of PBP1B^Ec-Dy647 particles (solid lines) in the absence (left) or presence (right) of lipid II were analysed using a Rayleigh model incorporating two populations of particles, a fast-diffusing one (grey dashed lines) and a slow-diffusing one (black dashed lines). In the absence of lipid II, only 8 ± 5% of the steps were classified into the slow fraction (121 ± 6 nm average displacement), while the majority of steps were of 257 ± 6 nm (fast fraction). The slow fraction increased upon addition of lipid II to 37 ± 5% of the steps, with an average displacement of 132 ± 16 nm.

Figure 4—source data 1. Numerical data to support graphs in Figure 4.

elife-61525-fig4-data1.xlsx^{(126.3KB, xlsx)}

Figure 4—figure supplement 1. — (A) Schematic illustration of the approach (not to scale). A single-cysteine version of PBP1B^Ec (MGC-⁶⁴PBP1B-his C777S C795S) labelled with fluorescent probe Dy647 in its single Cys residue (PBP1B^Ec-Dy647) was reconstituted into a polymer-supported lipid membrane formed with *E. coli* polar lipids, and its diffusion was monitored using TIRF microscopy in the presence or absence of substrate lipid II. (B) Single-molecule TIRF micrograph of PBP1B^Ec-Dy647 diffusing in the lipid membrane in the presence of 1.5 µM lipid II (corresponding to Video 1). Calculated particle tracks are overlaid. (C) Histograms of diffusion coefficients (D_coef) of PBP1B^Ec-Dy647 particles in the presence (red) or absence (black) of lipid II. The average D_coef decreased from 0.23 ± 0.06 µm²/s to 0.1 ± 0.04 µm²/s upon addition of lipid II. Values are mean ± SD of tracks from three independent experiments. (D) Representative tracks for diffusing PBP1B^Ec-Dy647 particles in the absence (black, top) or presence of lipid II (red, bottom), showing the absence of confined motion in the presence of lipid II. (E) Displacement distributions of PBP1B^Ec-Dy647 particles (solid lines) in the absence (left) or presence (right) of lipid II were analysed using a Rayleigh model incorporating two populations of particles, a fast-diffusing one (grey dashed lines) and a slow-diffusing one (black dashed lines). In the absence of lipid II, only 8 ± 5% of the steps were classified into the slow fraction (121 ± 6 nm average displacement), while the majority of steps were of 257 ± 6 nm (fast fraction). The slow fraction increased upon addition of lipid II to 37 ± 5% of the steps, with an average displacement of 132 ± 16 nm.

Figure 4—source data 1. Numerical data to support graphs in Figure 4.

elife-61525-fig4-data1.xlsx^{(126.3KB, xlsx)}

Video 1. Single-molecule imaging of PBP1B on supported lipid bilayers (SLBs).

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PBP1B^Ec-Dy647 was reconstituted in E. coli polar lipids SLBs at a 1:10⁶ (mol:mol) protein-to-lipid ratio and was tracked using single-molecule TIRF before or after the addition of 1.5 µM lipid II. Images were taken with a rate of 62 ms per frame.

Next, we wanted to confirm that PBP1B^Ec remained active to produce planar bilayer-attached PG. We incubated SLBs containing PBP1B^Ec with radioactive lipid II and digested any possible PG produced with a muramidase and analysed the digested material by HPLC. Due to the low density and amount of PBP1B^Ec on each SLB chamber, we expected a small amount of PG product; hence, we included LpoB(sol) to boost the activity of PBP1B^Ec. Under these conditions, about 12% of the added radiolabelled lipid II was incorporated into PG after an overnight incubation (Figure 4—figure supplement 2A). However, products of both the GTase and TPase activities of PBP1B^Ec were detected, and these products were absent in the presence of moenomycin (Figure 4—figure supplement 2B). After overnight PG synthesis reactions with radioactive lipid II, about 32% of the radioactivity remained in the membrane fraction after washing (PG products and unused lipid II) and 68% was in the supernatant. The analysis of the membrane and wash fractions by HPLC (Figure 4—figure supplement 2C, D) revealed that SLB-reconstituted PBP1B^Ec produced crosslinked PG while, importantly, the wash fraction contained no PG products, confirming that the PG synthesis occurred on the SLBs and this PG remained attached to the bilayer. The fraction of membrane-attached radioactivity was almost the same (33%) when PBP1B^Ec was not present in the bilayer, indicating that PBP1B^Ec did not affect lipid II binding to the bilayer.

FRET assay on supported bilayers

Next, we adapted the FRET assay to SLBs and TIRF microscopy, taking advantage of the photostability and brightness of the Atto550 and Atto647n probes. Our aim was to visualize PG synthesis by class A PBPs at high resolution as a first step towards understanding PG synthesis at a single molecule level. We used a similar approach as for liposomes, where both Atto550- and Atto647n-labelled lipid II were co-reconstituted with PBP1B^Ec on SLBs and PG synthesis was triggered by the addition of unlabelled lipid II (Figure 3A). To measure any change in FRET due to PG synthesis, we took advantage of the fact that upon photobleaching of the acceptor probe in a FRET pair the emitted fluorescence intensity of the donor increases as absorbed energy cannot be quenched by a nearby acceptor (Loose et al., 2011; Verveer et al., 2006). Indeed, we detected an increase in lipid II-Atto550 fluorescence intensity upon photobleaching of the Atto647n probe after the addition of unlabelled lipid II and LpoB(sol), indicating the presence of FRET (Figure 5A, Figure 5—figure supplement 1A). When we bleached the acceptor at different time points of the reaction, we found the FRET signal to increase after a lag phase of ~8 min. Importantly, there was no FRET increase in the presence of ampicillin (Figure 5B, Figure 5—figure supplement 1A, Video 2) or when a GTase-defective PBP1B^Ec version (E233Q) was used (Figure 5C). In addition, the FRET signal was abolished when the muramidase cellosyl was added after the PG synthesis reaction (Figure 5C). These results imply that the FRET signal detected by microscopy is primarily due to the transpeptidase activity of PBP1B^Ec, in agreement with the results obtained on liposomes (Figure 5C).

(A) FRET acquisition by TIRF microscopy. PBP1B^Ec was reconstituted into a polymer-supported lipid membrane to preserve its lateral diffusion. A supported lipid membrane was formed from *E. coli* polar lipid extract supplemented with 0.5 mol% of labelled lipid II (Atto550 and Atto647n at 1:1 ratio). To initiate peptidoglycan (PG) polymerization, unlabelled lipid II (10 µM) and LpoB(sol) (4 µM) were added from the bulk solution. An increase in FRET efficiency was recorded by dual-colour TIRF microscopy: the acceptor (lipid II-Atto647n) was photobleached, and the concomitant increase in the donor intensity (lipid II-Atto550) was recorded within a delay of 1 s. (B) FRET kinetics of PG polymerization and crosslinking. Inhibition of PBP1B^Ec TPase activity with 1 mM ampicillin did not produce any changes in the donor intensity, confirming that FRET signal is specific to crosslinked PG. A sigmoid (straight lines) was fitted to the data to visualize the lag in the increase of FRET signal. (C) FRET efficiency was measured after a round of PG synthesis before and after digestion with the muramidase cellosyl. After cellosyl digestion, FRET efficiency decreased by 2.5-fold, resulting in a FRET signal comparable to the one of a control surface with a GTase-defective PBP1B^Ec(E233Q), performed in parallel. Each dot corresponds to a different surface area within the same sample. (D) Quantification of the diffusion coefficient of lipid II-Atto647n over the time course of PG polymerization (left) from the experiment presented in B, calculated from the dynamics of the recovery of lipid II-Atto647n signal within the photobleached region of interest (ROI). (E) Quantification of the fraction of immobile lipid II-Atto647n from several experiments as the one depicted in B; each dot represents the value from a different experiment. (F) Diffusion of lipid II-Atto647n or a phospholipid bound probe labelled with Alexa 488 (supported lipid bilayer) was recorded in a FRAP assay using a 1 s delay and dual-colour imaging, 30 min after initiation of PG synthesis by addition of lipid II and LpoB(sol). Only the diffusion of lipid II, but not of a fluorescently labelled, His₆-tagged peptide attached to dioctadecylamine-tris-Ni²⁺-NTA, was affected by the presence of ampicillin during the PG synthesis reaction.

Figure 5—source data 1. Numerical data to support graphs in Figure 5.

elife-61525-fig5-data1.xlsx^{(31.1KB, xlsx)}

Figure 5—figure supplement 1. — (A) FRET acquisition by TIRF microscopy. PBP1B^Ec was reconstituted into a polymer-supported lipid membrane to preserve its lateral diffusion. A supported lipid membrane was formed from *E. coli* polar lipid extract supplemented with 0.5 mol% of labelled lipid II (Atto550 and Atto647n at 1:1 ratio). To initiate peptidoglycan (PG) polymerization, unlabelled lipid II (10 µM) and LpoB(sol) (4 µM) were added from the bulk solution. An increase in FRET efficiency was recorded by dual-colour TIRF microscopy: the acceptor (lipid II-Atto647n) was photobleached, and the concomitant increase in the donor intensity (lipid II-Atto550) was recorded within a delay of 1 s. (B) FRET kinetics of PG polymerization and crosslinking. Inhibition of PBP1B^Ec TPase activity with 1 mM ampicillin did not produce any changes in the donor intensity, confirming that FRET signal is specific to crosslinked PG. A sigmoid (straight lines) was fitted to the data to visualize the lag in the increase of FRET signal. (C) FRET efficiency was measured after a round of PG synthesis before and after digestion with the muramidase cellosyl. After cellosyl digestion, FRET efficiency decreased by 2.5-fold, resulting in a FRET signal comparable to the one of a control surface with a GTase-defective PBP1B^Ec(E233Q), performed in parallel. Each dot corresponds to a different surface area within the same sample. (D) Quantification of the diffusion coefficient of lipid II-Atto647n over the time course of PG polymerization (left) from the experiment presented in B, calculated from the dynamics of the recovery of lipid II-Atto647n signal within the photobleached region of interest (ROI). (E) Quantification of the fraction of immobile lipid II-Atto647n from several experiments as the one depicted in B; each dot represents the value from a different experiment. (F) Diffusion of lipid II-Atto647n or a phospholipid bound probe labelled with Alexa 488 (supported lipid bilayer) was recorded in a FRAP assay using a 1 s delay and dual-colour imaging, 30 min after initiation of PG synthesis by addition of lipid II and LpoB(sol). Only the diffusion of lipid II, but not of a fluorescently labelled, His₆-tagged peptide attached to dioctadecylamine-tris-Ni²⁺-NTA, was affected by the presence of ampicillin during the PG synthesis reaction.

Figure 5—source data 1. Numerical data to support graphs in Figure 5.

elife-61525-fig5-data1.xlsx^{(31.1KB, xlsx)}

Video 2. Förster resonance energy transfer (FRET) assay on supported lipid bilayers (SLBs).

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PBP1B^Ec was reconstituted in E. coli polar lipids SLBs at a 1:10⁵ (mol:mol) protein-to-lipid ratio along lipid II-Atto647 and lipid II-Atto550. Membranes were incubated with 5 µM lipid II in the presence or absence of 1 mM ampicillin. To detect FRET, the fluorescence of the acceptor Atto647n was bleached within a region. In the subsequent frame, the fluorescence of Atto550 increased, indicating the presence of FRET. In the presence of ampicillin, this increase did not happen.

PG synthesized on SLBs

As our experiments confirmed that the PG synthesized by PBP1B^Ec on SLBs remained attached to the bilayer, we next analysed the lateral diffusion of lipid II-Atto647n and its products during PG synthesis reactions. We first analysed the recovery of fluorescence intensity after photobleaching to monitor the diffusion of lipid II-Atto647n during PG synthesis (Figure 5D). Only when crosslinking was permitted (absence of ampicillin), the diffusion coefficient of lipid II-Atto647n decreased two- to threefold in a time-dependent manner. The time needed to reach the minimum diffusivity value (~10 min) was similar to the lag detected in the increase of FRET efficiency (Figure 5B). The fraction of immobile lipid II-Atto647n did not change significantly in the presence or absence of ampicillin (13 ± 2% or 18 ± 6%, respectively, p-value=0.15) (Figure 5E), indicating that the crosslinked PG was still mobile under these conditions, but diffused more slowly. We also compared the diffusion of lipid II-Atto647n during the PG synthesis reaction with that of an Alexa Fluor 488-labelled membrane-anchored peptide in the presence or absence of ampicillin (Figure 5F, Figure 5—figure supplement 1B). The inhibition of TPase by ampicillin only affected the diffusivity of lipid II (2.9 ± 0.4 µm²/s with ampicillin and 0.67 ± 0.1 µm²/s without), while that of the lipid probe remained unchanged (1.6 ± 0.65 µm²/s with ampicillin and 1.94 ± 0.62 µm²/s without). This shows that the membrane fluidity was not altered by the PG synthesis reaction and therefore was not the cause of the change in lipid II diffusivity upon transpeptidation. As the immobile fraction of labelled lipid II did not increase after PG synthesis and the diffusion was reduced only two- to threefold, we concluded that lipid II-Atto647n was incorporated into small groups of crosslinked glycan chains which can still diffuse on the bilayer.

In summary, we report the incorporation of active PBP1B^Ec into SLBs, where we could track a decrease in the diffusion of the protein and its substrate during PG synthesis reactions. Using this system, we detected an increase in FRET upon initiation of PG synthesis, only occurring when transpeptidation was not inhibited.

Discussion

Although class A PBPs are membrane proteins and PG precursor lipid II is embedded in the bilayer, few studies have provided information about the activity of these important enzymes in a membrane environment. Here, we developed a new assay that reports on PG synthesis by these enzymes in detergents, on liposomes, or on SLBs.

Intra-chain vs. inter-chain FRET

For all PBPs and conditions tested, FRET increased when only the GTase domain was active (i.e., when FRET occurred between probes incorporated along the same strand), but the FRET signal was always higher when transpeptidase was active (Figures 1–3, Figure 3—figure supplements 7 and 8). For detergent-solubilized PBP1B^Ec, the FRET curve closely followed the rate of the production of crosslinked PG as determined by HPLC analysis of the products (Figure 2C–E), and the FRET of PBP1B^Ec-produced labelled PG decreased substantially upon digestion with an endopeptidase (Figure 2A, B). These results indicate that inter-chain FRET (arising from both fluorophores present on different, adjacent glycan chains) was a main component of the total FRET signal. Why is this the case? FRET depends on the distance and orientation of the two probes. It might be sterically unfavourable that two large Atto550 and Atto647n containing lipid II molecules simultaneously occupy the donor and acceptor sites in the GTase domain (Van't Veer et al., 2016), preventing the incorporation of probes (and high FRET) at successive subunits on a single glycan chain. Indeed, for all PBPs tested either in detergents or liposomes, the incorporation of labelled lipid II into glycan chains was more efficient when unlabelled lipid II was present, and, for most enzymes, an activator was required to polymerize glycan chains using labelled lipid II in the absence of unlabelled lipid II. We thus hypothesize that the TPase activity brings glycan chains to close proximity, reducing the distance between probes sufficiently to produce high levels of FRET (Figure 6).

Figure 6. — (A) A mixture of Atto550-lipid II, Atto647n-lipid II, and unlabelled lipid II is utilized by a class A penicillin-binding protein (PBP) with or without inhibition of the TPase activity by a β-lactam. FRET can only occur between fluorophores within the same glycan strand in linear glycan chains produced in the presence of a β-lactam (left reaction, dashed arrows). When the TPase is active (right reaction), FRET can occur either between probes within the same strand (dashed arrows) or between probes on different strands of the crosslinked PG product (solid arrows). We hypothesize that at any time only one labelled lipid II molecule occupies the two binding sites in the GTase domain and that therefore two probes within the same strand are separated by at least one subunit. As a result, average distances between probes in different strands may be shorter than between probes within the same strand, and thus inter-chain FRET contributes stronger to the total FRET signal than intra-chain FRET. (B) Lipoprotein-stimulated PBPs produced short chains when labelled lipid II versions were incubated in the absence of unlabelled lipid II (e.g., Figure 1B, Figure 1—figure supplement 1C). In this situation, crosslinking does not occur due to the attachment of the probe to the mDAP residue in the pentapeptide. Within these short strands, intra-chain FRET is stronger than within the long glycan strands depicted in (A) due to a shorter average distance between the probes.

Limitation of the FRET assay

The FRET assay is sensitive and currently the only method that allows to follow PG synthesis continuously in the membrane. Naturally, there are also limitations with the assay. First, the overall FRET signal is a combination from intra-chain and inter-chain FRET, which both depend on the average distances and orientation of the fluorophore molecules on the growing glycan chains. We currently do not have a method to measure these parameters individually and determine whether and how they change during the process of PG synthesis, preventing the determination of absolute rates for GTase and TPase reactions. Second, different class A PBPs may produce slightly different distribution and density of fluorophores in the PG synthesized, hence differences in FRET signals may not always reflect different reaction rates. Third, an activator can potentially enhance the ability of a class A PBP to incorporate the fluorescent lipid II analogues, as we observed for LpoB^Ec and PBP1B^Ec, leading to an increase in intra-chain FRET. Due to these limitations, the assay is inherently semi-quantitative, but with appropriate control samples (β-lactams; only labelled lipid II) it is possible to determine whether the FRET signal follows more the GTase (intra-chain FRET) or TPase (inter-chain FRET) reaction.

Coupled reactions in class A PBPs and their activation

Our assay revealed the effect of Lpo activators on PBP1B analogues from three bacteria. P. aeruginosa uses LpoP to stimulate its PBP1B (Greene et al., 2018; Caveney et al., 2020). Here, we identified an LpoP homologue in A. baumannii and showed that it stimulated its cognate PBP1B. All three PBP1B homologues started the reaction after a lag phase, which was abolished by the addition of the cognate activator (Egan et al., 2014; Caveney et al., 2020) Considering the recently described role of PBP1B in repairing cell wall defects (Vigouroux et al., 2020; Morè et al., 2019), the slow start in polymerization and its acceleration by Lpo activators could be an important mechanism to start PG synthesis at gaps in the PG layer where the activators can contact the synthase.

To distinguish the effects of an activator on the TPase and GTase rates requires to use different assays to measure GTase only or GTase/TPase because ongoing glycan chain polymerization is required for transpeptidation to occur (Bertsche et al., 2005; Gray et al., 2015). An elegant recent report (Catherwood et al., 2020) described the use of a coupled D-Ala release assay to determine the kinetic parameters of the TPase activity of PBP1B^Ec and the effect of LpoB on this rate. Based on their observation that PBP1B^Ec had barely any TPase activity in the absence of LpoB, the authors concluded that the LpoB-mediated TPase activation explains the essentiality of LpoB for PBP1B function in the cell (Catherwood et al., 2020). However, the assay used an enzyme concentration that is too low to support GTase activity in the absence of LpoB, as demonstrated previously (Pazos et al., 2018; Müller et al., 2007). Therefore, the essentiality of LpoB can be readily explained by its primary effect, the greater than tenfold stimulation of the GTase rate (Egan et al., 2014). Our results provide an alternative explanation for PBP1B^Ec essentiality. Activation by LpoB was much more needed when PBP1B was embedded in the membranes of liposomes and supported bilayers, compared to detergent-solubilized enzyme, supporting the idea that cellular PBP1B strictly requires LpoB for GTase activity. In vitro, LpoB also stimulated the TPase causing PBP1B^Ec to produce a hyper-crosslinked PG (Typas et al., 2010; Egan et al., 2018) and the same was observed for LpoP^Pa and PBP1B^Pa (Caveney et al., 2020). The GTase and TPase contribute both to the signal in our FRET assay, and the relative contribution of intra-chain FRET (due to the GTase) and inter-chain FRET (due to the TPase) can be modified by an activator that enables the incorporation of two adjacent probe molecules on the same glycan chain. Therefore, to untangle the effects of activators on each of the activities requires a single quantitative model accounting for the GTase and TPase rates and including parameters for the initiation, elongation, and termination of glycan chain synthesis of membrane-embedded enzymes. Currently, such a model is not available and our assay could help to develop such a model in the future.

Class A PBP activities in the membrane

Remarkably, we found slower reaction rates in liposomes than in detergents for all enzymes tested. Several possible factors can explain this, including a slow incorporation of the added unlabelled lipid II into liposomes, a limited capacity of the liposomes to incorporate the unlabelled lipid II, or the accumulation of the undecaprenyl pyrophosphate by-product that has been showed to inhibit PBP1B activity (Hernández-Rocamora et al., 2018). None of these factors should change in the presence of LpoB. Hence, we favour the alternative explanation that the membrane-embedding of PBP1B hinders lipid II binding, slowing down the reaction. Remarkably, PBP1B^Ab showed higher TPase activity in liposomes than in detergents. This observation highlights again that detergents can affect the activity of membrane proteins and that experimental conditions in PG synthesis assays should be as close as possible at the physiological conditions.

Towards single-molecule PG synthesis

We also adapted the FRET assay to SLBs and super resolution microscopy to study how PBP1B^Ec polymerizes PG on SLBs (Figure 5). As with the liposome assays, we detected an increase in FRET signal upon triggering PG synthesis that correlated with transpeptidation. Importantly, we could follow the diffusion of the reaction products, which indicates that PBP1B^Ec does not completely cover the surfaces with a layer of PG but instead produced smaller patches of crosslinked glycan chains. We attribute this to the fact that PBP1B^Ec was reconstituted at a very low density in order to ensure the homogeneity and stability of the SLBs. Remarkably, we detected a reduction of PBP1B^Ec diffusivity in the presence of lipid II (Figure 4). Previous in vivo single-molecule tracking of fluorescent-protein tagged class A PBPs reported the presence of two populations of molecules, a fast diffusing one and an almost immobile one with a near-zero diffusing rate, which was assumed to be the active population (Cho et al., 2016; Lee et al., 2016; Vigouroux et al., 2020). Our result supports this interpretation, although more experiments are required to further explore this point.

Several real-time methods to study PG synthesis in vitro are described in the literature. However, most of these report on either the GTase or TPase reaction, but not both at the same time, and most available methods are not applicable to the membrane. The scintillation proximity assay by Kumar et al. reports on PG production in a membrane environment and in real time, but it is rather crude in that it uses membrane extract instead of purified protein and relies on the presence of lipid II-synthesizing enzymes present in the extract (Kumar et al., 2014). Moreover, it uses radioactivity detection and is not amenable to microscopy, in contrast to methods based on fluorescently labelled substrates. An important advantage of our new assay over other real-time PG synthesis assays is that it uses natural substrates for transpeptidation, that is, nascent glycan strands, instead of mimics of the pentapeptide, and its ability to measure the activities in a natural lipid environment.

Our new FRET assay can potentially be adopted to assay PG synthases in the presence of interacting proteins, for example, monofunctional class B PBPs in the presence of monofunctional GTases (cognate SEDS proteins or Mtg proteins) or interacting class A PBPs (Meeske et al., 2016; Bertsche et al., 2006; Sjodt et al., 2020; Derouaux et al., 2008; Banzhaf et al., 2012; Sjodt et al., 2018; Taguchi et al., 2019). In addition, our assay has the potential to be adopted to high-throughput screening for new antimicrobials.

Materials and methods

Key resources table.

Reagent type (species) or resource	Designation	Source or reference	Identifiers	Additional information
Strain, strain background (Escherichia coli)	BL21(DE3)	New England Biolabs	C2527
Recombinant DNA reagent	pDML219	Bertsche et al., 2006		Expression of N-terminal His-tagged E. coli PBP1B
Recombinant DNA reagent	pKPWV1B	This paper		Expression of N-terminal His-tagged Acinetobacter baumannii19606 (ATCC) PBP1B
Recombinant DNA reagent	pAJFE52	Caveney et al., 2020		Expression of N-terminal His-tagged Pseudomonas aeruginosa PBP1B
Recombinant DNA reagent	pMGCPBP1BCS1CS2	This paper		Expression of E. coli PBP1B version with a single Cys residue in the N-terminus and C-terminal His-tag
Recombinant DNA reagent	pET28His-LpoB(sol)	Egan et al., 2014		Expression of soluble version of E. coli LpoB with an N-terminal His-tag
Recombinant DNA reagent	pKPWVLpoP	This paper		Expression of N-terminal His-tagged A. baumannii 19606 (ATCC) LpoP
Recombinant DNA reagent	pAJFE57	Caveney et al., 2020		Expression of soluble version of P. aeruginosa LpoP with an N-terminal His-tag
Sequence-based reagent	PBP1B.Acineto-NdeI_f	This paper	PCR cloning primers	AGATATCATATGATGAAGTTTGAACGTGGTATC GGTTTCTTC
Sequence-based reagent	PBP1B.Acineto-BamHI_r	This paper	PCR cloning primers	GCGGGATCCTTAGTTGTTATAACTACCACTTGA AATG
Sequence-based reagent	Seq1_rev_PBP1B_Acineto	This paper	PCR cloning primers	AGGTTCTAAACGGGCAACTC
Sequence-based reagent	Seq2_fwd_PBP1B_Acineto	This paper	PCR cloning primers	TGGTTATGGATTGGCCTCTC
Sequence-based reagent	Seq3_fwd_PBP1B_Acineto	This paper	PCR cloning primers	CTGGGCAAGCCAGATTGAAG
Sequence-based reagent	Seq4_fwd_PBP1B_Acineto	This paper	PCR cloning primers	ACAATTACGCCAGACACCAG
Sequence-based reagent	PBP1B-MGC-F	This paper	PCR cloning primers	CATCATCCATGGGCTGTGGCTGGCTATGGCTACTGCTA
Sequence-based reagent	PBP1B-CtermH-R	This paper	PCR cloning primers	CATCATCTCGAGATTACTACCAAACATATCCTT
Sequence-based reagent	C777S-D	This paper	PCR mutagenesis primers	AACTTTGTTTCCAGCGGTGGC
Sequence-based reagent	C777S-C	This paper	PCR mutagenesis primers	GCCACCGCTGGAAACAAAGTT
Sequence-based reagent	C795S-D	This paper	PCR mutagenesis primers	CAATCGCTGTCCCAGCAGAGC
Sequence-based reagent	C795S-C	This paper	PCR mutagenesis primers	GCTCTGCTGGGACAGCGATTG
Chemical compound	[14C]GlcNAc-labelled lipid II (mDAP)	Breukink et al., 2003 Bertsche et al., 2005
Chemical compound	Lipid II (mDAP)	Egan et al., 2015
Chemical compound	Lipid II (Lys)	Egan et al., 2015
Chemical compound	Lipid II-dansyl	Egan et al., 2015
Chemical compound	Lipid II-Atto550	Mohammadi et al., 2014 Van't Veer, 2016
Chemical compound	Lipid II-Atto647n	Mohammadi et al., 2014 Van't Veer, 2016
Chemical compound	Polar lipid extract from E. coli (EcPL)	Avanti Polar Lipids	100600P
Chemical compound	1,2-Dioleoyl-sn-glycero-3-phosphocholine (DOPC)	Avanti Polar Lipids	850375P
Chemical compound	1-Palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1'-rac-glycerol) (POPG)	Avanti Polar Lipids	840457P
Chemical compound	Tetraoleoyl cardiolipin	Avanti Polar Lipids	710335P
Chemical compound	Dy647P1-maleimide probe	Dyomics	647P1-03
Chemical compound	Alexa Fluor 488 C5 Maleimide	ThermoFisher Scientific	A10254
Chemical compound	Alexa Fluor 555 C2 maleimide	ThermoFisher Scientific	A20346
Chemical compound	Triton X-100	Roche	10789704001
Chemical compound	Moenomycin	Sigma	32404
Chemical compound	Ampicillin	Sigma	A9518
Chemical compound	Methyl-β-cyclodextrin	Sigma-Aldrich	332615
Chemical compound	Poly(ethylene glycol) M_n8000	Sigma-Aldrich	1546605
Chemical compound	1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (DOPE-Rhodamine)	Avanti Polar Lipids	810150C
Chemical compound	Dioctadecylamine (DODA)-tris-Ni-NTA	Beutel et al., 2014
Chemical compound	cOmplete, EDTA-freeProtease Inhibitor Cocktail	Roche Molecular Biochemicals	5056489001
Chemical compound	Phenylmethylsulfonylfluoride (PMSF)	Sigma-Aldrich	P7626
Chemical compound	Ni-NTA superflow resin	Qiagen	1018142
Chemical compound	Bio-Beads SM-2 resin	Bio-Rad	1523920
Commercial assay, kit	Pierce BCA Protein Assay Kit	ThermoFisher Scientific	23227
Commercial assay, kit	HiTrap SP HP column, 1 mL	GE biosciences	17115101
Commercial assay, kit	HiTrap Desalting column, 5 mL	GE biosciences	17140801
Commercial assay, kit	Prontosil 120–3 C18 AQ reversed-phase column	BISCHOFF Chromatography	1204F184P3
Peptide, recombinant protein	DNase	ThermoFisher Scientific	90083
Peptide, recombinant protein	Cellosyl	Hoechst (Germany)		Mutanolysin from Sigma (M9901) can also be used
Peptide, recombinant protein	MepM	Federico Corona, following protocol in Singh et al., 2012
Chemical compound	His6-tagged (on the C-terminus) neutral peptide	BioMatik		CMSQAALNTRNSEEEVSSRRNNGTRHHHHHH
Software, algorithm	Fiji		https://fiji.sc
Software, algorithm	Matlab	MathWorks	https://www.mathworks.com
Software, algorithm	frap_analysis	Jönsson, 2020

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Chemicals

[¹⁴C]GlcNAc-labelled lipid II and the lysine or mDAP forms of lipid II were prepared as published (Breukink et al., 2003; Egan et al., 2015). Lipid II-Atto550 and Lipid II-Atto647n were prepared from the lysine form of lipid II, as described previously (Egan et al., 2015), and Atto550-alkyne or Atto647n-alkyne (Atto tec, Germany) in two steps: (1) conversion of lysine form of lipid II to azidolysine form and (2) labelling of azidolysine lipid II via click-chemistry. The protocol is extensively detailed elsewhere (Mohammadi et al., 2014). The advantage of using this methodology over directly attaching the probes to the amine group is the higher yield of click-chemistry reactions, allowing the use of a smaller excess of the reactive florescent probes (Van't Veer et al., 2016). All lipid II variants were kept in 2:1 chloroform:methanol at −20°C. Before enzymatic assays, the required amounts of lipid II were dried in a speed-vac and resuspended in water (for assays in detergents) or the appropriate buffer (for liposome and SLB assays). Polar lipid extract from E. coli (EcPL), 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1'-rac-glycerol) (POPG), and tetraoleoyl cardiolipin (TOCL) were obtained from Avanti Polar Lipids (USA). Lipids were resuspended in chloroform:methanol (2:1) at a concentration of 20 g/L, aliquoted, and stored at −20°C. Triton X-100, ampicillin, phenylmethylsulfonyl fluoride (PMSF), protease inhibitor cocktail (PIC), and β-mercaptoethanol were from Merck. n-Dodecyl-beta-D-maltopyranoside was purchased from Anatrace (USA). Moenomycin was purchased from Hoechst, Germany. All other chemicals were from Merck.

Cloning

Construction of overexpression vector pKPWV1B

The plasmid pKPWV1B was constructed for overexpression of full-length A. baumannii PBP1B (PBP1B^Ab: aa 1–798) with a cleavable N-terminal oligo-histidine tag (His₆ tag). Therefore, the gene mrcB was amplified using the Phusion high-fidelity DNA polymerase and the oligonucleotides PBP1B.Acineto-NdeI_f and PBP1B.Acineto-BamHI_r and genomic DNA of A. baumannii 19606 (ATCC) as template. The resulting PCR fragment and the plasmid DNA of the overexpression vector pET28a(+) (Novagen) were digested with NdeI and BamHI, ligated, and transformed into chemical-competent E. coli DH5α cells with kanamycin selection. Plasmid DNA of transformants was isolated and sent for sequencing using the following oligonucleotides: Seq1_rev_PBP1B_Acineto, Seq2_fwd_PBP1B_Acineto, Seq3_fwd_PBP1B_Acineto, and Seq4_fwd_PBP1B_Acineto.

Construction of overexpression vector pKPWVLpoP

The sequence of the hypothetical PBP1B activator of A. baumannii 19606 (LpoP^Ab: NCBI reference number: WP_000913437.1) contains a TPR fold and was found by blast analysis through its homology to P. aeruginosa LpoP (30% identity). The plasmid pKPWVLpoP was purchased from GenScript. The gene was synthesized without the first 51 nucleotides (encoding the 17 amino acids of the signal peptide) and with codon optimization for overexpression in E. coli. The codon-optimized gene was subcloned in the overexpression vector pET28a(+) using the cloning sites NdeI and BamHI, enabling the overexpression of the protein with an N-terminal oligo-histidine tag.

MGC-⁶⁴PBP1B-his C777S/C795S

This fusion protein contains PBP1B with the substitution of the N-terminal cytoplasmic tail for residues MGC and the addition of a hexahistine tag at the C-terminus. To obtain this construct, the regions coding for amino acids 64 to 844 of PBP1B were amplified from genomic DNA using oligonucleotides PBP1B-MGC-F and PBP1B-CtermH-R. The resulting product was cloned into pET28a+ vector (EMD Biosciences) after digestion with NcoI and XhoI. C777S and C795S mutations were introduced using the QuikChange Lightning mutagenesis kit (Agilent) through oligonucleotide primers C777S-D, C777S-C, C795S-D, and C795S-C. The resulting plasmid was called pMGCPBP1BCS1CS2.

Purification and labelling of proteins

The following proteins were purified following published protocols: PBP1B^Ec (Bertsche et al., 2006), LpoB(sol) (Egan et al., 2014), PBP1B^Pa (Caveney et al., 2020), LpoP^Pa(sol) (Caveney et al., 2020), and MepM (Singh et al., 2012). All chromatographic steps were performed using an AKTA PrimePlus system (GE Healthcare).

E. coli PBP1B

The protein was expressed as a fusion with an N-terminal hexahistidine tag in E. coli BL21(DE3) pDML924 grown in 4 L of autoinduction medium (LB medium supplemented with 0.5% glycerol, 0.05% glucose, and 0.2% α-lactose) containing kanamycin at 30°C for ~16 hr. Cells were harvested by centrifugation (10,000 × g, 15 min, 4°C) and the pellet resuspended in 80 mL of buffer I (25 mM Tris-HCl, 1 M NaCl, 1 mM EGTA, 10% glycerol, pH 7.5) supplemented with 1× PIC (Sigma-Aldrich), 100 µM PMSF (Sigma-Aldrich), and DNase I. After disruption by sonication on ice, membrane fraction was pelleted by centrifugation (130,000 × g for 1 hr at 4°C) and resuspended in buffer II (25 mM Tris-HCl, 1 M NaCl, 10% glycerol, 2% Triton X-100, pH 7.5) by stirring at 4°C for 24 hr. Extracted membranes were separated from insoluble debris by centrifugation (130,000 × g for 1 hr at 4°C) and incubated for 2 hr with 4 mL of Ni²⁺-NTA beads (Novagen) equilibrated in buffer III (25 mM Tris-HCl, 1 M NaCl, 20 mM imidazole, 10% glycerol, pH 7.5). Beads were washed 10 times with 10 mL of buffer III, and the protein was eluted with 3 mL buffer IV (25 mM Tris-HCl, 0.5 M NaCl, 20 mM imidazole, 10% glycerol, pH 7.5). His-PBP1B-containing fractions were pooled and treated with 2 U/mL of thrombin (Novagen) for 20 hr at 4°C during dialysis against dialysis buffer I (25 mM Tris-HCl, 0.5 M NaCl, 10% glycerol, pH 7.5). Protein was then dialyzed in preparation for ion exchange chromatography, first against dialysis buffer II (20 mM sodium acetate, 0.5 M NaCl, 10% glycerol, pH 5.0), then against dialysis buffer II with 300 mM NaCl, and finally against dialysis buffer II with 100 mM NaCl. Finally, the sample was applied to a 1 mL HiTrap SP column (GE Healthcare) equilibrated in buffer A (20 mM sodium acetate, 100 mM NaCl, 10% glycerol, 0.05% reduced Triton X-100, pH 5.0). The protein was eluted with a gradient from 0% to 100% buffer B (as A, with 2 M NaCl) over 14 mL PBP1B-containing fractions that were pooled and dialyzed against storage buffer (20 mM sodium acetate, 500 mM NaCl, 10% glycerol, pH 5.0) and stored at −80°C.

A. baumannii 19606 PBP1B

The protein was expressed in E. coli BL21 (DE3) freshly transformed with plasmid pKPWV1B using the same protocol as PBP1B^Ec. Cells were harvested by centrifugation (6,200 × g for 15 min at 4°C) and resuspended in 120 mL of PBP1B^Ab buffer I (20 mM NaOH/H₃PO₄, 1 M NaCl, 1 mM EGTA, pH 6.0) supplemented with DNase I, PIC (1:1000 dilution), and 100 µM PMSF. After disruption by sonication on ice, the membrane fraction was pelleted by centrifugation (130,000 × g for 1 hr at 4°C) and resuspended in PBP1B^Ab extraction buffer (20 mM NaOH/H₃PO₄, 1 M NaCl, 10% glycerol, 2% Triton X-100, pH 6.0) supplemented with PIC and PMSF by stirring at 4°C for 16 hr. Extracted membranes were separated from insoluble debris by centrifugation (130,000 × g for 1 hr at 4°C) and incubated with 4 mL of Ni²⁺-NTA beads equilibrated in PBP1B^Ab extraction buffer containing 15 mM imidazole. Beads were washed 10 times with 10 mL of PBP1B^Ab wash buffer (20 mM NaOH/H₃PO₄, 10% glycerol, 0.2% Triton X-100, 1 M NaCl, 15 mM imidazole, pH 6.0), and the protein was eluted with 3 mL buffer IV PBP1B^Ab elution buffer (20 mM NaOH/H₃PO₄, 10% glycerol, 0.2% Triton X-100, 1 M NaCl, 400 mM Imidazole, pH 6.0).

PBP1B^Ab-containing fractions were pooled and dialyzed in preparation for ion exchange chromatography, first against PBP1B^Ab dialysis buffer I (20 mM sodium acetate, 1 M NaCl, 10% glycerol, pH 5.0), then against PBP1B^Ab dialysis buffer II (20 mM sodium acetate, 300 mM NaCl, 10% glycerol, pH 5.0), and finally against PBP1B^Ab dialysis buffer III (10 mM sodium acetate, 100 mM NaCl, 10% glycerol, pH 5.0). The sample was centrifuged for 1 hr at 130,000 × g and 4°C, and the supernatant was applied to a 5 mL HiTrap SP HP column equilibrated in PBP1B^Ab buffer A (20 mM sodium acetate, 100 mM NaCl, 10% glycerol, 0.2% Triton X-100, pH 5.0). The protein was eluted from 0% to 100% PBP1B^Ab buffer B (20 mM sodium acetate, 2 M NaCl, 10% glycerol, 0.2% Triton X-100, pH 5.0) over 70 mL. PBP1B^Ab-containing fractions were pooled and dialyzed against PBP1B^Ab storage buffer (10 mM sodium acetate, 500 mM NaCl, 0.2% Triton X-100, 20% glycerol, pH 5.0) and stored at −80°C.

P. aeruginosa PBP1B

The protein was expressed on E. coli BL21(DE3) freshly transformed with plasmid pAJFE52, which encodes PBP1B^Pa as a fusion with an N-terminal hexahistidine tag in E. coli BL21(DE3). Cells were grown in 4 L of LB at 30°C, and expression was induced for 3 hr with 1 mM isopropyl β-D-1-galactopyranoside (IPTG) when the culture reached an OD₅₇₈ of 0.6. PBP1B^Pa was extracted and purified using the same protocol as for E. coli PBP1B, with the exception that only 2 mL of Ni²⁺ beads were used.

MGC-⁶⁴PBP1B-his C777S/C795S

This protein was expressed in E. coli BL21(DE3) freshly transformed with plasmid pMGCPBP1BCS1CS2 and subsequently purified using the same protocol as for the WT protein, except for the addition of 1 mM tris(2-carboxyethyl)phosphine (TCEP) to all purification buffers. The protein was labelled with Dy647-maleimide probe (Dyomics, Germany) following the manufacturer's instructions. Briefly, 10.2 µM protein was incubated with 100 µM probe and 0.5 mM TCEP for ~20 hr at 4°C, and free probe was removed by desalting using a 5 mL HiTrap desalting column (GE Healthcare).

LpoB(sol)

The protein was expressed on E. coli BL21(DE3) transformed with pET28His-LpoB(sol). Cells were grown in 1.5 L of LB plus kanamycin at 30°C to an OD₅₇₈ of 0.4–0.6, and expression was induced with 1 mM of IPTG for 3 hr at 30°C. Cells were pelleted and resuspended in buffer I (25 mM Tris-HCl, 10 mM MgCl₂, 500 mM NaCl, 20 mM imidazole, 10% glycerol, pH 7.5) plus DNase, PIC, and PMSF. Cells were disrupted by sonication on ice and centrifuged (130,000 × g, 1 hr, 4°C) to remove debris. The supernatant was applied to a 5 mL HisTrap HP column (GE Healthcare) equilibrated in buffer I. After washing with buffer I, the protein was eluted with a stepwise gradient with buffer II (25 mM Tris-HCl, 10 mM MgCl₂, 500 mM NaCl, 400 mM imidazole, 10% glycerol, pH 7.5). Fractions containing the protein were pooled and the His-tag was removed by addition of 2 U/mL of thrombin while dialyzing against buffer IEX-A (20 mM Tris-HCl, 1000 mM NaCl, 10% glycerol, pH 8.3). Digested protein was applied to a 5 mL HiTrap Q HP column (GE Healthcare) at 0.5 mL/min. LpoB(sol) was collected in the flow through, concentrated, and applied to size exclusion on a Superdex200 HiLoad 16/600 column (GE Healthcare) at 1 mL/min in a buffer containing 25 mM HEPES-NaOH, 1 M NaCl, 10% glycerol at pH 7.5. Finally, the protein was dialyzed against storage buffer (25 mM HEPES-NaOH, 200 mM NaCl, 10% glycerol at pH 7.5) and stored at −80°C.

A. baumannii 19606 LpoP(sol)

The protein was expressed on E. coli BL21(DE3) transformed with plasmid pKPWVLpoP. Cells were grown overnight at 30°C in 4 L of autoinduction medium. Cells were pelleted by centrifugation (6200 × g for 15 min at 4°C) and resuspended in 80 mL of buffer I (25 mM Tris/HCl, 10 mM MgCl₂, 1 M NaCl, 20 mM imidazole, pH 7.5) supplemented with DNase I, PIC (1:1000 dilution), and 100 µM PMSF. Cells were disrupted by sonication on ice and centrifuged (130,000 × g for 1 hr at and 4°C) to removed debris. The supernatant was incubated for 1 hr with 6 mL Ni-NTA beads preequilibrated in buffer I at 4°C with gentle stirring. The resin was split in two columns, each washed 10 times with 5 mL wash buffer (25 mM Tris/HCl, 10 mM MgCl₂, 1 M NaCl, 20 mM imidazole, pH 7.5), and the protein was eluted 7 times with 2 mL of elution buffer (25 mM Tris/HCl, 10 mM MgCl₂, 1 M NaCl, 400 mM imidazole, pH 7.5). The best fractions according to SDS-PAGE analysis were pooled and dialyzed stepwise against increasing percentage of dialysis buffer I (25 mM HEPES/NaOH, 10 mM MgCl₂, 200 mM NaCl, 10% glycerol, pH 7.5). Thrombin (nine units) was added to the protein to cleave the N-terminal His₆ tag overnight at 4°C. The successful cleavage of the N-terminal His₆ tag was confirmed by SDS-PAGE. The protein was diluted 2× with 25 mM HEPES/NaOH, 10 mM MgCl₂, 10% glycerol, pH 7.5 to reduce the amount of NaCl down to 100 mM. The protein was applied to a 5 mL HiTrap SP HP column and washed with buffer A (25 mM HEPES/NaOH, 10 mM MgCl₂, 100 mM NaCl, 10% glycerol, pH 7.5). The protein was then eluted with a gradient of 100 mM to 1 M NaCl over 50 mL at 1 mL/min using increasing percentage of buffer B (25 mM HEPES/NaOH, 10 mM MgCl₂, 1 M NaCl, 10% glycerol, pH 7.5). Fractions were collected and analysed by SDS-PAGE. The best fractions were pooled, dialyzed against 25 mM HEPES/NaOH, 200 mM NaCl, 10% glycerol, 10 mM MgCl₂, pH 7.5, and the protein was stored at −80°C.

P. aeruginosa LpoP(sol)

The protein was expressed on E. coli BL21(DE3) freshly transformed with plasmid pAJFE57, encoding His₆-LpoP^Pa(sol). Cells were grown on 1.5 L LB at 30°C to an OD₅₇₈ of 0.5, and expression was induced for 3 hr by addition of 1 mM IPTG. After harvesting, cells were resuspended in 80 mL of 25 mM Tris-HCl, 500 mM NaCl, 20 mM imidazole, 10% glycerol at pH 7.5. After addition of PIC and 100 µM PMSF, cells were disrupted by sonication on ice. Debris was removed by centrifugation (130,000 × g, 1 hr, 4°C) and the supernatant was applied to a 5 mL HisTrap column equilibrated in resuspension buffer. After washing with 25 mM Tris-HCl, 1 M NaCl, 40 mM imidazole, 10% glycerol at pH 7.5, the protein was eluted with 25 mM Tris-HCl, 500 mM NaCl, 400 mM imidazole, 10% glycerol at pH 7.5. Fractions containing His-LpoP^Pa(sol) were pooled and the His-tag was removed by addition of 4 U/mL of thrombin while dialyzing against 20 mM Tris-HCl, 200 mM NaCl, 10% glycerol at pH 7.5 for 20 hr at 4°C. The sample was concentrated and further purified by size exclusion column chromatography at 0.8 mL/min using a HiLoad 16/600 Superdex 200 column equilibrated in 20 mM HEPES-NaOH, 200 mM NaCl, 10% glycerol at pH 7.5. LpoP^Pa-containing fractions that were pooled, concentrated, aliquoted, and stored at −80°C.

PG synthesis assays in the presence of detergents

In vitro PG synthesis assay using radiolabelled lipid II in detergents

To assay the in vitro PG synthesis activity of PBP1B^Ec with radiolabelled lipid II substrate in the presence of detergents, we used a previously published assay (Banzhaf et al., 2012; Biboy et al., 2013). Final reactions included 10 mM HEPES/NaOH pH 7.5, 150 mM NaCl, 10 mM MgCl₂, and 0.05% Triton X-100. The concentration of PBP1B^Ec was 0.5 µM. Reactions were carried out for 1 hr at 37°C. Reactions were stopped by boiling for 5 min. Digestion with cellosyl, reduction with sodium borohydride, and analysis by HPLC were performed as described (Biboy et al., 2013).

FRET-based in vitro PG synthesis assay in detergents

For assays in detergents, samples contained 50 mM HEPES/NaOH pH 7.5, 150 mM NaCl, 10 mM MgCl₂, and 0.05% Triton X-100 in a final volume of 50 µL. PBP1B^Ec, PBP1B^Ab, or PBP1B^Pa were added at a concentration of 0.5 µM. When indicated, activators LpoB(sol), LpoP^Ab(sol), or LpoP^Pa(sol) were added at a concentration of 2 µM. Reactions were started by the addition of an equimolar mix of lipid II, lipid II-Atto550, and lipid II-Atto647n, each at 5 µM and monitored by measuring fluorescence using a Clariostar plate reader (BMG Labtech, Germany) with excitation at 540 nm and emission measurements at 590 and 680 nm. In controls containing unlabelled lipid II plus only one of the labelled lipid II versions (lipid II-Atto550 or lipid II-Atto647n) (Figure 1—figure supplement 4), the labelled lipid II was added at 5 µM along 10 µM of unlabelled lipid II. Reactions were incubated at the indicated temperature for 60 or 90 min. After the reaction, emission spectra from 550 to 740 nm were taken in the same plate reader with excitation at 522 nm. When indicated, ampicillin was added at 1 mM and moenomycin was added at 50 µM. After plate reader measurements, reactions were stopped by boiling for 5 min, vacuum-dried using a speed-vac desiccator, and analysed by Tris-Tricine SDS-PAGE as described previously (Van't Veer et al., 2016).

FRET reactions in the presence of radiolabelled lipid II described in Figure 1E, F were performed using the same buffer and substrate and enzyme concentrations as for the plate reader assay but in a final volume of 350 µL. Samples were incubated at 25°C with shaking using an Eppendorf Thermomixer. Also, 50 µL aliquots were taken out at the indicated times and reactions were stopped by addition of 100 µM moenomycin. Samples were then transferred to a 96-well plate to measure FRET as described above. Finally, samples were transferred back to Eppendorf tubes, digested with cellosyl, and reduced with sodium borohydride as described previously (Biboy et al., 2013).

Quantification of lipid II consumption after the FRET assay in detergents

For the assay in Figure 2—figure supplement 1, reactions were performed in the same conditions as described for the FRET assay in detergents, but four different molar proportions of fluorescent lipid II over the total amount of lipid II were used: 80%, 66.7%, 50%, and 20%. Radiolabelled lipid II was included to allow for quantification of non-fluorescent lipid II consumption at the end of reactions. The total concentration of lipid II was kept at 15 µM, and the molar ratio of lipid II-Atto550 to lipid II-Atto647n was 1:1. At each proportion of fluorescent lipid II, PBP1B^Ec reactions were measured in triplicate with and without LpoB (2 µM), and two control reactions with substrate but no enzyme were prepared to determine the amount of each type of lipid II consumed. Reactions were monitored using a plate reader for 1 hr at 25°C as described above and then stopped by boiling for 10 min. Next, 5 µL aliquots were taken from each reaction, dried in a speed-vac, and analysed by SDS-PAGE with fluorescent scanning as described previously (Van't Veer et al., 2016). Fluorescent lipid II consumption was calculated by comparing the intensity of the lipid II bands on reaction lanes and control lanes. To quantify the consumption of radiolabelled lipid II, the unused lipid II was extracted from the remaining aliquot of the reactions with butanol:pyridine (2:1, vol:vol) at pH 4.2, and the radioactivity was quantified by scintillation counting as described previously (Egan et al., 2015).

Analysis of FRET reaction curves

Reaction curves were obtained by calculating the ratio between fluorescence intensity at 680 and 590 nm monitored at every well. This maximizes the amount of information captured from the change in the spectrum due to FRET and normalizes the intensity removing any non-specific jumps in the signal due to bubbles in the reaction well or lamp instability. The slope of reaction curves obtained by the FRET assay was calculated when the ratio started to rise, avoiding the lag phase when present. Only the linear phase of each curve was used. For example, for PBP1B^Ec in detergents, slopes were calculated from 10 to 15 min in the absence of LpoB and within the first minute in the presence of activator. To compare our results with prior reports, we report the fold-change in the slope in the presence of the corresponding Lpo activator, that is, the ratio between the slope in a condition with activator and the slope at the same condition without activator.

Determination of FRET efficiency

For the determination of FRET efficiency, reactions were prepared in the same conditions as for the plate reader assays but they were incubated at 37°C for 1 hr in Eppendorf tubes instead and boiled for 5 min afterwards. For every antibiotic condition, four samples were prepared: sample 1 (DA reaction) contained 5 µM each of unlabelled lipid II, lipid II-Atto550, and lipid II-Atto647n; sample 2 (D reaction) contained 10 µM unlabelled lipid II plus 5 µM lipid II-Atto550; sample 3 (A reaction) contained 10 µM unlabelled lipid II plus 5 µM lipid II-Atto647n; and sample 4 (BG reaction) contained 15 µM unlabelled lipid II. For digestion with hydrolases, 50 µL of the PG synthesis reactions were prepared as described above and split into three aliquots. Either 5 µM MepM, 0.05 mg/mL cellosyl, or buffer was added to a final volume of 20 µL, and samples were incubated overnight at 37°C and boiled for 5 min to stop reactions.

Samples were measured in Cary Varian fluorimeter using a 1.5 mm light-path quartz cuvette. For samples 1, 3, and 4, two spectra were measured, one with excitation at 552 nm ( $λ_{e x}^{D}$ ) and emission collected from 560 to 750 nm (ds spectrum) and the other with excitation at 650 nm ( $λ_{e x}^{A}$ ) and emission collected from 660 to 750 nm (as spectrum). For sample 2, only the ds spectrum was measured. All spectra were taken with 5 nm slits for emission and excitation at the same detector voltage settings (850 V).

FRET efficiency (E) was calculated according to the (ratio)_A method described in Vámosi and Clegg, 1998. Briefly, (ratio)_A is a normalized measure of the enhancement of the acceptor emission due to FRET,

{(r a t i o)}_{A} = \frac{F_{A} (λ_{e x}^{D}, λ_{e m})}{F_{A} (λ_{e x}^{A}, λ_{e m})} = \frac{[ϵ_{D} (λ_{e x}^{D}) E + ϵ_{A} (λ_{e x}^{D})] Φ^{A} (λ_{e m})}{ϵ_{A} (λ_{e x}^{A}) Φ^{A} (λ_{e m})} = \frac{ϵ_{D} (λ_{e x}^{D}) E + ϵ_{A} (λ_{e x}^{D})}{ϵ_{A} (λ_{e x}^{A})}

(1)

where $Φ^{A} (λ_{e m})$ is a shape function of the acceptor emission spectrum, $F_{A} (λ_{e x}^{D}, λ_{e m})$ is the emission of the acceptor (only the acceptor) when excited at $λ_{e x}^{D}$ , and $F_{A} (λ_{e x}^{A}, λ_{e m})$ is the emission of the acceptor when excited at $λ_{e x}^{D}$ , both in the sample containing both donor and acceptor. FRET efficiency is normalized by the extinction coefficients of the donor at $λ_{e x}^{D}$ ( $ϵ_{D} (λ_{e x}^{D})$ = 120,000 M⁻¹cm⁻¹) and of the acceptor at both $λ_{e x}^{D}$ and $λ_{e x}^{A}$ ( $ϵ_{A} (λ_{e x}^{D})$ = 6000 M⁻¹cm⁻¹, and $ϵ_{A} (λ_{e x}^{A})$ = 150,000 M⁻¹cm⁻¹, respectively).

In order to calculate (ratio)_A from the ds spectrum, three spectral contributions ( $δ^{d s}$ , $α^{d s}$ , and $β^{d s}$ ) were fitted in the ds spectra, $F^{d s} (λ)$ , and two spectral contributions ( $α^{a s}$ and $β^{a s}$ ) were fitted in the as spectra, $F^{a s} (λ)$ :

F^{d s} (λ) = δ^{d s} F_{D r e f}^{d s} (λ) + α^{d s} F_{A r e f}^{d s} (λ) + β^{d s} F_{B r e f}^{d s} (λ)

(2)

F^{a s} (λ) = α^{a s} F_{A r e f}^{a s} (λ) + β^{a s} F_{B r e f}^{a s} (λ)

(3)

where $F_{D r e f}^{d s} (λ)$ is the background-free spectra from the donor-only reference sample exited at $λ_{e x}^{D}$ ; $F_{A r e f}^{d s} (λ)$ is the background-free spectra from the acceptor-only reference sample exited at $λ_{e x}^{D}$ ; $F_{A r e f}^{a s} (λ)$ is the background-free spectra from the acceptor-only reference sample exited at $λ_{e x}^{A}$ ; and $F_{B r e f}^{d s} (λ)$ and $F_{B r e f}^{a s} (λ)$ are the background spectra obtained at $λ_{e x}^{D}$ and $λ_{e x}^{A}$ , respectively. (ratio)_A was then calculated from Equation 4, integrating at wavelengths common in both $F_{A r e f}^{d s} (λ)$ and $F_{A r e f}^{d s} (λ)$ (from 660 to 750 nm).

{(r a t i o)}_{A} = \frac{α^{d s} F_{A r e f}^{d s} (λ)}{α^{a s} F_{A r e f}^{a s} (λ)}

(4)

All calculations were implemented in Excel.

Continuous GTase assay using dansylated lipid II

Continuous fluorescence GTase assays using dansylated lipid II and A. baumannii PBP1B were performed as described previously (Schwartz et al., 2001; Offant et al., 2010; Egan and Vollmer, 2016). Samples contained 50 mM HEPES/NaOH pH 7.5, 105 mM NaCl, 25 mM MgCl₂, 0.039% Triton X-100, and 0.14 µg/µL cellosyl muramidase in a final volume of 60 µL. PBP1B^Ab was added at a concentration of 0.5 µM. When indicated, LpoP^Ab(sol) was added at a concentration of 0.5 µM. Reactions were started by addition of dansylated lipid II to a final concentration of 10 µM and monitored by following the decrease in fluorescence over 60 min at 37°C using a FLUOstar OPTIMA plate reader (BMG Labtech, Germany) with excitation at 330 nm and emission at 520 nm. The fold-increase in GTase was calculated against the mean rate obtained with PBP1B^Ab alone at these reaction conditions, at the fastest rate.

Time-course GTase assay by SDS-PAGE followed by fluorescence detection

PBP1B^Ab at a concentration of 0.5 µM was incubated with 5 µM lipid II-Atto550 and 25 µM unlabelled lipid II in the presence or absence of 1.5 µM LpoP^Ab(sol). Reactions contained 20 mM HEPES, 150 mM NaCl, 10 mM MgCl₂, 0.06% TX-100, and 1 mM ampicillin to block transpeptidation. Aliquots were taken after 0, 2, 5, 10, 30, and 60 min incubation at 37°C, boiled for 10 min to stop reactions, and analysed by Tris-Tricine SDS-PAGE followed by fluorescence detection as described previously (Van't Veer et al., 2016).

Fluorescence scanning of SDS-PAGE gels with PG synthesis products

SDS-PAGE gels were scanned using either a Typhoon FLA9500 (GE) or a Typhoon 9400 (Amersham) fluorescence scanner. Atto550 fluorescence was scanned using a 532 nm laser and either a 590 nm, 30 nm-bandwidth bandpass filter (9400) or a 575 nm long-pass filter (FLA9500). Atto647n fluorescence was scanned using a 635 nm laser and either a 670 nm, 30 nm-bandwidth bandpass filter (9400) or a 665 nm long-pass filter (FLA9500). Voltage of the photodetector was carefully adjusted to avoid saturated pixels in the resulting images. For quantification, ImageJ was used utilizing the original files produced by the scanner. For visualization, images were exported using ImageQuant software (GE Healthcare). During this conversion, ImageQuant automatically adjusts contrast so that tenuous bands are easier to visualize. Unfortunately, this adjustment makes the most intense bands (unused lipid II, wells) appear saturated, which can be misleading when interpreting the images. Thus, we provide the original gel files produced by the scanner as source data files.

PG synthesis in liposomes

Reconstitution of class A PBPs in liposomes

Proteoliposomes containing class A PBPs were prepared as described previously with some modifications (Egan et al., 2015; Rigaud and Lévy, 2003; Hernández-Rocamora et al., 2018). The appropriate lipid or mixture of lipids was dried in a glass test tube under stream of N₂ to form a lipid film followed by desiccation under vacuum from 2 hr. When labelled lipid II was co-reconstituted with the indicated class A PBP, they were added at 1:200 mol:mol phospholipid to each lipid II-Atto550 and lipid II-Atto647n. Resuspension into multilamellar vesicles (MLVs) was achieved by addition of 20 mM Tris/HCl, pH 7.5 with or without 150 mM NaCl as indicated in each experiment and several cycles of vigorous mixing and short incubations in hot tap water. The final lipid concentration was 5 g/L. To form large unilamellar vesicles (LUVs), MLVs were subjected to 10 freeze–thaw cycles and then extruded 10 times through a 0.2 µm filter. LUVs were destabilized by the addition of Triton X-100 to an effective detergent:lipid ratio of 1.40 and mixed with proteins in different protein-to-lipid molar ratios (1:3000 for PBP1B^Ec and PBP1B^Pa, and 1:2000 for PBP1B^Ab). After incubation at 4°C for 1 hr, prewashed adsorbent beads (Biobeads SM2, BioRad, USA; 100 mg per 3 µmol of Triton X-100) were added to the sample to remove detergents. Biobeads were exchanged after 2 and 16 hr, followed by incubation with fresh Biobeads for a further 2 hr. After removal of Biobeads by short centrifugation at 4000 × g, liposomes were pelleted at 250,000 × g for 30 min at 4°C. The pellet containing proteoliposomes was resuspended using the appropriate buffer. The resuspension was done in a 43% smaller volume than the volume added of lipid II, so that the final concentration of lipids was 11.6 g/L. Samples were then centrifuged for 5 min at 17,000 × g and 4°C to remove any possible aggregates. The supernatant was then used in the appropriate assays. Liposomes were analysed by SDS-PAGE and, only for liposomes without labelled lipid II, also by bicinchoninic acid assay (Pierce BCA Assay Kit, ThermoFisher Scientific, USA) to determine protein concentration. The concentration of protein for liposomes with labelled lipid II was calculated by densitometry of the samples in SDS-PAGE gels after reactions were carried out.

PBP1B^Ec orientation assay

To assess the orientation of liposome-reconstituted PBP1B^Ec, MGC-⁶⁴PBP1B-his C777S C795S mutant containing a single cysteine in the N-terminal region was reconstituted in liposomes with EcPL as described above. The accessibility of the cysteine was determined using sulfhydryl-reactive fluorescent probe Alexa Fluor555-maleimide. Reactions containing 0.5 µM protein, 10 µM Alexa Fluor555-maleimide, and 0.2 mM TCEP were incubated for 16 hr at 4°C in the presence or absence 0.5% Triton X-100. Reactions were stopped by addition of 5 mM DTT and boiling for 5 min. Samples were loaded in a 10% acrylamide gel and, after electrophoresis, gels were first scanned using an Amersham Typhoon Trio with excitation at 533 nm and a 40 nm-wide band-pass emission filter at 580 nm. The gel was then stained by Coomassie.

In vitro PG synthesis assay using radiolabelled lipid II in liposomes

The same methodology as in detergents was used to assay the in vitro PG synthesis activity of PBP1B^Ec in liposomes, with minor modifications. To start reactions, 1.5 nmol [¹⁴C]-labelled lipid II were dried in a 0.5 mL glass tube using a vacuum concentrator, resuspended in 5 µL of the appropriate liposome buffer, and mixed with liposomes, buffer, and MgCl₂ to a total volume of 50 µL. Final reactions contained 0.5 µM PBP1B^Ec, 30 µM lipid II, and 1 mM MgCl₂ in 20 mM Tris/HCl pH 7.5 with or without 150 mM NaCl as indicated for each experiment. Samples were incubated for 90 min at 37°C with shaking at 800 rpm. Reactions were stopped by boiling for 5 min. Digestion with cellosyl, reduction with sodium borohydride, and analysis by HPLC were performed as described (Biboy et al., 2013).

FRET-based in vitro PG synthesis assay in liposomes

For assays with liposomes, samples contained 20 mM Tris pH 7.5, 1 mM MgCl₂ in a final volume of 50 µL. In this case, the same volume for each liposome preparation was added to the reactions, 10 µL, so that the total amount of labelled lipid II was present in every reaction. In these conditions, concentration of lipid II-Atto550 and lipid II-Atto647n would be 14.5 µM each, assuming no loss of lipids during sample preparation. The final concentrations of enzymes for the reactions shown in Figure 3, determined by densitometry of SDS-PAGE gels, were ~0.59 µM for PBP1B^Ec, ~0.81 µM for PBP1B^Ab, and ~0.53 µM for PBP1B^Pa. When indicated, activators LpoB(sol), LpoP^Ab(sol), or LpoP^Pa(sol) were added at a concentration of 2 µM. Reactions were started by the addition of lipid II at 12 µM and monitored by measuring fluorescence over a period of 60 min (or 90 min for PBP1B^Pa liposomes) at 37°C using a Clariostar plate reader (BMG Labtech, Germany), with emission measurements at 590 and 680 nm after excitation at 522 nm. When indicated, ampicillin was added at 1 mM and moenomycin was added at 50 µM. Activity assays were performed immediately after preparation of liposomes was finished as we noticed that some proteins could slowly start polymerization using the labelled lipid II. After reactions, samples were analysed by Tris-Tricine SDS-PAGE as indicated for detergents.

Analysis of FRET reaction curves

Slopes were calculated as indicated in the FRET assay in the presence of detergent. As it is not possible to precisely adjust the final amount of enzyme in different liposome preparations, there could be differences in activities measured due to different enzyme amounts. Therefore, we calculated the ratio of the slope with activator over the slope without activator for every liposome preparation and then averaged the values (instead of averaging the different measurements from every sample). At least two independent liposome preparations were prepared for every class A PBP.

Assays in SLBs

Preparation of small unilamellar vesicles (SUVs) and proteoliposomes for SLB formation

Liposomes of EcPL lipids and proteoliposomes with reconstituted PBP1B^Ec were prepared as described previously by addition of β-cyclodextrin to the solution of lipids and Triton X-100 detergent (Degrip et al., 1998; Roder et al., 2011). Briefly, a thin lipid film of EcPL extract was prepared by N₂-assisted chloroform evaporation. After 2 hr of drying under vacuum, the lipid film was rehydrated to 5 mM (total phosphorus concentration) in 150 mM NaCl, 10 mM Tris-HCl, pH 7.4 supplemented with 20 mM Triton X-100. The suspension of lipids/detergent was extensively vortex, freeze/thawed for five cycles, and sonicated using a water-bath sonicator for 10 min (on ice, to avoid lipids overheating upon sonication). To prepare proteoliposomes, full-length PBP1B produced as described above and containing 0.05% Triton X-100 was mixed with a lipid–detergent suspension at the indicated ratio, usually 1:25,000 (protein:lipids), and incubated for 10 min at room temperature (RT). Incorporation of PBP1B^Ec into liposomes was achieved by addition of 2× excess of β-cyclodextrin solution for 5 min (at RT) with subsequent twentyfold dilution in 20 mM HEPES, pH 7.4. The rapid depletion of detergent by addition of β-cyclodextrin leads to the formation of very small unilamellar vesicles with an average diameter of 18–25 nm and narrow size distribution (Roder et al., 2011).

To prepare liposomes with fluorescently labelled lipid II, the extract of EcPL was supplemented with 2 mol% solution of either lipid II-Atto550 or lipid II-Atto647n. The lipid film was treated similarly as the film for the preparation of proteoliposomes. Liposomes were also prepared by cyclodextrin-assisted extraction of Triton X-100.

Formation of polymer-SLBs and reconstitution of PBP1B^Ec into a supported lipid membrane

To form polymer-supported lipid membranes, the coverslips were functionalized beforehand with a dense PEG film, where the ends of the polymer brush were covalently modified with palmitic acid, which served as a linker to capture liposomes as described elsewhere (Roder et al., 2011). To perform a FRET assay on supported lipid membrane, empty EcPL liposomes (1), liposomes with 2 mol% of either lipid II-Atto550 (2) or lipid II-Atto647n (3), and PBP1B^Ec proteoliposomes (4) were mixed at equimolar ratio and diluted by twentyfold with the 10 mM Tris pH 7.5 buffer directly in the reaction chamber. After 30 min of incubation at 37°C, the reaction chamber was washed five times by solution exchange. Proteoliposomes adsorbed on the surface were fused by the addition of 10% (w/v) PEG 8 kDa solution (in water). The fusion reaction was carried out for 15 min at 37°C, afterwards PEG solution was rigorously washed out. Fluidity and homogeneity of the lipid membrane were checked either with PE-Rhodamine dye (Avanti) or by addition of a His₆-tagged (on the C-terminus) neutral peptide (CMSQAALNTRNSEEEVSSRRNNGTRHHHHHH) labelled with a single Alexa 488 fluorophore on its only Cys residue at the N-terminus to the EcPL membrane containing 0.1 mol% dioctadecylamine (DODA)-tris-Ni-NTA (Beutel et al., 2014).

FRET-based in vitro PG synthesis assay in SLBs using TIRF microscopy

PG synthesis reactions were carried out at 10 mM Tris pH 7.5 supplemented with 1 mM MgCl_2, with or without 1 mM ampicillin and in the presence of 4 µM LpoB(sol). The reaction was started by addition of 4 µM of unlabelled lipid II. TIRF microscopy, using a set up described elsewhere (Baranova et al., 2020), was used to monitor an increase in FRET efficiency and spatial reorganization of FRET signal over the time course of PG synthesis. To detect real-time FRET on supported lipid membranes, we used the so-called ‘acceptor photobleaching approach’ where a region of interest of about 10 × 10 µm was photobleached in the acceptor channel (lipid II-Atto647n) and the increase in fluorescence intensity of the donor (lipid II-Atto550) was recorded within a delay of 1 s. We found that in our experiments photobleaching of the acceptor dye was the only process that contributed to the recorded increase in the donor fluorescence signal. Accordingly, the relative increase in donor fluorescence can be used as a direct readout for the FRET efficiency and could therefore be calculated as described (Loose et al., 2011; Verveer et al., 2006). Briefly, donor intensity levels were calculated before (I^D) and after photobleaching (I^D,pb) using intensity measurements in ImageJ. FRET efficiency was calculated using Equation 5:

E = (I^{D, p b} - I^{D}) / I^{D, p b}

(5)

For time-course measurements (Figure 4D), the acceptor signal (lipid II-Atto647n) was photobleached every minute after the initiation of the reaction (the data point at time 0 corresponds to the addition of unlabelled lipid II). For each time point, a new region of interest in the same chamber was photobleached, and the change in the donor intensity was recorded to calculate FRET efficiency using Equation 1.

To have a control on the lipid membrane integrity during PG synthesis, the phospholipid DODA-tris-Ni-NTA (Beutel et al., 2014) was included during reconstitution at a 0.1 mol% ratio. DODA-tris-Ni-NTA was then visualized using a His₆-containing peptide (CMSQAALNTRNSEEEVSSRRNNGTRHHHHHH) labelled with Alexa488 on its single Cys residue, which we added in the same experiment in which we performed FRET analysis. To compare the fluidity and immobile fraction of lipid II-Atto647n before and after 1 hr of the synthesis reaction with the fluidity of phospholipids in the lipid membrane, the same region of interest was photobleached with a laser first at 640 nm and afterwards at 480 nm.

In vitro PG synthesis assay using radiolabelled lipid II on SLBs

To assay PG synthesis on SLBs using radioactively labelled lipid II, we first reconstituted PBP1B^Ec on SLBs containing EcPL extract and a 1:10⁵ PBP1B^Ec to lipid molar ratio, as described above. Due to the low density of the enzyme, several 1.1 cm² chambers were assayed for every condition in order to accumulate a measurable signal. In every chamber, reactions were started by addition of 10 µM [¹⁴C]-labelled lipid II and 4 µM LpoB(sol) in a total volume of 100 µL per chamber. The synthesis reaction was carried out in 10 mM Tris pH 7.5, 1 mM MgCl₂. The chambers were incubated overnight (~16 hr) at 37°C and covered with parafilm. Reactions were stopped by addition of 100 µM moenomycin. To digest the produced PG, cellosyl was added at 0.05 g/L in the presence of 0.3% Triton X-100. After 1 hr incubation at 37°C, samples from six chambers were pooled in an Eppendorf tube, concentrated using a speed-vac evaporator, reduced using sodium borohydride, and analysed by HPLC as described above. For the experiment to determine lipid II incorporation and the localization of the produced PG, before addition of moenomycin, chambers were washed by removal of 50 µL of buffer and addition of 50 µL of fresh buffer while mixing. This was repeated five times. The removed volume from each wash was pooled and treated the same as the samples left in the chamber.

Single-molecule tracking and analysis

To perform single-molecule tracking, MGC-⁶⁴PBP1B-his C77S C795S was labelled with the photostable far-red dye Dy647N as described above and then reconstituted into a polymer-supported lipid membrane as described elsewhere (Roder et al., 2011; Roder et al., 2014). Single-molecule tracking experiments were performed at a low protein-to-lipid molar ratio (1:10⁻⁶). At this ratio, supported lipid membrane was largely homogeneous with the lowest immobile fraction from all the ratios tested (Figure 3—figure supplement 1). The single-molecule motion of PBP1B was measured prior to and after the addition of 1.5 µM lipid II after 15 min ex situ incubation in the presence of 10 mM HEPES pH 7.4, 150 mM NaCl, 1 mM MgCl₂ buffer and in the absence of LpoB(sol). The localization and tracking of PBP1B particles were performed by the SLIMfast software (Roder et al., 2014). To ensure that non-specifically stuck PBP1B particles did not contribute to the measured diffusion coefficient, the immobile particles were excluded using the DBSCAN spatial clustering algorithm (Sander et al., 1998) with the following clustering parameters: a search area of 100 nm and a minimal time window of 30 frames at 65 ms/frame acquisition time. The displacement distribution for active PBP1B (in the presence of lipid II) was compared to the displacement distribution of PBP1B before lipid II addition by fitting the two-component Rayleigh distribution and comparing the weighted contribution of each population. The mean-squared displacement (MSD) was fitted to each individual trajectory longer than 650 ms (10 frames). Each MSD curve was fitted with a linear fit considering max 30% of the lag time for each trajectory.

FRAP analysis

To control membrane fluidity upon the reconstitution of the transmembrane PBP1B (Figure 3—figure supplement 1, Figure 4—figure supplement 1) and fluidity of lipid II Atto-647n during PG synthesis (Figure 4E, F), we used a Matlab-based GUI frap_analysis (Jönsson, 2020) in details described elsewhere (Jönsson et al., 2008). This code allows to quantify the contribution of the immobile fraction to the estimated diffusion coefficient and is particularly suitable for the analysis of 2D diffusion with the photobleaching contribution during the recovery.

Acknowledgements

We thank Alexander Egan (Newcastle University) for purified proteins LpoB(sol) and LpoP^Pa(sol), Federico Corona (Newcastle University) for purified MepM, and Oliver Birkholz and Jacob Piehler (Department of Biology and Center of Cellular Nanoanalytics, University of Osnabrück) for their help with PBP1B reconstitution into polymer-SLBs and initial guidance on single particle tracking. We also acknowledge Christian P Richter and Changjiang You (Department of Biology and Center of Cellular Nanoanalytics, University of Osnabrück) for providing SLIMfast software and tris-DODA-NTA reagent, respectively. This work was funded by the BBSRC grant BB/R017409/1 (to WV), the European Research Council through grant ERC-2015-StG-679239 (to ML), and long-term fellowships HFSP LT 000824/2016-L4 and EMBO ALTF 1163–2015 (to NB).

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Contributor Information

Martin Loose, Email: mloose@ist.ac.at.

Waldemar Vollmer, Email: w.vollmer@ncl.ac.uk.

Jie Xiao, Johns Hopkins University, United States.

Olga Boudker, Weill Cornell Medicine, United States.

Funding Information

This paper was supported by the following grants:

BBSRC BB/R017409/1 to Waldemar Vollmer.
European Research Council ERC-2015-StG-679239 to Martin Loose.
EMBO EMBO ALTF 1163-2015 to Natalia Baranova.
Human Frontiers Science Program HFSP LT 000824/2016-L4 to Natalia Baranova.

Additional information

Competing interests

No competing interests declared.

Author contributions

Conceptualization, Data curation, Formal analysis, Validation, Investigation, Methodology, Writing - original draft, Writing - review and editing.

Conceptualization, Data curation, Formal analysis, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing.

Data curation, Formal analysis, Investigation, Writing - original draft, Writing - review and editing.

Resources, Writing - review and editing.

Conceptualization, Data curation, Formal analysis, Funding acquisition, Validation, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing.

Conceptualization, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Methodology, Writing - original draft, Project administration, Writing - review and editing.

Additional files

Supplementary file 1. Table of oligonucleotides used in this study.

elife-61525-supp1.docx^{(22.5KB, docx)}

Transparent reporting form

elife-61525-transrepform.docx^{(253.8KB, docx)}

Data availability

All data generated or analysed during this study are included in the manuscript and supporting files. Source data files have been provided for Figures 1–5 and the corresponding figure supplements.

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eLife. doi: 10.7554/eLife.61525.sa1

Decision letter

Editor: Jie Xiao¹

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

In the work by Hernandez-Rocamora, a new FRET assay was developed to monitor the peptidoglycan synthesis of bifunctional PBPs. This new assay has the major advantages of being real-time, applicable to membrane-embedded enzymes, and able to monitor both GTasea and TPase activities at the same time. While there exist complications in the interpretation of the FRET signal quantitatively, with appropriate controls this assay will greatly help researchers in the field to determine and compare enzymatic activities and kinetics semi-quantitatively. The assay also has the potential to be developed further into single-molecule assays, allowing mechanistic studies of cell wall synthesis enzymes.

Decision letter after peer review:

Thank you for submitting your article "Real time monitoring of peptidoglycan synthesis by membrane-reconstituted penicillin binding proteins" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Olga Boudker as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

As the editors have judged that your manuscript is of interest, but as described below that the manuscript needs to be substantially revised to provide new biological insight if possible, with additional control experiments, critical analyses and clarifications.

We would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). First, because many researchers have temporarily lost access to the labs, we will give authors as much time as they need to submit revised manuscripts. We are also offering, if you choose, to post the manuscript to bioRxiv (if it is not already there) along with this decision letter and a formal designation that the manuscript is "in revision at eLife". Please let us know if you would like to pursue this option. (If your work is more suitable for medRxiv, you will need to post the preprint yourself, as the mechanisms for us to do so are still in development.)

Summary:

The manuscript by Hernández-Rocamora et al. describes a FRET-based real time assay that allows for continuous monitoring of peptidoglycan synthesis by aPBPs. The authors examine how the assay performs in detergent and in liposomes; they also assess whether the assay will be useful for studying regulators of peptidoglycan biosynthesis enzymes by applying it to previously studied lipoprotein activators of aPBPs, and a new lipoprotein activator that is similar to a previously studied one. The major advantage of the current assay in comparison to the original dansylated lipid II assay is that the FRET assay can monitor both GTase and TPase activities. Finally, the current FRET assay has the potential to become single-molecule study-compatible. The Vollmer lab is one of only a few that can carry out studies of PG biosynthetic enzymes, and he has teamed up with an expert in microscopy on this manuscript. The experiments are technically challenging and generally well done and the technical advances should be useful to the field. However, new biological/mechanistic insights are few. The work also needs to be significantly strengthened with more critically analyzed data, control experiments and better writing.

Reviewer #1:

1) FRET controls and calculations:

a) The authors should provide donor-only and acceptor-only controls under the same conditions for all experiments to allow the calculation of FRET efficiency quantitatively (preferably using the Ratio A method Clegg RM, Methods in Enzymology, 1992, 211, p353). This quantification is important because one major potential advantage of this method could be studying the kinetics of GTase and TPase activities, which may offer mechanistic insight on the enzyme's working. The Facceptor /Fdonor ratio measurement is convenient and can be useful for screening of a large sample size but is prone to artifacts such as fluorescence quenching due to reasons other than FRET, especially when the way the system is set up does not allow precise determination of the donor/acceptor ratio and attachment positions. These donor only or acceptor only experiments are also important for the FRAP measurement in the SLB assay.

b) In Figure 4 the authors used a ratiometric FRET equation Erel = IA/(IA+ID) in the semi-single-molecule assay to determine FRET efficiency. This measurement should be unified to allow direct comparison of FRET between different types of experiments (solubilized, liposomes, SLBs, bulk or single-molecule). As mentioned above, the acceptor intensity consists of the intensities from donor-excitation (FRET), direct-excitation, and bleed-through from the donor. Donor emission could be quenched due to reasons (such as the local environment as that in the dansyl-lipid II assay) other than FRET. More control experiments will allow the authors to determine how this impact the true acceptor signal resulting from FRET.

c) There appears to be a major difference in the fluorescence intensity spectra between the detergent-solubilized (Figure 1—figure supplement 1C) and liposome experiments (Figure 2—figure supplement 2A, C, D). In the detergent solubilized experiments, there appears to be little change in the intensity of the donor peak from PBP1B to +LpoB. However, in the liposome experiments the LpoB has a drastic loss of donor signal compared to that of PBP1B. This is another reason why calculating FRET efficiency instead of the relative ratio would allow for a better comparison between the experiments. This drastic loss of donor signal in the liposome experiments is seen in all 3 species. The authors should address why there is such a difference in the FRET profiles between these two experiments.

d) The authors rely on "Relative Slope" to show how the addition of LpoB impacts the PBP1B activity. Is the "Relative slope" calculated by normalizing the slope of a reaction with LopB to that without LopB? How is the time window determined? As this ratio does not really inform on the enzyme activity, it would be difficult for other groups to use the same method and to compare to this metric. It is also difficult to judge whether the GTase is faster or slower in the presence of TPase activity without the absolute values. Please provide the absolute measurement of the slope (or the initial velocity V0) using a better determined FRET efficiency value as in point 1a.

e) Additionally, perhaps the authors could determine the reaction velocities at different substrate concentrations to see if Michaelis-Menten kinetics could be applied? A reanalysis of the data with better quantifications with regard to enzyme's kinetics would benefit the community significantly by providing a more complete picture of enzyme's activity rather than only comparing PBP1B alone to +LpoB.

2) According to the authors' data and others' work, PBP1B can only synthesize short, uncrosslinked PG chains when its TPase is inhibited. When the TPase is not inhibited, the crosslinking could somehow help and accelerate the PG chains synthesis by the GTase. The entire signal change should be because of both GTase and TPase activities. The authors mentioned that the first slow phase reflects the GTase activity and the second fast phase reflects the TPase, but I do not find it convincing.

a) Please define which section is slow and which section is fast in the FRET curves-in the presence of LpoB, it does not appear to have a slow-phase if the initial lag phase of rise is the slow-phase the authors mentioned.

b) In the presence of Amp, the slope of the initial rise appears to be slower than that in the absence of Amp, which, could be the reason that the authors argue that TPase help accelerate GTase. Please provide absolute quantifications using FRET efficiency to show that this is indeed the case.

c) Can the authors add D,D-endopeptidase in the system after the end of the reaction to uncrosslink the peptides and see how much FRET signal remains, which should be contributed by the pure GTase activity? This signal could be compared with that in the presence of Amp to examine the authors' claim.

3) The lag of TPase activity behind GTase activity as shown in Figure 1F is intriguing. The lag appears to be on the order of 5-10 min in this system and suggest that the GTase and TPase activities are not synchronous. Does this mean that even though PBP1B has both activities in one single molecule, but the two reactions are catalyzed asynchronously by two different molecules?

4) The authors suggest that the decrease of FRET signal in Figure 2C, E and G is due to additional changes of fluorophore's spectroscopic properties. If so why such changes were not observed in the detergent or SLB experiments, if the product is the same? The authors should obtain parallel time points using radio-labeled Lipid II and run the products of both assays on HPLC to show that the decrease of FRET signal in these experiments did not correspond to changed product.

5) The SLB assay has great potentials to allow single-molecule measurement of glycan chain elongation and crosslinking kinetics. It is disappointing, though, to find out that no single-lipid II molecules in the polymerized PG can be visualized. Is this because the lipid II concentration too high on the SLB? Does it need to be that high?

6) To fully drive home the message that the new FRET is better in many ways than previous assays, the authors should compare it with the dansyl-lipid II assay side by side (i.e., not putting the other assay in the supplemental figures) and quantitatively.

7) Similarly, there appeared to be significant differences in FRET signals between the detergent, liposome and SLB assays with or without Amp, and it is difficult to compare the enhancement by LpoB among these assays. Please address these differences.

Reviewer #2:

1) It is stated that: "As expected, ampicillin did not prevent the stimulation of PBP1B by LpoB(sol), which accelerated the FRET increase by 10-20 times with or without ampicillin, consistent with the previously reported stimulation of both GTase and TPase activities." Although other results may suggest TPase activity is stimulated, one cannot draw the conclusion from the FRET assay results here that LpoB stimulates TPase activity – only that TPase activity boosts the FRET signal. There is some confusion in the manuscript about what can and cannot be concluded about mechanism from the FRET assays.

2) The authors use initial rates from the FRET assay to compare effects of LpoB in Figure 1 (and later do the same for other conditions/other activators). Initial rate conditions are used to quantify reaction rates for enzymes because under these conditions substrate depletion and product inhibition do not impact the rate. Initial rates are misleading when comparing a hyperbolic progress curve to a sigmoidal progress curve. The authors should comment on this, clarify the time frame over which the initial rates were measured (perhaps indicated with a figure), and also report the slopes of the "fast" phase of the reactions where activator is not present.

3) There is a pattern of low mass bands in the PAGE gel in Figure 1D for the LpoB reactions; these are barely visible in the reactions without LpoB. At the same time, the glycan strands are shorter in length for the LpoB reaction compared to the reaction without LpoB (lanes 2 and 6). (Shorter glycan strands were observed previously for LpoB - e.g, Paradis-Bleau et al.; Lupoli et al.). I am curious about whether these observations are connected (and therefore whether they are connected to how LpoB achieves activation).

4) The authors comment that the combined assays suggest that the total FRET signal emerges from two steps that have different rates. I recommend either refining or removing this comment. The statement suggests that it might be possible to extract rates for the two steps alluded to from the FRET signal. But there are more than two steps and two rates. For polymerization reactions, initiation and elongation rates are often substantially different and there is evidence in the literature that this is case for PG synthesis. (The slow rate of reaction for the "only labelled Lipid II" reactions does not report on the rate of polymerization with unlabeled Lipid II present because the products are very short – so the polymerization does not "take off" the way it does in the other reactions.) Moreover, there are likely two rates for crosslinking in vitro because the first crosslinks are formed intermolecularly and later crosslinks are formed intramolecularly. Finally, the FRET signal itself is different for uncrosslinked and crosslinked PG – as the authors note, it reaches a higher maximum for the latter.

5) Figure 2C: was the lane 7 sample in the gel the same as the one for which the fluorescence trace is shown? The fluorescence trace looks flat but lane 7 shows polymerization. In other gels, other lanes that show comparable levels of polymerization also show a FRET signal.

6) I already mentioned that the discussion could use some work, but wanted to elaborate. The explanation for how TPase activity increases the FRET signal is plausible and supported by the data. But the section on "coupled reactions in Class A PBPs" is confusing because it is not clear what the authors are trying to accomplish with it. There has been a lot of debate about what LpoB does to activate PBP1b and how it does it. That LpoB affects both GTase activity and TPase activity has been shown repeatedly to one degree or another, and the idea that one activity influences the other in this context has been discussed explicitly going back at least to 2014 (Egan et al., 2014; Lupoli et al., 2014). The authors discuss the idea again here, evidently in response to a recent paper (Catherwood et al.) that attributes LpoB's essentiality to its effects on TPase activity. They conclude that Catherwood's conditions may have overemphasized the TPase effect. I don't think arguing this point is the most important use of the discussion space. The authors should clarify how they think about what the FRET assays show with respect to mechanism of Lpo activators and try to connect their work to a path forward.

Summary: there is a lot of nice material in this manuscript and I believe the authors are pushing the field ahead with the FRET assays, particularly their use in SLMs. However, the analysis of the results is not at the same level as the technical innovations and this should be improved. Mainly text revisions could make the paper suitable for eLife, but it will take some work on the part of the authors.

Reviewer #3:

This manuscript describes studies of generation of improved in vitro assay systems for peptidoglycan synthesis, using liposomes and supported lipid bilayers. This work can be very influential as this synthetic process is a primer candidate for the development for additional antibiotics, and the detection methods utilized are potentially adaptable to high-throughput screens.

Overall the paper is well written and the data are convincing. I have a few comments below, primarily concerning the data presentation.

1) Figures 1A, 2B, and 5: These figures show some of the fluorescently-labeled peptides as being involved in crosslinks. I do not think there is evidence in this paper that these substrates are ever used by transpeptidases. They do not appear to show up in obvious radioactively-labeled HPLC peaks. Doesn't this suggest that they are NOT being crosslinked? All off the crosslinked material seen in HPLC peaks is simply the radiolabeled Lipid II.

2) Figure 1C: I don't understand why the slope with LpoB is greater in the presence of Amp? Can the authors comment?

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

As the editors have judged that your manuscript is of interest, but as described below that additional experiments are still required before it is published. We would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). First, because many researchers have temporarily lost access to the labs, we will give authors as much time as they need to submit revised manuscripts. We are also offering, if you choose, to post the manuscript to bioRxiv (if it is not already there) along with this decision letter and a formal designation that the manuscript is "in revision at eLife". Please let us know if you would like to pursue this option. (If your work is more suitable for medRxiv, you will need to post the preprint yourself, as the mechanisms for us to do so are still in development.)

Major comments:

The major advantages of the assay are three-fold, (1) it is a real-time; (2) it is applicable to membrane-embedded enzymes, and (3) it can monitor both GTase and TPase. The revised manuscript has provided evidence for all the three with improved clarity, new control experiments, and better FRET efficiency calculations, but it still lacks the rigor in the quantitative (or semi-quantitative) aspect of the assay. Specifically, the authors need to provide evidence that the F_acceptor/F_donor ratio indeed reflects proportionally to either just the GTase activity or a combined GTase and TPase activity. The authors did show qualitatively that this ratio is altered when the activator LpoB (or P) or drugs are used, but it is difficult to use this ratio to compare quantitatively (or semi-quantitatively) the rates of different reactions catalyzed by different enzymes, or the same enzyme but with different mutations and regulators. Being able to do so will greatly enhance the impact of the FRET assay and promote its wide adoption in the field.

It appears that the measured F_acceptor/F_donor ratio is not only dependent on the GTase and TPase activities of PBP1B, but some other effects that influence the fluorescence of the dyes. These other effects will make it difficult to use the FRET assay to quantitatively compare different reactions. For example, in Figure 3C, the reaction with Amp (VII) barely had any signal change in F_acceptor/F_donor, but the corresponding gel lane cleared showed polymerized, linear PG strands, suggesting that the polymerization reaction is not reflected in the fluorescence ratio measurement. The rise and decay of the +LpoB/Amp (III) reaction also look strange-this reaction had a similar PG ladder on the gel as reaction (-LpoB/Amp)VII, but the fluorescence ratio measurements are drastically different. Another example is the comparison between the three PBP1Bs: it appears that crosslinking activities have different contributions to the fluorescence ratio measurement: ampicillin completely abolished the fluorescence ratio signal for PBP1BEc but not for PBP1BAb. It is hard to imagine that these highly homologous enzymes would have very different mechanisms to synthesize PG.

A likely explanation for these different fluorescence signals in these reactions is probably what the authors suggested, that some spectroscopic properties of the dyes changed due to the incorporation and/or local environment. Another possibility is that these enzymes of different species may handle the dye-labeled LipidII differentially, one PBP1B may have more difficulty in accepting the dye-labeled LipidII than the other PBP1B. Yet another possibility could also be that the stimulation effect of LpoB is simply that now more dye-labeled LipidII v.s. unlabeled Lipid II can be incorporated, but not necessarily the rate at which Lipid II is incorporated.

It is understandable that these in vitro experiments are extremely challenging, but fluorescence measurements are sensitive and prone to artifacts, and the authors should perform control experiments to make sure that the measured signal indeed reflects the desired enzymatic activities. In particular, in calculating the fluorescence ratio, the donor-only and acceptor-only controls need to be run simultaneously to determine how much fluorescence change is due to the local environment change, and how much is due to FRET. The authors have performed these control experiments but only used them for the end-point measurements. These controls should also be performed for the real-time measurements (with LpoB, with or without Amp), which could help to explain some strange changes in the measured fluorescence ratio (such as that in Figure 3C).

The goal, hopefully, is that one can confidently interpret the ~ 25-fold higher initial slope of PBP1BEc with LpoB in the presence of ampicillin as a 25-fold (or proportionally) higher GTase activity of PBP1BEc stimulated by LpoB (such as that in Figure 3D). Similarly, one can use the smaller fold-increase of the initial slopes in the cases of PBP1BAb and PBP1BPa in the presence of their cognate activators to draw the conclusion that these activators do not stimulate these PBP1Bs to the same extent as that of PBP1BEc. If the fold enhancement of the initial slopes by LpoB in the presence and absence of ampicillin are similar to each other, one perhaps can say that here LpoB only stimulates the GTase activity, and so on and so forth.

The new FRET assay bears great potential in providing a sensitive and quantitative way to characterize both GTase and TPase activities in ways not possible before. The Vollmer laboratory also has the expertise to make this assay hugely impactful for the field. These experiments can become powerful, but they need critical controls.

eLife. 2021 Feb 24;10:e61525. doi: 10.7554/eLife.61525.sa2

Author response

Summary:

The manuscript by Hernández-Rocamora et al. describes a FRET-based real time assay that allows for continuous monitoring of peptidoglycan synthesis by aPBPs. The authors examine how the assay performs in detergent and in liposomes; they also assess whether the assay will be useful for studying regulators of peptidoglycan biosynthesis enzymes by applying it to previously studied lipoprotein activators of aPBPs, and a new lipoprotein activator that is similar to a previously studied one. The major advantage of the current assay in comparison to the original dansylated lipid II assay is that the FRET assay can monitor both GTase and TPase activities. Finally, the current FRET assay has the potential to become single-molecule study-compatible. The Vollmer lab is one of only a few that can carry out studies of PG biosynthetic enzymes, and he has teamed up with an expert in microscopy on this manuscript. The experiments are technically challenging and generally well done and the technical advances should be useful to the field. However, new biological/mechanistic insights are few. The work also needs to be significantly strengthened with more critically analyzed data, control experiments and better writing.

We thank the reviewers for their positive assessment of our manuscript and acknowledging the usefulness of the assay. We would like to stress that, in addition to being able to monitor both reactions, the other major advantage of the new assay is that it is capable of measure PG synthesis with membrane-reconstituted enzymes. The dansylated lipid II assay does not work with liposomes or supported bilayers. We are also grateful for the insightful points made by the reviewer. Based on these points we critically analyzed the data, added the suggested control experiments and improved the writing of the manuscript. We hope that the reviewers agree with us that these changes have significantly improved the manuscript.

Reviewer #1:

1) FRET controls and calculations:

a) The authors should provide donor-only and acceptor-only controls under the same conditions for all experiments to allow the calculation of FRET efficiency quantitatively (preferably using the Ratio A method Clegg RM, Methods in Enzymology, 1992, 211, p353). This quantification is important because one major potential advantage of this method could be studying the kinetics of GTase and TPase activities, which may offer mechanistic insight on the enzyme's working. The Facceptor /Fdonor ratio measurement is convenient and can be useful for screening of a large sample size but is prone to artifacts such as fluorescence quenching due to reasons other than FRET, especially when the way the system is set up does not allow precise determination of the donor/acceptor ratio and attachment positions. These donor only or acceptor only experiments are also important for the FRAP measurement in the SLB assay.

The reviewer is correct in that we did not provide donor-only and acceptor-only controls to calculate FRET efficiency quantitatively. To address this point, we have now measured apparent FRET efficiencies using the suggested ratio A method for the end products of the reactions of E. coli PBP1B in detergents in the presence ampicillin or moenomycin, or without antibiotic. We measured samples containing only lipid II-Atto647n+lipid II (acceptor reference) and samples containing only lipidII-Atto550+lipid II (donor reference). With these data we demonstrate that signals arising in these reactions are indeed due to FRET, and we provide the values for FRET efficiencies. The new data shown in Figure 1B-D.

We respectfully question the reviewer suggestion to calculate FRET efficiencies when monitoring reactions in real time, for several reasons. First, FRET efficiencies will not be useful to provide any meaningful distances between fluorophores, because probe molecules are unlikely to be positioned homogeneously within the PG produced. Very likely, probe molecules are separated by variable distances and with a range of different orientations, giving rise to a mixture of FRET between probes from the same glycan chain and different chains.

Second, calculating FRET efficiency in real time would require combining signals arising from different reactions in different wells (the references with donor-only or acceptor-only plus the sample with both) which, in practice, would substantially increase the noise in the final reaction curve. This problem would be exacerbated in the membrane system, which would require to use different liposome preparations for the different samples (because the labelled lipid versions need to be reconstituted together with the enzyme).

We believe that the F_acceptor/F_donor ratio is useful even if a small part of the signal might come from quenching. We demonstrated that the F_acceptor/F_donor ratio monitors PG synthesis reactions, because the rise in the ratio requires the incorporation of the probes into peptidoglycan, as verified by the controls with ampicillin and moenomycin.

To measure FRET within a PG layer on a supported lipid bilayer, we performed acceptor photobleaching or donor dequenching experiments, where the energy transfer from the donor to the acceptor is irreversibly blocked and its intensity is increased. The relative increase in donor fluorescence can be used a direct readout for the FRET efficiency as photobleaching of the acceptor dye is the only process that contributes to the recorded increase in the donor fluorescence signal (see for example Verveer et al., 2006). In rare cases, an increase in donor fluorescence can be recorded even in the absence of acceptor dye, which could result in an overestimation of the FRET efficiency. To our knowledge, this has so far only been found when fluorescent proteins were used (Karpova et al., 2003). Additionally, the donor signal could appear brighter due to spectral crosstalk. In this the FRET signal would be underestimated as the relative intensity increase after acceptor bleaching is decreased.

To test for these two opposing effects, we performed several control experiments. First, in the presence of Ampicillin, there was no increase in donor intensity and the calculated FRET efficiency was 0 (Figure 5B). Second, the signal was dramatically reduced after treatment with cellosyl and when the PBP1B E233Q mutant was used (Figure 5B-C). Additionally, we now also performed a control experiment, where only Lipid II Atto550 and Atto647n, and not PBP1B, are present in the assay (see Author response image 1). Also under these conditions, the donor intensity was unchanged upon acceptor bleaching and the calculated FRET efficiency was therefore 0. We therefore conclude that there is no contribution of acceptor fluorescence in the donor channel in the absence of PG synthesis.

Author response image 1. — After photobleaching of the acceptor dye (Att647N), we did not observe an donor increase in intensity in the bleached area.

Together with the controls performed in our spectroscopic assays in solution, we are confident that the increase in donor intensity was soley due to the activity of PBP1B and not caused by an unidentified photophysical process or spectral crosstalk. We added a comment on this issue to the revised text.

b) In Figure 4 the authors used a ratiometric FRET equation Erel = IA/(IA+ID) in the semi-single-molecule assay to determine FRET efficiency. This measurement should be unified to allow direct comparison of FRET between different types of experiments (solubilized, liposomes, SLBs, bulk or single-molecule). As mentioned above, the acceptor intensity consists of the intensities from donor-excitation (FRET), direct-excitation, and bleed-through from the donor. Donor emission could be quenched due to reasons (such as the local environment as that in the dansyl-lipid II assay) other than FRET. More control experiments will allow the authors to determine how this impact the true acceptor signal resulting from FRET.

The reviewer is correct that a different equation was used to calculate FRET efficiency 4 than for detergents or liposomes. However, the equation we used is not the one indicated by the reviewer but:

(1) E = (ID,pb – ID) / ID,pb = 1 – (ID/ID,pb)

Where ID,pb is the donor intensity before photobleaching and ID,pb is the donor intensity after photobleaching (Verveer et al., 2006). This equation uses the donor fluorescence to calculate FRET, the acceptor fluorescence is not relevant in this calculation.

Due to the very low concentration and density of PBP enzyme on the supported bilayer the comparison of reaction rates from these experiments with the ones from liposomes or detergents is not meaningful. For a description of our controls performed, please refer to our answer to question 1a

c) There appears to be a major difference in the fluorescence intensity spectra between the detergent-solubilized (Figure 1—figure supplement 1C) and liposome experiments (Figure 2—figure supplement 2A, C, D). In the detergent solubilized experiments, there appears to be little change in the intensity of the donor peak from PBP1B to +LpoB. However, in the liposome experiments the LpoB has a drastic loss of donor signal compared to that of PBP1B. This is another reason why calculating FRET efficiency instead of the relative ratio would allow for a better comparison between the experiments. This drastic loss of donor signal in the liposome experiments is seen in all 3 species. The authors should address why there is such a difference in the FRET profiles between these two experiments.

The activity of PBP1B without LpoB is lower in liposomes than in detergents, hence the difference is more prominent in the end-point spectra of PBP1B versus PBP1B+LpoB in liposomes. We now discuss this difference in the revised text. We also added further comments on the difference between the behaviour of PBPs in the membrane and in detergents to Discussion.

d) The authors rely on "Relative Slope" to show how the addition of LpoB impacts the PBP1B activity. Is the "Relative slope" calculated by normalizing the slope of a reaction with LopB to that without LopB? How is the time window determined? As this ratio does not really inform on the enzyme activity, it would be difficult for other groups to use the same method and to compare to this metric. It is also difficult to judge whether the GTase is faster or slower in the presence of TPase activity without the absolute values. Please provide the absolute measurement of the slope (or the initial velocity V0) using a better determined FRET efficiency value as in point 1a.

We thank the reviewer for making these points. The relative slope is indeed the ratio between the slope in the presence of LpoB and the slope without LpoB under the same conditions. This method has been used when measuring the activation by LpoB using the dansyl-Lipid II method to measure GTase activity (e.g. Egan et al., 2014, 2018) and we used this method in order to compare our results with those from the literature. We relabelled the plots from “Relative slope” to “Fold change in initial slope” to better describe them.

The slopes were calculated in the linear part of the curve after the signal started to increase. We now explain in the main text how slopes were calculated and provide more details in the experimental section. The absolute values for the slopes and the time window used to calculated them for each condition are now listed in the source files for the corresponding figures (Figures 1, 3, and Figure 3—figure supplement 5, Figure 3—figure supplement 6 and Figure 3—figure supplement 7).

e) Additionally, perhaps the authors could determine the reaction velocities at different substrate concentrations to see if Michaelis-Menten kinetics could be applied? A reanalysis of the data with better quantifications with regard to enzyme's kinetics would benefit the community significantly by providing a more complete picture of enzyme's activity rather than only comparing PBP1B alone to +LpoB.

To determine enzyme kinetics parameters was not our goal when developing this method although, in theory, the method should be useful for this purpose. Searching the literature we noticed that the kinetics of class A-PBP GTase and TPase activities have only been quantified separately, never both reactions together. We believe that developing a precise quantitative theoretical model for both activities is out of the scope of this paper and would require extensive new experiments and modelling.

We also noticed that the kinetics of the individual activities of E. coli PBP1B have been reported in the literature, but only for the detergent solubilized enzyme. Schwartz et al. (2002) established that glycosyltransferase activity followed Michaelis-Menten kinetics in the presence of the detergent decyl-PEG using an HPLC assay (Schwartz et al., 2002). Catherwood et al. (2020) used a D-Ala release assay to determined that PBP1B transpeptidase did not follow Michaelis-Menten in reactions with lipid II (mDAP) – same as here – and in the presence of Triton X-100. We now discuss these issues.

2) According to the authors' data and others' work, PBP1B can only synthesize short, uncrosslinked PG chains when its TPase is inhibited. When the TPase is not inhibited, the crosslinking could somehow help and accelerate the PG chains synthesis by the GTase. The entire signal change should be because of both GTase and TPase activities. The authors mentioned that the first slow phase reflects the GTase activity and the second fast phase reflects the TPase, but I do not find it convincing.

We would like to respectfully correct the statement made by the reviewer in the first sentence. TPase-inhibited PBP1B can indeed synthesise long chains (e.g., lane 2 on Figure 2C). The reviewer might have confused this fact with the results of control samples performed with only fluorescently-derivatised lipid II analogues (right panel in Figure 1E and lanes 4 and 8 in Figure 1—figure supplement 3B). We discuss that the binding of two labelled lipid II molecules to the two binding sites (donor and acceptor) in the GTase domain of PBP1B is probably hindered due to the big size of the Atto550 and Atto647n probes. Hence, in the absence of unlabelled lipid II, PBP1B on its own is unable to form glycan chains, but with LpoB PBP1B is stimulated to produce short glycan chains under these conditions.

We have now removed the statement cited by the reviewer, which was confusing and only intended for PBP1B reactions without LpoB. We did not mean to imply that all FRET signal was due to transpeptidation and only stated that transpeptidation was necessary for a substantial increase in FRET signal. Our model is that FRET arises from glycosyltransferase reactions (intra-chain FRET) and transpeptidation (inter-chain FRET), and the second contribution is higher in the absence of LpoB.

a) Please define which section is slow and which section is fast in the FRET curves-in the presence of LpoB, it does not appear to have a slow-phase if the initial lag phase of rise is the slow-phase the authors mentioned.

As the reviewer comments, there is a lag phase in the increase in FRET in reactions with only PBP1B, but this disappears in the presence of activator, LpoB (Figure 1E, left panel). This lag phase occurs in reactions with or without ampicillin (Figure 1E, middle panel) and it has been previously observed with the dansyl-lipid II method when measuring glycosyltransferase rates (Schwartz et al., 2002). The suppression of the lag phase by LpoB was also observed using the dansyl-lipid II method (Egan et al., 2014). Thus we hypothesize that LpoB accelerates the initiation of GTase, in addition to the rate of glycan chain elongation. In our case, we also have to take into account that LpoB allows PBP1B to incorporate two labelled lipid II molecules consecutively, as demonstrated in the controls with only Atto550- and Atto647n-labelled lipid II (Figure 1E, right panel). Thus, LpoB most likely enhances the intra-chain FRET, consistent with the observation that LpoB increased not only the reaction slope but also the final FRET signal in reactions with ampicillin when only intra-chain FRET is possible.

b) In the presence of Amp, the slope of the initial rise appears to be slower than that in the absence of Amp, which, could be the reason that the authors argue that TPase help accelerate GTase. Please provide absolute quantifications using FRET efficiency to show that this is indeed the case.

Sorry for the misunderstanding. We did not intend to claim that the TPase helps accelerate the GTase. In the statement we only meant to say that the FRET signal increased rapidly once transpeptidation occur in reactions without activator and without ampicillin. We therefore removed the confusing statement. We address the last sentence made by the reviewer in our reply to the next point.

c) Can the authors add D,D-endopeptidase in the system after the end of the reaction to uncrosslink the peptides and see how much FRET signal remains, which should be contributed by the pure GTase activity? This signal could be compared with that in the presence of Amp to examine the authors' claim.

Again, we did not intend to claim that the TPase helps accelerate the GTase. Our model was and still is that the in the absence of activator, inter-chain FRET is a major contributor to the FRET signal. This is now demonstrated in new data we present in the new Figure 1B-D in which we show that the addition of ampicillin (which blocks cross-link formation) causes a decrease of FRET efficiency to 30% of the value without antibiotic, and this result is commented in the revised text.

In addition, we carried out the experiment with a DD-endopeptidase, as suggested by the reviewer, and shown these new data in the new Figure 2A-B. We used the endopeptidase MepM to digest PG produced in the presence or absence of ampicillin. We found that MepM was unable to digest all cross-links in Atto-labelled PG, but caused a substantial reduction in the cross-linked PG (compare lanes M in the presence or absence of ampicillin on new Figure 2A). Consistent with this result, MepM digestion of the labelled PG reduced the FRET efficiency by 2-fold while, as expected, MepM did not significantly affect the FRET efficiency of the uncross-linked PG synthesised in the presence of ampicillin (new Figure 2D).

3) The lag of TPase activity behind GTase activity as shown in Figure 1F is intriguing. The lag appears to be on the order of 5-10 min in this system and suggest that the GTase and TPase activities are not synchronous. Does this mean that even though PBP1B has both activities in one single molecule, but the two reactions are catalyzed asynchronously by two different molecules?

Our results or published data do not suggest that both activities are catalysed “asynchronously”. Rather, PBP1B requires ongoing GTase reactions for efficient TPase activity (Bertsche et al., 2005) and Catherwood et al. (2020) recently demonstrated that PBP1B uses peptides in the growing glycan chain synthesised by the GTase domain as TPase donors (i.e. they link the D-Ala in fourth position in the pentapeptides of the growing chain to the mDAP group in an acceptor chain) (Catherwood et al., 2020).

One might expect a short lag between the start of the GTase and the beginning of the TPase because the growing chain has to grow at least to ~8 subunits to reach the TPase site (Sung et al., 2009). However, the experiment of Figure 1F (new Figure 2E) is not suited to measure a possible lag phase because of the sampling rate (every 5 min) and the possibility that, at the beginning of the reaction, small peaks are missed because of the detection limit and the inherent noise of radioactivity detection in chromatography. For example, in Figure 1F (new Figure 2E) at 5 min, the area of the Penta peak is only 459 CPM (over a peak width of 0.8 min) and a possible smaller TetraPenta peak would be barely detectable over the signal noise, which is ~90 CPM/min.

4) The authors suggest that the decrease of FRET signal in Figure 2C, E and G is due to additional changes of fluorophore's spectroscopic properties. If so why such changes were not observed in the detergent or SLB experiments, if the product is the same? The authors should obtain parallel time points using radio-labeled Lipid II and run the products of both assays on HPLC to show that the decrease of FRET signal in these experiments did not correspond to changed product.

Sorry, we are confused by the reviewer's question. We only see a downwards slope in the FRET curve of PBP1B reactions with LpoB and ampicillin in liposomes (old Figure 2C, middle panel), not in old Figure 2E or Figure 2G. We speculate in the text that this decrease could be due to the influence of the membrane on the fluorescence of the probe. If our speculation is correct, this decrease should not be observed on SLBs because due to the very low lipid II concentrations in those experiments, little PG is produced and, as inter-chain FRET is the major contributor to the FRET signal, very little or no FRET signal is observed with ampicillin. In assays with detergents there is no membrane that can affect fluorescence properties and, consistent with our speculation, we did not observe a negative slope (new Figure 1E, middle panel).

5) The SLB assay has great potentials to allow single-molecule measurement of glycan chain elongation and crosslinking kinetics. It is disappointing, though, to find out that no single-lipid II molecules in the polymerized PG can be visualized. Is this because the lipid II concentration too high on the SLB? Does it need to be that high?

The reviewer is correct that lipid II concentrations are too high in SLB for single molecule analysis, and this is so because concentrations low enough for single molecules do not support PG synthesis. We agree with the reviewer that single-molecule measurements of glycan chain elongation and cross-link formation is a worthy goal. We are working towards this goal but it is still a big step from where we are now with the current technology, which we believe already provides a major advance.

6) To fully drive home the message that the new FRET is better in many ways than previous assays, the authors should compare it with the dansyl-lipid II assay side by side (i.e., not putting the other assay in the supplemental figures) and quantitatively.

Both EcPBP1B and PaPBP1B have been studied by dansyl-lipid II assay in the literature (Schwartz et al., 2002, Egan et al., 2014, Caveney et al., 2020) and we did not repeat those published assays here. We performed the dansyl-lipid II assay for AbPBP1B and included it into the ms, because there was no prior information about this enzyme in the literature. We did not mean to claim that our assay is superior to the dansyl-lipid II because it provides better data, but because it allows to monitor transpeptidation and glycosyltransferase activity of the membrane-reconstituted enzymes (as is stated in the Discussion). Where relevant, we compared our results from the FRET assay with prior results in the literature obtained with dansyl-lipid II, i.e. when comparing the activation of the GTase rate by LpoB and LpoP regulators (“Coupled reactions in class A PBPs” section in the Discussion).

7) Similarly, there appeared to be significant differences in FRET signals between the detergent, liposome and SLB assays with or without Amp, and it is difficult to compare the enhancement by LpoB among these assays. Please address these differences.

The conditions in SLB are not comparable to the conditions in liposomes and detergents, due to the low density of PBP1B in SLBs. This difference is discussed in the text (Results section on SLBs). As for the comparison between liposomes and detergents, we may expect differences in behaviour of enzymes because of the following reasons: (1) possible effects of detergents on enzyme activity and (2) kinetics of enzymatic reactions occurring on the membrane are different from reactions occurring in solution. We comment about the specific difference in behaviour of the enzymes in membranes in the Discussion.

Reviewer #2:

1) It is stated that: "As expected, ampicillin did not prevent the stimulation of PBP1B by LpoB(sol), which accelerated the FRET increase by 10-20 times with or without ampicillin, consistent with the previously reported stimulation of both GTase and TPase activities." Although other results may suggest TPase activity is stimulated, one cannot draw the conclusion from the FRET assay results here that LpoB stimulates TPase activity – only that TPase activity boosts the FRET signal. There is some confusion in the manuscript about what can and cannot be concluded about mechanism from the FRET assays.

We thank the reviewer for this point. We agree that the FRET assay only does not allow the conclusion about TPase rate. Some activators enable PBPs to polymerise Atto-lipid II without unlabelled lipid II (thus reducing the distance between probes within the chains and increasing intra-chain FRET), hence we cannot conclude from FRET results alone that the activators stimulate the TPase. We have carefully edited the text to avoid this inaccurate statement.

2) The authors use initial rates from the FRET assay to compare effects of LpoB in Figure 1 (and later do the same for other conditions/other activators). Initial rate conditions are used to quantify reaction rates for enzymes because under these conditions substrate depletion and product inhibition do not impact the rate. Initial rates are misleading when comparing a hyperbolic progress curve to a sigmoidal progress curve. The authors should comment on this, clarify the time frame over which the initial rates were measured (perhaps indicated with a figure), and also report the slopes of the "fast" phase of the reactions where activator is not present.

The slopes were calculated from the linear part of the curve when the signal started to increase after the lag phase (when present). For example, for reactions with detergent-solubilised E. coli PBP1B, we measured the slope from 10-20 min without LpoB (after the lag phase), but from 0-1.3 min with LpoB (no lag). The calculation of initial rates is now explained briefly in the Results text and in detail in the Materials and methods section. We now provide the absolute values for the slopes and the time windows used to calculate them for each condition in the source files for the corresponding figures (new Figures 1, 3, and Figure 3—figure supplement 5, Figure 3—figure supplement 6 and Figure 3—figure supplement 7). We have also relabelled the figures as “Fold change in initial rate” to avoid confusion.

3) There is a pattern of low mass bands in the PAGE gel in Figure 1D for the LpoB reactions; these are barely visible in the reactions without LpoB. At the same time, the glycan strands are shorter in length for the LpoB reaction compared to the reaction without LpoB (lanes 2 and 6). (Shorter glycan strands were observed previously for LpoB - e.g, Paradis-Bleau et al.; Lupoli et al.). I am curious about whether these observations are connected (and therefore whether they are connected to how LpoB achieves activation).

We thank the reviewer for pointing to this observation. We would like to be hesitant to use the relatively crude method of SDS-PAGE to compare smaller differences in glycan chain lengths, as they might occur between samples with or without LpoB. Egan et al. (2018) observed an increase in the heterogeneity of chain lengths produced by PBP1B in the presence of LpoB, contradicting the results by Paradis-Bleu et al. and Lupoli et al. (Egan et al., 2018). Egan et al. also provided evidence that the solvent demethyl sulphoxide (DMSO) used in high concentration in the other studies, reduced the PBP1B-LpoB affinity and might itself has an effect on glycan chain length distribution. We believe it is outside the scope of the current manuscript to clarify this issue.

4) The authors comment that the combined assays suggest that the total FRET signal emerges from two steps that have different rates. I recommend either refining or removing this comment. The statement suggests that it might be possible to extract rates for the two steps alluded to from the FRET signal. But there are more than two steps and two rates. For polymerization reactions, initiation and elongation rates are often substantially different and there is evidence in the literature that this is case for PG synthesis. (The slow rate of reaction for the "only labelled Lipid II" reactions does not report on the rate of polymerization with unlabeled Lipid II present because the products are very short – so the polymerization does not "take off" the way it does in the other reactions.) Moreover, there are likely two rates for crosslinking in vitro because the first crosslinks are formed intermolecularly and later crosslinks are formed intramolecularly. Finally, the FRET signal itself is different for uncrosslinked and crosslinked PG – as the authors note, it reaches a higher maximum for the latter.

We thank the reviewer for this point. We agree that our statement was confusing and we removed it from the manuscript. We thank the reviewer also for their comments on the different rates for GTase and TPase. We have added a more extensive discussion on GTase and TPase rates of class A PBPs, citing the relevant literature, and commenting on what can be concluded from our data on this topic in the rewritten “Coupled reactions in class A PBPs and their activation” section of the Discussion.

5) Figure 2C: was the lane 7 sample in the gel the same as the one for which the fluorescence trace is shown? The fluorescence trace looks flat but lane 7 shows polymerization. In other gels, other lanes that show comparable levels of polymerization also show a FRET signal.

We double checked the gels and the lanes are the same. Although there are some short glycan chains on lane 7, most of the lipid II remained unpolymerised. Besides, for FRET to occur in the absence of crosslinking more than one label has to be incorporated per chain and they should be sufficiently close together. The very low FRET thus indicates that very few probes are incorporated per chain at distances close enough to produce FRET.

6) I already mentioned that the discussion could use some work, but wanted to elaborate. The explanation for how TPase activity increases the FRET signal is plausible and supported by the data. But the section on "coupled reactions in Class A PBPs" is confusing because it is not clear what the authors are trying to accomplish with it. There has been a lot of debate about what LpoB does to activate PBP1b and how it does it. That LpoB affects both GTase activity and TPase activity has been shown repeatedly to one degree or another, and the idea that one activity influences the other in this context has been discussed explicitly going back at least to 2014 (Egan et al., 2014; Lupoli et al., 2014). The authors discuss the idea again here, evidently in response to a recent paper (Catherwood et al.) that attributes LpoB's essentiality to its effects on TPase activity. They conclude that Catherwood's conditions may have overemphasized the TPase effect. I don't think arguing this point is the most important use of the discussion space. The authors should clarify how they think about what the FRET assays show with respect to mechanism of Lpo activators and try to connect their work to a path forward.

We thank the reviewer for these points. Based on the reviewer's suggestions we have extensively rewritten the part about “Coupled reactions in Class A PBPs” in the Discussion. We now discuss the effects of Lpo activators on activities of PBPs more broadly, with an emphasis to what our results can add to the ongoing discussions. In particular, we discuss the importance of assaying PBPs in the membrane environment based on our findings that LpoB is required for the activities of several PBPs we studied, when they were reconstituted into the membrane. We also discuss that this observation agrees with the absolute requirement of LpoB for the function of PBP1B in the cell.

Reviewer #3:

This manuscript describes studies of generation of improved in vitro assay systems for peptidoglycan synthesis, using liposomes and supported lipid bilayers. This work can be very influential as this synthetic process is a primer candidate for the development for additional antibiotics, and the detection methods utilized are potentially adaptable to high-throughput screens.

Overall the paper is well written and the data are convincing. I have a few comments below, primarily concerning the data presentation.

1) Figures 1A, 2B, and 5: These figures show some of the fluorescently-labeled peptides as being involved in crosslinks. I do not think there is evidence in this paper that these substrates are ever used by transpeptidases. They do not appear to show up in obvious radioactively-labeled HPLC peaks. Doesn't this suggest that they are NOT being crosslinked? All off the crosslinked material seen in HPLC peaks is simply the radiolabeled Lipid II.

We thank the reviewer for this point. Although it is technically possible that fluorescently-labelled pentapeptides participate as donors in the transpeptidase reaction, as depicted in our schemes, we found no evidence for this to happen in the HPLC assay. Thus, we have modified the schemes in the cited figures and now avoid showing Atto-labelled pentapeptides participating directly in cross-links.

2) Figure 1C: I don't understand why the slope with LpoB is greater in the presence of Amp? Can the authors comment?

In Figure 1C (Figure 1F in the revised manuscript) we present the ratio between the slope of reactions with or without LpoB (relative slope = slope LpoB / slope no activator). Hence, Figure 1C indicates that the presence of LpoB causes a larger change in reaction slopes in the presence of ampicillin than in its absence. This was the same for all pairs of PBP1B-activator pairs studied, except for AbPBP1B/AbLpoP in detergents. The slope changes produced by activators in the presence of ampicillin reflect the acceleration of the GTase, while the slope changes produced in the absence of antibiotic reflect the combined effect of regulators on GTase and TPase rates. As we cannot separate the TPase and GTase rates from the curves without ampicillin, it is difficult to know what exactly causes this difference. In the revised manuscript we discuss the challenge to separating the effect of an Lpo activator on the GTase rates from the effect on the TPase rates.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Major comments:

The major advantages of the assay are three-fold, (1) it is a real-time; (2) it is applicable to membrane-embedded enzymes, and (3) it can monitor both GTase and TPase. The revised manuscript has provided evidence for all the three with improved clarity, new control experiments, and better FRET efficiency calculations, but it still lacks the rigor in the quantitative (or semi-quantitative) aspect of the assay. Specifically, the authors need to provide evidence that the F_acceptor/F_donor ratio indeed reflects proportionally to either just the GTase activity or a combined GTase and TPase activity. The authors did show qualitatively that this ratio is altered when the activator LpoB (or P) or drugs are used, but it is difficult to use this ratio to compare quantitatively (or semi-quantitatively) the rates of different reactions catalyzed by different enzymes, or the same enzyme but with different mutations and regulators. Being able to do so will greatly enhance the impact of the FRET assay and promote its wide adoption in the field.

We thank the reviewers for appreciating the advantages of the new assay. The control experiment shown in Figure 2AB and other experiments with inhibitors (Figures 1E; Figure 3C,E,G; Figure 3—figure supplement 7A, Figure 3—figure supplement 7B), demonstrate that the increase in F_acceptor/F_donor ratio is proportional to the GTase or GTase-TPase rates. However, it is currently not possible to precisely quantify their respective contributions (see our responses below).

We agree with the reviewers that we cannot provide quantitative comparison of the catalytic rates of different enzyme based only on FRET increase. This is due to an inherent limitation of using (fluorescent) modified substrates in an enzymatic assay: different enzymes can potentially have different rates for utilising those substrates. We added a paragraph about the limitations of the assay to Discussion.

As for the effect of activators, we performed a new experiment, added the data to the manuscript (Figure 2—figure supplement 1) and present these in the section “Intra-chain versus Inter-chain FRET”. These data clarify the effect of activators on the final FRET signal and on the slopes of the reaction curves.

With the appropriate control reactions (β-lactams, using only labelled lipid II) it was possible to determine in every case whether the FRET signal follows more closely the GTase (intra-chain FRET) or the TPase (inter-chain FRET) activities.

In conclusion, we are convinced that our novel FRET assay can be used to measure PG synthesis activity of a wide variety of enzymes and that it offers a straightforward approach for the biochemical screening of inhibitors and paves the way for detailed mechanistic studies of PG synthases.

It appears that the measured F_acceptor/F_donor ratio is not only dependent on the GTase and TPase activities of PBP1B, but some other effects that influence the fluorescence of the dyes. These other effects will make it difficult to use the FRET assay to quantitatively compare different reactions. For example, in Figure 3C, the reaction with Amp (VII) barely had any signal change in F_acceptor/F_donor, but the corresponding gel lane cleared showed polymerized, linear PG strands, suggesting that the polymerization reaction is not reflected in the fluorescence ratio measurement. The rise and decay of the +LpoB/Amp (III) reaction also look strange-this reaction had a similar PG ladder on the gel as reaction (-LpoB/Amp)VII, but the fluorescence ratio measurements are drastically different. Another example is the comparison between the three PBP1Bs: it appears that crosslinking activities have different contributions to the fluorescence ratio measurement: ampicillin completely abolished the fluorescence ratio signal for PBP1BEc but not for PBP1BAb. It is hard to imagine that these highly homologous enzymes would have very different mechanisms to synthesize PG.

We agree that the results with PBP1B in the membrane require careful controls and that the interpretation is more difficult in part due to the lower activity of membrane-reconstituted PBP1B^Ec in the absence of LpoB. Therefore, in new experiments we performed reactions of PBP1B^Ec in liposomes for 90 min (instead of 60 min), as for the reactions with PBP1B^Pa and PBP1B^Ab in liposomes. We think this new data shows more clearly the differences in activity between PBP1B^Ec in the membrane with and without LpoB, and we have updated Figure 3C accordingly.

The reviewers correctly observed that the polymerization reaction seen by SDS-PAGE is not always reflected in the fluorescence ratio measurement (FRET). This is true for several reasons:

i) We noticed that the image software automatically adjusts the contrast of files before exporting them, making weaker bands (e.g. bands for short strands) easier to visualize but misrepresenting their contribution to the total signal. We added a section to Materials and methods about fluorescent scanning of gels and their analysis, in which we mention the image processing. For quantification of gel signals in Figure 1 —figure supplement 5 we used the original image files without contrast adjustment. We have now uploaded the original image files for all gels presented as source data files.

ii) Regarding the examples discussed by the reviewers: The reactions with PBP1B^Ec liposomes with ampicillin without LpoB (-LpoB/+Amp, VII in previous Figure 3C, II in current Figure 3C) contain much more unused lipid II in their corresponding gel lanes than reactions in which LpoB was present (+LpoB/+Amp, III in previous Figure 3C, VI in current Figure 3C), consistent with a smaller signal change in F_acceptor/F_donor in the corresponding reactions curves. We investigated the rise and decay of the signal in the +LpoB/+Amp reaction with new experiments (see below, point iv).

iii) As discussed in the section “Intra-chain FRET versus inter-chain FRET”, our data suggests that with LpoB, PBP1B produces PG chains with more densely packed fluorophores. This is consistent with the glycan chains produced in the presence of Ampicillin without LpoB showing less FRET than with LpoB.

iv) We were also puzzled by the decaying FRET signal for +LpoB/Amp condition, which we previously had tested for a single preparation of liposomes. Thus, we repeated the assay at the same conditions (LpoB/Amp) several times with two new liposome preparations and using ampicillin from two different suppliers. We did not consistently observe this decay in FRET during the reactions and instead observed that the reaction at these conditions had two possible outcomes even with the same liposome preparation and ampicillin supplier: (i) (most often): no FRET-decay during reactions and long PG chains produced; (ii) (less often) FRET-decay and short chains at the end of reactions. We cannot currently explain why reactions of PBP1B^Ec liposomes with LpoB/Amp (and only these reactions) can have the two outcomes. It appears that the decay in the FRET signal did not happen because of environmental effect on the probes, but because of different enzyme activity. We have updated the data on Figure 3C and added a comparison of the different possible outcomes of the assay in this particular condition on Figure 3—figure supplement 2A. We comment on these results in section “FRET assay to monitor PG synthesis in liposomes”.

We disagree with the reviewers’ statement that "It is hard to imagine that these highly homologous enzymes would have very different mechanisms to synthesize PG.". Actually, the similarity between different PBP1B homologues is not so high. Sequence comparison shows 33% identity and 45% similarity between PBP1B^Ab and PBP1B^Ec, and 41% identity and 52% similarity between PBP1B^Pa and PBP1B^Ec and, importantly, their activators have different structures. Such differences could be the reason why, for example, ∆lpoP ∆PBP1A is viable in P. aeruginosa (Greene et al. 2018, PNAS 115(12):3150-3155), while ∆lpoB ∆PBP1A is lethal in E. coli (Paradis-Bleau et al., 2010, Cell 143(7):1110-20.), indicating that PBP1B^Pa has higher residual activity in the cell in the absence of activator.

A likely explanation for these different fluorescence signals in these reactions is probably what the authors suggested, that some spectroscopic properties of the dyes changed due to the incorporation and/or local environment. Another possibility is that these enzymes of different species may handle the dye-labeled LipidII differentially, one PBP1B may have more difficulty in accepting the dye-labeled LipidII than the other PBP1B. Yet another possibility could also be that the stimulation effect of LpoB is simply that now more dye-labeled LipidII v.s. unlabeled Lipid II can be incorporated, but not necessarily the rate at which Lipid II is incorporated.

We appreciate the discussion of these possibilities by the reviewers and believe the new controls added to the paper clarify most of their hypothesises:

1) Changes in spectroscopic properties of the dyes due to incorporation to PG or local environment effects: the end point controls with donor-only and acceptor-only as fluorescent lipid II versions show that FRET occurs. In addition, the new real-time controls with donor-only or acceptor-only as fluorescent lipid II version in detergents or liposomes (Figure 1—figure supplement 4 and Figure 3—figure supplement 4) demonstrate that changes in donor or acceptor fluorescence are minimal unless there is PG synthesis and both fluorophores are present in the reaction. Finally, as discussed above, we demonstrate that the FRET decay observed in some reactions with PBP1B^Ec liposomes and LpoB/Amp reflects a change in the type of glycan chains produced (Figure 3—figure supplement 2A).

2) Different enzymes having different reactivity towards labelled lipid II: we always assumed that this can be the case. This is an inherent limitation of any assay that uses fluorescent labelled substrates and is now discussed in the text (Discussion).

3) LpoB might affect the rate of incorporation of dye-labelled lipid II versus incorporation of non-labelled lipid II: It is known that LpoB stimulates the GTase rate with non-fluorescent lipid II (e.g. Figure 4 in Lupoli et al., 2014 J Am Chem Soc. 136:52-55). The new data in Figure 2—figure supplement 1 commented in the section “Intra-chains versus Inter-chain FRET” show that LpoB does not increase the proportion of dye-labelled lipid II that gets incorporated into PG and suggests that the faster FRET increase in the presence of LpoB is not only caused by faster GTase rate but also reflects a more dense packing of the fluorophores in the final PG product. Presumably, LpoB allows PBP1B^Ec to incorporate labelled substrate molecules more densely along the glycan chain.

It is understandable that these in vitro experiments are extremely challenging, but fluorescence measurements are sensitive and prone to artifacts, and the authors should perform control experiments to make sure that the measured signal indeed reflects the desired enzymatic activities. In particular, in calculating the fluorescence ratio, the donor-only and acceptor-only controls need to be run simultaneously to determine how much fluorescence change is due to the local environment change, and how much is due to FRET. The authors have performed these control experiments but only used them for the end-point measurements. These controls should also be performed for the real-time measurements (with LpoB, with or without Amp), which could help to explain some strange changes in the measured fluorescence ratio (such as that in Figure 3C).

As requested by the reviewers and explained above, we incorporated these real-time controls for detergents and liposomes reactions in Figure 1—figure supplement 4 and Figure 3—figure supplement 4, respectively. In both cases, the changes in donor or acceptor fluorescence are minimal unless they are both present in the reaction with active enzyme.

The goal, hopefully, is that one can confidently interpret the ~ 25-fold higher initial slope of PBP1BEc with LpoB in the presence of ampicillin as a 25-fold (or proportionally) higher GTase activity of PBP1BEc stimulated by LpoB (such as that in Figure 3D). Similarly, one can use the smaller fold-increase of the initial slopes in the cases of PBP1BAb and PBP1BPa in the presence of their cognate activators to draw the conclusion that these activators do not stimulate these PBP1Bs to the same extent as that of PBP1BEc. If the fold enhancement of the initial slopes by LpoB in the presence and absence of ampicillin are similar to each other, one perhaps can say that here LpoB only stimulates the GTase activity, and so on and so forth.

We fully agree with the reviewer that an ideal assay would precisely quantify the GTase and TPase rates in detergents and the membrane. However, this was not our goal when we set up the FRET assay and, indeed, by its nature, the FRET assay cannot provide absolutely quantitative data for reaction rates for several reasons:

1) FRET depends on the average distances and orientation of the fluorophore molecules on the growing glycan chains and the resulting PG. We cannot know the distribution and orientation of the fluorophores on the growing glycan chain and whether these parameters remain constant during the synthesis of a glycan chain and there is currently no method to determine these parameters.

2) Different PBPs may produce PG chains with different distribution/density of fluorophores, depending on their affinities to the different lipid II variants.

3) There is currently no methodology available to validate our data, as there is no other published assay that allows quantification of absolute GTase rates.

We discuss these limitations in a new section “Limitation of the FRET assay”. The new real-time control with donor-only or acceptor-only as labelled substrates (Figure 1—figure supplement 4, Figure 3—figure supplement 4) don't support the idea that the fluorophores are highly sensitive to the environment and reinforce our conclusion that FI_acceptor/FI_donor ratio is specific for FRET. The new data in Figure 2—figure supplement 1 clarify what is the cause of the changes in FRET of the produced PG when LpoB is added, and help to interpret FRET curves with or without activators.

Our motivation to develop this assay was not to quantify the GTase rates for a detailed mechanistic comparison between different enzymes. Instead, we aimed to develop the first method to monitor PG synthesis continuously in a biomimetic membrane. Assays with fluorescently-labelled substrates do not report directly on the reaction rates of the enzymes for the unlabelled substrate, complicating precise comparisons between different enzymes (or combinations of enzyme and activator). However, these fluorescent substrates are the key to the specific and sensitive real time detection of PG synthesis in the membrane. In addition, as the assay does not require a complicated setup, it has the potential to be widely used in basic research and for screening for inhibitors. Hence, we believe that the assay will be widely used by the PG community despite its semi-quantitative nature.

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figure 1—source data 1. Numerical data to support graphs in Figure 1 and original gel images for Figure 1B.

elife-61525-fig1-data1.zip^{(2.2MB, zip)}

Figure 1—figure supplement 2—source data 1. Numerical data to support graphs in Figure 1—figure supplement 2.

elife-61525-fig1-figsupp2-data1.xlsx^{(77.2KB, xlsx)}

Figure 1—figure supplement 3—source data 1. Original gel images for Figure 1—figure supplement 3.

elife-61525-fig1-figsupp3-data1.zip^{(354.4KB, zip)}

Figure 1—figure supplement 4—source data 1. Numerical data to support graphs in Figure 1—figure supplement 4.

elife-61525-fig1-figsupp4-data1.xlsx^{(104KB, xlsx)}

Figure 2—source data 1. Numerical data to support graphs in Figure 2 and original gel images for Figure 2B.

elife-61525-fig2-data1.zip^{(722.8KB, zip)}

Figure 2—figure supplement 1—source data 1. Numerical data to support graphs in Figure 2—figure supplement 1 and original gel images used to quantify labelled lipid II consumption.

elife-61525-fig2-figsupp1-data1.zip^{(5.6MB, zip)}

Figure 3—source data 1. Numerical data to support graphs in Figure 3 and original gel images for Figure 3C, E and G.

elife-61525-fig3-data1.zip^{(1.4MB, zip)}

Figure 3—figure supplement 2—source data 1. Numerical data to support graphs in Figure 3—figure supplement 2 and original gel images for Figure 3—figure supplement 2A.

elife-61525-fig3-figsupp2-data1.zip^{(1.8MB, zip)}

Figure 3—figure supplement 3—source data 1. Numerical data to support graphs in Figure 3—figure supplement 3.

elife-61525-fig3-figsupp3-data1.xlsx^{(18KB, xlsx)}

Figure 3—figure supplement 4—source data 1. Numerical data to support graphs in Figure 3—figure supplement 4.

elife-61525-fig3-figsupp4-data1.xlsx^{(142.2KB, xlsx)}

Figure 3—figure supplement 6—source data 1. Numerical data to support graphs in Figure 3—figure supplement 6.

elife-61525-fig3-figsupp6-data1.xlsx^{(24.1KB, xlsx)}

Figure 3—figure supplement 7—source data 1. Numerical data to support graphs in Figure 3—figure supplement 7 and original gel images for gels in Figure 3—figure supplement 7C.

elife-61525-fig3-figsupp7-data1.zip^{(1MB, zip)}

Figure 3—figure supplement 8—source data 1. Numerical data to support graphs in Figure 3—figure supplement 8 and original gel images for gels in Figure 3—figure supplement 8C.

elife-61525-fig3-figsupp8-data1.zip^{(1MB, zip)}

Figure 4—source data 1. Numerical data to support graphs in Figure 4.

elife-61525-fig4-data1.xlsx^{(126.3KB, xlsx)}

Figure 5—source data 1. Numerical data to support graphs in Figure 5.

elife-61525-fig5-data1.xlsx^{(31.1KB, xlsx)}

Supplementary file 1. Table of oligonucleotides used in this study.

elife-61525-supp1.docx^{(22.5KB, docx)}

Transparent reporting form

elife-61525-transrepform.docx^{(253.8KB, docx)}

Data Availability Statement

All data generated or analysed during this study are included in the manuscript and supporting files. Source data files have been provided for Figures 1–5 and the corresponding figure supplements.

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PERMALINK

Real-time monitoring of peptidoglycan synthesis by membrane-reconstituted penicillin-binding proteins

Víctor M Hernández-Rocamora

Natalia Baranova

Katharina Peters

Eefjan Breukink

Martin Loose

Waldemar Vollmer

Roles

Abstract

Introduction

Results

Real-time assay for detergent-solubilized E. coli PBP1B

Figure 1. Förster resonance energy transfer (FRET) assay to monitor peptidoglycan synthesis in real time.

Figure 1—figure supplement 1. Fluorescent lipid II analogues to monitor peptidoglycan synthesis in real time.

Figure 1—figure supplement 2. Analysis of fluorescence spectra to calculate Förster resonance energy transfer (FRET) efficiency.

Figure 1—figure supplement 3. Förster resonance energy transfer assay to monitor peptidoglycan synthesis in real time.

Figure 1—figure supplement 4. Fluorescence intensity (FI) of lipid II-Atto550 and lipid II-Atto647n only changes significantly during reactions when both versions are present.

Intra-chain versus inter-chain FRET

Figure 2. The Förster resonance energy transfer (FRET) signal arises from both the glycosyltransferase and transpeptidase reactions.

Figure 2—figure supplement 1. Effect of proportion of labelled lipid II substrates on PBP1BEc activity in detergents.

FRET assay to monitor PG synthesis in liposomes

Figure 3. The Förster resonance energy transfer (FRET) assay for peptidoglycan synthesis can be adapted for reactions on liposomes.

Figure 3—figure supplement 1. Activity of membrane-reconstituted PBP1BEc is optimal in E. coli polar lipids at low ionic strength.

Figure 3—figure supplement 2. The Förster resonance energy transfer (FRET) assay for peptidoglycan synthesis can be adapted for reactions on liposomes.

Figure 3—figure supplement 3. Moenomycin does not affect Förster resonance energy transfer (FRET) on liposomes with lipid II-Atto550 and lipid II-Atto647n in the absence of class A penicillin-binding proteins.

Figure 3—figure supplement 4. Fluorescence intensity (FI) of lipid II-Atto550 and lipid II-Atto647n in the membrane only changes significantly during reactions when both species are present.

Figure 3—figure supplement 5. Amino acid sequence comparison between LpoP homologues from A.baumannii and P.aeruginosa.

Figure 3—figure supplement 6. LpoPAb stimulates the glycosyltransferase activity of PBP1BAb.

Figure 3—figure supplement 7. Peptidoglycan synthesis activity of A.baumannii PBP1B in the presence of Triton X-100 followed by Förster resonance energy transfer (FRET).

Figure 3—figure supplement 8. Peptidoglycan synthesis activity of P. aeruginosa PBP1B in the presence of Triton X-100 followed by Förster resonance energy transfer (FRET).

Activities of other membrane-bound class A PBPs

PG synthesis on supported lipid bilayers

Figure 4. Addition of lipid II slows down diffusion of PBP1B on supported lipid bilayers.

Figure 4—figure supplement 1. Control of membrane fluidity and integrity upon reconstitution of E. coli PBP1B.

Figure 4—figure supplement 2. E. coli PBP1B is active after reconstitution in supported lipid bilayers (SLBs).

Video 1. Single-molecule imaging of PBP1B on supported lipid bilayers (SLBs).

FRET assay on supported bilayers

Figure 5. Förster resonance energy transfer (FRET) assay on a planar lipid membrane.

Figure 5—figure supplement 1. Control of membrane fluidity and integrity during the Förster resonance energy transfer assay.

Video 2. Förster resonance energy transfer (FRET) assay on supported lipid bilayers (SLBs).

PG synthesized on SLBs

Discussion

Intra-chain vs. inter-chain FRET

Figure 6. Peptidoglycan (PG) synthesis with labelled lipid II versions and detection of Förster resonance energy transfer (FRET).

Limitation of the FRET assay

Coupled reactions in class A PBPs and their activation

Class A PBP activities in the membrane

Towards single-molecule PG synthesis

Materials and methods

Key resources table.

Chemicals

Cloning

Construction of overexpression vector pKPWV1B

Construction of overexpression vector pKPWVLpoP

MGC-64PBP1B-his C777S/C795S

Purification and labelling of proteins

E. coli PBP1B

A. baumannii 19606 PBP1B

P. aeruginosa PBP1B

MGC-64PBP1B-his C777S/C795S

LpoB(sol)

A. baumannii 19606 LpoP(sol)

P. aeruginosa LpoP(sol)

PG synthesis assays in the presence of detergents

In vitro PG synthesis assay using radiolabelled lipid II in detergents

FRET-based in vitro PG synthesis assay in detergents

Quantification of lipid II consumption after the FRET assay in detergents

Analysis of FRET reaction curves

Determination of FRET efficiency

Continuous GTase assay using dansylated lipid II

Time-course GTase assay by SDS-PAGE followed by fluorescence detection

Fluorescence scanning of SDS-PAGE gels with PG synthesis products

PG synthesis in liposomes

Reconstitution of class A PBPs in liposomes

PBP1BEc orientation assay

In vitro PG synthesis assay using radiolabelled lipid II in liposomes

FRET-based in vitro PG synthesis assay in liposomes

Analysis of FRET reaction curves

Assays in SLBs

Figure 2—figure supplement 1. Effect of proportion of labelled lipid II substrates on PBP1B^Ec activity in detergents.

Figure 3—figure supplement 1. Activity of membrane-reconstituted PBP1B^Ec is optimal in E. coli polar lipids at low ionic strength.

Figure 3—figure supplement 6. LpoP^Ab stimulates the glycosyltransferase activity of PBP1B^Ab.

MGC-⁶⁴PBP1B-his C777S/C795S

MGC-⁶⁴PBP1B-his C777S/C795S

PBP1B^Ec orientation assay

Formation of polymer-SLBs and reconstitution of PBP1B^Ec into a supported lipid membrane