Vibrio cholerae’s ToxRS bile sensing system

  1. Nina Gubensäk  Is a corresponding author
  2. Theo Sagmeister
  3. Christoph Buhlheller
  4. Bruno Di Geronimo
  5. Gabriel E Wagner
  6. Lukas Petrowitsch
  7. Melissa A Gräwert
  8. Markus Rotzinger
  9. Tamara M Ismael Berger
  10. Jan Schäfer
  11. Isabel Usón
  12. Joachim Reidl
  13. Pedro A Sánchez-Murcia
  14. Klaus Zangger
  15. Tea Pavkov-Keller  Is a corresponding author
  1. Institute of Molecular Biosciences, University of Graz, Austria
  2. Laboratory of Computer-Aided Molecular Design, Division of Medicinal Chemistry, Otto-Loewi Research Center, Medical University of Graz, Austria
  3. Institute of Chemistry / Organic and Bioorganic Chemistry, Medical University of Graz, Austria
  4. Diagnostic and Research Institute of Hygiene, Microbiology and Environmental Medicine, Medical University of Graz, Austria
  5. Biological Small Angle Scattering, EMBL Hamburg, Germany
  6. RedShiftBio, United States
  7. Institute of Molecular Biology of Barcelona, Spain
  8. ICREA, Institució Catalana de Recerca i Estudis Avançats, Spain
  9. BioHealth Field of Excellence, University of Graz, Austria
  10. BioTechMed-Graz, Austria

Abstract

The seventh pandemic of the diarrheal cholera disease, which began in 1960, is caused by the Gram-negative bacterium Vibrio cholerae. Its environmental persistence provoking recurring sudden outbreaks is enabled by V. cholerae’s rapid adaption to changing environments involving sensory proteins like ToxR and ToxS. Located at the inner membrane, ToxR and ToxS react to environmental stimuli like bile acid, thereby inducing survival strategies for example bile resistance and virulence regulation. The presented crystal structure of the sensory domains of ToxR and ToxS in combination with multiple bile acid interaction studies, reveals that a bile binding pocket of ToxS is only properly folded upon binding to ToxR. Our data proposes an interdependent functionality between ToxR transcriptional activity and ToxS sensory function. These findings support the previously suggested link between ToxRS and VtrAC-like co-component systems. Besides VtrAC, ToxRS is now the only experimentally determined structure within this recently defined superfamily, further emphasizing its significance. In-depth analysis of the ToxRS complex reveals its remarkable conservation across various Vibrio species, underlining the significance of conserved residues in the ToxS barrel and the more diverse ToxR sensory domain. Unravelling the intricate mechanisms governing ToxRS’s environmental sensing capabilities, provides a promising tool for disruption of this vital interaction, ultimately inhibiting Vibrio’s survival and virulence. Our findings hold far-reaching implications for all Vibrio strains that rely on the ToxRS system as a shared sensory cornerstone for adapting to their surroundings.

Editor's evaluation

This study provides important insights into the structure and mechanism of the sensory protein complex ToxR/S that is associated with the survival and virulence of the cholera pathogen. The structural studies are solid and supported by a series of biophysical experiments revealing a split, periplasmic protein binding interface for bile acid. Results are of interest to protein biochemistry and pharmacology where they may open new routes for the treatment of cholera disease.

https://doi.org/10.7554/eLife.88721.sa0

eLife digest

Cholera is a contagious diarrheal disease that leads to about 20,000 to 140,000 yearly deaths. It is caused by a bacterium called Vibrio cholerae, which can survive in harsh conditions and many environments. It often contaminates water, where it lives in an energy-conserving mode. But when humans consume Vibrio cholerae-contaminated water or food, the bacterium can sense its new environment and switch into a high-energy consuming state, causing fever, diarrhea, and vomiting.

Vibrio cholerae recognizes bile acid in the human stomach, which signals that the bacterium has reached ideal conditions for causing disease. So far, it has been unclear, how exactly the bacterium detects bile acid. Understanding how these bacteria sense bile acid, could help scientists develop new ways to prevent cholera outbreaks or treat infections.

Gubensäk et al. analysed two proteins from the Vibrio cholerae bacterium, called ToxR and ToxS, which are located below the bacteria’s protective membrane. More detailed analyses showed that the two proteins bind together, forming a bile-binding pocket. When correctly assembled, this bile-sensing machine detects bile concentrations in the body, allowing the bacterium to adapt to the local conditions. Using crystal structures, a series of interaction studies, and modeling software, Gubensäk et al. detailed step-by-step how the two proteins sense bile acid and help the bacteria adapt and thrive in the human body.

The results confirm the results of previous studies that implicated ToxR and ToxS in bile sensing and provide new details about the process. Scientists may use this information to develop new ways to interfere with the bacteria’s bile-sensing and gut adaptation processes. They may also use the information to screen for existing drugs that block bile sensing and then test as cholera treatments or prevention strategies in clinical trials. New cholera treatment or prevention approaches that don't rely on antibiotics may help public health officials respond to growing numbers of cholera outbreaks and to prevent the spread of antibiotic-resistant bacteria.

Introduction

The Gram-negative bacterium Vibrio cholerae is the causative agent of the diarrheal cholera disease, which is pandemic since 1960 (Hu et al., 2016). Its dangerousness is highlighted by its sudden outbreaks and environmental persistence causing 21 000–143,000 000 deaths worldwide per year (Ali et al., 2015) including long-lasting environmental and economic damage (Kanungo et al., 2022).

V. cholerae exhibits a life cycle between dormant and virulent state, enabled by its rapid adaption to changing environments (Almagro-Moreno et al., 2015b; Bari et al., 2013). This survival mechanism is maintained via sensory proteins reacting to environmental conditions and substances (DiRita et al., 1991; Hung and Mekalanos, 2005). For entero-pathogens like V. cholerae, bile acid represents one of the major components for virulence activation, pressuring survival strategies (Hung et al., 2006; Hung and Mekalanos, 2005; Li et al., 2016; Midgett et al., 2017).

ToxR is a transmembrane transcription factor (Figure 1—figure supplement 1) involved in the regulation of numerous genes, not only virulence associated, and can function as an activator, co-activator and repressor (Bina et al., 2003; Champion et al., 1997; Lee et al., 2000; Morgan et al., 2011; Skorupski and Taylor, 1997; Wang et al., 2002; Welch and Bartlett, 1998). ToxR periplasmic domain is proposed to act as environmental sensor, being able to bind bile acids (Midgett et al., 2017; Midgett et al., 2020) and consequently activate transcription with its cytoplasmic DNA binding domain (Gubensäk et al., 2021a; Morgan et al., 2011; Morgan et al., 2019; Pfau and Taylor, 1996; Withey and DiRita, 2006), thus inducing a switch of outer membrane proteins from OmpT to OmpU (Simonet et al., 2003; Wibbenmeyer et al., 2002). Since OmpU is more efficient in excluding bile salts due to its negatively charged pore (Duret and Delcour, 2006; Simonet et al., 2003), bile-induced ToxR activation enables V. cholerae survival in the human gut (Wibbenmeyer et al., 2002).

ToxS is built of a periplasmic, a transmembrane and a short cytoplasmic region (Figure 1—figure supplement 1). The periplasmic domain of ToxS (ToxSp) interacts with the periplasmic domain of ToxR (ToxRp) forming a stable heterodimer (ToxRSp; Gubensäk et al., 2021b) thereby protecting ToxR from periplasmic proteolysis and enhancing its activity (Almagro-Moreno et al., 2015a; Almagro-Moreno et al., 2015c; Gubensäk et al., 2021b; Lembke et al., 2018; Pennetzdorfer et al., 2019).

The exact mechanism of ToxR functionality is not clear yet, it was proposed that ToxR binds DNA as a homodimer at the so called ‘tox-boxes’ which represent direct repeat DNA motifs (Crawford et al., 1998; Goss et al., 2013; Krukonis and DiRita, 2003; Ottemann and Mekalanos, 1996; Pfau and Taylor, 1996; Withey and DiRita, 2006). Recently, it was shown that ToxR uses a topological DNA recognition mechanism by recognizing DNA structural elements rather than base sequences (Canals et al., 2023). Also, it is suggested that by multiple binding events of ToxR to promoter regions, membrane attached transcription regulation of ToxR is enabled (Canals et al., 2023). The versatile functionality of ToxR suggests a general role of ToxRS in Vibrio strains, for example the sensing and consequent adaption to changing environmental conditions (Chen et al., 2018; Provenzano et al., 2000).

Bile significantly alters the virulence factor production. Nevertheless, the exact mechanism remains unclear. Studies showed opposite outcomes in regard of up- or downregulation of virulence factors by bile (Bina et al., 2021; Gupta and Chowdhury, 1997; Hung and Mekalanos, 2005; Midgett et al., 2017; Xue et al., 2016; Yang et al., 2013). On the one hand a bile induced reduction of virulence factor production was observed (Gupta and Chowdhury, 1997) via inhibiting ToxT DNA binding ability (Plecha et al., 2015). On the other hand bile seems to enhance TcpP and ToxR activity (Yang et al., 2013) and even induce ToxR ability to activate ctx without ToxT in classical biotype V. cholerae strains (Hung and Mekalanos, 2005). Nevertheless, direct activation of ctx by ToxRS by bile acids is dependent on ctx promotors, differing between strains (Hung and Mekalanos, 2005). Concluding, the effect of bile on V. cholerae virulence seems to be complex and probably dependent on multiple other factors for example calcium concentration and oxygen levels (Hay et al., 2017; Sengupta et al., 2014).

The presented crystal structure of sensory domains of V. cholerae ToxR and ToxS reveals ToxS as a main environmental sensor in V. cholerae. The ToxRS complex exposes a bile binding pocket inside ToxS lipocalin-like barrel that is only properly built via stabilization by newly formed structural elements of ToxR. We performed multiple interaction experiments combined with extensive molecular dynamic MD simulations, to eventually present a bile binding ToxRSp complex, revealing contributions from both proteins to the interaction with bile acid. ToxRSp shows structural and functional similarities with bile sensing complex VtrAC from V. parahaemolyticus (Alnouti, 2009; Li et al., 2016; Tomchick et al., 2016) thus supporting a common superfamily of co-component signal transduction systems whose sensory function is strictly connected to an obligate heterodimer formation (Kinch et al., 2022). AlphaFold-Multimer (Evans et al., 2021; Jumper et al., 2021) structure predictions furthermore reveal a conserved fold of ToxS in different Vibrio species, in contrast to ToxR exhibiting a structural variability among different Vibrio strains.

Results

Periplasmic domains of V. cholerae ToxRS form an obligate dimer

The crystal structure of ToxRp and ToxSp reveals the formation of a heterodimer, with ToxRp contributing secondary structure elements to ToxSp otherwise unstable ß-barrel fold (Figure 1, Figure 2, pdb: 8ALO). The complex forms spontaneously upon addition of both proteins.

Figure 1 with 3 supplements see all
Heterodimer formation of ToxRSp (pdb: 8ALO).

ToxRp ß strand 5 completes main chain hydrogen bond network of ToxSp ß-barrel formation, by interaction with ToxSp ß1. ToxSp ß8 position is stabilized by main chain hydrogen bonds with ToxSp ß1, as well as side chain interactions with ToxRp ß5 and ß4. ToxRp ß5 and α2 are formed upon ToxSp interaction. ToxRSp crystal structure was determined by molecular replacement using an AlphaFold (Evans et al., 2021; Jumper et al., 2021) model as template.

Detailed description of ToxRSp interface.

(A) Overview of ToxRSp interface. The interaction is mainly established via ToxRp ß strands 3, 4 and 5, and ToxSp ß strands 6, 7 and 8. Additionally, an intermolecular salt bridge is formed between ToxRp R281 and ToxSp D142 located at the opening of ToxSp barrel. (B) The intermolecular main chain H-bond pattern of ToxRSp. The interprotein H-bond network is established between ToxRp ß5 and ToxSp ß1 thereby filling the gap between ToxSp ß1 and ß8, which interact only in the C-terminal region of the strands.

ToxRp in the complex has an αß fold which is formed by an α-helix flanked by an anti-parallel hairpin and a three-stranded anti-parallel ß-sheet, followed by a C-terminal helix (Figure 1). Together, the ß-strands form a five-stranded anti-parallel ß-sheet. The C-terminal Cys293ToxRp is involved in a solution-oriented disulphide bond with Cys236ToxRp of the N-terminal helix (Figure 3).

Comparison of ToxRp NMR structure (salmon) to ToxRp bound to ToxSp (green).

Upon binding to ToxSp, ToxRp intrinsically disordered C-terminal region forms ß5 and α2. (A) Topology diagram of ToxRp and ToxRp bound to ToxSp. Newly formed structural elements are highlighted in grey. (B) Structural alignment between ToxRp (PDB 7NN6) and ToxRp in complex with ToxSp (PDB 8ALO). The C-terminal disulphide bond is shown in sticks.

ToxSp forms a lipocalin-like fold consisting of an eight-stranded ß-barrel stabilized by intermolecular H-bond network and flanked by two α-helices located at the openings of the barrel (Figure 2, Supplementary file 2). For a correct barrel formation ToxSp ß1 and ß8 would need to be close enough for building main chain interactions. However, ToxSp ß8 contacts only the first three residues of nine residues long ToxSp ß1 (Figure 2). ToxRp ß5 is positioned into this gap and forms main chain H-bonds with remaining ToxSp ß1 residues. Additionally, close side-chain interactions between ToxRp ß4 and ToxSp ß8 further stabilize ToxSp ß8 positioning (Figure 2). ToxRp ß4 and ß3 are also involved in side-chain interactions with ToxSp ß7 and ß6 completing the strong complex formation. The residue Arg281 located in ToxRp ß5 forms an intermolecular salt bridge with Asp142 of ToxSp α2, which restricts the opening of the barrel (Figure 2).

DSSP secondary structure analysis (Kabsch and Sander, 1983; Touw et al., 2015) indicates a ten-stranded intermolecular ß-sheet consisting of ß1-ß5 of ToxSp and ß1-ß5 of ToxRp. ToxSp five stranded ß-sheet (ß1-ß5) and three-stranded ß-sheet (ß6-ß8), result in a ß-barrel formation, which is stabilized mainly via hydrophobic core interactions and interactions with ToxRp ß5 (Figure 1).

Mutational studies reveal that ToxSp mutant L33S results in increased proteolysis of ToxRp indicating a loss of function of ToxS (Pfau and Taylor, 1998). The crystal structure of ToxRSp shows that L33 is located at ToxSp α1 at the N-terminal membrane-oriented opening of the barrel of ToxSp (Figure 1—figure supplement 2). L33 forms central hydrophobic interactions with apolar residues of ToxSp ß strands stabilizing the core of the barrel (Figure 1—figure supplement 2.). A serine to leucine mutation probably disrupts the hydrophobic interactions and may result in a loss of structure, thus explaining the inability of ToxSp L33S mutant to protect ToxR from proteolysis (Pfau and Taylor, 1998).

There are currently no structures of proteins with significant sequence homology to the ToxRSp complex available in the protein data bank. However, despite the lack of sequence identity the VtrAC complex of V. parahaemolyticus (Tomchick et al., 2016; pdb code: 5KEV, 5KEW) has striking structural and functional similarities with ToxRSp from V. cholerae (Figure 1—figure supplement 3, all-atom RMSD 5.348 Å calculated for 836 atoms; sequence identity: ToxRp-VtrA: 8.1%; ToxSp-VtrC: 11.7%). Similar to ToxRS, the bile sensing functionality of VtrAC induces an essential step for virulence activation upon human gastro-intestinal infection. VtrAC activates the production of main virulence factor VtrB which enables the production of a type III secretion system 2 (T3SS2) for the injection of virulence factors into host cells (Kaval et al., 2023; Zou et al., 2023).

Intrinsically disordered ToxRp C-terminus folds into structural elements essential for the complex formation with ToxSp

Small-angle X-ray scattering (SAXS) experiments indicate a dimer formation of ToxSp in the absence of ToxRp (Figure 6—figure supplement 1, Figure 6—figure supplement 2, Supplementary file 1c) supporting the recently proposed HDOCK ToxS-ToxS dimer (Canals et al., 2023). Nevertheless, aggregation of ToxSp in solution occurs rapidly suggesting an unstable dimer formation (Gubensäk et al., 2021b; Figure 6—figure supplement 3). The instability of ToxSp can be explained by the inability of the first and the last ß strand of the barrel to form strong hydrogen bonding, thus resulting in an unstable hydrophobic core and consequently aggregation. Interaction with ToxRp likely protects otherwise exposed hydrophobic regions of ToxSp (Figure 1, Figure 2).

Comparison of the ToxRSp crystal structure with our previously determined NMR structure of ToxRp (pdb: 7NN6) (Gubensäk et al., 2021b), shows the formation of additional secondary structure elements upon complex formation (Figure 3). Unbound ToxRp has a long intrinsically disordered C-terminus in solution, containing the second cysteine Cys293ToxRp forming a disulphide bond with Cys236 ToxRp located in the middle of helix 1 (Gubensäk et al., 2021b). When bound to ToxSp, the unstructured region of the C-terminus of ToxRp performs a disorder to order transition, by forming two additional structural elements: short α helix 2 and ß strand 5 (Figure 3). The newly formed short α helix 2 contacts the turn between ToxSp ß1 and ß2, as well as residues Leu45ToxSp and Ile46ToxSp located at the C-terminal part of ToxSp ß1 thus increasing the interaction interface and further stabilizing the fold. Although the C-terminal region undergoes extreme structural changes upon complex formation, the orientation of the disulphide bond does not change drastically.

ToxRp ß strand 5 is only formed upon complex formation with ToxSp and forms the essential basis for the interaction by building a stabilizing hydrogen bonding network with ß strands 1 and 8 of ToxSp (Figure 2). Newly formed ToxRp ß5 also forms hydrogen bonds with ToxRp ß4. In unbound ToxRp, polar residues of ß4 are pointing towards solution, whereas hydrophobic parts contribute to the hydrophobic core. Nevertheless, ToxRp ß4 does not reveal significant conformational changes upon ToxRSp complex formation.

ToxSp in complex with ToxRp contains a bile binding pocket

ToxRSp bile interaction was tested using NMR (Becker et al., 2018) (saturation transfer difference STD Figure 6—figure supplement 3, Figure 6—figure supplement 4), chemical shift perturbation CSP (Figure 7, Figure 6—figure supplement 5) and diffusion ordered spectroscopy DOSY (Figure 6B, Figure 6—figure supplement 6), as well microfluidic modulation spectroscopy MMS (Figure 4, Figure 4—figure supplement 1), size exclusion chromatography (Figure 6C) and SAXS (Figure 6A, Figure 6—figure supplement 1, Figure 6—figure supplement 2, Supplementary file 1c). Interaction experiments were performed with bile acid sodium cholate hydrate S-CH. All mentioned experiments confirm an interaction of the ToxRSp complex with bile.

Figure 4 with 1 supplement see all
Analysis of microfluidic modulation spectroscopy MMS experiments.

MMS experiments of ToxRSp with different S-CH concentrations show conformational changes upon binding of bile to ToxRSp. Additional structural rearrangements could be detected upon higher additions of bile. (A) Bar chart of the relative abundance of secondary structural motifs based on Gaussian deconvolution of the corresponding spectra. (B) MMS spectra (baselined, inverted 2nd Derivative) of ToxRSp with and without bile, showing spectral changes upon higher bile additions.

The lipocalin-like fold of ToxSp forms a hydrophobic cavity (Figure 5, Figure 5—figure supplement 1), similar to the bile binding pocket of VtrC in complex with VtrA from V. parahaemolyticus (Tomchick et al., 2016). Using extensive MD simulations enabled a detailed analysis of the protein-ligand interface. Upon binding of bile acid mainly loop regions undergo major conformational changes (Figure 4, Figure 5). Comparison of ToxRSp apo and bile-bound state reveals that ToxSp β1/β2 loop and ToxRp α1/β2 loop experience strongest rearrangements, whereas loop β3/β4ToxSp, α1ToxSp, α2ToxSp and loop β3/β4ToxRp reveal only minor structural changes (Figure 5C). Similarly, MMS spectra show significant spectral changes in disordered, turn and helical regions (Figure 4).

Figure 5 with 1 supplement see all
ToxRSp binding pocket for bile acid.

(A) Superimposition of 100 structures of the ToxRSp complex (dark grey/light grey cartoons) with cholate (sticks, C-atoms in yellow) along one representative classical MD simulation. (B) Detailed view of the binding mode of cholate to ToxRSp. The residues with the larger contribution to the binding of the ligand are highlighted as well as residues 46–49 of ToxSp located on loop β1/β2. (C) Overlay of ToxRSp apo and bile-bound structures. Conformational changes mainly occur in ToxSp loop β1/β2 and ToxRp loop α1/β2. ToxSp helices α1 and α2 and ToxRp loop β3/β4, both close to the ToxRSp interface, experience minor conformational rearrangements upon bile interaction. (D) Hydrophobic properties of the bile binding cavity calculated with the CavMan (v. 0.1, 2019, Innophore GmbH plugin in PyMOL). The entrance of the cavity (blue to green) faces the solvent exhibiting a hydrophilic environment, in contrast to the buried hydrophobic areas (yellow to red). For the analysis of the hydrophobicity of the cavities the program VASCo (Steinkellner et al., 2009) was used; cavities were calculated using a LIGSITE algorithm (Hendlich et al., 1997).

The nature of the interactions of bile acid with the protein complex are mostly hydrophobic. The carboxylic moiety forms a strong interaction with Arg52ToxSp. Additionally, cholate interacts with residues Ser44ToxSp, Ile135ToxSp, Leu138ToxSp, Phe139ToxSp, Val283ToxRp, and Arg281ToxRp (Figure 5). Both proteins contribute to the bile acid interaction, although ToxSp residues are forming a major part of the interface.

ToxRSp has additional surface exposed bile binding patches

ToxRSp interaction experiments propose an increase of molecular weight upon bile addition as shown in Figure 6. SEC-SAXS experiments indicate an increase of molecular weight, radius of gyration and effective maximum particle dimension of ToxRSp upon addition of equimolar amounts of bile acid (Figure 6A, Figure 6—figure supplement 1, Figure 6—figure supplement 2, Supplementary file 1c). Similar outcomes are observed by SEC runs of ToxRSp with and without bile acid using a Superdex200 column resulting in a broadening of ToxRSp-bile acid peak and slightly decreased retention times (Figure 6C, Figure 6—figure supplement 7). NMR DOSY experiments of ToxRSp bound to bile acid clearly show a decrease of the diffusion coefficient, which is linked to an increase of size and weight (Figure 6B, Figure 6—figure supplement 5). Via MMS experiments conformational changes could be detected upon bile acid binding but clearly indicate that no severe structural rearrangements for example aggregation events occur (Figure 4, Figure 4—figure supplement 1).

Figure 6 with 8 supplements see all
ToxRSp bile interaction experiments propose additional bile interaction areas I-IV.

(A) SEC-SAXS experiments with ToxRSp and S-CH reveal an increase of the radius of gyration RG and the effective maximum particle dimension Dmax upon bile addition. (B) Superimposed NMR DOSY spectra of ToxRSp with and without bile acid S-CH. Addition of bile acid causes a decrease of the diffusion coefficient due to an increase of molecular weight. (C) SEC experiments with ToxRSp and S-CH reveal a slight decrease of retention volume upon bile acid presence from 16.19 ml to to 16.08 ml and a broadening of the peak. (D) MD determined surface exposed bile interacting regions of ToxRSp. Besides the bile binding cavity of ToxSp, four regions (marked as I-IV) on the ToxRSp surface could be mapped as additional bile binding areas. Region I and III involves both proteins, whereas region II is located on ToxSp and region IV on ToxRp. The representative structure of the ToxRSp complex (ToxRp: dark grey, ToxSp: light grey) is shown with cholate bound at the cavity (orange) and additional cholate molecules attach to surface exposed regions of ToxRSp.

Due to a loss of signals upon equimolar additions of bile acid caused by the previously mentioned increase of molecular weight, NMR titration experiments were performed using small additions of S-CH (Figure 7). Spectra were recorded with ToxRp and ToxSp both 15N isotopically labelled. An overlay of spectra of only ToxRp or ToxSp 15N labelled enabled the ascription of signals to each of the proteins (Figure 7A). Even small additions of bile (0.1 molar ratio) to ToxRSp results in a significant decrease of signal intensity. ToxSp seems to be mostly affected by the addition of bile acid, causing a disappearance of ToxSp signals indicating direct interaction events (Figure 7B–D). ToxRp signals are also affected by decreased signal intensities but to a lower extent when compared to ToxSp. Upon bile additions higher than a molar ratio of 0.6, mostly signals from ToxRp are visible (Figure 7D). In general, all ToxRSp peaks experience decreased signal intensities caused by fast T2 relaxation.

NMR titration of S-CH to 15 N labelled ToxRSp.

(A) Overlay of 15N-HSQC spectra of 15N-labelled ToxRSp (purple) with 15N-labelled ToxRp (green) and ToxSp (blue) for ascription of peaks to each of the proteins. (B–D) Overlay of 15N-HSQC spectra of 15N-labelled ToxRSp before (purple) and after the addition of increasing ratios of S-CH (orange): 0.3 (B), 0.5 (C), 0.7 (D). Upon bile acid addition signal intensities significantly decrease. Upon higher additions of bile acid mainly ToxRp peaks are visible indicated by green asterisk in (D), whereas most ToxSp signals disappear indicating direct interaction with S-CH. Remaining ToxSp signals are marked with blue asterisk.

Interestingly, MMS experiments show additional structural changes of ToxRSp upon higher bile acid concentrations, with maximum changes achieved at a double molar excess of bile acid suggesting the presence of additional binding sites on ToxRSp (Figure 4, Figure 4—figure supplement 1). Prompted by this evidences, further MD simulations of the complex ToxRSp were performed with one cholate molecule at the binding cavity in the presence of 14 additional cholate molecules randomly positioned in the solvent (Figure 6D). Four solvent exposed regions of the protein, which show a transient binding of cholate molecules could be identified (Figure 6D). Three are located on ToxSp, of which two implicate interactions with ToxRp. One interaction site is exclusively located on ToxRp. These results support the experimental finding of an increase of the molecular weight by binding of more than one cholate molecule.

The active state of ToxR is proposed to be dimeric due to its binding to direct repeat DNA sequences (Canals et al., 2023; Gubensäk et al., 2021a; Krukonis and DiRita, 2003; Withey and DiRita, 2006). Nevertheless, the increase of molecular weight observed in the interaction experiments is significantly lower than 30 kDa (Figure 6A–C), which corresponds to the molecular weight of ToxRSp. Interaction experiments therefore do not implement a bile induced dimerization or oligomerization event. A bile induced formation of trimers involving two ToxRp and one ToxSp or two ToxSp and one ToxRp molecules also seems unlikely. A described trimer formation would involve the disruption of ToxRSp heterodimer complexes, which exhibits a dissociation constant at nanomolar range (Gubensäk et al., 2021b), and includes aggregation of unstable ToxSp. Also, MMS experiments reveal subtle conformational changes of ToxRSp upon bile interaction and do not support extreme structural rearrangements. Instead of major structural changes like dimerization or loss of structure due to aggregation, the experimental results more likely indicate that the presence of bile leads to local conformational changes near the binding cavity of the ToxRSp barrel, similar to the VtrAC complex of V. parahaemolyticus (Li et al., 2016; Tomchick et al., 2016).

Taken together, we propose that ToxRSp bile sensing functions in a complex manner involving a binding site in ToxRSp barrel as well as surface exposed binding regions for bile acid. Bile acid interaction results in concentration dependent conformational changes of ToxRSp which may be related to ToxR transcriptional activity changes (Eichinger et al., 2011; Gubensäk et al., 2021a; Kenney, 2002; Martínez-Hackert and Stock, 1997a; Martínez-Hackert and Stock, 1997b).

ToxRp ß-sheet forms a low-affinity binding region for bile

The MD determined ToxRp binding region to bile (Figure 6D) is also confirmed experimentally via chemical shift mapping (Figure 8, Figure 8—figure supplement 1; Becker et al., 2018; Williamson, 2013). The bile binding area is located at the solution-oriented region of the ß-sheet of ToxRp. The determined dissociation constant of 2.6±1.42 mM (Figure 8, Supplementary file 1d) exhibits a weak binding affinity of ToxRp to bile but physiologically still relevant (Qin and Gronenborn, 2014; Sukenik et al., 2017) since bile acid concentrations in the small intestine vary between 2 and 10 mM (Hamilton et al., 2007). Therefore, we propose that ToxRp interaction with bile acid is only relevant upon high bile concentrations and could be linked to an additional bile sensing mechanism not connected to ToxRSp complex formation.

Figure 8 with 1 supplement see all
Binding studies with ToxRp and bile.

(A) Chemical shift perturbation experiments expose residues mostly affected upon binding of S-CH. (B) Residues which experience a change of their chemical shift upon bile addition are coloured according to a gradient from white to blue. Residues highlighted in dark blue are mostly affected upon bile addition and are therefore most likely located at the interaction area. The calculated dissociation constant of 2.6 mM suggests a low affinity binding of bile to ToxRp. (C) Binding area IV determined by MD simulations (Figure 6) is located at the ß-sheet of ToxRp. Taken together, MD simulation and CSP experiments reveal a bile interacting area located at the ß-sheet of ToxRp proposing a bile sensing function of ToxRp independent on ToxRSp complex formation relevant only at high bile salt concentrations.

Interaction experiments with ToxSp alone and S-CH did not reveal any binding event (Figure 6—figure supplement 4). Due to ToxSp structural instability (Gubensäk et al., 2021b) we propose that without stabilization of ToxRp, the binding pocket is not properly formed and therefore binding to bile cannot occur. Additionally, the presented bile binding complex of ToxRSp shows that residues from ToxRp are also involved in the bile interaction (Figure 5).

Discussion

Bile-induced ToxRS activation enables V. cholerae bile resistance

Sensory proteins represent an indispensable tool for V. cholerae to react to changes in its environment and consequently survive in harsh habitats like the human gut (Almagro-Moreno et al., 2015b). The interaction of inner membrane proteins ToxR and ToxS initiates a sensing function followed by signal transmission thereby causing immediate changes of the expression system of the bacterium (Bina et al., 2003; Childers and Klose, 2007; DiRita et al., 1991). In order to achieve bile resistance, ToxRS bile induced activation leads to a change of outer membrane protein production from OmpT to OmpU (Morgan et al., 2011; Provenzano and Klose, 2000; Simonet et al., 2003).

Our studies reveal a crucial heterodimer formation of the periplasmic sensory domains of V. cholerae thereby shaping a bile binding pocket which is only properly folded upon the complex formation. Interaction experiments show that correct heterodimer assembly of ToxRSp is critical for efficient bile sensing functionality. In vivo these findings suggest that bile-induced virulence regulation and OmpU production occurs only when ToxR and ToxS are both present for forming the obligate heterodimer, thus providing a regulatory restriction in the regulation process by bile. Although a single protein may be more efficient in sensory regulation, a co-component system offers a more strict regulation enabling fine-tuning of transcription levels according to small changes of the pathogen’s environment.

A distinctive feature of members of the ‘ToxR-like’ transcription factor family is the transduction of signals through the membrane without chemical modification, probably via conformational changes (Eichinger et al., 2011; Gubensäk et al., 2021a; Kenney, 2002; Martínez-Hackert and Stock, 1997a; Martínez-Hackert and Stock, 1997b). Thus, we suggest that the observed structural changes of the periplasmic domains of ToxRS upon bile recognition are passed on through the ToxR transmembrane domain to its cytoplasmic effector domain, thereby enhancing ToxR binding to recognition sequences and subsequently induction of transcription (Figure 9).

Model of bile induced activation of V. cholerae ToxRS.

When entering the human gut V. cholerae senses the presence of bile acids by binding of bile to the periplasmic domains of inner membrane proteins ToxR and ToxS. The bile binding pocket is formed by ToxS and stabilized by ToxR. Subsequently, the interaction with bile induces ToxR activation which leads to a change of outer-membrane proteins from OmpT to OmpU. OmpU then provides bile resistance and thus enables the bacterium’s survival in the human body. PDB accession codes: ToxRSp crystal structure: 8ALO, cytoplasmic domain of ToxR: 7NMB. The cytoplasmic domain of ToxR bound to ompU operon is a NMR guided HADDOCK model (Gubensäk et al., 2021a).

Higher additions of bile acid induce additional structural changes of ToxRSp as shown by MMS experiments (Figure 4). A bile acid concentration dependent mechanism could be an efficient tool for V. cholerae for sensing its preferred location of infection in the small intestine (SI). Bile acid concentrations also vary within the SI: postprandial bile acid concentrations are around 10 mM in the initial region of the SI, whereas the distal part contains 2 mM bile acids. (Hamilton et al., 2007). Still, the exact localization of V. cholerae within the SI is not clear (Millet et al., 2014). Previous studies revealed higher numbers of colony forming units (cfu) of the pathogen in the middle and distal regions of the SI (Barbieri et al., 1999). Nevertheless, V. cholerae colonization seems to depend on its motility and host factors e.g. mucins (Millet et al., 2014).

The presented experiments reveal that bile sensing of ToxRS functions according to a complex scheme dependent on the concentration of bile acids. ToxRSp interaction involves a binding site of ToxRSp barrel hosting one bile molecule and multiple surface exposed binding regions offering attachment sites for bile acid molecules. Bile interaction with ToxRSp induces concentration dependent conformational changes of ToxRSp probably influencing ToxR transcriptional activity changes (Eichinger et al., 2011; Gubensäk et al., 2021a; Kenney, 2002; Martínez-Hackert and Stock, 1997a; Martínez-Hackert and Stock, 1997b). The low affinity binding region of ToxR alone could be connected to another bile sensing mechanism, which is relevant at high concentrations of bile acid and independent on ToxRSp complex formation.

The expression system of V. cholerae is complex and sensitively regulated involving numerous factors and depending on environmental conditions and signals. The high-energy consuming virulent state of the pathogen is only switched on when its localization for colonization is reached. Sensing different bile acid concentrations could support the determination of the exact position of the bacterium in the human body and thereby adapting its gene expression according to environmental conditions. Bile resistant associated genes as well as factors essential for colonization are switched on at early stages in order to guarantee the survival of the bacteria, whereas other factors like the cholera toxin are produced later, enabling the spread of the bacteria and causing the main symptoms of the disease (Almagro-Moreno et al., 2015b; Srivastava et al., 1980).

The presented experiments do not indicate a bile-induced dimer- or oligomerization of the ToxRS complex, which is currently proposed as the active conformation of ToxR. Nevertheless, since experiments are performed with periplasmic domains only, a different behaviour of full-length proteins in their natural environment cannot be ruled out. Dimer- and oligomerization events may be dependent on the presence of DNA as suggested recently (Canals et al., 2023), and remain to be elucidated.

The strong interaction of periplasmic domains of ToxR and ToxS indicates that complex formation also occurs spontaneously in-vivo, and independent on the presence of ligands. Presented experiments were performed with already established ToxRSp complexes, to which bile acid was added after complex formation. Nevertheless, we cannot rule out that a folding-on-binding interaction of ToxRS and bile acid happens in-vivo, which remains to be elucidated.

Taken together we propose that in regard to V. cholerae’s virulence expression system ‘ToxR-regulon’ ToxR transcriptional activity is guided by ToxS sensory function. The heterodimer formation is therefore inevitable for the individual functionality of each protein forming a co-dependent system. Understanding the mechanistic details of the ToxRS complex provides a relevant basis for disruption of the crucial interaction interface and consequently inhibition of Vibrio’s adaption to its host and its virulent action.

ToxRS belongs to the superfamily of VtrAC-like co-component signal transduction systems

Vibrio pathogens have a complex regulatory mechanism that allows them to survive and cause disease. ToxRS and VtrAC (Tomchick et al., 2016) are both Vibrio protein complexes fulfilling an essential function since they induce virulence by sensing bile via direct interaction (Bina et al., 2021; Hay et al., 2017; Hung and Mekalanos, 2005; Tomchick et al., 2016). Bile sensing is inevitable for the survival of gastro-entero pathogens (Gunn, 2000; Tomchick et al., 2016). On one hand outer membrane proteins need to be adapted to bile stress in order to guarantee the survival of the bacteria (Sistrunk et al., 2016). On the other hand, the presence of bile signals the environment of the host and therefore the virulence activating state needs to be induced in order to effectively colonize the gut.

Structurally, the protein complexes share the ß-barrel lipocalin-like formation of ToxSp/VtrC completed by a ß strand of ToxRp/VtrA, together forming a bile binding pocket (Figure 1—figure supplement 3). Similar to ToxSp, VtrC is not stable without its interaction partner and tends to aggregate easily (Tomchick et al., 2016). There is no structure of unbound VtrA available yet, so we cannot compare if VtrA also forms new structural elements upon complex formation like ToxRp (pdb code: 7NN6). Another difference between the complexes is that VtrA does not contain cysteine residues like ToxRp. The disulphide linkage represents another option for regulation in V. cholerae. Also, bile interaction occurs in a different manner. Whereas in ToxRSp both proteins contribute to the direct interaction with bile, in VtrAC bile binding strictly involves only VtrC (Li et al., 2016; Tomchick et al., 2016; Zou et al., 2023). If VtrC, in contrast to ToxS, is capable of binding bile acid on its own remains to be elucidated.

Recently, V. cholerae ToxRS was proposed as a distant homolog to VtrAC, forming together a superfamily of co-component signalling systems which are not linked to sequence homology (Kinch et al., 2022). Bacterial two-component systems typically enable signal transduction via chemical modification for example phosphorylation of a response regulator (Mitrophanov and Groisman, 2008). In the case of VtrAC and ToxRS signal transduction is achieved via binding of signalling molecules like bile acid, thereby inducing conformational changes which likely facilitate transcription regulation (Eichinger et al., 2011; Gubensäk et al., 2021a; Kenney, 2002; Li et al., 2016; Martínez-Hackert and Stock, 1997a; Martínez-Hackert and Stock, 1997b; Tomchick et al., 2016).

Although the eight-stranded ß-barrel fold of ToxS and VtrC shows similarities with the calycin superfamily (including fatty acid binding proteins FABP and lipocalins; Supplementary file 1f; Kinch et al., 2022; Tomchick et al., 2016), the formation of an obligate heterodimer to fold the barrel including the binding pocket is unique for ToxRSp and VtrAC (Tomchick et al., 2016).

Despite the obligate heterodimerization and the structural similarities, another feature of this protein family is the arrangement of a two-gene cassette encoding the two proteins (Kinch et al., 2022). Using this information, genetic cluster analysis recently revealed additional members of the VtrAC-like superfamily including BqrP/BqrQ from enteric Burkholderia pseudomallei, as well as GrvA/FidL from pathogenic E. coli O157:H7 Sakai strain (Kinch et al., 2022).

The distinctive structural and functional features of ToxRS and VtrAC support a common superfamily of co-component signalling systems of VtrAC-like protein complexes, whose sensory function is strictly linked to dimerization and ligand binding (Kinch et al., 2022). So far VtrAC and ToxRSp remain the only experimentally solved structures of this relatively new defined superfamily.

ToxRSp obligate dimer formation is conserved among Vibrio strains

The predicted protein structures of ToxRSp proteins from different Vibrio strains (Supplementary file 1e) share the distinctive ToxRSp fold consisting of a ß-barrel ToxSp and an αß folded ToxRp (Figure 10, Fig. Figure 10—figure supplement 1). Although ToxRp structure varies among the strains, the barrel shaped lipocalin-like structure of ToxSp is consistent, clearly visible in the alignment of all structures (Figure 11, Figure 11—figure supplement 1, Figure 11—figure supplement 2).

Figure 10 with 1 supplement see all
Heterodimer structures of periplasmic sensory domains of ToxRS from Vibrio organisms calculated using AlphaFold-Multimer (Evans et al., 2021; Jumper et al., 2021).

ToxRSp dimer formation occurs in different Vibrio strains. The structural alignment of all models reveal that the ToxSp fold is highly similar in whereas ToxRp seems to be more diverse.

Figure 11 with 2 supplements see all
Sequential and structural analysis of sensory domains of ToxRS from different Vibrio strains.

ToxRSp from different Vibrio species exhibit similar heterodimer formations. Especially ToxSp ß-barrel structure, which forms the binding cavity, seems to be conserved. (A) Sequential analysis of conserved residues of ToxRSp. Sequential alignment was done using ToxRS amino acid sequences from Vibrio species (Supplementary file 1e). The sequence identity (%) of ToxR and ToxS proteins from different strains to V. cholerae is shown in the table above the structures. (B) Conserved residues are highlighted on the ToxRSp structure. Conserved residues are coloured in red and shown in sticks, residues which are conserved between groups of similar properties are coloured in gold. Most conserved regions could be found in ToxSp, especially at the openings of the barrel. ToxRp seems to have a higher sequential variability.

Sequence alignments (Figure 11—figure supplement 1, Figure 11—figure supplement 2) reveal numerous conserved residues of ToxSp compared to more diverse ToxRp (Figure 11). Most of the conserved residues of ToxSp are located at the openings of the barrel as well as ß strands 8, 7 and 6 which are involved in ToxRp interaction (Figure 11). Also, residues near the N-terminus, which is probably located near the membrane, seem to be highly conserved in Vibrio species. The aspartate residue which is involved in the intermolecular salt bridge with ToxRp is also one of the conserved residues of ToxSp (Figure 10—figure supplement 1). The high content of conserved residues in the binding pocket of ToxSp proposes that its general function is to bind to small hydrophobic molecules similar to bile acids.

Compared to ToxSp, ToxRp sequence seems to be only conserved between human colonizing strains (Figure 11—figure supplement 2). Most of ToxRp conserved residues are involved in the formation of the hydrophobic core, as well as the ToxSp interface (Figure 11). Additionally, two cysteine residues are conserved, as well as an arginine/lysine residue involved in the formation of the intermolecular salt bridge with the conserved aspartate residue of ToxSp. ToxRp residues involved in the binding to S-CH are also conserved (Val283ToxRp, Arg281ToxRp).

In conclusion, the sequential and structural alignments reveal that ToxSp is conserved among different Vibrio species, whereas ToxRp shows significant sequence similarities only among human colonizing strains. The high number of conserved residues involved in the direct interaction with bile acid furthermore proposes that ToxRSp complex has a common sensory function in Vibrio’s.

Conclusion

Cholera represents a dangerous disease which leads to sudden epidemic outbreaks due to the perseverance of the causative V. cholerae in aqueous environments (Ahmed and Nashwan, 2022; Ali et al., 2015). The bacterium has an immense ability to adapt to challenging conditions due to its sensitive sensory systems. Herein, we present insights into the regulatory bile binding protein complex of V. cholerae involving sensory proteins ToxR and ToxS. Our analysis reveals ToxS as a conserved environmental sensor in Vibrio strains, functionally effective only in complex with transcription factor ToxR. Targeting the disruption of this vital binding mechanism provides a powerful tool for inhibiting Vibrio’s adaption to its host and consequently its virulent action.

Materials and methods

Protein expression and purification

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All constructs listed in table Supplementary file 1a were generated by using standard procedures and verified via automated sequencing. For protein expression E. coli BL21 DE cells were used. Cells were grown under shaking at 180 rpm and induced with 1 mM IPTG when the OD600 reached 0.6–0.8. ToxRp producing cultures were incubated at 37 °C overnight, ToxSp was expressed at 37 °C before induction, after growth at 20 °C overnight. For production of non-isotopically labelled proteins LB media was used. For isotopically labelled proteins cells were grown in M9 minimal media containing 15N-labelled (NH4)2SO4 and 13C labelled glucose as the sole nitrogen and carbon sources, respectively.

After overnight expression, cell cultures were centrifuged, and the pellet dissolved in 20 ml of loading buffer containing either 8 M urea, 300 mM sodium chloride, 10 mM imidazole pH 8 for ToxRp, or 20 mM Tris, 300 mM sodium chloride, 10 mM imidazole pH 8 for ToxSp. Protease inhibitor mix (Serva, Protease Inhibitor Mix HP) was added to the loading buffer. The cells were disrupted via sonication. The lysate was centrifuged, and the supernatant was loaded on a gravity column containing 2 ml of Ni-NTA agarose beads. The column was washed with 15 column volumes (CV) of loading buffer followed by 5 CV of the loading buffers containing 1 M sodium chloride and 5 CV of the loading buffer containing 20 mM imidazole. His-tagged proteins were eluted with 5 CV elution buffer containing 330 mM imidazole. ToxRp constructs were refolded overnight by dialysis at 4 °C in 50 mM Na2HPO4, 300 mM sodium chloride, pH 8. The final purification step includes purification by FPLC using a HiLoad 26/600 Superdex 75 pg column in 50 mM sodium phosphate, 300 mM sodium chloride, pH 8.

Crystallization and data acquisition

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Periplasmic protein domains were expressed separately and added in a 1:2 ratio, with excess of ToxSp. Due to ToxSp instability saturation of ToxRp binding sites is only achieved when ToxSp is added in two molar excess. Unbound ToxSp was eliminated by SEC. The protein complex was dialyzed against crystallization buffer (20 mM Tris, 150 mM NaCl, 0.02% NaN3, pH 8) and concentrated to 30 mg/ml using an Amicon Ultra-15 Centricon (Millipore Merck, Darmstadt, Germany). Crystallization experiments were performed with ORYX8 pipetting robot (Douglas Instruments, Hungerford, UK) and sitting drop vapor-diffusion method using 96-well 3-drop plates (SwissCI AG, Neuheim, Switzerland). Initial crystallizations were setup using commercially available screens: Index (Hampton Research, United States), JCSG +and Morpheus (Molecular Dimensions, United states). Each drop was set up with 0.5 µl protein mix and 0.5 µl of the screen condition. Crystallization plates were incubated at 16 ° C and H11 condition JCSG +containing 0.2 M magnesium chloride hexahydrate, 0.1 M Bis-Tris pH 5.5 and 25 % w/v PEG3350 yielded the best diffracting single crystals after two months. The crystals were stored in liquid nitrogen until screening and data collection at the ID30A-3 beamline at the ESRF (Grenoble, France) using a PILATUS detector. Data were collected at 100° K at 0.96770 Å wavelength with 0.20° oscillation range and a total of 700 images.

Data were processed using XDS (Kabsch, 2010) and AIMLESS (v.0.7.7.) (Evans and Murshudov, 2013). Crystals diffracted up to 3 Å and belong to the space group P65 with cell constants of 72.5 Å, 72.5 Å, 79.4 Å and 90°, 90° and 120°. The structure was solved with molecular replacement using Phaser (2.8.3) (McCoy et al., 2007). A partial ab initio model of the assembled complex of ToxSp and ToxRp predicted by AlphaFold-Multimer (Evans et al., 2021) was used as a template. Employing the complete predicted model of the complex for phasing failed, hence only the larger part of it, ToxSp, was used for initial phasing with Phaser. Furthermore, the predicted model of the ToxSp part was trimmed down at parts below 0.7 predicted IDDT score. The trimmed partial model of ToxSp was suitable to position the model correctly, solve the phase problem and obtain phases that were sufficient enough to place the second molecule, ToxRp, using Phaser. The solved structure was rebuilt and improved in Coot (v.0.9.6.) (Emsley et al., 2010) and with ChimeraX (v.1.3.) (Pettersen et al., 2021) using the ISOLDE plugin (Croll, 2018). Waters were placed manually within Coot. Refinement was performed with REFMAC5 (Murshudov et al., 2011). The final refined model was analysed and validated with PISA (Krissinel and Henrick, 2007), MolProbity (Chen et al., 2010) and PDB. Data collection and refinement statistics (Supplementary file 1b) were generated within Phenix Table1 (Liebschner et al., 2019). The structure was deposited at the PDB with the DOI: https://doi.org/10.2210/pdb8ALO/pdb and the PDB accession code 8ALO.

Ab initio models of ToxRp, ToxSp and dimers with AlphaFold

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Ab initio models of ToxRp and ToxSp (V. cholerae) were calculated using an AlphaFold 2.1 (Jumper et al., 2021) installation with full databases in standard configuration for procaryotes. The search models used for molecular replacement of the heterodimer ToxRSp (V. cholerae) as well as the models of the proposed homodimer ToxSp (V. cholerae) were calculated on an AlphaFold-Multimer 2.1 (Evans et al., 2021) installation with full databases in standard configuration for procaryotes.

For calculation of ToxRSp Heterodimers from various Vibrio Species shown in Supplementary file 1e, an AlphaFold-Multimer 2.2 (31) installation in standard configuration with full databases and v2 model weights was used. For all species, 5 models were generated and ranked by the highest model confidence metric (0.8·ipTM + 0.2·pTM).

NMR experiments

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All NMR spectra were recorded on a Bruker Avance III 700 MHz spectrometer equipped with a cryogenically cooled 5 mm TCI probe using z-axis gradients at 25 ° C. All NMR samples were prepared in 90% H2O/10% D2O. The total sample volume was 600 µL. Spectra were processed with NMRPipe (Delaglio et al., 1995).

Diffusion-ordered NMR spectroscopy DOSY

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For DOSY experiments all samples were dissolved in the exact same buffer (20 mM Tris, 100 mM NaCl pH 6.5). Sodium cholate hydrate S-CH was added to ToxRSp samples from a 20 mM stock dissolved in the same buffer. The concentration of ToxRSp was 360 µM. The DOSY spectra were recorded using the Bruker dstebpgp3s pulse sequence employing a solvent suppression via presaturation and 3 spoil gradients (Johnson, 1999; Morris and Johnson, 1992). Relevant parameters include a gradient duration of 1.4ms, a diffusion time of 70ms and a linear 32-step gradient ramp profile from 1.06 to 51.84 G/cm. Due to phase errors in the first gradient steps, the initial measurement points could not be used, thus the series of points was truncated to afford only the stable signal attenuation. The spectra were processed with Bruker Dynamics Center using standard parameters.

Saturation transfer difference (STD)

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Samples for STD experiments (Mayer and Meyer, 1999) contained an excess of S-CH compared to protein (protein to ligand ratios: ToxRp 1:50, ToxRSp 1:20, ToxSp 1:20). Protein concentrations of samples were: 50 µM ToxRp, 300 µM ToxRSp, 30 µM ToxSp. Three protein selective regions were chosen for saturation: 6000 Hz (amide region), 5000 Hz (amide region) and –5000 Hz (negative control). Experiments with ToxRp and bile were done in two buffers: 20 mM Tris, 100 mM NaCl pH 6.5. Reference spectrum of S-CH was recorded in the exact same buffer at a concentration of 2 mM. Additionally, experiments were repeated in buffer 20 mM Kpi, 200 mM NaCl, pH 8 according to previously published NMR experiments (Midgett et al., 2017).

NMR titration experiments

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Titration experiments with ToxRSp (360 µM) and ToxRp (200 µM) were performed in 50 mM Na2HPO4, 100 mM NaCl at pH 6.5. Additionally, experiments were repeated in buffer 20 mM Kpi, 200 mM NaCl, pH 8 according to previously published NMR experiments (Midgett et al., 2017). S-CH was titrated using a 20 mM stock solution prepared in the exact same buffer. 1D 1H proton spectra as well as 2D 1H-15N-HSQC spectra (Davis et al., 1992) were recorded for each step of the titrations. Due to the disappearance of ToxRSp peaks upon bile addition, bile interaction experiments were performed using low amounts of S-CH. Following protein-to-ligand ratios of S-CH were used for ToxRSp experiments: 0.1, 0.2, 0.3, 0.4, 0.5, 0.7, 0.8, 1, 1.5, 2, 4. For Figure 7 only ratios 0.3, 0.5, and 0.7 are shown. Regarding ToxRp-bile titration following protein-to-ligand ratios of S-CH were used: 1, 3, 13, 50, 100, 150, 200, shown in Figure 8—figure supplement 1.

CSP analysis

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Spectra were processed with NMRPipe (NMRDraw v5.6 Rev) (Delaglio et al., 1995) and analyzed with CcpNmr Analysis 2.4.2. (Skinner et al., 2016). Molecular images were created with PyMOL (v2.0 Schrödinger, LLC). For ToxRp NMR experiments published NMR assignments were used (Gubensäk et al., 2021b).

Euclidean distances, also called d-values, were calculated as described by Williamson, 2013, (Becker et al., 2018):

(1) d=12[δH2+(αδN)2]

d…Euclidean distance, δN15N chemical shift changes, δH1H chemical shift changes,

α…scaling factor (glycines α= 0.2, all other amino acids α= 0.14)

A threshold value was determined according to the procedure described by Schumann et al., 2007 to exclude residues with non significant chemical shift changes.

The dissociation constant (Kd) was calculated for each amino acid individually using CcpNmr Analysis 2.4.2. (Skinner et al., 2016) and the following equation:

(2) Δδobs=Δδmax{([P]t+[L]t+Kd)([P]t+[L]t+Kd)24[P]tLt/2[P]t

Δδobs…change in observed shift, Δδmax…maximum shift change on saturation,

[P]t…total protein concentration, [L]t…total ligand concentration, Kd…dissociation constant

Size exclusion chromatography (SEC)

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SEC experiments were carried out on an ÄKTA pure 25 (Cytavi, Marlborough, USA) using a Superdex 200 Increase 10/300 column (Cytiva) with a flowrate of 0.4 ml/min. As sample and running buffer a phosphate buffer (50 mM Na2HPO4, 300 mM NaCl, pH 8, supplemented with 0.02% NaN3) was used. A total of 100 µl of ToxRS without and with bile acid S-CH were measured. Sample 1 was 1 mM of ToxRSp complex, sample 2 was 300 µM of ToxRSp with equimolar amount (1:1) of S-CH.

Microfluidic modulation spectroscopy (MMS)

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For initial MMS measurements three 1 ml ToxRSp samples of 5 mg/ml were prepared in 50 mM Na2HPO4, 300 mM NaCl pH 8.0. A 20 mM stock of S-CH was made in the exact same buffer and added to the protein complex for final protein to ligand ratios of 1:1 and 1:2. Also, a ToxRSp sample without ligand was measured. Results are shown in Figure 4. For each sample a reference buffer, containing the same amount of S-CH was used.

Additionally, measurements with increasing protein-to-ligand ratios were performed with 4.5 mg/ml ToxRSp complex in 50 mM Na2HPO4, 300 mM NaCl pH 8. A 20 mM stock of S-CH was made in the exact same buffer and added to the protein complex for final protein to ligand ratios of: 1:0, 1:1, 1:2, 1:3, 1:5, 1:20. For each sample, a reference buffer, containing the same amount of S-CH was used. Results are shown in Figure 4—figure supplement 1.

Samples were measured and analysed with a RedShiftBio AQS3pro MMS production system equipped with sweep scan capability (RedShiftBio, Boxborough, MA, USA). For background subtraction, chemically identical buffer and buffer-bile mixtures were loaded pairwise with the corresponding sample onto a 96-well plate. The instrument was run at a modulation frequency of 1 Hz and with a microfluidic transmission cell of 23.5 µm optical pathlength. The differential absorbance spectra of the sample against its buffer reference were measured across the amide I band (1714–1590 cm-1). For each spectrum, triplicate measurements were collected and averaged. The data were analysed on the RedShiftBio delta analysis software.

SEC-SAXS

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SEC-SAXS was performed at the EMBL P12 bioSAXS beamline (PETRA III, Hamburg, Germany). Samples were dissolved in 50 mM Na2HPO4, 300 mM NaCl pH 8.0 and 3% glycerol to prevent radiation damage. For the single proteins, ToxSp (10 mg/ml), ToxRp (10 mg/ml) and ToxRSp (14 mg/ml), a Superdex 75 Increase 10/300 column was used. For the ToxRS complex in the presence of bile acid S-CH (10 mg/ml), a Superdex 75 Increase 5/150 column was used. The column outlet was connected to the flow-through SAXS cell.

Throughout the elution, frames (I(q) vs q, where q=4πsinθ/λ, and 2θ is the scattering angle) were collected successively. The data were normalized to the intensity of the transmitted beam and radially averaged; the scattering of the solvent was subtracted from the sample frames using frames corresponding to the scattering of the solvent. For optimized selection of buffer and sample frames the program CHROMIXS was employed. Data were further analysed and prepared via PRIMUS from the ATSAS software (Manalastas-Cantos et al., 2021), model preparation was done using DAMMIF at the online ATSAS server provided by EMBL Hamburg (https://www.embl-hamburg.de/biosaxs/atsas-online/). Comparison to theoretical scattering curves was performed with CRYSOL. Figures were done using PyMOL (Schrodinger, 2010).

The data have been deposited in the SASBDB databank. Accession codes: SASDR25, SASDR35, SASDR45, SASDR55.

MD simulations

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The Cartesian coordinates of the complex between ToxRp and ToxSp were used as initial structure for our MD simulations. The minimum energy structure of the ligand cholate was first of all minimized with the GFN2-xTB (Bannwarth et al., 2019) using CREST, (Grimme, 2019), minimized with r2SCAN (Furness et al., 2020) and the RESP point charges for all atoms of cholate were derived from the electrostatic potential computed at the HF-6–31 G* level of theory with Gaussian16 v. C. (Gaussian 16 and Revision B.01, 2016). The protonation states of the titritable residues on the protein complex were calculated via the H++ web server (Anandakrishnan et al., 2012; Gordon et al., 2005; Myers et al., 2006) assuming a pH value of 8.0. Cholate was manually docked at the hydrophobic inner chamber of the ToxRSp complex guided by the superimposition of the crystallographic structure of Vibrio parahaemolyticus VtrA/VtrC complex in the presence of taurodeoxycholate (PDB id. 5KEW) (Alnouti, 2009; Li et al., 2016).

MD simulations were carried out using the suite of programs AMBER20 (Amber20, 2022) Protein residues and solvated ions were treated with the AMBER ff19SB force-field (Ponder and Case, 2003) and cholate was described as AMBER atoms types following the standard procedure in antechamber. We simulated two systems: the apo complex (ToxRSp) and the cholate-bound complex (ToxRSp +cholate). The two systems were then energetically minimized to avoid close contacts, and then placed in the center of a cubic box filled with OPC water molecules (Izadi et al., 2014). Then, the solvated systems were minimized in three consecutive steps (all protons, solvent and all system) and heated up in 50 ps to from 100 k to 300 K in a NVT ensemble using the Langevin thermostat (gamma friction coefficient of 1.0). Care was taken to constraint the solute during the heating step by imposition of a harmonic force on each atom of the solute of 40 kcal mol–1 Å–2. Afterwards, these harmonic constraints were gradually reduced up to a value of 10 kcal mol–1 Å–2 in 4 simulation stages (NVT, 300 K). Then, the systems were switched to constant pressure (NPT scheme, 300 K) and the imposed constraints during the heating step were totally removed. Finally, each of the systems was submitted to three independent MD simulations of 0.8 μs (total simulation time per system: 2.4 μs). Atom-pair distance cut-offs were applied at 10.0 Å to compute the van der Waals interactions and long-range electrostatics by means of Particle-Mesh Ewald (PME) method. SHAKE algorithm was applied to restrain the hydrogen atoms on water molecules. MD trajectory analysis was carried out using the CPPTRAJ (Roe and Cheatham, 2013) module from AMBER20 for monitoring the root-mean-square distance (RMSD) and root-mean-square fluctuation (RMSF), amongst other parameters.

Data availability

Diffraction data have been deposited in PDB under the accession code 8ALO. SAXS data have been deposited: ToxR - SASDR25 ToxS - SASDR35, ToxR:ToxS - SASDR45, ToxR:ToxS:bile - SASDR55. All data generated or analysed during this study are included in the manuscript and supporting files.

The following data sets were generated
    1. Gubensaek N
    2. Sagmeister T
    3. Pavkov-Keller T
    4. Zangger K
    5. Bulheller C
    6. Wagner GE
    (2023) RCSB Protein Data Bank
    ID 8ALO. Heterodimer formation of sensory domains of Vibrio cholerae regulators ToxR and ToxS.
    1. Graewert M
    (2023) SASBDB
    ID SASDR25. Periplasmic domain of cholera toxin transcriptional activator ToxR.
    1. Graewert M
    (2023) SASBDB
    ID SASDR35. Periplasmic domain of cholera transmembrane regulatory protein ToxS.
    1. Graewert M
    (2023) SASBDB
    ID SASDR45. A complex between the periplasmic domains of cholera toxin transcriptional activator ToxR and transmembrane regulatory protein ToxS.
    1. Graewert M
    (2023) SASBDB
    ID SASDR55. A complex between the periplasmic domains of cholera toxin transcriptional activator ToxR and transmembrane regulatory protein ToxS bound to bile salt.

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    3. Rivera-Cancel G
    (2016)
    Vibrio Parahaemolyticus VtrA/VtrC Complex
    PDB.

Decision letter

  1. Hannes Neuweiler
    Reviewing Editor; University of Würzburg, Germany
  2. Amy H Andreotti
    Senior Editor; Iowa State University, United States
  3. Hannes Neuweiler
    Reviewer; University of Würzburg, Germany

Our editorial process produces two outputs: (i) public reviews designed to be posted alongside the preprint for the benefit of readers; (ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Vibrio cholerae's ToxRS Bile Sensing System" for consideration by eLife. Your article has been reviewed by three peer reviewers, including Hannes Neuweiler as Reviewing Editor and Reviewer #1 and the evaluation has been overseen by Amy Andreotti as the Senior Editor.

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

The reviewers support the publication of your work as an article in eLife after their concerns have been addressed. Please address all concerns in a point-by-point response letter and a revised manuscript. The reviews and the concerns (detailed in the sections "recommendations for the authors") are below.

To improve the quality of your manuscript further, the reviewers suggest that you use the space provided by eLife (no limits on the number of display items) and move Figures that show important results from biophysical experiments (i.e., SEC-MALS, NMR, and ITC) from Supporting Information into the main article, together with improved Figure legends.

Reviewer #1 (Recommendations for the authors):

1) SEC-MALS experiments show an unreasonable increase of molecular weight of the ToxRSp complex by ~10 kDa upon binding of bile acid in 1:1 stoichiometry (Figure S11). On a side note: there is a mistake in the labelling of panels in Figure S11. Panel A is labelled ToxRSp but MALS data show a Mw of ~40 kDa. Panel B is labelled ToxRSp:bile acid and MALS data show a Mw of ~30 kDa, which is not reasonable. On page 6 the authors argue for additional binding patches that would explain the 10-kDa increase of Mw. But this is unreasonable from a 1:1 stoichiometry in the SEC-MALS experiment. Bile acid has a Mw of ~0.5 kDa. The interactions seen in MD simulations likely indicate transient, rather non-specific interactions and not specific binding sites. The 10-kDa increase of Mw in SEC-MALS experiments remains ambiguous. The shape of the peak in the chromatogram in panel B is not symmetric. A possible explanation could be sample heterogeneity induced by formation of the ternary complex, which would require further investigations and explanations.

2) With regard to point 1): Can the authors comment on the concentration of bile acid in the stomach, which has implications on their findings? The concentration is of relevance to the discussion regarding weak bile-ToxRSp interactions showing high mM Kd values.

3) The readability of the manuscript would benefit from putting some of the supplementary figures into the main manuscript. E.g., it would be helpful to show Figures S2, S3 and S5 next to main Figure 1. They address the interesting disorder-order transition and details on molecular interactions within the interface, which are described to on page 3-4.

4) Can the authors infer the order of events in the assembly of the ToxRSp-bile complex from their data? From the structure it looks like that the folding-on-binding interaction of ToxRp cages the ligand, which would indicate that ToxRp binds after or in cooperation with formation of a ToxSp-bile complex. The structure seems to show that the flexible b1/b2 loop folds over the binding pocket. Could this explain the observed 14% increase of disorder upon bile binding in MMS experiments (Figure 3)?

5) Can the authors expand their discussion on why a split binding interface for bile acid evolved? What is the benefit or the rational? Couldn't a conventional single-domain periplasmic binding interface do the same job? The recruitment of ToxS to ToxR seems to be a requirement for successful signal transduction.

Reviewer #3 (Recommendations for the authors):

Although the structural data are very interesting, some phenotypic assays with protein mutants will significantly enhance the significance of the study.

The provided biophysical experiments to explain the binding mechanism, raise more questions than answers. The data presented in the Supplementary Information section, are not convincingly supported. Below are some points regarding the interaction study.

– SEC-MALS: The difference of elution volume between the bound- and free ToxRSp (16.4 vs 16.3 ml) is too small to be significant. The elution peak in the presence of bile acid is not symmetrical, suggesting a mixture between a bound and a free-state. Did the author try to add more bile acid in order to saturate the complex and have a more significant shift of the elution volume?

– Figure S10: The ITC data appear to be inconsistent. How were the data fitted? The heat of interaction shows a negative value, whereas the fitted enthalpy is positive. The stoichiometry of 0.5 should be explained. Additionally, could the authors show the raw ITC data?

– Figure S14: The amount of added sodium cholate seems to be very high. How can we be certain that the spectral modifications are not caused by changes in viscosity? Have the authors attempted to add another hydrophobic compound within a similar concentration range?

How are the chemical shift perturbations (CSP) considered? Typically, CSP values greater than 1 or 2 times the standard deviation of the shift of all residues are significant.

– It would be helpful to include the 1D NMR spectra of Sodium cholate. This would help in understanding the STD experiments and Figure S8A. In Figure S8A, the shift observed in the 1D experiment may be due to a variation in pH. If the protein signals disappear in the 2D HSQC due to the very large molecular weight, as explained by the authors, why is the amide region still well-visible in the 1D experiment?

Figure 1: Regarding the sentence, "ToxSp ß8 position is stabilized by minor main chain hydrogen bonds," what is meant by "minor HBonds"?

Figure 2: Based on the figure, the conformational changes in the loops do not appear to be significant, as mentioned in the text (just above, line 16). Could you please highlight the conformational changes more clearly or provide a zoomed-in view?

Page 5, line 3: The sentence, "The conformation of ß4 shows only slight changes when interacting with ß5 or its aqueous environment," is not clear. Could you please specify what the slight conformational changes are?

Figure S6A and other figures, the legend needs to be completed, please explain different panels.

Figure 4: Panel C is not clear.

Overall, the figure legends and quality of the figures are currently not suitable for publication.

References: The references for Gaussian and Amber are not well formatted.

https://doi.org/10.7554/eLife.88721.sa1

Author response

Reviewer #1 (Recommendations for the authors):

1) SEC-MALS experiments show an unreasonable increase of molecular weight of the ToxRSp complex by ~10 kDa upon binding of bile acid in 1:1 stoichiometry (Figure S11). On a side note: there is a mistake in the labelling of panels in Figure S11. Panel A is labelled ToxRSp but MALS data show a Mw of ~40 kDa. Panel B is labelled ToxRSp:bile acid and MALS data show a Mw of ~30 kDa, which is not reasonable. On page 6 the authors argue for additional binding patches that would explain the 10-kDa increase of Mw. But this is unreasonable from a 1:1 stoichiometry in the SEC-MALS experiment. Bile acid has a Mw of ~0.5 kDa. The interactions seen in MD simulations likely indicate transient, rather non-specific interactions and not specific binding sites. The 10-kDa increase of Mw in SEC-MALS experiments remains ambiguous. The shape of the peak in the chromatogram in panel B is not symmetric. A possible explanation could be sample heterogeneity induced by formation of the ternary complex, which would require further investigations and explanations.

We appreciate the remark about the switched molecular weight assignment and agree with reviewer 1 that the molecular weight increase (10 kDa) determined by SEC-MALS experiments of ToxRSp with bile acid remains ambiguous. Our understanding is that bile acid may interfere with the absorption properties of the complex. Also, the interaction of bile acid with aromatic residues in the binding pocket of ToxRSp could induce a quenching effect resulting altogether in an incorrect normalization of the light scattering signal. Therefore, we decided to replace the graph and show the UV traces instead of the light scattering, which reveal clear differences between the samples with and without bile acid indicating an interaction between ToxRSp and bile acid. We agree with reviewer 1 about the non symmetrical shape of the ToxRSp-bile peak and agree about the possibility of a transient ternary complex formation. To gain more clarity, we performed additional experiments (NMR and MMS titrations) further supporting the entangled dynamic mechanism of ToxRSp bile sensing but excluding the hypothesis of the formation of (stable) bile induced ToxRSp multimers.

Although we performed multiple interaction experiments using several biophysical methods we cannot provide a clear explanation for the increase of size of ToxRSp upon bile addition. Instead we could show that ToxRSp and bile acid are interacting according to a complex scheme involving a binding cavity and surface attachment of bile acid molecules. The exact impact of bile acid on ToxRSp ternary structure remains to be elucidated.

Still, the experiments reveal an entangled concentration dependent bile sensing mechanism of ToxRSp involving subtle changes of ToxRSp secondary structure which may be relevant for transcriptional activity.

2) With regard to point 1): Can the authors comment on the concentration of bile acid in the stomach, which has implications on their findings? The concentration is of relevance to the discussion regarding weak bile-ToxRSp interactions showing high mM Kd values.

We appreciate the remark about the bile concentrations in the human gut and added a section on this topic to the results and discussion chapters.

3) The readability of the manuscript would benefit from putting some of the supplementary figures into the main manuscript. E.g., it would be helpful to show Figures S2, S3 and S5 next to main Figure 1. They address the interesting disorder-order transition and details on molecular interactions within the interface, which are described to on page 3-4.

We agree with reviewer 1 and added new figures (now: Figure 2 and Figure 3) to the manuscript.

4) Can the authors infer the order of events in the assembly of the ToxRSp-bile complex from their data? From the structure it looks like that the folding-on-binding interaction of ToxRp cages the ligand, which would indicate that ToxRp binds after or in cooperation with formation of a ToxSp-bile complex. The structure seems to show that the flexible b1/b2 loop folds over the binding pocket. Could this explain the observed 14% increase of disorder upon bile binding in MMS experiments (Figure 3)?

Experiments were performed with already established ToxRSp complexes. Bile acid was added after the ToxRSp complex formation. Nevertheless, we cannot rule out that the mentioned folding-on-binding interaction happens in-vivo. We appreciate the notion of reviewer 1 and included this topic in the Discussion section.

We agree with reviewer 1 about the importance of conformational changes upon bile interaction and added a new figure (Figure 5C) with a more detailed discussion regarding this topic. The conformational changes of ß1/ß2 may be an explanation for the increase of disorder, but since also other regions seem to be affected e.g. helical regions, bile acid binding could also influence the stability of helices. This remains to be elucidated.

5) Can the authors expand their discussion on why a split binding interface for bile acid evolved? What is the benefit or the rational? Couldn't a conventional single-domain periplasmic binding interface do the same job? The recruitment of ToxS to ToxR seems to be a requirement for successful signal transduction.

We agree with reviewer 1 that the formation of the complex is necessary for bile sensing and a single protein would be more efficient. We expanded the discussion accordingly.

Reviewer #3 (Recommendations for the authors):Although the structural data are very interesting, some phenotypic assays with protein mutants will significantly enhance the significance of the study.

We agree with reviewer 3 that phenotypic assays with ToxRSp mutants would be an interesting experiment, which we are considering for future studies. Nevertheless, mentioned experiments are time consuming and deliver considerable amount of data which would implicate another separate publication. We believe that the experiments shown in this manuscript already provide crucial and detailed information about ToxRS which we would like to make accessible for the scientific community.

The provided biophysical experiments to explain the binding mechanism, raise more questions than answers. The data presented in the Supplementary Information section, are not convincingly supported. Below are some points regarding the interaction study.

– SEC-MALS: The difference of elution volume between the bound- and free ToxRSp (16.4 vs 16.3 ml) is too small to be significant. The elution peak in the presence of bile acid is not symmetrical, suggesting a mixture between a bound and a free-state. Did the author try to add more bile acid in order to saturate the complex and have a more significant shift of the elution volume?

We agree with reviewer 3 that the difference in the elution volume between ToxRSp with and without bile acid is rather small, but in regard to the molecular weight increase observed by other methods (NMR, SAXS) we believe it is still relevant to mention. Binding of bile acid to ToxRSp induces a broadening of the ToxRSp SEC peak and a change of its shape indicating a binding event.

We agree with reviewer 3 about the non symmetrical shape of the ToxRSp-bile peak and agree with the possibility of a mixture of bile-induced states of ToxRSp. But since higher bile to protein ratios did not influence the values significantly, we assume that peak broadening occurs due to the complex dynamic mechanism of ToxRSp bile sensing involving a binding cavity and surface attachment of bile acid molecules. Although we performed multiple interaction experiments using several biophysical methods we cannot offer a clear explanation about the impact of bile acid on the ternary structure of ToxRSp. Instead we could show that ToxRSp bile sensing involves concentration dependent distinctive conformational changes which could be crucial for ToxR transcriptional activity.

– Figure S10: The ITC data appear to be inconsistent. How were the data fitted? The heat of interaction shows a negative value, whereas the fitted enthalpy is positive. The stoichiometry of 0.5 should be explained. Additionally, could the authors show the raw ITC data?

We agree that ITC data are inconsistent therefore we performed additional experiments which we present in the revised manuscript. ITC was the first experiment we performed for investigating ToxRSp bile sensing. Nevertheless, it turned out bile binding to ToxRSp is more complex than expected. To obtain preliminary information from the ITC we performed analysis with the single site independent model. However, since ITC data seems to raise more questions than it answers, we decided to remove the data from the manuscript and concentrate on the results obtained by other more detailed biophysical characterizations. Additionally, we added further experiments (NMR, MMS) supporting our theory of an entangled bile sensing mechanism of ToxRSp.

– Figure S14: The amount of added sodium cholate seems to be very high. How can we be certain that the spectral modifications are not caused by changes in viscosity? Have the authors attempted to add another hydrophobic compound within a similar concentration range?

The concentrations of ToxRSp are relatively high (~350µM) due to the high molecular weight of the complex (30 kDa). Lower concentrations of ToxRSp resulted in a significant decrease of spectra quality. Nevertheless, via NMR experiments (e.g. STD, 1D, CSP, DOSY) in combination with other biophysical methods like SAXS and MMS we could confirm an interaction of ToxRSp with bile acid. Also, we performed additional NMR experiments starting with 0.1 molar excess of bile acid over ToxRSp (corresponding to a final concentration of 35 µM bile acid) resulting in similar outcomes: line broadening and peak shifting. Furthermore, we tested the interaction of ToxRSp with the hydrophobic compound cFP which did not result in line broadening or peak shifting indicating no interaction between ToxRSp and cFP. We included these measurements in the supplementary material (Figure 6 —figure supplement 8).

How are the chemical shift perturbations (CSP) considered? Typically, CSP values greater than 1 or 2 times the standard deviation of the shift of all residues are significant.

The spectrum quality of ToxRSp decreases significantly with increasing amounts of bile acid and therefore hinder a reliable analysis of the chemical shift changes. Also, the high molecular weight of ToxRSp of 30kDa is problematic for NMR experiments, but could be improved by using methods like deuteration and TROSY experiments etc. Nevertheless, spectrum quality still remains a restriction in the analysis of peak shifting. To still gain information about the bile interaction we provide new experiments by which we are able to distinguish between ToxRp and ToxSp signals and could show that ToxSp signals are mainly influenced upon bile interaction indicating a direct interaction.

– It would be helpful to include the 1D NMR spectra of Sodium cholate. This would help in understanding the STD experiments and Figure S8A. In Figure S8A, the shift observed in the 1D experiment may be due to a variation in pH. If the protein signals disappear in the 2D HSQC due to the very large molecular weight, as explained by the authors, why is the amide region still well-visible in the 1D experiment?

We included a spectrum of sodium cholate hydrate in the supplementary information. In case of a change of the pH shift we would expect all signals to be affected. We also made sure to use exactly the same buffer during the NMR titration. The protein was dialyzed several days with multiple changes of the buffer to prevent any pH changes during titration. As shown in the experiments, only some signals experience changes of the chemical shift and the decrease of intensity is also differing between signals as for ToxSp signals the decrease is larger than for ToxRp signals. The line broadening due to increased molecular weight is also visible in 1D experiments. Signal loss due to fast T2 relaxation in high molecular weight complexes is much more severe during the much longer pulse-sequences of multi-dimensional experiments, rather than 1D 1H spectra.

Figure 1: Regarding the sentence, "ToxSp ß8 position is stabilized by minor main chain hydrogen bonds," what is meant by "minor HBonds"?

We appreciate the notification and rewrote the sentence accordingly.

Figure 2: Based on the figure, the conformational changes in the loops do not appear to be significant, as mentioned in the text (just above, line 16). Could you please highlight the conformational changes more clearly or provide a zoomed-in view?

We provided an additional figure (Figure 5C) comparing the apo and bile-bound state of ToxRSp and discussed it accordingly in the Results section.

Page 5, line 3: The sentence, "The conformation of ß4 shows only slight changes when interacting with ß5 or its aqueous environment," is not clear. Could you please specify what the slight conformational changes are?

We thank reviewer 3 for pointing out this issue. The sentence was rewritten accordingly. There are no significant conformational changes of ToxRp ß4 when interacting with newly formed ß5 (when bound to ToxSp) compared to its conformation when it is exposed to the solution in unbound ToxRp.

Figure S6A and other figures, the legend needs to be completed, please explain different panels.

Figure 4: Panel C is not clear.

Overall, the figure legends and quality of the figures are currently not suitable for publication.

We corrected the figure legends and generated new figures with higher quality.

References: The references for Gaussian and Amber are not well formatted.

We thank reviewer 3 for the notification and formatted the reference accordingly.

https://doi.org/10.7554/eLife.88721.sa2

Article and author information

Author details

  1. Nina Gubensäk

    Institute of Molecular Biosciences, University of Graz, Graz, Austria
    Contribution
    Conceptualization, Data curation, Supervision, Funding acquisition, Investigation, Writing - original draft, Project administration, Writing – review and editing
    For correspondence
    nina.gubensaek@uni-graz.at
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0415-4299
  2. Theo Sagmeister

    Institute of Molecular Biosciences, University of Graz, Graz, Austria
    Contribution
    Data curation, Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
  3. Christoph Buhlheller

    Institute of Molecular Biosciences, University of Graz, Graz, Austria
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  4. Bruno Di Geronimo

    Laboratory of Computer-Aided Molecular Design, Division of Medicinal Chemistry, Otto-Loewi Research Center, Medical University of Graz, Graz, Austria
    Contribution
    Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
  5. Gabriel E Wagner

    1. Institute of Chemistry / Organic and Bioorganic Chemistry, Medical University of Graz, Graz, Austria
    2. Diagnostic and Research Institute of Hygiene, Microbiology and Environmental Medicine, Medical University of Graz, Graz, Austria
    Contribution
    Conceptualization, Data curation, Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5704-3955
  6. Lukas Petrowitsch

    Institute of Molecular Biosciences, University of Graz, Graz, Austria
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  7. Melissa A Gräwert

    Biological Small Angle Scattering, EMBL Hamburg, Hamburg, Germany
    Contribution
    Data curation, Investigation, Visualization, Writing – review and editing
    Competing interests
    No competing interests declared
  8. Markus Rotzinger

    Institute of Chemistry / Organic and Bioorganic Chemistry, Medical University of Graz, Graz, Austria
    Contribution
    Investigation, Visualization
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0411-3403
  9. Tamara M Ismael Berger

    Institute of Molecular Biosciences, University of Graz, Graz, Austria
    Contribution
    Investigation, Visualization
    Competing interests
    No competing interests declared
  10. Jan Schäfer

    RedShiftBio, Boxborough, United States
    Contribution
    Data curation, Investigation, Visualization, Writing – review and editing
    Competing interests
    is affiliated with Redshift BioAnalytics, Inc which distributes the AQS3pro. Access to the AQS3pro instrument was provided to Nina Gubensäk as part of the RedShiftBio demo lab
  11. Isabel Usón

    1. Institute of Molecular Biology of Barcelona, Barcelona, Spain
    2. ICREA, Institució Catalana de Recerca i Estudis Avançats, Barcelona, Spain
    Contribution
    Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
  12. Joachim Reidl

    1. Institute of Molecular Biosciences, University of Graz, Graz, Austria
    2. BioHealth Field of Excellence, University of Graz, Graz, Austria
    3. BioTechMed-Graz, Graz, Austria
    Contribution
    Conceptualization, Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
  13. Pedro A Sánchez-Murcia

    Laboratory of Computer-Aided Molecular Design, Division of Medicinal Chemistry, Otto-Loewi Research Center, Medical University of Graz, Graz, Austria
    Contribution
    Data curation, Investigation, Visualization, Writing – review and editing
    Competing interests
    No competing interests declared
  14. Klaus Zangger

    1. Institute of Chemistry / Organic and Bioorganic Chemistry, Medical University of Graz, Graz, Austria
    2. BioHealth Field of Excellence, University of Graz, Graz, Austria
    3. BioTechMed-Graz, Graz, Austria
    Contribution
    Conceptualization, Data curation, Supervision, Funding acquisition, Validation, Investigation, Project administration, Writing – review and editing
    Competing interests
    No competing interests declared
  15. Tea Pavkov-Keller

    1. Institute of Molecular Biosciences, University of Graz, Graz, Austria
    2. BioHealth Field of Excellence, University of Graz, Graz, Austria
    3. BioTechMed-Graz, Graz, Austria
    Contribution
    Conceptualization, Data curation, Supervision, Funding acquisition, Validation, Investigation, Writing - original draft, Project administration, Writing – review and editing
    For correspondence
    tea.pavkov@uni-graz.at
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7871-6680

Funding

Austrian Science Fund (FWF T-1239)

  • Nina Gubensäk

Austrian Science Fund (FWF DK W09)

  • Klaus Zangger

Austrian Science Fund (FWF P 29405)

  • Joachim Reidl

Land Steiermark (1109)

  • Klaus Zangger

Ministerio de Ciencia e Innovación and European Union Regional Development Fund (MICINN/AEI/FEDER/UE) (PID2021-128751NB-I00)

  • Isabel Usón

Austrian Science Fund (Biomolecular Structures and Interactions DOC 130)

  • Tea Pavkov-Keller
  • Markus Rotzinger
  • Klaus Zangger

Austrian Science Fund (Molecular Metabolism DOC 50)

  • Theo Sagmeister
  • Klaus Zangger

Fundación Martínez Escudero (Postdoctoral grant)

  • Bruno Di Geronimo

University of Graz

  • Joachim Reidl
  • Klaus Zangger
  • Tea Pavkov-Keller

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

Acknowledgements

We are grateful for the beam time on ID30A-3 at ESRF (Grenoble, France) for intensive diffraction screening and data collection, as well as for staff support during data collection measurements (BAG mx-1740). JR, KZ and TPK acknowledge the support of the field of excellence BioHealth at the University of Graz. We also thank the interuniversity programs NAWI Graz and BioTechMed for financial support. Furthermore, we acknowledge the financial support by Austrian Science Fund FWF for projects: T-1239 for NG, DK W09 for KZ, P 29405 for JR, and doc.fund projects Molecular Metabolism (DOC 50 for TS and KZ) and Biomolecular Structure and Interactions (DOC 130 for MR, KZ and TPK). Financial contributions by the Land Steiermark infrastructure grant “Frontier NMR”, project number 1109 are also gratefully acknowledged. Ministerio de Ciencia e Innovación and European Union Regional Development Fund (MICINN/AEI/FEDER/UE) are acknowledged for grant No. PID2021-128751NB-I00 (for IU) and BDG thanks Fundación Martínez Escudero for a Postdoctoral grant.

Senior Editor

  1. Amy H Andreotti, Iowa State University, United States

Reviewing Editor

  1. Hannes Neuweiler, University of Würzburg, Germany

Reviewer

  1. Hannes Neuweiler, University of Würzburg, Germany

Version history

  1. Received: April 20, 2023
  2. Preprint posted: May 4, 2023 (view preprint)
  3. Accepted: September 27, 2023
  4. Accepted Manuscript published: September 28, 2023 (version 1)
  5. Version of Record published: November 3, 2023 (version 2)

Copyright

© 2023, Gubensäk et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Nina Gubensäk
  2. Theo Sagmeister
  3. Christoph Buhlheller
  4. Bruno Di Geronimo
  5. Gabriel E Wagner
  6. Lukas Petrowitsch
  7. Melissa A Gräwert
  8. Markus Rotzinger
  9. Tamara M Ismael Berger
  10. Jan Schäfer
  11. Isabel Usón
  12. Joachim Reidl
  13. Pedro A Sánchez-Murcia
  14. Klaus Zangger
  15. Tea Pavkov-Keller
(2023)
Vibrio cholerae’s ToxRS bile sensing system
eLife 12:e88721.
https://doi.org/10.7554/eLife.88721

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