In Vitro Metabolism and Analytical Characterization of SLU‐PP‐332 and SLU‐PP‐915: Novel Pan‐ERR Agonists With Doping Potential

ABSTRACT

Rationale

Estrogen‐related receptor (ERR) agonists such as the drug candidates SLU‐PP‐332 and SLU‐PP‐915 are currently being investigated as exercise mimetics, given their ability to trigger human physiological processes similar to those initiated by actual physical activity. This capability prompted the consideration of these compounds as drugs potentially relevant for sports drug testing programs.

Methods

The two pan‐ERR agonists SLU‐PP‐332 and SLU‐PP‐915 were characterized using liquid chromatography‐high resolution (tandem) mass spectrometry (LC–HRMS/MS). Furthermore, the in vitro metabolic transformation products of both compounds prepared by means of human liver S9 fraction (S9 fraction) and human liver microsomes (HLMs) were analyzed. In addition, selected metabolites of SLU‐PP‐915 were synthesized and their structures were analyzed by nuclear magnetic resonance (NMR) spectroscopy.

Results

A total of nine metabolites were identified for SLU‐PP‐332, consisting of six Phase‐I metabolites and three Phase‐II conjugates. Conversely, the analysis of SLU‐PP‐915 yielded only Phase‐I transformation products, with a total of seven metabolites identified. In both cases, an in‐depth structural elucidation was conducted to obtain a comprehensive overview of the detected metabolites. Furthermore, three metabolites of SLU‐PP‐915 were confirmed through chemical synthesis and NMR.

Conclusion

The results obtained in this study gave an in‐depth view into the analysis and in vitro metabolism of the newly developed pan‐ERR agonists SLU‐PP‐332 and SLU‐PP‐915. This may help to uncover the illicit use of these novel compounds as potential performance‐enhancing substances.

1. Introduction

The investigation of new physiological targets is a crucial aspect of drug research and development. One of these potential new targets is the class of estrogen‐related receptors (ERRs) [1]. These orphan receptors play an important role in energy homeostasis and lipid metabolism, and they have been associated with various diseases, such as metabolic or skeletal muscle disorders [2, 3, 4, 5]. Despite this receptor family's structural and sequence similarities to the estrogen receptors (ERs), they are not activated by their endogenous agonists [2, 6]. Therefore, potential ERR agonists have been investigated within recent years. Two of these newly developed agonists, SLU‐PP‐332 and SLU‐PP‐915, have been shown to target all three isoforms of the ERR (ERRα, ERRβ, and ERRγ), suggesting their potential for use in the treatment of various diseases [7]. Both compounds were investigated as pharmacological exercise mimetics, since these compounds were found to induce effects analogous to those of actual physical activity. SLU‐PP‐332 was demonstrated to promote an increase in type IIa oxidative skeletal muscle fibers and improve exercise endurance in animal models [8]. Similarly, SLU‐PP‐915 exhibited effects on gene expression and agonistic activity in vitro and in vivo [9]. These properties render them not only interesting as potential therapeutics to manage obesity or metabolic disorders, but also as potential substances of abuse in the context of sports. In order to maintain an effective anti‐doping system, it is vital that such compounds are analytically characterized prior to their market launch and, thus, availability to athletes. Non‐approved substances that exhibit the potential for performance enhancement can be subject of the World Anti‐Doping Agency (WADA) Prohibited List [10], and proactive analytical consideration in preventive anti‐doping research is critical.

When implementing new substances into existing doping control methods, it is essential to consider not only the intact compounds as target analytes but also their potential metabolites as these may allow for superior detection windows and improved sensitivity compared to approaches that exclusively consider the unmetabolized material [11]. In order to investigate the metabolic pathways of such compounds, in vitro metabolic approaches represent commonly employed first methods for the identification of potential metabolites. To date, limited data on the metabolism of SLU‐PP‐332 are available; for SLU‐PP‐915 no such studies are published [12]. Moreover, there is an absence of literature describing their mass spectrometric behavior, which would be of particular interest with regard to the evaluation of doping control urine samples.

This study presents the mass spectrometric characterization of SLU‐PP‐332 and SLU‐PP‐915. A thorough investigation was conducted into the in vitro metabolic behavior of both compounds. This investigation involved the identification and characterization of their potential metabolites. Furthermore, the chlorinated analog to SLU‐PP‐915 (referred to as SLU‐PP‐915‐Cl) was synthesized for its use as an internal standard (IS). In addition, the synthesis of selected metabolites of SLU‐PP‐915 was undertaken to validate their structures and to produce reference material for use in routine doping control analysis.

2. Methods

2.1. Chemicals and Reagents

SLU‐PP‐332 was purchased from Hycultec (Beutelsbach, Germany). 2‐chloroaniline, cesium carbonate, dimethyl formamide (DMF), 2‐fluoroaniline, hydrogen peroxide, magnesium chloride, N,N‐diisopropyl ethylamine (DIPEA), lithium hydroxide (LiOH), potassium dihydrogen phosphate, sodium hydroxide (NaOH), D‐saccharic acid‐1,4‐lactone (SL), and uridine diphosphate glucuronic acid (UDGPA) were obtained from Sigma Aldrich (St. Louis, MO, USA). 5‐bromothiophene‐2‐carboxylic acid, 1,4‐dioxane, human liver microsomes (HLMs), and human liver S9 fraction (S9 fraction) were obtained from Thermo Scientific (Bremen, Germany). Celite, magnesium sulfate (MgSO₄), and tetrahydrofuran (THF) were purchased from Carl Roth (Karlsruhe, Germany). Ammonium acetate (AcNH₄), dichloromethane (DCM), dimethyl sulfoxide (DMSO), TBTU, and tetrakis (triphenlyphosphine)palladium(0) ((PPh₃)₄Pd) were acquired from Merck (Darmstadt, Germany). Acetonitrile (ACN), acetic acid, cyclohexane, ethyl acetate (EtOAc), formic acid (FA), and n‐pentane were obtained from VWR Chemicals (Langenfeld, Germany). 1,3‐Phenyldiboronic acid was purchased from BLD Pharm (Reinbeck, Germany), and the nicotinamide adenine dinucleotide phosphate (NADPH) regenerating system was purchased from Promega (Madison, WIS, USA). MeOH was purchased from J.T.Baker (Phillipsburg, New Jersey, USA). Hydrogen gas (99.999%) was from Praxair (Düsseldorf, Germany). Ultrapure water was received from a Barnstead GenPure xCAD Plus from Thermo Scientific (Bremen, Germany).

Column chromatography was performed using silica gel (63–200 μm) from Supelco (Sigma Aldrich, St. Louis, Missouri, USA). For reaction control and control of the column chromatography, thin layer chromatography (TLC) plates were used from Merck (Darmstadt, Germany). Chromabond C18 6‐cc SPE cartridges were purchased from Macherey‐Nagel (Düren, Germany).

2.2. NMR Spectroscopy

For nuclear magnetic resonance (NMR) spectroscopy, a Bruker Avance I 300 and Bruker Avance III 499 system was used. ¹H NMR‐spectra were acquired at a frequency of 300.1 or 499.9 MHz, while ¹³C NMR‐spectra were acquired at a frequency of 125.7 MHz. Peak assignments were facilitated by two‐dimensional spectra (H,H‐COSY, H,C‐HMBC, H,C‐HMQC). The chemical shift σ and the coupling constant 3J or 4J are indicated in parts per million and in hertz, respectively, and the multiplicity is classified as singlet (s), doublet (d), triplet (t), doublet doublet (dd), triplet triplet (tt), and multiplet (m). Spectra of all synthesized products are presented in the Supporting Information.

2.3. Synthesis

General procedure for the amidation of the respective aniline halides (Figure 1a).

FIGURE 1.

Synthesis of SLU‐PP‐915 and IS SLU‐PP‐915Cl. Reaction conditions: (a) aniline halide, TBTU, DiPEA, DMF, RT, overnight; (b) 1,3 phenyldiboronic acid, Cs₂CO₃, (PPh₃)₄Pd, DMF, H₂O, 100°C, overnight.

In a baked‐out flask, 5‐bromothiophene‐2‐carboxylic acid 1 (1 eq) was dissolved in dry DMF (1 mL/mmol) followed by the addition of TBTU (1.0 eq) and DiPEA (2.5 eq). After 20 min, the respective aniline halide (1.2 eq) was added, and the mixture was stirred overnight at room temperature (RT). The reaction was stopped by addition of H₂O and was then extracted using EtOAc (3×). The combined organic phases were dried over MgSO₄, and afterwards, the solvent was removed under reduced pressure. The crude product was then purified using column chromatography (n‐pentane: EtOAc = 5:1).

5‐bromo‐N‐(2‐fluorophenyl)thiophene‐2‐carboxamide 2a was synthesized starting from 2‐fluoroaniline (0.64 g; 5.8 mmol) and was isolated as a gray solid with a yield of 0.89 g (61.8%).

¹H NMR (DMSO‐d₆; 500 MHz): δ 7.22 (m, 1H), 7.30 (m, 1H), 7.37 (d; J = 4.0 Hz, 1H), 7.58 (t; J = 7.7 Hz, 1H), 7.87 (d; J = 4.0 Hz, 1H), 10.25 (s, 1H).

¹³C NMR (DMSO‐d₆; 125 MHz): δ 116.28 (C_{quat .}), 118.50 (CH), 124.88 (C_{quat .}), 125.34 (CH), 127.74 (C_{quat .}), 130.80 (C_{quat .}), 132.24 (C_{quat .}), 141.736 (CH), 155.00 (CH), 157.45 (CH), 159.42 (CH).

5‐bromo‐N‐(2‐chlorophenyl)thiophene‐2‐carboxamide 2b was synthesized starting from 2‐chloroaniline (0.30 g; 2.3 mmol) and was isolated as a yellowish oil with a yield of 0.44 g (72.0%).

¹H NMR (DMSO‐d₆; 500 MHz): δ 7.32 (t J = 7.7 Hz, 1H), 7.39 (m, 2H), 7.55 (m, 2H), 7.86 (d; J = 3.9 Hz, 1H), 10.21 (s, 1H).

¹³C NMR (DMSO‐d₆; 125 MHz): δ 118.44 (CH), 128.04 (C_{quat .}), 128.39 (C_{quat .}), 129.28 (C_{quat .}), 130.13 (C_{quat .}), 130.23 (CH), 130.69 (C_{quat .}), 132.25 (C_{quat .}), 134.65 (CH), 141.35 (CH), 159.46 (CH).

General Procedure for the Suzuki Cross Coupling Reaction (Figure 1b)

The respective thiophene‐2‐carboamide (1.0 eq), 1,3 phenylenediboronic acid (1.3 eq), and Cs₂CO₃ (2.0 eq) were dissolved in 1,4‐dioxane (5 mL/mmol) and H₂O (2 mL/mmol). Following the addition of (PPh₃)₄Pd (1 mol‐%), the reaction mixture stirred under reflux overnight. Afterwards, the reaction was cooled to RT and was then filtered over Celite. The solvent was removed under reduced pressure, and the crude product was washed with cyclohexane and DCM until the triphenylphosphine oxide residue vanished (screened via LC‐HRMS).

(3‐(5‐((2‐fluorophenyl)carbamoyl)thiophen‐2‐yl)phenyl)boronic acid SLU‐PP‐915 was synthesized starting from 2a (0.20 g; 0.7 mmol) and was found to form a grayish solid with a yield of 0.17 g (72.3%).

¹H NMR (DMSO‐d₆; 500 MHz): δ 7.22 (m, 2H), 7.30 (m, 3H), 7.6 (m, 3H), 7.76 (d; J = 8.0 Hz, 1H), 7.80 (d; J = 3.9 Hz, 1H), 8.09 (m, 2H), 10.24 (s, 1H).

¹³C NMR (DMSO‐d₆; 125 MHz): δ 116.25 (CH), 116.44 (CH), 123.11 (CH), 124.80 (CH), 125.86 (CH), 126.05 (C_{quat .}), 126.45 (CH), 127.39 (CH), 127.77 (CH), 130.83 (CH), 131.17 (CH), 134.44 (C_{quat .}), 139.20 (C_{quat .}), 147.94 (C_{quat .}), 155.12 (C_{quat .}), 157.57 (C_{quat .}), 160.30 (C_{quat .}).

(3‐(5‐((2‐chlorophenyl)carbamoyl)thiophen‐2‐yl)phenyl)boronic acid SLU‐PP‐915‐Cl was synthesized starting from 2b (0.30 g; 1.0 mmol). It was isolated as a yellow solid with a yield of 0.23 g (64.7%).

¹H NMR (DMSO‐d₆; 500 MHz): δ 7.31 (m, 2H), 7.39 (m, 4H), 7.56 (m, 3H), 7.76 (d; J = 7.7 Hz, 1H), 7.88 (d; J = 3.9 Hz, 1H), 8.05 (s, 1H), 10.22 (s, 1H).

¹³C NMR (DMSO‐d₆; 125 MHz): δ 118.48 (C_{quat .}), 126.27 (CH), 127.81 (CH), 128.01 (CH), 128.33 (CH), 129.22 (CH), 129.74 (CH), 130.12 (CH), 130.22 (C_{quat .}), 130.69 (CH), 132.22 (CH), 132.22 (CH), 134.58 (CH), 134.69 (C_{quat .}), 138.43 (C_{quat .}), 141.40 (C_{quat .}), 149.09 (C_{quat .}).

5‐(3‐Hydroxyphenyl)thiophene‐2‐Carboxylic Acid M1

SLU‐PP‐915 (50 mg; 0.15 mmol; 1.0 eq) was dissolved in a solution of NaOH (6 M; 1 mL) and then H₂O₂ (30% in H₂O; 1 mL) was added. The reaction mixture was stirred for 2 h at RT. Afterwards, LiOH (31 mg; 0.74 mmol; 5 eq) as well as THF (1.2 mL) and MeOH (0.2 mL) were added. The reaction was heated up to 70°C and was then stirred for 4 h. After cooling to RT, the reaction mixture was diluted with H₂O and was then extracted with EtOAc (3×). The combined organic phases were dried over MgSO₄ and afterwards the solvent was removed in vacuo. The crude product was then purified using a Chromabond C18 SPE cartridge (6 cc) eluting with ACN/H₂O (30% to 80% ACN). After drying under a N₂ stream the final product was isolated as a beige solid in quant. yield.

¹H NMR (DMSO‐d₆; 500 MHz): δ 6.80 (m, J = 7.9 Hz, 1H), 7.08 (s, 1H), 7.16 (d; J = 7.6 Hz, 1H), 7.26 (m, 1H), 7.45 (d; J = 3.6 Hz, 1H), 7.70 (d; J = 3.5 Hz, 1H).

¹³C NMR (DMSO‐d₆; 125 MHz): δ 112.92 (C_{quat .}), 116.50 (C_{quat .}), 117.18 (C_{quat .}), 124.84 (C_{quat .}), 130.87 (C_{quat .}), 133.51 (CH), 134.44 (CH), 134.73 (C_{quat .}), 150.40 (CH), 158.42 (CH), 163.25 (CH).

5‐(3‐Boronophenyl)thiophene‐2‐Carboxylic Acid M3

To a solution of SLU‐PP‐915 (50 mg; 0.15 mmol; 1.0 eq) dissolved in THF (1.2 mL), MeOH (0.2 mL) and H₂O (0.6 mL), LiOH (62 mg; 1.47 mmol; 10 eq) was added and the mixture was stirred at 70°C for 6 h. Afterwards, the reaction was cooled down to RT and was then extracted with EtOAc (3×). The combined organic phases were dried using MgSO₄ and the solvent was removed under reduced pressure. The residue was further purified using a Chromabond C18 SPE cartridge (6 cc) eluting with ACN/H₂O (30% to 80% ACN). Finally, the product was isolated after drying under a N₂ stream and was found to produce a beige solid in quant. yield.

¹H NMR (D₂O; 500 MHz): δ 7.48 (m, 3H), 7.58 (d; J = 7.2 Hz, 1H), 7.84 (s, 1H), 8.38 (s, 1H).

¹³C NMR (D₂O; 125 MHz): δ 123.32 (CH), 124.50 (CH), 128.10 (CH), 128.73 (CH), 131.92 (CH), 132.07 (CH), 134.35 (C_{quat .}), 139.31 (C_{quat .}), 132.24 (C_{quat .}), 147.37 (C_{quat .}), 149.82 (C_{quat .}).

N‐(2‐Fluorophenyl)‐5‐(3‐Hydroxyphenyl)thiophene‐2‐Carboxamide M4

SLU‐PP‐915 (50 mg; 0.15 mmol; 1.0 eq) was dissolved in a solution of NaOH (6 M; 1 mL) and H₂O₂ (30% in H₂O; 1 mL). Following, the reaction mixture was stirred for 2 h at RT. Without further purification, the reaction mixture was loaded onto a Chromabond C18 SPE cartridge (6 cc) and eluted with ACN/H₂O (0% to 20% ACN). The solvent was removed using an N₂ stream to afford the final product as a brown solid in quant. yield.

¹H NMR (DMSO‐d₆; 500 MHz): δ 6.79 (d, J = 8.0 Hz, 1H), 7.13 (m, 2H), 7.27 (m, 5H), 7.52 (d; J = 3.8 Hz, 1H), 7.60 (t, J = 7.6 Hz, 1H), 7.98 (d; J = 3.6 Hz, 1H), 10.14 (s, 1H).

¹³C NMR (DMSO‐d₆; 125 MHz): δ 113.43 (CH), 115.67 (CH), 116.28 (CH), 116.47 (CH), 117.32 (CH), 124.54 (CH), 124.98 (CH), 125.22 (C_{quat .}), 127.57 (CH), 130.87 (CH), 131.36 (CH), 134.23 (C_{quat .}), 137.06 (C_{quat .}), 149.96 (C_{quat .}), 155.04 (C_{quat .}), 157.51 (C_{quat .}), 160.85 (C_{quat .}).

2.4. In Vitro Metabolic Assay

In vitro incubation experiments were conducted using a protocol slightly deviating from the protocol described by Kuuranne et al. [13]. HLMs and S9 fraction were used to perform both Phase‐I and Phase‐II metabolism and for NADPH supply a NADPH regenerating system (NADPH reg system) was used. SLU‐PP‐332 was diluted in MeOH, while SLU‐PP‐915 and SLU‐PP‐915‐Cl were diluted in DMSO to produce stock solutions with concentrations of 1 and 2 mg/mL, respectively. In order to obtain suitable working solution with a concentration of 200 μM, the stock solutions were diluted in 50 mM phosphate buffer (pH 7.4) containing 5 mM MgCl₂. All incubation experiments were conducted in triplicate, using 10 μL working solution, 10 μL NADPH regenerating system (50 mM), 5 μL HLM (20 mg/mL), and 5 μL S9 fraction (20 mg/mL) for Phase‐I incubation experiments. After addition of 20 μL phosphate buffer for a total volume of 50 μL, all samples were incubated at 37°C for 24 h.

For Phase‐II metabolism, additional 5 μL of HLM and S9 fraction, as well as 10 μL of UDGPA (50 mM), 10 μL SL (50 mM), and 10 μL PAPS (20 μM) were added and the mixture was again incubated at 37°C for 24 h. To verify the results obtained in this study, blank samples either excluding enzymes (enzyme blank) or excluding substrate (substrate blank) were prepared. These samples were also used to identify metabolites obtained from non‐enzymatic transformations. Metabolic reactions were stopped by the addition of 150 μL ice‐cold ACN. The supernatant was harvested after centrifugation (17 000 × g, 5 min) and transferred into a fresh tube. After drying using a vacuum centrifuge (45°C, 45 min), the samples were reconstituted in 100 μLH₂O:ACN (90:10 v/v).

2.5. Liquid Chromatography‐High Resolution (Tandem) Mass Spectrometry (LC–HRMS/MS)

LC‐HRMS/MS measurements were performed using a Vanquish UHPLC system coupled to an Orbitrap Exploris 480 mass spectrometer both manufactured by Thermo Fisher (Bremen, Germany). The HPLC system was equipped with an EC 4/3 Nucleoshell RP 18 Plus guard column (4 × 3 mm, 5‐μm particle size) from Macherey–Nagel (Düren, Germany) and a Poroshell 120 EC C18 column (3.0 × 50 mm, 2.7 μm) by Agilent (Santa Clara, California, USA). The chromatographic conditions were optimized for the different compounds analyzed in this study, using 50 mM AcNH₄‐buffer as eluent A in the case of SLU‐PP‐332 and 0.1% FA in H₂O in the case of SLU‐PP‐915 and SLU‐PP‐915‐Cl. In any case, 0.1% FA in ACN was used as eluent B. The gradient elution with each solvent system started at 0% B and was then increased to 100% B within 10 min, where it was held for 2 min. After returning to starting conditions within 0.01 min, the column was re‐equilibrated for 2 min. A flow of 0.3 mL/min was applied. The injection volume was 10 μL.

The HRMS parameters were optimized to obtain the best possible results for each analyte. A heated electrospray ionization (ESI) source was used that was either run in positive or negative ionization mode with a voltage of 3000 or −2600 V, respectively. Both full scan data and product ion scans were monitored. In full scan mode, the system operated at a resolution of 60 000 full width at half maximum (FWHM) and a scan range of m/z 80–800. Product ion scans were generated using parallel reaction monitoring (PRM) at a resolution of 30 000 FWHM. The isolation window was set to m/z 1.3. For higher energy collisional dissociation (HCD) in positive and negative ionization mode, normalized collision energies of 40% were used. In addition, pseudo MS³ experiments were conducted, using in‐source fragmentation. Nitrogen was used as collision gas and was generated by a CMC nitrogen generator (Eschborn, Germany). The HRMS was regularly calibrated using the Pierce Flex Mix calibration solution from Thermo Fisher (Bremen, Germany).

3. Results and Discussion

3.1. Synthesis of Reference Material and Mass Spectrometric Characterization

The mass spectrometric characterization is a crucial part of the implementation of new compounds into existing doping control methods or to establish novel methods for those substances. SLU‐PP‐332 has been shown to be detectable in both positive and negative ionization modes. However, with regard to the results obtained for the in vitro metabolic transformations, positive ionization was selected for further analysis.

In positive ionization mode, SLU‐PP‐332 was detectable as [M+H]⁺ (m/z 291.1142). Evaluating MS² data, m/z 121.0288 was identified as the most abundant peak and is most likely produced by α‐cleavage of the hydrazide group. Interestingly, the counterpart at m/z 171.0924 was also detected. Probably, this ion is produced by transferring a proton from the ion at m/z 121.0288 by forming a ion/neutral complex as described in the literature [14, 15]. This product ion is suggested to further eliminate ammonia (17 u) to produce a signal at m/z 154.0657. Another α‐cleavage, most likely occurring at the keto group, yielded the product ion at m/z 197.0718. The results obtained for SLU‐PP‐332 are summarized in Figure 2.

FIGURE 2.

(a) Extracted ion chromatogram (EIC) of SLU‐PP‐332; (b) MS² spectrum obtained for SLU‐PP‐332; (c) proposed dissociation pathway of SLU‐PP‐332. Measurements were conducted in positive ionization mode.

Due to the limited availability of SLU‐PP‐915 reference material, it was synthesized in‐house using a synthesis approach previously described in the literature [9]. In brief, 2‐fluoroaniline 1a was reacted with 5‐bromothiophene‐2‐carboxylic acid to produce intermediate 2a. Subsequently, SLU‐PP‐915 was formed by transforming 2a using a Suzuki cross coupling reaction. The IS was synthesized analogously, starting from the chloro analog 1b. Both reaction procedures are outlined in Figure 1.

For SLU‐PP‐915 and the IS, the negative ionization mode was chosen for characterization and further analysis of its in vitro metabolic behavior. As shown in Figure 3, both compounds display comparable chromatographic properties and a similar dissociation pattern. Consequently, the mass spectrometric characterization is addressed at unison. Both compounds produce a major product ion at m/z 203.0345 which is most probably attributed to the α‐cleavage at the keto function. Pseudo‐MS³ experiments further indicate that this product ion undergoes further dissociation by cleaving either H₂O (18 u) or by releasing its boronic acid function (44 u) to produce the product ions at m/z 185.0239 and m/z 159.0276, respectively.

FIGURE 3.

(a) EIC for SLU‐PP‐915 (black) and SLU‐PP‐915‐Cl (blue); (b) MS² spectrum obtained for SLU‐PP‐915 (black) and SLU‐PP‐Cl (blue); (c) proposed dissociation pathway of SLU‐PP‐915. Measurements were conducted in negative ionization mode.

3.2. In Vitro Metabolic Transformation

Within this study, the in vitro metabolic experiments were conducted using established methods involving the usage of HLMs and S9 fraction; further cofactors were added to promote Phase‐II metabolism [13]. However, the in vitro metabolic pathways of SLU‐PP‐332 were already described [12]. This study aimed to provide new insights and further information on the in vitro metabolism of this substance. The metabolic transformations cover a wide range of reactions, including hydroxylation (M1a‐b), bishydroxylation (M2a‐b), bishydroxylation, and reduction (M3a‐b), as well as Phase‐II transformations such as glucuronidation (M4), sulfation (M5), and the subsequent hydroxylation followed by sulfation (M6). In the literature, similar transformations such as hydroxylation, bishydroxylation, combined bishydroxylation and reduction, and glucuronidation were described. However, hydroxylation with subsequent glucuronidation was described beforehand but could not be identified in this study. In contrast to the literature, within this study the formation of different sulfates were accomplished. An overview of the transformation products of SLU‐PP‐332 observed in this study can be found in Table 1.

TABLE 1.

List of the metabolic transformations identified for SLU‐PP‐332, including the product ions obtained after MS² analysis. For identification, signals within a maximum mass error of 5 ppm were accepted. MS² experiments were conducted using a normalized collision energy of 40%. For each metabolite the most abundant signal is highlighted in bold.

compound	Metabolic transformation	[M + H]+ (theo.) (m/z)	Formula	RT (min)	Product ions (exp.)	Proposed formula
SLU‐PP‐332		291.1128	C18H15N2O2 +	6.71	197.0718	C12H9N2O+
					171.0924	C11H11N2 +
					154.0657	C11H8N+
					121.0288	C 7 H 5 O 2 +
332‐M1a	Hydroxylation	307.1077	C18H15N2O3 +	4.43	187.0874	C11H11N2O+
					170.0606	C11H8NO+
					138.0554	C7H8NO2 +
					121.0288	C 7 H 5 O 2 +
332‐M1b	Hydroxylation	307.1077	C18H15N2O3 +	6.36	197.0717	C12H9N2O+
					171.0924	C11H11N2 +
					154.0658	C11H8N+
					137.0238	C 7 H 5 O 3 +
					111.0444	C6H7O2 +
332‐M2a	Bishydroxylation	323.1026	C18H15N2O4 +	5.17	280.0978	C 17 H 14 NO 3 +
					203.0821	C11H11N2O2 +
					262.0872	C17H12NO2 +
					121.0289	C7H5O2 +
332‐M2b	Bishydro‐xylation	323.1026	C18H15N2O4 +	5.41	280.0978	C 17 H 14 NO 3 +
					262.0875	C17H12NO2 +
					121.0290	C7H5O2 +
332‐M3a	Reduction + Bishydro‐xylation	325.1183	C18H17N2O4 +	4.41	307.1089	C18H15N2O3 +
					188.0715	C11H10NO2 +
					170.0606	C11H8NO+
					161.0603	C10H9O2 +
					121.0288	C 7 H 5 O 2 +
332‐M3b	Reduction + Bishydro‐xylation	325.1183	C18H17N2O4 +	4.64	188.0717	C11H10NO2 +
					170.0608	C11H8NO+
					161.0603	C10H9O2 +
					121.0289	C 7 H 5 O 2 +
332‐M4	Glucuronidation	467.1449	C24H23N2O8 +	5.79	291.1141	C 18 H 15 N 2 O 2 +
					197.0718	C12H9N2O+
					171.0924	C11H11N2 +
					154.0658	C11H8N+
					121.0289	C7H5O2 +
332‐M5	Sulfation	371.0696	C18H15N2O5S+	6.51	291.1141	C18H15N2O2 +
					154.0658	C 11 H 8 N +
					121.0287	C7H5O2 +
332‐M6	Hydroxylation + Sulfation	387.0645	C18H15N2O6S+	6.61	307.1095	C18H15N2O3 +
					197.0718	C12H9N2O+
					171.0924	C11H11N2 +
					154.0658	C 11 H 8 N +
					137.0239	C7H5O3 +

For structural elucidation, the MS² data were further evaluated. As exemplified for metabolite group M1 (hydroxylation; m/z 307.1077) the extracted product ion spectra were used to determine the position of the hydroxy groups within the molecules. For metabolite M1a, the product ions at m/z 121.0288 and 187.0874 most likely result from α‐cleavage of the amide bond, thus indicating hydroxylation on the naphthalene function of the molecule (product ion m/z 187.0874 shown in Figure 1, left). In addition, the product ions m/z 138.0554 and 170.0607, proposed to result from cleavage of the hydrazine group (not shown), further corroborate this assumption. For metabolite M1b, similar results were obtained, showing again α‐cleavage of the amide bond to produce the signals at m/z 137.0238 and m/z 171.0924, indicating hydroxylation on the phenolic function of the molecule. Moreover, additional product ions at m/z 111.0444 and 197.0715, probably resulting from α‐cleavage of the benzaldehyde bond (not shown), substantiate this interpretation. The MS² spectra and the proposed structure of M1a and M1b can be found in Figure 4. The metabolites M2a‐b (bishydroxylation) were analyzed likewise. In both cases, the MS² spectra indicate both transformations on the naphthalene ring of the metabolites.

FIGURE 4.

MS² spectra and proposed structures of the hydroxylated metabolites (a) M1a and (b) M1b of SLU‐PP‐332.

The metabolite group M3a‐b presumably results from bishydroxylation together with a reductive transformation and is already mentioned in the literature [12]. However, due to its unusual nature, this transformation was further investigated within this study. Reviewing the MS² spectra, the metabolization can be located on the naphthalene ring within the molecules. Further, the MS² data support the assignment of losses of water (−18 u), which would contribute to re‐establish the metabolically interrupted aromatic system, supporting the proposed structure of the metabolites shown in Figure 5. As described in the literature, naphthalene itself is known to be transformed in vivo and in vitro to produce its corresponding epoxide, naphthalene oxide, which is then further transformed into the metabolite naphthalene dihydrodiol [16, 17]. A similar transformation pattern may thus be assumed for SLU‐PP‐332 as described here.

FIGURE 5.

MS² spectra and proposed structure of the reduced and bishydroxylated metabolites M3a‐b of SLU‐PP‐332. Losses of H₂O are indicated with blue arrows.

Besides phase‐I transformation products, also Phase‐II conjugates were identified. Both glucuronidation and sulfation of the parent compound were observed, yielding the metabolites M4 (m/z 467.1449) and M5 (m/z 371.0696), respectively. In addition, SLU‐PP332 underwent hydroxylation and subsequent sulfation to form a metabolite at m/z 387.0645 (M6). The dissociation pattern of M6, however, gave similar results to those obtained for metabolite M1b (see Table 1), therefore, M6 can be tentatively assigned to the sulfate conjugate of metabolite M1b. Altogether, a total of nine metabolites comprising six Phase‐I metabolites and three phase‐II conjugates were identified for SLU‐PP‐332. An overview of the metabolic pattern of SLU‐PP 332 is shown in Figure 6.

FIGURE 6.

Overview of the metabolic pattern of SLU‐PP‐332.

The in vitro metabolic behavior of SLU‐PP‐915 was investigated as described for SLU‐PP‐332. However, for this compound only phase‐I transformation products were identified. The metabolic reactions ranged from amide hydrolysis, oxidation of the boronic acid, and hydroxylation to form the metabolites M1 (amide hydrolysis and oxidation), M2 (amide hydrolysis, oxidation, and hydroxylation), M3 (amide hydrolysis), M4 (oxidation), M5a‐b (oxidation and hydroxylation), and M6 (hydroxylation). Remarkably, oxidation to convert the boronic acid moiety into a hydroxy‐group, as observed for the metabolites M4 and M5, is frequently described in the literature [18, 19]. However, it is important to mention that metabolites M1, M3, and M4 were also detectable in the enzyme blank, indicating a non‐enzymatic metabolic pathway. Nevertheless, these metabolites could also be formed in vivo and might thus be no less interesting as putative targets for doping control analysis. An overview of the transformation products of SLU‐PP‐915 observed in this study can be found in Table 2.

TABLE 2.

List of the metabolic transformations identified for SLU‐PP‐915, including the product ions obtained after MS² analysis. For identification, signals within a maximum mass error of 5 ppm were accepted. MS² experiments were conducted using a collision energy of 40%. For each metabolite, the most abundant signal is highlighted in bold.

Compound	Metabolic transformation	[M‐H]− (theo.) (m/z)	Formula	RT (min)	Product ions (exp.)	Proposed formula
SLU‐PP‐915		340.0620	C17H12O3NBFS−	7.11	203.0340	C 10 H 8 O 2 BS −
					185.0249	C10H6BOS−
					159.0284	C10H7S−
SLU‐PP‐915‐Cl (IS)		356.0325	C17H12O3NBClS−	7.76	203.0356	C 10 H 8 O 2 BS −
					185.0248	C10H6BOS−
					159.0284	C10H7S−
915‐M1	Amide hydrolysis + oxidation	219.0121	C11H7O3S−	5.97	175.0220	C 10 H 7 OS −
915‐M2	Amide hydrolysis + oxidation + Hydroxylation	235.0071	C11H7O4S−	5.19	191.0170	C 10 H 7 O 2 S −
915‐M3	Amide hydrolysis	247.0242	C11H8BO4S−	5.56	203.0341	C 10 H 8 O 2 BS −
					159.0272	C10H7S−
915‐M4	Oxidation	312.0500	C17H11FNO2S−	7.55	292.0436	C17H10NO2S−
					174.0220	C 10 H 7 OS −
915‐M5a	Oxidation + hydroxylation	328.0449	C17H11FNO3S−	6.49	308.0385	C 17 H 10 NO 3 S −
					152.0151	C7H3FNO2 −
					175.0232	C10H7OS−
915‐M5b	Oxidation + hydroxylation	328.0449	C17H11FNO3S−	6.83	308.0385	C17H10NO3S−
					191.0181	C 10 H 7 O 2 S −
915‐M6	Hydroxylation	356.0570	C17H12BFNO4S−	6.16	338.0490	C 17 H 10 BFNO 3 S −
					318.0424	C17H9BNO3S−
					152.0163	C7H3FNO2 −

For structural conformation and to provide reference material for routine doping control testing, the metabolites M1, M3, and M4 were synthesized using the reaction conditions shown in Figure 7a. In each case, the chromatographic and mass spectrometric properties of the reference material matched those of the in vitro generated metabolites, confirming the proposed structures. Upon collision‐induced dissociation, all metabolites showed α‐cleavage yielding the most abundant signal within the MS² spectra at m/z 175.0224 for M1, m/z 203.0348 for M3, and m/z 175.0232 for M4. Further, M3 showed a second product ion at m/z 159.0278, which can most likely be assigned to the chemical formula of C₁₀H₇S⁻. M4, however, showed the additional elimination of HF (20 u), generating a product ion at m/z 292.0453. The MS² spectra of each metabolite can be found in Figure 7b.

FIGURE 7.

a) Synthesis of the metabolites M1, M3 and M4. Reaction conditions: (a) NaOH, H₂O₂, H₂O, RT, 2 h; (b) LiOH, THF, MeOH H₂O, 70°C, 4–6 h. b) MS² spectra obtained for the metabolites M1, M3, and M4 of SLU‐PP‐915.

The MS² spectra of the remaining metabolites were further analyzed, to obtain information on the structures of the produced metabolites. As indicated in Table 2, the metabolite group M5 (oxidation and hydroxylation) included two metabolites with varying product ions, indicating different positions of the OH‐groups. As described for the synthesized metabolites M1, M3 and M4, both compounds showed α‐cleavage resulting in the product ions at m/z 175.0232 for M5a and m/z 191.0183 for M5b, that are most likely assigned to the 3‐(thiophen‐2‐yl)phenol ion and its hydroxylated counterpart, respectively. In case of M5a this interpretation can be further underlined by the product ion at m/z 152.0162 which can be attributed to the hydroxylated aniline ring within the molecule. Based on this data, the structures indicated in Figure 8 were proposed.

FIGURE 8.

MS² spectra and proposed structures of the oxidized and hydroxylated metabolites (a) M5a and (b) M5b of SLU‐PP‐915.

As described for M5a‐b, the MS² spectrum of metabolite M6 (hydroxylation) was analyzed and similar to metabolite M5a this compound exhibited a product ion (m/z 152.0163), indicating hydroxylation on the aniline ring of the molecule. Summarizing the results for SLU‐PP‐915, a total of seven metabolites, all of which are assigned to phase‐I transformations, were identified. An overview of the metabolic pattern of SLU‐PP‐915 is shown in Figure 9.

FIGURE 9.

Overview of the metabolic pattern of SLU‐PP‐915.

Within this study, MS² experiments were conducted to yield a deeper understanding on the possible position of introduced functionalities during in vitro metabolism. However, using such experiments is oftentimes insufficient to determine the exact position of the metabolic transformations rather than their approximate location. To address this issue, the suspected metabolite has to be synthesized to confirm its structure, as also exemplified in this work. Additionally, all transformations described in this paper are observed using an in vitro metabolic approach. However, while this approach provides an overview of possible metabolites, it might not fully reflect the authentic human metabolic pattern and, thereby, plasma or urinary metabolic profiles. More complex metabolism studies, such as organ‐on‐a‐chip models or clinical trials, are required to gain comprehensive knowledge of the metabolism of pan‐ERR agonists.

4. Conclusion

The investigation of novel substances that might be misused as doping agents is one of the most crucial tasks in preventive doping research. Within this work, the analytical characterization of the drug candidates SLU‐PP‐332 and SLU‐PP‐915, two compounds that are of particular interest for doping control purposes, was accomplished. The obtained results may help implementing these substances into existing doping control methods. In addition, both substances were metabolized in vitro, and an in‐depth structural elucidation was presented. Moreover, in the case of SLU‐PP‐915, a total of three metabolites were synthesized in order to confirm their proposed structures and to provide reference material. Gathering the results of this work, an important foundation has been laid for enabling the detection of the investigated pan‐ERR agonists and their (potential) metabolites in doping control samples. Further research is needed to address the metabolic behavior in vivo and to ensure effective detection of both substances in human urine samples.

Author Contributions

Mario Thevis: conceptualization, funding acquisition, writing – review and editing, methodology, supervision, project administration. Tristan Möller: conceptualization, conducted experiments, writing – original draft. Oliver Krug: conducted experiments.

Funding

This project was conducted with support of the Manfred‐Donike Institute for Doping Analysis (Cologne, Germany), the Ingeborg‐Gross‐Foundation, and the Federal Chancellery of the Federal Republic of Germany (Berlin, Germany).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: ¹H‐NMR of 2a

Figure S2: 13C‐APT NMR of 2a

Figure S3: 1H‐NMR of 2b

Figure S4: 13C‐APT NMR of 2

Figure S5: 1H‐NMR of SLU‐PP‐915

Figure S6: 13C‐APT NMR of SLU‐PP‐915

Figure S7: 1H‐NMR of SLU‐PP‐915‐Cl

Figure S8: 13C‐APT NMR of SLU‐PP‐915‐Cl

Figure S9: 1H‐NMR of M1

Figure S10: 13C‐APT NMR of M1

Figure S11: 1H‐NMR of M3

Figure S12: 13C‐APT NMR of M3

Figure S13: 13C‐APT NMR of M4

Figure S14: 1H‐NMR of M4

Möller T., Krug O., and Thevis M., “In Vitro Metabolism and Analytical Characterization of SLU‐PP‐332 and SLU‐PP‐915: Novel Pan‐ERR Agonists With Doping Potential,” Rapid Communications in Mass Spectrometry 40, no. 8 (2026): e70039, 10.1002/rcm.70039.

Data Availability Statement

Data supporting this study are available upon reasonable request from the corresponding author (M.T.).