Cellular uptake kinetics are key for understanding time-dependent chemical exposure in in vitro cell assays. Slow cellular uptake kinetics in relation to the total exposure time can considerably reduce the biologically effective dose. In this study, fluorescence microscopy combined with automated image analysis was applied for time-resolved quantification of cellular uptake of 10 neutral, anionic, cationic, and zwitterionic fluorophores in two reporter gene assays. The chemical fluorescence in the medium remained relatively constant during the 24-h assay duration, emphasizing that the proteins and lipids in the fetal bovine serum (FBS) supplemented to the assay medium represent a large reservoir of reversibly bound chemicals with the potential to compensate for chemical depletion by cell uptake, growth, and sorption to well materials. Hence FBS plays a role in stabilizing the cellular dose in a similar way as polymer-based passive dosing, here we term this process as serum-mediated passive dosing (SMPD). Neutral chemicals accumulated in the cells up to 12 times faster than charged chemicals. Increasing medium FBS concentrations accelerated uptake due to FBS-facilitated transport but led to lower cellular concentrations as a result of increased sorption to medium proteins and lipids. In vitro cell exposure results from the interaction of several extra- and intracellular processes, leading to variable and time-dependent exposure between different chemicals and assay setups. The medium FBS plays a crucial role for the thermodynamic equilibria as well as for the cellular uptake kinetics, hence influencing exposure. However, quantification of cellular exposure by an area under the curve (AUC) analysis illustrated that, for the evaluated bioassay setup, current in vitro exposure models that assume instantaneous equilibrium between medium and cells still reflect a realistic exposure because the AUC was typically reduced less than 20% compared to the cellular dose that would result from instantaneous equilibrium.

High-throughput in vitro reporter gene assays are increasingly applied to assess the potency of chemicals to alter specific cellular signaling pathways. Genetically modified reporter gene cell lines provide stable readouts of the activation of cellular receptors or transcription factors of interest, but such reporter gene assays have been criticized for not capturing cellular metabolism. We characterized the metabolic activity of the widely applied AREc32 (human breast cancer MCF-7), ARE-bla (human liver cancer HepG2), and GR-bla (human embryonic kidney HEK293) reporter gene cells in the absence and in the presence of benzo[a]pyrene (BaP), an AhR ligand known to upregulate cytochrome P450 in vitro and in vivo. We combined fluorescence microscopy with chemical analysis, real-time PCR, and ethoxyresorufin-O-deethylase activity measurements to track temporal changes in BaP and its metabolites in the cells and surrounding medium over time in relation to the expression and activity of metabolic enzymes. Decreasing BaP concentrations and formation of metabolites agreed with the high basal CYP1 activity of ARE-bla and the strong CYP1A1 mRNA induction in AREc32, whereas BaP concentrations were constant in GR-bla, in which neither metabolites nor CYP1 induction was detected. The study emphasizes that differences in sensitivity between reporter gene assays may be caused not only by different reporter constructs but also by a varying biotransformation rate of the evaluated parent chemical. The basal metabolic capacity of reporter gene cells in the absence of chemicals is not a clear indication because we demonstrated that the metabolic activity can be upregulated by AhR ligands during the assay. The combination of methods presented here is suitable to characterize the metabolic activity of cells in vitro and can improve the interpretation of in vitro reporter gene effect data and extrapolation to in vivo human exposure.