# Interplay of acidic residues in the proton channel of E. coli cytochrome bd-I oxidase to promote oxygen reduction and NO release

Raaif Siddeeque, Lucia Heger, Jan Kägi, Thorsten Friedrich, Frédéric Melin, Petra Hellwig

<figure><img src="image_1.png" /></figure>

► To cite this version:

Raaif Siddeeque, Lucia Heger, Jan Kägi, Thorsten Friedrich, Frédéric Melin, et al. Interplay of acidic residues in the proton channel of E. coli cytochrome bd-I oxidase to promote oxygen reduction and NO release. Biochimica biophysica acta (BBA) - Bioenergetics, 2025, 1866 (2), pp.149537. 10.1016/j.bbabio.2025.149537. hal-05081757

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Submitted on 23 May 2025

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# Interplay of acidic residues in the proton channel of *E. coli* cytochrome *bd*-I oxidase to promote oxygen reduction and NO release

Raaif Siddeeque<sup>a</sup>, Lucia Heger<sup>b</sup>, Jan Kägi<sup>b</sup>, Thorsten Friedrich<sup>b,*</sup>, Frédéric Melin<sup>a</sup>,
Petra Hellwig<sup>a,c,**</sup>

<figure><img src="image_3.png" /></figure>

ARTICLE INFO

Keywords:

Cytochrome *bd*

Oxygen reaction

NO

Proton transfer

Escherichia coli

Electrocatalysis

A B S T R A C T

The reduction of oxygen to water is crucial to life under aerobic conditions. Cytochrome *bd* oxidases perform this reaction with a very high oxygen affinity. Members of this protein family are solely found in prokaryotes and some archaea playing an important role in bacterial virulence and antibiotic resistance. Here, we combine mutagenesis, electrocatalysis, nitric oxide binding and release experiments as well as FTIR spectroscopy to demonstrate that proton delivery to the active site is essentially rate limiting in Cyt *bd*-I electrocatalysis. D58 and D105 of subunit CydB are crucial residues in this proton path and communicate via a hydrogen bond network. Oxygen reduction depends on proton delivery to the active site, which also influences NO release.

1. Introduction

Bacterial antibiotic resistance is an ever-growing global problem,
and it is expected that microbial infections will cause more deaths than
cancer within the next decades. When pathogenic bacteria colonize host
cells, they are exposed to unfavorable environmental factors such as
reactive oxygen and nitrogen species. To survive under these hostile
conditions, pathogens increase the production of cytochrome *bd* oxidase
(Cyt *bd*). [1] Members of this enzyme family are solely found in pro-
karyotes and some archaea making them excellent drug targets. [2,3] A
pre-requisite to finding selective and potent inhibitors of *bd* oxidases is
in-depth knowledge of their structure and function. Cyt *bd* is part of
aerobic respiratory chains reducing molecular oxygen to water at the
expense of the oxidation of hydrophobic quinols. More than one Cyt *bd*
oxidase can be found in several bacteria. [4] *Escherichia coli* contains two
homologous Cyt *bd* oxidases named *bd*-I and *bd*-II. Cyt *bd*-I is made up of
two large subunits CydA and B, and two small subunits X and H/Y. [5,6]
The redox cofactors heme *d*, heme *b*<sub>558</sub> and heme *b*<sub>595</sub> are located in
CydA, which also contains a hydrophilic loop, the Q-loop, contributing
to quinone binding [7–10] and stabilization of the entire enzyme

complex. [11] The heme groups are arranged in a triangle with hemes
b<sub>558</sub> and b<sub>595</sub> expected to transfer electrons from quinol to the heme
d active site. O<sub>2</sub> is reduced at the high spin d-heme that is also the
binding site for several cellular stressors including CO, H<sub>2</sub>S and NO.
[1,12–17] CydB adopts the same fold as CydA, however, it does not
carry any cofactor. Instead, the positions taken by the heme groups in
CydA are occupied by a quinone in CydB. [5,6] Remarkably, dioxygen
has no direct access to the active site but is guided by a long substrate
channel leading from CydB to heme d on CydA. The small subunits take
over different tasks. While CydX has a structural role in stabilizing the
active site, [18,19] CydH/Y shields access to heme b<sub>595</sub> from the
membrane. [3,6]

The mechanism of Cyt *bd*-I has been studied in detail and several
catalytic intermediates were described. [2,20] Since the reaction in-
volves the de-protonation of a quinol and a proton transfer to dioxygen,
it is not surprising that several conserved acidic amino acid residues play
an important role in the mechanism. For example, turnover is blocked
when heme *b*<sub>595</sub> is locked in its ferric form by mutation of E445<sup>A</sup> (the
superscript refers to the name of the *E. coli* subunit). [21,22] Further-
more, the redox potentials of hemes *d* and *b*<sub>595</sub> are pH dependent. [23]

<footnote><sup>a</sup> Laboratoire de Bioélectrochimie et Spectroscopie, UMR 7140, Chimie de la Matière Complexe, Université de Strasbourg-CNRS 4, Rue Blaise Pascal, 67081 Strasbourg, France</footnote>

<footnote><sup>b</sup> Institut für Biochemie, Albert-Ludwigs-Universität Freiburg, Albertstr 21, 79104 Freiburg, Germany</footnote>

<footnote><sup>c</sup> Institut universitaire de France (IUF), France</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>** Correspondence to: P. Hellwig, Laboratoire de Bioélectrochimie et Spectroscopie, UMR 7140, Chimie de la Matière Complexe, Université de Strasbourg-CNRS 4, Rue Blaise Pascal, 67081 Strasbourg, France.</footnote>

<footnote>E-mail addresses: friedrich@bio.chemie.uni-freiburg.de (T. Friedrich), hellwig@unistra.fr (P. Hellwig).</footnote>
A putative pathway was proposed for proton transfer from the cytoplasm to heme d. This pathway is located at the interface of CydA and CydB and comprises the strictly conserved residues E107A and E99A of CydA and D58B and D105B of CydB (Fig. 1). [24–27] Mutations at these positions strongly influence the kinetics of bd-I oxidase. [28]. Here, we probe the role of these residues for oxygen reduction by means of electrocatalytic studies. FTIR difference spectra give evidence on the protonation state of these residues. Furthermore, the NO release monitored by UV–vis spectroscopy points to the important role of the proton pathway for oxygen binding.

## 2. Materials and methods

### 2.1. Generation of mutants, protein production and purification

The CydB variants were constructed as described. [6] The mutagenic primers are listed in Supplementary Materials (Table S1). Generation of the D58E<sup>B</sup> and D58L<sup>B</sup> mutants is described in [26], all other variants were newly generated for this study. Cytochrome bd-I oxidase and its variants were prepared from *E. coli* strain CBO/pET28b(+):cydA<sub>H</sub>BX (with ‘h’ describing the position of the His-tag at the C-terminus of CydA) as previously described. [6] The protein was stored at a final concentration of 40–50 mg·mL<sup>-1</sup> in 20 mM MOPS, 20 mM NaCl, 30 µM PMSF, 0.003 % Lauryl Maltose Neopentyl Glycol (LMNG), pH 7.0.

### 2.2. Q₁H₂:dioxygen oxidoreductase activity

The oxidase activity was determined as reported previously [26], however, using a different substrate. In short, the Q₁H₂:dioxygen oxidoreductase activity of isolated *E. coli* Cyt bd-I and the variants (F63P<sup>A</sup>, W57A<sup>B</sup>, D58N<sup>B</sup>, D58E<sup>B</sup>, D58L<sup>B</sup>, D105A<sup>B</sup>, D105N<sup>B</sup>, and D105E<sup>B</sup>) was determined by monitoring the O₂ consumption at 30 °C with a Clark-type oxygen electrode (Oxyview System, Hansatech Instruments). The reaction chamber was filled with 2 mL buffer (20 mM MOPS, 20 mM

NaCl, 30 µM PMSF, 0.003 % LMNG, pH 6.0–9.0), 10 µL DTT (1 M) and 5 µL Q₁H₂ (reduced 1,4-benzoquinone with one isoprenoid moiety) in ethanol (100 mM) and incubated for 1 min. 10 µL protein in LMNG buffer (2 mg·mL<sup>-1</sup>) at a final concentration of 95.25 nM protein was added to the buffer to start the reaction. The rates were corrected for the non-enzymatic reaction. The pH value of the buffers was checked at the experiment's beginning and end.

### 2.3. UV-visible spectroscopy

To validate the presence of heme d, excess detergent of the preparations of Cyt bd-I and its variants were removed by ultra-filtration in Amicon Ultra centrifugal 50 kDa filters. The preparations were reduced by an excess of sodium dithionite in 50 mM KPi buffer pH 7, 0.05 % DDM and UV–vis spectra were recorded with a dual-beam diode array VARIAN Cary 300 spectrometer with a glass cuvette of 1 cm path length.

For the kinetic measurement of NO binding and release, UV–vis spectra of the reduced protein in 100 mM KPi, 100 mM KCl, 0.05 % DDM buffer at pH 6, 7 and 8, were recorded every 30 s for an hour (until a stable spectrum was achieved) after a single addition of a freshly prepared 20 mM MAHMA NONOate (from Sigma Aldrich) solution in 0.01 M NaOH, respectively. MAHMA NONOate releases two equivalents of NO, the ratio of the protein and nitric oxide in the cuvette were 5 µM and 500 µM (1:100), respectively. The pH of the reaction mixture was tested at the end of each experiment.

### 2.4. Protein film voltammetry for oxygen catalysis

The electrochemical characterization of immobilized protein was adapted from a previously reported method. [29,30] Gold nanoparticles of 15 nm average diameter were obtained by the procedure of Turkevich et al. and Frens. [31] In brief, a sodium citrate solution (39 mM, 13 mL) was added to a boiling solution of HAuCl₄ in water (1 mM, 125 mL). After the appearance of the red colour, the solution was maintained

<figure><img src="image_4.png" /><figcaption>**Fig. 1.** Structure of *E. coli* cytochrome bd-I oxidase (PDB code: 6RX4). All hemes are located in CydA (cyan). Oxygen is reduced at heme d (violet). The upper right inset shows the residues of CydB proposed to be involved in the proton pathway including residues D58B, W57B and D105B. Lower right inset depicts residues of CydA possibly comprised in the proton pathway including residues F63A, E107A and E99A. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>
under boiling for 15 min and then cooled to room temperature. The average size of the NPs was estimated from the absorbance ratio at 521 and 450 nm.

The colloidal gold solution was concentrated by centrifugation for 30 min at 13000 rpm, and removal of 99 % of the supernatant. Concentrated gold colloid solution (5 µL) were deposited three times on the cleaned gold electrode surface one after the other and were allowed to dry under air after each step. For cleaning, the polycrystalline surface of the gold electrode (RDE) was mechanically polished with 0.05 µm alumina paste and activated by subsequent oxidation at +2.0 V for 5 s, reduction at -0.35 V for 10 s, and then cycling 100 times between -0.35 V and 1.5 V at 4 V s⁻¹ in 0.1 M H₂SO₄. The modified nano-structured gold electrode was cycled between -0.35 V and 1.5 V at 0.1 V s⁻¹ until a stable voltammogram was obtained.

Before protein deposition, the modified gold electrode was immersed in a solution containing a 1:1 mixture of hexanethiol (0.5 mM) and 6-mercaptohexanol (0.5 mM), respectively. The monolayer was allowed to self-assemble for 1 h. The modified electrode was rinsed with ethanol, dried under an argon stream, and was cycled again between -0.6 V and 0.3 V first at 0.1 V s⁻¹ and then at 0.02 V s⁻¹ in 50 mM phosphate buffer. Finally, protein solution (10 µM, 5 µL with 0.025 eq of phosphatidyl-ethanolamine (PE), previously washed from the excess of detergent in Amicon Ultra centrifugal 50 kDa filters was deposited on the modified electrode surface and incubated in the refrigerator at 4 °C for 4 h.

All voltammetric measurements were performed in a standard three electrode cell connected to a Princeton Applied Research VERSASTAT 4 potentiostat. A silver/silver chloride in 3 M KCl electrode was used as reference, a platinum wire as counter electrode, and the modified gold disk electrode as working electrode. The working electrode was rotated at 1000 rpm.

## 2.5. Redox induced FTIR difference spectroscopy

The experiments were performed in the previously described thin-layer transmission electrochemical cell. [21,32] A small sheet of platinum contact and an aqueous Ag/AgCl 3 M KCl were used as counter and reference electrodes, respectively. A semi-transparent gold grid that served as a working electrode was modified with a 1:1 cysteamine and mercaptopropionic acid solution. Cyt *bd*-I and its variants were prepared in 100 mM KPi, 100 mM KCl, 0.025 % DDM, pH 7 at 0.6 mM concentration. Before each experiment, protein solutions were incubated with a cocktail of 19 mediators at a final concentration of 25 µM. [33] The cell was assembled as described previously. [21,32] FTIR spectra were obtained from fully reduced samples at -0.5 V and fully oxidized ones at +0.5 V vs Ag/AgCl using a Vertex 70 FTIR spectrometer from Bruker Optics, with an equilibration time of 3 min for each step. The redox reaction was cycled over 50–80 times and for each redox state two spectra (256 scans, 4 cm⁻¹ resolution) were averaged in each cycle.

# 3. Results and discussion

## 3.1. Characterization of the variants

Mutations have been introduced in potentially crucial positions along the proton pathway leading from the cytoplasm to heme *d*. Table 1 summarizes the specific Q₁H₂:dioxygen oxidoreductase activity of the preparations at pH 6, 7 and 8. Mutation of positions D58B and D105B clearly diminish the specific activity at the various pH (Table 1). As the only exception, the D105E variant exhibits the same activity as Cyt *bd*-I within the statistical variance at pH 7. The D58L and D58N replacements led to a strong reduction in activity, while the D58E substitution only slightly hampered activity (Table 1). This is in agreement with our earlier findings [26], although the absolute activity values differ due to the use of Q₁H₂ as substrate instead of duroquinol. Remarkably, the W57A variant features a higher activity than Cyt *bd*-I at pH 6 and 7 (Table 1).

<table><caption>**Table 1**
Q₁H₂:dioxygen oxidoreductase activity of cytochrome *bd*-I oxidase and its CydB variants at various pH values. All measurements were done in triplicates.</caption><thead><tr><th rowspan="2">Sample</th><th colspan="3">Specific activity (µmol/min*mg)</th></tr><tr><th>pH 6</th><th>pH 7</th><th>pH 8</th></tr></thead><tbody><tr><td>WT</td><td>47 (±10)</td><td>79 (±20)</td><td>103 (±1)</td></tr><tr><td>D58N</td><td>13 (±0.5)</td><td>14 (±0.4)</td><td>13 (±0.3)</td></tr><tr><td>D58E</td><td>28 (±3)</td><td>56 (±20)</td><td>79 (±3)</td></tr><tr><td>D58L</td><td>7 (±0.5)</td><td>9 (±0.2)</td><td>9 (±0.2)</td></tr><tr><td>D105N</td><td>8 (±0.6)</td><td>19 (±1)</td><td>8 (±1)</td></tr><tr><td>D105A</td><td>11 (±1)</td><td>5 (±0.1)</td><td>3 (±0.4)</td></tr><tr><td>D105E</td><td>28 (±13)</td><td>84 (±6)</td><td>97 (±2)</td></tr><tr><td>W57A</td><td>79 (±19)</td><td>102 (±10)</td><td>96 (±3)</td></tr></tbody></table>

The heme content of the variants was checked by UV-vis spectroscopy, with the heme cofactors showing distinct bands (Fig. 2). The W57A<sup>B</sup>, D58E<sup>B</sup>, D105A<sup>B</sup>, D105E<sup>B</sup> and D105N<sup>B</sup> variants exhibited a similar heme *d* / heme *b* ratio as Cyt *bd*-I. For the D58N<sup>B</sup> and D58L<sup>B</sup> variants, the amount of heme *d* was decreased to about one half and one third, respectively, whereas the F63P<sup>A</sup> mutant contained almost no heme *d*.

## 3.2. Protein film voltammetry

Protein film voltammetry provides insights into the sequence of catalytic events including electron transfer and coupled proton transfer. The electrical current is monitored, which is proportional to the protein coverage of the electrode and turnover rate, as a function of electrode potential and other experimental parameters including pH. [34–36] The onset potential, which refers to the potential where the catalytic reaction starts, provides information on the kinetic overpotential required to drive the reaction. [37] The variants in focus of this study were immobilized as described previously. [29,30,38] The voltammograms recorded under aerobic conditions at pH 6, 7 and 8 are shown in Fig. 3. Fig. S1 shows the voltammograms of the individual samples at these pH values plus those at pH 9.

As described before, [38,39] a sigmoidal curve with an onset potential at about -0.13 V and E₁/₂ at -0.38 V (vs Ag/AgCl, pH 7, 23 °C) is obtained for Cyt *bd*-I (red curves in Fig. 3) typical for the catalytic reduction of oxygen by the immobilized enzyme.

The catalytic curves are strongly altered for the D58N<sup>B</sup> and D58L<sup>B</sup> variants. In contrast, the D58E<sup>B</sup> variant with a conservative substitution retained most of the electrocatalytic activity (Fig. 3a,d,g). The lower activity of the D58<sup>B</sup> variants is consistent with their specific activities in solution (Table 1) and it is not only caused by a decreased heme *d* content (Fig. 2). This supports the assumption that an acidic residue in position 58 of subunit CydB is crucial for the reactivity, [26] probably acting as a key residue in proton transfer. Based on flow-flash kinetic studies, D58<sup>B</sup> was proposed to be the high pKa proton donor involved in the peroxy to ferryl transition during the reaction between the fully reduced enzyme and oxygen. [28]

Unexpectedly, all D105<sup>B</sup> variants (Fig. 3b,e,h) showed a high electrocatalytic activity in stark contrast to the much lower specific activity of the D105N<sup>B</sup> and D105A<sup>B</sup> variants in solution. The reason for this discrepancy could be mainly due to differences in the methodology. In the activity measurements in solution, electrons are provided by the redox partner Q₁H₂ to the enzyme at a specific site, namely the Q-binding site. In protein film voltammetry the electrons flow from the electrode to the enzyme with a driving force that can be tuned by scanning the potential towards more negative values. In addition, they may enter the protein at various sites and thus bypass the Q-binding site.

Interestingly, the D105E<sup>B</sup> variant exhibited a higher catalytic onset potential and a higher reduction current during electrocatalysis. It can thus be concluded that D105<sup>B</sup> participates in proton transfer proton to heme *d*, as proposed from flow-flash kinetic studies. [28] However, it is not put in place to guarantee fast proton delivery but rather to tune the
<figure><img src="image_5.png" /><figcaption>Fig. 2. UV-vis spectra of reduced Cyt *bd*-I and variants at pH 7. The spectra were normalized at 561 nm.</figcaption></figure>

<figure><img src="image_6.png" /><figcaption>Fig. 3. The pH-dependent oxygen catalysis of Cyt *bd*-I and its variants at pH 6, 7, and 8; scan rate 0.02 V s⁻¹ and rotation speed 1000 rpm.</figcaption></figure>

reaction. This is in line with the fact that its replacement by uncharged residues does not completely block the access of protons to the active site.

The W57A<sup>B</sup> variant also showed a higher catalytic onset potential and a higher catalytic current than Cyt *bd*-I at all pH values (Fig. 3). This is consistent with the higher activity of the variant in solution at pH 6 and 7, while at pH 8 it is in the same range as that of Cyt *bd*-I (Table 1). On one hand, the replacement of the bulky tryptophan by a small alanine residue improves the access of water and thus the transfer of protons to the active site. This assumption is supported by the much higher activity of the variant compared to Cyt *bd*-I at acidic conditions, while no significant differences in activity are observed at more alkaline conditions (Table 1). On the other hand, it is also discussed that W57B<sup>B</sup>

stabilizes the anionic form of D58<sup>B</sup> by a π-anion interaction. [26] The lack of the π-electron system in the variant might stabilize the protonated form of the carboxylic group leading to a slower proton transfer via this position. The voltammogram of the F63P<sup>A</sup> variant demonstrates the nearly complete loss of electrocatalytic activity with the oxidase lacking most of heme *d* (Fig. S1).

The electrocatalytic curve of Cyt *bd*-I shifts by approximately -60 mV per unit of pH (Fig. S2), which is the expected pH dependence for proton-coupled electron transfer involving an equal number of protons and electrons. The W57A<sup>B</sup> and D105<sup>B</sup> variants exhibit the same shift of the catalytic curves (Fig. S2). The maximum currents of *bd*-I oxidase as well as the W57A<sup>B</sup> and D105E<sup>B</sup> variants are similar and a high level of electrocatalytic activity is observed at pH 6 and 8. This activity is
diminished at pH 9, whereas that of the D105A<sup>B</sup> and D105N<sup>B</sup> variants
exhibits a rather gradual decrease in the investigated pH range. Unex-
pectedly, a significant shift of the signals of the D58N<sup>B</sup> and D58E<sup>B</sup> vari-
ants towards more negative potentials is observed when changing from
pH 6 to 7, while the influence was less pronounced at more alkaline pH.
These variants also exhibited similar and low catalytic currents at the
tested pHS (Fig. S2). This specific pH-dependency likewise points to the
key role of the D58<sup>B</sup> in proton transfer to the active site.

3.3. NO binding and release experiments

It is known that NO binds to *heme d* of Cyt *bd*-I leading to an inhi-
bition of the enzyme, but the dioxygen reductase activity is quickly and
fully recovered upon NO removal due to a unusually high NO dissoci-
ation rate. [1,13] This was suggested to represent the molecular cause of
the enhanced bacterial tolerance to nitrosative stress conditions. In a
previous report, we have shown that i) NO release kinetics are affected
by pH, i.e. it is slower at pH values above 8 and ii) NO is only partly
released from the reduced D58E<sup>B</sup> variant at all pH values. [27] We
considered that mutations in the putative proton pathway could thus
also affect binding and release of substrates from the heme *d* active site.
This prompted us to study NO binding and release in the proton pathway
variants that are in the focus of this study.

Binding of NO to the reduced heme *d* of Cyt *bd*-I at pH 6 shifts the maximum band from 630 nm to 642 nm and NO release leads to the backshift to 630 nm over time (Fig. 4a). NO release from Cyt *bd*-I shows similar rates at pH 6 and 7, but is much slower at pH 8 (Fig. 4b,c,d). The same holds true for the D105A<sup>B</sup> and W57A<sup>B</sup> variants indicating that binding and release of small molecules remain almost unaffected by these mutations.

In contrast, binding of NO to the D58<sup>B</sup> variants differed from binding to the other variants and to Cyt *bd*-I (Fig. 4). A pronounced shift of the maximum in the absorbance spectra of the D58N<sup>B</sup> variant from 631 nm

to 645 nm is found. A similar shift is observed with the D58L<sup>B</sup> variant,
however, this variant has a maximum absorbance at 633 nm in its
dithionite-reduced state that shifts to 645 nm upon NO addition and to
633 nm after 1 h incubation (Fig. 4). Furthermore, NO release is
significantly hampered at pH 6 in both variants. This supports the pro-
posal that mutation of D58<sup>B</sup> will either alter the access to or the archi-
tecture of the heme *d* binding site (or both). This may lead to differences
in stabilizing reaction intermediates thus affecting Cyt *bd*-I kinetics.

3.4. FTIR difference spectroscopy

Fig. 5 shows the oxidized minus reduced FTIR difference spectra of the D105A<sup>B</sup> variant, along with those of the D58E<sup>B</sup> and Cyt <i>bd</i>-I reported previously [26] for comparison. The spectrum obtained with the D58N<sup>B</sup> variant is shown in Fig. S3. Since the *d*-heme content is diminished to more than 60 % in the latter variant (Fig. 2), an interpretation of the data is extremely difficult.

The spectra shown in Fig. 5 reflect contributions from the backbone,
the hemes, the quinones and individual amino acids reorganizing upon
the redox reaction. The C=O, C=C and C-OCH3 vibrational modes from
the oxidized quinones are found at 1672, 1612 and 1264 cm⁻¹,
respectively. [40,41] Absorbances from reduced quinols are involved in
the broad features between 1500 and 1200 cm⁻¹. Importantly, proton-
ated acidic amino acid residues are seen in the region between 1780 and
1715 cm⁻¹, the deprotonated carboxylates are expected to contribute in
the 1550–1570 cm⁻¹ spectral range (COO⁻)ᵃᵉ, and around 1400 cm⁻¹
(COO⁻)ᵃᵉ. [25,42] Signals from several protonated acidic residues mainly
contribute to the spectral region above 1715 cm⁻¹. A pKa above 8 was
calculated for D58ᵇ using the PropKa 2.0 software. [27] Thus, the
carboxylate of D58ᵇ should contribute to the spectral region highlighted
in Fig. 5. Clearly, several signals are perturbed in the critical spectral
region of the variant's spectra. Importantly, the shifts observed with the
D105Aᵇ and D58Eᵇ variants are very similar. It is noted that the change

<figure><img src="image_7.png" /><figcaption>**Fig. 4.** pH-dependent NO-binding and release to heme *d* in reduced Cyt *bd*-I at pH 6 *a*; reduced protein (black), complex heme *d*-NO (red), final state 1 h after NO binding (green). Shift of the maximum wavelength of the heme *d*-NO complex over 1 h with 100 equivalents excess NO at pH 6 *b*), pH 7 *c*) and pH 8 *d*); *bd*-I oxidase (black), D105A<sup>B</sup> (red), W57A<sup>B</sup> (blue), D58N<sup>B</sup> (orange) and D58L<sup>B</sup> (dark green).</figcaption></figure>
<figure><img src="image_8.png" /><figcaption>**Fig. 5.** Oxidized minus reduced FTIR difference spectra of Cyt *bd*-I (a), variant D58E<sup>B</sup> (b), and variant D105A<sup>B</sup> (c) at pH 7 in the +500/-500 mV (vs Ag/AgCl) potential range. The insets show the double difference spectra obtained by subtracting D58E<sup>B</sup> minus Cyt *bd*-I (black line) and D105A<sup>B</sup> minus Cyt *bd*-I (red line). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

from an aspartate to a glutamate residue is expected to change the position of the COOH vibrational modes. The double difference spectra in the left inset of Fig. 5 show the decrease of a contribution around 1762 cm⁻¹ and the increase of a signal at about 1750 cm⁻¹ in both variants. E107A that is located close to heme *d*, was previously shown to be involved in the signature shown here. [25] However, this residue is too far away to be affected by the mutations in CydB reported here.

Here, we also used PropKa 2.0 [43,44] and calculated the pKa of D105B to 4. This value is close to the textbook value of 3.7 as for example found in the Dayhoff matrix. [45] This is not surprising as D105B points into the putative proton pathway that comprises five structurally resolved water molecules [3,6]. Due to the hydrophilicity of the residues forming this pathway, it is expected that it binds several additional, mobile water molecules. Thus, D105B is positioned in an aqueous environment resulting in a similar pKa as found in aqueous buffer solutions. Accordingly, the carboxylate group of D105B is deprotonated under the given experimental conditions. Indeed, a signal at 1558 cm⁻¹ that is characteristic for the (COO⁻)ᵃs vibrational modes is modified in the D105Aᵇ variant (Fig. 5, right inset). The spectral change in the region above 1710 cm⁻¹ is thus an indirect effect of the mutation in the proton path, i.e. due to the dislocation or removal of water molecules. We suggest that D58B is affected by the D105B mutations and vice versa,

leading to the observed spectral shifts. These spectral alterations in the region between 1780 and 1715 cm⁻¹ are indicative for acidic amino acid side chains and they are very similar for the D105Aᵇ and D58Eᵇ variants (Fig. 5). It is very likely that different spectra of the individual variants would result if the mutations led to isolated effects. The D105Aᵇ variant features only a very low activity (Table 1) and it was recently shown that mutations at this position change the kinetics of the catalytic reaction. [27] The perturbation of the protonation state of D58B induced by this mutation could possibly contribute to this observation.

## 4. Conclusions

Positions D58B and D105B are crucial residues for the proton uptake in E. coli Cyt *bd*-I. Electrocatalytic studies of the corresponding variants reveal that the pH dependency of oxygen reduction is associated with these residues. FTIR difference spectra demonstrate that mutations of D105B and D58B influence each other, most likely due to a connecting hydrogen bond network. In addition, NO release is found to be slower with the D58 variants than in Cyt *bd*-I. It is suggested that oxygen and NO bind competitively and the slower oxygen reaction in these variants privileges NO binding. In conclusion, the data allows us to trace a proton channel from D105B to D58B and then to the active site, helping to
further complete the picture of the enzyme mechanism.

### CRediT authorship contribution statement

**Raaif Siddeeque:** Methodology, Investigation, Formal analysis.
**Lucia Heger:** Investigation. **Jan Kägi:** Investigation. **Thorsten Friedrich:** Writing – review & editing, Project administration, Funding acquisition. **Frédéric Melin:** Writing – original draft, Supervision, Formal analysis, Conceptualization. **Petra Hellwig:** Writing – original draft, Validation, Supervision, Funding acquisition, Conceptualization.

### Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

### Acknowledgements

The work in the Friedrich lab was supported by the Deutsche Forschungsgemeinschaft (DFG) by grant 278002225/RTG 2202. The work in the Hellwig lab was supported by the Interdisciplinary Thematic Institute SysChem via the IdEx Unistra (ANR-10-IDEX-0002) within the program Investissement d'Avenir.

### Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.bbabio.2025.149537.

### Data availability

Data will be made available on request.

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