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. 2024 Feb 22;63(9):4176–4184. doi: 10.1021/acs.inorgchem.3c04076

O-versus S-Metal Coordination of the Thiocarboxylate Group: An NMR Study of the Two Tautomeric Forms of the Ga(III)-Photoxenobactin E Complex

Larissa Buedenbender †, Lucía Ageitos †, Marta A Lages ‡, Carlos Platas-Iglesias †, Miguel Balado ‡, Manuel L Lemos ‡, Jaime Rodríguez †,*, Carlos Jiménez †,*
PMCID: PMC10915793  PMID: 38387064

Abstract

graphic file with name ic3c04076_0005.jpg

Photoxenobactin E (1) is a natural product with an unusual thiocarboxylic acid terminus recently isolated from an entomopathogenic bacterium. The biosynthetic gene cluster associated with photoxenobactin E, and other reported derivatives, is very similar to that of piscibactin, the siderophore responsible for the iron uptake among bacteria of the Vibrionaceae family, including potential human pathogens. Here, the reisolation of 1 from the fish pathogen Vibrio anguillarum RV22 cultured under iron deprivation, its ability to chelate Ga(III), and the full NMR spectroscopic characterization of the Ga(III)-photoxenobactin E complex are presented. Our results show that Ga(III)-photoxenobactin E in solution exists in a thiol–thione tautomeric equilibrium, where Ga(III) is coordinated through the sulfur (thiol form) or oxygen (thione form) atoms of the thiocarboxylate group. This report represents the first NMR study of the chemical exchange between the thiol and thione forms associated with thiocarboxylate-Ga(III) coordination, including the kinetics of the interconversion process associated with this tautomeric exchange. These findings show significant implications for ligand design as they illustrate the potential of the thiocarboxylate group as a versatile donor for hard metal ions such as Ga(III).

Short abstract

Ga(III)-photoxenobactin E in solution exists in a thiol–thione tautomeric equilibrium, where Ga(III) is coordinated through the S or O atoms of the thiocarboxylate group. This is the first NMR study describing the chemical exchange between the thiol and thione forms in thiocarboxylate-Ga(III) coordination and the associated kinetics of interconversion. The results suggest that this motif can be useful for the preparation of new ligand families.

Introduction

The interest in metal complexes of thiocarboxylate ligands has increased in the last years not only from a structural point of view but also from synthetic aspects due to their interesting properties and their involvement in some biological processes.1 Because these ligands contain both “soft” sulfur and “hard” oxygen donor sites, they enable the incorporation of hard, soft, and borderline metal ions into a coordination compound, as well as the stabilization of unusual coordination numbers and geometries.2 Metal thiocarboxylate complexes are being used to prepare several types of sulfide materials that exhibit interesting optical, electronic, and conductive properties.3−5 They can be used to form polymeric compounds with electrical and luminescent features6,7 and to prepare new catalysts for organic synthesis8 such as the azide–alkyne cycloaddition reaction.9 Thiocarboxylate complexes have also shown to be effective in a variety of biological applications, such as enzymatic catalysis and drug delivery.10 Further applications of thiocarboxylate complexes include their use in a variety of environmental problems such as water purification and remediation of contaminated soils.11,12

Although the tautomerism of thiocarboxylic acids as a mixture of thiol and thione forms has been studied on a number of occasions,13−15 the tautomerism of their corresponding thiocarboxylate complexes has received less attention. Four coordination modes have been suggested for thiocarboxylate complexes of metals, depending on the nature of the metal ion and the solvent (Scheme 1).16,17 The coordination behavior of the thiocarboxylate ligand toward the metal was studied by spectral analysis using infrared, electronic, and Mössbauer spectra. However, NMR studies of tautomerism between some of the mentioned coordination modes have not been reported so far.

Scheme 1. Thiocarboxylate Coordination Modes with Metal Ions as Monodentate Ligand through (i) the Sulfur Atom or (ii) the Oxygen Atom, (iii) as Chelate Bidentate (μ1-(O,S)) or (iv) Bridging Bidentate (μ2-(O,S)) Ligand.

Scheme 1

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(M = metal ion).

There are a number of studies of the synthesis, structures, and applications of metal complexes bearing thiocarboxylate ligands;1 however, only a few examples of gallium–thiocarboxylate complexes have been reported so far. Nevertheless, the coordination chemistry of thiocarboxylate ligands remains underexplored, particularly in comparison with carboxylate analogues. The first dialkyl gallium thioacetate was published in 1971, but the first example of a structurally characterized gallium thiocarboxylate was methyl gallium thioacetate, Ga(SCOMe)2Me(3,5-dimethylpyridine) reported in 1996 as a novel single-source precursor for gallium thin films by aerosol-assisted chemical vapor decomposition (CVD).18 For the purpose of the present work, it is important to notice that the 13C NMR spectrum in bencene-d6 for this compound displays the carbonyl chemical shift at 201.95 ppm. Some years later, the synthesis and structural characterization of gallium thiobenzoates were reported. The treatment of thiobenzoic acid with one mol equivalent of trialkyl gallium, tBu3Ga or Me3Ga, yielded tBu2Ga[S(O)CPh] or Me2Ga[S(O)CPh], respectively. Their corresponding 13C NMR spectra in CDCl3 display the carbonyl chemical shifts at 221.98 and 218.81 ppm, respectively. In contrast, when Me3Ga is treated with three equivalents of thiobenzoic acid, a crystalline compound Ga[S(O)CPh]3 is obtained. Its crystal data, obtained by X-ray diffraction, showed a central hexacoordinate gallium atom bonded to three thiobenzoate moieties acting as bifunctional SO ligands. Its corresponding 13C NMR spectrum in CDCl3 showed a carbonyl chemical shift at 212.33 ppm.19

Natural thiocarboxylic acid-containing metabolites are extremely rare and so far, to the best of our knowledge, only nine natural products have been fully characterized due to the instability of these metabolites.20−25 Photoxenobactin E (1a) is such a natural product with an unusual thiocarboxylic acid terminus recently isolated by Shi and co-workers from an entomopathogenic bacterium.25 The pxb biosynthetic gene cluster (BGC) associated with photoxenobactin E (PxbE), and other reported derivatives, is very similar to the irp BGC encoding piscibactin (Pcb, 1b), the siderophore responsible for iron uptake in the fish pathogenic Gram-negative bacterium Photobacterium damselae subsp. piscicida.26 The irp BGC is widespread among bacteria of the Vibrionaceae family including potential human pathogens26−30 and contains an additional heterocyclization–adenylation–thiolation (Cy4-A2-T6) module. This module was thought to be either nonfunctional or to be involved in the synthesis of a cryptic metabolite containing an additional thiazoline, but to date has not been identified.26 An identical module was described for the pxb BGC encoding for photoxenobactin E,31 suggesting that Vibrionaceae might also be able to produce photoxenobactin-like metabolites. Siderophores are low-molecular-weight metabolites that are produced by microorganisms to acquire essential iron directly from the environment.32 The ability to scavenge iron from iron-deficient environments determines the growth and survival of the microorganism, and siderophores represent important virulence factors for pathogenic bacteria.33 Pcb was characterized as its Ga(III) complex due to its high instability in its apo-form.26 Surprisingly, the metal binding ability of photoxenobactin E was not detected when its isolation was reported.25

Reinvestigation of the fish pathogen Vibrio anguillarum RV22, cultured under iron deprivation, allowed us to isolate the siderophore photoxenobactin E as its Ga(III) complex (2). The coordination chemistry of Ga(III) and Fe(III) displays many similarities: they have the same charge (both 3+), similar ionic radii (Fe(III) = 0.65 Å, Ga(III) = 0.62 Å for coordination number six),34 preferred coordination number of six, and they are hard Lewis acids.35 The diamagnetic character of Ga(III), in contrast to the paramagnetic character of Fe(III), enables NMR studies of the Ga(III) thiocarboxylate complexes.

In this work, we report the reisolation of photoxenobactin E (1a) from the fish pathogen V. anguillarum RV22, its ability to chelate Ga(III), and the full NMR spectroscopic characterization of the Ga(III)-photoxenobactin E complex, via experimental and computational investigations. Our results showed that the Ga(III)-photoxenobactin E complex (2) exists as a tautomeric equilibrium mixture, where Ga(III) is bound either through the sulfur (2A) or through the oxygen (2B) of the thiocarboxylate (Figure 1). Moreover, we performed dynamic NMR to study the kinetic process of this tautomeric exchange. To the best of our knowledge, this is the first report in which the chemical exchange associated with thiocarboxylate-Ga(III) coordination of both the thiol and thione species was observed via NMR spectroscopy, allowing the analysis of kinetics of this tautomeric exchange.

Figure 1.

Figure 1

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Apo-forms of photoxenobactin E (1a) and piscibactin (1b) as well as holo-thiol (2A) and holo-thione (2B) forms of photoxenobactin E as the Ga(III)-photoxenobactin E complex (2).

Results and Discussion

The isolation of photoxenobactin E (1a) by Bode and co-workers from entomopathogenic bacteria and the similarity of its biosynthetic gene cluster25 to that of piscibactin (1b),26 motivated us to investigate whether pathogenic Vibrionaceae are also able to biosynthesize this compound and to assess its metal-chelating properties.

Cell-free culture supernatants of strain V. anguillarum RV22 grown under iron-restricted conditions were chelated with Ga(acac)3. Formation of Ga(III) complexes stabilizes the siderophores and allows spectroscopic analysis via NMR due to the absence of paramagnetism in this metal ion. Solid Phase Extraction (SPE) using HLB cartridges of the cell-free culture supernatants followed by HPLC separation allowed the isolation of the Ga(III) complex of photoxenobactin E (2). The molecular formula of 2 was established based on the isotopic cluster distribution due to the presence of gallium and the HRESIMS [M + H]+ ion at m/z 535.9719 (calculated for C19H21N3O3S469Ga m/z 535.9722 [M + H]+), which matched that of the Ga(III)-photoxenobactin E complex (Figure S1, Supporting Information).

Characterization of the Ga(III)-Photoxenobactin E Complex by NMR and IR Spectroscopy

Comparison of 1D and 2D NMR spectral data of 2 in DMSO-d6 with those reported for apo-photoxenobactin E (1a)25 allowed the confirmation of its structure. The carbon and proton chemical shifts were very similar in both compounds (Table 1). However, proton chemical shifts at positions near the Ga(III) ion: the methyl C-18 and methylene C-16 protons of the methylthiazoline moiety, the proton bonded to the nitrogen (10-NH) in the thiazolidine ring, and the proton at position C-12, exhibited duplicate signals in the 1H NMR spectrum of 2. More specifically, the chemical shift corresponding to CH3-18 resonated as two distinct methyl proton singlets at δH 1.51 and δH 1.61, which correlated in the HSQC experiment with carbons at δC 24.56 and δC 25.69, respectively (Figure S6). The 1H NMR methylene signals at C-16 were observed at δH 3.16/3.51 and δH 3.34/4.01, which correlated in the HSQC experiment with the carbon chemical shifts at δC 38.17 and δC 42.41, respectively. Two carbon resonances at higher chemical shifts were observed at δC 208.72 and δC 222.23 which correlated in the HMBC experiment to the CH3-18 protons at δH 1.51 and δH 1.61, respectively (Figure S7). Two proton chemical shift signals were observed for 10-NH of the thiazolidine ring at δH 6.37 and δH 6.64. Finally, the chemical shift of the methine proton at C-12 was observed as two multiplet signals at δH 3.70 and δH 3.62, which correlated in the HSQC experiment with carbons at δC 68.40 and δC 68.74, respectively. The carbon chemical shift of δC 208.72 agrees with the C=O resonance of the thiol form of the thiocarboxylate group as reported by Bode and co-workers,25 while the carbon chemical shift at δC 222.23 likely corresponds to the C=S resonance of the thione form of the thiocarboxylate group.36

Table 1. 1H and 13C NMR Data of apo-Photoxenobactin E (1, Reported by Bode and Co-Workers25) and for Ga(III)-Photoxenobactin E Tautomers (2A/B) at 298.15 K (Slow Exchange) and at 348.15 K (2T) (Fast Exchange) in DMSO-d6a.

Apo-photoxenobactin E
 Ga(III)-photoxenobactin E complex
  1a
 2A (thiol-form)
 2B (thione-form)
 2T
  298 K (800/200 MHz)25
 298 K (500/125 MHz)
 298 K (500/125 MHz)
 348 K (400/100 MHz)b
# δH (mult., J in Hz) δC, mult. δH (mult., J in Hz) δC, mult. δH (mult., J in Hz) δC, mult. δH (mult., J in Hz) δC, mult.
1   172.4, C   166.20, C   166.25, C   n.d.
2 6.50 (d, 8.5) 124.4, CH 6.67 (m) 123.25, CH 6.67 (m) 122.68, CH 6.68 (d, 8.8) 122.2, CH
3 7.09 (td, 8.5, 1.8) 133.9, CH 7.32 (m) 135.39, CH 7.33 (m) 135.67, CH 7.30 (m) 134.6, CH
4 6.32 (br t, 7.3) 112.1, CH 6.63 (m) 115.27, CH 6.63 (m) 114.73, CH 6.64 (t, 7.5) 114.6, CH
5 7.16 (dd, 7.9, 1.8) 132.3, CH 7.27 (m) 131.67, CH 7.27 (m) 131.71, CH 7.29 (d, 7.5) 130.7, CH
6   116.7, C   115.39, C   115.85, C   n.d.
7   172.1, C   174.83, C   176.41, C   n.d.
8 3.09 (ov) 33.7, CH2 3.34 (ov) 33.36, CH2 3.34 (ov) 33.05, CH2 3.36 (dd, 12.3, 11.3) 32.6, CH2
3.45 (ov) 3.54 (ov) 3.54 (ov) 3.57 (dd, 11.3, 8.6)
9 4.40 (ddd, 13.1, 10.3, 7.5) 78.7, CH 4.62 (m) 75.67, CH 4.61 (m) 75.72, CH 4.59 (ddd, 12.3, 9.9, 8.6) 75.3, CH
10 4.71 (dd, 10.3, 6.8) 70.6, CH 4.88 (dd, 9.9, 6.6) 68.99, CH 4.88 (dd, 9.9, 6.6) 68.85, CH 4.83 (dd, 9.9, 6.6) 68.6, CH
10-NH 5.31 (dd, 10.3, 6.9)   6.37 (dd, 9.5, 6.7)*   6.64*   6.30 (br t)  
11 3.45 (dd, 12.3, 7.6) 37.5, CH2 2.99 (m) 37.76, CH2 2.99 (m) 37.23, CH2 3.01 (ov) 37.0, CH2
3.03 (br t, 11.4) 3.50 (m) 3.50 (m) 3.52 (dd, 12.4, 7.3)
12 3.72 (dd, 18.1, 10.3) 66.3, CH 3.70 (m)** 68.40, CH 3.62 (m)** 68.74, CH 3.73 (m) 68.1, CH
13 3.99 (br s) 67.8, CH 4.08 (m) 70.07, CH 4.08 (m) 70.16, CH 4.14 (m) 69.8, CH
13-OH 7.52 (s)              
14 2.88 40.0, CH2 2.73 (brt, 14.4) 40.31, CH2 2.73 (br t, 14.4) 40.83, CH2 2.75 (d, 17.3) 40.3, CH2
3.08 2.92 (m) 2.92 (m) 2.95 (dd, 17.3, 5.1)
15   170.6, C - 180.21, C   181.92, C   n.d.
16 3.59 (d, 11.5) 39.4, CH2 3.16 (d, 11.5) 38.17, CH2 3.34 (ov) 42.41, CH2 3.25 (d, 11.2) 39.2, CH2
3.15 (d, 11.5) 3.51 (ov) 4.01 (d, 11.1) 3.79 (ov)
17   93.1, C   88.80, C   88.64, C   n.d.
18 1.56 (s) 25.6, CH3 1.51 (s) 24.56, CH3 1.61 (s) 25.69, CH3 1.60 (s) 24.7, CH3
19   212.1, C   208.72, C   222.23, C   n.d.
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a

ov = overlapped resonances; n.d. = not detected; *,** = interchangeable signals.

b

assigned by 1H,1H–COSY and 1H,13C-HSQC NMR experiments, hence quaternary carbons are not listed.

Closer inspection of the 13C NMR spectrum of 2, showed duplication of all carbon chemical shifts (Table 1 and Figure S4). Most of them displayed minor chemical shift differences that ranged between 0.1 to 1.5 ppm (Figure S4), but large differences were observed for carbons at C-16 (Δδ 4.24 ppm) and C-19 (Δδ 13.51 ppm) positions.

This behavior is characteristic of the presence of two complex species, where the variation in chemical shift is due to the different environments of the corresponding nuclei. The NOESY spectrum of 2 showed evidence of chemical exchange through cross-peaks between the two sets of 10-NH-signals of the thiazolidine ring (Figure S8). All of these data suggest that Ga(III)-photoxenobactin E complex (2) occurs as a tautomeric mixture of its thiol-2A and thione-2B forms, where the Ga(III) ion is coordinated to the sulfur or the oxygen, respectively (Figure 1). Integration of the CH3-18 resonances in the 1H NMR spectrum of 2 in DMSO-d6 indicated that the two tautomer species occurred in a ∼51:49 ratio at 298 K (Figure S3), with the thiol population being slightly higher compared to that of the thione form. Based on this ratio, we could differentiate the carbon chemical shifts of each tautomeric form in the 13C NMR spectrum of 2 (Table 1 and Figure S4).

Over time, we observed an increase of a singlet signal at 1.49 ppm in the 1H NMR spectra of 2 that was assigned to the methyl C-18 resonance of the Ga(III)-piscibactin complex. The presence of this compound was confirmed by the HRESIMS-(+) of an aliquot of the NMR tube, which shows the [M + H]+ ion at m/z 519.9948 corresponding to Ga(III)-piscibactin complex (2). We suggest that the Ga(III)-piscibactin complex must be the hydrolysis product of the Ga(III)-photoxenobactin E complex (2) since the hydrolysis of thiocarboxylic acids to carboxylic acids has been reported previously. Cortese et al. observed the hydrolysis of pyridine-2,6-bis(thiocarboxylic acid) in aqueous solutions through the stepwise release of H2S, particularly when the compound was chelated with Bi(III), Cr(III), and Pb(II).11 Recently, the transformation of thiocarboxylic acids into carboxylic acids via a visible-light-promoted atomic substitution was described with DMSO as the oxygen source.37

The absorption bands observed in the IR spectrum of 2 at 1606 cm–1 (very sharp and strong) and 982 cm–1 (weak) (Figure S2) were assigned to the ν(C=O) and ν(C=S) stretching vibration bands, respectively. The increase of both ν(C=O) and ν(C=S) in relation to those of sodium salts of monothiobenzoic acid (ν(C=O) at 1500 and ν(C=S) at 960 cm–1), used as a reference, confirmed coordination to gallium through the sulfur and oxygen atoms of the thiocarboxylate group.16,17

DFT Calculations of NMR Chemical Shifts of Tautomers 2A and 2B

The thiol-2A and thione-2B forms were optimized by using DFT at the B3LYP/6-31+G(d,p) level. Attempts were performed to compute different octahedral geometries involving a bidentate coordination of the thiocarboxylate group to Ga(III) through both S and O atoms, instead of the lone pair of the N atom of the thiazoline ring. However, we found that the thiocarboxylate group always coordinates to Ga(III) as either a thiol or a thione form, as shown in Figure 1 for 2A and 2B, respectively. Their 13C NMR chemical shifts were theoretically determined at the mPW1PW91/6-311+G(2d,p) level and then corrected by the computed Tantillo’s slope and intercept of −1.0490 and 186.6525, respectively.38 The 13C NMR chemical shifts obtained by this methodology were 210.3 ppm for the thiol- and 225.6 ppm for the thione forms, which were in very good agreement with the experimental carbon chemical shifts of 2 in DMSO-d6 at 208.72 and 222.23 ppm, respectively.

Dynamic NMR Studies

To confirm the existence of a tautomeric equilibrium between species 2A and 2B, we performed NMR experiments at variable temperatures (Figure 2). When the temperature was increased, the proton chemical shifts in the 1H NMR spectrum of the Ga(III)-photoxenobactin E complex (2) recorded in DMSO-d6 (400 MHz) exhibited a better resolution and the signal duplication disappeared (Figure 2, spectrum vi). For instance, slow exchange was observed at room temperature (298.15 K) for the signals of the C-18 methyl group for each tautomeric form as two sharp singlets separated by 25.33 Hz (Figure 2B, spectrum i). When the sample was warmed (303–313 K), the two singlet signals broadened, coalesced, and then started to sharpen and overlap (Figure 2B, spectra iv–vii). Fast exchange occurs at temperatures above 313.15 K and the C-18 methyl group is displayed as a single averaged peak in the 1H NMR spectrum.

Figure 2.

Figure 2

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(A) Dynamic 1H NMR of the tautomeric mixture of the Ga(III)-photoxenobactin E complex (2) measured in DMSO-d6 at variable temperatures: 298.15 to 348.15 K (400 MHz). (B) Expansion of regions from 1.5 to 1.7 ppm, showing the proton chemical shift corresponding to the methyl H-18 signal, and from 3.9 to 4.3 ppm, showing the proton chemical shift at δH 4.01 ppm corresponding to one of the methylene protons at C-16 (DMSO-d6, 400 MHz, 298.15–313.15 K).

The coalescence effect was also observed for the methylene protons at position C-16. At room temperature (298.15 K), the diastereotopic methylene protons in 2A and 2B resonated as two chemical shift pairs at δH 3.16/3.51 and δH 3.34/4.01 (Figure 2B, spectrum i), respectively. Interestingly, as we increased the temperature in the NMR experiments, the proton at δH 4.01 became broader and then disappeared at 313.15 K (Figure 2Bvii), indicating the presence of a dynamic process.

On the other hand, at higher temperatures, a new signal (broad doublet) appeared at δH 3.23 (Figure S10A), showing a similar coupling constant (11.2 Hz) as the one observed at δH 3.16 at 298.15 K. At 348.15 K, the selective irradiation of the signal at δH 3.23 in a 1D-TOCSY experiment revealed its diastereotopic partner at δH 3.79 (Figure S10B).

The assignment of the two methylene protons at δH 3.23 and 3.79 at C-16 (DMSO-d6, 400 MHz, 348.15 K, Figure S10B) was confirmed by the correlation between those protons observed in 1H–1H COSY spectrum of 2 (DMSO-d6, 400 MHz, 348.15 K, Figure S10C). These chemical shifts are the averaged values for both tautomeric forms 2A and 2B in a fast-interchanging regime of the two pairs of diastereotopic protons seen at 298.15 K. The 1H–13C-HSQC NMR spectrum correlation between δH 3.23 and δC 39.5 (Figures S10D) corroborated the assignment of C-16 as well as the rest of the NMR resonances as weight-averaged values of the two tautomeric forms (2T) in the fast chemical exchange process (Table 1).

Due to the high freezing point of DMSO-d6, we acquired the NMR experiments of Ga(III)-photoxenobactin E complex (2) at lower temperatures in a 9:1 deuterated solvent mixture of DMSO-d6:CD3OD (Figure S11). The separation of the two sharp signals (30.80 Hz) corresponding to the C-18 methyl groups of 2A and 2B observed in this solvent mixture at δH 1.55 and 1.63, respectively, was higher than that observed in DMSO-d6 (25.33 Hz). These resonances were subsequently used to characterize the tautomeric equilibrium defined as 2B ⇆ 2A. The relative intensities of the C-18 signals were obtained by deconvolution of the 1H NMR spectra in the temperature range 278.15–308.15 K, as above this temperature coalescence takes place. The relative populations of the two tautomers afforded the equilibrium constant at each temperature, which increases slightly from Ke = 1.37 at 273.15 K to 1.53 at 303.15 K, indicating that the relative population of 2A progressively increases with temperature. A plot of ΔG2B→2A = −RT ln Ke versus temperature gives a straight line (Figure 3A), with the corresponding linear fit yielding the reaction enthalpy (ΔH2B→2A = +2980 ± 570 J mol–1) and reaction entropy (ΔS2B→2A = +13.3 ± 2.0 J mol–1 K–1) for the tautomeric equilibrium. These data indicate that the higher population of the thiol form 2A around room temperature is the result of a favorable entropy contribution, which compensates for the unfavorable enthalpy term. Our DFT calculations, using DMSO as the solvent, display a very small free energy difference between the two local minima, favoring the thiol form by only 0.52 kJ mol–1. This is in agreement with the experimental NMR data.

Figure 3.

Figure 3

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(A) Free energy values versus temperature for the 2B ⇆ 2A tautomeric equilibrium obtained from the integration of C-18 signals in DMSO-d6:CD3OD (9:1) solution; (B–H) Experimental (black) and simulated (red) 1H NMR spectra and rate constants obtained at different temperatures; (I) Eyring plot for the 2B → 2A interconversion. (*) The signal at 1.49 ppm corresponds to the C-18 signal of Ga(III)-piscibactin as the hydrolysis product of Ga(III)-photoxenobactin E.

Kinetics of Tautomeric Exchange

The kinetics of chemical exchange involving the tautomerization process were investigated by simulating the 1H NMR spectra at varying temperatures with the freeware WinDNMR (Figure 3B–H).39 The line widths of the C-18 methyl groups were simulated assuming that they are the result of an exchange process involving two sites with different populations. The analysis was performed using a rather broad temperature range of 293.15 to 323.15 K, incorporating spectra below and above coalescence, which occurs at ∼313 K. The populations of the two forms were obtained from the reaction enthalpy (ΔH2B→2A) and reaction entropy (ΔS2B→2A) values described above. The experimental spectra could be satisfactorily simulated with the rate constants (k2B→2A + k2A→2B) shown in Figure 3. The rate constants for the 2B → 2A interconversion process (k2B→2A) were subsequently obtained by multiplying (k2B→2A + k2A→2B) by the molar fraction of 2A.40 The values of k2B→2A obtained at different temperatures were analyzed using an Eyring plot according to eq 1, where kb and h are Boltzmann and Planck constants, respectively, R is the gas constant, and ΔH‡ and ΔS‡ are the activation enthalpy and activation entropy.41

graphic file with name ic3c04076_m001.jpg 1

The Eyring plot affords a rather large ΔH‡ value of 83.9 ± 3.2 kJ mol–1, which is mainly associated with the energy cost required to break the Ga(III)-O bond in the 2B thione form and reach the transition state. Similar ΔH‡ values were reported for the rotation of carboxylate groups in complexes with the trivalent lanthanide ions.42,43 The large enthalpy cost is compensated for in part by a positive ΔS‡ value (+60.0 ± 2.4 J mol–1 K–1). The sign of ΔS‡ is in agreement with the sign of the reaction entropy for the 2B ⇆ 2A equilibrium, as described above. This entropy change likely arises from the gain in degrees of freedom of the solvent molecules upon exposure of the less electronegative S atom to the solvent. This hypothesis is supported by the fact that the rotation of carboxylate groups in lanthanide complexes proceeds with a negligible activation entropy.42

Solvent Effects

Since solvents are known to influence tautomerism, we proceeded to study this effect on the thione–thiol exchange process by running the NMR spectra in different deuterated solvents (Tables 1 and 2; Figures S12–S19).44 Although the best solubility was obtained in DMSO-d6 and in a DMSO-d6:CD3OD (9:1) mixture, the Ga(III)-photoxenobactin E complex (2) was also soluble in CDCl3 and partially soluble in THF-d8, but it was insoluble in CD3OD and D2O.

Table 2. 1H and 13C NMR Data Assignments of the Ga(III)-Photoxenobactin E Tautomers (2A/B) in DMSO-d6, CDCl3, and THF-d8 (500/125 MHz, 298.15 K)a.

  2A (thiol-form)
 2B (thione-form)
 2A (thiol-form)
 2B (thione-form)
 2A (thiol-form)
 2B (thione-form)
  DMSO-d6
 DMSO-d6
 CDCl3
 CDCl3
 THF-d8
 THF-d8
#-mult δH (mult., J) δC δH (mult., J) δC δH (mult., J) δC δH (mult., J) δC δH (mult., J) δC δH (mult., J) δC
1-C   166.20   166.25   166.7   166.7   167.4   167.4
2-CH 6.67 (m) 123.25 6.67 (m) 122.68 6.86 (d, 8.7) 124.3 6.86 (d, 8.7) 124.3 6.75 (m) 124.2 6.75 (m) 124.2
3-CH 7.32 (m) 135.39 7.33 (m) 135.67 7.32 (ddd, 8.7, 7.2, 1.7) 137.0 7.32 (ddd, 8.7, 7.2, 1.7) 137.0 7.22 (t, 7.8) 135.8 7.22 (t, 7.8) 135.8
4-CH 6.63 (m) 115.27 6.63 (m) 114.73 6.66 (dd, 8.0, 7.2) 116.9 6.66 (dd, 8.0, 7.2) 116.9 6.55 (m) 116.0 6.55 (m) 116.0
5-CH 7.27 (m) 131.67 7.27 (m) 131.71 7.29 (dd, 8.0, 1.7) 132.1 7.29 (dd, 8.0, 1.7) 132.1 7.27 (m) 132.0 7.27 (m) 132.0
6-C   115.39   115.85   115.1   115.1   116.1   116.1
7-C   174.83   176.41   177.9   177.9   177.3   177.3
8-CH2 3.34 (ov) 33.36 3.34 (ov) 33.05 3.19 (br m) 34.5 3.19 (br m) 34.5 3.25 (ov) 34.4 3.25 (ov) 34.4
3.54 (ov)   3.54 (ov) 3.53 (m) 3.53 (m) 3.52 (ov) 3.52 (ov)
9-CH 4.62 (m) 75.67 4.61 (m) 75.72 4.72 75.9 4.72 75.9 4.64 (br m) 76.7 4.64 (br m) 76.7
10-CH2 4.88 (dd, 9.9, 6.6) 68.99 4.88 (dd, 9.9, 6.6) 68.85 4.74 (br m) 70.6 4.74 (br m) 70.6 4.75 (br m) 70.7 4.75 (br m) 70.7
11-CH2 2.99 (m) 37.76 2.99 (m) 37.23 3.16 (br m), 3.42 (br m) 39.4 3.16 39.4 3.13 (ov), 3.42 (ov) 38.6 3.13 (ov), 3.42 (ov) 38.6
3.50 (m) 3.50 (m) 3.42 (br m)
12-CH 3.70 (m)** 68.40 3.62 (m)** 68.74 3.83 (br m) 69.0 3.83 (br m) 69.0 3.81(br m)* 69.0** 3.72 (br m)* 69.3**
13-CH 4.08 (m) 70.07 4.08 (m) 70.16 4.34 (d, 3.8) 71.8 4.34 (d, 3.8) 71.8 4.28 (br m) 71.1 4.28 (br m) 71.1
14-CH2 2.73 (brt, 14.4) 40.31 2.73 (br t, 14.4) 40.83 2.67 (d, 17.0), 3.15 (br m) 42.1 2.67 (d, 17.0) 42.1 2.71 (m) 40.9 2.71 (m) 40.9
2.92 (m) 2.92 (m) 3.15 (br m) 3.04 (ov) 3.04 (ov)
15-C   180.21   181.92   180.2   180.2   182.2   182.2
16-CH2 3.16 (d, 11.5) 38.17 3.34 (ov) 42.41 n.d. 37.8 n.d. 43.4 3.16 (ov) 38.9 3.37 (ov), 4.08 (d, 11.6) 43.5
3.51 (ov) 4.01 (d, 11.1) 3.50 (br m)
17-C   88.80   88.64   90.7   90.7   90.8   90.8
18-CH3 1.51 (s) 24.56 1.61 (s) 25.69 1.84 (s) 27.7 1.84 (s) 27.7 1.68 (s) 25.8 1.76 (s) 26.6
19-C   208.72   222.23   210.3   224.4   208.7   222.7
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a

ov = overlapped resonances; n.d. = not detected; *,** = interchangeable signals.

The NMR spectra of 2 in CDCl3 (Figures S12–S15) showed that the proton and carbon chemical shift of the CH3-18 group coalesce at room temperature at δH 1.84/δC 27.7. The presence of the two tautomeric forms in CDCl3 was suggested by the two C-19 carbon resonances at δC 210.3 and δC 224.4 observed in the HMBC experiment of 2 (Figure S15). This was confirmed by the HMBC correlations between the CH3-18 protons at δH 1.84 and both C-19 carbon resonances at δC 210.3 and δC 224.4. Relative intensities of the HMBC cross-peaks from the CH3-18 protons to the thiol and thione C-19 carbons suggested that the thione-form (2B) was predominant (Figure S15). CDCl3 is less polar than DMSO-d6, and thus the ligation through the oxygen atom appears to be favored in nonpolar solvents.

1D and 2D NMR experiments of 2 recorded in THF-d8 (Figures S16–S19) showed duplicate signals, indicating the presence of the two tautomeric forms in this solvent. For example, two carbon resonances for C-19 at δC 208.7 and δC 222.7, were observed in the HMBC spectrum of 2 that correlated to CH3-18 protons at δH 1.68 and at δH 1.76, respectively (Figure S19). Furthermore, the CH2–16 protons were observed in this solvent as two chemical shift pairs at δC 38.9/δH 3.16 and 3.50 and at δC 43.5/δH 3.37 and 4.08 (Table 2). We could not determine the tautomer ratio of the thiol- and thione forms in this case because the CH3-18 methyl chemical shifts were under the residual protonated species of the deuterated solvent. However, intensities of the cross-peaks between CH3-18 protons and the two carbon resonances for C-19 in the HMBC spectrum (Figure S19) suggest that the thiol-form (2A) in THF-d8 is slightly more abundant in this solvent than in CDCl3.

Thus, experiments in different solvents indicate that the thiol form is more favored in polar solvents. This is reasonable, as coordination through the sulfur atom leaves the more electronegative O atom of the thiocarboxylate group exposed to the solvent, and thus, most likely, the thiol form is more efficiently solvated by polar solvents compared with the thione tautomer.

Conclusions

In summary, in this study, we described for the first time the structure of the Ga(III)-photoxenobactin E complex (2) and its tautomeric equilibrium via NMR analysis where the Ga(III) can be coordinated through either the sulfur or the oxygen of the thiocarboxylate terminal. The thiol-form of Ga(III)-photoxenobactin E results in δC 208.7 (298.15 K, DMSO-d6, 500 MHz) while the thione form results in δC 222.2 (298.15 K, DMSO-d6, 500 MHz). These chemical shift assignments were supported by DFT calculations. 1H NMR line shape analysis of the signal at position C-18 of the Ga(III)-photoxenobactin E complex (2) estimated the temperature of coalescence in DMSO-d6 at 313.15 K. Using WinDNMR software and the Eyring equation, the kinetic parameters for the PxbE-Ga(III) tautomer were calculated. To the best of our knowledge, carbon NMR chemical resonances associated with the Ga(III)-bound thione-form have not been described in the literature. Bode and co-workers25 only observed thiol-associated NMR resonances (δC 212.1 in DMSO-d6) in the thiocarboxylate terminal of apo-photoxenobactin E.

The present work has important implications for ligand design, as we have demonstrated that the thiocarboxylate group can act as an efficient donor for hard metal ions, such as Ga(III), paving the way for the preparation of new ligand families containing this motif. Our results indicate that both O- and S-coordination can take place, with the interconversion among the two forms taking place in the ms time scale.

Acknowledgments

The work was supported by grants PID2021-122732OB-C22/C21 from MCIN/AEI/10.13039/501100011033/FEDER “A way to make Europe” (AEI, Spanish State Agency for Research and FEDER Programme from the European Union). M.B. was supported by grant PID2019-103891RJ-100 from MCIN/AEI/10.13039/501100011033 (Spain). Work at the University of Santiago de Compostela and University of A Coruña was also supported by grants GRC2018/018 and ED431C 2022/39, respectively, from Xunta de Galicia. L.B. thanks Horizon Europe Marie Skłodowska-Curie Actions Postdoctoral Fellowship funded by the European Union (ID: 101066127). L.A. thanks Xunta de Galicia (Spain) for a predoctoral fellowship.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.inorgchem.3c04076.

  • HRMS, IR, and NMR spectra of 2; Cartesian coordinates obtained with DFT calculations (PDF)

Author Contributions

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript.

The authors declare no competing financial interest.

Supplementary Material

ic3c04076_si_001.pdf (6.7MB, pdf)

References

  1. Singh S. Evolution of Metal-Thiocarboxylate Chemistry in 21st Century. J. Mol. Struct. 2021, 1234, 130184. 10.1016/j.molstruc.2021.130184. [DOI] [Google Scholar]
  2. Kato S.; Niyomura O. Group 1–17 Element (except Carbon) Derivatives of Thio-, Seleno- and Telluro-Carboxylic Acids. Top. Curr. Chem. 2005, 251, 13–85. 10.1007/b101006. [DOI] [Google Scholar]
  3. Deivaraj T. C.; Park J. H.; Afzaal M.; O’Brien P.; Vittal J. J. Novel Bimetallic Thiocarboxylate Compounds as Single-Source Precursors to Binary and Ternary Metal Sulfide Materials. Chem. Mater. 2003, 15 (12), 2383–2391. 10.1021/cm031027v. [DOI] [Google Scholar]
  4. Vittal J. J.; Meng T. N. Chemistry of Metal Thio- and Selenocarboxylates: Precursors for Metal Sulfide/Selenide Materials, Thin Films, and Nanocrystals. Acc. Chem. Res. 2006, 39 (11), 869–877. 10.1021/ar050224s. [DOI] [PubMed] [Google Scholar]
  5. Chaturvedi J.; Singh S.; Bhattacharya S.; Nöth H. The Chemistry of Cadmium-Thiocarboxylate Derivatives: Synthesis, Structural Features, and Application as Single Source Precursors for Ternary Sulfides. Inorg. Chem. 2011, 50 (20), 10056–10069. 10.1021/ic200927w. [DOI] [PubMed] [Google Scholar]
  6. Troyano J.; Castillo Ó.; Amo-Ochoa P.; Fernández-Moreira V.; Gómez-García C. J.; Zamora F.; Delgado S. A Crystalline and Free-Standing Silver Thiocarboxylate Thin-Film Showing High Green to Yellow Luminescence. J. Mater. Chem. C 2016, 4 (36), 8545–8551. 10.1039/C6TC02401G. [DOI] [Google Scholar]
  7. Yoshida T.; Izuogu D. C.; Zhang H. T.; Cosquer G.; Abe H.; Wernsdorfer W.; Breedlove B. K.; Yamashita M. Ln-Pt Electron Polarization Effects on the Magnetic Relaxation of Heterometallic Ho- and Er-Pt Complexes. Dalton Trans. 2019, 48 (21), 7144–7149. 10.1039/C8DT03338B. [DOI] [PubMed] [Google Scholar]
  8. Volkov A. A.; Bugaenko D. I.; Bogdanov A. V.; Karchava A. V. Visible-Light-Driven Thioesterification of Aryl Halides with Potassium Thiocarboxylates: Transition-Metal Catalyst-Free Incorporation of Sulfur Functionalities into an Aromatic Ring. J. Org. Chem. 2022, 87 (12), 8170–8182. 10.1021/acs.joc.2c00913. [DOI] [PubMed] [Google Scholar]
  9. Joshi D. K.; Mishra K. B.; Tiwari V. K.; Bhattacharya S. Synthesis, Structure, and Catalytic Activities of New Cu(i) Thiocarboxylate Complexes. RSC Adv. 2014, 4 (75), 39790–39797. 10.1039/C4RA05290K. [DOI] [Google Scholar]
  10. Liu A.; Si Y.; Dong S.-H.; Mahanta N.; Penkala H. N.; Nair S. K.; Mitchell D. A. Functional Elucidation of TfuA in Peptide Backbone Thioamidation. Nat. Chem. Biol. 2021, 17 (5), 585–592. 10.1038/s41589-021-00771-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Cortese M. S.; Paszczynski A.; Lewis T. A.; Sebat J. L.; Borek V.; Crawford R. L. Metal Chelating Properties of Pyridine-2,6-Bis (Thiocarboxylic Acid) Produced by Pseudomonas spp. and the Biological Activities of the Formed Complexes. Biometals 2002, 15 (2), 103. 10.1023/A:1015241925322. [DOI] [PubMed] [Google Scholar]
  12. Lewis T. A.; Paszczynski A.; Gordon-Wylie S. W.; Jeedigunta S.; Lee C.-H.; Crawford R. L. Carbon Tetrachloride Dechlorination by the Bacterial Transition Metal Chelator Pyridine-2, 6-Bis (Thiocarboxylic Acid). Environ. Sci. Technol. 2001, 35 (3), 552–559. 10.1021/es001419s. [DOI] [PubMed] [Google Scholar]
  13. Delaere D.; Raspoet G.; Nguyen M. T. Thiol - Thione Tautomerism in Thioformic Acid: Importance of Specific Solvent Interactions. J. Phys. Chem. A 1999, 103, 171–177. 10.1021/jp983298c. [DOI] [Google Scholar]
  14. Huang G.; Xia Y.; Li Y. Substituent Effects on the Tautomerism of Monochalcogenocarboxylic Acids XC({double Bond, Long}O)YH (X = H, F, NH2, OH, CN, and CH3; Y = S, Se, and Te): A Theoretical Study. J. Mol. Struct.: THEOCHEM 2009, 896 (1–3), 80–84. 10.1016/j.theochem.2008.11.003. [DOI] [Google Scholar]
  15. Kaur G.; Vikas The Mechanism of Tautomerisation and Geometric Isomerisation in Thioformic Acid and Its Water Complexes: Exploring Chemical Pathways for Water Migration. Phys. Chem. Chem. Phys. 2014, 16 (44), 24401–24416. 10.1039/C4CP03481C. [DOI] [PubMed] [Google Scholar]
  16. Savant V. V.; Gopalakrishnan J.; Patel C. C. Metal Monothiobenzoates. Inorg. Chem. 1970, 9 (4), 748–751. 10.1021/ic50086a011. [DOI] [Google Scholar]
  17. Baranwal B. P.; Gupta T. Synthesis and Spectral Characterization of Some Oxo-Centered, Trinuclear Mixed-Valence Iron Thiocarboxylates. Spectrochim. Acta, Part A 2003, 59 (4), 859–865. 10.1016/S1386-1425(02)00232-9. [DOI] [PubMed] [Google Scholar]
  18. Shang G.; Hampden-Smith M. J.; Duesler E. N. Synthesis and Characterization of Gallium Thiocarboxylates as Novel Single-Source Precursors to Gallium Sulfide Thin Films by Aerosol-Assisted CVD. Chem. Commun. 1996, 15, 1733–1734. 10.1039/cc9960001733. [DOI] [Google Scholar]
  19. Jaśkowska E.; Dobrzycki Ł.; Rzepiński P.; Ziemkowska W. Aluminum, Gallium and Indium Thiobenzoates: Synthesis, Characterization and Crystal Structures. J. Sulfur Chem. 2015, 36 (3), 326–339. 10.1080/17415993.2015.1025073. [DOI] [Google Scholar]
  20. Lee C.-H.; Lewis T. A.; Paszczynski A.; Crawford R. L. Identification of an Extracellular Catalyst of Carbon Tetrachloride Dehalogenation from Pseudomonas stutzeri Strain KC as Pyridine-2, 6-Bis (Thiocarboxylate). Biochem. Biophys. Res. Commun. 1999, 261 (3), 562–566. 10.1006/bbrc.1999.1077. [DOI] [PubMed] [Google Scholar]
  21. Matthijs S.; Tehrani K. A.; Laus G.; Jackson R. W.; Cooper R. M.; Cornelis P. Thioquinolobactin, a Pseudomonas Siderophore with Antifungal and Anti-Pythium Activity. Environ. Microbiol. 2007, 9 (2), 425–434. 10.1111/j.1462-2920.2006.01154.x. [DOI] [PubMed] [Google Scholar]
  22. Liu Y.-P.; Cai X.-H.; Feng T.; Li Y.; Li X.-N.; Luo X.-D. Triterpene and Sterol Derivatives from the Roots of Breynia fruticosa. J. Nat. Prod. 2011, 74 (5), 1161–1168. 10.1021/np2000914. [DOI] [PubMed] [Google Scholar]
  23. Dong L.-B.; Rudolf J. D.; Kang D.; Wang N.; He C. Q.; Deng Y.; Huang Y.; Houk K. N.; Duan Y.; Shen B. Biosynthesis of Thiocarboxylic Acid-Containing Natural Products. Nat. Commun. 2018, 9 (1), 2362. 10.1038/s41467-018-04747-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Zheng C.-J.; Kalkreuter E.; Fan B.-Y.; Liu Y.-C.; Dong L.-B.; Shen B. PtmC Catalyzes the Final Step of Thioplatensimycin, Thioplatencin, and Thioplatensilin Biosynthesis and Expands the Scope of Arylamine N-Acetyltransferases. ACS Chem. Biol. 2021, 16 (1), 96–105. 10.1021/acschembio.0c00773. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Shi Y.-M.; Hirschmann M.; Shi Y.-N.; Ahmed S.; Abebew D.; Tobias N. J.; Grün P.; Crames J. J.; Pöschel L.; Kuttenlochner W.; Richter C.; Herrmann J.; Müller R.; Thanwisai A.; Pidot S. J.; Stinear T. P.; Groll M.; Kim Y.; Bode H. G. Global Analysis of Biosynthetic Gene Clusters Reveals Conserved and Unique Natural Products in Entomopathogenic Nematode-Symbiotic Bacteria. Nat. Chem. 2022, 14, 701–712. 10.1038/s41557-022-00923-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Souto A.; Montaos M. A.; Rivas A. J.; Balado M.; Osorio C. R.; Rodríguez J.; Lemos M. L.; Jiménez C. Structure and Biosynthetic Assembly of Piscibactin, a Siderophore from Photobacterium Damselae Subsp. Piscicida, Predicted from Genome Analysis. Eur. J. Org. Chem. 2012, 2012 (29), 5693–5700. 10.1002/ejoc.201200818. [DOI] [Google Scholar]
  27. Naka H.; Dias G. M.; Thompson C. C.; Dubay C.; Thompson F. L.; Crosa J. H. Complete Genome Sequence of the Marine Fish Pathogen Vibrio Anguillarum Harboring the PJM1 Virulence Plasmid and Genomic Comparison with Other Virulent Strains of V. Anguillarum and V. ordalii. Infect. Immun. 2011, 79 (7), 2889–2900. 10.1128/IAI.05138-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Balado M.; Lages M. A.; Fuentes-Monteverde J. C.; Martínez-Matamoros D.; Rodríguez J.; Jiménez C.; Lemos M. L. The Siderophore Piscibactin Is a Relevant Virulence Factor for Vibrio Anguillarum Favored at Low Temperatures. Front. Microbiol. 2018, 9, 1766. 10.3389/fmicb.2018.01766. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Ruiz P.; Balado M.; Fuentes-Monteverde J. C.; Toranzo A. E.; Rodríguez J.; Jiménez C.; Avendaño-Herrera R.; Lemos M. L. The Fish Pathogen Vibrio ordalii under Iron Deprivation Produces the Siderophore Piscibactin. Microorganisms 2019, 7 (9), 313. 10.3390/microorganisms7090313. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Galvis F.; Ageitos L.; Rodríguez J.; Jiménez C.; Barja J. L.; Lemos M. L.; Balado M. Vibrio Neptunius Produces Piscibactin and Amphibactin and Both Siderophores Contribute Significantly to Virulence for Clams. Front. Cell. Infect. Microbiol. 2021, 11, 750567. 10.3389/fcimb.2021.750567. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Shi Y.-M.; Hirschmann M.; Shi Y.-N.; Bode H. B. Cleavage Off-Loading and Post-Assembly-Line Conversions Yield Products with Unusual Termini during Biosynthesis. ACS Chem. Biol. 2022, 17 (8), 2221–2228. 10.1021/acschembio.2c00367. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Hider R. C.; Kong X. Chemistry and Biology of Siderophores. Nat. Prod. Rep. 2010, 27 (5), 637–657. 10.1039/b906679a. [DOI] [PubMed] [Google Scholar]
  33. Lemos M. L.; Balado M. Iron Uptake Mechanisms as Key Virulence Factors in Bacterial Fish Pathogens. J. Appl. Microbiol. 2020, 129 (1), 104–115. 10.1111/jam.14595. [DOI] [PubMed] [Google Scholar]
  34. Shannon R. D. Revised Effective Ionic Radii and Systematic Studies of Interatomic Distances in Halides and Chalcogenides. Acta Crystallogr., Sect. A 1976, 32 (5), 751–767. 10.1107/S0567739476001551. [DOI] [Google Scholar]
  35. Chitambar C. R. Gallium and Its Competing Roles with Iron in Biological Systems. Biochim. Biophys. Acta, Mol. Cell Res. 2016, 1863 (8), 2044–2053. 10.1016/j.bbamcr.2016.04.027. [DOI] [PubMed] [Google Scholar]
  36. Kato S.; Kawahara Y.; Kageyama H.; Yamada R.; Niyomura O.; Murai T.; Kanda T. Thion (RCSOH), Selenon (RCSeOH), and Telluron (RCTeOH) Acids as Predominant Species. J. Am. Chem. Soc. 1996, 118 (6), 1262–1267. 10.1021/ja953035l. [DOI] [Google Scholar]
  37. Wang R.; Xie K. J.; Fu Q.; Wu M.; Pan G. F.; Lou D. W.; Liang F. S. Transformation of Thioacids into Carboxylic Acids via a Visible-Light-Promoted Atomic Substitution Process. Org. Lett. 2022, 24 (10), 2020–2024. 10.1021/acs.orglett.2c00481. [DOI] [PubMed] [Google Scholar]
  38. Pierens G. K. 1H and 13C NMR Scaling Factors for the Calculation of Chemical Shifts in Commonly Used Solvents Using Density Functional Theory. J. Comput. Chem. 2014, 35 (18), 1388–1394. 10.1002/jcc.23638. [DOI] [PubMed] [Google Scholar]
  39. Reich H. J. WinDNMR: Dynamic NMR Spectra for Windows. J. Chem. Educ. 1995, 72 (12), 1086. 10.1021/ed072p1086.1. [DOI] [Google Scholar]
  40. Březina V.; Hanyková L.; Velychkivska N.; Hill J. P.; Labuta J. NMR Lineshape Analysis Using Analytical Solutions of Multi-State Chemical Exchange with Applications to Kinetics of Host–Guest Systems. Sci. Rep. 2022, 12 (1), 17369. 10.1038/s41598-022-20136-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Laidler K. J.; King M. C. Development of Transition-State Theory. J. Phys. Chem. A 1983, 87 (15), 2657–2664. 10.1021/j100238a002. [DOI] [Google Scholar]
  42. Fusaro L.; Luhmer M. An Oxygen-17 Dynamic NMR Study of the Pr-DOTA Complex. Dalton Trans. 2014, 43 (3), 967–972. 10.1039/C3DT52533C. [DOI] [PubMed] [Google Scholar]
  43. Mayer F.; Platas-Iglesias C.; Helm L.; Peters J. A.; Djanashvili K. 17O NMR and Density Functional Theory Study of the Dynamics of the Carboxylate Groups in DOTA Complexes of Lanthanides in Aqueous Solution. Inorg. Chem. 2012, 51 (1), 170–178. 10.1021/ic201393n. [DOI] [PubMed] [Google Scholar]
  44. Bahadoor A.; Watt S.; Rajotte I.; Bates J. Tautomerization and Isomerization in Quantitative NMR: A Case Study with 4-Deoxynivalenol (DON). J. Agric. Food Chem. 2022, 70 (8), 2733–2740. 10.1021/acs.jafc.1c08053. [DOI] [PubMed] [Google Scholar]

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