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. 2018 Mar 27;2:156–167. doi: 10.1016/j.isci.2018.03.015

Bioprinted 3D Primary Human Intestinal Tissues Model Aspects of Native Physiology and ADME/Tox Functions

Lauran R Madden 1, Theresa V Nguyen 2, Salvador Garcia-Mojica 1, Vishal Shah 1, Alex V Le 1, Andrea Peier 3, Richard Visconti 3, Eric M Parker 3, Sharon C Presnell 1, Deborah G Nguyen 1, Kelsey N Retting 1,4,∗
PMCID: PMC6135981  PMID: 30428372

Summary

The human intestinal mucosa is a critical site for absorption, distribution, metabolism, and excretion (ADME)/Tox studies in drug development and is difficult to recapitulate in vitro. Using bioprinting, we generated three-dimensional (3D) intestinal tissue composed of human primary intestinal epithelial cells and myofibroblasts with architecture and function to model the native intestine. The 3D intestinal tissue demonstrates a polarized epithelium with tight junctions and specialized epithelial cell types and expresses functional and inducible CYP450 enzymes. The 3D intestinal tissues develop physiological barrier function, distinguish between high- and low-permeability compounds, and have functional P-gp and BCRP transporters. Biochemical and histological characterization demonstrate that 3D intestinal tissues can generate an injury response to compound-induced toxicity and inflammation. This model is compatible with existing preclinical assays and may be implemented as an additional bridge to clinical trials by enhancing safety and efficacy prediction in drug development.

Subject Areas: In Vitro Toxicology Including 3D Culture, Bioengineering, Tissue Engineering

Graphical Abstract

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Highlights

  • •

    Bioprinted 3D human intestinal tissues enable complex modeling of ADME/Tox in vitro

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    3D intestinal tissues develop barrier function and polarized transporter expression

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    Key cytochrome P450 enzymes are expressed, metabolically active, and inducible

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    GI toxicants can trigger barrier disruption and cytotoxicity in 3D intestinal tissues

In Vitro Toxicology Including 3D Culture; Bioengineering; Tissue Engineering

Introduction

Systemic availability, diminished efficacy, and off-target effects remain challenges to the successful clinical prediction of candidate drugs and contribute to attrition in drug development, highlighting the need for better preclinical tools to model intestinal function in vitro (Alqahtani et al., 2013, Jones et al., 2016, Peters et al., 2016). Oral delivery is the most common method for drug administration, and in addition to absorption and first-pass metabolism, the intestine serves directly as a site of off-target toxicity for compounds such as non-steroidal anti-inflammatory drugs (NSAIDs) (Boelsterli et al., 2013) and chemotherapeutic agents (Aprile et al., 2015) and indirectly as a site of drug-drug interactions (Bentz et al., 2013, Peters et al., 2016). Animal models are frequently used to estimate compound bioavailability; however, species differences can lead to disparity in the expression of metabolic enzymes and transporters, resulting in poor clinical prediction (Jones et al., 2016, Musther et al., 2014, Peters et al., 2016). Human intestinal slices can provide correct cellular architecture and physiological complexity; however, limited viability ex vivo (about 24 hr) and incompatibility with cryopreservation restrict use to short-term studies (Li et al., 2016). Predominant in vitro systems used to study intestinal bioavailability and toxicity include intestinal microsomes and two-dimensional (2D) monolayers. Microsomes are a convenient tool for initial assessment of metabolism, but cannot model cellular level outcomes. Physiological conditions lacking in 2D culture may be improved by microfluidic chip models; however, maintenance of cell-cell and cell-matrix interactions and scale-up for high-throughput applications remain challenging (Bhatia and Ingber, 2014, Chi et al., 2016). Standard 2D monolayers and more recent gut-on-a-chip systems commonly include cell lines originating from colorectal and duodenal tumor tissue (Alqahtani et al., 2013, Kim et al., 2012) that may exhibit altered metabolic profiles and expression patterns of xenobiotic-metabolizing enzymes and transporters potentially leading to misrepresentation of the native response (Prueksaritanont et al., 1996, Yamaura et al., 2016). The colorectal tumor-derived Caco-2 cell line is the most established model to study permeability and predict intestinal absorption. Known limitations of the Caco-2 model include reduced CYP450 activity (Prueksaritanont et al., 1996, Yamaura et al., 2016), variability with passage number, and inconsistencies between subclones of the cell line (Bentz et al., 2013).

Limitations of common in vitro models have sparked the development of methods utilizing primary human intestinal cells. Although primary monocultures more closely resemble in vivo tissue compared with cell lines (Kauffman et al., 2013, Takenaka et al., 2016), separation of the epithelium from supportive cells may impair function and limit the ability to study complex cell interactions including interplay with interstitial cells or inflammatory responses. The discovery of organoids to expand primary cells (Sato et al., 2011, Yin et al., 2014) or differentiate pluripotent stem cells (Kauffman et al., 2013, Watson et al., 2014) revealed another path to model the intestine in vitro. Organoids derived from all regions of the intestinal tract (Wang et al., 2015) can be applied to intestinal research including organ development, disease modeling, and regenerative medicine (Fatehullah et al., 2016, Sinagoga and Wells, 2015), although they may lack in vivo organ physiology. Notably, the closed lumen of the organoid structure with inward epithelial orientation makes the apical surface relatively inaccessible for absorption, distribution, metabolism, and excretion (ADME) assays. Approaches to create accessible compartments can require a complex bioreactor setup (Schweinlin et al., 2016) and produce monolayers with non-physiological transepithelial electrical resistance (TEER) (Kauffman et al., 2013, VanDussen et al., 2015). Cell-seeded engineered three-dimensional (3D) tubular scaffolds have recently been utilized with hollow lumens to promote a more native-like microenvironment (Chen et al., 2017); however, functional characterization is restricted to the apical compartment by geometry, making most standard ADME/Tox assays that require access to both apical and basal compartments incompatible.

To overcome existing limitations of the current in vitro systems, a proprietary bioprinting platform was utilized to develop a multicellular 3D primary human intestinal tissue model to better mimic native tissue structure and function (Nguyen et al., 2016, King et al., 2017). The fully human primary cell-derived tissue designed with laminar architecture incorporates a polarized intestinal epithelium supported by an interstitial tissue layer allowing compatibility with standard histological and biochemical ADME/Tox readouts. Tissues maintained viability over 2 weeks in culture, developed barrier function, and demonstrated selective permeability. Presence of functional key CYP450 metabolic enzymes and transporters was confirmed with endogenous expression similar to native intestine. Last, tissue injury was evaluated in response to known toxicants indomethacin and tumor necrosis factor alpha (TNF-α) revealing barrier disruption and cytotoxicity at the biochemical and histological levels. The human tissue combined with the reproducibility of the automated 3D bioprinting platform and compatibility with standard assay approaches make this system a practical in vitro tool for ADME/Tox applications across drug development.

Results

Histological Characterization of 3D Human Intestinal Tissue

The Organovo 3D NovoGen Bioprinter system was used to create an in vitro model of the human intestine. The tissue was generated with bilayered architecture, consisting of a human intestinal myofibroblast (IMF) interstitium supporting an epithelial layer containing human intestinal epithelial cells (hIEC) and maintained up to 21 days in culture. Tissues were printed on cell culture inserts (Figure 1A) allowing access to both apical and basolateral surfaces for direct compound testing. Histological analysis at day 17 showed that tissues exhibited polarized columnar epithelial morphology and secondary structure formation (Figures 1B–1I). The epithelial and interstitial tissue compartments are in direct contact with each other but remained distinct, with correct expression of epithelial (CK19) and myofibroblast (vimentin) cell-specific markers (Figure 1C). E-cadherin and ZO-1, key proteins involved in tight junction formation and barrier function, were expressed between epithelial cells of the hIEC layer (Figure 1D). Correct polarization of the hIEC and brush border formation were seen by positive staining for brush border protein villin at the apical surface (Figure 1E). Periodic acid-Schiff (PAS)/Alcian blue staining confirmed an apical brush border and suggested the presence of a subpopulation of goblet cells as well as the excretion of mucus (Figure 1F). Immunohistochemistry for mucin-2 confirmed the presence of goblet cells and mucous secretion (Figure 1G), a feature similar to native tissue and an indication of normal intestinal function. In addition to goblet cells, other specialized cell types present in the intestinal epithelium included lysozyme-positive Paneth cells and chromogranin-expressing enteroendocrine cells (Figures 1H and 1I). Patterning of 3D intestinal tissues was similar to that observed in native intestinal tissue and consistent between multiple donors (Figure S1). Tissue architecture of the 3D intestinal model was also compared with that of standard Caco-2 monolayer culture (Figure S2). Caco-2 monolayers achieved a polarized epithelial phenotype with tight junction formation, although epithelial morphology appeared less columnar and lacked secondary structure formation. Subpopulations of specialized cells and evidence of mucus production were also absent in Caco-2 monolayers (Figure S2). Substitution of hIEC with Caco-2 cells in the 3D bioprinted model produced a bilayered structure with increased thickening of the epithelial layer and cyst formation over time, akin to the cancer origin (Figure S3A). However, this phenotype change did not improve transporter or enzyme gene expression to the level of 3D intestinal tissues fabricated with primary hIECs (Figure S3B). Thus the standard Caco-2 monolayer was used as the main comparison to bioprinted tissues composed of primary hIECs.

Figure 1.

Figure 1

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Architecture of 3D Intestinal Tissue

(A) A bilayered architecture is achieved by bioprinting an interstitial layer containing adult human intestinal myofibroblasts (IMF) followed by adult human intestinal epithelial cells (hIEC). The vimentin-expressing interstitial cells and CK19-expressing epithelial compartments remain separate over 17 days in culture (B, C). Epithelial cells express markers necessary for barrier formation such as E-cadherin (D), ZO-1 (D, inset), and apically expressed villin (E). Mucus production from goblet cells (* denotes secreted mucus, arrow indicates goblet cells) is observed throughout culture (F, G). Lysozyme-stained Paneth cells (H) and chromogranin-expressing enteroendocrine cells (I) are also present.

Characterization of Gene Expression

Gene expression analysis was utilized to further evaluate the expression of key intestinal epithelial tissue markers, metabolic enzymes, and transporters in the 3D intestinal tissue and was compared with both native donor intestinal tissue and standard Caco-2 monolayers (Figure 2). Although five native donor tissues were processed, challenges in obtaining high-quality donor material resulted in the utilization of three of five donors for gene expression analysis. To specifically study differential expression in the epithelium and to remove any variance in the total cell number, genes were analyzed relative to the expression of epithelial-specific marker CK19. Heatmap visualization shows close clustering of 3D intestine to the native intestine compared with more disparate Caco-2 monolayers (Figure 2A). In support of histological findings, tight junction marker E-cadherin (CDH1) and brush border marker villin (VIL1) were highly expressed, as well as markers for specialized cell subpopulations including Paneth cells (LYZ), enteroendocrine cells (CHGA), and goblet cells (MUC2) (Figure 2B). Key xenobiotic-activated nuclear receptors involved in drug metabolism and disposition including VDR, PXR (NR1I2), and CXR (NR1I3) were expressed in 3D intestinal tissue at comparable levels to native intestine. Major intestinal Phase I CYP450 metabolic enzymes including CYP3A4, CYP2C9, CYP2C19, CYP2D6, and CYP2J2 were also detected. Clinically important CYP3A4 was highly expressed in the 3D intestinal model, whereas it was notably not detectable in Caco-2 monolayers (Figure 2B). Key intestinal Phase II metabolic enzymes GSTP1 and UGT1A1 were expressed, as well as regulators of fatty acid metabolism, including DGAT1, MOGAT2, and MTTP. Major efflux transporters P-gp (ABCB1, MDR1) and BCRP (ABCG2), key uptake transporters PEPT1 (SLC15A1) and OATP2B1 (SLCO2B1), and intestinal bile acid-related transporters ASBT (SLC10A2), OSTa (SLC51A), and OSTb (SLC51B) were expressed in 3D intestinal tissues with levels comparable with those in native intestine (Figure 2B). Collectively, gene expression levels in 3D intestinal tissues corresponded to endogenous levels observed in native tissue, whereas Caco-2 monolayers exhibited reduced, overexpressed, or absent expression of several genes suggesting that the bioprinted model more closely resembles native tissue.

Figure 2.

Figure 2

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Gene Expression Comparisons of Native Intestine, 3D Intestinal Tissue, and Caco-2 Monolayers

(A) Heatmap clustering shows a closer relationship between native intestine and 3D intestinal tissue than Caco-2 monolayers (native intestine N = 3 donors, 3D intestine n = 3 biological replicates per donor, and Caco-2, n = 3 biological replicates).

(B) General markers for intestinal barrier function (CDH1, VIL1) are similar among the three groups. Epithelial subtype markers are highest for native and 3D intestinal tissue. Drug-inducible transcription factor NR1I2 (PXR) is similar for native intestine and 3D intestinal tissue. Native intestine and 3D intestinal tissue express all metabolic enzymes analyzed, including Phase I cytochrome P450s, the majority of which Caco-2 cells lack. Transporters are expressed by all three groups, with variation in the level. For efflux transporters, 3D intestine is most similar to native intestine, whereas Caco-2 cells underexpress ABCB1 (P-gp) and ABCG2 (BCRP). Uptake transporters SLC15A1 (PEPT1) and SLCO2B1 (OATP2B1) are similarly expressed by native and 3D intestine, but not by Caco-2 monolayers. Data in (B) are expressed as mean ± SD.

Demonstration of Cytochrome P450 Metabolic Function in 3D Intestinal Tissue

Functional assays were performed to evaluate the activity and specificity of CYP3A4 and CYP2C9, two major intestinal Phase I cytochrome P450 metabolic enzymes profiled in gene expression studies (Figure 3). CYP2C9 activity was readily detected in 3D intestinal tissue by luminogenic P450 substrate conversion and could be significantly inhibited by sulfaphenazole (Figure 3A). CYP3A4 activity and specific inhibition by ketoconazole were confirmed by both a luminogenic P450 substrate conversion (Figure 3B) and midazolam metabolite formation (Figures 3D and 3E). Luminogenic substrate conversion occurred at a higher level in 3D intestine compared with Caco-2 monolayers (Figure 3C), and only 3D tissues were induced by rifampicin treatment (Figures 3C and 3E). Rifampicin treatment was associated with significantly higher turnover of CYP3A substrates in treated tissues and significant upregulation of PXR-inducible genes including CYP2C9, CYP3A4, CYP3A5, P-gp, and UGT1A1 (Figures 3E and 3F). Variation in human CYP3A-dependent metabolism is well established (Paine et al., 1997). Comparison of CYP3A4 activity by midazolam turnover for 3D intestinal tissues fabricated from three separate donors showed expected interindividual donor variation (Donor 1: 19.3 ± 1.9; Donor 2: 2.85 ± 0.59; Donor 3: 3.6 ± 3.6 pmol/min/mg) consistent with gene expression data (Figure 2A) as well as demonstrated reproducibility of ketoconazole inhibition and rifampicin induction between donors (Table S1). The degree of variability in CYP3A4 activity was consistent with observations of native intestine gene expression in the current study (6.3-fold among donors, Figure 2A) and with previous studies evaluating intestinal CYP3A4 activity (18-fold among donors) (Paine et al., 1997).

Figure 3.

Figure 3

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Cytochrome P450 Metabolism in 3D Intestinal Tissue

(A) CYP2C9 basal activity was validated by a luciferin activity assay and was reduced by inhibitor sulfaphenazole (n = 6). CYP3A4 activity was shown by luciferin activity assay (n = 3–4) (B, C) and midazolam hydroxylation (n = 5) (D, E), both of which were inhibited by ketoconazole. (C, E) CYP3A4 induction via rifampicin treatment was detected by increased luciferin activity and midazolam metabolism in 3D intestine (n = 5). (C) A comparison of CYP3A4 activity in 3D intestinal tissue with 2D Caco-2 monolayer shows low activity of CYP3A4 in Caco-2 monolayers with no rifampicin induction (n = 4–11 for 3D intestine, n = 3 for 2D Caco-2). (F) Rifampicin treatment increased gene expression of PXR-regulated CYPs, ABCB1 (P-gp), and UGT1A1 but not CK19 and ECAD (n = 4). Level of significance: ****p < 0.0001, ***p < 0.001, **p < 0.01 by two-way ANOVA. Data are expressed as mean ± SD.

Demonstration of Barrier Function

The intestine is a selectively permeable barrier, regulating absorption of both nutrients and xenobiotics. TEER was utilized to measure barrier function in 3D intestinal tissues over a 21-day culture period. Measurements demonstrated that the tissues developed and maintained barrier function between days 10 and 21 of culture, exhibiting values within a physiological range (50–100 Ω*cm2) comparable to native human intestine (Srinivasan et al., 2015) (Figure 4A). Representative compounds with high and low permeability were used to further validate 3D intestinal tissue barrier function (Figure 4B, Table S2). Paracellular transport marker lucifer yellow correctly demonstrated low permeability in 3D intestine (Papp = 0.45 ± 1.5 × 106 cm/s), suggesting the presence of an intact physical barrier for drug transport. The 3D intestinal tissues correctly distinguished between low-, intermediate-, and high-permeability compounds. Mitoxantrone (Papp = 0.2 ± 0.03 × 106 cm/s) is a low-permeability compound with absorption mainly mediated by ABCG2 (BCRP) efflux transporter. Topotecan and digoxin showed Papp values in the intermediate range (3.1 ± 0.63 × 106 cm/s and 8.53 ± 2.02 × 106 cm/s, respectively), whereas propranolol, a passive transcellular transport reference compound, showed the highest permeability (Papp = 14.8 ± 1.96 × 106 cm/s).

Figure 4.

Figure 4

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Barrier Function of 3D Intestinal Tissue

(A) Transepithelial electrical resistance (TEER) was measured for 3D intestinal tissue from day 6 to day 21, showing increase early in culture, with maintenance of barrier function within physiological levels (dotted lines) after day 10 (n = 24).

(B) Permeability of test compounds for 3D intestine, measured in the apical to basal direction, shows distinction between low- (mitoxantrone [n = 4]), moderate- (topotecan [n = 4], digoxin [n = 6]), and high- (propranolol [n = 5]) permeability compounds.

(C) Comparison of TEER values shows more physiological TEER values for 3D intestine (n = 24) and 3D Caco-2 (n = 6) versus 2D Caco-2 (n = 5), whereas IMF alone does not show barrier function (n = 19).

(D) Comparison of lucifer yellow permeability values between models shows low permeability for all epithelial-cell-containing models and high permeability for IMF only (n = 4 for 2D and 3D Caco-2 and IMF only, n = 28 for 3D intestine). Data are expressed as mean ± SD.

In comparison, Caco-2 monolayers demonstrated significantly higher TEER measurements (840 ± 55 Ω*cm2) (Figure 4C). When substituted for hIEC in the 3D bioprinted model, 3D Caco-2 tissues had reduced TEER (254 ± 45 Ω*cm2) compared with their 2D counterparts, a possible effect of cyst-like secondary structure formation, but the values remained above a physiological range. Both 2D and 3D Caco-2 models had low lucifer yellow permeability (Papp = 0.46 ± 0.25 × 106 cm/s and 1.05 ± 0.1 × 106 cm/s, respectively) confirming barrier formation, whereas IMF tissue alone did not show barrier function (5 ± 3 Ω*cm2) and exhibited high permeability (163 ± 29 × 106 cm/s) (Figure 4D).

Transporter Localization and Function in 3D Intestinal Tissue Model

Intestinal efflux and uptake transporters can be both sites of drug-drug interactions and limiting factors for drug absorption. Immunohistochemical staining confirmed the correct polarized epithelial expression of key efflux transporters at the apical (P-gp [ABCB1], BCRP [ABCG2], MRP2 [ABCC2]) and basolateral (MRP3 [ABCC3]) surfaces in a continuous pattern similar to native intestine (Figures 5A, 5B, and S4). To assess the capability of 3D intestinal tissues to predict active efflux, P-gp and BCRP function were evaluated by measuring bidirectional transport with and without inhibition (Figures 5C and 5D). Under control conditions, asymmetric permeability of P-gp substrate digoxin was observed with an efflux ratio greater than 2. Inhibition of P-gp by zosuquidar (Dantzig et al., 1996) decreased the rate of B to A transport, reducing the efflux ratio to 1.2, to confirm the activity of P-gp. BCRP function was tested by efflux of topotecan (Figure 5D) and mitoxantrone (Figure S5). BCRP/P-gp substrate topotecan and BCRP substrate mitoxantrone were preferentially transported in the B to A direction with efflux ratios of 8.8 and 190, respectively. Furthermore, subsequent inhibition of topotecan transport by the BCRP inhibitor Ko143 (Allen et al., 2002) reduced the efflux ratio to 3.6 and dual inhibition with Ko143 and zosuquidar further decreased the efflux ratio to 1.4. Collectively, these results demonstrate that the 3D intestinal tissue model expresses clinically relevant P-gp and BCRP transporters with proper localization and function. Compounds assessed demonstrate greater efflux ratio similarity to other models composed of primary human intestinal cells than to Caco-2 models (Table S3). Histological analysis showed apical P-gp and BCRP expression in patches across Caco-2 monolayers, whereas MRP2 and MRP3 appeared to be overexpressed (Figures 5A, 5B, and S4). Transporter expression patterning was not rescued by substitution of Caco-2 cells into the 3D bioprinted model (Figure S3).

Figure 5.

Figure 5

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P-gp and BCRP Transporter Function in 3D Intestinal Tissue

(A and B) (A) P-gp and (B) BCRP are apically localized in 3D intestinal tissue, similar to native intestine. Expression in Caco-2 monolayers was in patches at the apical surface.

(C) P-gp substrate digoxin had greater permeability in the B to A direction with efflux ratio of 2.1. In the presence of P-gp inhibitor zosuquidar, the efflux ratio of digoxin reduced to 1.2 (n = 6).

(D) BCRP/P-gp substrate topotecan had an efflux ratio of 8.8 under control conditions. BCRP inhibition by Ko143 reduced B to A transport and the efflux ratio decreased to 3.6. Dual inhibition of P-gp and BCRP resulted in ablation of transport with an efflux ratio of 1.4 (n = 3–4). Level of significance: ****p < 0.0001 by two-way ANOVA. Data in (C) and (D) are expressed as mean ± SD.

Characterization of 3D Intestinal Tissues as a Model for Gastrointestinal Toxicity

The utility of the 3D intestinal model for compound toxicity applications was evaluated by the NSAID indomethacin, a prostaglandin E2 (PGE2) oxygenase inhibitor and known gastrointestinal (GI) toxicant that can reduce intestinal epithelial barrier function through enterocyte apoptosis and necrosis (Boelsterli et al., 2013). The 3D intestinal tissue showed a dose-dependent decrease in barrier function as measured by TEER in response to 24-hr indomethacin treatment (Figure 6A). Injury to the intestinal cells was detected by lactate dehydrogenase (LDH) release and was significantly increased in the presence of indomethacin doses above 0.25 mM (Figure 6B), consistent with previous reports modeling indomethacin injury in vitro (Tang et al., 1993). All doses of indomethacin demonstrated inhibition of prostaglandin E2 secretion, confirming the known mechanism of activity (Figure 6C). Histological analysis further illustrated dose-dependent disruption of epithelial morphology with decreased E-cadherin expression, consistent with loss in barrier function (Figure 6D). Release of inflammatory cytokines may also damage the intestine, and TNF-α is a major contributor to disease-related inflammation and intestinal toxicity (Neurath, 2014, Peyrin-Biroulet, 2010). To evaluate the susceptibility of the model to pro-inflammatory cytokines, tissues were dosed with TNF-α for 24 hr and evaluated for changes in morphology, LDH release, and gene expression (Figure 7). TNF-α altered the epithelial morphology and resulted in the dissociation of cells from the interstitial layer accompanied by a significant increase in LDH release, suggesting a cytotoxic response and enterocyte death. Gene expression for several known targets of the TNF-α/nuclear factor (NF)-κB pathway including interleukin (IL)-6, IL-8, CCL2, CXCL10, ICAM1, and TNF-α were upregulated, demonstrating activation of an inflammatory cascade (Figure 7C) in response to tissue injury.

Figure 6.

Figure 6

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Indomethacin Toxicity in 3D Intestinal Tissue

(A) TEER measurements of tissues following 24-hr incubation with ethanol vehicle or varying doses of indomethacin show dose response decrease in TEER with increasing indomethacin.

(B) LDH activity increased with increasing indomethacin dose, suggesting increased cytotoxicity.

(C) Prostaglandin E2 synthesis decreased to similar levels for all indomethacin doses tested, confirming drug activity (n = 6–7, A–C).

(D) Histology of indomethacin-treated tissues shows disruption of the epithelium and distorted nuclear staining at higher doses of indomethacin, accompanied by a reduction in E-cadherin, a marker of barrier function. Level of significance: ****p < 0.0001 by one-way ANOVA. Data in (A–C) are expressed as mean ± SD.

Figure 7.

Figure 7

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TNF-α-Induced Toxicity in 3D Intestinal Tissue

(A) Tissues treated with TNF-α for 24 hr showed increased epithelial disorganization compared with controls.

(B) Increased LDH activity correlated with changes in cell morphology following TNF-α treatment (n = 5–6).

(C) A subset of genes related to inflammation, TNF-α, IL-6, IL-8, CCL2, and ICAM, were upregulated in response to TNF-α treatment (n = 3). Level of significance: ***p < 0.001 by t test in (B); **p < 0.01, ***p < 0.001, and ****p < 0.0001 by two-way ANOVA in (C). Data in (B) and (C) are expressed as mean ± SD.

Discussion

Current preclinical models are limited in the ability to capture complexities and function of human intestinal tissue, which can lead to poor predictability in drug development (Alqahtani et al., 2013, Aprile et al., 2015, Boelsterli et al., 2013, Jones et al., 2016, Peters et al., 2016). This study incorporated primary cells into a fully human 3D bioprinted intestinal tissue model to recapitulate multiple facets of native intestinal biology and function in vitro. Histological analysis of the 3D intestinal tissues confirmed development and maintenance of a polarized epithelium with columnar epithelial morphology, tight junctions, and apical brush border formation. Stability of tissue architecture and function for greater than 2 weeks in culture suggests that the model may be suitable for extended compound studies.

In this study, the 3D intestinal model was compared with Caco-2 cells cultured as a standard 2D monolayer with results suggesting that the 3D tissues more closely mimic native tissue in gene and protein expression patterning and function. The divergence of Caco-2 cells from native tissue, consistent with previous reports (Prueksaritanont et al., 1996, Takenaka et al., 2014), may be due in part to their cancer origin and variability in expression patterns of metabolic enzymes and transporters along the GI tract (Paine et al., 1997, Peters et al., 2016). Both native donor tissue and the 3D intestinal model were derived from the ileum, whereas Caco-2 cells are derived from the colon. Substitution of primary intestinal epithelial cells with Caco-2 cells in the 3D system produced phenotypes more consistent with native tissue, including secondary structure formation, indicating a benefit to the 3D microenvironment that may be in part influenced by cross talk with interstitial cells, but did not rescue aberrant expression patterning.

Histology and gene expression analyses demonstrated the presence of specialized cell subpopulations in 3D intestinal tissues that are notably absent in cell lines including the Caco-2 model. The presence of mucin-2-expressing goblet cells, evidence of mucus production, and lysozyme-expressing Paneth cells suggests the potential for modeling mucosal barrier function and antimicrobial or microbiome function. Chromogranin-positive cells suggest the potential to study enteroendocrine function in the gut, including glucagon-like peptide (GLP)-1 signaling. Cell subpopulations indicate that primary cells in 3D intestinal tissues may be able to undergo directed differentiation by modification of culture conditions similar to maturation observed in organoid cell culture (Sato et al., 2011, Spence et al., 2011, Yin et al., 2014). Incorporation of induced pluripotent stem cells may be considered as an alternative cell source to primary cells, although maturity can resemble a fetal stage phenotype (Sinagoga and Wells, 2015) with inconsistent barrier function over time (Kauffman et al., 2013).

The 3D intestinal tissue demonstrated the expression and function of key enzymes involved in xenobiotic metabolism including CYP3A4, an enzyme known to be absent in Caco-2 models (Yamaura et al., 2016), consistent with our findings. Enzyme activity was confirmed by modulation of midazolam hydroxylation through both inhibition and induction—CYP3A4 activity and PXR-regulated genes were induced in response to rifampicin. The 3D intestinal model demonstrated reproducible midazolam metabolism in tissues separately fabricated from three different donors with interindividual donor variation as expected (Paine et al., 1997). Values were similar to those shown for intestinal slices (van de Kerkhof et al., 2006) and much higher than those reported for 2D systems (Takenaka et al., 2014, Yamaura et al., 2016), suggesting suitability of this model for drug-induced metabolic and transporter studies, which cannot be achieved by previous primary or Caco-2 models (Takenaka et al., 2014, Yamaura et al., 2016).

A differentiating factor of this 3D model over other 3D primary cell systems including organoids and more recent cell-seeded tubular scaffolds (Chen et al., 2017) is the laminar architecture that provides both apical and basal access and compatibility with standard barrier function and directional transport assays. Physiological barrier function was successfully demonstrated by TEER (74.4 ± 24.5 Ω*cm2) in 3D intestinal tissues at day 10 and maintained through day 21 of culture. Furthermore, tissues could successfully differentiate between low-, intermediate-, and high-permeability substrates such as paracellular marker lucifer yellow and transcellular marker propranolol. With the advantage of physiological TEER values, models with hIECs may yield a better correlation with in vivo permeability compared with Caco-2 monolayers with artificially high measurements (840 ± 55 Ω*cm2), which may be due in part to observed differences in E-cadherin expression (Srinivasan et al., 2015). Barrier measurements in the 3D intestinal model were consistent with reported monolayer cultures of adult intestinal epithelial cells (70–120 Ω*cm2) (Takenaka et al., 2016). These monolayer models, however, are typically limited to <11 days in culture, can suffer from low CYP expression similar to Caco-2 cells, and have not yet been assessed for toxicity or inflammatory response (Kauffman et al., 2013, Takenaka et al., 2014, Takenaka et al., 2016).

Intestinal efflux and influx transporters are key mediators of absorption for low-permeability drugs. Clinically relevant P-gp and BCRP efflux transporters, which can significantly affect the net fraction of compound absorbed (Bentz et al., 2013, Peters et al., 2016), were correctly expressed in the apical epithelium and functional in response to known substrates digoxin and topotecan, respectively. The level of efflux in the 3D intestinal tissue model was more similar to that previously observed in 2D human intestinal cell monolayers than with Caco-2 monolayers (Kauffman et al., 2013, Li et al., 2008, Takenaka et al., 2014). Expression of functional transporters suggests that this system could be applied to assess the relative contributions of efflux transporters to drug disposition, or could be used as a potential model for increased absorption by targeting uptake transporters such as PEPT1 and OATP2B1. The dual presence of transporters and enzymes in the 3D intestinal tissue model suggests that it could be used to shed light on complex drug-drug interactions, such as those seen with overlapping P-gp/CYP3A4 substrates (Kim et al., 1999). Furthermore, high expression of enzymes involved in fatty acid metabolism in the 3D intestinal tissue indicates a potential application for evaluating compounds targeting these enzymes to combat obesity (Shi and Cheng, 2009).

GI toxicity is a common clinical adverse event that cannot be accurately predicted or characterized with current in vitro or in vivo models (Aprile et al., 2015, Boelsterli et al., 2013, Peyrin-Biroulet, 2010). The NSAID indomethacin demonstrated a toxicity response in 3D intestinal tissue with a dose-dependent decrease in barrier function by TEER and corresponding epithelial cell disruption, correlating with previously reported in vitro (Tang et al., 1993, Tomisato et al., 2001) and in vivo outcomes (Boelsterli et al., 2013). The 3D intestinal tissue also responded to the toxic inflammatory stimulus TNF-α, resulting in epithelial cell death and upregulation of inflammatory genes, consistent with human data (Christophi et al., 2012) and previous Caco-2 studies (Cui et al., 2010, Treede et al., 2009). These data combined with long-term viability suggest that the 3D intestinal model may be applied to characterize other known classes of compounds that have off-target toxicity in the intestine, such as chemotherapeutics (Aprile et al., 2015), to evaluate inflammatory pathways, and to model chronic diseases, such as inflammatory bowel disease (IBD), Crohn disease, and colitis (Aprile et al., 2015, Boelsterli et al., 2013, Peyrin-Biroulet, 2010). Future applications could increase complexity by incorporating additional cell types such as immune cells or cancer cells, and by utilizing healthy and diseased donor material (Fatehullah et al., 2016, VanDussen et al., 2015), to better understand the contribution of each cell type to the disease phenotype and to better characterize candidate modulators for therapeutic intervention on disease-relevant backgrounds.

In conclusion, we provide evidence for a fully human 3D intestinal tissue model composed of primary cells with increased complexity and function compared with standard in vitro models. The 3D tissue recapitulates major intestinal features and physiological functions and is designed to provide a flexible platform compatible with barrier function, permeability, metabolism, transport, and toxicity studies.

Methods

All methods can be found in the accompanying Transparent Methods supplemental file.

Acknowledgments

This work was funded by Organovo and Merck.

Author Contributions

Conceptualization, L.R.M., T.V.N., A.P., R.V., E. M.P., S.C.P., D.G.N., K.N.R.; methodology, L.R.M, S.G.-M., V.S., D.G.N., K.N.R.; formal analysis, L.R.M., S.G.-M., V.S., A.V.L., K.N.R.; investigation, L.R.M., S.G.-M., V.S., A.V.L.; Resources, S.C.P.; writing: original draft, L.R.M., K.N.R.; writing: review and editing, L.R.M., T.V.N., V.S., A.P., R.V., E.M.P., S.C.P., D.G.N., K.N.R.; visualization, L.R.M., S.G.-M., A.V.L.; supervision, L.R.M., S.C.P., D.G.N., K.N.R.; project administration, L.R.M., T.V.N., A.P., R.V., E.M.P., S.C.P., D.G.N., K.N.R.; funding acquisition, T.V.N., A.P., R.V., E.M.P., S.C.P., D.G.N., K.N.R.

Declaration of Interests

L.R.M., S.G.-M., V.S., A.V.L., S.C.P., D.G.N., and K.N.R. are currently or have been employed by and may be shareholders of Organovo, Inc. T.V.N., A.P., R.V., and E.M.P. are currently or have been employed by and may be shareholders of Merck.

D.G.N. is also affiliated with Cellular Approaches; T.V.N. is also affiliated with Thermo Fisher Scientific; R.V. is also affiliated with Celgene Corporation; and E.M.P. is also affiliated with Synergy Partners R&D Solutions.

S.C.P. is the Chief Scientific Officer of Organovo, Inc., and is a member of its Executive Committee.

K.N.R., D.G.N., L.R.M., and Organovo, Inc., have a patent related to this work: PCT/US17/61016, ENGINEERED INTESTINAL TISSUE AND USES THEREOF, 10-NOV-2017.

Published: April 27, 2018

Footnotes

Supplemental Information includes Transparent Methods, five figures, and three tables and can be found with this article online at https://doi.org/10.1016/j.isci.2018.03.015.

Supplemental Information

Document S1. Transparent Methods, Figures S1–S5, and Tables S1–S3
mmc1.pdf (1.5MB, pdf)

References

  1. Allen J.D., van Loevezijn A., Lakhai J.M., van der Valk M., van Tellingen O., Reid G., Schellens J.H., Koomen G.J., Schinkel A.H. Potent and specific inhibition of the breast cancer resistance protein multidrug transporter in vitro and in mouse intestine by a novel analogue of fumitremorgin C. Mol. Cancer Ther. 2002;1:417–425. [PubMed] [Google Scholar]
  2. Alqahtani S., Mohamed L.A., Kaddoumi A. Experimental models for predicting drug absorption and metabolism. Expert Opin. Drug Metab. Toxicol. 2013;9:1241–1254. doi: 10.1517/17425255.2013.802772. [DOI] [PubMed] [Google Scholar]
  3. Aprile G., Rihawi K., De Carlo E., Sonis S.T. Treatment-related gastrointestinal toxicities and advanced colorectal or pancreatic cancer: a critical update. World J. Gastroenterol. 2015;21:11793–11803. doi: 10.3748/wjg.v21.i41.11793. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Bentz J., O’Connor M.P., Bednarczyk D., Coleman J.A., Lee C., Palm J., Pak Y.A., Perloff E.S., Reyner E., Balimane P. Variability in p-glycoprotein inhibitory potency (ic(50)) using various in vitro experimental systems: implications for universal digoxin drug-drug interaction risk assessment decision criteria. Drug Metab. Dispos. 2013;41:1347–1366. doi: 10.1124/dmd.112.050500. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bhatia S.N., Ingber D.E. Microfluidic organs-on-chips. Nat. Biotechnol. 2014;32:760–772. doi: 10.1038/nbt.2989. [DOI] [PubMed] [Google Scholar]
  6. Boelsterli U.A., Redinbo M.R., Saitta K.S. Multiple NSAID-induced hits injure the small intestine: underlying mechanisms and novel strategies. Toxicol. Sci. 2013;131:654–667. doi: 10.1093/toxsci/kfs310. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Chen Y., Zhou W., Roh T., Estes M.K., Kaplan D.L. In vitro enteroid-derived three-dimensional tissue model of human small intestinal epithelium with innate immune responses. PLoS One. 2017;12:e0187880. doi: 10.1371/journal.pone.0187880. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Chi C.W., Ahmed A.R., Dereli-Korkut Z., Wang S. Microfluidic cell chips for high-throughput drug screening. Bioanalysis. 2016;8:921–937. doi: 10.4155/bio-2016-0028. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Christophi G.P., Rong R., Holtzapple P.G., Massa P.T., Landas S.K. Immune markers and differential signaling networks in ulcerative colitis and Crohn's disease. Inflamm. Bowel Dis. 2012;18:2342–2356. doi: 10.1002/ibd.22957. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Cui W., Li L.X., Sun C.M., Wen Y., Zhou Y., Dong Y.L., Liu P. Tumor necrosis factor alpha increases epithelial barrier permeability by disrupting tight junctions in Caco-2 cells. Braz. J. Med. Biol. Res. 2010;43:330–337. doi: 10.1590/S0100-879X2010007500020. [DOI] [PubMed] [Google Scholar]
  11. Dantzig A.H., Shepard R.L., Cao J., Law K.L., Ehlhardt W.J., Baughman T.M., Bumol T.F., Starling J.J. Reversal of P-glycoprotein-mediated multidrug resistance by a potent cyclopropyldibenzosuberane modulator, LY335979. Cancer Res. 1996;56:4171–4179. [PubMed] [Google Scholar]
  12. Fatehullah A., Tan S.H., Barker N. Organoids as an in vitro model of human development and disease. Nat. Cell Biol. 2016;18:246–254. doi: 10.1038/ncb3312. [DOI] [PubMed] [Google Scholar]
  13. Jones C.R., Hatley O.J., Ungell A.L., Hilgendorf C., Peters S.A., Rostami-Hodjegan A. Gut wall metabolism. Application of pre-clinical models for the prediction of human drug absorption and first-pass elimination. AAPS J. 2016;18:589–604. doi: 10.1208/s12248-016-9889-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Kauffman A.L., Gyurdieva A.V., Mabus J.R., Ferguson C., Yan Z., Hornby P.J. Alternative functional in vitro models of human intestinal epithelia. Front Pharmacol. 2013;4:79. doi: 10.3389/fphar.2013.00079. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Kim H.J., Huh D., Hamilton G., Ingber D.E. Human gut-on-a-chip inhabited by microbial flora that experiences intestinal peristalsis-like motions and flow. Lab Chip. 2012;12:2165–2174. doi: 10.1039/c2lc40074j. [DOI] [PubMed] [Google Scholar]
  16. Kim R.B., Wandel C., Leake B., Cvetkovic M., Fromm M.F., Dempsey P.J., Roden M.M., Belas F., Chaudhary A.K., Roden D.M. Interrelationship between substrates and inhibitors of human CYP3A and P-glycoprotein. Pharm. Res. 1999;16:408–414. doi: 10.1023/a:1018877803319. [DOI] [PubMed] [Google Scholar]
  17. King S.M., Higgins J.W., Nino C.R., Smith T.R., Paffenroth E.H., Fairbairn C.E., Docuyanan A., Shah V.D., Chen A.E., Presnell S.C. 3D proximal tubule tissues recapitulate key aspects of renal physiology to enable nephrotoxicity testing. Front. Physiol. 2017;8:123. doi: 10.3389/fphys.2017.00123. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Li H., Jin H.E., Kim W., Han Y.H., Kim D.D., Chung S.J., Shim C.K. Involvement of P-glycoprotein, multidrug resistance protein 2 and breast cancer resistance protein in the transport of belotecan and topotecan in Caco-2 and MDCKII cells. Pharm. Res. 2008;25:2601–2612. doi: 10.1007/s11095-008-9678-0. [DOI] [PubMed] [Google Scholar]
  19. Li M., de Graaf I.A., Groothuis G.M. Precision-cut intestinal slices: alternative model for drug transport, metabolism, and toxicology research. Expert Opin. Drug Metab. Toxicol. 2016;12:175–190. doi: 10.1517/17425255.2016.1125882. [DOI] [PubMed] [Google Scholar]
  20. Musther H., Olivares-Morales A., Hatley O.J., Liu B., Rostami Hodjegan A. Animal versus human oral drug bioavailability: do they correlate? Eur. J. Pharm. Sci. 2014;57:280–291. doi: 10.1016/j.ejps.2013.08.018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Neurath M.F. New targets for mucosal healing and therapy in inflammatory bowel diseases. Mucosal Immunol. 2014;7:6–19. doi: 10.1038/mi.2013.73. [DOI] [PubMed] [Google Scholar]
  22. Nguyen D.G., Funk J., Robbins J.B., Crogan-Grundy C., Presnell S.C., Singer T., Roth A.B. Bioprinted 3D primary liver tissues allow assessment of organ-level response to clinical drug induced toxicity in vitro. PLoS One. 2016;11:e0158674. doi: 10.1371/journal.pone.0158674. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Paine M.F., Khalighi M., Fisher J.M., Shen D.D., Kunze K.L., Marsh C.L., Perkins J.D., Thummel K.E. Characterization of interintestinal and intraintestinal variations in human CYP3A-dependent metabolism. J. Pharmacol. Exp. Ther. 1997;283:1552–1562. [PubMed] [Google Scholar]
  24. Peters S.A., Jones C.R., Ungell A.L., Hatley O.J.D. Predicting drug extraction in the human gut wall: assessing contributions from drug metabolizing enzymes and transporter proteins using preclinical models. Clin. Pharmacokinet. 2016;55:673–696. doi: 10.1007/s40262-015-0351-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Peyrin-Biroulet L. Anti-TNF therapy in inflammatory bowel diseases: a huge review. Minerva Gastroenterol. Dietol. 2010;56:233–243. [PubMed] [Google Scholar]
  26. Prueksaritanont T., Gorham L.M., Hochman J.H., Tran L.O., Vyas K.P. Comparative studies of drug-metabolizing enzymes in dog, monkey, and human small intestines, and in Caco-2 cells. Drug Metab. Dispos. 1996;24:634–642. [PubMed] [Google Scholar]
  27. Sato T., Stange D.E., Ferrante M., Vries R.G., Van Es J.H., Van den Brink S., Van Houdt W.J., Pronk A., Van Gorp J., Siersema P.D. Long-term expansion of epithelial organoids from human colon, adenoma, adenocarcinoma, and Barrett's epithelium. Gastroenterology. 2011;141:1762–1772. doi: 10.1053/j.gastro.2011.07.050. [DOI] [PubMed] [Google Scholar]
  28. Schweinlin M., Wilhelm S., Schwedhelm I., Hansmann J., Rietscher R., Jurowich C., Walles H., Metzger M. Development of an advanced primary human in vitro model of the small intestine. Tissue Eng. Part C Methods. 2016;22:873–883. doi: 10.1089/ten.TEC.2016.0101. [DOI] [PubMed] [Google Scholar]
  29. Shi Y., Cheng D. Beyond triglyceride synthesis: the dynamic functional roles of MGAT and DGAT enzymes in energy metabolism. Am. J. Physiol. Endocrinol. Metab. 2009;297:E10–E18. doi: 10.1152/ajpendo.90949.2008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Sinagoga K.L., Wells J.M. Generating human intestinal tissues from pluripotent stem cells to study development and disease. EMBO J. 2015;34:1149–1163. doi: 10.15252/embj.201490686. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Spence J.R., Mayhew C.N., Rankin S.A., Kuhar M., Vallance J.E., Tolle K., Hoskins E.E., Kalinichenko V.V., Wells S.I., Zorn A.M. Directed differentiation of human pluripotent stem cells into intestinal tissue in vitro. Nature. 2011;470:105–109. doi: 10.1038/nature09691. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Srinivasan B., Kolli A.R., Esch M.B., Abaci H.E., Shuler M.L., Hickman J.J. TEER measurement techniques for in vitro barrier model systems. J. Lab. Autom. 2015;20:107–126. doi: 10.1177/2211068214561025. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Takenaka T., Harada N., Kuze J., Chiba M., Iwao T., Matsunaga T. Human small intestinal epithelial cells differentiated from adult intestinal stem cells as a novel system for predicting oral drug absorption in humans. Drug Metab. Dispos. 2014;42:1947–1954. doi: 10.1124/dmd.114.059493. [DOI] [PubMed] [Google Scholar]
  34. Takenaka T., Harada N., Kuze J., Chiba M., Iwao T., Matsunaga T. Application of a human intestinal epithelial cell monolayer to the prediction of oral drug absorption in humans as a superior alternative to the Caco-2 cell monolayer. J. Pharm. Sci. 2016;105:915–924. doi: 10.1016/j.xphs.2015.11.035. [DOI] [PubMed] [Google Scholar]
  35. Tang A.S., Chikhale P.J., Shah P.K., Borchardt R.T. Utilization of a human intestinal epithelial cell culture system (Caco-2) for evaluating cytoprotective agents. Pharm. Res. 1993;10:1620–1626. doi: 10.1023/a:1018976804403. [DOI] [PubMed] [Google Scholar]
  36. Tomisato W., Tsutsumi S., Rokutan K., Tsuchiya T., Mizushima T. NSAIDs induce both necrosis and apoptosis in Guinea pig gastric mucosal cells in primary culture. Am. J. Physiol. Gastrointest. Liver Physiol. 2001;281:G1092–G1100. doi: 10.1152/ajpgi.2001.281.4.G1092. [DOI] [PubMed] [Google Scholar]
  37. Treede I., Braun A., Jeliaskova P., Giese T., Fullekrug J., Griffiths G., Stremmel W., Ehehalt R. TNF-alpha-induced up-regulation of pro-inflammatory cytokines is reduced by phosphatidylcholine in intestinal epithelial cells. BMC Gastroenterol. 2009;9:53. doi: 10.1186/1471-230X-9-53. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. van de Kerkhof E.G., Ungell A.L., Sjoberg A.K., de Jager M.H., Hilgendorf C., de Graaf I.A., Groothuis G.M. Innovative methods to study human intestinal drug metabolism in vitro: precision-cut slices compared with ussing chamber preparations. Drug Metab. Dispos. 2006;34:1893–1902. doi: 10.1124/dmd.106.011148. [DOI] [PubMed] [Google Scholar]
  39. VanDussen K.L., Marinshaw J.M., Shaikh N., Miyoshi H., Moon C., Tarr P.I., Ciorba M.A., Stappenbeck T.S. Development of an enhanced human gastrointestinal epithelial culture system to facilitate patient-based assays. Gut. 2015;64:911–920. doi: 10.1136/gutjnl-2013-306651. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Wang X., Yamamoto Y., Wilson L.H., Zhang T., Howitt B.E., Farrow M.A., Kern F., Ning G., Hong Y., Khor C.C. Cloning and variation of ground state intestinal stem cells. Nature. 2015;522:173–178. doi: 10.1038/nature14484. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Watson C.L., Mahe M.M., Munera J., Howell J.C., Sundaram N., Poling H.M., Schweitzer J.I., Vallance J.E., Mayhew C.N., Sun Y. An in vivo model of human small intestine using pluripotent stem cells. Nat. Med. 2014;20:1310–1314. doi: 10.1038/nm.3737. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Yamaura Y., Chapron B.D., Wang Z., Himmelfarb J., Thummel K.E. Functional comparison of human colonic carcinoma cell lines and primary small intestinal epithelial cells for investigations of intestinal drug permeability and first-pass metabolism. Drug Metab. Dispos. 2016;44:329–335. doi: 10.1124/dmd.115.068429. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Yin X., Farin H.F., van Es J.H., Clevers H., Langer R., Karp J.M. Niche-independent high-purity cultures of Lgr5+ intestinal stem cells and their progeny. Nat. Methods. 2014;11:106–112. doi: 10.1038/nmeth.2737. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Document S1. Transparent Methods, Figures S1–S5, and Tables S1–S3
mmc1.pdf (1.5MB, pdf)

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