Synthesis of Small‐Molecule Fluorescent Probes for the In Vitro Imaging of Calcium‐Activated Potassium Channel K_Ca3.1

Abstract

Small‐molecule probes for the in vitro imaging of K_Ca3.1 channel‐expressing cells were developed. Senicapoc, showing high affinity and selectivity for the K_Ca3.1 channels, was chosen as the targeting component. BODIPY dyes 15–20 were synthesized and connected by a Cu^I‐catalyzed azide–alkyne [3+2]cycloaddition with propargyl ether senicapoc derivative 8, yielding fluorescently labeled ligands 21–26. The dimethylpyrrole‐based imaging probes 25 and 26 allow staining of K_Ca3.1 channels in NSCLC cells. The specificity was shown by removing the punctate staining pattern by pre‐incubation with senicapoc. The density of K_Ca3.1 channels detected with 25 and by immunostaining was identical. The punctate structure of the labeled channels could also be observed in living cells. Molecular modeling showed binding of the senicapoc‐targeting component towards the binding site within the ion channel and orientation of the linker with the dye along the inner surface of the ion channel.

Imaging channels: Small‐molecule probes for the in vitro imaging of K_Ca3.1 channel‐expressing cells were designed and developed. Using senicapoc as the targeting component, the BODIPY‐labeled probes could selectively stain the channels in living cells and the channel punctate structure could be observed.

Introduction

Ion channels play a critical role in the progression of cancer and contribute to features of essentially all “cancer hallmarks”.1 The calcium‐activated potassium channel 3.1 (K_Ca3.1) is an intensively studied ion channel in this context. It is involved in critical steps of the metastatic cascade, such as proliferation, migration, invasion, and extravasation.2, 3 Inhibition of the K_Ca3.1 channel leads in many different tumor entities to reduced proliferation, migration, and metastasis.4, 5, 6 Overexpression of this channel directly correlates with tumor grade and metastatic status and is often related to poor prognosis for tumor patients and high lethality rates.7 Moreover, expression of K_Ca3.1 channels is dysregulated in many tumor entities. When patients are stratified according to K_Ca3.1 channel expression, patient survival is worse for those with elevated K_Ca3.1 expression. This points to the predictive power of analyzing K_Ca3.1 channel expression with respect to patient survival.8, 9

Therefore, developing tools for the visualization of K_Ca3.1‐expressing cells would be a great asset for a better understanding of its (diagnostic) role and predictive value in cancer. Currently, only indirect immunoflourescence staining is available for optical imaging of the K_Ca3.1 channel, making the process time‐consuming and expensive. Herein, we present the development and synthesis of novel small‐molecule probes, which show promising results in in vitro imaging of the K_Ca3.1‐expressing cells by fluorescence microscopy. Simple, fast, and efficient staining protocols are the advantages of these novel imaging probes, which can be instrumental for elucidating the mechanisms by which K_Ca3.1 channels contribute to cancer progression and metastasis.

Results and Discussion

Synthesis of the Probes

For the development of small‐molecule fluorescently labeled probes of the K_Ca3.1 channel, senicapoc (1) served as starting point. Senicapoc (1) has a high affinity towards the K_Ca3.1 channel in the low‐nanomolar range (IC₅₀=11±2 nm; measured on human erythrocytes) and excellent selectivity over related ion channels.10 In order to introduce a fluorescent dye one F‐atom of senicapoc should be replaced by a hydroxy moiety. The OH moiety of the senicapoc derivative 2 should allow broad chemical modifications including the introduction of a fluorescent dye (Figure 1).

Figure 1.

Chemical structure of senicapoc (1) and its derivative 2.

The phenol 2 was obtained in five steps starting with the protection of 4‐bromophenol (3) with tert‐butyldiphenylsilyl chloride (TBDPS‐Cl). The protected 4‐bromophenol 4 was reacted with n‐butyllithium in THF to obtain the lithiated intermediate, which was trapped with 4‐fluorobenzophenone yielding the tertiary alcohol 5. Trimethylsilyl cyanide and catalytic amounts of InCl₃ were used to convert the alcohol 5 into the nitrile 6. The silyl protective group was cleaved under basic conditions resulting in the intermediate 7 with a free hydroxy group in p‐position. In the last step, the nitrile was hydrolyzed partially under basic conditions to obtain the senicapoc derivative 2. Although the partial hydrolysis of the nitrile to an amide has been described for the synthesis of senicapoc in high yields, in case of the phenol 7 the transformation was rather low and incomplete resulting in a yield of the amide 2 of 23 %. However, a large amount of the starting material 7 could be re‐isolated, which increased the efficiency of the last reaction step considerably (Scheme 1).

Scheme 1.

Synthesis of the senicapoc derivative 8. Reagents and reaction conditions: a) imidazole, TBDPS‐Cl, DMF, 24 h, rt, 96 %. b) 1. n‐ butyllithium, THF, 1 h, −78 °C; 2. 4‐fluorobenzophenone, THF, 3 h, rt, 58 %. c) 1. InCl₃, trimethylsilyl cyanide, CH₂Cl_2, 1 h, 20–40 °C, 1.5 h, 40–50 °C; 2. KOH, KNa‐tartrate, H₂O, 12 h, 0 °C—rt, 89 %. d) KOH, KNa‐tartrate, 2‐methylbutan‐2‐ol,5 h, 100 °C, 88 %. e) KOH, EtOH, H₂O, 90 °C, 48 h, 23 %, f) propargyl bromide, Cs₂Co₃, DMF, 30 min, rt, 90 %.

For the introduction of the fluorescent label a Cu^I‐catalyzed azide–alkyne [3+2]cycloaddition was envisaged. Therefore, an alkyne moiety had to be introduced into the phenol 2. For this purpose, phenol 2 was alkylated with propargyl bromide in the presence of Cs₂CO₃ to afford the propargyl ether 8 (Scheme 1).

In order to visualize the senicapoc‐derived targeting component by the introduction of a fluorescent dye, different boron dipyrromethene (BODIPY) dyes were designed bearing an azide moiety suitable for the planned [3+2]cycloaddition reaction with the alkyne 8.

The photochemical properties of the BODIPY dyes are easily adjustable by variation of the substitution pattern of the dipyrromethene core.11, 12 In terms of medicinal research, the BODIPY dyes show some important advantages; namely, the emission intensity derived from high quantum yields is almost independent from the pH value and the polarity of the solvent. Moreover, low toxicity was reported. On the other hand, low solubility of some dyes in water and physiological water‐based buffer systems may cause problems concerning their application in biological systems.

For the synthesis of BODIPY‐based dyes, a general one‐pot three‐step approach was pursued, which includes the acid catalyzed condensation of two equivalents of a pyrrole with one equivalent of a benzaldehyde followed by oxidation of the dipyrrolylmethane with 2,3,5,6‐tetrachloro‐1,4‐benzoquinone (p‐chloranil). Then, the oxidized product was deprotonated with a base and complexed with BF₃⋅OEt₂ to give the BODIPY dye13 (Scheme 2).

Scheme 2.

General concept for the synthesis of BODIPY dyes. a) H⁺. b) oxidant, for example, p‐chloranil. c) NEt₃, BF₃⋅OEt₂. (modified according to literature13).

Due to the limited conjugation with the dipyrrolomethyne core, the phenyl moiety has only minor effects on the excitation and emission wavelength of the resulting BODIPY dye. Herein, benzaldehyde derivatives were used bearing an azide moiety within the substituent in p‐position. This azido moiety should be used for the connection of the BODIPY dye with the targeting component through a Cu^I‐catalyzed azide–alkyne [3+2]cycloaddition. Consequently, the benzaldehyde derivatives contain the linker to the targeting component. Benzaldehyde derivatives with different lengths and polarity of the linker, that is, an ethoxy‐ (9), a 3,6,9‐trioxaundecyloxy (10) and a dodecyloxy linker (11) between the phenyl ring and the azido moiety were synthesized as described in literature13, 14, 15 and further used for the synthesis of the BODIPY dyes.

The dipyrromethene core, which defines the optical properties of the dye, can be varied by starting from different pyrrole derivatives. An enlarged conjugated π‐electron system resulting from annulation of an aromatic ring at the pyrrole ring shifts the emission wavelength towards the near‐infrared region. To achieve this effect, the annulated thienopyrrole 12 was reacted with benzaldehyde 9 according to the general one‐pot procedure outlined in Scheme 3 to obtain BODIPY dye 15. Due to stability issues with 15, containing the thienopyrrole system, alkyl substituents were taken into consideration. In general, most of the reactive pyrrole positions should be blocked by suitable substituents to avoid side reactions during the condensation with benzaldehydes and to enhance the stability of the final dye. Therefore, 3‐ethyl‐2,4‐dimethylpyrrole (13) was reacted with the benzaldehydes 9–11 to provide the stable and storable BODIPY dyes 16–18. In order to reduce the size and lipophilicity of the dyes, the reaction of 2,4‐dimethylpyrrole (14) with benzaldehydes 9 and 10 was carried out to afford two additional stable and storable BODIPY dyes 19 and 20.

Scheme 3.

A) Structures of benzaldehyde derivatives 9–11 and pyrrole derivatives 12–14 used for the BODIPY synthesis. B–D) Synthesis of BODIPY dyes with azide moiety in the side chain. Series (B) is based on the thiophene annulated pyrrole 12, series (C) is based on the ethyldimethylpyrrole 13 and series (D) on the dimethylpyrrole 14. Reagents and reaction conditions: a) 1. TFA, CH₂Cl₂, molecular sieves 3 Å, rt, 24 h; 2. p‐chloranil, CH₂Cl₂, rt, 30 min; 3. NEt₃, CH₂Cl₂, rt, 15 min; 4. BF₃⋅OEt₂, CH₂Cl₂, rt, 12 h. Yields: 15: 7 %. 16: 14 %. 17: 30 %. 18: 22 %. 19: 16 %. 20: 27 %. Yields were calculated over three reaction steps for >90 % purity compounds.

Since fluorescence microscopy should be performed with the designed fluorescently labeled ligands, the synthesized dyes need to be suitable for this imaging technique. Due to lack of conjugation with the dipyrrolomethyne core, electronic effects were not expected regarding the photophysical properties in relation to the conjugation of the phenyl moiety, its substituents, or the targeting component. Therefore, the optical characteristics of the six BODIPY dyes 15–20 were analyzed before attachment of the targeting component. High quantum yields are desirable. Excitation and emission wavelengths suitable for commercially available filter sets (FITC, TRITC) are necessary. The photophysical properties of the BODIPY dyes 15–20 are summarized in Tables 1–3 and discussed thereafter (see below).

Table 1.

Absorption and emission maxima of the dyes 15–20.

compd.	λ abs, max [nm][a]	λ em, max [nm][b]	λ abs, max [nm][c]	λ em, max [nm][d]
15	553	571	559	574
16	521	533	524	537
17	521	533	524	537
18	521	532	524	535
19	497	507	500	510
20	497	508	500	510

[a]  Absorption in CH₃CN. [b] Emission in CH₃CN. [c] Absorption in CH₂Cl₂/CH₃OH. [d] Emission in CH₂Cl₂/CH₃OH.

Table 3.

Photoluminescence quantum yields (Φ _F), fluorescence lifetimes (τ _F), fluorescence (k _F) and non‐radiative (k _nr) rate constants in CH₂Cl₂/CH₃OH (1:1) of compounds 15–20.

compd.	Φ F [a], [b]	τ F [ns][b]	k F [×106  s−1][b]	k nr [×106  s−1][b]	Φ F [c], [d]	τ F [ns][d]	k F [×106  s−1][d]	k nr [×106  s−1][d]
15	0.05	0.818±0.004	61±30	1160±40	0.1	1.252±0.002	<160	720±80
16	0.73	6.00±0.02	122±4	45±5	0.9	5.99±0.02	150±20	<30
17	0.63	6.54±0.03	96±4	57±5	0.9	5.46±0.02	160±20	<40
18	0.64	5.83±0.03	110±4	62±6	0.9	5.6±0.2	160±20	<40
19	0.41	3.810±0.016	108±6	154±8	0.9	5.06±0.02	180±20	<40
20	0.44	4.408±0.018	100±5	127±8	0.9	5.8±0.2	160±20	<30

[a] (±0.02). [b] 298 K. [c] (±0.1). [d] 77 K.

The wavelengths of the FITC and TRITC filter sets are as follows: λ _excitation(FITC)=450–490 nm, λ _emission(FITC)=515–565 nm; and λ _excitation(TRITC)=546/12 nm (bandpass filter), λ _emission(TRITC)=590 nm (longpass filter). Thus, the absorption and emission wavelengths of BODIPY dyes 16–20 fit to the FITC filter set. The TRITC filter set can be used for fluorescence microscopy of BODIPY 15.

The solvatochromic shift in the absorption and in the emission maxima is modest for all dyes when increasing the polarity of the solvent, which points to a predominant ππ* character of the excited states that also accounts for the small Stokes shifts and the vibrational shoulders observed in all cases (see Table 1 and the corresponding spectra in the Supporting Information).

However, the absorption and the emission spectra both in CH₃CN and in the more polar CH₂Cl₂/CH₃OH mixture clearly show a bathochromic shift for compound 15 and a blue‐shift for 19 and 20 compared with 16–18, indicating that the energy of the excited states can be fine‐tuned by judicious choice of the substitution pattern. In particular, the fused thiophene rings extend the delocalization of the π system, causing the observed red‐shifted absorption and emission maxima. Interestingly, the fluorescence rate constants in CH₃CN are comparable for all exemplars except for compound 15, which is significantly smaller (Table 2).

Table 2.

Photoluminescence quantum yields (Φ _F), fluorescence lifetimes (τ _F), fluorescence (k _F) and non‐radiative (k _nr) rate constants in fluid CH₃CN at 298 K of compounds 15–20.

compd.	Φ F	τ F [ns]	k F [×106  s−1]	k nr [×106  s−1]
15	0.04	0.891±0.006	40±20	1077±30
16	0.81	6.32±0.05	128±4	30±5
17	0.81	6.37±0.05	127±4	30±5
18	0.82	6.60±0.06	124±4	27±6
19	0.53	3.96±0.03	134±6	120±8
20	0.50	4.09±0.03	122±6	122±8

[a] (±0.02)

Moreover, the radiationless deactivation rates are drastically higher for the latter and significantly enhanced for 19 and 20, if compared with 16–18. This is surprising since 16–18 have additional side chains providing extra degrees of freedom and therefore a higher density of rotovibrational states that could favor non‐radiative relaxation pathways. This points to radiationless transitions that are mostly governed by electronic effects associated to the variable substitution patterns. The latter is evident for 19–20 but particularly for 15, as can be also observed in the more polar CH₂Cl₂/CH₃OH mixture in which all cases the radiative relaxation rates are slightly slower than in CH₃CN while the radiationless processes are somewhat faster than in the less polar environment (Table 3). Interestingly, in frozen matrices at 77 K, all dyes show practically identically high radiative rate constants, whereas the radiationless process is only (yet drastically) boosted for 15. This observation points to an enhanced intersystem crossing that could be ascribed to the additional lone pairs from the sulfur atoms at the thiophene rings (El Syed rule). It is also evident that the substitution at the methyne bridge has no significant effect on the excited state energies and their relaxation rate constants.

In order to obtain the desired fluorescently labeled ligands, the synthesized BODIPY dyes 15–20 were coupled with the alkyne 8 by a Cu^I‐catalyzed azide–alkyne [3+2]cycloaddition reaction (Scheme 4).

Scheme 4.

Synthesis of fluorescently labeled tool compounds. Reagents and reaction conditions: a) CuSO₄, sodium ascorbate, DMF, H₂O, rt, 24 h, 21: 58 %, 22: 57 %, 23: 61 %, 24: 2 %, 25: 19 %, 26: 68 %.

The six fluorescently labeled ligands 21–26 shown in Figure 2 were tested in first simple staining experiments to analyze their suitability for in vitro imaging of the K_Ca3.1 channel.

Figure 2.

Fluorescently labeled ligands 21–26 designed to image K_Ca3.1 channels.

In Vitro Imaging of NSCLC Cells

The A549 cell line, used for all experiments, is a hypotriploid epithelial cell line from a non‐small‐cell lung cancer (NSCLC) taken from a 58‐year‐old Caucasian in 1972. The suffix “3R” refers to the repeated process of intravenous administration of parental cells (also initially “0R”) into immunocompromised mice to obtain lung metastases. Cells from these metastases were isolated and cultured in vitro followed by another round of reinjection to select for tumor cells with high metastatic potential.16

A549‐3R cells overexpress the K_Ca3.1 channel.7 Thus, they were selected for testing of the novel small‐molecule imaging probes. We aim at the development of a fast and simple staining protocol. Therefore, a drop (30 μL) of a previously prepared staining solution (10 μm in PBS) of the respective fluorescently labeled ligand was pipetted onto Parafilm. The cover slip with the adherent cells was placed carefully upside down onto the drop and the NSCLC cells were incubated in a humidified dark chamber for 10 min (Figure 3).

Figure 3.

Setup for the staining of NSCLC cells.

The NSCLC cells are ready for microscopy after only 10 min incubation time and subsequent washing with PBS‐buffer. This newly developed protocol using the fluorescently labeled probes is very fast and simple in comparison to the much more complicated indirect immunofluorescence assay (ca. 4 h). An inverted fluorescence microscope was used to image NSCLC cells labeled with the fluorescent senicapoc derivative.

After performing the staining protocol with the fluorescently labeled thiophene annulated ligand 21, staining of the A549‐3R cells could not be observed. To ensure that this result is not due to the low quantum yield of BODIPY 15, a drop of the staining solution of 21 was examined under the microscope and an orange‐red color was observed. Thus, imaging probe 21 is not suitable for the visualization of the K_Ca3.1 channels.

The same staining protocol was performed with the fluorescently labeled ligands 22 and 23 bearing an ethyl and two methyl moieties at the pyrrole rings and the NSCLC cells were observed. Unfortunately, the staining was not specific and no staining pattern of the K_Ca3.1 channels was found (see Figure S1 in the Supporting Information).

Problems occurred while preparing the staining solution of fluorescently labeled ligand 24 with the undecyloxy linker. Due to its high lipophilicity, it was not possible to dilute the DMSO stock solution with PBS‐buffer. The molecule precipitated immediately and neither ultrasound nor heating of the suspension led to a clear solution. Thus, it was not possible to perform the staining experiments with imaging probe 24.

In contrast, the fluorescently labeled dimethylpyrrole‐based ligands 25 and 26 showed promising results. In both cases the typical punctate staining pattern of the K_Ca3.1 channel was observed7, 17 (Figure 4 A–D).

Figure 4.

A) NSCLC cells incubated for 10 min with a 10 μm staining solution of imaging probe 25. B) Magnification of (A) (white box). C) NSCLC cells incubated for 10 min with a 10 μm staining solution of imaging probe 26. D) Magnification of (C) (white box). E) NSCLC cells blocked for 5 min with unlabeled senicapoc (30 μm) and subsequently stained for 10 min with a 10 μm staining solution of imaging probe 25. F) NSCLC cells blocked for 5 min with unlabeled senicapoc (30 μm) and subsequently for 10 min with a 10 μm staining solution of imaging probe 26.

As a proof of principle, the NSCLC cells were pre‐incubated with unlabeled senicapoc (30 μm) 5 min before staining. Consequently, the typical punctate staining pattern could not be observed any more (Figure 4 E,F).

Since all the binding regions were blocked with senicapoc, the fluorescently labeled probes 25 and 26 were no longer able to bind specifically to the K_Ca3.1 channel. This experiment indicates that the developed fluorescently labeled ligands bind to the same region of the K_Ca3.1 channel as senicapoc does. Therefore, specific binding of the new probes to the K_Ca3.1 channel was confirmed. Thus, the two novel imaging probes 25 and 26 are suitable for fast and simple visualization of the K_Ca3.1 channels, which appear as bright spots in the fluorescence images.

Since the best results were obtained using the fluorescently labeled ligand 25 (best signal‐to‐noise ratio), this imaging probe was further evaluated. In a control experiment, HEK293 cells not expressing the K_Ca3.1 channel were incubated with a 10 μm staining solution of 25 following the same protocol as used for the NSCLC cell‐staining experiments. Staining of the HEK293 cells could not be observed (see Figure S2 in the Supporting Information).

Further staining experiments with the fluorescently labeled ligand 25 in living NSCLC cells showed more background signals owing to the incomplete permeation of the cell membrane. However, the punctate staining pattern of the K_Ca3.1 channel was still visible (see Figure S3 in the Supporting Information).

In order to compare the results obtained by K_Ca3.1 channel imaging with fluorescently labeled ligand 25 with the results obtained by the indirect immunofluorescence assay, both methods were performed and the K_Ca3.1 channel density was determined for each experiment. (Figure 5) Therefore, ten squares with a side length of 50 pixels, corresponding to an area of 9 μm², were randomly located in one cell. Signals were analyzed with a linescan (MetaVue software) and were counted as one K_Ca3.1 channel if the full width at half maximum (FWHM) was ≤5 pixels (ca. 300 nm). In total, 15 cells (150 squares) were analyzed for each staining method.

Figure 5.

Signal analysis with linescans (blue lines in magnification boxes). 1 pixel≈60 nm. A) NSCLC cells incubated for 10 min with a 10 μm staining solution of imaging probe 25 with 50×50 pixel square, magnification of this square with linescan and 16 signals for K_Ca3.1 channels. B) Linescan of (A) Intensity profile [arbitrary units] of signal 1 and signal 2. C) NSCLC cells after performing the indirect immunofluorescence assay with 50×50 pixel square, magnification of this square with linescan and 16 signals for K_Ca3.1 channels. D) Linescan of (C) Intensity profile [arbitrary units] of signal 1.

For both the imaging with fluorescently labeled ligand 25 and the indirect immunofluorescence assay, an average K_Ca3.1 channel density of 1.79 μm⁻² was found (compare Figures S4 and S5 in the Supporting Information). This analysis shows, that the same results can be achieved with 25 compared to the indirect immunofluorescence assay but staining with 25 is more than six‐fold faster and more convenient than performing the indirect immunofluorescence assay.

In order to investigate the binding mode of the fluorescently labeled ligand 25, molecular modelling of 25 in the available K_Ca3.1 Cryo‐EM structure (pdb 6cn0)18 was performed (Figure 6).

Figure 6.

A model of 25 (green) in the K_Ca3.1 Cryo‐EM structure (pdb 6cn0)18 in the bound state II (light blue) with bound calmodulin (khaki). A,B) Overview of 25 binding in the inner pore. C) Surface representation of 25. D) Molecular representation of 25 and surface representation of ion channel. The given distance of 7.86 Å, indicates the distance between the O‐atoms and thus the length of the linker and the distance between senicapoc moiety and the fluorescent dye.

The modelling of 25 indicates a linker length that is nearly perfect to span the distance between the senicapoc moiety and the dye. Figure 6 A,B shows the binding of 25 in the inner pore of the K_Ca3.1 channel. Figure 6 C,D shows a more detailed view and how 25 fits onto the surface of the K_Ca3.1 channel. The senicapoc moiety (in the upper part) is binding in a similar region as described for a rosetta homology model of K_Ca3.1 channel. In this model senicapoc binds also to the upper inner pore of the channel.19 The dye is lying outside of the inner pore (lower part) directly on the surface. Therefore, the linker shows a suitable length to connect both moieties through the inner pore. (Figure 6 D) The distance between both O‐atoms is circa 7.86 Å. This analysis confirms a similar binding mode of the fluorescently labeled probe 25 and senicapoc in the inner pore of the K_Ca3.1 channel.

Conclusion

A series of fluorescently labeled ligands targeting the K_Ca3.1 channel was designed, synthesized, and evaluated for their suitability for imaging K_Ca3.1 channels in vitro in NSCLC cells by fluorescence microscopy. Ligands 25 and 26 labeled with dimethylpyrrole‐based BODIPY dyes and different linkers showed promising results in staining experiments. The typical punctate staining pattern was observed, which was reversed after pre‐incubation with the competitor senicapoc. Imaging of the K_Ca3.1 channels with the indirect immunofluorescence assay and ligand 25 led to the same density of signals, but imaging with 25 was more than six‐fold faster. Additionally, modelling in the available K_Ca3.1 Cryo‐EM structure showed similar binding modes for 25 and senicapoc. Compounds 22 and 23 labeled with ethyldimethylpyrrole‐based BODIPY dyes revealed unspecific staining of NSCLC cells without any structure. Whereas ethyl and 3,6,9‐trioxaundecyl moieties are suitable linkers for fluorescently labeled probes, a dodecyl linker (e.g., 24) led to highly lipophilic compounds resulting in low water solubility, which caused problems in terms of preparing staining solutions. Compound 21 labeled with the BODIPY bearing the thienopyrrole moiety was not suitable as a small‐molecule imaging probe owing to lack of staining the NSCLC cells. The low quantum yields of the connected dye 15, resulting in low intensity of emitted light is an additional disadvantage of this compound. In summary, dimethylpyrrole‐based BODIPY dyes connected with the targeting senicapoc moiety through an ethyl linker (25) represents a promising new small‐molecule probe for selective imaging of K_Ca3.1 channels of NSCLC cells in vitro and in vivo using a simple, fast, and efficient staining protocol.

Conflict of interest

The authors declare no conflict of interest.

Supporting information

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Supplementary

K. Brömmel, S. Maskri, I. Maisuls, C. P. Konken, M. Rieke, Z. Pethő, C. A. Strassert, O. Koch, A. Schwab, B. Wünsch, Angew. Chem. Int. Ed. 2020, 59, 8277.