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Abstract

Inhibition of the K_Ca3.1 potassium channel has therapeutic potential in a variety of human diseases, including inflammation-associated disorders and cancers. However, K_Ca3.1 inhibitors with high therapeutic promise are currently not available. This study aimed to establish a screening assay for identifying inhibitors of K_Ca3.1 in native cells and from library compounds derived from natural products in Thailand. The screening platform was successfully developed based on a thallium flux assay in intestinal epithelial (T84) cells with a Z′ factor of 0.52. The screening of 1352 compounds and functional validation using electrophysiological analyses identified 8 compounds as novel K_Ca3.1 inhibitors with IC₅₀ values ranging from 0.14 to 6.57 µM. These results indicate that the assay developed is of excellent quality for high-throughput screening and capable of identifying K_Ca3.1 inhibitors. This assay may be useful in identifying novel K_Ca3.1 inhibitors that may have therapeutic potential for inflammation-associated disorders and cancers.

Keywords

assay development potassium channel high-throughput screening natural compound

Introduction

The intermediate-conductance Ca²⁺-activated potassium channel (K_Ca3.1; also known as KCNN4, IK1, IK_Ca1, SK4, and Gárdos channel) is ubiquitously expressed in many tissues throughout the body, including cells of hematopoietic lineages, epithelial tissues, vascular endothelia, fibroblasts, and smooth muscle cells.¹ The function of K_Ca3.1 was first described in human erythrocytes by Gárdos in 1958.² The efflux of K⁺ through this channel alters the osmotic balance and causes shrinkage of erythrocytes, establishing K_Ca3.1 as a regulator of erythrocyte cell volume.³ In addition, K_Ca3.1 is functionally coupled with calcium channels. Calcium influx through the Ca²⁺ channels elevates the intracellular Ca²⁺ level, causing K_Ca3.1 activation. Potassium efflux through K_Ca3.1 in turn induces membrane hyperpolarization, which counteracts membrane depolarization caused by Ca²⁺ influx, maintaining the electrical gradient and sustaining Ca²⁺ channel activity.⁴ Due to this role, K_Ca3.1 is an important player in cellular calcium signaling involved in the activity of various immune cells.⁵ The activation of T lymphocytes is associated with the upregulation of K_Ca3.1, whereas loss of K_Ca3.1 function is associated with the reduction of activated phenotypes such as cytokine production.⁶ Similar to T cells, K_Ca3.1 is involved in functions of B lymphocytes, macrophages, microglia, immature dendritic cells, mast cells, and neutrophils.^5,7 Furthermore, K_Ca3.1 is expressed in most epithelial cells,⁸ where it functions to protect against hypotonic stress^9,10 and provide a driving force for anion secretion.^11,12

K_Ca3.1 is regarded as a promising drug target for many diseases. Its role in erythrocyte volume regulation leads to the first potential therapeutic application of K_Ca3.1 inhibitor in sickle cell disease characterized by deformation of erythrocytes into a sickle-like shape because of self-polymerizing sickle hemoglobin (HbS). This “sickling” process is highly sensitive to intracellular HbS concentration and exacerbated by cell dehydration.¹³ The roles of K_Ca3.1 in immune cell activation established K_Ca3.1 inhibitors as anti-inflammatory agents.⁶ In intestinal epithelial cells, K_Ca3.1 has been proposed to be a drug target for the treatment of diarrheal diseases caused by excessive Cl⁻ secretion, such as rotavirus-induced diarrheas¹⁴ and cholera.^15,16 In addition, inhibition of K_Ca3.1 has been shown to increase the migration of intestinal epithelial cells in a PI3K/Akt-dependent manner, which may be useful for promoting wound healing.¹⁷ Importantly, several lines of evidence suggest that K_Ca3.1 inhibitors may be useful in the treatment of cancer. K_Ca3.1 overexpression is associated with poor prognosis of lung cancer,¹⁸ renal cell carcinoma,¹⁹ and ovarian cancer.²⁰ K_Ca3.1 inhibition reduces cancerous phenotypes in disease models of pancreatic cancer, prostate cancer, breast cancer, melanoma, glioblastoma, hepatocellular carcinoma, colorectal cancer, lymphocytic leukemia, and endometrial carcinoma.^21–23

Despite these diverse potentials, there is currently no K_Ca3.1 inhibitor available for therapeutic use. TRAM-34, a K_Ca3.1 inhibitor widely used in research, has a short half-life and poor oral bioavailability,¹ rendering it unfavorable as a drug candidate. In addition, Senicapoc, another K_Ca3.1 inhibitor that entered a clinical trial for sickle cell disease, ultimately failed the phase 3 study due to its lack of efficacy in reducing vaso-occlusive crises in patients.²⁴ In order to identify novel classes of K_Ca3.1 inhibitors from a large number of molecular scaffolds, the implementation of high-throughput screening technologies is required. The screening method using thallium (Tl⁺) flux assays in K_Ca3.1-transfected HEK293 cells has been reported.²⁵ Since K_Ca3.1 function depends not only on intracellular Ca²⁺ but also on posttranslational modifications, including serine²⁶ and histidine²⁷ phosphorylation as well as protein trafficking,²⁸ assays using exogenously introduced K_Ca3.1 might not fully recapitulate the complex regulatory mechanisms of endogenously expressed K_Ca3.1. Therefore, an assay with an endogenous expression system would be preferable in this regard. Intestinal epithelial cells have been shown to express K_Ca3.1, and functional measurement of K_Ca3.1 activity in these cells is readily achievable by basolateral K⁺ current analyses. Therefore, this study was aimed at developing a cell-based assay for identifying K_Ca3.1 inhibitors in an endogenous expression system, that is, intestinal epithelial cell lines, and using this assay for the initial screening of compounds derived from Thai natural products. We have successfully established a Tl⁺ flux assay in T84 cells and several novel K_Ca3.1 inhibitors have been identified by this assay.

Materials and Methods

Chemical Reagents

Fetal bovine serum (FBS), culture media, trypsin, penicillin, streptomycin, and FluxOR II Green Potassium Ion Channel Assay were purchased from Thermo Fisher Scientific Inc. (Waltham, MA). Other chemicals were purchased from Sigma-Aldrich (St. Louis, MO). The natural compound library was provided by the Excellent Center for Drug Discovery, Faculty of Science, Mahidol University (Bangkok, Thailand).

Cell Culture

T84 (CCL-248), HT-29 (HTB-38), and HCT 116 (CCL-247) cells were obtained from the American Type Culture Collection (Manassas, VA) (passages 10–30) and cultured in a mixture of Dulbecco’s modified Eagle’s medium and Ham’s F-12 medium (1:1) supplemented with 10% FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin. The cells were grown in a humidified incubator under an atmosphere of 5% CO₂ at 37 °C. The culture medium was changed every other day and cells were split every 7 days.

Thallium Flux Assay

The cells were plated at a density of 2.5 × 10⁵ cells/well in black, flat-bottom, 96-well plates (Costar, New York, NY) and cultured overnight until confluence. The cells were then incubated for an hour with FluxOR II reagent dye in loading buffer, followed by washing and 30 min of exposure to 50 µL of assay buffer containing DMSO (vehicle control), TRAM-34 (K_Ca3.1 inhibitory control), or test compounds. Then, baseline fluorescence intensity was obtained before the injection of 50 µL of stimulus buffer containing NS309 (K_Ca3.1 activator) and 5 mM Tl₂SO₄, immediately followed by measurements of fluorescence kinetics for 1–2 min. The fluorescence intensity was measured at excitation/emission wavelengths of 495 nm/519 nm using a microplate reader installed with a dispenser unit for injection of the stimulus buffer (EnVision; PerkinElmer, Waltham, MA). The fluorescence intensity of each well was normalized with an average baseline fluorescence intensity value from 10 repeated measurements prior to stimulus buffer injection. Normalized data points after Tl⁺ injection were fitted to quadratic regression. If R² for quadratic regression was more than 0.9, the initial rate of the quadratic curve was used to represent the rate of Tl⁺ influx. If data fitted poorly to the quadratic curve (R² < 0.9), simple linear regression would be performed instead, and the slope of the regression line was used to indicate the Tl⁺ flux rate. All regression models were performed in Microsoft Excel 2019 (Redmond, WA). The K_Ca3.1 activator and inhibitor control wells were situated as the first and last columns of the 96-well plate with the instrument set to read the signal by row, ensuring the data from control groups were evenly interspersed across the plate during measurements. To calculate the percentage of Tl⁺ flux inhibition, the Tl⁺ flux rate of each well was compared against the means of the K_Ca3.1 activator and K_Ca3.1 inhibitor control groups from the same plate, using the following equation:

% T l^{+} f l u x i n h i b i t i o n = \frac{μ_{N S 309} - T l^{+} f l u x r a t e}{μ_{N S 309} - μ_{T R A M - 34}} \times 100

(1)

To evaluate the assay quality, the Z′ factor was calculated from data of the K_Ca3.1 activator and K_Ca3.1 inhibitor control groups from multiple separated experiments.

Z^{'} - f a c t o r = 1 - \frac{3 \times (σ_{N S 309} + σ_{T R A M - 34})}{| μ_{N S 309} - μ_{T R A M - 34} |}

(2)

Measurement of Basolateral K⁺ Current (I_K⁺)

T84 cells were plated on Snapwell polycarbonate inserts (Costar, New York, NY) at a density of 5 × 10⁵ cells/insert. The cells were cultured with a daily replacement of media for 2 weeks until the transepithelial electrical resistance was >1000 Ω.cm², as measured by a Millicell ERS-2 volt-ohm meter (Merck Ltd., an affiliate of Merck KGaA, Darmstadt, Germany). For basolateral I_K⁺ measurement, solutions with high K⁺ and low K⁺ concentrations (pH 7.4) were added into apical and basolateral hemichambers, respectively, to generate an apical-to-basolateral K⁺ gradient. The apical high K⁺ solutions contained 145 mM potassium gluconate, 5 mM KCl, 1 mM MgCl₂, 1 mM CaCl₂, 5 mM HEPES, and 10 mM glucose. In the basolateral low-K⁺ solution, 145 mM potassium gluconate was replaced with 145 mM sodium gluconate. To induce apical membrane permeabilization, amphotericin B was added into the apical hemichamber 30 min before basolateral I_K⁺ measurement. The solutions were maintained at 37 °C and bubbled with 100% O₂. I_K⁺ was recorded using a DVC-1000 voltage-clamp (World Precision Instruments, Sarasota, FL) with Ag/AgCl electrodes and 3M KCl agar bridges.

Statistical Analysis

The statistical difference was tested using Student t tests for pairs of measurements or one-way analysis of variance (ANOVA) followed by Bonferroni’s post hoc test for multiple groups of measurements, with P < 0.05 considered significantly different. GraphPad Prism software (version 5; La Jolla, CA) was used for all statistical analyses.

Results

Development of a Screening Assay Based on the Tl⁺ Flux Assay

A Tl⁺ flux potassium channel assay was used in this study. This assay utilized two main components, fluorometric membrane-permeable Tl⁺ indicator dye and Tl⁺ ion.²⁹ Since Tl⁺ could pass through K_Ca3.1, the presence of high extracellular [Tl⁺] would generate a driving force for Tl⁺ influx into the cells. If the K_Ca3.1 was opened, Tl⁺ entered into the cells and intracellular Tl⁺ interacted with the Tl⁺ indicator dye, yielding a fluorescence signal (Fig. 1A left). In contrast, if the K_Ca3.1 was closed, Tl⁺ could not enter the cells and no change in fluorescence intensity would be observed (Fig. 1A right). The potassium channel activity could be determined from the rate of fluorescence increment. The screening protocol was depicted in Figure 1B . Briefly, the cells were loaded with the reporter dye before treatment with test compounds. Fluorescence signals before and after the addition of buffers containing both Tl⁺ and K_Ca3.1 activator were recorded. Since cells with native ion channel expression might also contain Ca²⁺-activated K⁺-permeable channels other than K_Ca3.1, a pharmacological activator NS309 was used instead of stimulation by intracellular Ca²⁺ to increased the selectivity of the assay to K_Ca3.1 channels.

Figure 1.

Overview of Tl⁺ flux assay. (A) Principle of fluorescent cell-based Tl⁺ flux assay for KCa3.1 inhibitor. Extracellular Tl⁺ ions enter the cells through the K_Ca3.1 channel and activate intracellular fluorescent reporter dye (left). K_Ca3.1 inhibition prevents Tl⁺ influx and fluorescence signal increase (right). (B) Simplified workflow of Tl⁺ flux assay. The fluorescent reporter was loaded into the cells, followed by washing and test compound addition. Baseline fluorescence intensity was measured before addition of Tl⁺ ions and K_Ca3.1 activator to initiate Tl⁺ influx for measurement of fluorescence kinetics.

To find the intestinal epithelial cell line suitable for this assay, we compared K_Ca3.1-mediated fluorescence signal induced by 10 µM NS309 in intestinal cell lines expressing K_Ca3.1, including HT-29, T84, and HCT 116. We found that the maximal signal at 1 min after Tl⁺ addition from HT-29 cells was significantly smaller than those from the other two cell lines. In addition, assays using T84 produced a more consistent signal than those using HCT 116 (Fig. 2A). Therefore, T84 cells were chosen as the K_Ca3.1-expressing model for Tl⁺ flux assay. Next, the kinetics of the fluorescence signal was investigated. As depicted in Figure 2B , in the absence of reporter dye, a very low level of signal was detected, which was not changed by addition of either Tl⁺ or NS309. In contrast, with dye loading, the fluorescence signal was steadily increased after Tl⁺ addition. Interestingly, NS309 treatment caused a sharp initial increase in fluorescence intensity, which eventually returned to a steady slow rate similar to basal Tl⁺ flux without the addition of K_Ca3.1 activator. These results indicated that the detected signal from the fluorescence indicator corresponds to Tl⁺ flux, and the initial rate of an increase in fluorescence intensity reflects K_Ca3.1 channel activity. To assess the effect of DMSO, which was used as a solvent for test compounds, on assay validity, the Tl⁺ flux assay was performed under various concentrations of DMSO. We found that up to 2% DMSO had no effect on NS309-induced Tl⁺ flux in T84 cells (Fig. 2C).

Figure 2.

Tl⁺ flux assay development. (A) Robustness of signal stimulated with NS309 (10 µM) in different intestinal epithelial cell lines. The Tl⁺ flux assay was performed in intestinal cell lines HT-29, T84, and HCT 116. The normalized maximum fluorescence signal was evaluated 1 min after Tl⁺ addition. Data are displayed as mean ± SEM (n = 5). (B) Representatives of background and assay reporter fluorescence kinetics in T84 cells. The NS309-inducible signal was assessed in the Tl⁺ flux assay with or without K_Ca3.1 activator NS309. Background signal was determined by performing the Tl⁺ flux assay without the dye-loading step. (C) Solvent tolerance of the NS309 signal. The Tl⁺ flux assay was performed with varying concentrations of DMSO from 0.125% to 2% in T84 cells. Data are displayed as mean ± SEM (n = 5). ns, not significant. ***P < 0.001.

Validation and Optimization of Tl⁺ Flux Assay

Since NS309 was previously reported to activate both K_Ca3.1 and small-conductance K_Ca2 channels and Tl⁺ flux can occur through K⁺ channels other than K_Ca3.1,^30,31 the assay specificity to K_Ca3.1 activity was determined. A panel of K⁺ channel inhibitors was used to identify ion channels contributing to NS309-induced Tl⁺ flux. We found that Tl⁺ flux was sensitive to only TRAM-34 (K_Ca3.1 inhibitor) and ouabain (Na⁺/K⁺-ATPase inhibitor), but not apamin (K_Ca2 inhibitor), BaCl₂ (K_cAMP inhibitor), or bumetanide (Na⁺-K⁺-2Cl⁻ co-transporter inhibitor), as shown in Figure 3A . These findings indicate that Tl⁺ flux was mainly mediated by K_Ca3.1 and the constitutively active Na⁺/K⁺-ATPase. The contribution of K_Ca3.1 and K_Ca2 to the NS309-stimulated signal was also evaluated using the electrophysiological method. We found that the NS309-induced I_K⁺ was sensitive to TRAM-34 but not apamin, confirming that K_Ca2 did not contribute to the NS309-induced K⁺ flux in T84 cells (Fig. 3B).

Figure 3.

Specificity and optimization of screening assay. (A) Responsiveness of NS309-induced signal to inhibitors of K_Ca3.1 (TRAM-34, 30 µM), K_Ca2 (apamin, 1 µM), K_ir (BaCl₂, 5 mM), Na⁺/K⁺-ATPase (ouabain, 1 mM), and NKCC (bumetanide, 100 µM) channels in the Tl⁺ flux assay (n = 9). (B) Validation of NS309-induced current specificity. Basolateral I_K⁺ in the T84 cell monolayer was measured with the Ussing chamber electrophysiological method. NS309 (10 µM), apamin, and TRAM-34 were added to both apical and basolateral chambers. A representative I_K⁺ tracing is shown (n = 3). (C) Effect of NS309 incubation time on Tl⁺ flux. Tl⁺ ion addition and fluorescence kinetics measurement were performed immediately (0 min) and 30 or 60 min after NS309 (10 µM) was added to the cells (n = 12). (D) Relationship between NS309 concentration and fluorescent signal kinetics. The Tl⁺ flux assay was performed with varying concentrations of NS309 (n = 3). (E) Relationship between TRAM-34 concentration and fluorescent signal kinetics. The Tl⁺ flux assay was performed with 10 µM NS309 and varying concentrations of TRAM-34 (n = 6). (F) Concentration–response relationship of TRAM-34 in the Tl⁺ flux assay (n = 4) and electrophysiological I_K⁺ analysis (n = 5) under stimulation of 10 µM NS309. Data are displayed as mean ± SEM. ns, not significant. *P < 0.05, **P < 0.01, ***P < 0.001 when compared with control. ^###P < 0.001 when compared with NS309 group.

Subsequently, experimental parameters of our assay were optimized to achieve the best fluorescence signal for the screening. First, the optimal incubation time of NS309 activator was determined. Prolonged NS309 incubation time before kinetics measurement up to 60 min did not affect the Tl⁺ flux rate (Fig. 3C). Therefore, NS309 was included in the stimulus buffer to minimize the steps and time of assays. Then, we investigated the optimal concentration of NS309 by varying its concentration from 0.5 µM to 30 µM. We found that NS309 significantly stimulated Tl⁺ flux to a similar rate at concentrations ranging from 1 to 10 µM (Fig. 3D). In addition, using NS309 at 10 µM and TRAM-34 from 10 µM to 50 µM treatment significantly reduced Tl⁺ flux, with the maximal effect being observed at 30 µM (Fig. 3E). Therefore, we used NS309 at 10 µM and TRAM-34 at 30 µM (positive control) for the analysis of assay quality. To evaluate whether the assay was capable of producing quantitative data of K_Ca3.1 activity comparable to the data from the electrophysiological method, the effect of TRAM-34 at varying concentrations from 0.01 µM to 30 µM in the presence of 10 µM NS309 was assessed between both methods. Concentration–response analysis of TRAM-34 yielded IC₅₀ values of 0.125 ± 0.03 µM in the Tl⁺ flux assay and 0.129 ± 0.01 µM in electrophysiological analysis, indicating the assay’s capability for quantitative analysis of K_Ca3.1 inhibition, consistent with the electrophysiology data.

Evaluation of Assay Quality

To verify the ability of the assay to differentiate the inhibitor control group from the activator control group, changes in fluorescence signal were compared between both groups. Similar to Figure 2B , NS309 produced an initial sharp rise in fluorescence signal, while co-treatment with TRAM-34 only allowed a steady slow increase in the signal (Fig. 4A), demonstrating that both control groups displayed distinct fluorescence kinetics. Moreover, the NS309-treated group showed significantly higher Tl⁺ flux than the inhibitor-treated group (Fig. 4B), suggesting an ability of our assay to detect the difference in K_Ca3.1 activity with or without the K_Ca3.1 inhibitor TRAM-34. In addition, there was no difference in data obtained from the beginning, middle, and end of the assay in each plate, indicating the robustness of signal across the plate. However, there was still a considerable variation in the data of each control group, which may be a result of plate-to-plate variation. To minimize this inconsistency between each round of experiment, the data from each plate were normalized by the means of both control groups from that plate using eq 1. Transformed data still retained the same level of significant difference, but with lower variation within each group (Fig. 4C). The quality of the assay was assessed by calculation of the Z′ factor as in eq 2. Our Tl⁺ flux assay yielded a Z′ factor of 0.52 ± 0.08 from seven separate experiments after data normalization, indicating an excellent screening assay quality.

Figure 4.

Evaluation of assay quality. (A) Representative plots of activator and inhibitor controls. The Tl⁺ flux assay was performed with NS309 as a control for active K_Ca3.1 function, and with co-treatment of NS309 plus TRAM-34 for the inhibited K_Ca3.1 channel. (B) Data distribution of controls displayed as individual data of each well with mean ± SD (n = 55/group). (C) Data distribution of controls after transformation with mean ± SD (n = 55/group). ***p < 0.001.

Screening of the Compound Library

To evaluate the practicality of our assay, the Tl⁺ flux assay was used to identify potential K_Ca3.1 inhibitors from the small set of the compound library. The screening library contained 1352 samples of 679 isolated natural compounds, 273 synthetic derivatives, and 400 crude extracts from various plants and fungi in Thailand. Compounds in the library were prepared as 100× working stock at 5 mM in DMSO. Primary screening with Tl⁺ flux assay was performed at a concentration of 50 µM with a final DMSO concentration of 1.2%, using criteria of more than 80% Tl⁺ flux inhibition for compound selection. A total of 23 compounds passed the selection criteria of the Tl⁺ flux assay. Next, K_Ca3.1 inhibitory properties of compounds obtained from screening were validated by measuring the effect of the compounds on basolateral I_K⁺ stimulated by NS309. A natural compound, resveratrol, was previously reported to inhibit K_Ca3.1 with an IC₅₀ of approximately 10 µM,³² which is a relatively low potency compared with other small-molecule K_Ca3.1 inhibitors with submicromolar IC₅₀.²² Therefore, a threshold of at least 50% inhibition of K_Ca3.1-mediated I_K+ at 10 µM was used to identify effective inhibitor compounds and to exclude molecules with low potency. With this, we verified eight inhibitors of the K_Ca3.1 channel to undergo a more detailed concentration–response relationship analysis, summarized in Table 1 . Examples of a concentration–response relationship of compounds N15, N18, and N602 are demonstrated in Figure 5A . Notably, we have identified several compounds with submicromolar IC₅₀ (N134, S267, and S269) and others with low micromolar potency (IC₅₀ ~ 1–6 µM). The preliminary evaluation of potential nonspecific inhibitory effects was also performed. All eight K_Ca3.1 inhibitor compounds did not significantly alter basal Tl⁺ flux without NS309 stimulation, indicating a negligible effect on other constitutive K⁺-permeable channels in T84 cells (data not shown).

Table 1.

Summary of Identified K_Ca3.1 Inhibitors.^a

Compound Code	Maximum I_K⁺ Inhibition	IC₅₀ (µM)
N15	94.80% at 10 µM	3.34 ± 1.37
N18	92.27% at 30 µM	6.57 ± 1.56
N134	87.40% at 5 µM	0.14 ± 0.07
N602	92.94% at 10 µM	3.84 ± 0.96
N689	85.08% at 5 µM	1.05 ± 0.34
S267	94.84% at 10 µM	0.54 ± 0.10
S269	90.95% at 10 µM	0.28 ± 0.21
S270	98.92% at 10 µM	1.02 ± 0.07

The maximum I_K⁺ inhibition and IC₅₀ values of K_Ca3.1 inhibitor compounds were obtained from concentration–response analysis by the electrophysiological method. Data were fitted to the Hill equation. IC₅₀ values are displayed as mean ± SEM (n = 3–4).

Figure 5.

Examples of concentration–response data. (A) Concentration–response relationship of N15, N18, and N602 (n = 3). Basolateral I_K⁺ in the T84 cell monolayer was obtained from Ussing chamber analysis. NS309 (10 µM) was used to stimulate the KCa3.1 current. All compounds were added to both apical and basolateral chambers. Data were fitted to the Hill equation and are displayed as mean ± SEM.

Discussion

To establish a screening assay targeting the K_Ca3.1 channel, numerous approaches were considered as a basis for its methodology. The nonfunctional ligand-binding assay was excluded since molecular binding does not always confer functional modulation. On the other hand, highly sensitive direct electrophysiological assays were not suitable due to the necessity of specialized equipment and operation skills. Therefore, a fluorescence flux-based Tl⁺ flux assay was used because it requires only a standard fluorescence plate reader and because of its optimizable components.

During the selection of suitable cell lines, we have observed differential functional expression of K_Ca3.1 between cell lines. Interestingly, HCT 116 cells showed noticeably higher signal variability between wells, possibly due to greater heterogeneity in cell population compared with the other two cell lines. While it is ideal to activate K_Ca3.1 via a native mechanism using Ca²⁺, this method of activation is not specific to K_Ca3.1 channels. A rise in intracellular Ca²⁺ would activate not only K_Ca3.1 but also other Ca²⁺-activated channels, like large-conductance K_Ca1/BK. Moreover, NS309 acts by increasing the opening probability of K_Ca3.1 at a given Ca²⁺ concentration,³³ allowing the basal intracellular Ca²⁺ level to stimulate the channels. Therefore, the activation of K_Ca3.1 by NS309 is still dependent on stimulation by Ca²⁺. Another thing of note is that even though NS309 could activate both intermediate-conductance K_Ca3.1 and small-conductance K_Ca2 channels through interaction with their shared calmodulin-binding domain required in intracellular Ca²⁺ sensing,²² we found that NS309 activated only K_Ca3.1 in T84 cells during both the Tl⁺ flux assay and electrophysiological measurement. We hypothesized that T84 cells might express a very low level of K_Ca2 channels, which therefore contribute very little to NS309-stimulated signal. We also found that Tl⁺ flux was significantly reduced upon administration of ouabain, implicating a role of Na⁺/K⁺-ATPase in mediating Tl⁺ flux. The main function of Na⁺/K⁺-ATPase is the active transport of Na⁺ efflux and K⁺ influx. Since Tl⁺ could be transported via any K⁺-permeable transporters, Na⁺/K⁺-ATPase would uptake Tl⁺ into the cell as well, a phenomenon that has been previously reported in myocardial cells.³⁰ In addition, Na⁺/K⁺-ATPase is responsible for maintaining negative resting membrane potential, and therefore its function would be important in providing the electrical driving force for the entry of positively charged Tl⁺ into the cells.

Unexpectedly, prolonged exposure or excessive concentration of the K_Ca3.1 activator NS309 apparently diminished Tl⁺ flux, an effect not seen with the K_Ca3.1 inhibitor TRAM-34. This might be due to a desensitization mechanism of the K_Ca3.1 channel, which thereby prevents overactivation by NS309. Moreover, TRAM-34 showed considerably lower potency (IC₅₀ = 129 nM) in our electrophysiological analysis than previously reported in other publications (IC₅₀ ~ 10–25 nM).²² However, it is important to note that these studies often use different methodologies or cell models to determine the potency of TRAM-34, which might contribute to this discrepancy. The fluorescence signal pattern elicited by NS309 showed a rapid increase that slowed down as time went on, corresponding to the driving force for Tl⁺ flux from the concentration gradient. The difference in [Tl⁺] would be highest immediately after Tl⁺ addition into the extracellular buffer, resulting in a strong initial Tl⁺ flux. With the passage of time, Tl⁺ influx would gradually increase intracellular [Tl⁺] and reach equilibrium, where the concentration gradient of Tl⁺ was diminished, eliminating the driving force of the Tl⁺ flux. This stage would manifest as late steady state of the signal, where the signal retained a constant rate of change, comparable to those without NS309 stimulation or with K_Ca3.1 inhibited by TRAM-34. Since Na⁺/K⁺-ATPase was shown to contribute to Tl⁺ flux, we hypothesized that this residual baseline Tl⁺ flux might result from Na⁺/K⁺-ATPase-mediated active transport of Tl⁺. Nevertheless, it is important to emphasize that our experimental design did not exclude the possibility of saturation of the fluorescent reporter dye or precipitation of excess Tl⁺ with intracellular Cl⁻ as the cause of signal plateau.

Our Tl⁺ flux screening assay had identified several new K_Ca3.1 inhibitors validated by electrophysiological measurements from the natural compound library, demonstrating the effectiveness of a screening assay based on endogenous expression of the K_Ca3.1 channel in the detection of K_Ca3.1 inhibitors. A total of 23 compounds from a library of 1352 compounds passed the selection criteria of the Tl⁺ flux assay, and only 8 compounds were classified as K_Ca3.1 inhibitors after validation by electrophysiology. Therefore, the initial hit rate and validated hit rate for our assay were 1.7% and 0.6%, respectively. Among eight discovered compounds, only the bioactivities of N15 and N602 have been previously reported. A depsidone N15 from marine-derived fungus has been shown to inhibit aromatase and xanthone oxidase enzymes³⁴ and reduce lipopolysaccharide (LPS)-induced nitric oxide production in vitro.³⁵ A flavonoid N602 was also reported to have antifungal,³⁶ antimicrobial,³⁷ and anticancer^38,39 effects. However, none of the eight compounds have been described for their inhibitory effect on the K_Ca3.1 channel. In addition, none of the identified K_Ca3.1 inhibitors shared structural similarity with well-known inhibitor scaffolds: triaryl-methanes, phenyl-pyrans, and cyclohexadienes.²² According to the structural similarity, these compounds could be loosely categorized into four groups: N15 and N18; N134 and N602; N689; and S267, S269, and S270. Among compounds S267, S269, and S270, an amine-attached cyclic hydrocarbon chain seems to greatly affect the potency of the compound in this family. Introduction of nitrogen atom into the end of a benzene ring increased the potency of S269 almost twofold compared with S267. Substitution of a benzyl group in S267 to cyclohexenyl ethyl in S270 reduced its potency by half. Nevertheless, further comprehensive analysis of the structure–activity relationship is required to ascertain these observations. It is important to note that, while we have confirmed the effects of these compounds in K_Ca3.1-mediated I_K⁺ inhibition, further studies are needed to determine whether they inhibit K_Ca3.1 via direct interaction with the channel or inhibition of K_Ca3.1 was a secondary effect from interaction with other target molecules within the regulatory system of the K_Ca3.1 channel. For instance, the compounds could modulate the function of nucleoside diphosphate kinase B (NDPK-B) and mammalian protein histidine phosphatase (PHPT1), which regulate phosphorylation at the histidine 358 residue of K_Ca3.1 that is required for channel activation.^40,41 In addition, a test of specificity against related channels within the K_Ca family and other critical ion channels such as the cardiac human ether-à-gogo-related gene (hERG) is still needed to ensure the specificity and safety of these K_Ca3.1 inhibitors. Moreover, detailed pharmacokinetics and pharmacodynamics data of these compounds are necessary for further in vivo testing in disease models to progress further toward therapeutic agents in humans.

In summary, a high-throughput screening assay targeting the K_Ca3.1 channel inhibitor was successfully established based on the Tl⁺ flux assay in the T84 intestinal epithelial cell line. This assay was capable of identifying potent K_Ca3.1 inhibitors verified by the electrophysiological method from the natural compound library. Further refinement and upscaling of our assay may result in a streamlined high-throughput screening process for rapid drug discovery. Further study of these newly discovered inhibitory compounds might lead to a new class of K_Ca3.1 inhibitors applicable for human use.

Footnotes

Declaration of Conflicting Interests

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding

The authors received the following financial support for the research, authorship, and/or publication of this article: This work was supported by the Thailand Research Fund (DBG5980001, DBG6180029); Faculty of Science, Mahidol University; and Medical Scholars Program, Mahidol University.

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Development of a Cell-Based Assay for Identifying K Ca 3.1 Inhibitors Using Intestinal Epithelial Cell Lines

Abstract

Keywords

Introduction

Materials and Methods

Chemical Reagents

Cell Culture

Thallium Flux Assay

Measurement of Basolateral K+ Current (IK+)

Statistical Analysis

Results

Development of a Screening Assay Based on the Tl+ Flux Assay

Validation and Optimization of Tl+ Flux Assay

Evaluation of Assay Quality

Screening of the Compound Library

Discussion

Footnotes

Declaration of Conflicting Interests

Funding

References

Measurement of Basolateral K⁺ Current (I_K⁺)

Development of a Screening Assay Based on the Tl⁺ Flux Assay

Validation and Optimization of Tl⁺ Flux Assay