Intestinal enteroids are stem cell-based “mini-guts” that mimic many aspects of the corresponding epithelial barrier in vivo. Here, we established and characterized differentiated apical-out (AO) and basal-out (BO) jejunal enteroids in suspension and followed their differentiation by quantitative global proteomics and different microscopic techniques. The barrier integrity and function and subcellular location of nutrient and clinically important drug transporters were investigated in the matured enteroids using live-cell microscopy. The presystemic metabolism of two drugs by CYP3A4 was determined and the results were used to predict the pharmacokinetics after oral administration by a PBPK population model. The differentiated AO enteroids displayed a protein profile that overlapped both qualitatively and quantitatively with that of freshly isolated jejunal enterocytes and tissue. They exhibited a morphology that recapitulates the mature villus enterocyte in vivo, formed an intact barrier with a well-developed glycocalyx and are impermeable to the hydrophilic low molecular weight compound lucifer yellow and transported a medium chain fatty acid derivative by FAT4 into lipid deposits. The clinically important ABC-transporters Pgp and BCRP were expressed at near in vivo levels, had the correct subcellular localization and effluxed their substrates. Terfenadine and midazolam were metabolized by CYP3A4 and the results were used to predict the clinical pharmacokinetics of the drugs after oral administration with good accuracy. We conclude that suspended 3D AO enteroids provide a physiologically relevant model for studies of intestinal function that offers convenient access to the apical surface and is easy to dispense in multi-wells formats for large scale experimentation.

Enterocytes, the primary absorptive cells in the small intestinal epithelium, play a critical role in the presystemic biotransformation of orally administered drugs and other ingested chemicals. The assessment of intestinal drug metabolism is essential for determining the pharmacological efficacy and safety of new drug candidates. However, current in vitro models fall short in accurately mimicking human intestinal metabolic processes. Our study investigates the use of freshly isolated enterocytes from human jejunal tissue for drug metabolism research. Enterocytes were isolated from human jejunum using a non-enzymatic method, resulting in enterocyte isolations with high yield, high viability, and low apoptosic activity. ADME-relevant enzyme and transporter protein levels in enterocytes showed a strong correlation (R=0.82) with human jejunal mucosa. The metabolic activity of selected Phase I enzymes, including CES1/2, CYP3A4, CYP2C9, CYP1A2, CYP2D6, and CYP2C19, was evaluated, demonstrating their functionality in the isolated enterocytes. Of these CES1/2, CYP2C19, CYP2D6, and CYP3A4 seemed to display the highest activity. We conclude that freshly isolated human enterocytes are a promising alternative for in vitro studies of intestinal presystemic metabolism. 

Intestinal drug permeability is commonly studied in vitro using monolayer forming cells such as the Caco-2 or MDCK cell lines grown on a permeable support. These models allow both passive and active transport of drugs in solution to be studied. However, in the small intestine different processes, e.g. release, dissolution and digestion, are occurring simultaneously with permeation which can affect the resulting measured flux. This is especially true for drugs in need of enabling formulations and advanced drug delivery systems.

In this study we therefore established a co-culture model of two monolayer-forming MDCK cell lines, that overexpressed human multidrug resistance protein 1 (hMDR1) and human breast cancer resistance protein (hBCRP). These are the two most important efflux transporters expressed in the small intestine, of importance to drug absorption and drug-drug interactions at the intestinal level. We further validated the co-culture model by assessing the transport of substrates specific for, or shared between, these two transporters. Specific substrates were ritonavir, verapamil and labetalol (hMDR1) and dantrolene and fluvastatin (hBCRP), whereas the shared hMDR1/hBCRP substrate used was prazosin. Further, to evaluate the efficacy of this co-culture for assessing advanced drug delivery systems, we used it as the absorption layer in an in house developed enabling absorption (ENA) device. This is a device for simultaneous in vitro release/dissolution/digestion and permeation studies, and we have here used our co-culture in the ENA for investigating the dissolution and absorption of amorphous ritonavir.

Our results show that the co-culture enabled interactions with both hMDR1 and hBCRP to be studied. Further, using ENA we showed that this co-culture allowed both the dissolution and the absorption of ritonavir to be studied in a setup capturing the more dynamic processes occurring in vivo in the small intestine. In contrast to the dissolved ritonavir administered in a traditional transwell system on top of the co-culture monolayer, the ENA model included the impact of dissolution and supersaturation on absorption and its potential to influence the efflux observed in the system. We anticipate that this co-culture can be further optimized to mimic transporter expression levels found in different intestinal segments and thereby provide an in vitro model for studying the role of active efflux in absorption of drugs in need advanced formulations.