Regulatory mechanisms of c‐di‐AMP synthase from Mycobacterium smegmatis revealed by a structure: Function analysis

Abstract

Cyclic‐di‐nucleotide‐based secondary messengers regulate various physiological functions, including stress responses in bacteria. Cyclic diadenosine monophosphate (c‐di‐AMP) has recently emerged as a crucial second messenger with implications in processes including osmoregulation, antibiotic resistance, biofilm formation, virulence, DNA repair, ion homeostasis, and sporulation, and has potential therapeutic applications. The contrasting activities of the enzymes diadenylate cyclase (DAC) and phosphodiesterase (PDE) determine the equilibrium levels of c‐di‐AMP. Although c‐di‐AMP is suspected of playing an essential role in the pathophysiology of bacterial infections and in regulating host‐pathogen interactions, the mechanisms of its regulation remain relatively unexplored in mycobacteria. In this report, we biochemically and structurally characterize the c‐di‐AMP synthase (MsDisA) from Mycobacterium smegmatis. The enzyme activity is regulated by pH and substrate concentration; conditions of significance in the homoeostasis of c‐di‐AMP levels. Substrate binding stimulates conformational changes in the protein, and pApA and ppApA are synthetic intermediates detectable when enzyme efficiency is low. Unlike the orthologous Bacillus subtilis enzyme, MsDisA does not bind to, and its activity is not influenced in the presence of DNA. Furthermore, we have determined the cryo‐EM structure of MsDisA, revealing asymmetry in its structure in contrast to the symmetric crystal structure of Thermotoga maritima DisA. We also demonstrate that the N‐terminal minimal region alone is sufficient and essential for oligomerization and catalytic activity. Our data shed light on the regulation of mycobacterial DisA and possible future directions to pursue.

Short abstract

PDB Code(s): EMDB-33540;PDB-7Y0D

1. INTRODUCTION

Signaling cascades perceiving the environment and its perturbations are crucial to all organisms. In bacteria, several secondary messengers have been identified that coordinate signal transduction pathways during alterations in conditions, such as temperature, nutrient concentration, and pH. These molecules control cellular physiology by interacting with proteins or riboswitches (Abdul‐Sater et al., 2013; da Aline Dias et al., ²⁰²⁰; Gupta et al., ²⁰¹⁵; Gupta et al., ²⁰¹⁶; Hauryliuk et al., ²⁰¹⁵; Kalia et al., ²⁰¹³; Petchiappan et al., ^2020a; Petchiappan et al., ^2020b; Romling et al., ²⁰¹³; Stulke & Kruger, ²⁰²⁰; Yin et al., ²⁰²⁰). They include cyclic AMP (cAMP), cyclic GMP (cGMP), cyclic di‐GMP, cyclic di‐AMP (c‐di‐AMP), cyclic GAMP (cGAMP), and noncyclic pGpp or (p)ppGpp. The regulatory functions of cAMP, c‐di‐GMP, and (p)ppGpp signaling have been extensively studied, and they act as global regulators of bacterial lifestyle during stress. In bacteria, c‐di‐GMP coordinates the transition from a motile to a sessile state (Romling et al., 2013) and is also known to regulate cellular functions during biofilm formation, motility, virulence, cell cycle, antibiotic resistance, and starvation (Romling et al., 2013). The (p)ppGpp pathway primarily targets the transcription process to regulate cellular physiology (Hauryliuk et al., 2015). Functions critically controlled by (p)ppGpp include survival under amino acid starvation, biofilm formation, cell cycle, and virulence. Unlike c‐di‐GMP or (p)ppGpp, there have been limited reports regarding the regulatory mechanisms of c‐di‐AMP and cGAMP (Corrigan & Gründling, 2013; Stulke & Kruger, ²⁰²⁰). Although, recent advancements in c‐di‐AMP signaling studies elucidated its importance in many regulatory processes like osmoprotection, ion homeostasis, host‐pathogen interaction, etc. (Stülke & Krüger, 2020). Current data indicate that chemotaxis, virulence, and exoelectrogenesis are the major processes influenced by the cGAMP pathway (da Aline Dias et al., ²⁰²⁰).

The first report on c‐di‐AMP came from structural investigations of the bacterial DNA integrity scanning protein (DisA) from Thermotoga maritima (Witte et al., 2008). Since its discovery, the molecule has been linked to several signaling processes, including osmoregulation, DNA integrity maintenance, sporulation, cell‐wall homeostasis, cell‐wall biosynthesis, ion‐channel homeostasis, antibiotic resistance, virulence gene expression, acid resistance, and carbon metabolism (Corrigan et al., 2013; Fahmi et al., ²⁰¹⁷; Stulke & Kruger, ²⁰²⁰; Zeden et al., ²⁰¹⁸). In mycobacteria, c‐di‐AMP regulates fatty acid synthesis and DNA repair (Zhang, Li, & He, 2013). Bacterial c‐di‐AMP secretion into the host cytosol has been reported in intracellular pathogens like Listeria monocytogenes and Mycobacterium tuberculosis (Dey et al., 2015; Woodward et al., ²⁰¹⁰), and bacteria‐derived c‐di‐AMP is known to trigger the expression of inflammatory molecules like interferon‐1 (INF‐1) (Abdul‐Sater et al., 2013; Burdette et al., ²⁰¹¹; Manzanillo et al., ²⁰¹²; Sauer et al., ²⁰¹¹; Zhang et al., ²⁰¹¹; Zhang, Shi, et al., ²⁰¹³).

The synthesis of c‐di‐AMP is catalyzed from two molecules of ATP by a di‐adenylate cyclase (DAC) domain‐containing protein. c‐di‐AMP is hydrolyzed into pApA or two molecules of AMP by a phosphodiesterase (PDE). In bacteria, five classes of DAC domain‐containing proteins (DisA, CdaA, CdaS, CdaM, and CdaZ) have been reported (Petchiappan et al., ^2020a; Zhang & He, ²⁰¹³). Among these, DisA is an octameric protein that is considered bi‐functional. For instance, in Bacillus subtilis it has been shown to bind to Holliday junction DNA, and to synthesize c‐di‐AMP (Witte et al., 2008). DisA consists of three domains; an N‐terminal DAC domain connected to the C‐terminal DNA‐binding domain by a specific linker domain (Domain‐2) (Witte et al., 2008). The enzyme's active site is at the interface between two DAC domains (Witte et al., 2008), and the synthesis of c‐di‐AMP depends on metal ions such as Mg²⁺ and Mn²⁺. The cellular homeostasis of c‐di‐AMP is modulated by a specific PDE containing DHH‐DHHA1 or HD (His‐Asp) domain and encoded in a different operon (Commichau et al., 2019; He et al., ²⁰¹⁶; Tang et al., ²⁰¹⁵).

In gram‐positive bacteria, such as Staphylococcus aureus, B. subtilis, and Listeria monocytogens, c‐di‐AMP is essential for the growth under stress conditions and deletion of disA and pde shows a lethal phenotype (Corrigan & Gründling, 2013; Fahmi et al., ²⁰¹⁷; Whiteley et al., ²⁰¹⁵). In contrast, in Mycobacterium smegmatis disA deletion barely affected bacterial survival but negatively influenced bacterial C_12—C₂₀ fatty acids production. Conversely, the M. smegmatis pde deletion mutant showed higher intracellular C_12—C₂₀ fatty acids (Tang et al., 2015). The radiation‐sensitive gene A (radA) in M. smegmatis is located in the same operon as disA. It has been reported that RadA protein physically interacts with the DisA and negatively modulates c‐di‐AMP synthesis (Zhang & He, 2013). However, in Mycobacterium tuberculosis it has also been reported that pde deletion resulted in reduced virulence in an infection model (Yang et al., 2014). M. smegmatis is primarily a soil bacterium often used as a surrogate model to study M. tuberculosis biology. As a soil bacterium, M. smegmatis has evolved in the constant presence of environmental stresses like pH (acidic and alkaline), starvation, osmolytic stress, and temperature. On the other hand, adaptations to challenges, such as pH and starvation would be crucial to M. tuberculosis in its life cycle as an intracellular pathogen. Thus, second messenger signaling is likely to play essential roles in such adaptations in both these bacteria.

As a recently discovered second messenger, the role of c‐di‐AMP in stress management is not elucidated clearly in the mycobacterial system. Hence, we systematically analyzed the c‐di‐AMP synthase protein MsDisA (MSMEG_6080) from M. smegmatis mc ² 155 (Petchiappan et al., ^2020a; Tang et al., ²⁰¹⁵). This protein shares 83.66% identity with M. tuberculosis c‐di‐AMP specific‐DAC, making MsDisA an excellent model for studying c‐di‐AMP regulation in Mycobacterium. Our biochemical data reveal that MsDisA has higher activity at alkaline pH, and it rapidly converts two molecules of ATP to c‐di‐AMP without forming any intermediates. At neutral pH, MsDisA activity is reduced, and c‐di‐AMP is produced via intermediates, ppApA and pApA. We also found that increasing concentrations of ATP inhibited c‐di‐AMP synthesis by MsDisA and the kinetics followed the substrate‐inhibited model. An earlier report showed that B. subtilis DisA (BsuDisA) binds to the DNA and negatively regulates c‐di‐AMP synthesis (Witte et al., 2008). Interestingly, the DNA‐protein interaction assay revealed that MsDisA is unable to bind any type of DNA and c‐di‐AMP synthesis continues uninhibited. Biophysical characterization and electron microscopy (EM) image analysis revealed a substrate‐induced change in MsDisA structure. Electron cryomicroscopy (CryoEM) reconstruction of MsDisA at an overall resolution of 3.1 Å demonstrated an asymmetric assembly of the protein. A minimal N‐terminal region of MsDisA was identified by mutational studies to be sufficient for the activity and oligomerization. This report attributes a potential novel function to M. smegmatis c‐di‐AMP in regulating the alkaline stress response and gives new insights into the regulation of c‐di‐AMP synthesis facilitated by a flexible catalytic core in MsDisA.

2. RESULTS

2.1. Biochemical and functional characterization of MsDisA proteins

The disA gene from M. smegmatis mc²155 was cloned in pET28a and expressed in Escherichia coli BL21 (DE3) and the recombinant MsDisA protein was purified by affinity chromatography. The monomeric molecular weight estimated from SDS‐PAGE analysis of purified His‐tagged MsDisA matches the expected size of the protein. The identity of the protein was confirmed by peptide mass fingerprinting (PMF) analysis, and the molecular weight was found to be 41.3 kDa. Size exclusion chromatography‐multi angle light scattering (SEC‐MALS) study revealed the molecular mass of MsDisA to be 334.5 kDa (Figure S1A), suggesting an octameric assembly. This was also supported by negative stain EM analysis (discussed below).

We assayed the c‐di‐AMP synthesis activity of recombinant MsDisA, which contains DAC domain using ATP as substrate by high‐performance liquid chromatography (HPLC) with the detector set to 254 nm to analyze the nucleotide products (Bai et al., ^2012a; Manikandan et al., ^2014a). Schematic representation of c‐di‐AMP synthesis by DisA is shown in Figure 1a. Commercially available AMP, ATP, and c‐di‐AMP were used as controls. These controls eluted from the C‐18 HPLC column at 12.2, 17.4, and 23.7 min, respectively (Figure 1b). The propensity of MsDisA to synthesize c‐di‐AMP was assessed at different pH (5.4, 7.5, and 9.4) and salt concentrations (75–500 mM). The incubation of MsDisA with ATP for 4 h at pH 5.4 and 7.5 resulted in two distinct product peaks eluting at 20.3 and 23.7 min (Figure 1c,d). However, when the reaction was conducted at pH 9.4, a single peak eluted at 23.7 min (Figure 1e). The area under the curve (AUC) was measured for each peak to quantify the product formed. Under these conditions, the maximum activity for MsDisA was observed at a pH of 9.4 and 75 mM NaCl (Figure S1B, C). A significant decrease in activity was observed at other pH and salt conditions. The resolved peaks at 23.7 and 20.3 min were collected and subjected to LC–MS analysis to confirm the molecular weight of the product compounds. The molecular weight of peak at 23.7 min is 657.06 Da (Figure S1D), which matches the theoretical molecular mass of 658.4 Da of c‐di‐AMP. We obtained molecular masses of 677.1 [M + H] + and 755 [M−H]−, for the peak at 20.3 min, which corresponds to the molecular weight of pApA and ppApA, respectively (Figure S1D). These two molecules could be either intermediates or by‐products of the reaction, and we next sought to differentiate between these possibilities.

FIGURE 1.

Biochemical analysis of the c‐di‐AMP synthase (MsDisA). (a) Schematic representation of c‐di‐AMP synthesis by di‐adenylate cyclase (DAC)‐domain containing proteins using ATP/ADP as a substrate. (b) HPLC profiles of commercial AMP, ATP, and c‐di‐AMP molecules (250 μM each). The elution volume of the nucleotide is 12.2, 17.4, and 23.7 min, respectively. (c and d) Analysis of the MsDisA (1 μM) with ATP (500 μM) in reaction products at 50 mM MES/Tris‐Cl and 75 mM NaCl for 4H. HPLC profile showing the c‐di‐AMP synthetic intermediates formation in the MsDisA reactions at pH 5.4 and 7.5. (e) HPLC profile showing the c‐di‐AMP synthetic without any intermediates formation in the MsDisA reactions at pH 9.4. (f) Substrate specificity of MsDisA in the presence of other nucleotides (500 μM). (g) Steady‐state enzyme kinetics plot for c‐di‐AMP synthesis at different conditions (pH 9.4, 7.5, and 5.4).

Upon incubation of the MsDisA at pH 7.5 for 12 h, the intermediate peak at 20.3 min was undetectable in the products and the HPLC profile contained a single product peak at 23.7 min representing c‐di‐AMP. This allowed us to infer that the compounds (pApA and ppApA) corresponding to the peak at 20.3 min are likely formed as synthetic intermediates that are converted to c‐di‐AMP by the enzyme over a longer period (Bai et al., ^2012a; Manikandan et al., ^2014a). We propose that the synthesis of the c‐di‐AMP goes through intermediates pApA and ppApA, although our attempts to resolve the peak at 20.3 min to pApA and ppApA were not successful. However, on probing the substrate specificity of MsDisA using a variety of adenosine nucleotides, it was found that ATP was by far the best substrate for MsDisA, and the enzyme hardly hydrolyzed AMPCPP or AMPPCP (Figure 1f). This suggests a direct participation of α‐β and β‐γ phosphates of ATP at the active site of the enzyme, and further strengthens the hypothesis that pApA and ppApA are indeed reaction intermediates.

2.2. Substrate concentration modulates MsDisA activity

The c‐di‐AMP molecule has been referred to as an “essential poison” as it is indispensable for bacterial survival under stress, and toxic when over‐accumulated (Gundlach et al., 2015). Hence, an accurate balance between its synthesis and physiological requirements is imperative in bacterial homeostasis. M. smegmatis is a fast‐growing soil bacterium, and the activity of its various proteins may change differently according to the changing environment.

Similarly, intracellular pathogen M. tuberculosis gets exposed to various stress inside the host cell, for instance, it encounters pH stress in the macrophage. Thus, we first performed diverse functional analyses to determine the optimum condition of MsDisA activity. Then, we decided to estimate the steady‐state kinetic parameters to follow the rate of synthesis of c‐di‐AMP by MsDisA in different conditions. Using the AUC values of the HPLC peaks at different nucleotide concentrations, standard curves were prepared for c‐di‐AMP and ATP (data not shown).

To calculate the kinetic parameters of MsDisA, we plotted V_max against substrate concentrations for reactions carried out at pH 5.4, 7.5, and 9.4, at a range of substrate concentrations. The kinetic data followed a substrate inhibition model following the “equation 1” shown in the material and method section, with an R ² value of 0.95 rather than the standard Michaelis–Menten analysis or Lineweaver–Burk plot. V_max, K_m, and K_i values for the enzyme were obtained from the equation (Table 1). Interestingly, the in vitro activity of MsDisA was significantly inhibited by high ATP concentrations (K_i > 2.8 μM) at pH 9.4, suggesting a regulatory potential for the cellular ATP pool over MsDisA enzymatic activity. The activity of MsDisA also followed the substrate inhibition model at pH 5.4 and 7.5 (Figure 1g). However, the inhibition was less dramatic at these pH, given the lower rate of synthesis of c‐di‐AMP at pH 5.4 and 7.5.

TABLE 1.

Enzyme kinetics of c‐di‐AMP synthase (MsDisA) protein at different pH of the buffer

MsDisA	pH 5.4	pH 7.5	pH 9.4
Vmax (mM/min)	0.15	0.21	1.44
Km (mM)	1.10	0.48	0.94
Ki (mM)	3.30	1.43	2.81

2.3. MsDisA neither binds DNA nor is its enzymatic activity influenced by DNA

Experiments with B. subtilis DisA have shown that the enzyme binds to 4‐way DNA junctions with high affinity in vitro, and this interaction leads to allosteric inhibition of the c‐di‐AMP synthesis activity (Witte et al., 2008). The sequence alignment between the C‐terminal DNA‐binding domains of BsuDisA and MsDisA showed 40% identity at the HhH region (Heger et al., 2014) (Figure 2a). Therefore, we were curious about the DNA‐binding characteristics of MsDisA and the regulatory influence, if any, on the enzyme's catalytic activity. To assay DNA‐protein interaction, we employed electrophoretic mobility shift assays (EMSA) using MsDisA with Holliday junction DNA (HLDNA). In the conditions tested, we did not observe a shift corresponding to DNA‐protein complex formation indicating that MsDisA does not bind DNA whereas control BsuDisA did show DNA‐protein complex formation (Torres et al., 2021) (Figure 2b). Furthermore, HPLC analysis indicated that the presence of DNA in the reaction had no influence on c‐di‐AMP synthesis rates compared to the control without DNA (Figure 2c).

FIGURE 2.

DNA‐binding assay (EMSA) and activity of c‐di‐AMP synthase (MsDisA) in the presence of DNA. (a) Multiple sequence alignment of DisA protein from Mycobacterium smegmatis, Mycobacterium tuberculosis, Thermotoga maritima, Bacillus subtilis, and Clostridium botulinum at the C‐terminal DNA‐binding region. The secondary structure of MsDisA is shown on top of the alignment, and conserved residues are shaded in red. (b) EMSA was performed with varying concentrations of BsuDisA at pH 7.5 and MsDisA at both pH's (7.5 and 9.4) in the presence of 1 μM holliday junction DNA (HLDNA). Lanes 1, 6, and 11 are only DNA controls, whereas Lanes 2–5 are BsuDisA protein incubated with HLDNA. BsuDisA shows an evident shift in the presence of HLDNA, which is used as a control. In the case of MsDisA, HLDNA did not show any shift in both pH conditions (Lane 7–10 is pH 7.5 and Lanes 12–15 is pH 9.4). (c) HPLC profile of MsDisA activity assay with or without DNA (dsDNA and HLDNA) in the presence of 500 μL ATP and Mg²⁺. The primary change in the enzyme assay was that no EDTA was used.

2.4. MsDisA protein undergoes substrate‐induced conformational changes

The first reported crystal structure of TmDisA shows that c‐di‐AMP is tightly bound to the active site during the purification of the enzyme (Witte et al., 2008). A similar observation was made with MsDisA, where c‐di‐AMP co‐purifies, which was confirmed with mass spectrometry (Figure S1E,F). Given the substrate‐inhibited mode of action of MsDisA, we were interested in identifying the likely structural changes in the protein upon substrate binding. Toward this end, we performed CD spectroscopy and negative staining transmission electron microscopy (TEM) of MsDisA protein with and without substrate (ATP). CD data suggested that the product‐bound MsDisA protein is predominantly α‐helical (Figure 3a). Incubation of MsDisA with ATP at pH 7.5 led to a shift in the minima at 222 to 208 nm (Figure 3a). This phenomenon was specific to the cognate substrate as when the CD spectrum of MsDisA was recorded with GTP, no changes in the far‐UV CD spectra were observed (data not shown). In order to map the substrate‐induced structural alterations further, we conducted microscopic analysis of MsDisA and MsDisA + ATP complex at pH 7.5. TEM images of MsDisA with and without ATP were collected. Reference‐free two‐dimensional (2D) class averages were generated with EMAN 2.0, and length and width of the protein were calculated from the raw micrographs (Figure 3b,c). The average length and width of the MsDisA protein with c‐di‐AMP bound were 17.58 ± 1.07 and 7.3 ± 0.7 nm (number of particles from raw micrographs [N] = 1050), respectively (Figure 3b). However, upon incubation with ATP, the length and width of the protein, presumably MsDisA + ATP complex, was significantly increased to 19.73 ± 0.95 and 8.3 ± 0.8 nm (N = 1100), respectively (Figure 3c). We propose that incubation with ATP displaces the existing c‐di‐AMP from the DAC domain of MsDisA protein accompanied by structural alteration in the protein, which is implicated in the CD analysis and TEM data.

FIGURE 3.

Substrate‐induced structural changes of the c‐di‐AMP synthase (MsDisA). (A) Far‐UV CD spectra of the MsDisA and MsDisA + ATP (500 μM) complex. (b and c) transmission electron microscopy (TEM) raw images and two‐dimensional (2D) class averages of the MsDisA co‐purified with c‐di‐AMP (particle length: 17.58 ± 1.07 nm) and MsDisA +500 μM ATP complex (particle length: 19.73 ± 0.95 nm) (box size is 240 pixels).

2.5. Cryo‐EM structures of the MsDisA

To determine the structure of MsDisA and the arrangement of the subunits, we performed single‐particle cryo‐EM analysis of the purified protein. Images of the DisA were collected on ice, and on holey carbon grid coated with a thin carbon layer. While the data set on ice revealed predominantly top/bottom views, the data on carbon adopted predominantly side views. We chose the latter for further data processing. Best 2D classes were selected to generate the initial model, followed by non‐uniform refinement with no symmetry imposed using cryoSPARC (Punjani et al., 2017) (Figure S2A). We also attempted refinement/reconstruction with D4 symmetry, but this did not lead to meaningful maps. The final three‐dimensional (3D) reconstruction is asymmetric, has an estimated overall resolution of 3.1 Å (Figure S2B, Table S1), and revealed the structure of distinct subunits (Figure 4a). For each subunit, the map appears better resolved at the core DAC domain when compared with the periphery. This visual observation is substantiated by the local resolution plot which shows that the core (DAC domain) was better resolved (resolutions between 2.8 and 3.8 Å), while the periphery of the molecule was less resolved between 4.3 and 7.3 Å (Figure S2C). The EM maps at periphery show a certain degree of “breathing” of the MsDisA assembly. The quality of the map also varies across the different subunits. Due to the differential resolution across the map, and the single B‐factor‐sharpening resulting in the smearing of the density, we used multiple other maps. These included maps calculated with deepEMhancer (Sanchez‐Garcia et al., 2021), unsharpened map and maps sharpened with different B‐factors for model building. The monomeric structure of MsDisA predicted using AlphaFold was used as the starting model (Jumper et al., 2021) (Figure 4b). The α‐helices and β‐strands docked accurately near the core of the enzyme (Figure S2D). The model was manually inspected and rebuilt with Coot (Emsley & Cowtan, 2004) (Figure 4c). As expected, the fitting of the refined model is better at the core of the enzyme and varies within the protomers (Figure S3A and Table S2). Fourier shell calculation of the map against the model (at 0.5) indicates an overall resolution of 4.4 Å (Table S2).

FIGURE 4.

Cryo‐EM structure of Mycobacterium smegmatis DNA integrity scanning protein. (a) The cryo‐EM map of the octameric MsDisA with each subunit colored individually and shown in different views. (b) Alphafold2 predicted protomer model of MsDisA (Uniport ID‐ A0R564). (c) Final models of individual protomer fit into the cryo‐EM map (gray) and the refined final model of MsDisA. The subunits are colored as in panel (a).

The octameric model is shown with each protomer colored differently in Figure 4c. Further analysis of the MsDisA structure shows that each protomer could make polar contacts with more than two protomers through the DAC domain and the linker domain (Figure S3B). However, The N—N terminal DAC domain interaction is seen only in six protomer chains but absent in the other two protomers, thus, giving rise to an open structure where C‐terminal regions are splayed apart. In this open structure, two opposing protomers are separated by a distance of ~31 Å, disrupting the symmetry (Figure 5a,b). Consequently, this structure differs from the published T. martima DisA structure (Witte et al., 2008) which is founded upon a symmetric model.

FIGURE 5.

Intermolecular chain interaction of c‐di‐AMP synthase (MsDisA) compared to that of TmDisA. (a) Two opposing protomers are separated by ~31 Å (chain CE and chain BF), whereas adjacent chain EF and chain BC are separated by ~28 Å in MsDisA octamer. The rest of six opposing protomers (Chain AB, DF, and GH) participate in similar interaction as TmDisA. (b) Octameric TmDisA (PDB‐ 3C1Y) forms same dimeric interface across the molecule.

To highlight the differences between TmDisA and MsDisA proteins, we colored the surface model of TmDisA and the cryo‐EM map of MsDisA with a color scheme correlated to the domain architecture (Figure 6a). Both the proteins comprise of an N‐terminal conserved diadenylate cyclase DAC domain and a C‐terminal DNA‐binding domain connected by a long α‐helical linker domain. The N‐terminal of MsDisA (DAC domain) is defined until residue 155 (19–155aa) and it is composed of five α‐helices and five β‐sheets as shown in blue, followed by a linker domain consisting of five α‐helices (156–316aa) colored orange, and a C‐terminal DNA‐binding domain consisting of four α‐helices (317–364aa) colored green (Figure 6a–c). The differences between the DAC domains of MsDisA and TmDisA are shown in the enlarged view (Figure 6b,c). The superimposition of MsDisA octameric assembly with that of TmDisA shows deviation with an RMSD of 32.15 Å (Figure 6d). The protomers of MsDisA and TmDisA were superimposed with an overall RMSD of 3.09 Å revealing maximum deviation at the HhH and DAC domains. In contrast, the linker domain superimposes much better. However, individual domains superimposed with an average RMSD of 1.10 Å, suggesting that the individual domains have a common fold (Figure 6d). Individual DAC domain when superimposed shows that the catalytic residues present at the nucleotide‐binding site are indeed similar in both MsDisA and TmDisA structure (Figure 6d). The one major difference is the presence of H137 in MsDisA in the place of R128 of TmDisA.

FIGURE 6.

Structural comparison of c‐di‐AMP synthase (MsDisA) and TmDisA. (A) Domain architecture of MsDisA. (b and c) Structural comparisons of MsDisA and TmDisA (PDB‐ 3C1Z) show the interchain interactions' differences. An enlarged view of DAC domain is shown within red box (Goddard et al., 2018). (d) Overlay of MsDisA (green) and TmDisA (red) octamer, monomer, and their DAC domains (boxed) (DeLano, ^2002b). Catalytic residues are shown in ball and stick model. MsDisA residues are boxed in green.

2.6. Molecular docking reveals the substrate‐recognizing residues

We also observed nonprotein densities at the dimeric interface of the protomer N terminal regions (Figure 7a, b). The residues that form interactions with extra density correspond to those in the nucleotide‐binding sites as shown in T. martima (Müller et al., 2015). The density quality does not allow for confident model building of the ligand molecule. However, this is likely to be c‐di‐AMP (Figure 7b) as deduced from mass spectrometry (Figure S1E,F) and supported by the observation in TmDisA, where the secondary messenger co‐purified with the enzyme (Witte et al., 2008). This asymmetrical structure is particularly relevant as it shows a potential formation of a dynamic, catalytically competent DAC‐DAC domain interface within DisA octamer. We used a molecular docking approach based on Schrödinger Glide to analyze the critical active site residues involved in substrate (ATP) binding (Friesner et al., 2004). The ligand ATP was docked in the cryo‐EM MsDisA model and with TmDisA as control (Figure 7c,d). The docking studies identified highly conserved residues crucial for ATP binding in MsDisA, such as D84, Q102, L103, R117, H118, and H137 (docking score − 9.41, glide energy −47.18 kcal/mol).

FIGURE 7.

The extra density at the interface and molecular docking. (a) The additional density observed in the putative nucleotide binding site of 6 monomers and the surrounding regions are shown. The EM map from deepEMhancer is in blue and the difference map from Servalcat (Yamashita et al., 2021) is shown in green. The arrow points to the extra density. Two other monomers do not have this extra density as these regions are poorly resolved. (b) The dimer interface of c‐di‐AMP synthase (MsDisA) with the difference density for the ligand in green indicates that the c‐di‐AMP could have been co‐purified as observed for TmDisA (Witte et al., 2008), but the c‐di‐AMP has not been modeled. (c) Molecular docking analysis of MsDisA active site with ATP (Friesner et al., 2004). (d) Molecular docking analysis of TmDisA (3C1Z) active site and ATP as a control (Friesner et al., 2004).

As a proof of principle experiment, selected critical residues for ATP binding or c‐di‐AMP release were mutated, and activity analyses of the resultant proteins were carried out. These studies involved four single‐point mutations in MsDisA. The mutant MsDisA proteins D84A, D84E, and H137A were inactive and completely failed to synthesize c‐di‐AMP, corroborating the predictions from the docking studies (Figure S4A–D). Residue D84 likely acts as a nucleophile and attacks the 3′OH of ATP molecule to form phosphodiester bond with neighboring ATP molecule. Even when D84 was mutated to glutamic acid, the protein showed no c‐di‐AMP activity, suggesting the side chain position also plays a crucial role. We anticipate that H137 is involved in base stacking with the substrate, which gets hampered when alanine is introduced in H137A. Another residue H118, when mutated to alanine did not abolish activity but reduced the catalytic efficiency. (Figure S4E). According to the docking study, R117 and H118 play an essential role in recognizing the substrate through hydrogen bonding with the phosphate groups of substrates.

2.7. N‐terminal domain of MsDisA is sufficient for c‐di‐AMP synthesis and octamerization

To check for the functional relevance of the three domains of MsDisA in oligomerization and catalysis, we generated truncates of MsDisA; C‐terminal deletion truncate (MsDisA_1–316)_, C‐terminal and linker domain deletion truncate (MsDisA_1–158), and N‐terminal deletion truncate (MsDisA_132–372) based on the domain architecture (Figure 6a). These truncated constructs were expressed and purified using the same protocol as full‐length (FL) MsDisA. The truncated proteins were analyzed for oligomerization status (SEC‐MALS and TEM) and their c‐di‐AMP synthesis ability (HPLC) compared using wild‐type FL MsDisA. C‐terminal deletion truncate (MsDisA_1–316) forms an octameric assembly with a molecular mass of 287 kDa (Figure 8a), and the TEM study confirms the same (Figure 8b). C‐terminal deleted protein actively synthesizes c‐di‐AMP at pH 9.4 suggesting no role of the C‐terminal in diadenylate cyclase (DAC) activity (Figure 8c). The mutant MsDisA_1–158 also assembled as an octamer with a molecular mass of 145 kDa. It also synthesized c‐di‐AMP at pH 9.4 similar to wild‐type protein suggesting the linker and C‐terminal domain has no role in DAC activity (Figure 8d–f). Interestingly, both the mutants co‐purify with c‐di‐AMP present in the active site, strengthening the hypothesis that alone the DAC domain can synthesize the c‐di‐AMP (Figure S5A–D). The third mutant lacking the N‐terminal DAC domain (MsDisA132–372) was a dimer with a molecular mass of 55 kDa (Figure 8g). As expected, MsDisA_132–372 lacking the DAC region, showed no catalytic activity (Figure 8h). From this, we conclude that the nucleotide‐binding domain (DAC domain) is sufficient and essential for catalysis of c‐di‐AMP synthesis and oligomerization of the order same as the full‐length protein.

FIGURE 8.

Analysis of the MsDisA domain mutants. (a) Size exclusion chromatography‐multi angle light scattering (SEC‐MALS) profile of MsDisA_1–316 (0.5 mg/mL) indicates octameric molecular mass of 287.32 kDa. (b) TEM images of the MsDisA_1–316 (box size of 220 pixels, length: 15.77 ± 0.78). (c) HPLC profile of the activity analysis of the MsDisA_1–316 and the oligomeric model cartoon. (d) SEC‐MALS profile of MsDisA_1–158 (0.5 mg/mL) indicates octameric molecular mass of 145.76 kDa. (e) TEM images of the MsDisA_1–158 (box size of 200 pixels, length‐ 10.72 ± 2.49). (f) HPLC profile of the activity analysis of the MsDisA_1–158 with an oligomeric model. (g) SEC‐MALS profile of MsDisA_132–372 (0.5 mg/mL) indicates dimer molecular mass of 55.70 kDa. (h) HPLC profile of the activity analysis of the MsDisA_132–372 along with the dimeric model. The synthesis activity in each case was carried out as shown in Figure 1.

3. DISCUSSION

As a multi‐functional second messenger molecule, c‐di‐AMP has been implicated in functions such as potassium transport, osmotic stress, acid stress, cold stress, DNA repair, sporulation, drug resistance, genetic competence, central metabolism, day‐night cycle in cyanobacteria, fatty acid biosynthesis and biofilm formation (Bowman et al., 2016; Gándara & Alonso, ²⁰¹⁵; Gibhardt et al., ²⁰¹⁹; Kim et al., ²⁰¹⁵; Oppenheimer‐Shaanan et al., ²⁰¹¹; Peng et al., ²⁰¹⁶; Pham et al., ²⁰²¹; Rubin et al., ²⁰¹⁸; Tang et al., ²⁰¹⁵; Whiteley et al., ²⁰¹⁷) Besides facilitating bacterial adaptation to various environmental stress, c‐di‐AMP has also been shown to influence the virulence of several pathogenic bacteria (Fahmi et al., 2017) and the host immune responses (Dey et al., 2017; Woodward et al., ²⁰¹⁰). These diverse functions are mediated via c‐di‐AMP binding to a multitude of receptors belonging to various protein families, and riboswitches. It is thus a crucial alarmone in bacteria and archaea (Corrigan & Gründling, 2013).

In this work, we report the synthesis of c‐di‐AMP by M. smegmatis DisA, an enzyme orthologous to the putative di‐adenylate cyclases Rv3586 (MtbDacA) of M. tuberculosis, and DisA of B. subtilis. The c‐di‐AMP formation process by MtbDac has been reported to be a two‐step process via intermediates pppApA or ppApA at pH 8.5 (Manikandan et al., ^2014b). MsDisA produces c‐di‐AMP via two intermediates, ppApA, and pApA, at pH 7.5. However, the reaction is more rapid at pH 9.4, and forms c‐di‐AMP without yielding any detectable intermediates. The strikingly higher preference of the enzyme for ATP as a substrate over other nucleotides in our experiments strongly suggests the participation of α‐β and β‐γ phosphates of ATP at the active site of the enzyme.

Substrate‐induced inhibition of MsDisA by ATP that we observed is intriguing and alludes to the significance of balancing the second messenger levels as a function of cell growth. The inhibition by substrate was observed at all pH tested, albeit to varying degrees. This is akin to several classical instances of cellular homeostasis of second messengers (Christen et al., 2006), where the synthetic activity is regulated by substrates, or products, or both. The di‐guanylate cyclase (DGC) of Pseudomonas aeruginosa, responsible for c‐di‐GMP synthesis, is regulated by an allosteric binding site in the enzyme for c‐di‐GMP resulting in non‐competitive product inhibition (Dey et al., 2017). Similar enzyme inhibition was also reported in the case of Rv3586 (MtbDisA), where the diadenylate cyclase activity was negatively regulated by ATP or ADP (Manikandan et al., ^2014b). The MsDisA kinetic data suggest that the cellular ATP pool most probably plays a crucial role in regulating the c‐di‐AMP concentration in the cells. We presume that the higher ATP concentration during the exponential growth phase down regulates the c‐di‐AMP synthesis rate by allosterically inhibiting the enzyme activity, whereas the lower ATP pool at stationary growth phase may induce a higher c‐di‐AMP synthesis.

The cryo‐EM structure of MsDisA differs from that of the T. maritima DisA and we propose that the relatively open complex in the former might be necessary to accommodate ATP into active site for continuous synthesis of c‐di‐AMP in the cells (Figure 6b). Note that, the sample of MsDisA cryoEM was at pH 7.9, where the protein showed comparatively weaker activity than at pH 9.4. The effect of pH on the structure (or asymmetry) is an interesting line of research to be pursued. The C‐terminus of MsDisA (316‐372aa) appears to be a DNA‐binding domain, and the homologous enzyme BsuDisA binds to DNA (Witte et al., 2008). The sequence alignment of the C‐terminal DNA‐binding domains (Stothard, 2000) of BsuDisA and MsDisA showed 40% identity at the HhH region. Surprisingly, our experiments could not detect any DNA‐binding activity of MsDisA protein to Holliday junction DNA. In congruence, c‐di‐AMP synthesis was unaffected by the presence of DNA in the reaction mixture. According to literature, conformational changes at the HhH domain of BsuDisA octamer are important for binding to branched nucleic acids (Witte et al., 2008). Perhaps, MsDisA binds to other proteins such as MsRadA (Zhang & He, 2013), which might assist in DNA binding. Further analysis is required to pinpoint the exact role of the C‐terminal domain and if this is involved in binding to other molecules.

The results presented above also prompted us to study the function of individual domains of the MsDisA, in particular, the importance of the N‐terminal domain in oligomerization and the catalytic activity of MsDisA (Figure 8). We propose that octamerization of MsDisA occurs via interactions between N‐terminal domains, and the interface of N‐terminal domains in a tetramer acts as the substrate‐binding region of the enzyme. It has been shown earlier that the deletion of the C‐terminal domain makes the M. tuberculosis Rv3586 (DacA) a tetramer (Bai et al., ^2012a), while truncated MsDisA lacking the C‐terminal domain remains octameric. Thus, our biochemical data of the MsDisA deletion constructs differ from previous observations (Bai et al., ^2012a), and the minimum N‐terminal domain is essential and sufficient for both octamerization and c‐di‐AMP synthesis in MsDisA.

A number of factors render c‐di‐AMP signaling an attractive area of research. These include its near‐ubiquitous presence in several microorganisms and the multiple phenotypes associated with essential physiological functions. In addition, given that changes in c‐di‐AMP homeostasis affect virulence and overall fitness in pathogens such as M. tuberculosis (where DisA is the only DAC), the topic is an attractive target for further investigations from the standpoint of infections but also from mechanistic perspective.

4. EXPERIMENTAL PROCEDURES

4.1. Bacterial strains and growth condition

M. smegmatis mc²155 and E. coli strains used in this study are described in Table S3. M. smegmatis mc²155 strain was grown in MB7H9 medium (Difco) or MB7H9 medium solidified with 1.5% (w/v) agar with additional 2% glucose (vol/vol) and 0.05% Tween 80 (Bharati et al., 2018). E. coli strains DH5α or BL21 (DE3) were grown in Luria‐Bertani (LB) medium or in LB medium containing 1.5% (w/v) agar (Russell & Sambrook, 2001). Different antibiotics were used as required at the following concentrations: kanamycin (35 μg/mL) or ampicillin (100 μg/mL) for the respective E. coli strains.

4.2. Cloning and purification of MsDisA protein

To purify MsDisA as a C‐terminally histidine‐tagged protein, a DNA fragment (carrying disA gene/MSMEG_6080 / 1119 bp) was prepared using primers DisA1 and DisA2 (Table S4) and the M. smegmatis mc ² 155 genomic DNA as a template. The resulting PCR fragment was digested with NcoI and NotI and then ligated into the predigested pET28a plasmid. The pET28a plasmid carrying the disA gene was transformed into E. coli DH5α. The clone was confirmed by DNA sequencing (Sigma).

MsDisA (372 aa) protein was purified using standard Ni‐NTA column chromatography as described before (Bharati et al., 2012; Bharati et al., ²⁰¹⁸). Briefly, E. coli BL21 (DE3) cells containing plasmids for disA was inoculated in LB medium (supplemented with 35 μg/mL of kanamycin), followed by their growth overnight at 37°C. Secondary cultures were prepared by inoculating 1% of the primary culture and grown at 37°C with shaking till the OD₆₀₀ reached ~0.6. The cultures were then induced with 1 mM isopropyl β‐d‐thiogalactopyranoside (IPTG) for 3 h at 37°C. The cultures were harvested by centrifugation at 6000 rpm. The cells were then resuspended in lysis buffer (50 mM Tris‐Cl; pH 7.9, 300 mM NaCl and 1 mM phenyl‐methylsulfonyl fluoride [PMSF] and lysed using probe sonication). The lysate was centrifuged at 14,000 rpm to remove the cell debris. Then, the supernatant was loaded onto a Ni‐NTA column and the recombinant proteins were allowed to bind to the Ni‐NTA beads and then washed with 100 column volumes of wash buffer containing 40 mM imidazole. Finally, MsDisA protein was eluted with the help of elution buffer containing 50 mM Tris‐Cl; pH 7.9, 300 mM NaCl, and 300 mM imidazole. Different fractions collected during protein purification were analyzed by 10% SDS‐PAGE. The eluted proteins were dialyzed against Tris‐Cl buffer at pH 7.5, 300 mM NaCl/KCl for 12–16 h at 4°C or injected and purified on SEC (size exclusion chromatography) column Superose 200/12 10/300 (GE Health care) against a buffer containing 50 mM Tris‐Cl (pH 7.9) and 300 mM NaCl and stored at −80°C for future use.

4.3. Cloning and purification of domain variants and point mutant proteins

Using MsDisA plasmid as the template, single‐point mutants of MsDisA including D84A, D84E, H118A, and H137A were generated. The primers for mutations are listed in Table S4. Site‐directed mutagenesis (SDM) of Asp84, His118, and H137 residues was performed by standard procedures (Krishnan et al., 2016). The PCR products were transformed into E. coli DH5α and a plasmid containing the mutation were confirmed by sequencing. The plasmids were transformed into E. coli BL21 (DE3) cells and proteins were purified as described above.

Based on domain boundaries, we cloned and express the respective domains of MsDisA proteins, DNA fragments (for specific MsDisA domain) were generated by PCR using primers disA1‐158A1/disA1‐158A2 (only N‐Terminal domain‐1), disA1‐316A1/disA1‐316A2 (domains 1 and 2) and disA132‐372A1/ disA132‐372A2 (domains 2 and 3), respectively (Table S4). The amplicons for disA1‐158aa, disA1‐316aa, and disA132‐372aa were digested with NcoI/NotI, and cloned into the plasmid pET28a predigested with the same set of enzymes. The resulting plasmids pETdisA_1–158, pETdisA_1–316, and pETdisA_132–372 were transformed into E. coli BL21 (DE3) to express and purify as MsDisA mutant proteins. The protein purification was performed as described for the full‐length DisA with some modifications. Briefly, E. coli BL21 (DE3) cells containing different plasmids pET‐DisA_1–158 and pET‐DisA_1–316 were grown in LB‐broth to an OD₆₀₀ at ~0.6, and the cultures were induced with 1 mM IPTG, 3 h at 37°C, and then harvested at 6000 rpm and stored at −20°C for further use. For pET‐DisA_132–372, E. coli BL21 (DE3) containing plasmids were grown until OD reached 0.6 at 37°C, and cells were transferred to 18°C for 16 h and induced with 0.1 mM IPTG. Purification steps of the mutant proteins were followed per the protocol described for wild‐type full‐length MsDisA protein.

4.4. Size exclusion chromatography‐multi angle light scattering

SEC‐MALS experiment was performed to estimate the molecular mass and oligomerization of the purified proteins (MsDisA and other mutant proteins) with a standard procedure. Briefly, a Superdex 200 10/300 GL column (GE Health Care) was equilibrated with 50 mM Tris‐Cl buffer at pH 7.9 and 300 mM NaCl. Then, 0.5 mg/mL of purified proteins were injected into the column separately, and the flow rate was fixed at 0.5 mL/min. To determine the molecular mass of the proteins, a triple angle MALS detector (mini DawnTreos, Wyatt Technology), refractive index detector and UV detectors were used. Data analysis was finally done with the help of ASTRA software.

4.5. Activity assay of MsDisA and mutant proteins

The enzymatic assay of MsDisA and other mutant proteins was adapted from the protocol described previously with modifications (Bai et al., ^2012b; Bharati et al., ²⁰¹²; Christen et al., ²⁰⁰⁵). The assays were mainly aimed at studying the synthesis of c‐di‐AMP by MsDisA and the mutant MsDisA enzymes. Briefly, to follow c‐di‐AMP synthesis by MsDisA and the mutant proteins, samples were desalted in different buffer compositions (Buffer 1–50 mM MES pH 5.4, 300 mM NaCl/KCl; Buffer 2–50 mM Tris pH 7.5, 300 mM NaCl/KCl; and Buffer 3–50 mM Tris pH 9.4, 300 mM NaCl/KCl). MsDisA protein (1 μM) was incubated with 0.5 mM ATP and 5 mM MgCl₂ at 37°C. The reactions were monitored for c‐di‐AMP synthesis, and reactions were stopped by adding EDTA (10 mM), followed by centrifugation at 12 000 rpm for 30 min at 4°C. Supernatants from the reaction samples were collected and subjected to HPLC analysis or stored at −20°C for further use. We performed the activity assay separately at different salt and pH levels to determine the optimum pH and salt concentration of MsDisA activity. All the reactions were carried out following the same protocol as stated earlier.

4.6. HPLC analysis

The synthesis of c‐di‐AMP by MsDisA protein in different activity reactions was detected by HPLC analysis of the samples using previously described protocols (Bharati et al., 2012; Ryjenkov et al., ²⁰⁰⁵). A reverse phase C‐18 column (4.6 × 150 mm, Agilent Eclipse XDB‐C‐18) was used to separate the reaction mixture containing nucleotides by an HPLC using buffer A (100 mM KH₂PO₄, 4 mM tetrabutylammonium hydrogen sulfate, pH 5.9) and buffer B (75% (v/v) buffer A with 25% (v/v) Methanol) (Agilent 1200). Different concentration gradients of c‐di‐AMP were used to prepare a standard curve. AUC values in the HPLC for each concentration of c‐di‐AMP was plotted against respective c‐di‐AMP concentration to prepare a c‐di‐AMP standard curve. AUC value is an integrated measurement of a measurable effect or phenomenon. It is used here as a cumulative measurement to compare peaks in chromatography analysis (Gagnon & Peterson, 1998). The product formation of MsDisA and mutant proteins were calculated from the standard curve. Each sample peak area was determined independently three times.

4.7. Enzyme kinetics

The enzyme activities of the MsDisA protein were measured with varying substrate concentrations. MsDisA activity assay was performed for 4 h with variable ATP concentration. The reactions were performed using a standard protocol described earlier in the activity section (Bharati et al., 2012). The reactions were stopped with the addition of EDTA (10 mM) and followed by centrifugation of the samples at 12000 rpm at 4°C for 30 min. Samples were collected and subjected to HPLC analysis. The amount of c‐di‐AMP synthesized by MsDisA was calculated from the standard c‐di‐AMP curve at different concentrations. The kinetic parameters like K_m, K_i, and V_max were determined from the substrate inhibition model of enzyme kinetics(Copeland, 2000) using GraphPad Prism (version 5.02).

$$\mathsf{Y}={{\mathsf{V}}_{\mathsf{max}}}^{*}\mathsf{X}/({\mathsf{K}}_{\mathsf{m}}+{\mathsf{X}}^{*}\left(\mathsf{1}+\mathsf{X}/{\mathsf{K}}_{\mathsf{i}}\right)$$ (1)

V_max is the maximum enzyme velocity, expressed in the same units as the Y‐axis. K_m is the Michaelis–Menten constant, expressed in the same units as the X‐axis. K_i is the dissociation constant for substrate binding so that two substrates can bind to an enzyme. It is expressed in the same units as X, where X is the substrate concentration.

4.8. Electrophoretic mobility shift assay

Synthetic holiday junction DNA (HL‐DNA) (Table S5) is used to understand MsDisA protein‐DNA interaction via fluorescence‐based EMSA. The DNA fragment used in this study was used previously (Guy & Bolt, 2005; Witte et al., ²⁰⁰⁸). This experiment was performed using the EMSA assay kit (Procured from Thermofisher Scientific) according to the manufacturer protocol. Briefly, 1–10 μM of BsuDisA and MsDisA were incubated with HL‐DNA molecules (50 mM Tris pH 7.5 and 9.4, 2 mM EDTA, 2 mM DTT, and 75 mM NaCl) at 37°C for 10 min. To stabilize the complexes before loading, 0.2% glutaraldehyde (Sigma) was added to the reaction mixture(Torres et al., 2021) and resolved on 7% native polyacrylamide gel electrophoresis (PAGE) for 8 h at 30 mV in 1X Tris‐EDTA buffer. The protein concentration was varied from 0–10 μM.

4.9. CD spectroscopic analysis

The secondary structural elements of MsDisA and other mutant proteins were determined from their far‐UV CD spectra (200–260 nm). The CD spectra of these proteins were recorded at room temperature by a standard procedure using a JASCO J815 spectro‐polarimeter (Bharati et al., 2018). The buffer values were subtracted from the corresponding spectra of protein samples. To determine the substrate‐induced structural alteration of the MsDisA protein, far‐UV CD spectra of the MsDisA + ATP complex were recorded. For MsDisA + ATP, ATP was incubated with MsDisA for 30 min before the experiment.

4.10. Mass‐spectroscopy and LC–MS analysis

A MALDI‐TOF instrument was used to determine the molecular masses of the proteins and nucleotide molecules according to the manufacturer's protocol (BurkerDaltonics, Germany). In‐gel tryptic digestion was performed to confirm the MsDisA and their domain variants by a standard protocol with some modifications (Henzel et al., 2003; Suckau et al., ²⁰⁰³). The HPLC eluted fractions from the MsDisA reaction were analyzed by the LC–MS method (BurkerDaltonics, Germany) (Bharati et al., 2018). MS–MS analysis (negative or positive ion mode) of eluted molecular masses was performed to verify the presence of nucleotides. A standard protocol was carried out for LC–MS and MSMS analysis, and the HPLC gradient composition was used as reported before (Bharati et al., 2012).

4.11. TEM analysis of the proteins

FPLC‐purified MsDisA, MsDisAD84A, MsDisAH118A, and the domain mutant proteins (MsDisA_1–158, MsDisA_1–316, and MsDisA_132–372) were used for TEM grid preparation. The procedure for TEM grid preparation was described earlier in Ohi et al., (2004). Briefly, carbon‐coated copper grids (CF300‐CU, Electron Microscopy Sciences) were glow discharged at 20 mA for 90 s. Wild‐type MsDisA and its variant proteins (0.05 mg/mL) were applied (3.5 μL) to the grid and let it stand for 2 min at room temperature, followed by removal of the excess buffer using filter paper and then stained with 0.5% of uranyl acetate solution, then allowed to air dry. Grids were imaged with a Tecnai‐T12, and Talos 120 electron microscope Thermo Fischer Scientific operated with 120 kV at room temperature. Images were collected at a pixel of 2.54 Å/pixel on the specimen level on a side‐mounted Olympus VELETA (2Kx2K) CCD camera. Protein particles were manually picked and extracted with a box size of 160 Å from the raw micrographs using e2boxer.py. The two‐dimensional reference‐free classification of the extracted particles was performed using e2projectmanager.py (EMAN2.1 software) (Tang et al., 2007).

4.12. Grid preparation and Cryo‐EM data collection

Purified MsDisA protein (50 mM Tris buffer pH 7.9 and 300 mM NaCl) was applied to (3.0 μL of 0.1 mg/mL) glow discharged Quantifoil Cu 2.0/2.0, 300 mesh grids covered with a thin layer of home‐made carbon. Grids were further incubated for 12 s at >95% relative humidity and blotted for 3.5 s. The grids were plunge‐frozen in liquid ethane using ThermoFisher Scientific Vitrobot Mark IV.

The dataset was collected on a Titan Krios G3 transmission electron microscope equipped with a FEG at 300 kV with the automated data collection software EPU (Thermo Fisher Scientific) at the National Cryo‐EM facility, Bangalore. Images of the MsDisA were collected with a Falcon III detector operating in counting mode at a nominal magnification of 75,000X and a calibrated pixel size of 1.07 Å. Table S1 contains the details on defocus range, total electron dose, exposure time, frame number, and data processing parameters.

4.13. Data processing and model building

A total of 1587 cryo‐EM movie frames were motion‐corrected by an algorithm inbuilt in Relion 3.0 (Scheres, 2012). CTF estimation was performed with patch CTF (Zhang, 2016) on the full‐dose weighted motion‐corrected movies. A total of 9,73,755 particles were automatically selected using the cryoSPARC template‐picker (Punjani et al., 2017). The particles were extracted with a box size of 440 pixels. Three rounds of reference‐free 2D classification were carried out with 240 Å particle diameter to remove bad particles. A total of 2,78,281 particles were selected, and ab initio initial model generation into six classes without any symmetry. Of the six ab initio models, 4 classes had the whole protein's density, which were selected for nonuniform refinement. The map was further refined by global CTF refinement. The resolution of the 3D map was estimated at FSC 0.143. The local resolution of the map was estimated with Relion. For the model building of MsdisA, the monomeric model, which AlphaFold predicted, was used as a template (Jumper et al., 2021; Waterhouse et al., ²⁰¹⁸). The protomer of the MsDisA protein was manually fit into the cryo‐EM map using UCSF Chimera and further inspected manually using Coot (Emsley et al., 2010). The model of MsDisA was refined with Phenix using real‐space refinement (Afonine et al., 2018). Figures were made with Chimera (Pettersen et al., 2004) and Pymol (DeLano, ^2002a). The Q scores for the molecule with different maps was calculated with MapQ within chimera (Pintilie et al., 2020).

4.14. Structural modeling, protein sequence alignment, docking and statistical analysis

A structural model of MsDisA protein obtained from cryo‐EM with side chains was used for the GLIDE program's flexible automated molecular docking analysis (Schrödinger Release 2018–3). DisA protein sequence alignment of different organisms was carried out by using ESPript 3.0 (Heger et al., 2014). The protein model and the ligand (ATP/c‐di‐AMP) were first prepared in low‐energy conformation. The ligand was docked in the protein grid with extra precision (XP) docking mode. Results were checked based on ligand‐protein interaction, docking score, and glide energy (kcal/mol). All experimental analyses were performed in three different biological replicates (n = 3).

AUTHOR CONTRIBUTIONS

Sudhanshu Gautam: Conceptualization (equal); data curation (equal); formal analysis (equal); investigation (equal); methodology (equal); software (equal); validation (equal); visualization (equal); writing – original draft (equal); writing – review and editing (equal). Avisek Mahapa: Conceptualization (equal); data curation (equal); investigation (equal); methodology (equal); visualization (equal); writing – original draft (equal); writing – review and editing (equal). Lahari Yeramala: Methodology (equal). Apoorv Gandhi: Investigation (equal). Sushma Krishnan: Formal analysis (equal); writing – review and editing (supporting). Vinothkumar Kutti R.: Data curation (equal); formal analysis (equal); funding acquisition (equal); investigation (equal); methodology (equal); resources (equal); supervision (equal); validation (equal); visualization (equal); writing – review and editing (equal). Dipankar Chatterji: Conceptualization (equal); data curation (equal); formal analysis (equal); funding acquisition (equal); resources (equal); supervision (equal); validation (equal); visualization (equal); writing – original draft (equal); writing – review and editing (equal).

FUNDING INFORMATION

Sudhanshu Gautam acknowledges UGC, Government of India for his fellowship. Avisek Mahapa acknowledges support from the “DBT‐RA Program in Biotechnology & Life Sciences” for fellowship. Dipankar Chatterji, Avisek Mahapa, and Sudhanshu Gautam thank the Indian Institute of Science (IISc) for providing laboratory facility. Dipankar Chatterji acknowledges J C Bose fellowship and Honorary Professorship funding this work. Vinothkumar Kutti R acknowledges SERB, India for the Ramanujan Fellowship (RJN‐094/2017), and the support of the Department of Atomic Energy, Government of India, under Project Identification No. RTI4006. The authors acknowledge the Department of Biotechnology, which support the National CryoEM facility, Bangalore (DBT B‐Life grant DBT/PR12422/MED/31/287/2014).

CONFLICT OF INTEREST STATEMENT

The authors declare no conflicts of interest.

Supporting information

Data S1: Supporting information

Gautam S, Mahapa A, Yeramala L, Gandhi A, Krishnan S, Kutti R. V, et al. Regulatory mechanisms of c‐di‐AMP synthase from Mycobacterium smegmatis revealed by a structure: Function analysis. Protein Science. 2023;32(3):e4568. 10.1002/pro.4568

DATA AVAILABILITY STATEMENT

The data generated and analyzed in this study are included within the manuscript and supplementary data. The cryoEM map and the coordinates have been deposited at EMDB‐33540 and PDB‐7Y0D.