Introduction

Bacterial cell wall defines cell shape and provides mechanical protection required for cell integrity. It notably encompasses a mesh-like and rigid polymer (the peptidoglycan) made of glycan strands cross-linked by peptide bonds that surrounds the whole cell1. In firmicutes, a major phylum of Gram-positive bacteria, the cell wall also contains secondary cell wall polymers named teichoic acids (TA). These compounds are anchored either to the peptidoglycan layer (wall teichoic acids, WTA) or to the cell membrane (lipoteichoic acids, LTA)2,3,4,5. WTA and LTA are generally synthesized by separated pathways and are composed of repeated units of polyol-phosphate, the best characterized structures consisting in ribitolphosphate and glycerolphosphate6. They both contribute to the functionality of the cell wall mediating very diverse functions ranging from cation homeostasis, autolysis, phage binding, cell division, cell wall maintenance, biofilm formation as well as adhesion to the host cells and virulence4,7. Being present in both pathogenic and commensal Gram-positive bacteria, TAs are often involved in host-microbe interactions, in particular they can define the immune-stimulatory potential of the bacteria8,9.

WTA and LTA can undergo different modifications, thus providing a structural diversity that modulates their function. Beside glycosylation and phosphocholine decoration10, a preponderant modification is d-alanylation which consists in the formation of an ester bond between a d-alanyl and the ribitolphosphate or the glycerolphosphate of TAs11,12. This modification modulates the negative charge of TAs and consequently modifies the cell surface charge and electrochemical properties. The amount of d-alanine esterified to TAs is variable and is an important mechanism by which bacteria tune teichoic acid functions11. Specifically, L. plantarum regulates the production of inflammation-related cytokines and protect the host (mice) from intestinal disorder through the d-alanine content on its cell envelope13. In the bacterial pathogen Staphylococcus aureus, the presence of d-Alanine residues on TAs changes the strain susceptibility to vancomycin and the ability to persist on human skin through evasion of cutaneous innate defenses14,15. d-alanylation is performed by proteins of the Dlt pathway16,17,18,19. This pathway begins in the cytoplasm with the acyl-carrier protein synthase AcpS that transfers a phosphopantetheine (Ppant) moiety from the coenzyme A onto the apo form of the d-alanyl carrier protein DltC20. Then, Ppant-DltC (holo form) interacts with the d-alanine ligase DltA to give rise to d-alanyl-Ppant-DltC21,22. The latter eventually interacts with the protein DltB that belongs to the membrane-bound-O-acetyltransferase (MBOAT) protein family23. Although this cytoplasmic pathway is supported by numerous studies, the molecular details underlying this suite of interactions remain largely unknown. While the structure of the AcpS-DltC complex has been solved at high resolution in Bacillus subtilis, providing a first description of the interaction between these two proteins, the contribution of Ppant has not been investigated24. The same is true regarding the mode of interaction between DltC and DltA that has never been clearly established. A recent study has even reported an absence of interaction between these two proteins23. In the same study, the mode of interaction between d-alanyl-Ppant-DltC and DltB of Bacillus subtilis has however been clearly determined. Concerning the extracellular step, the way d-alanine is transferred onto TAs remains highly debated. Indeed, it is proposed that DltB would either directly transfer d-alanine onto LTA, or to a lipid intermediate (phosphatidylglycerol or undecaprenyl phosphate) that would be further processed by the DltD protein for both WTA and LTA decoration25. Another hypothesis suggests that DltB could also directly transfer d-alanine to DltD23.

Lactiplantibacillus plantarum (former Lactobacillus plantarum), a nomadic species often found in the animal oro-gastro-intestinal tract26, confers health benefits to its host by supporting nutrition, gut epithelial homeostasis and protection against infections11,27,28,29,30,31. This bacterium encodes for all the conserved enzymes of the TA d-alanylation pathway, including AcpS and the Dlt proteins (DltA, DltB, DltC1 and DltD). The latter are encoded by a unique gene cluster (the pbpX2-dlt operon). This operon also encodes for the less conserved protein DltX that consists of a single transmembrane helix. DltX was shown to be required for the decoration of TA by d-alanine in Bacillus thuringiensis32 but its role within the Dlt pathway remains largely unknown. Interestingly, a second copy of the dltC gene (nc8_1214), named here dltC2, is also encoded elsewhere in the genome of L. plantarum but its potential role in TA d-alanylation has never been investigated. Recently, evidence was provided that deletion of the dltXABCD genes prevents the decoration of TAs by d-alanine33. In addition, and most importantly, it was also demonstrated that the dltXABCD genes are involved in the ability of L. plantarum to promote Drosophila juvenile growth under chronic undernutrition33. d-alanyl decoration of TA is therefore a cardinal step for the molecular dialog underlying the mutualistic interaction between Drosophila and L. plantarum.

In this study, we first showed that individual deletion of the acpS, dltA, dltB genes and double deletion of dltC1 and dltC2 abolish TA d-alanylation as well as the beneficial interplay between L. plantarum and Drosophila. We next disentangled the suite of interactions between the cytoplasmic components of the Dlt pathway in L. plantarum. To this end, we solved the X-ray structures of DltC1 in complex with AcpS, and of DltA. Using NMR spectroscopy, microscale thermophoresis and molecular modeling, we characterized for the first time the molecular details of the interactions between AcpS and DltC1, DltC1 and DltA in absence or presence of Ppant, and the mode of interaction between DltC1 and DltB. Altogether, these data show that a unique surface of DltC1 is required for the sequential interaction with AcpS, DltA and finally DltB, providing thus the molecular mechanism for the cytoplasmic steps of the d-alanylation pathway.

Results

AcpS, DltA, DltB and DltC proteins are essential for the TA d-alanylation in L. plantarum

We first aimed at confirming that AcpS and the Dlt proteins DltA, DltC1 and DltB were individually required for the cytoplasmic steps of the TA d-alanylation pathway in L. plantarum. To this end, we generated the ∆acpS, ∆dltA, ∆dltC1 and ∆dltB deletion mutants in the LpNC8 strain by homology-based recombination and determined the amount of d-Ala esterified to TAs in each strain (Fig. 1a and b). Compared to the d-Ala level measured in the WT strain (LpNC8 strain), the levels of d-Ala released from these mutants were almost undetectable, except for the ∆dltC1 mutant in which only a decrease of 25% is measured (Fig. 1b). This confirms that AcpS, DltA and DltB are required in this pathway but suggests that a complementary DltC activity should contribute to TA d-alanylation. In addition to dltC1 (nc8_1734, henceforth named dltC1) located within the dlt operon, careful inspection of the genome of L. plantarum identified a second gene, named dltC2 (nc8_1214), present in another place in the genome (Fig. 1a), which encodes a protein sharing 88.75% identity and 96.25% similarity with DltC1 (Supplementary Fig. 2a). The two proteins actually differ only in 9 residues. We therefore hypothesized that DltC2 could also contribute to TA d-alanylation. While we did not detect any decrease of TA-alanylation in LpNC8 devoid of the dltC2 gene, we observed that TA d-alanylation was completely abolished in the double deletion mutant ∆dltC1dltC2 (Fig. 1b). Together, these results show that DltC1 is the main enzyme at play in the TA d-alanylation pathway but also suggest that both DltC1 and DltC2 could work in synergy to allow d-alanylation of TA.

Figure 1
figure 1

Characterization of L. plantarum mutants in Drosophila’s growth. (a) Genetic organization of Lp engineered strains: WT corresponds to LpNC8 strain; ∆dltA corresponds to LpNC8 deleted for dltA gene; ∆dltB corresponds to LpNC8 deleted for dltB gene; ∆dltC1 corresponds to LpNC8 deleted for dltC1 gene; ∆dltC2 corresponds to LpNC8 deleted for dltC2 gene; ∆dltC1C2 corresponds to LpNC8 deleted for dltC1 and dltC2 genes; ∆acpS corresponds to LpNC8 deleted for acpS gene. in-frame deletions were constructed through homology-based recombination with double-crossing over, in such a way that the two first triplets of the sequence are fused with the two last. (b) Amount of d-Ala released from whole cells of NC8 and derivative mutants by alkaline hydrolysis and quantified by HPLC. Error bars represent the standard deviations. (c) Larval longitudinal length after inoculation with strains LpNC8, ∆dltA, ∆dltB, ∆dltC1, ∆dltC2, ∆dltC1dltC2, ∆acpS or PBS (for the GF condition). Larvae were collected 6 days after association and measured as described in the Methods section. Green asterisks illustrate statistically significant difference with LpNC8 larval size; ****: p < 0.0001. Center values in the graph represent means and error bars represent SD. Representative graph from one out of three independent experiments. (d) Day when fifty percent of pupae emerge during a developmental experiment (D50) for GF eggs associated with strains LpNC8, ∆dltA, ∆dltB, ∆dltC1, ∆dltC2, ∆dltC1dltC2, ∆acpS or PBS (for the GF condition). Center values in the graph represent means. Green asterisks illustrate statistically significant difference with LpNC8 D50; ns represent absence of statistically significant difference with LpNC8 D50. ***: 0.0001 < p < 0.001; **: 0.001 < p < 0.01; *: p < 0.05.

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The pbpX2-dlt operon of L. plantarum is required to promote the Drosophila juvenile growth under chronic undernutrition33. To analyze the respective contribution of each gene in this functional setting, we assessed the impact of each mutant strain on the ability of L. plantarum to support the growth of Drosophila larvae. To this end, we determined both the larval size at six days post-inoculation and the developmental timing of the germ free (GF) individuals and individuals mono-associated with the wild-type or ∆acpS, ∆dltA, ∆dltB or ∆dltC1dltC2 Lp strains (Fig. 1c). ∆acpS, ∆dltA, ∆dltB or ∆dltC1dltC2 associated larvae were significantly smaller than those associated with the wild type LpNC8 strain. In addition, their larval development was also severely delayed (Fig. 1d). Of note, single deletion of dltC1 or dltC2 induced only slight defects reinforcing the notion that dltC genes encode redundant and synergistic activities. These results therefore showed that AcpS, DltA, DltB and, DltC1 or DltC2, are needed for TA d-alanylation and contribute to L. plantarum beneficial activity towards its animal host exemplified here by the promotion of Drosophila juvenile growth.

Structural characterization of the AcpS/DltC1 complex

The first step in the d-alanylation mechanism of TA is the transfer of the phosphopantetheinyl moiety of coenzyme A (Ppant) (Supplementary Fig. 2) to the DltC protein by the Phosphopantetheinyl transferase AcpS. During this process, the Ppant is covalently attached to a conserved Serine residue (Ser38) (Supplementary Fig. 1a) converting DltC1 or DtC2 from an apo-form to the active holo-form. The high sequence identity between DltC1 and DltC2 suggested that the mode of interaction with AcpS is likely conserved (Supplementary Fig. 1a). Therefore, and as the dltC1 gene is located in the dlt operon, we decided to focus our structural analyses on the AcpS/DltC1 complex and to transpose our observations to DltC2 to characterize its mode of interaction with AcpS.

To study the transfer of Ppant, we overexpressed and purified L. plantarum AcpS and DltC1 fused to a His-tag from Escherichia coli cells. Then, by mixing the two Ni-affinity purified proteins, we performed size exclusion chromatography to isolate the AcpS-DltC1 complex (Supplementary Fig. 3). The fractions containing the AcpS-DltC1 complex were then pooled to conduct the crystallization trials. High quality diffracting crystals were obtained and the crystal structure was solved at 1.88 Å resolution. The data collection and refinement statistics are summarized in Table 1. The AcpS-DltC1 complex clearly adopts a 3:3 stoichiometry with a 3D arrangement similar to that of the AcpS-DltC complex from B. subtilis24 with a rmsd (root mean square deviation) of 1.062 Å on 341 residues. This arrangement forms a 3-branch AcpS star that is decorated with 3 DltC1 molecules, each of them being at the interface between two AcpS subunits (Fig. 2a).

Table 1 Data collection and refinement statistics.
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Figure 2
figure 2

The L. plantarum AcpS-DltC1 complex structure. (a) Cartoon representation of the L. plantarum AcpS-DltC1 3D structure with the 3-branch AcpS star shown as a cyan-green color gradient and the DltC1 molecules in pink. The Ppant cofactor bound to one DltC1 molecule is shown as black sticks and highlighted by a black dashed-rectangle. The main contact surfaces, 1 and 2 are depicted as red dashed line and circle respectively. (b) Close-up on the contact surface 1. The interacting residues are shown in stick. Comparison between L. plantarum (left—AcpS in blue and DltC1 in pink) and B. subtilis (right—AcpS in light grey and DltC in dark grey—PDB entry 1F80) complex structures. (c) Close-up on the contact surface 2. The interacting residues are shown in stick. Comparison between L. plantarum (left—AcpS in green and DltC1 in pink) and B. subtilis (right—AcpS in light grey and DltC in dark grey).

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Compared to the available structures of homologous proteins23,24,34,35,36,37,38, the global fold of AcpS and DltC1 is not modified upon complex formation (Supplementary Figs. 4 and 5). Indeed, each AcpS monomer adopts an α/β fold and shows high structural similarity with already published AcpS structures with a rms difference of Cα atoms of 0.7 Å compared to AcpS structure from B. subtilis24. The longest α-helix (α4 ranging from residues 44 to 65), is wrapped on one side by a three-stranded antiparallel β-sheet (β1, β4 and β5) and a β-ribbon composed of two strands (β2 and β3) and on the other side by three small α-helices and a 3–10 helix (Fig. 2a and Supplementary Fig. 4). The β-sheet of each monomer is positioned at the center of the trimer and contributes to the trimerization via hydrophobic and a few hydrophilic interactions.

Regarding DltC1, its structure exhibits a right-handed helix bundle topology, a typical fold shared with the other DltC proteins whose structure has been solved23,35,37. The rms differences of the Cα atoms ranged from 1 to 1.2 Å. While the 3 main α-helices (ranging from residues 3–18, 38–52 and 69–80) are well conserved, the region connecting the α-helices are less structured in DltC1 than those in other DltC homologs. A long loop is connecting α1 and α2 while in B. subtills or in L. rhamnosus an additional 6 residues helix is observed35,37. Between α2 and α3 the short perpendicular helix usually described in homologous structures is replaced by a 3–10 helix between residues 58–61(Fig. 2a and Supplementary Fig. 5).

The DltC1 catalytic Ser38 is the first residue of helix α2. A clear electron density, corresponding to a phosphate moiety of Ppant, was observed covalently linked to the Ser38 side chain in only one of the 3 subunits of DltC1 of the complex (Fig. 2a). Although the electron density of the remainder of Ppant moiety was weaker, presumably due to its intrinsic flexibility, the distal end of the prosthetic group could be modeled (Supplementary Fig. 6a). While Ppant folds back on DltC at the interface between helix α2 and α3 in the AcpS-DltC complex from B. subtilis, its position differs in our structure (Supplementary Fig. 6b). Indeed, although covalently linked to Ser38, it points toward a neighboring AcpS molecule and ends in the vicinity of the end of β2 and the beginning of α4. In this position, the thiol group at the extremity of the Ppant is stabilized via H-bond interactions with Glu49, Asp82 and the well conserved Arg46 of AcpS (Supplementary Fig. 6a). This Ppant location is however similar to that of the CoA molecule co-crystallized with AcpS from B. subtilis24. Considering the sequence conservation between DltC1 and DltC2 (Supplementary Fig. 1a), we used the 3D DltC1 structure to generate a 3D model of DltC2 and showed that all the structural features and binding analysis described for DltC1 are conserved in DltC2 (Supplementary Fig. 1b and c).

Binding interface of DltC1 to AcpS

The interface between two adjacent AcpS monomers serves as a docking area for DltC1 binding (Fig. 2a). Two main contact surfaces, annotated 1 and 2 on Fig. 2a, were identified between AcpS and DltC1. The first one is the largest and locates at the interface between the helix α2 and the loop connecting helices α2 and α3 of DltC1 and the helix α1 of one adjacent AcpS monomer. It is composed of hydrophobic and ionic interactions (Fig. 2b). The two main hydrophobic contacts involve Met39 and Leu46 of DltC1. Met39 binds in a pocket formed by the side chains of Ile16 and Ile 19 of AcpS helix α1 and to the side chain of Phe55 while Leu46 interacts only with Leu26 of AcpS. Two similar clusters of hydrophobic interactions were also described in the homologous complex structure described for B. subtilis but amino acids are not conserved24 (Fig. 2b). In this contact surface 1, key ionic interactions also contribute to the stabilization of the complex. One is strictly conserved in B. subtilis and is established between the well conserved Arg15 of AcpS and Asp37 of DltC1, the latter being located just before the reactive Serine. A second salt bridge is formed between Glu47 of DltC1 and Lys22 of AcpS (replaced by an Arg residue in B. subtilis) and is the only interaction observed at the terminal end of helix α2 of DltC1. The second contact surface (contact surface 2 on Fig. 2a) involves a salt bridge between DltC1 and the second neighboring AcpS molecule and further stabilizes the L. plantarum complex. The Asp32 of DltC1 located in the long loop between helices α1 and α2 forms a strong ionic interaction with Arg46 of AcpS and an H-bond with Tyr42. Although Asp32 from DltC1 and Arg46 from AcpS are conserved, these interactions were not described in B. subtilis complex, the loop region between α1 and α2 adopting different conformations in both structures (Fig. 2c).

Facing these structural features, we wondered what could trigger the dissociation of the AcpS-DltC1 complex after the Ppant has been transferred to DltC1, an event necessary prior to the transfer of d-Ala as a thiol ester to the Ppant by DltA ligase. We hypothesized that Ppant transfer to each DltC1 molecule could be key by decreasing the stability of the complex leading to the release of holo-DltC1. To test this hypothesis, we produced both the apo- and holo-forms of LpDltC1. The apo form of DltC1 was obtained by mutating the catalytic Ser in Ala. On the other hand, the production of the holo-form (full Ppanted DltC1) was achieved by co-expressing both LpAcpS and DltC1 from a pRSF-Duet1 vector in E. coli. The presence of the Ppant modification in DltC1 was confirmed after its purification by mass spectrometry analysis (Supplementary Fig. 7). Next, purified AcpS was mixed with either apo- or holo-DltC1 and the mixtures were subjected to size exclusion chromatography. In the case of apo-DltC1 a large elution peak containing the AcpS-DltC1 complex is obtained followed by a small peak of unbound DltC1 (Supplementary Fig. 3a and c). By contrast, mixing of AcpS with Ppant-modified DltC resulted in two large separate peaks corresponding to unbound AcpS and holo-DltC1, respectively (Supplementary Fig. 3b and c). No complex formation was therefore detected demonstrating that the AcpS-DltC1 complex can no longer form when DltC1 is fully Ppanted. Microscale thermophoresis (MST) experiments also confirmed that AcpS do not interact with holo-DltC1 (Supplementary Fig. 8).

Structural characterization of DltA

The d-alanine-d-alanyl carrier protein ligase (DltA) catalyzes a two-step reaction: the activation of d-Ala using ATP to form a d-alanyl-AMP intermediate and then the transfer of d-Ala as a thiol ester to the Ppant group of DltC. To investigate this mechanism, we produced and purified L. plantarum DltA and solved its crystal structure at 2.1 Å resolution. The data collection and refinement statistics are summarized in Table 1. The asymmetric unit of the crystal contains two DltA molecules which are largely similar (rmsd of 0.5 Å on 392 residues). Out of 508 residues of full-length DltA, the final model contains residues 3–401 in one molecule and residues 3–400 in the second molecule of the asymmetric unit. The last 100 aminoacids constituting the C-terminal minor domain are not visible in the electron density although present in the purified sample. The loop region ranging from residues 152 and 160 could only be partially modeled in one copy due to poor electron densities for residues 153 to 157 but was fully visible in the second molecule. The overall structure of the large N-terminal domain (residues 3–395) is composed of three subdomains (referred in previous studies as subdomains A, B and C39) two of which, A and B, sharing a similar topology with a β-sheet flanked on both sides by α -helices (Fig. 3). The last subdomain C (residues 321–400) forms a distorted β -barrel. Superimposition of the large domain of L. plantarum DltA with homologous DltA structures from B. subtilis39, B. cereus40,41,42 and S. pyogenes (PDB entries 3L8C, 3LGX, unpublished data) reveals high structural similarity with root mean square deviations (r.m.s.d.) in the range of 0.7–0.9 Å for 300 residues (Supplementary Fig. 9).

Figure 3
figure 3

The L. plantarum DltA structure. (a) Cartoon representation of the N-terminal domain of DltA structure. The subdomains A (3–48, 188–320), B (49–187) and C (321–400) are respectively colored in orange, yellow and wheat. The d-Ala-AMP substrate is shown as blue stick. The P-loop is encircled with dashed black line. (b) Surface representation of the N-terminal domain of DltA colored according to sequence conservation. The color ram** from white (low score) to red (identity) locate areas of weak and strong sequence conservation. The figure was generated with ENDscript (https://endscript.ibcp.fr/) (c) Close-up on the substrate binding site of DltA, the d-Ala-AMP (in blue) and the residues involved in the interaction are shown as sticks (d) Sequence alignment between L. plantarum DltA (LpDltA) and DltA from B. cereus (BcDltA—PDB entry 3DHV). The secondary structures extracted from the X-ray structures are depicted above. The position of the LpDltA residues interacting with the d-Ala-AMP substrate are highlighted by a blue triangle. The position of the P-loop is shown with a dashed black line. The figure was generated by ESPript (https://espript.ibcp.fr/).

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Interaction between DltA and d-Ala-AMP

As crystallization was achieved in the presence of d-Ala and ATP, one molecule of d-Ala-AMP could be observed in the electron density of one DltA molecule of the asymmetric unit (Fig. 3). This position is similar to that described in the B. cereus DltA structure40 (Supplementary Fig. 9a). In the other molecule, the clear electron density allowed to position with confidence the adenine ring moiety and the d-Ala in the same position as in the other DltA molecule. However, the weaker density observed at the ribose and phosphate positions did not support the presence of the covalent bond expected between the d-Ala and the phosphate group. This suggests either a certain degree of flexibility or that we manage to trap an intermediate during the d-Ala activation by an ATP molecule. The d-Ala-AMP binding cleft of DltA is defined at the interface between loop 270–274, the β strands β13 (residues 290–295), β14 (residues 305–308) and β21 (residues 394–399) and the P-loop (phosphate binding loop or Walker A motif) comprised between residues 152 and 160 (Fig. 3b and c). More precisely, the adenine ring is sandwiched between residues Gly271 and Glu272 on one side and Tyr295 on the other side. The ribose forms three H bonds with the carboxylate group of Asp384, the side chain of Tyr396 and the hydroxyl group of Gly271. The phosphate group interacts with the side chain and the amide group of Thr298. The d-alanyl amino group appears anchored by three interactions with the carboxylate group of Asp198 and with the hydroxyl groups of Gly296 and Val302. As already described for other DltA structures mentioned above, a conserved cysteine residue 270 is positioned close to the side chain of the d-alanyl moiety participating in the d-Ala versus L-Ala specificity of the enzyme40 (Fig. 3c and d). Altogether, this analysis shows that this interaction web is largely similar to that of the d-Ala-adenylate described in the DltA structure from B. cereus40 with the exception that the interaction observed with the small C-terminus domain could not be confirmed due to its absence in our crystal structure. As a result, the binding cleft appears more widely opened. However, and very interestingly, our structure showed that the P-loop clearly participates to the sha** of the binding cleft whereas it was not observed in the DltA structure of B. cereus (see below) (Fig. 3b and c and Supplementary Fig. 9).

While most of the surface residues are poorly conserved in DltA proteins and homologous proteins forming AMP intermediates, the P-loop is highly conserved (nine residues from Thr152 to Lys 160)43,50 to allow for Gibson Assembly. PCR amplifications were made using the following primers: OL01/OL02 and OL03/OL04 (dltA), OL25/OL26 and OL27/OL28 (dltB), OL07/OL08 and OL09/OL10 (dltC1), OL13/OL14 and OL15/OL16 (acpS), OL19/OL20 and OL21/OL22 (dltC2) listed in Supplementary Table 2. The resulting plasmids obtained by Gibson Assembly (NEB) were transformed into L. plantarumNC8 electrocompetent cells and selected at the permissive temperature (28 °C) on MRS plates supplemented with 5 µg/mL of erythromycin. Overnight cultures grown under the same conditions were diluted and shifted to the non-permissive temperature (41 °C) in the presence of 5 µg/mL of erythromycin to select single crossover integrants. Plasmid excision by a second recombination event was promoted by growing integrants at the permissive temperature without erythromycin. Deletions were confirmed by PCR followed by sequencing. The strain deleted for dltC1 and dltC2 was obtained by the sequential deletion of dltC1 followed by dltC2.

Knock-in of dltA modified version in L. plantarum NC8

L. plantarumNC8 strains carrying a modified version of the dltA gene was built by knocking-in the modified sequences on ∆dltA strain constructed in this study. dltA modified sequence (dltA∆P-loop and dltA5m) were synthetized by Twist Bioscience. In order to perform Gibson Assembly (NEB) the overlap** regions were added by PCR with OL32 and OL33 on the respective plasmid provided by Twist Bioscience. The 5′- and 3′-terminal regions of dltA region were PCR-amplified with Q5 High-Fidelity 2X Master Mix (NEB) from L. plantarumNC8 chromosomal DNA using primers OL01/OL31 and OL34/OL4. The 3 fragments were assembled with pG + host951. The resulting plasmids were transformed into ∆dltA electrocompetent cells and selected at the permissive temperature (28 °C) on MRS plates supplemented with 5 µg/mL of erythromycin. Overnight cultures grown under the same conditions were diluted and shifted to the non-permissive temperature (41 °C) in the presence of 5 µg/mL of erythromycin to select single crossover integrants. Plasmid excision by a second recombination event was promoted by growing integrants at the permissive temperature without erythromycin. dltA5m and dltA∆P-loop knock-ins were confirmed by PCR followed by sequencing.

Quantification of d-alanine by high-performance liquid chromatography (HPLC)

d-alanine esterified to teichoic acids was detected on whole bacterial cells and quantified as described previously by52. Briefly, L. plantarum wild-type and mutants were grown overnight in 100 ml MRS. Bacteria were harvested, washed twice with 20 mM ammonium acetate, pH 4.7, resuspended in the same buffer, heat-inactivated for 10 min at 100 °C and finally lyophilized. d-alanine was released from whole heat-inactivated bacteria (10 mg) by mild alkaline hydrolysis with 0.1 N NaOH (150 µl) for 1 h at 37 °C. After neutralization with 0.1 N HCl, the extract was incubated with Marfey's reagent (1-fluoro-2,4-dinitrophenyl-5-L-alanine amide; Sigma). This reagent reacts with the optical isomers of amino acids to form diastereomeric N-aryl derivatives, which can be separated by HPLC. Separation of the amino acid derivatives was performed on a C18 reversed-phase column (Zorbax Eclipse Plus C18 RRHD 2.1 × 50 mm 1.8 µm Agilent) with an Agilent UHPLC 1290 system with a linear elution gradient of acetonitrile in 20 mM sodium acetate buffer (pH 5.0). The eluted compounds were detected by UV absorbance at 340 nm. Quantification was achieved by comparison with d-alanine standards in the range of 50 to 2000 pmol derived with Marfey’s reagent. Mean values were obtained from three independent cultures.

Larval size measurements

Axenic adults were put overnight in breeding cages to lay eggs on sterile poor yeast diet. Fresh axenic embryos were collected the next morning and seeded by pools of 40 in tubes containing fly food. 1 × 108 CFUs or PBS were then inoculated homogenously on the substrate and the eggs. Petri dishes are incubated at 25 °C until larvae collection. Drosophila larvae, 7 days after inoculation, were randomly collected and processed as described by30. Individual larval longitudinal length was quantified using ImageJ software53.

Developmental timing determination

Axenic adults were placed in breeding cages overnight to lay eggs on sterile poor-yeast diet. Fresh axenic embryos were collected the next morning and seeded by pools of 40 in tubes containing fly food. A total of 1 × 108 CFUs of each strain or PBS was then inoculated homogeneously on the substrate and the eggs and incubated at 25 °C. The emergence of pupae was scored every day until all pupae had emerged. D50 (day when 50% of the pupae emerged) was determined using D50App33.

Escherichia coli plasmid construction

DNA fragments were amplified by polymerase chain reaction using L.plantarum cDNA as a template and oligonucleotides listed in Supplementary Table 3. The DNA encoding the full-length wild-type L. plantarum DltC1, and the DltC1-S38A mutant were cloned into the NcoI and XhoI sites of the pET-28a(+) vector that expresses proteins fused to a C-terminal hexahistidine (His)6 tag (Supplementary Table 4). To construct plasmids producing the full-length wild-type L. plantarum AcpS, the full-length wild-type L. plantarum DltA and the DltA-∆P-loop mutant (missing the loop region comprised between T152 and K160), the corresponding DNA fragments were cloned into the NdeI and PstI sites of the pT7-7 vector that expresses proteins fused to a TEV (tobacco etch virus) cleavage site and a C-terminal Hexahistidine (His)6 tag. An untagged LpAcpSWT construct was also prepared in the pT7-7 vector by introducing a stop codon before the TEV cleavage site and the C-terminal hexahistidine tag (Supplementary Table 4).

The gene of the DltA-5 M harboring the K366A, K369A, D401A, Y435A and K440A mutations was synthesized by Twist Bioscience in a pET-28a(+) vector that contains a a TEV (tobacco etch virus) cleavage site followed by a C-terminal Hexahistidine (His)6 tag (Supplementary Table 4). The nucleotide sequences encoding the wild type DltC1 fused to a C-terminal His6 tag and the wild type AcpS were also inserted in pRSF-Duet1 vector for co-expression (Supplementary Table 4). The final constructs were verified by DNA sequencing.

Protein production and purification

DltA, DltA-∆P-loop, DltA-5 M, untagged and C-terminal 6His tagged AcpS, and the DltC1-6His-AcpS construct cloned in pRSF-Duet1were expressed in E. coli BL21 (DE3) cells. DltC1 and DltC1-S38A mutant were expressed in E. coli BL21 (DE3)-RIPL cells. Cells were grown in LB media at 37 °C and induced with 0.5 mM isopropyl b-d-1-thiogalactopyranoside (IPTG) overnight at 18 °C. The cells were then harvested by centrifugation and resuspended in lysis buffer (50 mM Tris, pH 7.5, 500 mM NaCl, 10% Glycerol, 1 mM Dithiothreitol (DTT) 0.01 mg/ml Lysozyme, 0.006 mg/ml Dnase/RNase, 1 × antiprotease CLAPA). The resuspended cells were disrupted by sonication and centrifuged at 14,000 g for 45 min. Proteins were all purified by a first step of Ni–NTA affinity chromatography with the elution buffer (50 mM Tris, pH 7.5; 300 mM NaCl; 1 mM Dithiothreitol (DTT); 250 mM Imidazole). The eluted fractions were then concentrated and buffer-exchange was performed in a centrifugal filter unit with gel filtration buffer (50 mM Tris, pH7.5; 100 mM NaCl and 1 mM Dithiothreitol (DTT)). The proteins were finally applied to a Superdex 200 10/300 GL size exclusion column (GE Healthcare) and eluted with gel filtration buffer.

In order to obtain the AcpS-DltC1 complex, the DltC1 containing fractions eluted from the first Ni–NTA chromatography were mixed with lysate from cells overexpressing untagged AcpS. The mix was concentrated and buffer-exchange was performed in a centrifugal filter unit with lysis buffer. The protein mix was loaded onto a nickel-NTA (Qiagen) column and recovered with elution buffer. The eluted fractions (containing DltC1 and AcpS-DltC1 complex) were then concentrated and buffer-exchange was performed in a centrifugal filter unit with gel filtration buffer. The sample was applied to a Superdex 200 10/300 GL size exclusion column and eluted with gel filtration buffer. Isotopically labelled proteins for NMR spectroscopy were prepared by growing cells in M9 minimal media containing 15NH4Cl and/or [13C]-glucose. Purifications were performed as described above except that the gel filtration buffer was replaced by an NMR buffer composed of 20 mM MES, pH6.5; 150 mM NaCl.

Crystallization, data collection and structure determination

Crystallization screenings were performed by the sitting-drop vapor-diffusion method at 293 K using crystallization kits Crystal Screen 1 and 2, PEG/Ion PEG/Ion 2 (Hampton Research). The crystallization drops (0.2µL protein solution and 0.2µL reservoir solution) were set up using a Mosquito crystallization robot. The DltA wild type protein was concentrated to 24 mg/ml and mixed with 1 mM ATP and 1 mM d-Ala prior to crystallization. Diffraction quality crystals grew after around 3 months from a solution of the PEG Ion crystallization kit containing 20% PEG3350 and 0.2 M KCl. The AcpS/DltC1 complex was concentrated to 4.8 mg/ml and crystallized in 3 days in one of the PEG Ion crystallization kit conditions containing 20% PEG3350 and 0.2 M Sodium Malonate. After rapid soaking in mother liquor supplemented with 20% (v/v) glycerol, crystals were flash-cooled in liquid nitrogen prior to diffraction experiments. Diffraction data were collected at cryogenic temperature (100 K) on beamline ID30A-3 at the European Synchrotron Radiation Facility (ESRF, Grenoble, France) from the AcpS/DltC1 complex crystals and on beamline PROXIMA-1 at SOLEIL synchrotron (Gif sur Yvette, France) for the DltA crystals.

Data were processed using the XDS package54. The structures were solved by molecular replacement using Phaser implemented in PHENIX55. The PDB entries 3DHV and 1F80 were used as starting models respectively to solve DltA and the AcpS/DltC1 complex. Both structures were refined using iterative rounds of COOT56 and PHENIX55. The quality of the final structure was assessed with MOLPROBITY before deposition at the PDB under the code 7R27 (DltA) and 7R49 (AcpS-DltC1). Sequence alignments and structure images were generated with PyMOL (Schrödinger, LLC), ESPript and ENDscript57.

NMR resonance assignments

The 2D- and 3D-NMR experiments were collected on a 3 mm NMR tube filled with 150 µL of a 268 μM 13C, 15N-labeled apo-DltC1 (S38A) sample prepared in 20 mM HEPES, 150 mM NaCl buffer at pH 6.5 and containing 7%D2O. Backbone resonance assignments were carried out using a combination of 2D 1H-15N-BEST-TROSY and 3D BEST-HN(CO)CACB, BEST-HNCACB, BEST-HNCO, BEST-HN(CA)CO, BEST-H(NCACO)NH58,59. A 2D 1H-15N-BEST-TROSY were acquired on a 150 μM 15N-labeled holo-DltC1 prepared in the same buffer and was assigned by comparison with the data acquired on 13C, 15N-labeled apo-DltC1 sample. All the spectra were acquired at 25 °C on Bruker Avance III 700 MHz spectrometer equipped with a xyz-gradient a TCI cryoprobe. The NMR spectra were processed using the TopSpin™ software by Bruker in its 4.1 version and were analyzed using the CcpNmr Analysis software in its version 3.060. The 1H chemical shifts were referenced to the internal standard 4,4-dimethyl-4-silapentane-1- sulfonic acid (DSS) methyl resonance. 13C and 15N chemical shifts were referenced indirectly using the IUPAC-IUB protocol61.

NMR titration experiments

Interaction studies of apo-DltC1 and holo-DltC1 with DltA, DltA-5M and DltA-∆P-loop were performed using 3 mm NMR tubes with 15N-labeled apo-DltC1 and holo-DltC1 at concentration varying from 150 to 300 μM and prepared in a buffer containing 20 mM HEPES buffer, pH 6.5, 150 mM NaCl and 7% (vol/vol) D2O. Unlabelled DltA samples were prepared in the same buffer at a concentration of 342 μM for titration with apo-DltC1 and 430 μM for titration with holo-DltC1 titration. DltA-5M and DltA-∆P-loop mutants were also prepared in the same buffer at a concentration of 781 μM and 139 μM, respectively. Unlabeled DltA proteins were successively added to the 15N-labelled DltC1 proteins to reach a protein-to-protein ratio of 0, 0.5, 1 and 1.3 for DltA/apo-DltC1, a ratio of 0, 0.2, 0.4, 0.6, 0.8, 1.0 and 1.2 for DltA/holo-DltC1, a ratio of 0, 0.2, 0.4, 0.6, 0.8, 1.0 and 1.8 for DltC1/DltA-5 M and holo-DltC1/DltA-5M and a ratio of 0, 0.1, 0.2, 0.5, 1.0 and 1.8 for holo-DltC1/DltA-∆P-loop. [1H, 15N]-BEST-TROSY spectra were collected at 25 °C for each protein ratio using Bruker AVANCE spectrometers equipped with a TCI cryoprobe and operating at 700 MHz proton frequency. Analysis software CcpNmr 3.0 was used to monitor protein chemical shift perturbations for every assigned amide resonance by superimposition of the 15N-BEST-TROSY spectra and automatic peak picking. Chemical shift perturbations (Δδ) were calculated on a per-residue basis for the highest substrate-to-protein ratio as described previously62.

Docking

Models of holo-DltC1 in complex with DltA were built with the version HADDOCK2.2 of “The HADDOCK web server for data-driven biomolecular docking”49. As starting structures, we used the X-ray crystallography structure of L. plantarum holo-DltC1 (this study, PDB ID 7R49), a model of full length DltA obtained with Phyre63 and the substrate position from the DltA N-terminal domain structure in complex with d-Ala-AMP (this study, PDB ID 7R27). Residues of DltC1 that showed chemical shift perturbations above the threshold in Fig. 4g were considered as active Ambiguous Restraints (AIR). Calculations were performed with 2000 structures during the HADDOCK rigid body energy minimization, 200 structures during the refinement, and 200 structures during the refinement in explicit water. The output model structures were sorted with the HADDOCK built-in clustering tool using the Fraction of Common Contacts (FCC) method64 with a 0.60-Å cutoff and a minimum of 4 structures per cluster. To improve the convergence during the HADDOCK run, a 2.5-Å unambiguous restraint was introduced between the sulfur atom of the Ppant and the phosphorus of the AMP molecule.

Microscale thermophoresis assays

Protein–protein interactions were analyzed by microscale thermophoresis (MST)65. Buffer of purified and concentrated protein samples was exchanged on a desalting PD-10 column to labeling buffer containing Hepes 25 mM pH7.5, NaCl 300 mM, Tween20 0.05% (w/v). Proteins were then labeled with NHS red fluorescent dye according to the instructions of the RED-NHS Monolith NT Protein Labeling kit (NanoTemper Technologies GmbH, Munchen, Germany). After a short incubation of target-partner complex, the samples were loaded into MST standard-treated glass capillaries and measurements were performed at 22 °C. The assays were repeated three times for each affinity measurement. Data analyses were performed using NanoTemper Analysis software provided by the manufacturer.

Statistics and reproducibility

Data representation and analysis of d-alanine quantification and Drosophila larval size measurements and developmental timing determination were performed using Graphpad PRISM 6 software (www.graphpad.com). A total of 3 to 5 replicates were used for all experiments performed in this study in order to ensure representativity and statistical significance. All samples were included in the analysis. Experiments were done without blinding. Two-sided Mann Whitney’s test was applied to perform pairwise statistical analyses between conditions for Drosophila larval size measurements and developmental timing determination experiments.

All MST assays were performed in triplicate and Kd represent the mean values with corresponding standard deviations.