Prediction of human exposure and first-pass metabolism of the fungicide trifloxystrobin using an in vitro intestinal pancreatin and Caco-2/HT29-MTX model.

Prediction of human exposure and first-pass metabolism of the fungicide trifloxystrobin using an in vitro intestinal pancreatin and Caco-2/HT29-MTX model - PMC Skip to main content An official website of the United States government Here's how you know Here's how you know Search Log in Primary site navigation Search Log in Search PMC Full-Text Archive Search in PMC Journal List User Guide As a library, NLM provides access to scientific literature. Inclusion in an NLM database does not imply endorsement of, or agreement with, the contents by NLM or the National Institutes of Health. Learn more: PMC Disclaimer | PMC Copyright Notice Arch Toxicol . 2026 Mar 10;100(6):2381–2395. doi: 10.1007/s00204-026-04340-z Prediction of human exposure and first-pass metabolism of the fungicide trifloxystrobin using an in vitro intestinal pancreatin and Caco-2/HT29-MTX model Eileen Hallscheidt 1, 2 , Kathrin Bothe 3 , Maria Hahn 2 , Kerstin Hartmann 3 , Mario Thevis 1 , Marc Lamshoeft 2, ✉ Author information Article notes Copyright and License information 1 Center for Preventive Doping Research, Institute of Biochemistry, German Sport University Cologne, 50933 Cologne, Germany 2 Metabolism and Kinetics, Environmental Safety, Division Crop Science, Bayer AG, 40789 Monheim, Germany 3 Regulatory Toxicology, Human Safety, Division Crop Science, Bayer AG, 40789 Monheim, Germany ✉ Corresponding author. Received 2025 Oct 24; Accepted 2026 Feb 12; Issue date 2026. © The Author(s) 2026 Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if you modified the licensed material. You do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by-nc-nd/4.0/ . PMC Copyright notice PMCID: PMC13221433  PMID: 41807794 Abstract Conducting toxicological risk assessment of agrochemicals for human health and safety lacks human data and is predominantly based on experimental data obtained from in vivo animal studies. In alignment with the 3R approach (reduce, refine, replace animal testing) and in support of advancing next generation risk assessment of pesticides for human safety, there is a growing demand of implementing new approach methodologies. Therefore, the fungicide trifloxystrobin was investigated concerning human exposure and intestinal first-pass metabolism for the first time using intestinal new approach methodologies. Simulated intestinal fluid containing pancreatin, along with an optimized static in vitro human gut-model (Caco-2/HT29-MTX cell barriers), were established to study the radiolabeled fungicide. A single dose of 10 µM was applied to the in vitro systems, resembling an estimated exposure to humans upon oral uptake from food and feed crops. Trifloxystrobin underwent mainly phase I metabolism to its carboxylic acid metabolite, before entering the systemic circulation consistent with findings from in vivo studies with rodents. The results agree with published in vivo and in vitro data and indicate that the carboxylic acid metabolite is a primary metabolite formed before hepatic phase II conjugation. Moreover, minimal passage through the in vitro gut barriers, facilitated by extensive metabolism indicated low oral bioavailability in humans. The presented in vitro methodologies provide valuable insights into the interplay between human intestinal metabolism and oral bioavailability, exemplified by the active ingredient trifloxystrobin. Moreover, the opportunity of transferring the models to more complex microphysiological technologies such as organ-on-a-chip systems, represents an additional value of these models regarding future studies on oral absorption, metabolism, and organ interactions in vitro. Supplementary Information The online version contains supplementary material available at 10.1007/s00204-026-04340-z. Keywords: New approach methodologies, Regulatory toxicology, Pesticides, Human in vitro gut-model, Next generation risk assessment, Metabolism Introduction Trifloxystrobin (methyl (E)-methoxyimino-{(E)-α-[1-(α,α,α-trifluoro-m-tolyl)ethylidene-aminooxy]-o-toly}acetate, IUPAC) is a broad-spectrum foliar fungicide belonging to the strobilurin class, registered for application on numerous agricultural food and feed crops (Hollingworth 2001 ). It inhibits mitochondrial respiration in plant pathogenic fungi (EPA 1999 ) by targeting mitochondrial complex III (EFSA 2017a ). In toxicokinetic studies with mammals trifloxystrobin showed limited oral absorption (60%), extensive metabolism and rapid excretion (94 – 96% within 48 h) (EFSA 2017a ). One of the main metabolites is the carboxylic acid form of trifloxystrobin (CGA321113, Fig. 1 c) that has been elucidated in rats, laying hens and lactating goats. Additionally, in poultry and ruminants, parent compound residues were detected in milk, muscle, and fat, whereas the metabolite was found in liver, kidney, milk, and egg white, which indicates oral absorption in livestock animals (EFSA 2025 ). In an extensive rat metabolism study, animals were dosed with two uniformly radiolabeled forms of trifloxystrobin ([glyoxyl-phenyl-(U)- 14 C] and [trifluoromethyl-phenyl-(U)- 14 C]) and pooled bile was analyzed to characterize hepatic biotransformation (EFSA 2017b ). Ten metabolites were isolated from bile and partially elucidated. Major hepatic metabolites were glucuronide and sulfate conjugates. Several of these conjugates were derived from hydroxylated parent metabolites that originated from the gut microflora. One biliary metabolite was identified as the carboxylic acid derivative of trifloxystrobin (CGA321113) (EFSA 2017b ). At least four additional biliary metabolites were not fully elicitable. CGA321113 appeared to serve as the principal precursor for subsequent hepatic phase II conjugation reactions in rats. Additionally, a substantial fraction of the administered dose (31 – 47% at high dose levels) was recovered in the feces as unchanged parent compound. Hydrolyzed trifloxystrobin formed in the gut also largely escaped absorption and was eliminated via feces. The absorbed portion underwent extensive degradation and was predominantly eliminated via bile following enterohepatic circulation (EFSA 2017b ). In comparative in vitro metabolism studies using rat and human liver microsomes, no unique metabolites were produced by human microsomes after exposure to trifloxystrobin (EFSA 2017a ). Based on a comprehensive mammalian toxicity study package, an acceptable daily intake (ADI) of 0.1 mg/kg body weight per day has been proposed for humans, applicable for the parent compound and the carboxylic acid metabolite (EFSA 2025 ). However, given the inherent uncertainties in extrapolating mammalian in vivo data to humans, there is a current emphasis on developing new approach methodologies aimed at more accurately predicting human outcomes for next generation risk assessment (NGRA) based on in silico and in vitro approaches. One of the major challenges in the application of NGRA to chemicals is the extrapolation from the in vitro points of departure (PoDs) to human exposure levels associated with the intended use of the chemical. Sufficient exposure to target tissues is a pre-requisite for the elicitation of toxic effects in vivo and its estimation is thus of crucial importance for performing human safety assessments under regulatory aspects. To bridge the gap, physiologically based kinetic (PBK) modelling is routinely used to predict systemic exposure (C max or AUC) from external exposures. However, the predictivity of such a model depends on the reliability of its input parameters. For the first time, human oral ingestion of the fungicide trifloxystrobin has been investigated with respect to first-pass metabolism and oral bioavailability using in vitro intestinal new approach methodologies. Specifically, trifloxystrobin was investigated by simulated intestinal fluid (SIF) and an advanced compartmentalized human in vitro gut-model, comprising Caco-2 and HT29-MTX cells (Fig. 1 a and b). Pancreatin, a mixture of amylases, proteases, lipases, trypsin, and ribonucleases was used to mimic digestive processes before entering the small intestine. The Caco-2 model is well-established and extensively characterized, having been used for decades to predict the transport of orally administered pharmaceuticals and to assess the membrane permeability of drugs and xenobiotics (Artursson 1990 ; Hidalgo et al. 1989 ; Artursson and Borchardt 1997 ). However, the Caco-2 model has notable limitations regarding the absorption of compounds that rely on paracellular transport routes and the replication of intestinal drug metabolism (Artursson 1990 ; Artursson and Karlsson 1991 ; Sun et al. 2008 ). In this study, an advanced Caco-2 model was developed and characterized to better reflect physiological conditions of the human intestine. In addition to the differentiated epithelial cell layer (Caco-2), the model incorporates additional barriers, such as a mucosal gel layer established by co-culturing Caco-2 cells with mucus-producing goblet cells (HT29-MTX) and a lamina propria component represented by collagen I. Mucin, produced by goblet cells, can bind to drugs and reduce their absorption (Artursson 1990 ). While subepithelial barriers such as the lamina propria and capillary walls are generally not considered rate-limiting for the absorption of small molecules (Artursson 1990 ), incorporating extracellular matrices has been shown to improve cell attachment, barrier integrity and cellular differentiation (Linz et al. 2020 ; Swiderek et al. 2013 ). Studies integrating HT29-MTX co-cultures have demonstrated modified permeability barriers by altering paracellular resistance and secretory transport, yielding improved physiological relevance and enhanced robustness with respect to the reproducibility of the results (Hilgendorf et al. 2000 ). These advanced in vitro ADME models, which address both absorption and metabolism in toxicokinetic assessment, represent physiologically human relevant systems and align with the ongoing shift towards integrating and adopting new approach methodologies into next generation risk assessment frameworks (Escher et al. 2022 ). Furthermore, the herein investigated cell model shows strong potential for adaptation to more complex and microfluidic in vitro systems, such as organ-on-a-chip platforms. These complex systems will be the focus of future research aimed at further elucidating intestinal absorption and metabolism of additional pesticides, thereby supporting EFSA’s proposal to prioritize ‘advanced cell culture models including organ on a chip’ as one of seven key scientific areas requiring further development for the implementation of NAMs in EFSA’s scientific assessment processes (Escher et al. 2022 ). In addition, EFSA’s progressive move towards more pragmatic, weight-of-evidence-based regulatory decision-making offers promise for supporting the 3Rs principle (reduction, refinement, and replacement of animal testing) through the application of the in vitro models presented here (Escher et al. 2022 ). Fig. 1. Open in a new tab a Schematic illustration of the pancreatin test method, with 14 C-trifloxystrobin as the test item and resorufin-labeled casein as the control substrate. b Design of the static compartmentalized intestinal gut-model composed of Caco-2/HT29-MTX monolayers co-cultured on collagen I-coated transwell inserts placed within standard multiwell plates. c Metabolism of [trifluoromethylphenyl-UL- 14 C] trifloxystrobin to [trifluoromethylphenyl-UL- 14 C] carboxylic acid trifloxystrobin (CGA321113, main metabolite). TFS = trifloxystrobin, */UL = uniformly labeled [ 14 C] Materials and methods Pancreatin assay Enzymatic activity control and digestive fate analysis of 14 C-trifloxystrobin The pancreatin assay validation involved ensuring the proteolytic activity of a pancreatin solution used as simulated intestinal fluid (SIF). Initially, 0.5% (w/v) pancreatin [Pancreatin from porcine pancreas (≥  3 USP specifications), Sigma-Aldrich, St. Louis, Missouri] was freshly prepared by dissolving pancreatin in 50 mM monobasic potassium phosphate [pH 7.5] through gentle mixing for 15 min, followed by chilling the solution on ice. For enzymatic activity control, SIF was further diluted to 0.025% (w/v) and used to digest a 0.4% (w/v) resorufin-labeled casein solution [Universal protease substrate (Casein, resorufin-labeled), Roche Holding AG, Basel, Switzerland]. The control experiment was conducted by incubating the diluted SIF with resorufin-labeled casein in an incubation buffer containing 200 mM Tris and 20 mM CaCl 2 [pH 7.8]. To investigate the digestive fate of 14 C-trifloxystrobin, the 0.5% (w/v) SIF solution was used for incubation with the radiolabeled fungicide. The assay was performed by incubating SIF with 14 C-trifloxystrobin, which was dissolved in deionized water, along with an incubation buffer containing 20 mM CaCl 2 [pH 7.8]. The final concentration of 14 C-trifloxystrobin in the reaction mixture was adjusted to 10 µM to comply with the estimated exposure of the fungicide by human ingestion through food and feed crops. Additionally, the chosen dose was aligned to experimental bioavailability data from animal in vivo studies showing maximum blood concentrations (C max ) of 10 µM. Moreover, the concentration is suitable for analytical measurements (LC–MS/MS). All samples were incubated and processed in Protein LoBind™ tubes [Eppendorf, Hamburg, Germany]. The incubations were carried out under kinetic conditions (15 and 30 min, 2, 4, 24 and 48 h) at 37 °C ± 2 °C, with blank reactions using potassium phosphate buffer instead of pancreatin to account for non-enzymatic degradation of casein and buffer-mediated degradation of 14 C-trifloxystrobin (stability control), respectively. Reactions were stopped by adding 5% (w/v) trichloroacetic acid yielding a 2:1 ratio for control samples and 100% acetonitrile [Acetonitrile, ≥  99.9%, Sigma-Aldrich, St. Louis, Missouri] (ratio 3:1), followed by a 10-min incubation step at 37 °C ± 2 °C. Precipitates were removed by centrifugation at maximum speed for 5 min. Supernatants of enzymatic control samples were transferred to an assay buffer (500 mM Tris [pH 8.8], ratio 2:3). Absorption values were captured spectrophotometrically at 574 nm [Cary UV–Vis Compact Photometer, Agilent Technologies, Inc., Santa Clara, California]. Digestive fate supernatants were analyzed using liquid chromatography and high-resolution quantitative mass spectrometry (LC–MS). All experiments were conducted in biological triplicates ( n = 3 ). Independent experiments showing similar results were not explicitly reported in detail. The proteolytic activity of SIF (0.5%) was calculated using specific equations. A reference range derived from an internal standard operating procedure (Bayer AG, Monheim, Germany) was employed to verify the proteolytic activity of the simulated intestinal fluid (SIF) to meet predefined acceptance criteria. U = units, ∆A 574 = mean absorption value normalized to mean blank, min = minutes of incubation, 1 activity unit = 0.001 increase in A 574 per min at 37 °C Proteolytic activity = enzymatic activity of 0.5% SIF, mL = volume of 0.5% SIF used in the assay Gut-model assay Collagen I coating Microporous polycarbonate membranes [0.4 µm pore size, Merck Millipore, Darmstadt, Germany] were coated with 10 µg/cm 2 collagen I [Collagen Type I, rat tail, Corning Inc., Corning, New York] dissolved in 17.5 mM acetic acid for 1 h at room temperature. Following incubation, the membranes were rinsed twice with prewarmed PBS to remove residual acetic acid. Prior to cell seeding, the transwells were preincubated with Caco-2/HT29-MTX medium for 15 min at 37 °C in a humidified atmosphere containing 5% CO 2 . Cultivation, seeding, model validation Caco-2 and HT29-MTX cells were precultured separately in T75 and T25 flasks, respectively, for 5 – 7 days until reaching 80% confluency. Caco-2 cells [CACO-2, ACC 169, human colon adenocarcinoma cell line, DSMZ German Collection of Microorganisms and Cell Cultures GmbH, Leibniz Institute, Braunschweig, Germany] were precultured in Minimum Essential Medium [MEM, Life technologies, Carlsbad, California] supplemented with 20% heat inactivated fetal bovine serum [FBS h.i., Sigma-Aldrich, St. Louis, Missouri], 1 × non-essential amino acids [MEM NEAA (100x), Life technologies, Carlsbad, California], 450 mg dL −1 glucose [45% Glucose solution, Corning Inc., Corning, New York] and 1 × penicillin/streptomycin [PenStrep, Thermo Fisher Scientific, Waltham, Massachusetts]. HT29-MTX cells [HT29-MTX-E12, 12040401, human methotrexate-resistant colorectal adenocarcinoma cell line, Merck KgaA, Darmstadt, Germany] were precultured in Dulbecco’s modified Eagle Medium [DMEM, Life technologies, Carlsbad, California] supplemented with 20% heat inactivated FBS, 1 × non-essential amino acids, 450 mg dL −1 glucose, 1 × penicillin/streptomycin and 2 mM L-glutamine [Corning® Glutagro™ Supplement, Corning Inc., Corning, New York]. At  ~ 80% confluency, cells were washed with PBS [PBS, Sigma-Aldrich, St. Louis, Missouri] and detached using 0.05% trypsin in PBS containing 0.02% EDTA [Trypsin-EDTA-solution, Sigma-Aldrich, St. Louis, Missouri]. Trypsin was neutralized by adding an eight-fold excess of complete medium. Cells were then centrifuged at 200× g for 5 min at 37 °C, resuspended in a 1:1 mixture of Caco-2/HT29-MTX medium, and counted using trypan blue [Trypan-blue, Sigma-Aldrich, St. Louis, Missouri] exclusion and a hemocytometer [C-Chip Neubauer Improved, Incyto, Hongcheondanggok-gil, Korea]. Cells were seeded onto collagen I-coated standing cell culture inserts [0.4 µm pore size, Merck Millipore, Darmstadt, Germany], at a Caco-2/HT29-MTX ratio of 9:1. Co-cultures were established in mixed medium and maintained for 22 days to allow the formation of confluent monolayers. Medium was partially replaced (50%) every 2 – 3 days, followed by measurement of transepithelial electrical resistance (TEER) using an epithelial volt/ohm meter [Evom3, WPI, Sarasota, Florida] to monitor monolayer integrity. Additionally, single cell monolayers cultivated for 22 days were exposed to lucifer yellow to assess the dye’s paracellular flux and thereby evaluate monolayer integrity. The lucifer yellow assay was applied to barrier models with different TEER values, and a correlation between the lucifer yellow paracellular flux and the TEER values was established, indicative of integrated, confluent and intact monolayers. The threshold of 1% paracellular flux corresponded to a TEER value of 500 Ω cm 2 (Online Resource 8). Cell-free collagen I-coated inserts were used to account for background resistance. Only monolayers with TEER values above 500 Ω cm 2 were used in experiments. Replaced apical medium was collected and stored for subsequent biomarker analysis (glucose, alkaline phosphatase [ALP], and lactate dehydrogenase [LDH]) to confirm endogenous metabolic activity, functional differentiation and exclude cytotoxicity. Monolayers from day 21 were processed for immunofluorescence and Alcian blue staining to visualize cellular structures and mucus production. Gut-exposure Confluent monolayers, confirmed by TEER measurements, were treated with 10 µM atenolol [Atenolol, 98%, Fisher Scientific, Waltham, Massachusetts] and diclofenac [Diclofenac, 98%, Combi-Blocks, San Diego, CA] on day 21 to control and evaluate monolayer permeability. Additionally, cells were exposed to 20 µM testosterone [Testosterone C-IIIN,  ≥  99.0%, Sigma-Aldrich, St. Louis, Missouri] to assess endogenous metabolic activity. The permeability and biotransformation of the fungicide 14 C-trifloxystrobin [[trifluoromethylphenyl-UL- 14 C] Trifloxystrobin,  >  98%, Bayer AG Crop Science, Wuppertal, Germany] were investigated and evaluated by applying 10 µM of the compound onto the cell layers. Dose selection was decided upon same justifications as described earlier. All test compounds were prepared in mixed medium and administered as single doses to the apical compartments, considering the unbound concentration in the application solution for further calculations e.g., permeability coefficients (P app ). Atenolol and diclofenac specimens were collected from basolateral compartments after 2 h of incubation. Testosterone specimens were collected from both compartments at 30 min, 2, 4 and 24 h. 14 C-trifloxystrobin specimens were collected from both compartments at 10 and 30 min, 2, 4, and 24 h. Following exposure, cell lysates were collected post-experimentally from monolayers treated with testosterone and 14 C-trifloxystrobin. Therefore, application solutions were removed, cells were washed with PBS and lysed with 80% acetonitrile [Acetonitrile,  ≥  99.9%, Sigma-Aldrich, St. Louis, Missouri]. Lysed cells were sonicated for 5 min. Supernatants were transferred to glass vials after centrifugation for 5 min at maximum speed. All samples were analyzed by high resolution mass spectrometry to quantify parent compounds and corresponding metabolites, and results were subjected to statistical analysis. Independent experiments were not explicitly reported in detail. All experiments were conducted in biological triplicates ( n = 3 ). Gut-model characterization: immunofluorescence staining To visualize the cellular architecture of the co-culture, particularly the expression and organization of tight junctions as confirmation of monolayer integrity, cells were subjected to immunofluorescence staining on culture day 21. For this purpose, excess culture medium was removed from the transwell inserts, and the cells were fixed with 4% formaldehyde in PBS (pH 7.3) for 10 min at room temperature. The cells were rinsed with PBS and then permeabilized using 0.25% Triton X-100 in PBS for 10 min at room temperature. Following an additional PBS wash, a blocking step was performed by incubating the cells with 1% BSA in PBS for 30 min. Fixation, permeabilization and blocking chemicals were obtained from the Image-iT™ fixation/permeabilization kit from Life Technologies [Carlsbad, California]. After removing the blocking solution, the cells were incubated with a mixture of primary antibody and phalloidin (1:1 ratio v/v) to label F-actin and the tight junction protein zonula occludens (ZO)-1. Specifically, 4 U mL −1 phalloidin conjugated to Alexa Fluor 488, derived from Amanita phalloides [Alexa Fluor™ 488 Phalloidin, Life technologies, Carlsbad, California] and 10 µg mL −1 of mouse anti-human ZO-1/TJP1 antibody conjugated to Alexa Fluor 594 [ZO1 monoclonal antibody (ZO1-1A12), Alexa Fluor™ 594, Life technologies, Carlsbad, California] were diluted in 0.1% BSA in DPBS. The cells were incubated with this staining solution for 30 min at room temperature in the dark. After a final PBS wash, the membranes were excised and mounted on glass slides. The cells were covered with VECTASHIELD® antifade mounting medium containing DAPI [Vectashield® Antifade Mounting Medium with DAPI, Biozol, Hamburg, Germany] and sealed with coverslips. Cellular structures were then imaged using immunofluorescence microscopy [Compact fluorescence microscope BZ-X810, Keyence, Osaka, Japan]. Gut-model characterization: chemical Alcian blue staining To characterize and evaluate the cell model, acidic mucins secreted primarily by goblet cells were detected in monolayers cultured for 21 days. For this purpose, the monolayers were fixed with 4% paraformaldehyde for 10 min, washed with PBS, and subsequently stained with a filtered alcian blue solution (pH 2.5; 0.45 µm syringe filter) [Alcian blue 8GX solution, Sigma-Aldrich, St. Louis, Missouri]. Following a 20-min incubation at room temperature, the monolayers were thoroughly rinsed with PBS. The membranes were then excised from the insert holders, mounted onto glass slides with a drop of PBS, and examined by phase-contrast microscopy [Microscope Primovert, Carl Zeiss Suzhou Co., Ltd., Suzhou, China]. Analytics Following sample collection, acetonitrile [Acetonitrile,  ≥  99.9%, Sigma-Aldrich, St. Louis, Missouri] was added to apical and basolateral specimens at a 1:1 (v/v) ratio. Following one freeze–thaw cycle, the samples were transferred to Protein LoBind™ tubes [Eppendorf, Hamburg, Germany]. The precipitates were pelleted by centrifugation at maximum speed (≥ 16,000× g ) for 7 min. The resulting supernatants were then measured by LC–MS. Diclofenac, testosterone and 14 C-trifloxystrobin samples were analyzed using a Q Exactive™ Plus hybrid quadrupole-Orbitrap™ mass spectrometer [Thermo Scientific™, Waltham, Massachusetts], while atenolol samples were analyzed using an Orbitrap Exploris™ 480 mass spectrometer [Thermo Scientific™, Waltham, Massachusetts]. Injection volumes were set to 2 µL (diclofenac, testosterone and 14 C-trifloxystrobin) and 5 µL (atenolol). Samples were separated using a YMC-Triart C 18 column [1.9 µm, 100 × 2.0 mm; YMC Co., Ltd., Kyoto, Japan] with a 0.2 mL min −1 gradient flow, coupled to the ESI (electrospray ionization) ion source of the mass spectrometer. The mobile phase gradient (solvent A: water + 0.1% formic acid, solvent B: acetonitrile + 0.1% formic acid) was programmed as follows: 5% B at 1 min, ramping to 95% B at 12 min, 95% B at 13.5 min, with re-equilibration at 15 min. Electrospray ionization was performed in positive ion mode, with capillary temperatures set to 325 °C and 320 °C, respectively. LC–MS measurements were conducted in full scan mode (75 – 1100 m/z) with a mass resolution of 140,000 and 240,000 at 200 m/z, respectively. Dissociation (MS/MS) experiments were performed using optimized collision energies specific to each compound yielding 5% parent ion [M + H] + intensities. The mass tolerance was set to  < 5 ppm. Spectral data were acquired and analyzed both qualitatively and quantitatively using Xcalibur™ software version 2.2 [Thermo Fisher Scientific, Waltham, Massachusetts]. Retention times, exact mass to charge (m/z) ratios and specific radiolabel patterns as well as dissociation patterns were used to identify applied parent compounds and corresponding metabolites. Integrated data were further processed using excel software [v. 2503, Microsoft Excel, Redmond, Washington]. Statistics Mean values were statistically analyzed using a one-tailed paired t -test two samples for means and significance levels were set at p ≤ 0.05 or p ≤ 0.01 . Statistical analyses were performed using excel software [v. 2503, Microsoft Excel, Redmond, Washington]. Results Pancreatin assay The proteolytic activity of simulated intestinal fluid (SIF) containing 0.5% pancreatin demonstrated a gradual decline over time. This decline can be attributed to the limited availability of control substrates within the reaction mixture (Online Resource 7). Nevertheless, measurable enzymatic activity persisted for up to 48 h of incubation, indicating the sustained functionality of the SIF preparation. This finding is further corroborated by the progressive increase in the concentration of the carboxylic acid metabolite of trifloxystrobin upon digestive fate analysis by SIF. As illustrated in Fig. 2 , the relative concentration (%) of the primary metabolite in enzymatic samples was compared to enzyme-free stability controls, employing a kinetic approach. Between 15 min and 24 h, the concentration of the acid metabolite was approximately double that observed in the stability controls. After 48 h, this concentration increased threefold ( p ≤ 0.01 ). Environmental fate studies have indicated that trifloxystrobin undergoes rapid degradation to its acid metabolite (CGA 321113; Fig. 1 c) in aquatic environments. This degradation occurs through various processes, including metabolism, photolysis, and hydrolysis, with reported half-lives ranging from a few hours to several days (EPA 1999 ). Notably, the free acid metabolite, which is inactive regarding the fungicidal mode of action, degrades more slowly than the parent compound and is characterized by its mobility and persistence (APVMA 2000 ). Under conditions simulating intestinal digestion, trifloxystrobin exhibited partial degradation in a buffered aquatic environment. However, this degradation was significantly enhanced in the presence of enzymatic activity. This confirms that pancreatic enzymes convert trifloxystrobin into its acid metabolite prior to its entry into the small intestine. Fig. 2. Open in a new tab Formation of the primary metabolite 14 C-carboxylic acid trifloxystrobin upon enzymatic digestion of. 14 C-trifloxystrobin by simulated intestinal fluid. Metabolite concentrations were normalized to the parent concentration added at timepoint zero and are presented as percentages (%). The mean concentrations from enzymatic samples were statistically compared to those from enzyme-free stability controls using a significance level of p ≤ 0.01***. Data are presented as mean ± standard deviation ( n = 3 ) Gut-model assay Monolayer integration & model characterization The integrity of the investigated cell monolayers was monitored by measuring transepithelial electrical resistance (TEER) (Fig. 3 a). From day 19 onwards, TEER values consistently exceeded the reference threshold of 500 Ω cm 2 , which was established through paracellular flux analysis using lucifer yellow (Online Resource Table 1, Fig. 8), thereby confirming monolayer integrity. Monolayers employed in the exposure experiments exhibited TEER values ranging from 600 – 950 Ω cm 2 on day 21. Additionally, confluency and monolayer integrity were verified by TEER measurements both four hours post compound application and post-experimental (day 22). Various TEER values for Caco-2 monoculture models have been reported in the literature, with many exceeding 1000 Ω cm 2 , an indicator of intact Caco-2 monolayers suitable for transport studies (Hubatsch et al. 2007 ; Hahn et al. 2022 ; Lechanteur et al. 2017 ). However, these values are higher than the reported physiological in vivo resistances observed in humans, which are approximately 300 – 400 Ω cm 2 for the colon and 50 – 100 Ω cm 2 for the small intestine (Srinivasan et al. 2015 ). Co-culture models comprising Caco-2 and HT29-MTX cells displayed lower resistance values, more closely reflecting the physiological conditions of the intestinal epithelium (Hilgendorf et al. 2000 ; Srinivasan et al. 2015 ), which is consistent with the values obtained in this study. Further characterization of the co-culture model was performed using immunofluorescence and alcian blue staining (Fig. 3 b and c). The dense and compressed cell layers observed are typical of differentiated, confluent Caco-2 monolayers (Artursson 1990 ) and morphological visualization confirmed the expression of tight junctions, further indicating intact monolayer integrity (Fig. 3 b). Moreover, the presence of acidic mucus was verified by alcian blue staining (Fig. 3 c). Mucus coverage was distributed across the entire cell layer, consistent with previous observations by Hilgendorf et al. ( 2000 ). In this study, mucus secretion was confirmed for cell layers cultured on semi-permeable transwell membrane supports. Fig. 3. Open in a new tab a Transepithelial electrical resistance (TEER) values of Caco-2/HT29-MTX co-cultures. Monolayer integrity was further confirmed via lucifer yellow paracellular flux analysis to establish the depicted reference value (500 Ω cm 2 ). TEER values were recorded prior to compound application (day 21), 4 h post-application (day 21), and post-experimental (day 22). Negative TEER values resulting from background resistance normalization were set to zero (day 2). Data are presented as mean ± standard deviation ( n = 8 ). b Overlay image of immunofluorescence-stained Caco-2/HT29-MTX co-cultures. Monolayer formation and expression of tight junctions were morphologically confirmed on culture day 21 by immunofluorescence labeling of nuclei (DAPI, blue), F-actin filaments (GFP, green) and ZO-1 as the tight junction marker (TxRed, red). c Phase-contrast image of Alcian blue-stained Caco-2/HT29-MTX co-cultures, highlighting mucus secretion. The depicted monolayer exhibited a transepithelial electrical resistance (TEER) value of 728 Ω cm 2 , indicative of barrier integrity. 20 × objectives were used to visualize cellular structural components and the secreted mucus layer (dark blue regions) of cells cultured on transwell membrane supports Endogenous gut-metabolism of testosterone To evaluate the endogenous metabolic activity in the in vitro co-cultured gut-model, testosterone was applied to the apical compartment. Previous ex vivo studies using homogenates extracted from adult male duodenal mucosa, have reported the formation of androstenedione, while no phase II metabolites were detected (Nienstedt et al. 1980 ). Similarly, in human ileum and colon precision-cut intestinal slices, androstenedione, 6 β - and 2 β -hydroxy testosterone were identified in the ileum, whereas 15 α -hydroxy testosterone was observed in the colon. Again, no phase II metabolites were detected (Iswandana et al. 2018 ). Additionally, 17 β -hydroxysteroid-dehydrogenase type 2 has been shown to be expressed throughout the human gastrointestinal tract, with immunoreactivity localized almost exclusively in the absorptive epithelium, emphasizing its central role in intestinal sex steroid metabolism (Sano et al. 2001 ). In the present experiment, using Caco-2/HT29-MTX co-cultures, testosterone levels in the apical compartment declined rapidly over time, while basolateral concentrations increased over the first 4 h of incubation, followed by a decline at 24 h (Fig. 4 a and b). In contrast to cell-free controls, mass balance for testosterone could not be achieved, indicating cell-dependent metabolic conversion. Androstenedione was detected in both compartments and increased progressively over time (Fig. 4 c), and with the absence of this metabolite in cell-free controls, the functional expression and enzymatic activity of 17 β -hydroxysteroid-dehydrogenase was confirmed in the co-culture model. No phase II testosterone metabolites or other phase I metabolites were detected in all specimens, which agrees with previous ex vivo observations. Statistical analysis of androstenedione concentrations revealed a significant time-dependent increase in both compartments up to 4 h of incubation. At 24 h, no further increase was observed, assuming redistribution into the basolateral compartment. These findings demonstrate substantial endogenous first-pass intestinal metabolism of testosterone by the in vitro gut-model. In humans, steroids are primarily absorbed via the intestinal tract. However, their oral bioavailability is limited by extensive intestinal and hepatic first-pass metabolism (Schedl 1965 ; Frey et al. 1979 ). The results presented here recapitulate the intestinal absorption and phase I endogenous first-pass metabolism of testosterone, supporting the relevance and suitability of this in vitro model for investigating human intestinal steroid metabolism. Fig. 4. Open in a new tab a Testosterone distribution in the apical compartment of Caco-2/HT29-MTX cell barriers and cell-free blanks. b Testosterone distribution in the basolateral compartment of Caco-2/HT29-MTX cell barriers and cell-free blanks. c Androstenedione distribution in apical and basolateral compartments of Caco-2/HT29-MTX cell barriers and cell-free blanks. Average androstenedione concentrations per compartment were statistically compared between conditions using significance levels of p ≤ 0.05* or p ≤ 0.01*** . All data are reported as mean ± standard deviation ( n = 3 ). n.d.  =  not detectable In vitro gut-permeability towards references and 14 C-trifloxystrobin To evaluate and control the in vitro gut-barrier permeability of the model, atenolol and diclofenac were employed as negative and positive reference compounds, respectively, and their apical-to-basolateral permission was assessed. In addition, the permeability of the in vitro cell layers to 14 C-trifloxystrobin was investigated. Figure 5 presents the apparent permeability coefficients (P app (*10 –6 cm s −1 )), calculated based on basolateral concentrations after 2 h of incubation for the tested compounds, along with cell-free blank controls. The fraction absorbed (F a , %) in humans is commonly categorized into three groups: F a < 50% (low), F a = 50 – 84% (intermediate), F a ≥ 85% (high) (FDA 2017 ). A drug is classified as highly permeable if the absolute bioavailability is  ≥ 85% or if  ≥ 85% of administered dose is recovered in urine as parent drug or as the sum of the parent drug and its phase I and II metabolites (Volpe 2022 ). Reported human F a values are approximately 50% for atenolol (Varma et al. 2010 ) and 82 – 100% for diclofenac (Velicky et al. 2011 ; Zhu et al. 2002 ). These compounds have consistently demonstrated distinct P app values in the Caco-2 monoculture model. Historically, a sigmoidal correlation between drug absorption rates in the Caco-2 model and human intestinal absorption has been described (Artursson and Karlsson 1991 ). In the literature, atenolol is widely used as a low-permeability control with typical P app values around or below 1*10 –6 cm s −1 (Zhu et al. 2002 ; Yang et al. 2007 ; Yamashita et al. 2000 ), while diclofenac demonstrated values  ≥ 10*10 –6 cm s −1 and serves as a high-permeability control in this study (Fig. 5 ). For Caco-2 monolayers, compounds with complete absorption exhibit high P app values (> 1*10 –6 cm s −1 ), whereas incompletely absorbed compounds tend to have lower P app values (< 1*10 –6 cm s −1 ) (Artursson and Karlsson 1991 ; Artursson et al. 2001 ). In another study, threshold P app values have been used to differentiate between high- and low-permeable drugs, with P app > 14*10 –6 cm s −1 indicative of high permeability and  < 5*10 –6 cm s −1 indicative of low permeability (Volpe et al. 2007 ). However, the inclusion of the HT29-MTX cell line can reduce monolayer resistance and thus increase permeability, as demonstrated in Fig. 3 a. Several studies have shown that the permeability coefficients of various compounds, including atenolol, increase with higher proportions of HT29-MTX co-cultured with Caco-2 cells (Hilgendorf et al. 2000 ; Lozoya-Agullo et al. 2017 ). Additional adaptations, such as the use of human plasma in transport assays, increased and enhanced P app values of lipophilic compounds, better reflecting physiological conditions than conventional HBSS transport buffers (Katneni et al. 2018 ). Therefore, the permeability coefficients obtained from conventional monoculture models may not be directly applicable to this adapted co-culture gut-model. Consequently, individual reference P app values must be established for this system. Figure 5 shows the measured P app values of atenolol (0.92 ± 0.41 *10 –6 cm s −1 ) and diclofenac (17.15 ± 1.98 *10 –6 cm s −1 ), confirming their suitability as low and high permeable controls, respectively, for this model. However, to define robust reference ranges, additional compounds with well-characterized human F a values should be evaluated in this advanced in vitro gut-model. The permeability of 14 C-trifloxystrobin yielded an average P app value of 2.21 ± 0.28 *10 –6 cm s −1 . Although this value must be interpreted in the context of the adapted model’s increased permeability (due to serum and HT29-MTX co-culture), it aligns more closely with the low-permeability reference atenolol. Based on this result, 14 C-trifloxystrobin can tentatively be categorized as low-permeability compound (F a < 50%). However, it is important to note that this conclusion does not account for intestinal metabolism, which will be addressed in the following section on xenobiotic in vitro gut-metabolism . Fig. 5. Open in a new tab Apical-to-basolateral permeability (P app (*10 –6 cm s −1 )) of Caco-2/HT29-MTX co-cultures towards reference compounds (atenolol and diclofenac) and 14 C-trifloxystrobin. Data are presented as mean ± standard deviation ( n = 3 ) Xenobiotic in vitro gut-metabolism of 14 C-trifloxystrobin To assess the in vitro intestinal metabolism of trifloxystrobin, the fungicide was applied to the apical compartment and investigated in apical and basolateral compartments as well as in cell lysates. In apical specimens (Fig. 6 a), the concentration of 14 C-trifloxystrobin decreased markedly over time (from 10 min to 24 h). After 24 h of incubation, only trace amounts (< 1% of the initial dose) were detectable, indicating substantial absorption and/or metabolic transformation. In the basolateral compartment, however, 14 C-trifloxystrobin was detected only in low concentrations (< 1% at 10 and 30 min and at 24 h;  < 2% at 2 and 4 h of the initial dose) at all measured timepoints (Fig. 6 b). This supports the observation of limited permeability of trifloxystrobin across the in vitro intestinal barrier, as discussed previously. Notably, trifloxystrobin was not detectable in the cell lysates (data not shown), assuming its minimal passage through the barrier occurs predominantly via paracellular routes rather than (active) transcellular or transcytotic pathways. Importantly, the primary metabolite, 14 C-carboxylic acid-trifloxystrobin (CGA321113) was detected in substantial amounts, cell-dependent in the apical compartment (Fig. 6 a) after 2, 4 and 24 h (> 20% of the initial dose at 2 h,  > 30% at 4 and 24 h). At 24 h, a significant decline in the apical metabolite concentration compared to 4 h concludes subsequent passage through the cellular barrier. Correspondingly, the metabolite was detected at increasing concentrations in the basolateral compartment, reaching nearly 20% of the initially applied dose by 24 h (Fig. 6 b). Given the low trifloxystrobin levels in the basolateral samples, the observed metabolite abundance is likely attributable to transport across the in vitro intestinal barrier. Moreover, transcellular transport cannot be excluded, as the carboxylic acid metabolite was qualitatively detected intracellularly (data not shown). Figure 6 c illustrates the proposed transport and metabolic pathways of trifloxystrobin and its carboxylic acid metabolite across the static Caco-2/HT29-MTX co-culture barrier-model. The primary pathway involves enzymatic conversion of trifloxystrobin to its carboxylic acid metabolite within the apical compartment, followed by transport across the cellular barrier. Minor pathways include medium-mediated hydrolysis of trifloxystrobin to the main metabolite in both apical and basolateral compartments (Online Resource 10c and 10d) and the generally low direct absorption of the parent compound across the cellular layer. To investigate the enterohepatic fate of trifloxystrobin, an in vitro study comparing metabolism in human and rat liver microsomes was submitted to the European Commission (EFSA 2017b ). After 1 h, incubations of trifloxystrobin with rat and human liver microsomes yielded similar metabolic profiles. Trifloxystrobin was essentially completely metabolized by rat microsomes, yielding four metabolites, each of which accounted for more than 5% of the applied dose. In human liver microsomes, trifloxystrobin was likewise completely metabolized within 1 h, yielding two predominant metabolites that accounted for 24.4% and 73.3% of the applied dose, respectively. In the comparative profiling, those two major human metabolites were also detected in rat microsomal incubations but at lower relative abundances (14.2% and 41.5%), indicating conserved metabolic pathways with quantitative differences in metabolite distribution between species. Rat microsomes produced a larger number of distinct metabolites, reflecting greater qualitative diversity of hepatic biotransformation compared to human microsomes. All metabolites formed by human microsomes were also formed by rat microsomes, and no human-unique metabolites were observed. Although rat microsomes generated more metabolite species overall, the two principal human metabolites were formed at higher relative abundance in human microsomes, indicating quantitatively greater metabolism along those specific pathways in humans (EFSA 2017b ). Combined with in vivo rat metabolic data, the intestinal carboxylic-acid metabolite CGA321113 appears to function as a major precursor for subsequent extensive hepatic (phase II) biotransformation in humans, thereby diminishing systemic exposure to the parent compound and reducing overall oral bioavailability. Fig. 6. Open in a new tab a Apical concentrations (µM) of 14 C-trifloxystrobin and its carboxylic acid metabolite in static Caco-2/HT29-MTX intestinal barrier models. b Basolateral concentrations (µM) of 14 C-trifloxystrobin and its carboxylic acid metabolite in static Caco-2/HT29-MTX intestinal barrier models. Metabolite concentrations were corrected for background levels observed in blank controls (data not shown) to account for medium-dependent hydrolysis of trifloxystrobin. All data are presented as mean ± standard deviation ( n = 3 ). Average 14 C-carboxylic acid trifloxystrobin concentrations were statistically compared between experimental conditions using a significance level of p ≤ 0.05* or p ≤ 0.01***. c Schematic illustration of the primary and secondary transport and metabolic pathways of trifloxystrobin and its carboxylic acid metabolite across static Caco-2/HT29-MTX barrier models. Major transport routes are indicated by dark blue arrows, while minor routes are represented by the light blue arrow. Orange arrows indicate medium mediated hydrolysis. The metabolite was also qualitatively detected in the intracellular space. TFS = trifloxystrobin, M = carboxylic acid metabolite Together, these results provide an initial indication of the potential intestinal absorption and metabolic fate of trifloxystrobin in humans. Although further data such as refined permeability coefficient reference values are needed to fully interpret these findings, this study presents human-derived intestinal in vitro data of trifloxystrobin for the first time, supporting human next generation risk assessment. Discussion The Caco-2 model is well-established and extensively characterized, having been used for decades to predict the transport of orally administered pharmaceuticals and to assess the membrane permeability of drugs and xenobiotics (Artursson 1990 ; Hidalgo et al. 1989 ; Artursson and Borchardt 1997 ). Numerous studies have demonstrated good correlations between human oral drug absorption or fraction absorbed (F a ) and apparent permeability coefficients determined using Caco-2 cells for compounds with diverse structural properties, such as varying lipophilicity, molecular weight, charge, and human absorption profiles (Artursson 1990 ; Artursson and Karlsson 1991 ; Kamiya et al. 2020 ). These correlations have often proven to be more reliable than the sole use of octanol/water partition coefficients alone for predicting passive absorption. However, the Caco-2 monoculture model has notable limitations. It tends to underestimate the absorption of compounds that rely on paracellular transport due to its tight junctions, which more closely resemble those found in the colon rather than the small intestine. Additionally, its capacity to replicate intestinal drug metabolism is limited (Artursson 1990 ; Artursson and Karlsson 1991 ; Sun et al. 2008 ). Several strains of heterogenous Caco-2 cell lines have been described resulting in variations in gene expression and transport activities, which are further influenced by interlaboratory differences such as culture conditions (Hayeshi et al. 2008 ). Another drawback lies in the diminished presence of key cytochrome P450 (CYP) isoforms, specifically CYP3A4 and CYP2C members, which are dominant in the epithelial lining of the human intestine (Lampen et al. 1998 ; Prueksaritanont et al. 1996 ). Despite these drawbacks, many studies have demonstrated that the Caco-2 model remains a reliable and predictive tool for studying drug transport across the intestinal epithelium. It is widely used to identify compounds with potential absorption issues and to select candidates with favorable passive absorption properties (Artursson et al. 2001 ). Moreover, Caco-2 cells exhibit increased expression of brush border hydrolytic enzymes (Küblbeck et al. 2016 ) such as carboxylesterase 1 (CES1) (Yamada et al. 2023 ). Human intestinal metabolism can account for more than 20% of the loss of drug bioavailability, highlighting its importance in bioavailability predictions (Varma 2010 ). Despite its limitations, the Caco-2 model remains a cornerstone of intestinal absorption research due to its predictive accuracy, reproducibility, and broad regulatory acceptance (Panse and Gerk 2022 ). In this study, we successfully established an advanced Caco-2/HT29-MTX co-culture model with pre-intestinal metabolism facilitated by pancreatic enzymes to overcome limitations observed in the Caco-2 monoculture system. Integrating the HT29-MTX cell line in our model led to less pronounced tight junctions, as indicated by lower TEER values, aligning the model more closely with human small intestine physiology. Furthermore, we verified the abundance of mucin, which supports the model’s physiological relevance and is capable in altering binding capacities of xenobiotics and reducing intestinal absorption. We focused on investigating the intestinal permeability and the xenobiotic metabolism of the fungicide trifloxystrobin. Incubations with pancreatin revealed a significant portion of the metabolized active ingredient even before it reached the intestine. Further, the hormone testosterone served as endogenous metabolic control to verify the metabolic competence of the used cell lines. This led to the discovery of carboxylic acid trifloxystrobin and androstenedione, indicating partial metabolism in the presented intestinal models. However, the absence of hydroxylation e.g., hydroxy-testosterone and phase II conjugates highlighted reduced expression of metabolic enzymes, such as key CYP450 isoforms, in Caco-2 cell lines. Addressing these limitations, genetically modified Caco-2 cell lines expressing human metabolic enzymes could enhance model accuracy, as previously discussed by Küblbeck et al. ( 2016 ). To improve physiological relevance further, integrating endothelial cells to simulate subepithelial capillary structures, increasing HT29-MTX cell density in co-culture, or incorporating the intestinal microbiota could enhance the model’s capabilities. The human gut microbiota has a larger metabolic repertoire compared to human cells. However, it often opposes or even reverses host metabolism as a self-preservation mechanism, rather than facilitating excretion of xenobiotics from the human body. Nevertheless, microbiota can actively modify the toxicokinetic and toxicodynamic of xenobiotics (Koppel et al. 2017 ). Despite its profound impact, this study excluded microbiota due to its reliance on the unique functional composition of microorganisms and its intricate integration into sterile cell cultures. To our knowledge, this study reports investigations of trifloxystrobin using intestinal in vitro models to explore human absorption and metabolism for the first time. We consider our data to demonstrate the feasibility of these assays and models in pesticide research, particularly in predicting human oral bioavailability and systemic exposure, thereby supporting human NGRA and the 3R principle. However, regarding intestinal permeability (P app ) our cell-model is not recommended as a standalone approach, emphasizing the critical role of metabolism in such contexts. Additionally, our experiments did not account for metabolites generated in the liver and subsequently transported to the intestine via enterohepatic circulation. Therefore, transferring these models to microphysiological systems (e.g., organ-on-a-chip) and combining in vitro organoids in one approach will be the subject of our future research. However, the presented static models provide comprehensive insights into human pesticide absorption, metabolism, and preliminary assumptions regarding transport mechanisms (transcellular vs. paracellular). Furthermore, broader investigations should encompass active ingredients from all major pesticide types (herbicides, fungicides, insecticides). Additionally, the pesticide’s mode of action may alter metabolic capacity in the cells. For instance, the mitochondrial activity inhibition of trifloxystrobin in fungi and mammalian cells could potentially impact endogenous and xenobiotic metabolism, although this specific toxicological effect was not addressed in our current study. Notably, trifloxystrobin is characterized by poor absorption from the gastrointestinal tract resulting in significant excretion of unchanged compound in feces, while the fraction absorbed undergoes extensive metabolism in rats, yielding approximately 35 metabolites limiting the systemic exposure to the parent compound (Hollingworth 2001 ). Studies using human and rat liver microsomes, together with analyses of rat bile from in vivo experiments, show that the carboxylic acid derivative of trifloxystrobin is the primary metabolite formed prior to hepatic phase II conjugation. These results, together with the data presented here, provide key insights into the intestinal and hepatic metabolism and the potential bioavailability of trifloxystrobin in humans (EFSA 2017a , EFSA b ). In regulatory in vivo toxicokinetic studies conducted in accordance with OECD Test Guideline 417, only total sample radioactivity (the sum of parent compound and its metabolites) is quantified. For rapidly metabolized pesticide active ingredients such as trifloxystrobin, this approach provides limited information on systemic exposure. This limitation is particularly relevant because metabolites may have differing toxicological profiles, underscoring the need for further investigation to inform risk assessment. For example, CGA321113 has been reported to exhibit lower toxicity than trifloxystrobin (EFSA 2025 ). Although current studies provide valuable toxicokinetic information, the complexity of trifloxystrobin’s metabolic pathways and the potential for variable toxicological effects among its numerous metabolites highlight the need for continued research. A deeper understanding of individual metabolites, particularly their systemic exposure and biological interactions in humans, is crucial for accurate risk assessment and protection of human health. Future investigations should therefore prioritize characterization and quantification of metabolites and evaluation of their toxicological properties, ensuring that regulatory decisions are based on comprehensive and up-to-date scientific evidence. Supplementary Information Below is the link to the electronic supplementary material. Supplementary Material 1 (2.5MB, docx) Acknowledgements The authors would like to thank Michael Kubicki (Bayer AG Division Crop Science, Monheim, Germany) for his expertise and support regarding LC-MS/MS measurements. Author contributions [Eileen Hallscheidt] and [Marc Lamshoeft] contributed to the study conception and design. Material preparation, data collection and analysis was performed by [Eileen Hallscheidt]. The first draft of the manuscript was written by [Eileen Hallscheidt] and all authors commented on previous versions of the manuscript. All authors read and approved the final manuscript. Funding This work was financially supported by Bayer AG, Crop Science Division, Monheim, Germany. Data availability All data generated or analyzed during this study are included in this published article (and its supplementary information files). Declarations Conflict of interest The authors declare they have no competing financial interest. Employment Kerstin Hartmann is not employed at Bayer AG anymore, but was at the time of fi rst submission of the manuscript. Integrity of research and reporting, ethical standards The manuscript does not contain clinical studies or patient data. Footnotes Publisher’s Note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. References APVMA, National Registration Authority for Agricultural and Veterinary Chemicals (2000) Evaluation of the new active Trifloxystrobin in the product Flint Fungicide. https://www.apvma.gov.au/sites/default/files/publication/14081-prs-trifloxystrobin.pdf . Accessed 21 Jul 2025 Artursson P (1990) Epithelial transport of drugs in cell culture. I: a model for studying the passive diffusion of drugs over intestinal absorptive (Caco-2) cells. J Pharm Sci. 10.1002/jps.2600790604 [ DOI ] [ PubMed ] [ Google Scholar ] Artursson P, Borchardt R (1997) Intestinal drug absorption and metabolism in cell cultures: Caco-2 and beyond. Pharm Res 14:1655–1658. 10.1023/a:1012155124489 [ DOI ] [ PubMed ] [ Google Scholar ] Artursson P, Karlsson J (1991) Correlation between oral drug absorption in humans and apparent drug permeability coefficients in human intestinal epithelial (Caco-2) cells. Biochem Biophys Res Commun 3:880–885. 10.1016/0006-291x(91)91647-u [ DOI ] [ PubMed ] [ Google Scholar ] Artursson P et al (2001) Caco-2 monolayers in experimental and theoretical predictions of drug transport. Adv Drug Deliv Rev 46:27–43. 10.1016/s0169-409x(00)00128-9 [ DOI ] [ PubMed ] [ Google Scholar ] EFSA, European Food Safety Authority (2017) Peer review of the pesticide risk assessment of the active substance trifloxystrobin. EFSA J 15(10):8. 10.2903/j.efsa.2017.4989 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] EFSA, European Food Safety Authority (2017b) Draft (renewal) assessment report for trifloxystrobin prepared according to the commission regulation (EU) N° 1107/2009. 3-B.6 EFSA, European Food Safety Authority (2025) Reasoned OPINION, MODIFICATION of the existing maximum residue levels and setting of import tolerances for trifloxystrobin in various crops. EFSA J 23(4):e9387. 10.2903/j.efsa.2025.9387 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] EPA, United States Environmental Protection Agency (1999) Pesticide fact sheet trifloxystrobin. US EPA-Pesticides-Fact Sheet for Trifloxystrobin. Accessed 24 Jul 2025 Escher S et al (2022) Development of a roadmap for action on new approach methodologies in risk assessment. External Scientific report EFSA supporting publication. EN-7341. 10.2903/sp.efsa.2022.EN-7341 FDA, Food and drug administration (2017) Waiver of in vivo bioavailability and bioequivalence studies for immediate-release solid oral dosage forms based on a biopharmaceutics classification system. Guidance for industry, U.S. Department of Health and Human Services, Food and drug administration, Center for drug evaluation and research. Waiver of In Vivo Bioavailability and Bioequivalence Studies for Immediate-Release Solid Oral Dosage Forms Based on a Biopharmaceutics Classification System Guidance for Industry Accessed 28 Jul 2025 Frey H et al (1979) Bioavailability of oral testosterone in males. Eur J Clin Pharmacol 16(5):345–349. 10.1007/bf00605634 [ DOI ] [ PubMed ] [ Google Scholar ] Hahn M et al (2022) Permeability of dopamine D2 receptor agonist hordenine across the intestinal and blood-brain barrier in vitro. PLoS ONE 17(6):e0269486. 10.1371/journal.pone.0269486 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Hayeshi R et al (2008) Comparison of drug transporter gene expression and functionality in Caco-2 cells from 10 different laboratories. Eur J Pharm Sci 35:383–396. 10.1016/j.ejps.2008.08.004 [ DOI ] [ PubMed ] [ Google Scholar ] Hidalgo I et al (1989) Characterization of the human colon carcinoma cell line (Caco-2) as a model system for intestinal epithelial permeability. Gastroenterology 96:736–749 [ PubMed ] [ Google Scholar ] Hilgendorf C et al (2000) Caco-2 versus Caco-2/HT29-MTX Co-cultured cell lines: permeabilities via diffusion, inside-and outside-directed carrier-mediated transport. J Pharm Sci 89:63–75. 10.1002/(SICI)1520-6017(200001)89:1<63::AID-JPS7>3.0.CO;2-6 [ DOI ] [ PubMed ] [ Google Scholar ] Hollingworth RM (2001) Inhibitors and uncouplers of mitochondrial oxidative phosphorylation. In: Krieger RI, Krieger WC (ed). 2 edn. Handbook of pesticide Toxicology, Elsevier, Amsterdam, pp 1169–1261 Hubatsch I, Ragnarsson EGE, Artursson P (2007) Determination of drug permeability and prediction of drug absorption in Caco-2 monolayers. Nat Protoc 2:2111–2119. 10.1038/nprot.2007.303 [ DOI ] [ PubMed ] [ Google Scholar ] Iswandana R et al (2018) Regional differences in human intestinal drug metabolism. Drug Metab Dispos 46:1879–1885. 10.1124/dmd.118.083428 [ DOI ] [ PubMed ] [ Google Scholar ] Kamiya Y et al (2020) Determination and prediction of permeability across intestinal epithelial cell monolayer of a diverse range of industrial chemicals/drugs for estimation of oral absorption as a putative marker of hepatotoxicity. Toxicol Rep 7:149–154. 10.1016/j.toxrep.2020.01.004 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Katneni K et al (2018) Using human plasma as an assay medium in Caco-2 studies improves mass balance for lipophilic compounds. Pharm Res 35(11):210. 10.1007/s11095-018-2493-3 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Koppel N et al (2017) Chemical transformation of xenobiotics by the human gut microbiota. Science 356(6344):eaag2770. 10.1126/science.aag2770 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Küblbeck J et al (2016) Genetically modified caco-2 cells with improved cytochrome P450 metabolic capacity. J Pharm Sci 105:941–949. 10.1016/s0022-3549(15)00187-2 [ DOI ] [ PubMed ] [ Google Scholar ] Lampen A et al (1998) Catalytic activities, protein- and mRNA-expression of cytochrome P450 isoenzymes in intestinal cell lines. Xenobiotica 28:429–441. 10.1080/004982598239362 [ DOI ] [ PubMed ] [ Google Scholar ] Lechanteur A et al (2017) Elucidation of the impact of cell culture conditions of Caco-2 cell monolayer on barrier integrity and intestinal permeability. Eur J Pharm Biopharm 119:137–141. 10.1016/j.ejpb.2017.06.013 [ DOI ] [ PubMed ] [ Google Scholar ] Linz G et al (2020) Cell barrier characterization in transwell inserts by electrical impedance spectroscopy. Biosens Bioelectron 165:112345. 10.1016/j.bios.2020.112345 [ DOI ] [ PubMed ] [ Google Scholar ] Lozoya-Agullo I et al (2017) Usefulness of Caco-2/HT29-MTX and Caco-2/HT29-MTX/RajiB coculture models to predict intestinal and colonic permeability compared to Caco-2 monoculture. Mol Pharm 14(4):1264–1270 [ DOI ] [ PubMed ] [ Google Scholar ] Nienstedt W et al (1980) Metabolism of testosterone and androstenedione by human adult and fetal gastrointestinal tissues in vitro. J Steroid Biochem 13:1421–1425. 10.1021/acs.molpharmaceut.6b01165 [ DOI ] [ PubMed ] [ Google Scholar ] Panse N, Gerk P (2022) The Caco-2 model: modifications and enhancements to improve efficiency and predictive performance. Int J Pharm 624:122004. 10.1016/j.ijpharm.2022.122004 [ DOI ] [ PubMed ] [ Google Scholar ] Prueksaritanont T et al (1996) Comparative studies of drug-metabolizing enzymes in dog, monkey, and human small intestines and in Caco-2 cells. Drug Metab Dispos 24:634–642 [ PubMed ] [ Google Scholar ] Sano T et al (2001) 17 beta-hydroxysteroid dehydrogenase type 2 expression and enzyme activity in the human gastrointestinal tract. Clin Sci 101(5):485–491 [ PubMed ] [ Google Scholar ] Schedl H (1965) Absorption of steroid hormones from the human small intestine. J Clin Endocrinol Metab 25(10):1309–1316. 10.1210/jcem-25-10-1309 [ DOI ] [ PubMed ] [ Google Scholar ] Srinivasan B et al (2015) TEER measurement techniques for in vitro barrier model systems. SLAS Technology 20(2):107–126. 10.1177/2211068214561025 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Sun H et al (2008) The Caco-2 cell monolayer: usefulness and limitations. Expert Opin Drug Metab Toxicol 4:395–411. 10.1517/17425255.4.4.395 [ DOI ] [ PubMed ] [ Google Scholar ] Swiderek M et al (2013) Technical bulletin #421 effects of ECM proteins on barrier formation in Caco-2 cells. Effects of ECM Proteins on Barrier Formation in Caco-2 Cells. Accessed 30 Jul 2025 Varma M (2010) Role of intestinal transporters and metabolism in the oral absorption of drug and prodrugs. Curr Drug Metab 11:715. 10.2174/138920010794328878 [ DOI ] [ PubMed ] [ Google Scholar ] Varma M et al (2010) Physicochemical space for optimum oral bioavailability: contribution of human intestinal absorption and first-pass elimination. J Med Chem 53(3):1098–1108. 10.1021/jm901371v [ DOI ] [ PubMed ] [ Google Scholar ] Velicky M et al (2011) In situ artificial membrane permeation assay under hydrodynamic control: correlation between drug in vitro permeability and fraction absorbed in humans. Eur J Pharm Sci 44(3):299–309. 10.1016/j.ejps.2011.08.007 [ DOI ] [ PubMed ] [ Google Scholar ] Volpe D et al (2007) Classification of drug permeability with a Caco-2 cell monolayer assay. Clin Res Regul Aff 24(1):39–47. 10.1080/10601330701273669 [ Google Scholar ] Volpe D. (2022) BCS methodology: solubility, permeability & dissolution. FDA (Food and drug administration), Division of applied regulatory Science, Center for drug evaluation and research. BCS Methodology: Solubility, Permeability & Dissolution. Accessed 01 Aug 2025 Yamada N et al (2023) Generation of Caco-2 cells with predictable metabolism by CYP3A4, UGT1A1 and CES using the PITCh system. Drug Metab Pharmacokinet 50:100497. 10.1016/j.dmpk.2023.100497 [ DOI ] [ PubMed ] [ Google Scholar ] Yamashita S et al (2000) Optimized conditions for prediction of intestinal drug permeability using Caco-2 cells. Eur J Pharm Sci 10:195–204. 10.1016/s0928-0987(00)00076-2 [ DOI ] [ PubMed ] [ Google Scholar ] Yang Y et al (2007) Biopharmaceutics classification of selected β-blockers: solubility and permeability class membership. Mol Pharm 4(4):608–614. 10.1021/mp070028i [ DOI ] [ PubMed ] [ Google Scholar ] Zhu C et al (2002) A comparative study of artificial membrane permeability assay for high throughput profiling of drug absorption potential. Eur J Med Chem 37:399–407. 10.1016/s0223-5234(02)01360-0 [ DOI ] [ PubMed ] [ Google Scholar ] Associated Data This section collects any data citations, data availability statements, or supplementary materials included in this article. Supplementary Materials Supplementary Material 1 (2.5MB, docx) Data Availability Statement All data generated or analyzed during this study are included in this published article (and its supplementary information files). Articles from Archives of Toxicology are provided here courtesy of Springer ACTIONS View on publisher site PDF (5.1 MB) Cite Collections Permalink RESOURCES Similar articles Cited by other articles Links to NCBI Databases Follow NCBI NCBI on X (formerly known as Twitter) NCBI on Facebook NCBI on LinkedIn NCBI on GitHub NCBI RSS feed Connect with NLM NLM on X (formerly known as Twitter) NLM on Facebook NLM on YouTube National Library of Medicine 8600 Rockville Pike Bethesda, MD 20894 Web Policies FOIA HHS Vulnerability Disclosure Help Accessibility Careers NLM NIH HHS USA.gov Back to Top

Related documents

• Investigate the Safety, Tolerability and Pharmacokinetics of CHF6523 in Healthy and in COPD Subjects#1096100
• A Study to Evaluate How Multiple Doses of Itraconazole Interacts With Nacresertib in Healthy Adult Participants#841398
• In vivo and in vitro assessment of absorption and disposition kinetics of bis[2-(perfluorohexyl)ethyl] phosphate in rats: food effect, physiologically-based modeling, and human exposure estimation#1002095
• Toxicokinetic Study of Lambda-cyhalothrin Biomarkers of Exposure#1140780
• Katritzky, A. R.; Kasemets, K.; Slavov, S.; Radzvilovits, M.; Tämm, K.; Karelson, M. Estimating the toxicities of organic chemicals in activated sludge process. Water Res. 2010, 44, 2451–2460.#933094