An optogenetic assay method for electrogenic transporters using Escherichia coli co‐expressing light‐driven proton pump

Abstract

In organisms, nutrients and wastes move across the cellular membrane, in which membrane‐embedded transporters facilitate and inhibit the movement. Despite the physiological significances, the currently used assay methods for transporter activities require tedious preparation and analytical processes. In this study, we report the isotope‐free and label‐free measurement system for the transport activities of electrogenic transporters. In the system, two molecules, a light‐driven inward proton pump rhodopsin, xenorhodopsin (XeR), and a representative of an electrogenic transporter, an oxalate transporter (OxlT), were co‐expressed in Escherichia coli cells. The light illumination of the cells co‐expressing XeR and OxlT showed an increase in the pH of the bulk solution and that the extent of the pH change is significantly enhanced by adding the oxalate, suggesting the light‐induced inward proton transport by XeR coupled to the negative electrogenic transport by OxlT. Such a pH increase was dependent on the oxalate concentration, but not on the XeR expression level. Of note, pH increase was not observed for the nonfunctional mutants of OxlT, R272A, and K355Q, supporting the validity of the system. Thus, we successfully developed an optogenetic assay method for electrogenic transporters using E. coli co‐expressing light‐driven proton pump.

1. INTRODUCTION

Transporters exist in the biological membranes of organisms in all kingdoms, with critical roles in maintaining their lives.¹ These integral membrane proteins catalyze the translocation of a wide array of chemicals such as ions, nutritious or hazardous substances, and signaling molecules as neurotransmitters, across the membranes. Their activities are responsible for cell growth and intercellular communications and thus are essential to various physiological functions.², ³, ⁴ The functions of the transporters mainly consist of two processes: the recognition of substrates and the permeation of substrates across the membrane. These processes are analyzed by various assay methods in life science studies.

As analytical methods to address the substrate recognition processes of the transporters, several assay methodologies for substrate binding have been reported. For example, binding assays using radioisotope‐ or fluorescence‐labeled chemicals, fluorometric analysis on substrate binding using labeled or intrinsic fluorescence of transporter molecules, and thermal shift assays using thiol‐reactive fluorescent dyes are widely used in various transporter studies.⁵, ⁶, ⁷, ⁸ If the purified samples are used, these assay methods provide accurate parameters for substrate binding, such as the dissociation constant. In addition, these methods are generally well suited for high‐throughput analyses. However, sample preparation through solubilization and purification processes sometimes fails because of sample instability. In addition, appropriate reagents for the binding assays as labeled substrates with radioisotope or fluorescence are not always available for all transporters of interest. Furthermore, the most critical limitation for binding assays is that they cannot provide information about substrate permeation activities, an essential process for transporter functions.

In contrast, transport assays provide information about the entire process of transporter functions, including both substrate binding and permeation. However, despite the significance of transporter studies, methodologies for analyzing transport activities are limited. Except for a few classes of transporters for which transporter assay methods using specific fluorescent indicators are established,⁷, ⁸, ⁹ the most common method used to general transporters is the proteoliposome assay. In the method, the solubilized and purified transporter molecules are reconstituted into liposomes and subjected to the uptake analysis of typically radioisotope‐labeled substrates.¹⁰, ¹¹ Nevertheless, the assay needs tedious processes for sample preparation, such as the solubilization and purification of the proteins of interest and their reconstruction into liposomes. It also needs appropriate radiolabeled chemicals or reagents, enabling specific quantification of transported substrates in the liposomes. Under these situations, transport assays with feasibility and general applicability are required.

Of note, visible light is useful as a stimulus for triggering the reaction of biological molecules. Among the various photoreactive molecules, microbial rhodopsins are widely distributed into various organisms and work as light‐driven ion transporters.¹² Microbial rhodopsins consist of 7‐transmembrane helices and the retinal chromophore, resulting in the absorption of the visible light.¹² Upon light absorption, microbial rhodopsins show a cyclic photoreaction called photocycle in the time domain from milliseconds to seconds.¹³ They are widespread in the microbial world with their diverse protein functions, including light‐driven ion pumping and channeling, during the photocycle.¹⁴ The ion‐transporting rhodopsins are used as tools for optogenetics, controlling neural activity by light.¹⁵

In this study, we proposed a novel transport assay method using Escherichia coli cells recombinantly co‐expressing a transporter protein and a light‐driven proton pump rhodopsin. In many cases, transporter activities are electrogenic: the translocation of charged substrates or symport/antiport of substrates and ions across the membrane consequently constructs membrane potentials.¹⁶ In the method, the electrogenic transport activities of the transporter of interest expressed in E. coli are optogenetically coupled to light‐dependent proton translocation across the membrane by the co‐expressed proton pump rhodopsin, allowing the evaluation of the transporter functions without purification from the cells. As a proof‐of‐concept study of the proposed method, we report an evaluation of the activity of oxalate transporter (OxlT), which catalyzes the antiport of oxalate and formate,¹⁷, ¹⁸ using xenorhodopsin (XeR), a light‐dependent inward proton pump.¹⁹

2. RESULTS

2.1. Assay design

In this study, we aimed to construct a new assay system to evaluate the electrogenic transport activity of a transporter by coupling a light‐dependent proton movement through a proton pump. We used an inward proton pump XeR from Rubricoccus marinus as a model for a light‐dependent proton translocator because of several reasons¹⁹: (i) The transporter function of R. marinus XeR is proton‐selective, whereas channelrhodopsins, the most general optogenetic tools, conduct not only proton but also other cations²⁰, ²¹; (ii) R. marinus XeR expresses well in E. coli cells as a recombinant protein, whereas full‐length channelrhodopsins, generally used for eukaryotic cells, barely express as functional forms in E. coli ²¹, ²²; and (iii) R. marinus XeR is known to be functional under a wide array of conditions, such as in the extracellular fluids with various kinds of salts. As a case study, the transport activity of OxlT is evaluated by the magnitude of the pH increase caused by the irradiation of E. coli co‐expressing OxlT and XeR with light before and after the addition of oxalate (Figure 1a). OxlT is a member of the major facilitator superfamily²³ existing in the membrane of an oxalate‐degrading bacterium Oxalobacter formigenes in the human gut microbiota.¹⁷ OxlT is responsible for oxalate uptake into the bacterial cells and the counter‐transport of formate, a decomposed oxalate product, from the cells.¹⁷, ¹⁸, ²⁴ The assay system is based on the assumption that the inward proton transport by XeR is promoted by the formation of a negative membrane potential across the membrane constructed by the exchange of divalent anion oxalate and monovalent anion formate by OxlT¹⁷, ¹⁸ (Figure 1b). According to the E. coli Metabolome Database,²⁵ the intracellular concentration of formate in E. coli is estimated to be 184 ± 25 μM. On the other hand, no relevant data are registered for oxalate, implying that no significant detection of the latter compound was reported in previous metabolomic studies. Therefore, an addition of oxalate from the outside of living E. coli cells expressing OxlT is expected to result in the counter‐transport of formate in the cytosol.

FIGURE 1.

Assay design. (a) Schematic diagram of the experimental setup of the transporter assay system proposed in this study. The pH change of the suspension solution of E. coli co‐expressing OxlT and XeR during the light irradiation is measured. The light‐driven inward proton transport of XeR is assumed to be promoted by the negative membrane potential formed by the oxalate/formate antiport of OxlT. (b) Speculative pH change expected as a result of the proposed assay method. The light‐dependent pH change in the presence of oxalate (ΔpH_S) is expected to be enhanced than that in the absence of oxalate (ΔpH₀). (c,d) Representative results of pH changes measured on E. coli suspended to 50 mM K₂SO₄ illuminated with a Xe lamp at a power of 150 mW/cm² with a long‐pass filter (≥ ~420 nm, yellow bars). The initial pH ranged from 6.05 to 6.35 (Figures S1–S3). The traces of pH changes in the presence (orange) and absence (gray) of 5 mM oxalate were shown. (c) E. coli co‐expressing OxlT and XeR. (d) E. coli only expressing XeR

The previous transport assays for OxlT using reconstituted proteoliposomes have been performed under conditions without sodium and chloride ions.²⁶, ²⁷ In particular, chloride ion has been reported to inhibit the transport of OxlT.¹⁷ Since it has been confirmed that XeR‐expressing E. coli exhibits photo‐dependent proton transport activity even when the Na₂SO₄ or KCl solution is used as the extracellular fluid,¹⁹ we chose the K₂SO₄ solution as an extracellular fluid in this study. In addition, oxalate was added in the form of potassium salt as reported.²⁶, ²⁷ Under this condition, E. coli cells expressing XeR exhibited a light‐dependent pH increase of the extracellular fluid, resulting in the proton transport from the outside to the inside of the cells (Figures 1c,d and S1A). Notably, the extent of pH increase was enhanced after adding oxalate to the E. coli cells co‐expressing OxlT (Figure 1c), whereas no such enhancement was observed on the cells without OxlT expression (Figure 1d). These results suggested that, as expected, oxalate transport‐coupled proton transport can be measured under the condition. The oxalate‐dependent enhancement of the pH increase was detected more sensitively than the difference of the initial velocity of the pH increase (Figure 1c). Therefore, we focused on the extent of the pH increase in the following analyses.

We confirmed that the extent of light‐dependent pH increase observed on the suspension of E. coli cells co‐expressing OxlT and XeR, but not that only expressing XeR, was dependent on oxalate concentration (Figure 2a). The result suggested that the observed pH increases are attributed to the transporter activity of OxlT. Therefore, pH increase enhancement by adding oxalate, ΔΔpH defined as the difference between the pH increases in the presence (ΔpH_S) and the absence (ΔpH₀) of oxalate, is considered to serve as an appropriate measure of OxlT transport activity.

FIGURE 2.

Verification of the proposed assay system. (a) The light‐dependent pH increase (ΔpH) of E. coli suspension solution in the presence of a different oxalate concentration (0, 1, 10 mM). The values after 3‐min irradiation of the light at a wavelength of 550 ± 10 nm and a power of 8 mW/cm² are shown. +OxlT and −OxlT indicate the results from E. coli cells expressing XeR with or without OxlT, respectively. (b) The pH increases of suspension solution of E. coli co‐expressing OxlT and XeR at different light intensities (60, 100, 150 mW/cm² at 550 nm). The line graph shows the light‐dependent pH increases (ΔpH) after 5‐min light irradiation, where the gray and orange lines indicate the results in the absence (ΔpH₀) and presence (ΔpH_S) of 5 mM oxalate, respectively, and the black dots indicate the ΔpH₀ observed from two independent measurements. The bar graph shows the enhancement of the pH increase by addition of 5 mM oxalate (ΔpH_S – ΔpH₀; ΔΔpH). (c,d) Correlation analysis between the XeR expression and the pH increase of suspension solution of E. coli co‐expressing OxlT and XeR. (c) Correlation between the expression levels of retinal‐integrated XeR and pH increases in the absence of oxalate (ΔpH₀). (d) Correlation between the expression levels of retinal‐integrated XeR and enhancement of the pH increase by addition of 5 mM oxalate (ΔΔpH), the measure of OxlT activity, of the relevant experimental lots. In panels (c) and (d), the samples with varied expression levels of XeR were obtained from nine different experimental lots prepared under the same condition described in Materials and Methods

We further tested whether the ΔΔpH value is affected by the other experimental conditions. The enhancement of the pH rise (ΔΔpH) is not significantly affected by the light intensities for irradiation, whereas the extent of pH rise itself (ΔpH) increased following light intensification (Figure 2b). The result suggested that light irradiation does not suppress the OxlT transport activity, at least up to 150 mW/cm² at 550 nm. We also verified the effect of the XeR activity on the measured values of OxlT transporter activity. The relative amount of active XeR molecules integrating retinal, that is, holo‐XeR, expressing in E. coli cells was evaluated by quantifying the reddish‐purple density of the cells attributed to holo‐XeR¹⁹ by image analysis (see Materials and Methods). The estimated holo‐XeR expression levels of E. coli used for the nine independent experiments showed a strong correlation with their pH increases in the absence of oxalate (ΔpH₀, Figure 2c), with the correlation coefficient R of 0.838, indicating that the ΔpH₀ values attribute to XeR activity in the cells as expected. In contrast, the ΔΔpH values, the measure of OxlT transport activity, showed much less correlation to the estimated holo‐XeR expression levels of the relevant experimental lot (R = −0.401). These results suggested that the level of XeR expression and its inward proton transport activity unlikely serve as a limiting factor for the ΔΔpH values to evaluate the OxlT activity, and thus the OxlT activities of the different samples can be assessed regardless of the variation of holo‐XeR levels under the tested conditions.

Collectively, the proposed assay system is expected to properly evaluate the transport activity of OxlT.

2.2. Verification of the assay method by OxlT mutant analysis

In OxlT, two basic residues, Arg272 and Lys355, are known to be critical for binding and transporting oxalate dianion, whereas R272A and K355Q mutants are reported to lose oxalate transport activity.²⁶, ²⁷ To assess the validity of the assay method constructed in this study, we analyzed the transport activity of R272A and K355Q mutants using the system (Figures S1–S3).

For evaluating the transport activities of the OxlT mutants, the measured values were corrected and standardized with that of the wild‐type OxlT measured on the same day of the experiment as follows. Firstly, we chose the oxalate concentration at 5 mM to maximize the ΔΔpH of the OxlT‐expressing samples and minimize the oxalate effects independent of OxlT, such as an effect on XeR activities or a buffering action by oxalate. Furthermore, to eliminate the effect of oxalate addition independent of the OxlT activities, the ΔΔpH values were corrected by the subtraction with that measured on the cells only expressing XeR. The corrected ΔΔpH values of the mutant OxlT were normalized to that of the wild‐type OxlT measured on the same day of the experiment. In addition, to cancel out the differences in expression levels, the normalized ΔΔpH values for mutant OxlT were further corrected using the relative expression levels analyzed by western blotting with those of wild‐type OxlT (Figures S1–S3). The derived values were defined as the normalized transport activity (%; see Materials and Methods).

As shown in Figure 3, the R272A and K355Q OxlT mutants exhibited almost no transport activities using the assay system, as expected. In summary, we concluded that the constructed assay system is applicable to evaluate the activity of an electrogenic transporter, OxlT.

FIGURE 3.

Analysis of transport activities of mutant OxlT by the constructed assay system. Mutant OxlT transport activities are shown as relative values with that of the wild‐type set as 100%

3. DISCUSSION

In this study, we reported a transport assay method using living E. coli cells that recombinantly expressed a transporter of interest. Since the electrogenic transport activity of the target molecule is coupled to proton translocation across the membrane catalyzed by microbial rhodopsin with a proton pump function, the activity is feasibly evaluated as a pH change of the solution outside of the cells using a pH electrode equipped in most of the biochemical laboratories. A pivot of this method is the use of a light‐driven proton pump that enables an optogenetic control of the proton translocation. This is effective for the specific detection of the pH change and assures the direction of the proton movement across the membrane matching to the direction of the membrane potential created by the transporter. The optogenetic approach is advantageous compared with the transporter assay with a similar concept, using a voltage‐sensitive fluorescent indicator for tracking membrane potential changes,²⁸ for assuring the specificity of detection devoid of other factors affecting the fluorescent intensities. In this study, we used XeR, an inward proton pump, as a coupling probe protein for negative‐electrogenic transport by OxlT. If an outward proton pump, such as archaerhodopsin‐3,²⁹ is used instead of an inward proton pump, evaluation of the activities of positive electrogenic transporters is likely achieved. Light‐gated proton channel rhodopsin converted from pump rhodopsin, which is amenable to express in E. coli,³⁰ is also expected to serve as a suitable tool for this system.

As a transport assay, the proposed method has several advantages compared with the proteoliposome assays. Firstly, the method proposed in this study does not require the solubilization and purification process of the target transporter. It dramatically improves the feasibility of the assay. More notably, the sample preparation without a solubilization process is expected to increase the success ratio for evaluating the activities of unstable transporters or mutants losing stability in the detergents. Another advantage is general applicability. The method is applicable with a conventional pH electrode and an illumination light source. It does not require specific labeled reagents and detection setups, such as radioisotope‐labeled substrates and a radioisotope laboratory equipped with a scintillation counter. A high‐throughput analysis can also be achieved by constructing an experimental setup allowing parallel multiple sample analysis, such as with multiple pH electrodes and a light source capable of simultaneous multisample illumination. The other notable characteristic of the method is that it generally achieves a unified orientation of the transporter in the membrane, most likely with a physiological orientation, because the transporter molecules are expected to be integrated into the membrane by an intrinsic physiological system in the cells, that is, co‐translational translocation mediated by ribosomes and translocons. The situation contrasts with the case of proteoliposome reconstruction, in which transporter molecules are spontaneously inserted into the lipid bilayer with a random orientation in many cases.¹⁰

It should be noted that the proposed method has some limitations. Firstly, although the method is applicable to a wide array of electrogenic transporters, it cannot be applied to non‐electrogenic transporters and proton‐coupled transporters. In addition, since it uses living E. coli cells, the intrinsic proteins in the E. coli might affect the measured activities. For example, if the substrate of the transporter also serves as a substrate for other proteins in the E. coli membrane, the knockout of the corresponding genes might be required for the accurate analysis. Even if the transporter activity is specific to the cells, other macromolecules in the system might give high background signal compared with the case using the purified proteins, sometimes resulting in a determination of kinetic parameters with higher errors. Although we confirmed that XeR activity did not restrict OxlT activity under the tested conditions in this study, it might not always be a case for a combination of other transporters and proton pumps unless the experimental condition is carefully designed. In addition, the measure of the transporter activity employed in this study, ΔΔpH, might be affected by the environmental change during the measurement. Thus, the quantitative accuracy of the method might be limited, and the assay is likely more suitable for use to compare relative activities, such as wild‐type and mutant transporters, rather than to determine their accurate kinetic parameters. If the transporter of interest is misfolded in E. coli, which may happen especially on eukaryotic membrane proteins in some cases,³¹, ³² the assay method is inapplicable. Conversely, by examining whether a significant signal can be detected using this method, it is possible to assess whether E. coli serves as a host for the functional expression of those transporters. On the other hand, the applicability of eukaryotic cells instead of E. coli as an expression platform for this system is currently unclear. It probably needs to optimize the assay condition because the current protocol requires high cell density to detect a significant pH change of the solution outside of the cells.

Nevertheless, the proposed method is expected to serve as an effective alternative to the proteoliposome assay for evaluating transporter activity. Because of its feasibility and general applicability, methodological development based on the current protocol is expected to establish a high‐throughput assay system to investigate transporter functions.

4. MATERIALS AND METHODS

4.1. Vector construction

For the construction of the E. coli expression vector for OxlT, pRSF‐OxlT, the gene encoding wild‐type OxlT with a C‐terminal nona‐His‐tag was amplified from pBDOxlTHis³³ by conventional PCR and inserted into the isopropyl β‐d‐thiogalactopyranoside (IPTG)‐inducible plasmid vector, pRSF‐1b (Novagen), between the NcoI and AvrII by Gibson Assembly (New England Biolabs).³⁴ The expression vectors for R272A and K355Q mutant OxlT were constructed by overlapped PCR using PrimeSTARMax (Takara Bio) using the wild‐type pRSF‐OxlT as a template. For the expression of XeR from Rubricoccus marinus, we used the vector pET21a(+)‐RmXeR, in which the gene for the six‐histidine‐tagged XeR (protein accession no. WP_094549673) was inserted into the IPTG‐inducible plasmid vector, pET21a(+), using NdeI and XhoI restriction enzyme sites.¹⁹

4.2. Transport assay

The expression vectors for OxlT and XeR, pRSF‐OxlT and pET21a(+)‐RmXeR, were transformed into E. coli strain BL21 (DE3) cells. The cells were grown at 37°C in LB medium supplemented with 20 μg/ml kanamycin and 100 μg/ml carbenicillin. When the cell density reached an absorbance of 0.8–0.9 at 600 nm, the protein production was induced by adding IPTG and all trans‐retinal (SIGMA) at the final concentration of 1 mM and 10 μM, respectively. After further culture at 20°C for 20 hr, the cells were collected by centrifugation of 3,500g for 5 min at 4°C. The cells were gently washed with 50 mM K₂SO₄ three times and centrifuged as above in 50‐ml polypropylene conical tubes. At this stage, images of the tubes were taken for the expression analysis of active XeR described below. The cells were then resuspended into 50 mM K₂SO₄ to the cell density at an absorbance of ~10 at 660 nm and subjected to the pH measurement as below. For the experiments using the E. coli cells only expressing XeR (mock‐transformed), BL21 (DE3) cells were transformed with the vector pRSF‐1b without the insertion of the OxlT gene and pET21a(+)‐RmXeR and were subjected to cell culture and protein production as described.

The pH change of the E. coli suspension solution prepared above was monitored with a pH electrode (LAQUA F‐72 pH meter, HORIBA). The cell suspension was kept at 25°C and continuously stirred while measuring. Before the measurement, the cell suspension was placed in the dark until the pH became stable.

In the evaluation of the transport activity of OxlT, the light‐induced pH change was measured as follows, otherwise stated in the text. The cell suspension was illuminated with a Xe lamp (Max‐303, Asahi Spectra) through a Sharp Cut Filter Y44 (a long‐pass filter at ≥~420 nm, HOYA) for 10 min. The light intensity was adjusted to approximately 150 mW/cm² at 550 nm using an optical power meter (#3664, Hioki) and an optical sensor (#9742, Hioki). First, the accumulated light‐induced pH change was measured in the absence of oxalate for 10 min (ΔpH₀). After the pH of the illuminated sample became stable, 5 mM potassium oxalate was added to the same sample. Then, 10 min after the addition of potassium oxalate, the sample was again illuminated, and the accumulated light‐induced pH change was measured for 10 min (ΔpH_S). The transport activity was evaluated by the difference of the pH change (ΔΔpH) between ΔpH_S and ΔpH₀ (Equation (1)), followed by the correction with subtracting the background differential pH change (ΔΔpH) measured using mock‐transformed E. coli only expressing XeR to eliminate the oxalate effect independent of OxlT (Equation (2)). The activities of each mutant were normalized by the corrected ΔΔpH and the relative expression level, analyzed using western‐blotting described below, of the wild‐type OxlT measured on the same day of the experiment (Equation (3)).

$$∆∆\mathrm{pH}=\left(∆{\mathrm{pH}}_S-∆{\mathrm{pH}}_0\right)$$ (1)

$$\text{Corrected}∆∆\mathrm{pH}=∆∆\mathrm{pH}\left(\text{mutant or wild type OxlT}\right)-∆∆\mathrm{pH}\left(\text{mock transformed}E.\mathit{\text{coli}}\text{only expressing}\mathrm{XeR}\right)$$ (2)

$$\text{Normalized OxlT transport activity}\left(\%\right)=\frac{\text{corrected}∆∆\mathrm{pH}\text{of mutant OxlT}}{\text{corrected}∆∆\mathrm{pH}\text{of wild type OxlT}}\times \frac{\text{expression level of wild type OxlT}}{\text{expression level of mutant OxlT}}\times 100$$ (3)

5. EXPRESSION ANALYSIS

The samples used for pH measurements were centrifuged at 3,500g at 4°C for 5 min. Then, the precipitated cells were resuspended with TBS buffer (10 mM Tris–HCl, 150 mM NaCl, pH 7.5) to the cell density at an absorbance at 660 nm of 10. An aliquot of the cell suspension (6 μl sample for one lane of SDS‐PAGE gel) was lysed with SDS‐PAGE sample buffer (0.05 M Tris–HCl, 2% SDS, 10% glycerol, 0.003% bromophenol blue, pH 6.8) including 100 mM dithiothreitol and 0.042 U/ml Benzonase Nuclease (Novagen). After incubation at room temperature for 60 min, the samples were subjected to SDS‐PAGE and then transferred to Amersham Hybond PVDF membrane (GE Healthcare) by electroblotting. The membrane was blocked with Blocking One (Nacalai Tesque) at 4°C overnight or at room temperature for 60 min. The membrane was washed with TBS‐T buffer (10 mM Tris–HCl, 150 mM NaCl, 0.5% Tween 20, pH 7.5) three times and incubated with Penta‐His HRP Conjugate (1:2,000 dilution, QIAGEN, no. 1014922) at room temperature for 60 min. After the membrane was washed with TBS‐T buffer, the chemiluminescent signals were detected with Immobilon Western Chemiluminescent HRP Substrate (Millipore) and Molecular Imager ChemiDoc XRS+ Imaging System (Bio‐Rad). The band intensities blotted on the same membrane were quantified using the ImageJ software,³⁵ and the average values of the triplicated samples were used for calculation to determine the relative expression level of the OxlT in the samples (expression level of wild type OxlT/ expression level of mutant OxlT) measured on the same day of the experiment.

In the case of XeR, the bands detected using western blotting comprise both active XeR integrating retinal, holo‐XeR, and inactive XeR without retinal, apo‐XeR. Holo‐XeR, but not apo‐XeR, has an absorption peak at a wavelength of 550 nm; thus, E. coli cells expressing holo‐XeR exhibit the complementary color, that is, reddish purple.¹⁹ The expression levels of holo‐XeR in the cells used for pH measurements were evaluated by the image analysis exploiting the absorption property of XeR using ImageJ. The cells corresponding to a 300 ml culture volume were collected in a 50 ml polypropylene conical tube (VIOLAMO, 1‐3500‐22) as described above. After supernatant removal, the tubes were placed on a blank sheet of paper, and the images of the cells in the tubes were obtained using a camera integrated with a mobile phone Galaxy Note 9 (SCV40, Samsung Electronics) from an angle of about 45° under fluorescent lights on the ceiling. Since the absorption attributed to holo‐XeR well matches the green color, we evaluated the amount of holo‐XeR in the cells with the color density of reddish‐purple by removing the green component from the image. For this purpose, the obtained images were first separated into three images: a channel either in red, green, or blue by “Split Channels” in ImageJ. The images were then recomposed by the “Image Calculator” with the equation: (the red channel image – the green channel image) + (the blue channel image – the green channel image) to eliminate the green and background light components. In this image composition, the background light component was assumed to be white light consisting of the same intensity of the red, green, and blue components; that is, the background component in red or blue is the same as that in green. Finally, the threshold of the recomposed image was set to 1–255, and the average brightness in this range was used as the expression level of holo‐XeR in each sample.

CONFLICT OF INTEREST

The authors declare no conflict of interest.

AUTHOR CONTRIBUTIONS

Masahiro Hayashi: Formal analysis; writing‐original draft; writing‐review & editing. Keiichi Kojima: Formal analysis; writing‐review & editing. Yuki Sudo: Conceptualization; funding acquisition; methodology; resources; supervision; writing‐original draft; writing‐review & editing. Atsuko Yamashita: Conceptualization; formal analysis; funding acquisition; resources; supervision; writing‐original draft; writing‐review & editing.

Supporting information

Appendix S1: Supporting Information

Hayashi M, Kojima K, Sudo Y, Yamashita A. An optogenetic assay method for electrogenic transporters using Escherichia coli co‐expressing light‐driven proton pump. Protein Science. 2021;30:2161–2169. 10.1002/pro.4154