A Peptoid-Based Fluorescent Sensor for Cyanide Detection

Abstract

Peptoids, N-substituted glycine oligomers, are versatile peptidomimetics with diverse biomedical applications. However, strategies to the development of novel fluorescent peptoids as chemical sensors have not been extensively explored, yet. Here, we synthesized a novel peptoid-based fluorescent probe in which a coumarin moiety was incorporated via copper(I)-catalyzed azide-alkyne cycloaddition reaction. Fluorescence of the newly generated coumarin-peptoid was dramatically quenched upon coordination of the Cu²⁺ ion, and the resulting peptoid-Cu²⁺ complex exhibited significant Turn-ON fluorescence following the addition of CN⁻. The rapid and reversible response, combined with cyanide selectivity of the synthesized peptoid, reflects a multistep photo-process and supports its utility as a new type of CN⁻ sensor.

1. Introduction

Peptoids, N-substituted glycine oligomers, are versatile mimics of peptides and are known to interact with various biological targets [1,2]. The proteolytic stability of peptoids, along with improved cell permeability relative to peptides, has led to their wide application as ligands for biological targets in various biomedical fields [3,4,5,6,7,8]. However, their significant conformational flexibility, combined with lack of hydrogen bonding ability, impose a major challenge for exploitation of peptoids in the drug discovery program.

Extensive studies have been devoted to enhancing the structural and functional capabilities of peptoids (including foldamers) [9,10,11]. Construction of peptoids with homogeneously-populated conformers remains a difficult task due to the low energy barrier of cis/trans-isomerization [12,13,14]. In particular, the development of new strategies to constrain flexible peptoids that adopt certain secondary structures is still challenging. To expand the applications of peptoids, understanding the physicochemical propensity that controls optimal conformations is essential. One of the methods to accomplish this goal is to attach fluorophores to the peptoid backbone and monitor interactions [15].

The 1,2,3-triazolyl group synthesized by the copper(I) catalyzed azide-alkyne [3+2] cycloaddition reaction coordinates with metal ions, with N₂ or N₃ directly involved in coordination [16,17]. Owing to their metal binding capacity [9,18,19,20,21,22,23,24,25,26,27], combined with ease of incorporation of fluorophore modules using solid-phase synthesis, peptoids present a good platform for the development of novel fluorescence sensors or functional peptidomimetics. In addition, understanding metal binding interactions with proteins is critical because human bodies are under the tight regulation of metal concentration inside and outside cells [28]. As a proof of concept to demonstrate a fluorescent peptoid sensor that can mimic the copper binding motif in many proteins, a simplified form of scaffold with functional diversities needs be explored.

Coumarins are one of the extensively used fluorophores because of their small size, high quantum yield, and efficient membrane permeability. Metal-complexed coumarin derivatives are versatile sensor molecules [29,30,31,32,33,34]. Recently, increasing numbers of triazolyl coumarin motifs have been utilized for various biological applications, such as the development of anti-inflammatory [35], anticancer [36], and antibacterial agents [37]. These compounds have also been explored as selective chemosensors [35,38,39,40,41]. In addition to the various advantages, including fast response and high sensitivity, development of new fluorescence sensor molecules for selective detection of ions is an active focus of biochemical research. Among the anions, efficient detection of the cyanide anion is particular interest due to severe toxicity issues related to human health [42,43,44,45]. In addition to wide industrial applications, the release of CN⁻ is harmful to the environment. So far, limited studies have explored the development and utility of peptoid-derived fluorescence sensors [15,46,47].

In this study, we have reported the synthesis and characterization of a novel peptoid-based fluorescence probe. The coumarin moiety was incorporated into the peptoid backbone via copper(I) catalyzed azide-alkyne [3+2] cycloaddition reaction. The fluorescence of the coumarin-peptoid was quenched by coordination with Cu²⁺ ion. The peptoid-Cu²⁺ complex exhibited significant Turn-ON fluorescence upon the addition of CN⁻. In view of its utility in rapid, selective, and reversible detection of CN⁻, our newly-generated peptoid molecule may serve as a novel type of CN⁻ sensor.

2. Results

2.1. Synthesis of Coumarin-Attached Peptoids

The synthesis of coumarin-attached peptoids is depicted in Figure 1. We expected that the coumarin unit at residue 1 would have a π-π interaction with any aromatic unit attached at residue 3. We limited the size of the peptoid as a trimer in our design, which would be a minimal length in terms of the number of atoms for metal complexation while avoiding the complicated NMR characterization of peptoids due to the presence of many rotamers.

The amine building block containing the acetylene group was incorporated using the standard solid phase method to accommodate alkyne peptoid 1 [48]. The peptoid was subsequently connected with azido-coumarin 2 via Cu(I)-catalyzed Huisgen 1,3-dipolar cycloaddition (CuAAC) using copper sulfate and sodium ascorbate in tert-butyl alcohol/water solution (v/v = 1:1) [49].

2.2. Spectroscopic Features of Free CP3

Figure 2 shows the absorption and emission spectra of the free form of CP3. UV spectra disclosed maximum absorption at around 350 nm in most solvents, with additional absorption bands at longer wavelengths, as shown in

Table 1. Spectroscopic values of CP3.

Wavelength	CHCl3	MeOH	DMF	DMSO	H2O
λex (nm)	357	347	347/441	347/441	344
λem (nm)	425	423	429/492	429/492	475
Stokes shift (nm)	68	76	82/51	82/51	131

. In aprotic polar solvents, such as DMF and DMSO, typical coumarin emission observed at ~430 nm decreased and a new emission band at longer wavelengths was observed. CP3 displayed dual emission. Excitation at 347 nm produced emission spectra with a maximum wavelength at 429 nm whereas excitation at 441 nm led to an additional emission band maximized at ~492 nm. The emission band at 429 nm represents 7-OH* of 7-hydroxycoumarin, while that at ~492 nm is likely to be 7-O⁻* of coumarin in the excited state. This dual emission was previously reported to occur due to the equilibrium between 7-OH* and 7-O⁻* of coumarin derivatives in the excited states [50].

2.3. Fluorescence Changes with Formation of the CP3-Cu²⁺ Complex

First, we explored the metal coordination ability of the coumarin-peptoid CP3 in various solvents using fluorescence spectroscopy. The changes in emission spectra induced upon the addition of Cu²⁺ are shown in Figure 3. The maximal emission band at 492 nm was markedly decreased in the presence of increasing concentrations of Cu²⁺ in DMF. A similar trend was observed upon addition of Cu²⁺ to CP3 solution in DMSO. This decrease in fluorescence appears to be related to photoinduced electron transfer (PET) based on dramatic fluorescent intensity changes with no spectral shift [51,52].

In the presence of protic solvents, such as H₂O and methanol, the quenching effect upon addition of Cu²⁺ was different. No spectral changes were observed in water, while the addition of Cu²⁺ resulted in a decrease of emission at 423 nm in methanol, but to a lesser extent than in DMF/DMSO. Even excess amounts of Cu²⁺ ion (35 equiv.) did not induce fluorescence quenching in water, indicating a constrained peptoid structure, presumably due to π-π stacking between coumarin and the benzene unit (Figure S1).

Considering the chelation-induced fluorescence changes in CP3 upon addition of Cu²⁺, we determined the stoichiometry of the peptoid-Cu²⁺ complex. Figure 4 shows Job plot analysis demonstrating a maximum at a mole fraction of 0.5, suggesting a 1:1 stoichiometry for the CP3-Cu²⁺ complex. The binding constant (K_d) of the peptoid-Cu²⁺ complex was 0.64 ± 0.093 μM in DMF and 0.65 ± 0.36 μM in MeOH:CHCl₃ (v/v = 1:5), as measured from an equilibrium binding titration experiment (Figure S2, Table S1). Changes in absorption spectra between free and Cu²⁺-complexed CP3 are shown in Figure 4b.

2.4. The CP3-Cu²⁺ Complex as a Cyanide Sensor

Next, we examined whether the peptoid-Cu²⁺ complex is dissociated in the presence of anions. As shown in Figure 5a the peptoid-Cu²⁺ complex exhibited Turn-ON fluorescence upon the addition of cyanide ions. Emission at 492 nm (solid black line) quenched in the presence of Cu²⁺ (dotted black line) was recovered by the addition of increasing amounts of cyanide. Turn-ON fluorescence is attributable to the formation of a [Cu(CN)_x]ⁿ⁻ complex [53], resulting in release of the free form of fluorescent coumarin-peptoid. Interestingly, this fluorescence change was reversed when Cu²⁺ ion was re-added to the same solution (Figure 5b). Notably, the ON/OFF switch response of CP3 was reproducible and rapid. All fluorescence emission spectra were recorded after each addition of the ion solutions was equilibrated for 1 min. Figure 5c demonstrates the kinetic profile of the fluorescence intensity increase at 492 nm when 5 μM of the cyanide anion was added to the 1 μM of CP3-Cu²⁺ complex. The kinetic trace was best fit to a single exponentional rate equation. The rate constant was measured as the increase in fluorescence intensity was 0.028 s⁻¹, which corresponds to a half-life (t_1/2) of 25.2 s. The data demonstrate a rapid sensing of CP3-Cu²⁺ complex for cyanide ion.

To determine the selectivity of peptoid-Cu²⁺ in anion sensing, we measured the recovered fluorescence increase following binding to various anions, including halogens, phosphate, azide, and sulfur anions (Figure 6). Among these, only the cyanide anion induced a fluorescence increase for CP3-Cu²⁺. The observed selectivity for the cyanide anion along with the rapid and reversible sensing properties of peptoid-Cu²⁺ supports its potential utility as a new class of cyanide sensor.

The 1-D NMR spectra of free CP3 and CP3-Cu²⁺ complex were compared, as shown in Figure S3a. In the complex, peaks in the aromatic proton region were shifted upfield. The data support the formation of 2-hydroxy coumarin via tautomerization, which may be promoted by Cu²⁺ coordination at the 2-hydroxy position.

2.5. CP4 Displays Different Spectroscopic Features to CP3

We additionally examined the spectroscopic features of the 7-methoxy analogue, CP4, to gain further insights into the mechanism of coumarin-peptoid. As shown in Figure 7, the emission peak at 492 nm observed with CP3 in DMF/DMSO was absent, resulting in a sole emission band at 418 nm. Furthermore, no fluorescence changes were observed, even upon addition of 10 equiv. of Cu²⁺ ion either in DMF or DMSO solvent (Figure 7). This result supports our hypothesis that the hydroxy group is critical for metal coordination with Cu²⁺, which is facilitated by the electron density redistribution within CP3.

3. Discussion

The unique photophysical properties of hydroxycoumarin have been documented, both spectroscopically and theoretically [54,55,56,57]. Here, we constructed a peptoid coordinated with Cu²⁺ where a fluorescent coumarin module was attached via click chemistry.

Figure 8 depicts the proposed mechanism of fluorescence sensing based on our observations with CP3. Excited state proton transfer (ESPT) appears to be related to the shift in electron density from hydroxyl to carbonyl group in 7-hydroxycoumarin. Redistribution of the electron density within CP3 upon photo-excitation explains the enhanced acidity of 7-hydroxycoumarin moiety, triggering photoinduced proton transfer to the solvent [54]. The pK_a of the 7-hydroxy group of coumarin is ~7.7 in the ground state, while photoexcitation enhances the acidity of the hydrogen-bonded proton to ~0.45 in the S₁ state [56]. The resulting anion of CP3 is highly emissive with a maximum peak at 492 nm.

Photoinduced electron transfer (PET) additionally occurs in the (tautomerized, 2O⁻) anionic coumarin fluorophore in the presence of Cu²⁺, thereby inducing fluorescence quenching at 492 nm. PET within the peptoid-Cu²⁺ complex was featured by large fluorescence intensity changes with no spectral shift. This OFF response with Cu²⁺ was subsequently reversed by the addition of cyanide ion, leading to an OFF-ON fluorescent response. This phenomenon is attributable to the formation of a stable [Cu(CN)_x]ⁿ⁻ complex [58,59]. The observed photo-driven processes were rapid, switchable, and reproducible.

Hydroxycoumarin-based chemosensors have been reported previously [40,55], but our molecule utilizes the unique photophysical characteristic of photo-excited enol-to-keto conversion in 7-hydroxycoumarin. In the previous report, the anion of 7-hydroxycoumarin was shown to be directly involved in metal coordination [55], whereas photo-excited enol-to-keto conversion resulted in metal coordination with the hydroxy anion at the C2 position in our molecule. Coumarin was used as a fluorescence reporter as well as coordination ligand. The excited state of 7-hydroxy-4-methylcoumarin is reported to favor the keto form, with the S₁ potential energy minima of the enol/keto form separated by a 17–20 kcal/mol energy barrier. In the ground state, the enol form is more stable whereas the keto form is more stable by 0.6 kcal/mol in the S₁ state [56]. Our NMR results were consistent with the reported enol-to-ketone transformation in the presence of Cu²⁺ ion, revealing upfield-shifted peaks in the aromatic region (Figure S3a).

Cu²⁺ is known to have strong paramagnetic properties due to the faster electron-relaxation time (Ts) of Cu²⁺. Therefore, the Cu²⁺ complex is hard to characterize by NMR. Various studies have reported severe line broadening of the Cu²⁺ binding to metalloproteins and used other experimental methods than NMR (including CD, EPR, UV-Vis, X-ray crystallography, etc.) [28,60,61]. Initially, we tried to get a crystal for X-ray crystallography, which will unequivocally reveal the metal-complexed peptoid, but the crystal was not obtained. The absorption spectra are, in general, critical to verify the differences in the binding of ligands to metal. However, the absorption spectra of Cu²⁺ solution were overlapped with those of CP3 in DMSO and DMF (λ_max = 300 nm in DMSO; λ_max = ~280 in DMF), which imposed difficulties in obtaining reliable titration data based on the absorbance. Even though we observed absorption spectral changes between free CP3 and the CP3-Cu²⁺ complex (Figure 4b), the only binding stoichiometry that we determined for the CP3-Cu²⁺ complex was from the fluorescence data. This issue may be resolved once the structural information of the CP3-metal complex is obtained.

CP3 in the aqueous system does not exhibit fluorescence quenching by Cu²⁺, possibly due to the constrained conformation resulting from π-π stacking interactions, which does not afford optimal geometry for coordination with Cu²⁺. We performed a simulated annealing analysis followed by energy minimization of each conformer. An ensemble of energy-minimized structures with the lowest E values is depicted in Figure S1. The trans-trans rotamer showed the lowest energy (9.7 kcal/mol), furnishing π-π stacking interactions between the benzene ring and coumarin unit.

In summary, we have developed peptoid-Cu²⁺ complex that effectively acts as a CN⁻ sensor. Photo induced proton transfer resulted in conversion of the 7-hydroxycoumarin moiety to its tautomeric form, facilitating metal coordination within the peptoid. The current form of the peptoid-Cu²⁺ complex only working in aprotic polar solvents represents a drawback for practical applications, which requires more systematic approaches to develop peptoid sensors working in aqueous solution. Although detailed mechanistic analyses are yet to be conducted, our understanding on the peptoid-based sensor design can be enriched by the knowledge obtained in the present work with a newly developed coumarin-attached peptoid-Cu²⁺ probe.

4. Experimental Section

4.1. General

Unless specified, all reagents and starting materials were ACS grade or higher and were used without further purification. All reactions were monitored by thin-layer chromatography on a TLC silica gel 60 F254 plate (Merck, Darmstadt, Germany), and compounds were visualized under UV light (254, 365 nm, VL-4.LC, Vilber Lourmat, Eberhardzell, Germany). Flash column chromatography was performed using ZEOprep silica gel (230~400 mesh, Zeochem, Lake Zurich, Switzerland) with hexane, ethyl acetate, dichloromethane, and methanol as eluents. ¹H (300, 800 MHz) and ¹³C-NMR (75, 200 MHz) spectra were recorded on a GEMINI 2000 (VARIAN, Palo Alto, CA, USA) and FT-NMR Avance III HD (Bruker, Billerica, MA, USA). Chemical shifts (δ) are reported in parts per million (ppm) and coupling constants (J) are given in hertz (Hz). All ESI-MS were undertaken on a 6130 Single Quadrupole LC/MS (Agilent Technologies, Santa Clara, CA, USA) and high-resolution mass spectra (HR-MS) were acquired under fast atom bombardments (FAB) condition on a JMS-700 MStation (JEOL, Tokyo, Japan). UV-Vis and fluorescence spectra were obtained using a Lambda 25 (Perkin Elmer, Waltham, MA, USA) and FP-6500 (Jasco, Tokyo, Japan). High-performance liquid chromatography (HPLC) analysis was performed on a YL9100 reversed-phase HPLC (Younglin, Anyang, South Korea).

4.2. Synthesis of N-(2-Amino-2-oxoethyl)-2-(2-(benzylamino)-N-(2-methoxyethyl)acetamido)-N-(prop-2-yn-1-yl)acetamide (Solid Phase Peptide Synthesis) (1)

Peptoid oligomer was synthesized on MBHA resin (0.43 mmol/g) using a conventional peptoid synthesis protocol to generate an amide group at the C-terminus of the peptoids [2]. The Fmoc protected resin was swollen in N,N-dimethylformamide (DMF) for 30 min before starting oligomer synthesis. The Fmoc group was removed with 20% piperidine in DMF. Acylations using 1 M bromoacetic acid (BrAA, 10 equiv.) and 1 M N,N′-diisopropylcarbodiimide (DIC, 10 equiv.) in DMF followed by nucleophilic displacement step using 0.5–2 M amine (propargyl amine, 2-methoxyethylamine, benzylamine) in DMF were repeated until desired peptoid was obtained. The resin was cleaved with 95% trifluoroacetic acid (TFA) in H₂O for 1 h with occasional agitation (180 rpm). The filtered solution was purged with N₂ to remove TFA. The crude compounds were dissolved in 50% acetonitrile/H₂O and lyophilized two times to remove residual trifluoroacetic acid (HPLC purity: 93.6%). ¹H-NMR (300 MHz, DMSO-d₆): δ 9.32 (bs, s, 2H), 7.51–7.40 (m, 5H), 4.46/4.42* (2 × s, 1H), 4.27–4.22 (m, 2H), 4.13–4.07 (m, 6H), 3.93/3.89* (2 × s, 1H), 3.8 (bs, s, 1H), 3.46–3.40 (m, 4H) 3.30/3.25* (2 × t, 1H), 3.19 (m, 3H) (rotamer peaks*). ¹³C-NMR (75 MHz, DMSO-d₆): δ 169.7, 169.5*, 169.3*, 169.1*, 168.4, 168.0*, 167.8*, 167.4*, 166.5, 165.9*, 158.8, 158.4*, 158.0*, 157.5*, 131.6, 131.5*, 130.3, 130.23*, 130.17, 129.1, 128.7, 79.1, 78.9*, 78.8*, 78.5*, 76.1, 75.7*, 75.2*, 74.9*, 69.5, 69.4*, 69.3*, 69.2*, 58.34, 58.30*, 58.1, 58.0*, 50.1, 49.9*, 49.0*, 48.9*, 48.6*, 48.1, 48.0*, 47.4, 46.9*, 46.7*, 46.5, 46.2*, 37.3, 37.2*, 35.7*, 35.4* (rotamer peaks*). HR-MS (m/z): 375.2034 (calculated for C₁₉H₂₇N₄O₄ [M + H]⁺, 375.2032).

4.3. Synthesis of 3-Azido-7-hydroxycoumarin (2a) and 3-Azido-7-methoxycoumarin (2b)

The compounds 2a and 2b were synthesized according to the reported method [49].

3-Azido-7-hydroxycoumarin (2a). ¹H-NMR (300 MHz, MeOH-d₄): δ 7.40 (s, 1H). 7.37 (d, J = 2.1 Hz, 1H), 6.80 (d, J = 2.4 Hz, 1H), 6.77 (d, J = 2.4 Hz, 1H), 6.72 (d, J = 2.1 Hz, 1H).

3-Azido-7-methoxycoumarin (2b). ¹H-NMR (300 MHz, CDCl₃): δ 7.31 (d, J = 8.4 Hz, 1H), 7.18 (s, 1H), 6.89–6.84 (m, 2H), 3.87 (s, 3H).

4.4. Synthesis of N-(2-Amino-2-oxoethyl)-2-(2-(benzylamino)-N-(2-methoxyethyl)acetamido)-N-((1-(7-hydroxy-2-oxo-2h-chromen-3-yl)-1h-1,2,3-triazol-4-yl)methyl)acetamide (CP3)

Copper (II) sulfate pentahydrate (0.3 M in water, 35 μL, 0.0104 mmol) and sodium ascorbate (1 M in water, 42 μL, 0.0416 mmol) were added to a mixture of alkyne peptoid, 1 (39 mg, 0.104 mmol) and 3-Azido-7-hydroxycoumarin, 2a (20 mg, 0.104 mmol) in water and tert-butyl alcohol (v/v = 1:1, 4 mL). The reaction mixture was stirred at room temperature for 24 h in the dark. After removing the solvent under reduced pressure, the crude material was purified by column chromatography (MeOH/DCM, 1:10) afforded CP3 as a yellow solid (30 mg, 50%). ¹H-NMR (800 MHz, DMSO-d₆): δ 11.01 (bs, s, 1H), 9.24 (bs. s, 2H), 8.71/8.67*/8.47*/8.41* (4 × s, 1H), 8.61/8.59*/8.58* (4 × s, 1H), 7.77–7.75 (m, 1H), 7.50–7.39 (m, 5H), 6.92 (d, J = 8.4 Hz, 1H), 6.87 (d, J = 8.7 Hz, 1H), 4.72/4.71* (2 × s, 1H), 4.68/4.62* (2 × s, 1H), 4.61/4.59* (2 × s, 1H), 4.30/4.26* (2 × s, 1H), 4.13 (bs, s, 2H), 4.08 (m, 2H), 3.88 (d, J = 7.4 Hz, 1H), 3.85 (bs, m, 1H), 3.50–3.46 (m, 2H), 3.44–3.40 (m, 2H), 3.19 (dd, J₁ = 13.3, J₂ = 2.7 Hz, 3H) (rotamer peaks*). ¹³C-NMR (200 MHz, DMSO-d₆): δ 169.88, 169.65*, 169.57*, 169.39*, 168.67, 168.11*, 168.12*, 167.57*, 166.47, 166.46*, 165.80*, 165.79*, 162.59, 162.57*, 162.54*, 162.51*, 158.05, 157.89*, 157.73*, 157.58*, 156.25, 156.23*, 156.21*, 154.66, 154.65*, 154.61*, 154.59*, 143.31, 143.14*, 143.10*, 142.85*, 136.33, 136.15*, 136.10*, 131.52, 131.43*, 131.39*, 130.98, 130.96*, 130.93*, 130.22, 130.18*, 130.10*, 129.01, 128.64, 128.61*, 128.59*, 124.64, 124.7*, 124.38*, 124.24*, 119.17, 119.16*, 119.13*, 114.34, 114.33*, 110.27, 110.26*, 110.25*, 102.17, 102.14*, 69.31, 69.28*, 69.27*, 69.21*, 58.29, 58.25*, 57.97, 57.95*, 50.16, 50.13*, 49.88*, 49.86*, 49.09, 48.99*, 48.84*, 48.59, 48.21*, 47.29, 47.26*, 46.95*, 46.58*, 46.52*, 46.50*, 46.25*, 46.22*, 42.34, 41.87*, 41.59* (rotamer peaks*). HR-MS (m/z): 578.2368 (calculated for C₂₈H₃₂N₇O₇ [M + H]⁺, 578.2363). ¹H- and ¹³C-NMR spectra of CP3 are available in the Supplementary Materials (Figure S4).

4.5. Synthesis of N-(2-Amino-2-oxoethyl)-2-(2-(benzylamino)-N-(2-methoxyethyl)acetamido)-N-((1-(7-methoxy-2-oxo-2h-chromen-3-yl)-1h-1,2,3-triazol-4-yl)methyl)acetamide (CP4)

The synthetic procedure described above was used for the preparation of CP3. 3-Azido-7-methoxycoumarin, 2b (14.5 mg, 0.067 mmol) and alkyne peptoid, 1 (25 mg, 0.067 mg) were used. The reaction mixture was purified by column chromatography to obtain 23 mg (58%) of CP4 as a pale yellow solid. ¹H-NMR (800 MHz, DMSO-d₆): δ 9.21 (bs, s, 2H), 8.74/8.70*/8.50*/8.44* (4 × s, 1H), 8.67/8.66*/8.65* (4 × s, 1H), 7.87–7.85 (m, 1H), 7.49–7.39 (m, 5H), 7.18–7.16 (m, 1H), 7.09 (d, J = 8.6 Hz, 1H), 4.73/4.72* (2 × s, 1H), 4.68/4.63* (2 × s, 1H), 4.62/4.60* (2 × s, 1H), 4.30/4.27* (2 × s, 1H), 4.13 (d, J = 4.4 Hz, 2H), 4.08 (dd, J₁ = 11.9, J₂ = 3.9 Hz, 2H), 3.91 (d, J = 2.7, 3H), 3.88 (d, J = 8.3 Hz, 1H), 3.85 (d, J = 10.2 Hz, 1H), 3.50–3.46 (m, 2H), 3.44–3.40 (m, 2H), 3.19 (dd, J₁ = 13.6, J₂ = 2.9 Hz, 3H) (rotamer peaks*). ¹³C-NMR (200 MHz, DMSO-d₆): δ 169.87, 169.64*, 169.55*, 169.38*, 168.86, 168.12*, 168.02*, 167.57*, 166.51, 166.84*, 163.47, 163.45*, 163.43*, 163.40*, 157.68, 157.53*, 156.10, 156.09*, 154.53, 154.52*, 154.48*, 154.46*, 143.37, 143.20*, 143.16*, 142.91*, 135.80, 135.79*, 135.62*, 135.55*, 131.59, 131.47*, 130.64, 130.62*, 130.60*, 130.19, 130.15*, 130.08*, 129.00, 128.64, 128.60*, 128.59*, 124.62, 124.46*, 124.36*, 124.22*, 120.08, 120.07*, 120.05*, 113.57, 113.54*, 111.45, 111.44*, 111.43*, 100.73, 100.72*, 100.69*, 69.31, 69.27*, 69.21*, 58.29, 58.25*, 57.97, 57.95*, 56.20, 50.18, 50.15*, 49.90*, 49.88*, 49.11, 49.07*, 49.01*, 48.83*, 48.59*, 48.20*, 47.25, 46.94*, 46.51*, 46.49*, 46.27*, 46.25*, 42.35, 42.33*, 41.88*, 41.59* (rotamer peaks*). HR-MS (m/z): 592.2510 (calculated for C₂₉H₃₄N₇O₇ [M + H]⁺, 592.2520). ¹H- and ¹³C-NMR spectra of CP4 are available in the Supplementary Materials (Figure S5).

4.6. Spectroscopic Measurements

Fluorescence emission spectra were obtained at 20 °C using a JASCO FP-6500 spectrofluorometer (Jasco). The equilibrium binding titration experiments of peptoid with Cu²⁺ or other anions were performed as follows. 0.5–2 µL of metal or anion stock solutions (10 mM in DMSO) were added into 1 µM peptoid solutions in quartz cuvette, and the mixture was equilibrated for 1 min to ensure full binding before measurements. The slit widths used for measurement were 3 nm for both excitation and emission with medium sensitivity, but was adjusted depending on the strength of the fluorescence signal. The kinetic measurement was performed on the same instruments. The fluorescence intensity was recorded at 492 nm with 0.01 s time intervals without incubation time. For spectroscopic measurement, all stock solutions of peptoids and the sodium or potassium salts of anions (NaCN, NaHSO₃, Na₂HPO₄, NaN₃, Na₂SO₄, NaSCN, Na₂S₂O₅, NaNO₃, NaClO₄, NaNO₂, KF, KCl, KBr, KI) were prepared in DMSO (1% water). Absorption spectra were obtained at room temperature using a Lamda 20 UV-Vis spectrometer (Perkin Elmer, MA, USA) with 1.0 cm quartz cells. All measurements were triplicated. The CP3-Cu²⁺ complex for the absorption spectra was obtained by adding CuSO₄ to CP3 solution with stirring for 12 h at ambient temperature followed by purification described above. The ESI-MS spectrum of CP3-Cu²⁺ is available in Figure S6.