Recombinant Cytosolic Truncations of Histidine Kinases Retain Function for Targeted In Vitro Investigations

TL;DR

Researchers created functional parts of histidine kinases that work in lab tests, helping study bacterial signaling and drug development.

Contribution

The study introduces improved methods to produce functional cytosolic truncation mutants of histidine kinases from multiple bacterial species.

Findings

Cytosolic truncation mutants of histidine kinases retain autophosphorylation and phosphotransfer capabilities.

Functional mutants were successfully produced from Escherichia coli, Klebsiella pneumoniae, and other species.

These mutants are suitable for in vitro biochemical investigations and inhibitor screening.

Abstract

Histidine kinases are an integral component of bacterial two-component systems (TCSs), playing a pivotal role in signal transduction pathways, resulting in both resistance and virulence. However, their inherent membrane-bound nature often results in poor solubility, making them difficult to isolate and rendering them incompatible with most in vitro biochemical techniques. Consequently, much of the research on two-component systems has centered on response regulators, limiting both drug discovery efforts and our broader understanding of key signal transduction mechanisms. To address these challenges, we sought to straightforwardly generate cytosolic truncation mutants of histidine kinases that retain their autophosphorylation and phosphotransfer capabilities. Previously, we successfully developed a cytosolic truncation mutant of PmrB (PmrBc) that maintained these critical functions, demonstrating its suitability as a viable surrogate for in vitro investigations, including inhibitor compound screening. Building upon this foundation, we have refined our methods and here demonstrate these improvements by producing functional histidine kinase truncation mutants from the following diverse bacterial species: Escherichia coli; PhoQ, BasS and Klebsiella pneumoniae; and PmrB and PhoQ.

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1. Introduction

In 1962, histidine phosphorylation was first documented [1]. Since then, histidine kinases have been documented extensively throughout the prokaryotic, eukaryotic, and archaic domains [2]. Histidine kinases describe a broad superfamily of protein kinases, comprising at least 11 families determined by phylogenetic analysis, as well as the HWE (Histidine–Tryptophan–Glutamate) family of histidine kinases [3,4]. These proteins have garnered significant attention due to their function as key signal transduction proteins across the domains of life [5,6]. Histidine kinases are a core element in the critical regulatory systems known as two-component systems (TCS) [7]. Two-component systems typically function according to the following scheme: a stimulus is sensed by a structurally dimerized histidine kinase, upon which autophosphorylation on a conserved histidine residue occurs. Following this, the phosphoryl group from the conserved histidine on the kinase is transferred to a conserved aspartate residue on its cognate response regulator. The phosphorylated response regulator is then able to achieve the appropriate cellular response, typically by participating in transcriptional regulation [7]. Over 300,000 two-component systems have been identified [7]. In bacteria, they govern processes from virulence and motility to biofilm development and antibiotic resistance [8,9,10,11,12,13,14,15,16]. In fungi, they regulate several processes from cell wall biosynthesis and adherence to morphogenesis, and in plants, they play critical roles in both the signaling and developmental processes [17,18]. The number of two-component systems found in a single organism varies greatly, with bacterial species relying on tens to even hundreds of these systems, while their prevalence in yeast, fungi, plants, and archaea is significantly diminished [19,20]. Most notable is their apparent absence in the animal kingdom [2]. While histidine phosphorylation itself has been directly demonstrated, two-component system identification by genetic or structurally related means is absent [3,4]. This factor, combined with their regulation of several virulence and resistance-associated pathways, has made them key targets for antimicrobial therapeutics [21]. Recently, a class of histidine kinase inhibitors was shown to be effective in reducing colistin efficacy in Acinetobacter baumannii and was shown to do so by targeting the ATP-lid of the histidine kinase PmrB [22].

The structure and mechanistic details governing histidine kinase function are generally known [5,23]. Structurally, transmembrane histidine kinases contain both a membrane-bound sensor domain and a cytoplasmic domain that is responsible for exerting autophosphorylation, phosphotransfer, and phosphatase activity (Figure 1a) [23]. The membrane-bound portions are formed by various organizations of transmembrane, helical bundles [6]. In many histidine kinases, this sensor domain is followed by a HAMP (Histidine kinases, Adenylyl cyclases, Methyl-accepting chemotaxis proteins, and Phosphatases) domain, named after protein classes that frequently contain this motif (Figure 1b) [24]. This domain aids in signal transduction from the sensor domain to the cytoplasmic portion [24]. The HAMP domain is followed by the dimerization and histidine phosphotransfer domain (DHp), which contains the conserved site of phosphorylation (Figure 1b) [5,23]. This domain comprises a four-helical bundle, with each monomeric unit contributing two helices [23,24]. To complement this domain and form the catalytic core for these proteins, a catalytic ATP-binding (CA) domain is also found (Figure 1b) [24]. Histidine kinases are part of the GHKL (DNA Gyrase, Heat shock protein 90 (Hsp90), Histidine Kinase, and Mut L) protein family, which contain a characteristic Bergerat fold in their ATP-binding domains [25]. This domain contains the conserved N-, G1-, F, and G2- sequence motifs responsible for ATP-binding and hydrolysis. The CA domain is responsible for transferring the γ-phosphate to the histidine residue contained within the DHp domain [6]. Additionally, histidine kinases may contain Per-ARNT-Sim (PAS) and cGMP-specific phosphodiesterases, adenylyl cyclases and Fhla (GAF) domains that share a large degree of structural and functional similarity and serve to mediate signal transduction [26].

Despite the amount known about their structure and function, this class of proteins has, historically, been difficult to study due to their membrane-bound nature. The periplasmic domain contributes to solubility, as well as production issues that have hampered progress for high-throughput, in vitro drug-screening. Previous strategies have included the usage of either high amounts of glycerol or detergents to aid solubility, lipid nanodiscs, or protein truncation to remove the sensor domain [23,27,28,29]. While beneficial for solubilizing the sensor domain, detergents may disrupt the native structure of the cytosolic domain and therefore interfere or produce erroneous results in a high-throughput drug screening setting [30,31]. The usage of lipid nanodiscs is complex and requires considerable optimization to produce stable and homogenous results [32]. Truncation of the sensor domain has been implemented sporadically, yet it has been claimed that truncation in this manner produces artifacts in activity assays [27]. Contrary to this final claim, we have found that by implementing a structure-guided truncation approach using AlphaFold predicted structures for the histidine kinases, we are able to preserve their critical functions. Following truncation of the sensor domain, they retain the expected secondary structure, multimerization state, and ATP-binding characteristics, as well as the ability to autophosphorylate and to phosphotransfer to their cognate response regulators. To demonstrate the broad applicability of this approach, here, we show this for PmrB from Klebsiella pneumonia, PhoQ from both Escherichia coli and Klebsiella pneumonia, and for BasS from Escherichia coli.

2. Materials and Methods

2.1. Reagents

Buffer components were obtained from Chem-Impex (NaCl and Isopropyl-β-D-thiogalactopyranoside) and Thermo Fisher Scientific (Tris-HCl and MgCl_2_). TNP-ATP was purchased from Thermo Fisher Scientific (Waltham, MA, USA). [γ^−32^] ATP was obtained from Revvity (Waltham, MA, USA).

2.2. Vectors

The vectors used for recombinant protein expression in this study were purchased from Genscript (Piscataway, NJ, USA). The genes of interest were cloned into pET28a vectors using BamHI and NdeI restriction sites, placing the genes of interest directly following a His_6_ tag and a thrombin cleavage sequence found in the backbone of the pET28a vector.

2.3. AlphaFold Models

Primary sequences for the selected histidine kinases from the specified organisms were obtained from the Uniprot web database. The primary sequences for the full-length proteins were submitted directly to the AlphaFold web server, with the specification for two copies of each monomeric sequence to allow for visualization of the structural dimer. For each model depicting the His_6_-tagged histidine kinase truncation, the sequence was obtained by first translating the DNA sequence for the pET28a vector purchased from Genscript, using the translate tool found in the collection of web-based tools provided by Expasy. Following identification of the translated product of interest, the protein sequence, including the N-terminal His_6_ tag and thrombin cleavage sequence, was submitted directly to the AlphaFold web server with the specification for two copies of each monomeric sequence.

2.4. Recombinant Protein Expression and Purification

BL21(DE3) cells (Invitrogen) containing the desired vector were grown at 37 °C, 200 rpm to an OD600 of 0.6–0.8, prior to induction with 1 mM Isopropyl-β-D-thiogalactopyranoside (IPTG). Following induction, the bacterial growths were incubated at 18 °C, 120 rpm overnight. Cells were harvested by centrifugation at 5000 rpm. The resultant pellets were stored at −80 °C prior to purification. To isolate the desired histidine kinase truncation or response regulator, the bacterial pellet was resuspended in 20 mM Tris pH 7.5, 400 mM NaCl and lysed by sonication. The lysate was clarified by centrifugation at 15,000 rpm and the supernatant was loaded onto an equilibrated Ni-NTA column to allow for isolation of the His_6_-tagged protein of interest via immobilized metal affinity chromatography (Qiagen). The immobilized recombinant protein was washed first with 20 mM Tris pH 7.5, 1 M NaCl, prior to washing with 20 mM Tris pH 7.5, 400 mM NaCl, 5 mM imidazole. Elution was achieved using a linear gradient from 5 to 350 mM imidazole. Fractions were analyzed by SDS-PAGE on a 14% polycacrylamide gel at 200 V to provide an effective resolution for the kinases, which range from 32.7 to 33.0 kDa, as well as for the smaller response regulators, which range from 25.0 to 25.5 kDa. These gels were then Coomassie-stained to visualize the protein species present in the elution fractions. The identity was determined by comparison to a molecular weight ladder, while the purity was assessed by evaluating % composition by lane in ImageLab (BioRad, Hercules, CA, USA). Fractions containing > 95% protein of interest were pooled and dialyzed against 4 L of 20 mM Tris pH 7.5, 400 mM NaCl, 5 mM MgCl_2_, prior to storage at 4 °C. Representative purification gels are shown in the supplementary data (Supplementary Figure S1).

2.5. Circular Dichroism Spectroscopy

The 20 µM samples for each of the truncated histidine kinases were prepared in 20 mM Tris pH 7.5, 400 mM NaF, 5 mM MgCl_2_. Spectra were recorded from 180 to 260 nm with a 0.5 step size and 1 s read intervals on a Chirascan V100 (Applied Photophysics, Leatherhead, Surrey, UK). These spectra were recorded in triplicate. Data analysis was performed in ProDataViewer (Applied Photophysics, 4.5). Background spectra for the 20 mM Tris pH 7.5, 400 mM NaF, 5 MgCl_2_ were averaged and subtracted from the sample spectra prior to conversion to molar ellipticity. Final figures were prepared in Graphpad (Prism, 10.4.1).

2.6. Fluorescent ATP-Binding Assay

(2′-(or-3′)-O-(Trinitrophenyl) Adenosine 5′-Triphosphate (TNP-ATP) was utilized to determine the ATP-binding affinity for the truncated kinases. The 10 µM final concentrations for each histidine kinase were incubated with the indicated concentrations of TNP-ATP for 10 min prior to reading on a Synergy H1 (BioTek, Winooski, VT, USA) plate reader. For each indicated concentration of substrate, a sample lacking protein was also prepared. Samples were excited at 403 nm and fluorescent emission at 538 nm was recorded and plotted against the substrate concentration. The readings for the background samples at each concentration of substrate were subtracted from the protein containing samples to account for the weakly fluorescent nature of the probe in water. The processed data were plotted in Graphpad (Prism, 10.4.1) and Kd was determined by nonlinear regression, using a one-site specific binding model.

2.7. Analytical Ultracentrifugation

Samples of each histidine kinase were prepared to achieve final A280 readings of 0.3, 0.5, and 0.8 when adjusted for the 1.2 cm path length of the centerpiece used. Centrifugation was performed at 42,000 rpm on an Optima AUC (Beckman-Coulter). Scans were taken at 60 s intervals for a total of 300 scans per cell. The parameters for buffer viscosity, protein partial specific volume, and buffer density were calculated using SEDNTERP (RASMB, 20130813). Every third scan was loaded into SEDFIT for molecular weight determination, using a continuous c(s) distribution model. The resultant peaks with calculated molecular weights were exported and plotted in Graphpad (Prism, 10.4.1) to generate the final figures.

2.8. [γ−32] ATP Autophosphorylation

To assess autophosphorylation activity, 20 µM histidine kinases were incubated with 15 µM ATP and 66.6 nM radiolabeled ATP. Time points were taken at 0 min, 30 min, 1 h, and 2 h and added to 1× SDS loading dye (62.5 mM Tris, 1% SDS, 8% glycerol, 1.5% 14.7 M 2-mercaptoethanol, and 0.005% Bromophenol blue) supplemented with 10 mM EDTA to quench the reactions. The samples were analyzed by SDS-PAGE on a 14% polyacrylamide gel (Invitrogen, Carlsbad, CA, USA). Following electrophoresis, the gels were dried in 30% methanol and 3% glycerol between cellophane sheets. The dried gels were exposed on a phosphor-imaging screen for 1 h prior to visualization, using an Amersham Typhoon imager (GE Healthcare, Chicago, IL, USA).

2.9. [γ−32] ATP Phosphotransfer

To determine whether the phosphotransfer function was retained in our truncated histidine kinases, 20 µM histidine kinase was first incubated with 66.6 nM radiolabeled ATP for 1 h at 37 °C. Following this incubation period, a response regulator was added to a final concentration of 20 µM, as well as 500 µM unlabeled ATP to reduce the propensity for phosphatase activity by the truncated kinases. Time points were taken as indicated and analyzed by SDS-PAGE on a 14% polyacrylamide gel, following quenching in 1× SDS loading dye supplemented with 10 mM EDTA. The electrophoresed gels were dried in 30% methanol and 3% glycerol between cellophane sheets prior to exposure on a phosphor-imaging screen for 1 h. The exposed screen was analyzed using an Amersham Typhoon imager (GE Healthcare) to visualize the protein species carrying the radiolabel.

3. Results

3.1. Determination of Truncation Sites

To aid in solubilizing the histidine kinases, cytosolic truncations were generated (Figure 1c). To determine where the truncations would occur, a predictive, structure-guided approach was taken. AlphaFold3 was implemented to produce models for the full-length versions of each kinase (Figure 1c). The resultant structures showed cytosolic domains that were similar to those seen in the previously crystallized HK853 from Thermotoga maritima (PDB: 2C2A) and a sensor domain resembling that seen in Geobacter sulfurreducens HK29s (PDB: 3H7M).

TMHMM was initially used to determine the first residues located within the cytosol (Figure 2a–d). Following the identification of these residues, an inspection of the primary sequence was performed within that region to identify the helix-terminating residues that were most proximal to those residues. For the proteins analyzed in this study, the helix in question belonged to the HAMP domain. We utilized the primary sequence to search for helix-disrupting residues, such as proline or glycine associated with the beginning of the HAMP domain (Figure 1c). We utilized AlphaFold3 to provide additional confirmation that the helix-disrupting nature of these residues was reflected in the structural models. In the absence of proline or glycine residues in this region, AlphaFold3 would be utilized to determine the helix-terminating residues. Once the terminal residues were determined, an additional two to five residues preceding the helix-terminators were left to aid in appropriate helical capping, while avoiding helix destabilization due to entropic factors caused by a lengthy loop [33,34]. The additional residue length is primarily determined by the hydrophobicity of the residues within that area, with the goal of limiting the number of hydrophobic residues. The helix-disrupting residues are highlighted with a blue rectangle, with the selected cut sites shown by red bars (Figure 3).

3.2. Evaluation of Secondary Structure

To assess whether the expected folds for each histidine kinase were adopted following recombinant expression and affinity purification, circular dichroism (CD) spectroscopy was used. The secondary structural elements were determined by far-UV CD display negative minima at 222 and 208 nm, as well as displaying a positive maxima trending towards 192 nm (Figure 4). Each of the selected histidine kinase truncations display a high level of helical content, which aligned with the predicted spectra, as determined using the PDBMD2CD web server based on AlphaFold3 generated models for each truncation.

3.3. Multimerization State for Recombinant Cytosolic Truncations

The multimerization state for the recombinantly produced histidine kinase truncations was assessed by analytical ultracentrifugation under sedimentation-velocity conditions (SV-AUC). The monomeric weights for each truncated kinase, as well as the experimentally determined MW for the monomeric and dimeric populations, are shown in Table 1. There is good agreement between the theoretical and experimentally determined sizes for both the monomers and dimers for all the truncated kinases. These results show notable monomeric and dimeric populations, indicating the presence of monomer–dimer equilibriums in these truncations. The Kp PhoQc and Ec PhoQc dimeric populations are increased when compared to those of Ec BasSc and Kp PmrBc and show few changes in the distribution between populations as the concentration is increased (Figure 5a–l). Kp PmrBc shows marginal increases in its dimeric population as the concentration is increased (Figure 5a–c).

3.4. ATP-Binding Affinity

The histidine kinase truncations were assessed for their ability to bind ATP via a (2′-(or-3′)-O-(Trinitrophenyl) Adenosine 5′-Triphosphate (TNP-ATP) binding assay. TNP fluorescence is readily quenched in an aqueous, polar environment, yet readily exhibits fluorescence when stabilized by the hydrophobic ATP-binding cleft of an ATP-binding protein [35]. This allows for dissociation of constant determination between ATP and the truncated histidine kinase. For each of the kinases studied, the determined K_d_ values were in the low micromolar range (Figure 6a–d). These values fall within the expected range for a histidine kinase, which are reported between low- and mid-micromolar [36,37,38,39].

3.5. Autophosphorylation and Phosphotransfer

Due to the critical function of histidine kinases as signal transduction modules to their cognate response regulators, their ability to both autophosphorylate at their conserved histidine and transfer the phosphate to their cognate response regulator is necessary. The histidine kinases show time-dependent autophosphorylation when incubated with radiolabeled [γ^−32^] ATP in the absence of a response regulator (Figure 7). Following the addition of the response regulator, the transfer of the phosphoryl group is seen from the truncated histidine kinase to the response regulator (Figure 8a–d).

4. Discussion

Transmembrane histidine kinases have proved difficult to study in vitro and, as such, progress in developing effective inhibitors for these critical signaling modules has been hindered. Difficulties in studying these proteins in vitro arise largely due to the sensor domain, which exhibits significant hydrophobicity. The hydrophobicity of this domain often necessitates complicated purification strategies and may require the addition of detergents, further hindering their compatibility with many in vitro investigations, including high-throughput drug-screening.

Truncation has been implemented previously, with success in achieving soluble forms of the histidine kinases that display both autophosphorylation and phosphotransfer capacities [23,28,40]. Despite this, it has been posited that truncation for this class of proteins may produce artifacts in activity assays [27]. We sought to address this point by thoroughly characterizing several histidine kinase truncations produced using our design strategy.

Given the abundance of known histidine kinase sequences, the ability to analyze sequences for their likely locations with respect to a membrane, and the availability of protein structure prediction software, we had previously utilized a combination of sequence alignments, TMHMM predictions, and AlphaFold models to produce a truncated form of PmrB from Acinetobacter baumannii. This truncated form allowed for screening against several kinase inhibitors and allowed for binding site determination and indication behind its mechanism of action [22].

To test the broad applicability of our truncation design strategy, histidine kinases were selected from both E. coli and K. pneumoniae to cover aberrant characteristics in structure or sequence alignment within the truncation region. Despite sequence variation and minor structural deviations between these proteins, we confirmed that our truncation strategy produced high yields of soluble histidine kinases that remain soluble in common, inexpensive, and highly compatible laboratory reagents. Additionally, these truncations minimally affected the protein structure and function as demonstrated by circular dichroism, analytical ultracentrifugation, TNP-ATP binding assays, and radiolabeled autophosphorylation and phosphotransfer reactions.

Secondary structures deviated only slightly from the predicted structures for all the studied truncations. These minor deviations likely result from the Tris and Cl^−^ ions found in the buffer solution. The multimerization state, as determined by AUC, shows that all the truncations can form the dimeric species required for autophosphorylation and subsequent phosphotransfer capacities. Without previously existing multimerization studies for the full-length forms of these proteins, it is possible that the monomer–dimer equilibrium is a typical feature for this class of proteins. These proteins rely on dimer formation to form the required four-helix bundle in the HAMP domain. It is clear from our autophosphorylation and phosphotransfer data that the presence of these monomers does not impede the typical catalytic functions expected for these proteins (Figure 7 and Figure 8). It is also possible that the sedimentation-velocity parameters used in these AUC studies favor detection of the monomeric form for these proteins. The hydrostatic pressure gradient within the AUC cell may place additional pressure on the dimer as it sediments lower in the cell, with the pressure gradient encouraging dimer dissociation if the dimer interface is weak or if the equilibrium is rapid. There is also the consideration for dilution at the sedimentation boundary that may promote the monomeric state. As the dimer population sediments faster, there is a localized dilution effect as the monomer and dimer populations separate. The monomer–dimer equilibrium would then be affected in favor of the monomer as the dimeric population attempts to re-establish the equilibrium at the boundary. TNP-ATP binding assays revealed a similar ATP-binding affinity for each of the truncations, ranging from 11.31 to 34.46 µM. Typically, ATP-binding studies utilizing TNP-ATP have been shown to display an increased affinity for the TNP-ATP when compared to ATP. However, taking the modest difference in affinity into account, the ATP-binding affinity remains within the expected range for histidine kinases and is similar to affinity constants determined using TNP-ATP for other histidine kinases [36,37,38,39]. The autophosphorylation profiles, as determined through radiolabeled autophosphorylation assays, show remarkable similarity across the truncations, with increasing saturation of the radiolabeled population until the 2 h mark for each. Our ability to assess effects on autophosphorylation with these truncations has demonstrated its uses for assessing the kinase inhibitor efficacy for Ab PmrBc and will be extremely valuable for future investigations into inhibitors for many more of these kinases [22]. While the overall intensities of the phosphorylated kinases and response regulators appear to show differences across the time points, certain considerations must be accounted for prior to analysis of phosphotransfer kinetics. Due to the monomer–dimer equilibrium and the dimeric form being the active state for these proteins, the concentration of dimeric protein under the experimental conditions used must be known to allow for appropriate normalization. Additionally, the rate of combined spontaneous dephosphorylation by the response regulator as well as phosphatase activity by the truncated kinase must also be known. Given that the truncated kinases demonstrate capacity for phosphotransfer, accounting for the abovementioned factors would make them suitable for future kinetic studies. Variations in phosphotransfer rates between different histidine kinases has previously been documented [40,41]. Furthermore, the significance of varying phosphotransfer rates to mutant response regulators has been implicated in enhancing colistin resistance in Acinetobacter baumannii. Utilizing a truncated histidine kinase generated following these truncation guidelines, it was determined that phosphotransfer was impeded in a number of mutants of the response regulator PmrA from Acinetobacter baumannii that were noted to have increased colistin resistance [42]. Considering the number of two-component systems found in bacterial species and the considerable level of crosstalk that occurs between these systems, this truncation strategy would aid our ability to evaluate how these differences in phosphotransfer rates affect bacterial signaling by providing a simple method for producing the extensive number of kinases existing in these organisms while maintaining their functional integrity. While protein insolubility imparted by the presence of the sensor domain renders full-length kinases incompatible with many in vitro techniques, this issue is further exacerbated in the case of structural determination techniques. With protein concentrations typically ranging from 5 to 20 mg/mL for evaluation by X-ray crystallography and NMR, the solubility of the protein of interest is critically important. The truncated histidine kinases generated by this strategy allow for these high concentrations to be achieved. This factor, combined with the substantial overall yield of protein generated through the expression protocol, allows for both high volumes and concentrations of the truncated histidine kinase to be achieved. These histidine kinase truncations would allow for the expansion of histidine kinase structures found in the PDB, providing the information needed to enhance accuracy in models for this class of proteins, as well as improve our ability to accurately evaluate these proteins in silico.

Due to the incompatibility between full-length histidine kinases and most biochemical techniques, we are unable to directly compare the truncation mutants to their full-length counterparts. However, until this incompatibility is resolved, we provide this alternative approach with confidence in its ability to expand our investigative ability in the realms of bacterial histidine kinases and two-component systems.