A cytoplasmic chemoreceptor and reactive oxygen
species mediate bacterial chemotaxis to copper

Gwennaëlle Louis, Pauline Cherry, Catherine Michaux, Sophie
Rahuel-Clermont, Marc Dieu, Françoise Tilquin, Laurens Maertens, Rob van
Houdt, Patricia Renard, Eric Perpete, et al.

► To cite this version:

Gwennaëlle Louis, Pauline Cherry, Catherine Michaux, Sophie Rahuel-Clermont, Marc Dieu, et al. A cytoplasmic chemoreceptor and reactive oxygen species mediate bacterial chemotaxis to copper. Journal of Biological Chemistry, 2023, 299 (10), pp.105207. 10.1016/j.jbc.2023.105207. hal-04308375

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# A cytoplasmic chemoreceptor and reactive oxygen species
mediate bacterial chemotaxis to copper

Received for publication, March 10, 2023, and in revised form, August 16, 2023. Published, Papers in Press, September 1, 2023, https://doi.org/10.1016/j.jbc.2023.105207

Gwennaëlle Louis¹, Pauline Cherry¹, Catherine Michaux² ID, Sophie Rahuel-Clermont³ ID, Marc Dieu⁴ ID, Françoise Tilquin¹, Laurens Maertens¹,⁵ ID, Rob Van Houdt⁶ ID, Patricia Renard⁴ ID, Eric Perpete², and Jean-Yves Matroule¹,* ID

From the ¹Research Unit in Biology of Microorganisms (URBM), Department of Biology, Namur Research Institute for Life Sciences (NARILIS), and ²Laboratoire de Chimie Physique des Biomolécules, Namur Research Institute for Life Sciences (NARILIS) and Namur Institute of Structured Matter (NISM), University of Namur, Namur, Belgium; ³CNRS, IMoPA, Université de Lorraine, Nancy, France; ⁴MaSUN, Mass Spectrometry Facility, University of Namur, Namur, Belgium; ⁵Microbiology Unit, Interdisciplinary Biosciences, Belgian Nuclear Research Centre (SCK CEN), Mol, Belgium

Reviewed by members of the JBC Editorial Board. Edited by Ursula Jakob

Chemotaxis is a widespread strategy used by unicellular and multicellular living organisms to maintain their fitness in stressful environments. We previously showed that bacteria can trigger a negative chemotactic response to a copper (Cu)-rich environment. Cu ion toxicity on bacterial cell physiology has been mainly linked to mismetallation events and reactive oxygen species (ROS) production, although the precise role of Cu-generated ROS remains largely debated. Here, using inductively coupled plasma optical emission spectrometry on cell fractionates, we found that the cytoplasmic Cu ion content mirrors variations of the extracellular Cu ion concentration. ROS-sensitive fluorescent probe and biosensor allowed us to show that the increase of cytoplasmic Cu ion content triggers a dose-dependent oxidative stress, which can be abrogated by superoxide dismutase and catalase overexpression. The inhibition of ROS production in the cytoplasm not only improves bacterial growth but also impedes Cu chemotaxis, indicating that ROS derived from cytoplasmic Cu ions mediate the control of bacterial chemotaxis to Cu. We also identified the Cu chemoreceptor McpR, which binds Cu ions with low affinity, suggesting a labile interaction. In addition, we demonstrate that the cysteine 75 and histidine 99 within the McpR sensor domain are key residues in Cu chemotaxis and Cu coordination. Finally, we discovered that *in vitro* both Cu(I) and Cu(II) ions modulate McpR conformation in a distinct manner. Overall, our study provides mechanistic insights on a redox-based control of Cu chemotaxis, indicating that the cellular redox status can play a key role in bacterial chemotaxis.

Cell motility is widely distributed across the three domains of life. For instance, in animals, it is involved in fertilization, and in neutrophils recruitment to an inflammatory area. It also allows archaea and bacteria to seek nutrients in scarce environments and to flee from toxic compounds while it supports to some extent host tropism in the frame of symbiotic relationships.

Most of the time, cell motility is driven by chemical gradients and is therefore named chemotaxis. In flagellated bacteria, positive (attraction) and negative (repulsion) chemotactic responses imply stochastic direction changes triggered by the inversion of flagellar rotation (1, 2). This switch is caused by the binding of the phosphorylated response regulator, CheY~P, to the flagellar motor. Spontaneous or phosphatase-mediated CheY~P dephosphorylation by the CheZ phosphatase or by a member of the CheC/CheX/FliY phosphatase family triggers CheY release from the flagellar motor (3–5). The release of CheY promotes a straight bacterial swimming pattern until CheY is phosphorylated again and triggers a new direction change. CheY phosphorylation relies on the histidine kinase CheA, which is recruited and autophosphorylated within a chemotaxis cluster composed of cytoplasmic or inner membrane-spanning methyl-accepting chemotaxis proteins (Mcp), also named chemoreceptors (6). Chemoreceptors are classically composed of an N-terminal sensing domain, transmembrane (TM) domains (which are absent in cytoplasmic chemoreceptors), a HAMP domain, and the C-terminal (Mcp) signaling domain. Depending on their cellular topology, chemoreceptors monitor chemical variations in the cytosol or in the periplasm via their N-terminal sensing domain. Upon substrate binding to their sensing domain, chemoreceptors undergo a conformational change leading to the increase (repellent response) or the decrease (attractant response) of CheA histidine kinase activity (7, 8). In *Escherichia coli* Tar and Tsr chemoreceptors, this conformational change consists in a 2 Å shift of one of the TM helices (TM2), causing a “piston-like” movement in the direction of the membrane that impacts the HAMP domain (9). The methylation of the Mcps by the CheR methyltransferase results in an increase of CheA activity, and their demethylation by the CheA-activated CheB methylesterase leads to the decrease of CheA activity (9). This adaptation response sets back the Mcps in a prestimulus state.

Copper (Cu) has been used for centuries as an antimicrobial and antifouling strategy in medicine and in various

<footnote>* For correspondence: Jean-Yves Matroule, jean-yves.matroule@unamur.be.</footnote>
## Bacterial chemotaxis to copper and oxidative stress

anthropogenic activities including agriculture and industry. Cu toxicity in solution mainly results from its high redox potential causing the displacement of native metallic ions from metalloproteins and the disruption of Fe–S clusters (10). Cu ability to generate reactive oxygen species (ROS) has been shown in vitro (11), but the role of these ROS in vivo yet remains unclear. In a previous study, we identified an original bimodal strategy to cope with Cu stress where the flagellated morphotype (swarmer [SW] cell) of *Caulobacter crescentus* triggers a prompt negative chemotaxis from the Cu source (12). In addition to the negative Cu chemotaxis, *C. crescentus* exhibits a positive chemotactic behavior toward xylose (13) and $O_2$ (14), but no dedicated Mcp could be identified so far.

*C. crescentus* genome harbors two chemotaxis operons. The major chemotaxis operon encodes McpA and McpB, CheYI and CheYII, CheAI, one CheW, CheRI, CheBI, and the CheY-like CheA. This operon also contains the *cheD*, *cheU*, and *cheE* genes, which have been hardly described. CheYII and CheBI are likely the major CheY and CheB in *C. crescentus* chemotaxis (15, 16). The alternate chemotaxis operon encodes McpG and McpK, two CheY, CheAII, one CheW, CheBII, and CheRII. The other *cheY* genes and the third *cheW* are dispersed within the genome. Only the first chemotaxis operon seems to be essential for chemotaxis, whereas both operons are involved in biofilm formation and holdfast production, by regulating the expression of the *hfiA* holdfast inhibitor (16, 17).

No CheZ homolog has been found in *C. crescentus* genome, but the multiple CheY homologs are proposed to act as phosphate sinks to allow the termination process, such as in *Rhizobium meliloti* (18). Five of the CheY have been redefined as Cle proteins (CheY-like c-di-GMP effectors) and renamed CheA to CheE. CheA and CheD could compete with CheYII to prevent the rotational switch of the flagellum and to promote straight run (15). The *C. crescentus* Mcp array exhibits a higher-ordered hexagonal structure constituted of approximately 1000 to 2000 Mcps per array (19). Among the *C. crescentus* Mcps, only the inner membrane McpA and the cytoplasmic McpB encoded by the major chemotaxis operon were described as polar chemoreceptors, although their exact function remains unknown (20).

In the present study, we aimed to decipher the sensing mechanism underlying *C. crescentus* chemotaxis to a toxic Cu-rich environment. We provide evidence that an increased intracellular Cu concentration triggers a dose-dependent production of ROS, which is a prerequisite to Cu chemotaxis. We also demonstrate that the redox status of the Cu ions bound to the sensor domain of the newly identified McpR modulates McpR conformation *in vitro*. This original redox sensing mechanism may ensure a continuous monitoring of the intracellular Cu status, providing a real-time control of the chemotactic response.

## Results

To further dissect the molecular mechanisms underlying the negative Cu chemotaxis of *C. crescentus* SW cells (12), we first sought to identify the chemoreceptor involved in Cu

chemotaxis. The genome of *C. crescentus* is predicted to encode 19 chemoreceptors (named McpA to McpS) based on the presence of an Mcp domain binding CheA/CheW (21). No canonical Cu ion-binding site, including the CXXC motif or the βαββαβ fold (22), could be found by running manual and JPred4-based analyses of the 19 chemoreceptor protein sequences (23). Therefore, we proceeded to single in-frame deletions of the 19 *mcp* genes, and their respective chemotactic behavior to Cu ions was assessed by using the previously designed live chemotaxis imaging (LCI) (12). Most of the *Δmcp* mutants displayed a WT-like behavior with a flight percentage after 25 min varying between 15% and 51%. Only *ΔmcpA*, *ΔmcpB*, and *ΔmcpR* mutants showed a significant decrease of Cu chemotaxis relative to the WT strain (Fig. 1A), suggesting that these chemoreceptors might be involved in Cu chemotaxis. The *ΔmcpA*, *ΔmcpB*, and *ΔmcpR* mutants could be complemented by the expression of an ectopic version of *mcpA*, *mcpB*, and *mcpR* on a low copy plasmid, respectively, ruling out any polar effect of the deletions (Fig. S1). Owing to the position of the *mcpA* gene within the major chemotaxis cluster (16), McpA is likely involved in chemotaxis to a less specific extent. Consistent with this hypothesis, the *ΔmcpA* mutant displays a reduced motility halo in a Cu ion-free swarming assay, whereas the *ΔmcpB* and *ΔmcpR* mutants exhibit a WT-like chemotaxis profile in the same condition (Fig. S2A). The presence of the terminal sequence of the CheB/ CheR docking site in McpB (absent from McpR) led us to the idea that McpB might be necessary for the adaptive response as in *E. coli* (24), which will not be considered in this study. Therefore, we decided to determine how McpR modulates Cu chemotaxis.

The ability of the *ΔmcpR* mutant to form a motility halo in a Cu ion-free swarming assay (Fig. S2A) and to sustain negative chemotaxis to other chemicals, such as zinc (Zn) ions (Fig. S2B), indicates that the loss of Cu chemotaxis in the *ΔmcpR* mutant does not result from a loss of motility and/or a disruption of the general chemotactic machinery.

McpR harbors the canonical domain organization found in chemoreceptors including an N-terminal sensing domain followed by a HAMP domain and a C-terminal Mcp domain. McpR sequence analysis using the TMHMM 2.0 (DTU Health Tech) (25) and SignalIP 6.0 (DTU Health Tech) (26) tools did not reveal any McpR TM domain and signal peptide, respectively, arguing for a cytoplasmic localization for McpR, and therefore a sensing of the cytoplasmic Cu content.

Chemoreceptors can sense attractants or repellents in a direct manner or an indirect manner *via* their sensing domain (27). The 3D structure prediction of McpR by Alphafold2 (28, 29) highlights a cysteine (C75) in the vicinity of a histidine (H99) in the McpR sensing domain (Fig. 1B). Cysteine and histidine residues have often been described as key residues in Cu coordination (30), suggesting that McpR may bind Cu ions directly.

Therefore, we used intrinsic fluorescence spectroscopy to assess the direct binding of Cu to the purified McpR. Upon the addition of either Cu(I) ions or Cu(II) ions, the intrinsic McpR fluorescence at 346 nm is quenched in a dose-dependent
Bacterial chemotaxis to copper and oxidative stress

<figure><img src="image_2.png" /><figcaption>**Figure 1. McpR is involved in Cu chemotaxis and Cu binding.** A, percentage of WT and Δ*mcpA* (mcpA to mcpS) SW cells remaining in the vicinity of the Cu plug after 25 min. B, 3D structure prediction of McpR by AlphaFold. H99 and C75 residues are highlighted. C, McpR titration by Cu(I) (full squares) or Cu(II) (circles) monitored by protein intrinsic fluorescence intensity quenching, fitted to a single-site hyperbolic model by nonlinear regression analysis (solid line). The fluorescence intensity changes are normalized to the maximum fluorescence intensity change. D, number of Cu atoms per spheroplast from SW cells exposed to various Cu concentrations for 5 min. Mean ± SD, at least three biological replicates. *p* Values were calculated using an ANOVA (*p* < 0.05, **p** < 0.01, and ***p** < 0.001) (Table S1). Cu, copper; SW, swarmer cell.</figcaption></figure>

manner, reflecting a modification of the McpR aromatic residue environment (Fig. 1C). The dissociation constants ($K_d$) (see Experimental procedures section for calculation method) of 48 µM ± 10 µM for Cu(I) ions and 37 µM ± 9 µM for Cu(II) ions attest to a similar affinity of both ionic forms for McpR. However, the micromolar range of Cu(I) ion and Cu(II) ion $K_d$'s exceeds from several logs the $K_d$ of well-known cuproproteins (31, 32), arguing for a rather labile interaction between McpR and Cu. Nevertheless, McpR seems to preferentially bind Cu ions since Zn, cadmium (Cd), manganese, and nickel ions had no effect on purified McpR intrinsic fluorescence (Fig. S3).

To determine whether the cytoplasmic Cu ion content could reflect variations of the extracellular Cu ion concentration when the SW cell swims away from a Cu ion source, we measured by inductively coupled plasma optical emission spectrometry (ICP-OES) the Cu ion content in cytoplasmic fractions from SW cells subjected to increasing Cu ion concentrations. Although the measurement cannot discriminate between free and complexed Cu ions, Cu(I) and Cu(II), we measured a dose-dependent accumulation of Cu ions in the cytoplasm (Fig. 1D). This observation is in accordance with previous data from our laboratory, showing that an artificial decrease of Cu ion content via the overexpression of the PcoB
Bacterial chemotaxis to copper and oxidative stress

efflux pump impeded Cu chemotaxis (12). Together, these
data suggest that the sensing of cytoplasmic Cu ions by McpR
could be used as a proxy by the SW cells to mirror variations of
the concentration of Cu ions in C. crescentus environment to
control Cu chemotaxis.

The transient maintenance of a high intracellular Cu ion
content in the SW cells may potentially lead to an oxidative
stress *via* the production of superoxide anion (O₂⁻), hydrogen
peroxide (H₂O₂), and hydroxyl radical (OH⁺) (Fig. S4A). In line
with this idea, the relative fluorescence intensity of the ROS-
sensitive turn-on CellROX Deep Red (ThermoFisher) probe
is significantly increased in SW cells exposed to Cu ions, and
H₂O₂, the latter being used as a positive control (Fig. 2A).
Accordingly, the fluorescence of the cytoplasmic turn-off
rxYFP redox biosensor is partially quenched under the same
conditions (Fig. 2B), mirroring an imbalance of the cyto-
plasmic GSH-GSSG pool upon oxidative stress (Fig. S4B).
These findings are reinforced by the differential expression of
genes involved in oxidative stress response upon Cu stress,
including those encoding the catalase KatG and the superoxide
dismutases SodA and SodB (33). As observed for the cyto-
plasmic Cu ion content (Fig. 1D), the amount of ROS is
increased when the SW cells are exposed to increasing
extracellular Cu ion concentrations, indicating that the cyto-
plasmic ROS level mirrors the cytoplasmic Cu ion content
(Fig. 2C).

We envisioned that boosting the antioxidant defense system
would limit ROS production by Cu ions. Therefore, we
ectopically expressed the *sodB* gene or the *katG* gene in the
WT background from a low copy plasmid under the control of
the strong and constitutive *lac* promoter. The resulting *sodB*⁺
and *katG*⁺ strains prevent the increase of the CellROX Deep
Red fluorescence upon SW cell exposure to Cu ions, reflecting
a potentiation of antioxidant defenses (Fig. 2D). In support of
these data, the growth of these two optimized *sodB*⁺ and *katG*⁺
strains is improved under moderate Cu stress (Fig. 2E), arguing
for a role of O₂⁻ and H₂O₂ in Cu toxicity.

Interestingly, the sodB⁺ and katG⁺ strains are impeded in Cu
chemotaxis similarly to the ΔcheAI mutant (Fig. 2F, SI Ap-
pendix, and Fig. S5), indicating that the ROS resulting from the
accumulation of cellular Cu ions play a role in Cu chemotaxis.
It is worth mentioning that the loss of Cu chemotaxis in the
ΔmcpR mutant is not because of a loss of ROS production
under Cu stress since the ΔmcpR mutant exhibits a similar
decrease of the rxYFP fluorescence to the WT strain upon Cu
ion exposure (Fig. S6). These data support a major role of the
ROS resulting from Cu ion accumulation in Cu chemotaxis
and indicate that Cu ions binding to McpR are solely not
sufficient to sustain Cu chemotaxis.

An LC-MS analysis of McpR revealed a significant
oxidation of the McpR sensor domain (residues 1-161) and
more specifically of the H99 residue when purified McpR is
incubated *in vitro* with Cu(I) ions + H₂O₂ and with Cu(I)
ions, although at a lower extent (Fig. 3A, left panel). In Cu(I)
ions + H₂O₂ condition, about 30% of the H99 residues ac-
quired an additional oxygen atom, most likely yielding
2-oxoHis. Metal ion-catalyzed oxidation of histidines *via* the

generation of OH• has been previously described in the *Ba-
cillus subtilis* PerR repressor (34), yeast SOD1 (35), and
amyloid beta peptide (36). The exact role of this modification
remains unknown, and no redox cycling involving 2-oxoHis
has been described so far, questioning the relevance of H99
oxidation in Cu/ROS sensing. Metal ion-catalyzed oxidation
mostly targets the metal-binding residues in a caged process
(37), reinforcing the Cu binding to McpR (Fig. 1C) and
suggesting that H99 may coordinate Cu. To test this hy-
pothesis, we purified an McpR<sub>H99A</sub> mutant protein and
calculated the *K*<sub>d</sub> values of the McpR<sub>H99A</sub>/Cu(I) ion and
McpR<sub>H99A</sub>/Cu(II) ion complexes (Figs. 3B and S7A for the
*fitting curves*). H99A mutation triggers a 9.4- and 5.8-fold
increase of Cu(I) ion and Cu(II) ion *K*<sub>d</sub>s, respectively, rela-
tive to McpR, suggesting a key role of H99 in Cu(I) ion and
Cu(II) ion coordination. The important variability associated
with the McpR<sub>H99A</sub> *K*<sub>d</sub> values likely reflects a potential
instability of this mutant in solution upon Cu(I) ion and
Cu(II) ion exposure, resulting in the presence of several
subpopulations of McpR<sub>H99A</sub>.

The protocol used for the LC-MS analysis performed on McpR *in vitro* did not allow us to monitor the metal-catalyzed oxidation of the neighboring C75 residue as observed for the H99 residue. To assess the potential implication of the C75 residue in Cu ion coordination, we purified the McpR<sub>C75S</sub> mutant protein and calculated the *K*<sub>d</sub> values of the McpR<sub>C75S</sub>/Cu(I) ion and McpR<sub>C75S</sub>/Cu(II) ion complexes. As for the H99A mutation, the C75S mutation also impeded Cu(I) ion binding as attested by a 3.5-fold *K*<sub>d</sub> increase. Cu(II) ion binding to the McpR<sub>C75S</sub> mutant remained unchanged (Figs. 3B and S7B for the *fitting curves*), suggesting that McpR C75 is required to coordinate Cu(I) ions but not Cu(II) ions. Consistent with this observation, *in vitro* H99 oxidation by Cu(I) ions ±H<sub>2</sub>O<sub>2</sub> is fully abolished in the McpR<sub>C75S</sub> variant (Fig. 3A, *right panel*). The *K*<sub>d</sub> values calculated for the McpR<sub>C75S</sub> and McpR<sub>H99A</sub> variants are in agreement with the known selective binding of Cu(I) and Cu(II) ions to cysteine and histidine residues (30, 38).

To determine whether Cu coordination by McpR H99 and
C75 residues is essential to Cu chemotaxis, we decided to
express an ectopic *mcpR*<sub>H99A</sub> allele or *mcpR*<sub>C75S</sub> allele in the
Δ*mcpR* mutant and to monitor the chemotactic behavior of
both strains to Cu ions. None of the mutant allele was able to
complement the Cu-chemotaxis defect of the Δ*mcpR* mutant
(Fig. 3C). One could argue that the McpR<sub>H99A</sub> and McpR<sub>C75S</sub>
variants are unstable *in vivo*, explaining the Δ*mcpR* mutant
phenotype. In accordance with the pleiotropic sensing ability
of some Mcps, we discovered that McpR is also playing a key
role in Cd chemotaxis (Fig. S8). Interestingly, the *mcpR*<sub>H99A</sub>
and *mcpR*<sub>C75S</sub> mutant strains were not impeded in Cd
chemotaxis (Fig. S8). This latter observation attests the
integrity of the McpR<sub>H99A</sub> and McpR<sub>C75S</sub> variants *in vivo* and
suggests that Cd sensing by McpR does not rely on the H99
and C75 residues.

These data led us to the idea that Cu(I) ion coordination by
McpR H99 and C75 residues is decisive for McpR-mediated
Cu chemotaxis. Given the structural changes elicited by the
A

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E

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B

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D

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F

<figure><img src="image_8.png" /><figcaption>**Figure 2. Cu-induced ROS sustain Cu chemotaxis.** A, raw fluorescence of SW cells incubated with 5 µM CellROX Deep Red probe and exposed to 175 µM Cu or 400 µM H₂O₂. B, relative fluorescence intensity of SW cells expressing the rxYFP biosensor and exposed to 175 µM Cu or 400 µM H₂O₂ for 20 min. C, relative fluorescence intensity of SW cells expressing the rxYFP biosensor and exposed to various Cu concentrations for 20 min. D, relative fluorescence intensity of WT SW cells and SW cells overexpressing SodB (sodB+) and KatG (katG+) incubated with 5 µM CellROX Deep Red probe and exposed to 175 µM Cu. E, growth profiles at an absorbance at 660 nm of WT, sodB+, and katG+ strains in PYE (orange) and exposed to 175 µM Cu (blue). F, percentage of WT, ΔcheA1, sodB+, and katG+ SW cells in the vicinity of the Cu plug after 25 min. Mean ± SD, at least three biological replicates, except for 300 µM Cu (two biological replicates). p Values were calculated using an ANOVA when appropriate (*p* < 0.05, **p** < 0.01, and ***p** < 0.001) (Table S1). Cu, copper; PYE, peptone yeast extract; ROS, reactive oxygen species; SW, swarmer cell.</figcaption></figure>
Bacterial chemotaxis to copper and oxidative stress

<figure><img src="image_9.png" /><figcaption>**Figure 3.** Cu cycling modulates its coordination by McpR. A, percentage of oxidation of the H99 residue from purified McpR (left panel) and McpRC75S (right panel) incubated with Cu(II), Cu(I), H₂O₂, and Cu(I) + H₂O₂ at a 1:100 ratio. B, measure, by intrinsic fluorescence, of the K_d values of purified McpR, McpRH99A, and McpRC75S (1.79 µM) incubated with 0 to 500 µM Cu(I) and Cu(II). Mean ± SD. C, percentage of WT, Δ*mcpR*, and Δ*mcpR* overexpressing McpR, McpRH99A, or McpRC75S SW cells remaining in the vicinity of the Cu plug after 25 min. Mean ± SD, at least three biological replicates. *p* Values were calculated using an ANOVA (*p* < 0.05, **p** < 0.01, and ***p** < 0.001) (Table S1). D, CD spectra of purified McpR (8.95 µM) incubated with Cu(I) and Cu(II) at a 1:100 ratio. Cu, copper.</figcaption></figure>

binding of a ligand to a cognate chemoreceptor (39), we envisioned that Cu(I) ions binding to McpR could modulate its conformation. To test this idea, far-UV CD spectroscopy was used to monitor the impact of Cu(I) ions and Cu(II) ions on McpR secondary structure *in vitro*. Accordingly, Cu(I) ions has a more significant impact than Cu(II) ions on McpR secondary structure (Fig. 3D), increasing the α-helix percentage from 23% to 44%.

Based on our findings, we propose that Cu exposure triggers a dose-dependent Cu accumulation within the cytoplasm triggering a dose-dependent production of ROS that will sustain Cu chemotaxis. The increase in cytoplasmic Cu ion concentration will likely favor a rather labile Cu ion binding to the McpR sensor domain. Cu(I) ion coordination by McpR C75 and H99 residues triggers McpR conformational change. Interestingly, Cu(II) ions seem to preferentially bind McpR H99 residue and does not modify the apo form-like conformation. These observations suggest that a redox cycling of the McpR-bound Cu atom could modulate McpR conformation (Fig. 4).

Discussion

The role of ROS in Cu toxicity and Cu stress response remains largely unknown in bacteria. We sought to address this gap in the frame of negative chemotaxis to toxic Cu in the dimorphic bacterial model *C. crescentus*.

Our study identified the first Cu sensing chemoreceptor hereafter named McpR. McpR is devoid of any TM domains or

signal peptide and is therefore predicted to localize in the cytoplasm. Cytoplasmic chemoreceptors have been shown to monitor the cellular metabolic state (40), whereas inner membrane chemoreceptors usually sense the periplasmic environment, as a proxy for extracellular changes. The cytoplasmic Cu content varies in a dose-dependent manner when the SW cells are exposed to Cu, suggesting that the sensing of cytoplasmic Cu could provide a sensitive monitoring of subtle variations of Cu concentrations when the SW cell swims down a Cu gradient.

In a previous study, we showed that the maintenance of a high cellular Cu concentration is critical for Cu chemotaxis (12). Consistent with this observation, we provide evidence that McpR binds Cu with low affinity. Although aspecific Cu binding to noncognate residues cannot be ruled out, the significant difference in K_d values measured between the WT McpR and the McpR<sub>H99A</sub> and McpR<sub>C75S</sub> variants argues for a Cu binding to these residues. Considering that Cu binding to McpR is a prerequisite for Cu chemotaxis, one could propose that when the SW cell is sufficiently far from the Cu source, the resulting low cytoplasmic Cu content impedes Cu binding to McpR and in turn the chemotactic response.

We also demonstrate that the cellular Cu accumulation triggers a dose-dependent production of ROS. This dose dependency in ROS production argues that the increase of extracellular Cu ion content triggers an increase of the cytoplasmic Cu ion labile pool, which is likely involved in the labile Cu binding to McpR. ROS production can be prevented by the
Bacterial chemotaxis to copper and oxidative stress

<figure><img src="image_10.png" /><figcaption>**Figure 4.** Hypothetical model of Cu sensing by McpR. SW cell exposure to Cu increases the cytoplasmic Cu concentration, leading to the generation of ROS (left). In addition, high cytoplasmic Cu content favors Cu(I) coordination by the C75 and H99 residues within the cytoplasmic McpR, triggering a conformational change of the chemoreceptor. Cu(II) binding to H99 residue has no impact on McpR apo-like conformation, suggesting a redox-dependent modulation of McpR conformation. This process would be maintained along the extracellular Cu gradient until the SW cell reaches a Cu-poor environment (right). Under these conditions, the cytoplasmic Cu concentration together with the ROS level would return to their basal level, and the labile Cu binding to McpR would be hindered, bringing McpR to its apo form conformation. Cu, copper; ROS, reactive oxygen species; SW, swarmer cell.</figcaption></figure>

overexpression of SodB superoxide dismutase and KatG
catalase, suggesting that at least O₂⁻ and H₂O₂ are generated
under Cu stress. The contribution of the ROS derived from
cellular Cu ions in Cu toxicity is still largely debated (41, 42).
Here, we demonstrate that ROS derived from cellular Cu ions
not only impede cell growth to some extent but also play a key
role in Cu chemotaxis. The exact target of ROS in Cu
chemotaxis was not identified in this study, but one could
hypothesize that O₂⁻ is rapidly converted into H₂O₂. The
latter would feed a local Fenton-like reaction catalyzed by the
Cu(I) atom coordinated by McpR C75 and McpR H99 resi-
dues. This would lead to a local OH⁻ production and to the
oxidation of McpR-bound Cu(I) into McpR-bound Cu(II).
Owing to the reducing cytoplasmic environment, McpR-
bound Cu(II) could in turn be reduced by cytoplasmic elec-
tron donors such as GSH, NADH, and thioredoxin to restore
the Cu(I) ionic form. Thereby, the McpR-bound Cu ion would
undergo a redox cycling involving the Cu(I) and Cu(II) ionic
forms, which would be maintained as long as H₂O₂ is pro-
duced by cytoplasmic Cu ions. Cu-generated ROS would
therefore act as second messengers in a bacterial stress
response.

The modulation of the CheA–CheY two-component system
results from a conformational change of the chemoreceptor
occurring upon ligand binding (9). We demonstrated here that
McpR conformation is modulated by the ionic status of the Cu
atom bound to McpR. It should be emphasized that chemo-
receptors are usually organized in trimers of dimers and

assembled in large arrays in cells (43). Therefore, it is possible
that Cu coordination by the McpR C75 and H99 residues
within the McpR array implies two McpR monomers from one
or two distinct dimers. Nevertheless, McpR conformational
change likely regulates the downstream effectors such as
CheAI and the resulting chemotactic response.

The diversity of chemoreceptors produced by a single bac-
terial species varies from 5 to 60 chemoreceptors and is often
correlated to its lifestyle (40). Nevertheless, chemoreceptors
are not always specific to one single substrate and can also
perform indirect sensing *via* a substrate transporter (27). For
instance, the *E. coli* Tar chemoreceptor directly senses aspar-
tate and performs an indirect sensing of maltose through a
periplasmic maltose-binding protein (44).

The extrapolation of our findings to other motile bacterial
species based on sequence analyses is complicated by (i) the
large diversity of Cu coordination mechanisms and thereby the
low predictability of Cu-binding sites by *in silico* analyses
and (ii) the low conservation of the Mcp sensor domains.
Nevertheless, it is very likely that the use of ROS as second
messengers is widely conserved in bacterial response to Cu.

**Experimental procedures**

Bacterial strains, plasmids, and growth conditions

C. crescentus (NA1000 strain) was grown at 30 °C in peptone
yeast extract (PYE) medium (45) with 5 µg/ml kanamycin, 1 µg/
ml chloramphenicol, and/or an appropriate concentration of
## Bacterial chemotaxis to copper and oxidative stress

CuSO₄·5H₂O when required. Cultures in exponential phase were used for all experiments. Strains, plasmids, and primers used in this study are listed in Table S2, and the strategies for their construction are available upon request. SW cell isolation was achieved accordingly (46). Briefly, the bacteria were collected at an absorbance of 0.4 at 660 nm and centrifuged in Ludox LS colloidal silica gradient. After the centrifugation, SW cells are separated from the other cellular forms (ST and PD) based on a density difference. The SW cells were collected and washed several times with PBS prior to the experiments.

### Growth curve measurements

*C. crescentus* exponential phase cultures were diluted in PYE medium to a final absorbance of 0.05 at 660 nm and inoculated in 96-well plates with appropriate CuSO₄ concentration when required. Bacteria were then grown for 24 h at 30 °C under continuous shaking in an Epoch 2 Microplate Spectrophotometer from BioTek. The absorbance at 660 nm was measured every 10 min.

### Motility assay

*C. crescentus* cultures grown in PYE were diluted to an absorbance of 0.4 at 660 nm and inoculated with sterile toothpicks on a PYE swarming plate (PYE + 0.25% agar). The plates were incubated at 30 °C for 3 days and then imaged.

### Determination of cytoplasmic Cu content

*C. crescentus* cells were fixed for 20 min in 2% paraformaldehyde at 4 °C and then washed three times with an ice-cold wash buffer (10 mM Tris-HCl [pH 6.8] and 100 µM EDTA). The cells were then incubated for 10 min with zwitterionic detergent (0.0125% Zwittergent 3-14 detergent and 200 mM Tris-HCl [pH 7.6]) to disrupt the outer membrane but not the inner membrane. A final centrifugation at 14,500g for 15 min separated the spheroplasts (cytoplasm and inner membrane fraction) from the periplasm (and outer membrane fraction). The spheroplasts were lysed under 2.4 kbar by using a cell disrupter (Cell Disruption System, one shot model, Constant). Cell debris were removed by centrifugation at 10,000g for 10 min, and cell lysates were diluted in 1% HNO₃. Samples were finally analyzed by ICP-OES with an Optima 8000 ICP-OES from PerkinElmer. Cellular Cu concentrations were calculated as described (12). Briefly, 10 µl of the samples after fixation were diluted 1:100 in PBS and counted with a BD FACSVerse flow cytometer. Data were analyzed with the BD FACSuite V1.0.5 software. A ratio between the number of bacteria and the absorbance at 660 nm was determined. The combination of both flow cytometry and ICP-OES analyses allows to determine the number of Cu atoms in one spheroplast. Cu concentration was calculated using the following formula;

$$ \frac{\text{Amount of Cu (mg)}}{\text{Molecular weight of Cu (63,546 mg/L)}} \times \text{Avogadro constant} (6.022 \times 10^{23}) $$

Number of bacteria

### Oxidative stress measurement with the CellROX Deep Red probe

An exponential phase culture of *C. crescentus* was synchronized to isolate the SW cells. The isolated SW cells were diluted to a final absorbance of 0.2 at 660 nm and incubated for 30 min at 30 °C under moderate shaking with 5 µM of CellROX Deep Red probe and appropriate concentrations of CuSO₄ or H₂O₂. The bacteria were then washed three times with 20 mM phosphate buffer (12.5 mM Na₂HPO₄, 8 mM KH₂PO₄, pH 7) to remove the non-internalized probe. The bacteria were finally transferred into a black bottom 96-well plate, and the fluorescence was monitored at emission/excitation of 640/690 nm with a SpectraMax iD3 (Molecular Devices). The relative fluorescence intensity was determined by defining 100% as the fluorescence from control condition.

### Oxidative stress measurement with the rxYFP biosensor

The SW cells were synchronized from an exponential phase culture overexpressing the rxYFP biosensor (47). The SW cells were diluted to a final absorbance of 0.4 at 660 nm in a black bottom 96-well plate and mixed with different concentrations of CuSO₄ and H₂O₂. The rxYFP fluorescence was then monitored every minute for 20 min at emission/excitation of 510/556 nm with a SpectraMax iD3. The relative fluorescence intensity was determined by defining 100% as the initial measured fluorescence at time 0.

### LCI

The LCI assay was adapted from Ref. (12). Briefly, chemotaxis devices were made by casting solubilized 10:1 polydimethylsiloxane (PDMS Slygard 184; Dow Corning) in a small glass pot (d = 50 mm; h = 30 mm; glassware from Lenz laborglass instrument) in which coverslips were initially installed to mold the future bacterial chamber. The PDMS was left overnight, allowing degassing and curing. The different channels were drilled into the PDMS devices. A microscope slide and the PDMS cube were washed successively with acetone (only the slide), isopropanol, methanol, and rinsed with Milli-Q H₂O. The materials were blown dry between each wash. The PDMS cube was sealed to the slide by pressure. Eight microliters of melted 1.5% agarose H₂O with appropriate concentration of CuSO₄, ZnSO₄, or CdSO₄ were loaded into the chamber through the external inlet channels to generate the plug. About 150 µl of isolated SW cells were in turn injected into the bacterial chamber through the central inlet channel. Images were collected every minute for 25 min in one focal plan (near the liquid surface) in the vicinity of the plug with a Nikon Eclipse Ti2-E inverted microscope equipped with a Nikon 20x/0.5 Ph2 objective and with a Hamamatsu C13440 digital camera. Quantitative analysis of the time-lapse images was performed with MicrobeJ, an ImageJ plug-in (48). The number of bacteria in a same focal plane was determined at each time point; the number of bacteria at the beginning of the experiment is considered as 100%.
## Protein expression and purification

A BL21 pET28a-McpR<sub>His</sub> exponential phase culture was incubated for 3 h under shaking at 37 °C with 1 mM IPTG and then harvested by centrifugation for 30 min at 4000g at 4 °C. McpR<sub>His</sub> induction was verified by 12% tricine-SDS-PAGE, followed by Coomassie Blue staining and by Western blot using an anti-His antibody (MBL; PM032). To purify McpR<sub>His</sub>, the IPTG-induced culture was washed with binding buffer (20 mM Tris-HCl (pH 8.0), 500 mM NaCl, 10% glycerol, and 10 mM MgCl<sub>2</sub>) supplemented with 400 mg lysozyme (Sigma), complete EDTA-free protease cocktail inhibitor (Roche), 5 mg DNase I (Roche), and 0.4 g SDS and centrifuged for 15 min at 9000g at 4 °C. Bacterial cells were lysed at 2.4 kbar using a Cell disruptor (Cell Disruption System—one shot model, Constant). As McpR<sub>His</sub> was mainly forming inclusion bodies, the pellet fraction was resuspended for 1 h under shaking at 4 °C in the binding buffer containing 6 M guanidine. The solubilized inclusion bodies were centrifuged for 30 min at 9000g at 4 °C. The collected supernatant was dialyzed overnight in a 0.1 M phosphate buffer (12.5 mM Na<sub>2</sub>HPO<sub>4</sub>, 8 mM KH<sub>2</sub>PO<sub>4</sub>, pH 8) to remove guanidine, allowing the refolding of McpR. The same methodology was applied for the C75S and H99A mutants.

McpR protein concentration was determined (i) by measuring the absorbance of the sample at 280 nm with the SpectraMax iD3 or (ii) with a Bradford protein assay.

## Fluorescence quenching by Cu and other metal ions

Purified McpR at a concentration of 3.58 µM was incubated with different ratios of CuSO<sub>4</sub>, Tetrakis(acetonitrile)Cu(I) hexafluorophosphate, ZnSO<sub>4</sub>, CdSO<sub>4</sub>, MnSO<sub>4</sub>, and NiSO<sub>4</sub> in black bottom 96-well plates. The fluorescence emission spectra were recorded upon 280 nm excitation with the SpectraMax iD3.

## K<sub>d</sub> determination

The titration of McpR by Cu(I) or Cu(II) monitored by the protein intrinsic fluorescence intensity quenching was fitted to a single-site hyperbolic model by nonlinear regression using the SciDAVis 2.7 software (https://scidavis.sourceforge.net/index.html) to determine the affinity constants, according to the following equation:

Fluorescence intensity = F0 + A*[Cu]/(K<sub>d</sub> + [Cu]), with F0 corresponding to the fluorescence intensity of the apo McpR, A to the total fluorescence amplitude change upon Cu binding and K<sub>d</sub> to the affinity dissociation constant. Dissociation constants are reported as the mean value ± SD obtained for a minimum of two independent experiments performed on distinct protein productions.

## Identification of McpR oxidation by mass spectrometry

### Sample preparation and protein digestion

About 15 µg of purified McpR or McpR<sub>C75S</sub> were incubated with Cu(II), Cu(I), H<sub>2</sub>O<sub>2</sub>, and Cu(I) + H<sub>2</sub>O<sub>2</sub> at a 1:100 ratio for 10 min at room temperature. McpR samples were treated

using filter-aided sample preparation using the following protocol. To first wash the filter, 100 µl of 1% formic acid were placed in each Millipore Microcon 30 MRCFOR030 Ultracel PL-30 before a 15 min centrifugation at 14,100g. For each sample, 40 µg of protein adjusted in 150 µl of 8 M urea buffer (8 M urea in 0.1 M Tris buffer at pH 8.5) were placed individually in a column and centrifuged for 15 min at 14,100g. The filtrate was discarded, and the columns were washed three times by adding 200 µl of urea buffer followed by a 15 min centrifugation at 14,100g. For the reduction step, 100 µl of DTT were added and mixed for 1 min at 400 rpm with a thermomixer before a 15 min incubation at 24 °C. The samples were then centrifugated for 15 min at 14,100g, the filtrate was discarded, and the filter was washed by adding 100 µl of urea buffer before another 15 min centrifugation at 14,100g. An alkylation step was performed by adding 100 µl of iodoacetamide ([IAA], in urea buffer) in the column and mixing for 1 min at 400 rpm in the dark followed by a 20 min incubation in the dark and a 10 min centrifugation at 14,100g. To remove the excess of IAA, 100 µl of urea buffer were added and the samples were centrifuged for 15 min at 14,100g. To quench the rest of IAA, 100 µl of DTT were placed on the column, mixed for 1 min at 400 rpm, and incubated for 15 min at 24 °C before a 10 min centrifugation at 14,100g. To remove the excess of DTT, 100 µl of urea buffer were placed on the column and centrifuged for 15 min at 14,100g. The filtrate was discarded, and the column was washed three times by adding 100 µl of 50 mM sodium bicarbonate buffer ([ABC], in ultrapure water) followed by a 10 min centrifugation at 14,100g. The remaining 100 µl were kept at the bottom of the column to avoid any evaporation in the column. The digestion process was performed by adding 80 µl of mass spectrometry (MS) grade trypsin (1/50 in ABC buffer) in the column and mixed for 1 min at 400 rpm followed by an overnight incubation at 24 °C in a water saturated environment. The Microcon columns were placed in a 1.5 ml LoBind tube and centrifuged for 10 min at 14,100g. Forty microliters of ABC buffer were placed on the column before a 10 min centrifugation at 14,100g. Ten percent of TFA in ultrapure water were added into the LoBind Tube to obtain 0.2% TFA. The samples were dried in a SpeedVac up to 20 µl and transferred into an injection vial.

### MS

The digest was analyzed using nano-LC-ESI-MS/MS time-of-flight (TOF) Pro (Bruker) coupled with an UHPLC nanoElute (Bruker).

Peptides were separated by nanoUHPLC (nanoElute; Bruker) on a 75 µm ID, 25 cm C18 column with integrated CaptiveSpray insert (Aurora, IonOpticks) at a flow rate of 400 nl/min, at 50 °C. LC mobile phase A was water with 0.1% formic acid (v/v) and B acetonitrile with 0.1% formic acid (v/v). Samples were loaded directly on the analytical column at a constant pressure of 800 bar. The digest (1 µl) was injected, and the organic content of the mobile phase was increased linearly from 2% B to 15% in 36 min, from 15% B to 25% in 19 min, from 25% B to 37% in 5 min, and from 37% B to 95% in 5 min. Data acquisition on the time-of-flight Pro was performed
## Bacterial chemotaxis to copper and oxidative stress

using Hystar 5.1 and timsControl 2.0. timsTOF Pro (Bruker) data were acquired using 100 ms TIMS accumulation time, mobility (1/K0) range from 0.6 to 1.6 Vs/cm². Mass spectrometric analysis was carried out using the parallel accumulation serial fragmentation acquisition method. One MS spectra followed by ten parallel accumulation serial fragmentation MSMS spectra per total cycle of 1.1 s.

Data analysis was performed using PEAKS Studio X Pro with ion mobility module (Bioinformatics Solutions, Inc). Protein identifications were conducted using PEAKS search engine with 15 ppm as parent mass error tolerance and 0.05 Da as fragment mass error tolerance. Carbamidomethylation was allowed as fixed modification, oxidation of methionine, histidine, tryptophan, and acetylation (N-term) as variable modification. Enzyme specificity was set to trypsin, and the maximum number of missed cleavages per peptide was set at 3. The peak lists were searched against the *E. coli*-K12 (11,002 sequences) and *Caulobacter* NA1000 (3863 sequences) from UNIREF 100. Peptide spectrum matches and protein identifications were normalized to less than 1.0% false discovery rate.

The identification results were processed with the PTM profile tool from PEAKS, which provides quantitative information of modified peptides compared with unmodified peptides for the modification sites of the protein across all MS samples (49, 50).

The MS proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE (51) partner repository with the dataset identifier PXD044512 and 10.6019/PXD044512.

## CD

Purified McpR at a concentration of 8.95 µM was incubated with different CuSO₄ and *Tetrakis(acetonitrile)Cu(I) hexa-fluorophosphate* concentrations. Measurements were performed with an MOS-500 spectropolarimeter in the far-UV region (190–260 nm), using a 1 mm pathlength cell. Four scans were averaged, and base lines were subtracted. Secondary structure analyses were performed on the CD data with the BeStSel server.

## Data availability

All data are contained within the article and supporting information section. The raw data can be shared upon request to jean-yves.matroule@unamur.be.

**Supporting information**—This article contains supporting information (16, 17, 52–54).

**Acknowledgments**—We thank Jakob R. Winther for providing the rxYFP construct. We are also grateful to Valérie Charles and Carmela Aprile (CMI laboratory, NISM, UNamur) for the ICP measurements and to Virgile Neyman for technical support during the protein purification. Finally, we acknowledge Jean-François Collet and Xavier De Bolle for critical reading of this article and the URBM members and Johan Wouters (Chemistry Department, UNamur) for fruitful discussions.

**Author contributions**—G. L. and J.-Y. M. conceptualization; G. L. methodology; G. L. validation; G. L., P. C., C. M., S. R.-C., M. D., R. V. H., P. R., E. P., and J.-Y. M. formal analysis; G. L., P. C., C. M., M. D., F. T., L. M., and E. P. investigation; G. L. writing—original draft; G. L. visualization; J.-Y. M. writing—review & editing; J.-Y. M. supervision; J.-Y. M. funding acquisition.

**Funding and additional information**—This work was supported by the University of Namur. G. L. was supported by the Belgian Fund for Industrial and Agricultural Research Associate (FRIA/FNRS). Catherine Michaux and Eric Perpète thank the Belgian National Fund for Scientific Research (FNRS) for their respective associate and senior research associate positions.

**Conflict of interest**—The authors declare that they have no conflicts of interest with the contents of this article.

**Abbreviations**—The abbreviations used are: ABC, sodium bicarbonate buffer; Cd, cadmium; Cu, copper; H₂O₂, hydrogen peroxide; IAA, iodoacetamide; ICP-OES, inductively coupled plasma optical emission spectrometry; LCI, live chemotaxis imaging; MCP, methyl-accepting chemotaxis protein; MS, mass spectrometry; PDMS, polydimethylsiloxane; PYE, peptone yeast extract; ROS, reactive oxygen species; SW, swarmer cell; TM, transmembrane domain; Zn, zinc.

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