Inhibition of CdaA by GlmM in L. monocytogenes

To evaluate whether GlmM also inhibits the activity of CdaA in L. monocytogenes in vivo, we constructed strains allowing the IPTG-dependent overproduction of the wild type GlmM and the GlmM F154I variants. As shown above, the GlmM F154I variant does not inhibit CdaA as strong as the wild type GlmM enzyme (see above) (FIG 7B).

To assess whether a GlmM from a bacterium that does not produce c-di-AMP is capable of inhibiting CdaA, we replaced the native glmM gene with the glmM gene from E. coli. The wild type L. monocytogenes strain carrying the integrated empty vector served as a control. As shown in FIG 7C, growth of the strains was not affected when the GlmM variants were over-produced after the addition of IPTG. Surprisingly, the strain carrying the inducible E. coli glmM allele grew also in the absence of IPTG. It is tempting to speculate that GlmM from E.

coli is more active than the L. monocytogenes enzyme, thereby allowing the bacteria to produce sufficient amounts of precursors for cell wall biosynthesis in the absence of IPTG. By contrast, the strains in which the native GlmM variants were depleted grew much slower than the wild type strain (FIG 7C). To conclude, the native GlmM can be replaced by the F154I variant and by the homolog from E. coli in L. monocytogenes without affecting bacterial growth.

Next, we determined the intracellular c-di-AMP levels in the L. monocytogenes strains produing the different GlmM variants durproduing growth in LSM medium. As shown in FIG 7D, the c-di-AMP levels were only slightly reduced in the strains overproducing the GlmM variants.

Thus, the overproduction of GlmM per se is not sufficient for inhibiting CdaA in vivo. To assess whether the GlmM variants modulate the production of c-di-AMP in L. monocytogenes during adaptation to osmotic stress, we cultivated the strains in LSM medium until an OD600

of 0.5-0.6 and samples for determining the cellular c-di-AMP levels were taken. The cultures were split and diluted with pre-warmed LSM medium containing sodium chloride to a final concentration of 0.5 M, or with equal amounts of standard LSM medium as a control. The cultures were further incubated for 25 minutes and the c-di-AMP levels were compared with those obtained from culture samples prior to the dilution step.

As shown in FIG 7D, in all strains the c-di-AMP levels did not change when the cultures were diluted with LSM medium. By contrast, the cellular c-di-AMP levels strongly decreased in the strains overproducing the native GlmM enzyme after salt stress (FIG 7D). The decrease of the c-di-AMP level was less pronounced in a strain producing the GlmM F154I variant, indicating that the phenylalanine residue at position 154 in GlmM is important for controlling CdaA ac-tivity in vivo.

The E. coli GlmM enzyme did not inhibit the production of c-di-AMP in L. monocytogenes during adaptation to osmotic stress. To conclude, GlmM also inhibits CdaA in L. monocyto-genes and the GlmM-dependent control of c-di-AMP synthesis depends on osmotic stress.

3.4 Discussion

In the present study, we have confirmed that CdaR directly interacts with CdaA and modulates the activity of the diadenylate cyclase of L. monocytogenes (FIG 2). As previously reported, CdaR and CdaA form a complex in B. subtilis and L. monocytogenes (Gundlach et al., 2015;

Rismondo et al., 2016; Zhu et al., 2016). However, due to the fact that the membrane topology of CdaR was unknown, it was unclear which domain of the regulatory protein interacts with CdaA. Our topology analysis suggests that the YbbR domains of CdaR are exposed to the peptidoglycan layer of the cell envelope (FIG 1, FIG 8). Therefore, the interaction between CdaR and CdaA, and thus the control of the diadenylate cyclase, is very likely to be mediated through the transmembrane helices. Indeed, the cellular c-di-AMP levels were strongly reduced in a L. monocytogenes cdaR mutant strain overproducing only the transmembrane helix of CdaR (FIG 6D). The molecular details underlying the inhibition of CdaA by the transmem-brane helix and the extracytoplasmic signal perceived by the YbbR domains remain to be elu-cidated.

Previously, we have reported that CdaR inhibits CdaA in L. monocytogenes cells that were cultivated in BHI rich medium (Rismondo et al., 2016). The inhibition of CdaA by CdaR was also shown in S. aureus (Bowman et al., 2016). Interestingly, the inhibitory effect of CdaR was increased under acidic conditions, a phenomenon that remains to be resolved. CdaR of S. au-reus is also capable of reducing c-di-AMP production by CdaA in the environment of an E.

coli cell (FIG 6F) (Zhu et al., 2016).

Here we have observed that the overproduction of CdaR may also stimulate c-di-AMP synthe-sis in L. monocytogenes cells that were cultivated in LSM defined medium (FIG 6D). Moreo-ver, for B. subtilis it has been shown that CdaR stimulates the activity of CdaA when the cdaR and cdaA genes are co-expressed in E. coli (Mehne et al., 2013). Thus, depending on the growth conditions and the cellular environment, CdaR either inhibits or activates the diadenylate cyclase CdaA. Since CdaR is facing towards the peptidoglycan layer of the cell envelope it is tempting to speculate that the YbbR domains perceive mechanical shear forces arising as a result of a displacement of the membrane and the peptidoglycan in response to an osmotic up- or downshift (FIG 8). This model implies that the YbbR domains of CdaR interact with the peptidoglycan, or with proteins that are embedded in the cell wall, or that the reported self-interaction of the YbbR domains effect the self-interaction of CdaA via the transmembrane domains (Rismondo et al., 2016).

However, so far, we were unable to detect an interaction between the YbbR domains and the peptidoglycan layer (unpublished data). Thus, the signals that are perceived by the YbbR do-mains of CdaR and transmitted by the transmembrane helix to CdaA remain to be identified.

Structural studies of the YbbR domains I and IV of the CdaR protein from Desulfitobacterium hafniense Y51 uncovered similarities to the C-terminal domains of the TL5 and L25 ribosomal proteins (Barb et al., 2011). However, the potential interaction partners of C-terminal domains,

which are exposed to the surface of the ribosome are currently unknown. It will be interesting to find out whether the YbbR domains of different proteins respond to similar or different stimuli.

In the present study, we have also demonstrated that the phosphoglucosamine mutase GlmM directly interacts with CdaA and inhibits the activity of the cyclase in vitro and in vivo (FIG 7B and 7D). Furthermore, we could show that only the native phosphoglucosamine mutase is capable of inhibiting CdaA when the cells encounter hyperosmotic growth conditions (FIG 7D). Since the formation of a CdaA-GlmM complex has also been reported to occur in bacteria like B. subtilis, L. lactis and S. aureus, which are phylogenetically related to L. monocytogenes (Gundlach et al., 2015; Zhu et al., 2016; Tosi et al., 2019), the GlmM-dependent control of CdaA seems to be specific and a conserved among species possessing the cdaA-cdaR-glmM module. The specificity of the interaction between GlmM and CdaA is further supported by the observation that the replacement of the phenylalanine (F) at position 154 in GlmM by either isoleucine (I) or alanine (A) decreases the ability of the enzyme to inhibit CdaA (FIG 7B). The amino acid at the position 154 was previously reported to be important for the GlmM-depend-ent control of CdaA activity in the L. lactis strain MG1363 (FIG 7A) (Zhu et al., 2016). How-ever, albeit to a lesser extent, the GlmM F154I and F154A variants of L. monocytogenes still inhibited CdaA, indicating that additional residues of the phosphoglucosamine mutase are in-volved in the formation of the protein complex. Indeed, a recent structural model of the CdaA-GlmM complex from S. aureus revealed that other amino acid residues surrounding the phe-nylalanine 154 might be important for the regulatory protein-protein interaction (Tosi et al., 2019). However, a more comprehensive mutational study and a structural analysis of the CdaA-GlmM complex might provide insights into the molecular details of the interaction.

Given the fact that the GlmM enzyme has an additional function beside its role in providing precursors for cell wall biosynthesis, the phosphoglucosamine mutase can be assigned to the class of “moonlighting proteins” (Jeffery, 2019). In general, moonlighting proteins perform two or more distinct and physiologically relevant biochemical or biophysical functions in the cell (Jeffery, 2019). Research in recent years has shown that the phenomenon of moonlighting is widespread among bacterial proteins. For instance, the B. subtilis and E. coli UDP-glucose diacylglycerol glucosyltransferases UgtP and OpgH, respectively, which are active in lipid me-tabolism, act as metabolic sensors coupling the nutritional availability to cell division and thus cell size (Weart et al., 2007; Hill et al., 2013). The glutamate dehydrogenases RocG and GudB1 of B. subtilis are active in glutamate degradation and in controlling de novo synthesis of gluta-mate (Commichau et al., 2007a; Stannek et al., 2015). For many moonlighting enzymes, the signals controlling the secondary function of the enzymes have been identified (Commichau and Stülke, 2008). However, this is not the case for GlmM, which controls the activity of CdaA and thus the uptake of osmolytes via transporters whose activities are regulated by c-di-AMP

(Stülke and Krüger, 2020). Even though osmoregulation has been intensively studied in bacte-ria, it is still rather unclear how the cell senses the environmental osmolarity to adjust the turgor accordingly. The present study and a previous report revealed that GlmM alone is sufficient to inhibit CdaA (Tosi et al., 2019) and the membrane bound CdaR protein rather plays a minor role in modulating the CdaA activity in these coccoid bacteria. This idea is supported by the finding that some bacterial isolates like the L. lactis strain MG1363, which does not produce a functional CdaR protein, still adjusts the cellular c-di-AMP levels by employing GlmM (Zhu et al., 2016).

So how does a cell synthesizing CdaA and GlmM sense osmotic up- and downshifts and how is c-di-AMP production regulated by employing the phosphoglucosamine mutase? Over the past years it has been observed that in many bacteria c-di-AMP is controlling the uptake and efflux of potassium ions to adjust the cellular turgor (Stülke and Krüger, 2020). Thus, the cel-lular potassium concentration could control the regulatory interaction between GlmM and CdaA. However, in addition to potassium, other osmolytes like glycine betaine and carnitine are taken up by c-di-AMP-regulated transporters (Commichau et al., 2018; Stülke and Krüger, 2020). Therefore, it can be assumed that a mechanism, which is independent of a specific os-molyte, ensures the adjustment of the cellular turgor.

It would be an attractive idea to assume that the volume changes in response to an osmotic up- or downshift would result in a transient change in the cellular GlmM concentration, which in turn could affect c-di-AMP synthesis. The adjustment of the c-di-AMP levels would then lead to an inhibition and activation of osmolyte uptake and export, respectively, to adjust the cellular turgor to the environment. However, the most exciting question of how the cell uses c-di-AMP to adapt the turgor to the environmental osmolarity remains to be answered.