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Surrogate Wnt agonists that phenocopy canonical Wnt/β-catenin signaling
research-article
Author(s):
Claudia Y. Janda
1 ,
Luke T. Dang
2 ,
Changjiang You
3 ,
Junlei Chang
4 ,
Wim de Lau
5 ,
Zhendong A. Zhong
6 ,
Kelley S. Yan
4 ,
Owen Marecic
7 ,
Dirk Siepe
1 ,
Xingnan Li
4 ,
James D. Moody
2 ,
Bart O. Williams
6 ,
Hans Clevers
5 ,
Jacob Piehler
3 ,
David Baker
2 ,
Calvin J. Kuo
4 ,
K. Christopher Garcia
1
Publication date (Electronic):
03 May 2017
Journal:
Nature
There is no author summary for this article yet. Authors can add summaries to their articles on ScienceOpen to make them more accessible to a non-specialist audience.
Abstract
Wnts modulate cell proliferation, differentiation and stem cell self-renewal, by inducing β-catenin dependent signaling through Frizzled (Fzd) and Lrp5/6 to regulate cell fate decisions, and the growth and repair of a multitude of tissues 1 . The 19 mammalian Wnts interact promiscuously with the 10 Fzds, which has complicated the attribution of specific Fzd/Wnt subtype interactions to distinct biological functions. Furthermore, Wnts are post-translationally modified by palmitoylation, which is essential for Wnt secretion and functions as a critical site of interaction with Fzd 2– 4 . As a result of their acylation, Wnts are very hydrophobic proteins requiring detergents for purification, which presents major obstacles for the preparation and application of recombinant Wnts. This has hindered the delineation of the molecular mechanisms of Wnt signaling activation, understanding of the functional significance of Fzd subtypes, and the use of Wnts as therapeutics. Here we developed surrogate Wnt agonists, water-soluble Fzd-Lrp5/6 heterodimerizers, consisting of Fzd5/8-specific and broadly Fzd-reactive binding domains, that elicit a characteristic β-catenin signaling response in a Fzd-selective fashion, enhance osteogenic lineage commitment of primary mesenchymal stem cells (MSCs), and support the growth of a broad range of primary human organoid cultures comparably to Wnt3a. Furthermore, we demonstrate that the surrogates can be systemically expressed and exhibit Wnt activity in vivo, regulating metabolic liver zonation and promoting hepatocyte proliferation, resulting in hepatomegaly. These surrogates demonstrate that canonical Wnt signaling can be activated simply through bi-specific ligands that induce receptor heterodimerization. Furthermore, these easily produced non-lipidated Wnt surrogate agonists offer a new avenue to facilitate functional studies of Wnt signaling and the exploration of Wnt agonists for translational applications in regenerative medicine.
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Most cited references40
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- Abstract: found
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Long-Term Culture of Genome-Stable Bipotent Stem Cells from Adult Human Liver
Meritxell Huch, Helmuth Gehart, Ruben van Boxtel … (2015)
Introduction The liver is mainly composed of two epithelial cell types, hepatocytes and ductal cells. Hepatocytes synthesize essential serum proteins, control metabolism, and detoxify a wide variety of endogenous and exogenous molecules (Duncan et al., 2009). Despite their considerable replication capacity in vivo (Michalopoulos, 2014), hepatocytes have resisted long-term expansion in culture (Mitaka, 1998). Indeed, a recent study describes a human liver hepatocyte culture system for a period of ∼1 week with only 10-fold expansion (Shan et al., 2013). As an alternative, human embryonic stem (hES) cells and human induced pluripotent stem (hiPS) cells have been differentiated toward hepatocyte-like cells. However, recent reports imply that genetic and epigenetic aberrations occur during the derivation and reprogramming processes (Liang and Zhang, 2013; Pera, 2011; Lund et al., 2012). These range from chromosomal abnormalities (Laurent et al., 2011),“de novo” copy number variations (CNVs) (Hussein et al., 2011), and point mutations in protein-coding regions (Gore et al., 2011). Such changes may complicate their use for regenerative medicine purposes (Bayart and Cohen-Haguenauer, 2013). We have recently described a culture system that allows the long-term expansion (>1 year) of single mouse adult intestine (Sato et al., 2009), stomach (Barker et al., 2010), liver (Huch et al., 2013b), and pancreas (Huch et al., 2013a) stem cells. Lgr5, the receptor for the Wnt agonists R-spondins (Carmon et al., 2011; de Lau et al., 2011), marks adult stem cells in these mouse tissues (Barker et al., 2007, 2010; Huch et al., 2013a, 2013b). These cultures remain committed to their tissue of origin. We have recently adapted the technology to allow culturing of human intestinal stem cells (Jung et al., 2011; Sato et al., 2011) and have shown that patient-derived intestinal organoids recapitulate the pathology of hereditary intestinal diseases (Bigorgne et al., 2014; Dekkers et al., 2013; Wiegerinck et al., 2014). Here, we pursue the establishment of an organoid culture system for human liver. Results Optimization of Human Liver Stem Cell Culture Our defined mouse liver medium (ERFHNic [Huch et al., 2013b]) supported the growth of human liver cells only for 2–3 weeks (Figure 1A and 1B and Figure S1A, top, available online). Gene expression profiles of human liver cultures that were maintained for 2 weeks in “mouse liver medium” revealed highly active Tgf-β signaling. Tgf-β target genes such as CTGF, PLAT, TIMP1, and TIMP2 were highly expressed, whereas Tgf-β sequesters (LTBP2 and LTBP3) and Smad4 inhibitors (SMURF1 and SMURF2) (Massagué et al., 2005) were virtually absent (Figure S1B). Tgf-β signaling induces growth arrest and epithelial-to-mesenchymal transition (Xu et al., 2009). Specific inhibition of Tgf-β receptors Alk4/5/7 by the small molecule inhibitor A8301 downregulated CTGF, TIMP2, and PLAT (Figure S1C), extended the time in culture (∼6–7 weeks, six to seven splits) (Figure 1B), and enhanced colony-forming efficiency (Figure 1D). Still, the cultures eventually deteriorated (Figures 1B and 1C, left). Expression of the stem cell marker LGR5 decreased over time, whereas differentiation markers such as Albumin (ALB) or CYP3A4 were upregulated (data not shown), indicating that our conditions were promoting differentiation. We then tested additional compounds to induce proliferation and/or LGR5 expression (Table S1). Proliferating bile-duct progenitor cells occur both during homeostasis (Furuyama et al., 2011) and after damage (Dorrell et al., 2011; Huch et al., 2013b; Shin et al., 2011). As Forskolin (FSK), a cAMP pathway agonist, induces proliferation of biliary duct cells in vivo (Francis et al., 2004), we asked whether cAMP would support the human liver cultures. FSK addition upregulated LGR5 and the ductal marker KRT19, whereas ALB and CYP3A4 decreased (Figure S1D). Colony-forming efficiency was essentially unchanged (Figure 1D), yet the cultures expanded as budding organoids for many months in culture (>6 months) at a weekly split ratio of 1:4–1:6 (Figures 1B and 1C, right). Similar results were observed with other cAMP agonists (8-BrcAMP, Cholera toxin or NKH477) (Figure S1E). Removal of cAMP agonists resulted in rapid deterioration (Figures S1F and S1G). Similarly, removal of the Wnt agonist R-spo or blocking Wnt secretion by porcupine inhibition (IWP-2) resulted in rapid loss of the cultures (Figures S1F–S1H). This effect was rescued by exogenous addition of Wnt (Figure S1H). Twelve additional healthy human donor liver biopsies were cultured in the improved medium, with a consistent doubling time of ∼60 hr, independent of the age of the culture (Figures 1E and 1F and Table S2). EdU incorporation confirmed that the cells maintained their proliferative state in vitro (Figure 1G) for >3 months. Cultures could be readily frozen and thawed (data not shown). Thus, Wnt signals, cAMP activation, and Tgf-β inhibition were essential for long-term expansion. Organoids Originate from Ductal cells Collagenase perfusion of donor livers yields high numbers of fresh, viable, and functional human hepatocytes (Gramignoli et al., 2012) (Figure S2A). We employed EpCAM to differentially sort hepatocytes (EpCAM−) from ductal EpCAM+ ductal cells (Figures 1H, S2B, and S2C) (Schmelzer et al., 2007; Yoon et al., 2011). Although hepatocytes formed no organoids, EpCAM+ bile duct cells developed into organoids with a striking efficiency of 28.4% ± 3.2% (Figures 1H, S2D, and S2E). Crude liver cell preparations grew into organoid structures with an efficiency that equaled the number of EpCAM+ cells (Figures S2F and S2G). In our culture system, ductal cells rather than hepatocytes initiate organoids. Clonal Organoids Are Genetically Stable Organoids cultured for 3 months maintained normal chromosome numbers (Figures 3A and S4A). From two donors, we obtained biopsy samples, which we dissociated and cultured in bulk for 7 days. Subsequently, we isolated single cells and established two independent clonal lines for each of the two livers (cultures A and B). After 3 months of expanding these cultures, a second cloning step was performed. We could thus determine all genomic variation accumulated in a single cell during life, derivation, and 3 months of culture (Figures 2A and 2B). We observed 720–1,424 base substitutions per cultures, of which 63–139 were introduced during the 3 months culture (Figure 2C). Therefore, the majority of the base substitutions identified had been incorporated in vivo (during life) or introduced during organoid derivation, but not during culture. How do these numbers compare to published data? iPS cells contain 1,058–1,808 de novo base substitutions (determined at passage numbers between 15 and 25) compared to their parental somatic cells (Cheng et al., 2012). Of note, the numbers from these studies do not include the variation acquired in vivo in the parental somatic cells. Thus, 3 months of in vitro expansion of liver organoids introduces 10-fold fewer base substitutions than iPS cell reprogramming. Of the total number of base substitutions, only few were located in protein-coding DNA (seven to nine base substitutions per culture; Figures 2D and S3). With the exception of one synonymous mutation in culture A from donor 2 (Table S3), all mutations were already present in the early passage clonal cultures, indicating that they were not incorporated during the 3 months of expansion. None of the mutated genes occurs in COSMIC databases (Table S3). In iPS cells, an average of six base substitutions per line affect protein-coding DNA (Cheng et al., 2012; Gore et al., 2011). Next, we searched for structural aberrations in the WGS data. We did not observe any gross chromosomal aberrations (Figure 3B). We observed two copy number variants (CNVs), heterozygous gains, in one of the liver organoid cultures (Figures 3C). In the other cultures, we did not detect any CNV (Figures 3D and S4B–S4D). Moreover, these two CNVs were already present in the early passage cultures and therefore did not result from long-term culturing. ES cell cultures routinely show abnormal karyotypes (Baker et al., 2007), and iPS cells can harbor considerable numbers of somatic CNVs (Hussein et al., 2011; Laurent et al., 2011; Martins-Taylor et al., 2011; Mayshar et al., 2010; Abyzov et al., 2012). Differentiation into Functional Hepatocytes In Vitro and upon Transplantation The stem cell markers PROM1 and LGR5, as well as ductal (SOX9, OC2) and hepatocyte markers (HNF4a) were readily expressed (Figures 4A, S5A, and S5B). Histologically, liver organoids displayed a duct-like phenotype presenting either as: (1) a single-layered epithelium, expressing the cytokeratin markers KRT19 and KRT7, or (2) a pseudo-stratified epithelium with nonpolarized E-Cadherin+ HNF4a+ and some KRT7+ cells (Figures 4B–4D). SOX9 (Figure 4E) and EPHB2 (Figure 4F) were detectable in almost all cells, whereas LGR5 was detectable within the EPHB2+ population (Figure 4F). The organoids failed to express markers of mature hepatocytes, such as Albumin or CYP3A4 (Figures 4A and 5C, EM bars). Therefore, we defined a human differentiation medium (DM) (Table S1). Removal of the growth stimuli R-spo and FSK resulted in upregulation of Albumin and CYP3A4 (Figure S5C). To this medium, we then added the Notch inhibitor DAPT (Huch et al., 2013b), FGF19 (Wu et al., 2011), and dexamethasone (Rashid et al., 2010) (Figure S5D). BMP7 reportedly accelerates hepatocyte proliferation in vivo (Sugimoto et al., 2007). Addition of BMP7 slightly facilitated the expression of hepatocyte markers ALB and CYP3A4 even during expansion medium (data not shown). Therefore, 5–7 days prior to the start of differentiation, we added 25 ng/ml BMP7 to the expansion medium (EM) (Figure 5A). When cultured in this differentiation medium (DM), the cells acquired pronounced hepatocyte morphologies, including polygonal cell shapes (Figure 5B). Gene expression profiles revealed high levels of hepatocyte markers such as ALB, cytochromes, Apolipoproteins (APOB), and complement factors (C3) (Figures 5C, 5D, and S5E). Cells with high levels of ALB and MRP4 were detected by immunofluorescence (Figure 5B). Similar results were obtained with cultures derived from EpCAM+-sorted ductal cells (Figures S5F and S5G). Immunohistochemical analysis indicated that the cells accumulate glycogen (Figure 6A) and take up LDL (Figure 6B). Albumin was secreted into the medium (Figure 6C). The cultures exhibited similar CYP3A4 activity as fresh isolated hepatocytes (Figure 6D, compare to Figure S2A). Differentiated organoids hydroxylated midazolam, another indication of functional CYP3A3/4/5 activity (Wandel et al., 1994), and glucuronidated hydroxy-midazolam, thereby showing evidence of both phase I and II detoxifying reactions (Figure 6E). Bile acid salts were readily secreted into the medium (Figure 6F). Finally, the organoids detoxified ammonia at similar levels to HepaRG cells (Figure 6G). In all cases, the expanded human liver organoids showed stronger hepatocyte functions when compared to the standard/reference cell line HepG2 cells (Figure 6). To test the ability of the organoids to engraft as functional hepatocytes in vivo, we treated Balb/c nude mice with CCl4-retrorsine to induce acute liver damage. This treatment allows engraftment of hepatocytes (Guo et al., 2002; Schmelzer et al., 2007). Using human-specific antibodies (Figure S6A), we initially detected KRT19-positive, ductal-like cells at 2 hr and 2 days after transplantation, distributed throughout the liver parenchyma (Figure S6B). At later time points, we observed ALB + , KRT19 − human cells as singlets/doublets or, more rarely, in larger hepatocyte foci (Figures 6H and S6C). Of note, our damage model provides no stimulus for expansion of the transplant after engraftment. Human Albumin and α-1-antitrypsin were found in serum of recipient mice within 7–14 days (Figures 6I, S6D, and S6E) at a level that remained stable for more than 60 days in five out of six mice and for more than 120 days in two out of five animals. Although transplantation of primary human hepatocytes initially yielded higher levels of human Albumin (Figure 6I), the levels approximated those of transplanted organoids within a month. Patient Organoids Model Disease Pathogenesis α1-antitrypsin (A1AT) deficiency is an inherited disorder that predisposes to chronic obstructive pulmonary disease and chronic liver disease (Stoller and Aboussouan, 2005). A1AT is secreted from the liver to protect the lung against proteolytic damage from neutrophil elastase. The most frequent mutation is the Z allele (Glu342Lys) of the SERPINA1 gene, which causes accumulation of misfolded A1AT in hepatocytes. The ZZ mutant phenotype is characterized by a ∼80% reduction of the protein in plasma, which subsequently causes lung emphysema (Stoller and Aboussouan, 2005). Biopsies from three patients diagnosed with A1AT deficiency (Table S2 and Figure S7A) were subjected to histological characterization, RNA, and DNA isolation and expansion in culture. Organoids were grown for >4 months in culture and behaved normally. Gene expression analysis demonstrated that the cells differentiated normally in DM (Figure S7B). Functional tests revealed that the differentiated cells from A1AT patients secreted high levels of Albumin and take up LDL similar to that of healthy donor-derived organoid cultures (Figures 7B–7D). In A1AT deficiency, the molecular pathogenesis of the liver disease relates to the aggregation of the protein within the endoplasmic reticulum of hepatocytes (Lawless et al., 2008). A1AT protein aggregates were readily observed within the cells of the differentiated organoids (Figure 7H), similar to what was found in the original biopsy (Figure 7G). A1AT ELISA confirmed reduced protein secretion (Figure 7I) (Table S2 indicates the A1AT secretion per patient), and supernatants from differentiated mutant organoids showed reduced ability to block elastase activity (Figure 7J). Protein misfolding is one of the primary causes that drive hepatocytes apoptosis in PiZZ individuals (Lawless et al., 2008). Differentiated liver organoids from A1AT-D patients mimicked the in vivo situation and showed signs of ER stress, such as phosphorylation of eIF2α (Figure 7K) and increased apoptosis in the differentiated state (Figures S7C and S7D). Using a biopsy from an Alagille syndrome (AGS) patient, we tested whether structural defects of the biliary tree can also be modeled. AGS is caused by mutations in the Notch-signaling pathway, which results in partial to complete biliary atresia (Kamath et al., 2013). Patient organoids resembled their healthy counterparts in the undifferentiated state. However, upon differentiation to the biliary fate by withdrawal of R-spondin, Nicotinamide, TGFbi, and FSK, AGS patient organoids failed to upregulate biliary markers such as KRT19 and KRT7 (Figure S7E). Staining for KRT19 revealed that biliary cells were scarce and unable to integrate into the epithelium. Rather, they rounded up and underwent apoptosis in the organoid lumen (Figure S7F). In AGS mouse models, JAGGED-1/NOTCH2 is dispensable for biliary lineage specification but is required for biliary morphogenesis (Geisler et al., 2008; McCright et al., 2002). Thus, AGS liver organoids constitute the first human 3D model system to study Alagille syndrome. Discussion Liver diseases (ranging from genetic inherited disorders to viral hepatitis, liver cancer, and obesity-related fatty liver disease) account for the twelfth-leading cause of death in the United States (Heron, 2012). Failure in the management of liver diseases can be attributed to the shortage of donor livers (Vilarinho and Lifton, 2012) as well as to our poor understanding of the mechanisms behind liver pathology. The value of any cultured cell as a disease model or as a source for cell therapy transplantation depends on the fidelity and robustness of its expansion potential as well as its ability to maintain a normal genetic and epigenetic status (Pera, 2011). The possibility of differentiating hESC or reprogrammed fibroblasts (iPS) into almost any differentiated cell type, from neurons to hepatocytes, has allowed modeling of many human genetic diseases, including A1AT-D (Rashid et al., 2010). However, the genetic instability of cultured stem cells raises concerns regarding their safe use in cell therapy transplantation (Bayart and Cohen-Haguenauer, 2013). Here, we show that primary human bile duct cells can readily be expanded in vitro as bipotent stem cells into 3D organoids. These cells differentiate into functional hepatocyte cells in vitro and generate bona fide hepatocytes upon transplantation. Extensive analysis of the genetic stability of cultured organoids in vitro demonstrates that the expanded cells preserve their genetic integrity over months in culture. These results agree with our previous observations in the mouse (Huch et al., 2013b) yet are in striking contrast to recent publications in which, utilizing several lineage tracing approaches, ductal/resident stem cells have been described as not contributing to mouse liver regeneration (Schaub et al., 2014; Yanger et al., 2014; Yanger et al., 2013). Our results resemble what has been elegantly shown in zebrafish and rat models: in the event of an almost complete hepatocyte loss or blockage of hepatocyte proliferation, biliary epithelial cells convert into hepatocytes (Choi et al., 2014) (Michalopoulos, 2014). Our data are further corroborated in human fulminant hepatic failure, in which, upon 80% loss of hepatocyte compartment, huge numbers of proliferating EpCAM+ biliary epithelial cells are observed (Hattoum et al., 2013). Organoids from A1AT-deficiency patients can be expanded in vitro and mimic the in vivo pathology. Similarly, organoids from an Alagille syndrome patient reproduce the structural duct defects present in the biliary tree of these patients. Repair by homologous recombination using CRISPR/Cas9 technology is feasible in organoid cultures, as we recently demonstrated in colon stem cells of cystic fibrosis patients (Schwank et al., 2013). A variety of monogenic hereditary diseases affect the liver specifically, and these should all be amenable to a comparable in vitro approach of gene repair in clonal liver progenitor cells. Overall, our results open up the avenue to start testing human liver material expanded in vitro as an alternative cell source for studies of human liver regeneration, human liver disease mechanism, cell therapy transplantation, toxicology studies, or drug testing. Experimental Procedures Human Liver Organoid Culture Liver biopsies (0.5–1 cm3) were obtained from donor and explant livers during liver transplantation performed at the Erasmus MC, Rotterdam. The Medical Ethical Council of the Erasmus Medical Center approved the use of this material for research purposes, and informed consent was provided from all patients. For EpCAM sorting experiments and hepatocyte isolation, primary human liver tissue was obtained with informed consent and approval by the Regional Ethics Board, from the CLINTEC division of Karolinska institute (Dnr: 2010/678-31/3) (Jorns et al., 2014). Liver cells were isolated from human liver biopsies (0.5–1 cm3) by collagenase-accutase digestion, as described in the Extended Experimental Procedures. The different fractions were mixed and washed with cold Advanced DMEM/F12 and spun at 300–400 × g for 5 min. The cell pellet was mixed with Matrigel (BD Biosciences) or reduced growth factor BME 2 (Basement Membrane Extract, Type 2, Pathclear), and 3,000–10,000 cells were seeded per well in a 48-well/plate. Non-attaching plates were used (Greiner). After Matrigel or BME had solidified, culture medium was added. Culture media was based on AdDMEM/F12 (Invitrogen) supplemented with 1% N2 and 1% B27 (both from GIBCO), 1.25 mM N-Acetylcysteine (Sigma), 10 nM gastrin (Sigma), and the growth factors: 50 ng/ml EGF (Peprotech), 10% RSPO1 conditioned media (homemade), 100 ng/ml FGF10 (Peprotech), 25 ng/ml HGF (Peprotech), 10 mM Nicotinamide (Sigma), 5 uM A83.01 (Tocris), and 10 uM FSK (Tocris). For the establishment of the culture, the first 3 days after isolation, the medium was supplemented with 25 ng/ml Noggin (Peprotech), 30% Wnt CM (homemade prepared as described in Barker et al. [2010]), and 10 uM (Y27632, Sigma Aldrich) or hES cell cloning recovery solution (Stemgent). Then, the medium was changed into a medium without Noggin, Wnt, Y27632, hES cell cloning recovery solution. After 10–14 days, organoids were removed from the Matrigel or BME, mechanically dissociated into small fragments, and transferred to fresh matrix. Passage was performed in a 1:4–1:8 split ratio once every 7–10 days for at least 6 months. To prepare frozen stocks, organoid cultures were dissociated and mixed with recovery cell culture freezing medium (GIBCO) and frozen following standard procedures. When required, the cultures were thawed using standard thawing procedures and cultured as described above. For the first 3 days after thawing, the culture medium was supplemented with Y-27632 (10 μM). Growth curves and expansion ratios were performed and calculated as described in the Extended Experimental Procedures. Isolation of EpCAM+ Cells and Single-Cell Culture Cell suspensions prepared as described in the Extended Experimental Procedures were stained with anti-human CD326 (EpCAM), sorted on a MoFlo (Dako Cytomation) sorter, and cultured as described above with medium supplemented with Y-27632 (10 μM, Sigma Aldrich) for the first 4 days. Passage was performed in split ratios of 1:4–1:8 once per week. For clonogenic assays, single-cell suspensions were sorted using FSC and pulse width to discriminate single cells. Propidium iodide staining was used to label dead cells and FSC: pulse-width gating to exclude cell doublets (MoFlow, Dako). Sorted cells were embedded in Matrigel and seeded in 96-well plates at a ratio of 1 cell/well. Cells were cultured as described above. Hepatocyte Differentiation and In Vitro Functional Studies Liver organoids were seeded and kept 7–10 days under the liver medium explained above (EM, expansion medium) supplemented with BMP7 (25 ng/ml). Then, the cultures were split and seeded accordingly in this EM supplemented with BMP7 for at least 2–4 days. Then, medium was changed to the differentiation medium (DM): AdDMEM/F12 medium supplemented with 1% N2 and 1% B27 and containing EGF (50 ng/ml), gastrin (10 nM, Sigma), HGF (25 ng/ml), FGF19 (100 ng/ml), A8301 (500 nM), DAPT (10 uM), BMP7 (25 ng/ml), and dexamethasone (30 uM). Differentiation medium was changed every 2–3 for a period of 11–13 days. To assess hepatocyte function, culture medium was collected 24 hr after the last medium change. Functional studies were performed in the collected supernatant or in whole organoids, as described in the Extended Experimental Procedures. Transplantation We used a modified version of the protocol used by Guo et al. (Guo et al., 2002). In short, female BALB/c nude mice (around 7 weeks of age) were pretreated with two injections of 70 mg/kg Retrorsine (Sigma) at 30 and 14 days before transplantation. One day prior to transplantation, mice received 0.5 ml/kg CCl4 and 50 mg/animal anti-asialo GM1 (Wako Pure Chemical Industries) via IP injection. Furthermore, animals received 7.5 ug/ml FK506 in drinking water until the end of the experiment due to the reported positive effects on liver regeneration (He et al., 2010). On the day of transplantation, mice were anaesthetized, and suspensions of 1–2 × 106 human liver organoid cells derived from four independent donors (p6–p10) or fresh isolated hepatocytes (two donors) were injected intrasplenically. Transplanted mice received weekly injections of 50 mg/animal anti-asialo GM1 (Wako Pure Chemical Industries) to deplete NK cells. To monitor the transplantation state, blood samples were taken in regular intervals from the tail vein and were analyzed for the presence of human albumin and human α1-antitrypsin using respective human specific ELISAs (Assaypro). Karyotyping and Genetic Stability Analysis Organoid cultures in exponential growing phase were incubated for 16 hr with 0.05 μg/ml colcemid (GIBCO). Then, cultures were dissociated into single cells using TrypLE express (GIBCO) and processed using standard karyotyping protocols. DNA libraries for WGS analysis were generated from 1 μg of genomic DNA using standard protocols (Illumina). The libraries were sequenced with paired-end (2 × 100 bp) runs using Illumina HiSeq 2500 sequencers to a minimal depth of 30× base coverage (average depth of ∼36.9× base coverage). As a reference sample, liver biopsies was sequenced to equal depth for the different donors. Analysis of the sequence reads, calling of CNVs, and base substitutions are described in detail in the Extended Experimental Procedures. The data for the whole-genome sequencing were deposited to the EMBL European Nucleotide Archive with accession number ERP005929. Immunohistochemistry, Immunofluorescence, and Image Analysis Tissues and organoids were fixed o/n with formalin or 4% PFA, respectively, and stained and imaged as described in the Extended Experimental Procedures. A1AT-D Functional Experiments Elastase inhibition assay and detection of phosphorylated eIF2α were performed as described in the Extended Experimental Procedures. Microarray For the expression analysis of human liver cultures, total RNA was isolated from liver biopsies or from organoid cultures grown in our defined medium, using QIAGEN RNAase kit following the manufacturer’s instructions. Five hundred ng of total RNA were labeled with low RNA Input Linear Amp kit (Agilent Technologies). Universal human reference RNA (Agilent) was differentially labeled and hybridized to the tissue or cultured samples. A 4X 44 K Agilent whole human genome dual color microarray (G4122F) was used. Labeling, hybridization, and washing were performed according to Agilent guidelines. Microarray signal and background information were retrieved using Feature Extraction software (V.9.5.3, Agilent Technologies). Hierarchical clustering analysis was performed in whole-liver tissue or organoid arrays. A cut-off of 3-fold differentially expressed was used for the clustering analysis. Data Analysis All values are represented as mean ± SEM. Man-Whitney nonparametric test was used. p < 0.05 was considered statistically significant. In all cases, data from at least three independent experiments was used. All calculations were performed using SPSS package. Extended Experimental Procedures Human Liver Isolation Liver cells were isolated by collagenase digestion as follows: tissue (0.5-1cm3) was minced, rinsed 2x with DMEM (GIBCO) 1%FCS and incubated with the digestion solution (2.5 mg/ml collagenase D (Roche) + 0.1 mg/ml DNase I (Sigma) in EBSS (Hyclone, Thermoscientific), for 20-40 at 37°C. The digestion was stopped by adding cold DMEM 1%FCS and the suspension was then filtered through a 70 um Nylon cell strainer and spun 5 min at 300-400 g. The pellet was resuspended in DMEM 1%FCS and kept cold. Any material retained on the strainer was further digested for 10 min in Accutase (GIBCO) at 37C. Then, the digestion was stopped and the cells were collected as before. The different fractions (collagenase and accutase) were seeded and cultured as described in Experimental Procedures. In Vitro Growth Curves Expansion ratios were calculated from human liver cultures as follows: 3x103 cells were grown in our defined medium for 7 or 10 days. Then, the cultures were dissociated by incubation with TrypLE Express (GIBCO) until single cells. Cell numbers were counted by trypan blue exclusion at the indicated time points. From the basic formula of the exponential curve y(t) = y 0 x e (growth rate x t) (y = cell numbers at final time point; y 0 = cell numbers at initial time point; t = time) we derived the growth rate. Then, the doubling time was calculated as doubling time = ln(2)/growth rate for each time window analyzed. Isolation of EpCAM+ Cells from Primary Human Liver Human liver cells were isolated according to standard protocol at the liver cell laboratory at the unit for transplantation surgery, CLINTEC, Karolinska Institute (Jorns et al., 2014). Cell suspensions were shipped overnight on ice. Suspensions were diluted in 2 volumes of cold Advanced DMEM/F12 (GIBCO) and washed 3 times in the same medium. Viable cells were counted with Trypan blue and split into 3 parts for EpCAM sorting, Percoll purification (see Hepatocyte Percoll purification) and direct seeding into matrigel. For sorting, liver cells were stained with 1:100 Anti-Human CD326 (EpCAM) Alexa Fluor® 488 (eBioscience) for 30 min at 4°C. Subsequently cells were washed and sorted on a MoFlo (Dako Cytomation) cell sorter. Sorted cells were spun down, resuspended in Matrigel and grown into organoids according to standard human liver organoid culture procedure (see Human liver organoid culture). After 14 days in culture the number of organoids larger than 100 μm in diameter was scored. Hepatocyte Percoll Purification and Cyp3a4 Measurement Human hepatocyte suspensions (see Isolation of EpCAM+ cell from primary human liver) were washed as described, spun down and resuspended in 35 ml Advanced DMEM/F12 (GIBCO) + 13.5 ml Percoll (GE healthcare, density 1.130 g/ml) + 1.5 ml 10x HBSS (GIBCO). Cells were pelleted at 100 g for 10 min and washed 3 times in Advanced DMEM/F12 (GIBCO). Viable cells were counted with Trypan blue and 10.000 viable cells per 50 ul drop were seeded into matrigel. Remaining cells were stained for EpCAM as described above (see Isolation of EpCAM+ cells from primary human liver) or seeded onto collagen coated tissue culture plates for subsequent determination of cytochrome 3A4 activity. To measure Cyp3a4 in primary hepatocytes, the seeded cells were cultured in Williams E medium (GIBCO) containing Hepatocyte plating supplement pack (GIBCO) for 4 days with daily medium changes. On day 0 and day 4 the cells were incubated with Luciferin-PFBE substrate (50 μM) and Cytochrome P450 activity was measured using the P450-Glo Assay Kit (Promega) according to manufacturer’s instructions and normalized to the number of cells in the plate. HepG2 cells cultured in the same medium served as controls. Genetic Analysis DNA libraries for WGS analysis were generated from 1 μg of genomic DNA using standard protocols (Illumina). The libraries were sequenced with paired-end (2 × 100 bp) runs using Illumina HiSeq 2500 sequencers to a minimal depth of 30 x base coverage (average depth of ∼36.9 x base coverage). As reference sample, liver biopsies was sequenced to equal depth for the different donors. Sequence reads were mapped against human reference genome GRCh37 using Burrows-Wheeler Aligner (BWA) 0.7.5a with settings ‘bwa mem -c 100 -M’ resulting in sample-specific BAM files. To predict CNVs, BAM files were analyzed using Control-FREEC (Boeva et al., 2012) and DELLY (Rausch et al., 2012). To obtain somatically acquired CNVs, we filtered called CNVs for occurrence in the reference samples (liver biopsies). Single nucleotide variants (SNVs) were multi-sampled called using the Genome Analysis Toolkit (GATK) v2.7.2 UnifiedGenotyper (DePristo et al., 2011). We only considered positions at autosomal chromosomes, which were covered at least 20x in all liver stem cell samples and corresponding biopsy from the same donor. Candidate somatic SNVs were further filtered using the following criteria: no evidence in reference samples; minimal alternative allele frequency of 0.3 to exclude sequencing artifacts and potential substitutions that occurred after the clonal step; a minimal GATK quality score of 100; no overlap with single nucleotide polymorphisms (SNPs) in the Single Nucleotide Polymorphism Database (dbSNP 137.b37); and no overlap with SNVs in the other tested individual (Figure S2). Ultimately, SNVs with evidence in both clonal and subclonal cultures were considered as in vivo acquired somatic variation, and SNVs with evidence in only subclonal cultures were considered as variation accumulated during in vitro culturing. Immunohistochemistry, Immunofluorescence, and Image Analysis Tissues and organoids were fixed o/n with formalin or 4% PFA respectively, and stained washed and transferred to tissue cassettes and paraffin blocks using standard methods. Tissue sections (4 μM) were prepared and stained with antibodies, H&E or PAS using standard techniques. The antibodies and dilutions used are listed in Table S5. Stained tissues were counterstained with Mayer’s Hematoxylin. Pictures were taken with a Nikon E600 camera and a Leica DFDC500 microscope (Leica). For whole mount immunofluorescence staining, organoids were processed as described in Barker et al., (Barker et al., 2010). Nuclei were stained with Hoechst33342 (Molecular Probes). Immunofluorescence images were acquired using a confocal microscope (Leica, SP5). Images were analyzed and processed using Leica LAS AF Lite software (Leica SP5 confocal). All phase contrast pictures were acquired using a Leica DMIL microscope and a DFC420C camera. RT-PCR and qPCR Analysis RNA was extracted from organoid cultures or freshly isolated tissue using the RNeasy Mini RNA Extraction Kit (QIAGEN), and reverse-transcribed using reverse-transcribed using Moloney Murine Leukemia Virus reverse transcriptase (Promega). All targets were amplified (40 cycles) using gene-specific primers and MiIQ syber green (Bio-Rad). Data were analyzed using BioRad CFX manager. For Figure S7, cDNA was amplified in a thermal cycler (GeneAmp PCR System 9700; Applied Biosystems, London, UK) as previously described (Huch et al., 2009). Primers used are listed in Table S4. Functional Hepatocyte Studies To assess glycogen storage and LDL uptake, liver organoids grown in EM or DM for 11 days were stained by Periodic acid-Schiff (PAS, Sigma) and DiI-Ac-LDL (Biomedical Technologies), respectively, following manufacturer’s instructions. To determine albumin and A1AT secretion, liver organoids were differentiated as described. Culture medium was changed every 3-4 days and culture supernatant was collected was collected 24h after the last medium change. HepG2 (ATCC number 77400) and HEK293T (ATCC number CRL-3216) cells were cultured for 24h in the same medium without growth factors and were used as positive and negative control respectively. The amount of albumin and A1AT in culture supernatant was determined using a human specific Albumin or human specific A1AT ELISA kit (both from Assay Pro). To measure Cyp3a activity the cultures were differentiated as described and the day of the experiment the cells were removed from the matrigel and cultured with the Luciferin-PFBE substrate (50 μM) in Hepatozyme medium supplemented with 10% FBS (GIBCO). As controls, HepG2 and HEK293Tcells were cultured for 24h in DMEM 10%FBS and the day of the experiment transferred to Hepatozyme medium supplemented with 10% FBS (GIBCO) and Luciferin-PFBE substrate (50 μM). Cytochrome P450 activity was measured 8h later using the P450-Glo Assay Kit (Promega) according to manufacturer’s instructions. Concentrations of midazolam, 1-hydroxymidazolam (1-OH-M) and 1-hydroxymidazolam-glucuronide (1-OH-MG) were determined in 50 μl using LC-MS/MS. Analysis was carried out at the Clinical Pharmaceutical and Toxicological Laboratory of the Department of Clinical Pharmacy of the University Medical Center Utrecht, the Netherlands. All experiments were performed on a Thermo Fisher Scientific (Waltham, MA) triple quadrupole Quantum Access LC-MS/MS system with a Surveyor MS pump and a Surveyor Plus autosampler with an integrated column oven. Analytes were detected via MS/MS, with an electrospray ionization-interface in selected reaction monitoring-mode, by their parent and product ions. The method showed linearity over the range of 0.02 – 1.50 mg/L for MDZ and OHM and over the range of 0.10 – 10.0 mg/L for HMG. The analytical accuracy and precision were within the maximum tolerated bias and CV (20% for LLOQ, 15% for the other concentrations). Since a 1-OH-MG standard was not available, a Gold Standard was used. The Gold Standard consisted of the urine from two adult intensive care patients with a high dose of intravenous midazolam and good renal function. Total bile acids were measured on an AU5811 routine chemistry analyzer (Beckman Coulter, Brea, California) with an enzymatic colorimetric assay (Sentinel Diagnostics, Milano, Italy). Ammonia elimination was analyzed as follows: organoid cultures were expanded and differentiated in DM medium for 8 days. On day 8 CAG was added to the medium and 3 days later the organoids were remouved from the matrigel, washed with Williams’ medium and subsequently incubated with 1 ml of test medium (Williams’ E medium (Lonza, Basel, Switzerland) with 10% fetal bovine serum (Lonza), 5 μg / mL insulin (Sigma, St. Louis, U.S.), 50 μM hydrocortisone hemisuccinate (Sigma), 2mM glutamine (Lonza), 50 U / mL penicilline and 50 μg / mL streptomycin (penicilline/streptomycine mix (Lonza), 1.5 mM NH4Cl (Sigma), 2.27 mM D-galactose (Sigma), 2 mM L-lactate (Sigma) and 2 mM ornithine hydrochloride (Sigma)). Then 0.25 ml samples were taken after 45 min, 7 and 24 hr and stored at −20°C for further analysis. Subsequently, all cultures were washed twice with PBS, trypsinized and cell number was counted by tripan blue exclusion. Concentrations of ammonia were assessed in all samples by using the Ammonia (rapid) kit (Megazyme International, Wicklow, Ireland). The rates of ammonia elimination were established by calculating the changes in absolute molecular amounts of ammonia in the medium and corrected for time and cell number. SERPINA1 Sequencing All 4 SERPINA1 exons were amplified from genomic DNA using Phusion High-fidelity DNA polymerase (Thermo Scientific) and specific primersets (see Table S4). PCR products were purified using QIAquick PCR purification kit (QIAGEN) and sequenced on an ABI 3730XL capillary sequencer. Enzymatic Elastase Inhibition Assay For measurement of the inhibitory action of α1-antitrypsin in organoid supernatants, donor and patient organoids were differentiated for 11 days. Culture medium was changed every 2-3 days and culture supernatant was collected 24h after the last medium change. For the assay, 160 ul of supernatant are mixed with 20 ul of a 2 mg/ml N-Succinyl-Ala-Ala-Ala-p-nitroanilide (Sigma) 100 mM Tris pH 8.0 solution in a clear-bottom 96-well plate. After addition of 6x10-4 U of Elastase (porcine pancreas, Sigma) in 100 mM Tris pH 8.0, the increase in absorbance at 410 nm is measured continuously over 30 min. Elastase inhibition by supernatants is measured as the decreased inclination of absorbance over time in comparison to uninhibited controls (plain medium) and compared to a dilution series of purified human α1-antitrypsin (Zemaira) in medium. Detection of eIF2α Phosphorylation Donor and α1-antitrypsin deficient patient organoids were differentiated for 11 days. Culture medium was changed every 2-3 days and organoids were lysed in Lysis buffer (50 mM Tris pH 7.5, 50 mM NaCl, 0.5% Triton X-100, 0.5% NP40 substitute, 5 mM EGTA, 5 mM EDTA, 1x Complete protease inhibitor (Roche), 1x PhosStop (Roche)). Using standard techniques lysates were resolved by SDS-Page and blotted on PVDF membranes (Millipore). Antibodies against are listed in Table S5. Author Contributions M.H., H.G., and H.C. designed and, together with K.H., performed and analyzed experiments. M.H. designed and developed and, with K.H., performed all experiments and analyzed all data that characterized the human liver culture system. M.H., R.v.B., E.C., and H.C. designed the genetic studies. M.H. and H.G. designed and M.H., H.G., and K.H. performed A1AT experiments. H.G. and H.C. designed and H.G. and K.H. performed ductal origin, transplantation, and AGS experiments. R.v.B. performed the genetic stability studies, supervised the next-gen sequencing, and set up the filtering pipeline. F.B. adjusted and applied pipeline. J.d.L. performed the CNV analysis. M.H., M.v.W., R.H., S.A.F., S.J.B., and H.K. performed functional in vitro experiments and analyzed the data. M.v.d.W. and N.S. performed FACS. M.M.A.V., J.N.M.I., S.S., E.E. and L.J.W.v.d.L. provided Ethical Aproval, human liver donor biopsies, isolated hepatocytes, and patient material. E.E.S.N. and R.R.G.V. provided METC. R.R.G.V provided helpful discussions. M.H., H.G., R.v.B., E.C., and H.C. wrote the manuscript. All authors commented on the manuscript.
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A functional CFTR assay using primary cystic fibrosis intestinal organoids.
We recently established conditions allowing for long-term expansion of epithelial organoids from intestine, recapitulating essential features of the in vivo tissue architecture. Here we apply this technology to study primary intestinal organoids of people suffering from cystic fibrosis, a disease caused by mutations in CFTR, encoding cystic fibrosis transmembrane conductance regulator. Forskolin induces rapid swelling of organoids derived from healthy controls or wild-type mice, but this effect is strongly reduced in organoids of subjects with cystic fibrosis or in mice carrying the Cftr F508del mutation and is absent in Cftr-deficient organoids. This pattern is phenocopied by CFTR-specific inhibitors. Forskolin-induced swelling of in vitro-expanded human control and cystic fibrosis organoids corresponds quantitatively with forskolin-induced anion currents in freshly excised ex vivo rectal biopsies. Function of the CFTR F508del mutant protein is restored by incubation at low temperature, as well as by CFTR-restoring compounds. This relatively simple and robust assay will facilitate diagnosis, functional studies, drug development and personalized medicine approaches in cystic fibrosis.
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- Abstract: found
- Article: found
Unlimited in vitro expansion of adult bi-potent pancreas progenitors through the Lgr5/R-spondin axis
Meritxell Huch, Paola Bonfanti, Sylvia F Boj … (2013)
Introduction As first demonstrated for intestinal crypts (Korinek et al, 1998), Wnt signalling plays a crucial role in the regulation of multiple types of adult stem cells and progenitors (Clevers and Nusse, 2012). The Wnt target gene Lgr5 marks actively dividing stem cells in Wnt-driven, continuously self-renewing tissues such as small intestine and colon (Barker et al, 2007), stomach (Barker et al, 2010) and hair follicles (Jaks et al, 2008). However, expression of Lgr5 is not observed in endodermal organs with a low rate of spontaneous self-renewal, such as liver or pancreas. In the liver, we have recently described that Wnt signalling is highly activated during the regenerative response following liver damage. Lgr5 marks an injury-induced population of liver progenitor cells capable of regenerating the tissue after injury (Huch et al, 2013). In the adult pancreas, Wnt signalling is inactive (Pasca di Magliano et al, 2007), yet it is essential for its development during embryogenesis (Murtaugh et al, 2005; Heiser et al, 2006). The embryonic pancreas harbours multipotent progenitor cells that can give rise to all pancreatic lineages (acinar, duct and endocrine) (Zaret and Grompe, 2008). Injury to the pancreas can reactivate the formation of new pancreatic islets, called islet neogenesis, by mechanisms still not entirely understood but that resemble development of the embryonic pancreas (Bouwens, 1998; Gu et al, 2003). Lineage tracing studies have demonstrated that these ‘de novo beta cells' can be derived from pre-existing beta cells (Dor et al, 2004), or by conversion of alpha cells, after almost 90% beta-cell ablation (Thorel et al, 2010). Also, severe damage to the pancreas, by means of partial duct ligation (PDL) or acinar ablation, can stimulate non-endocrine precursors, such as duct cells, to proliferate and differentiate towards acinar (Criscimanna et al, 2011; Furuyama et al, 2011), duct (Criscimanna et al, 2011; Furuyama et al, 2011; Kopp et al, 2011) and also endocrine lineages (including beta cells) (Xu et al, 2008; Criscimanna et al, 2011; Pan et al, 2013; Van de Casteele et al, 2013), suggesting the existence of a pancreas progenitor pool within the ductal tree of the adult pancreas. The development of a primary culture system based on the adult, non-transformed progenitor pancreas cells would represent an essential step in the study of the relationships between pancreas progenitor cells, their descendants and the signals required to instruct them into a particular lineage fate. Also, the production of an unlimited supply of adult pancreas cells would facilitate the development of efficient cell replacement therapies. Most of the available pancreas adult stem cell-based culture protocols yield cell populations that undergo senescence over time unless the cells become transformed. It is fair to say that no robust, long-term culture system exists today that is capable of maintaining potent, clonal expansion of adult non-transformed pancreas progenitors over long periods of time under defined conditions. Recently, endoderm progenitors derived from embryonic stem cells (ESCs) (Cheng et al, 2012; Sneddon et al, 2012) or induced pluriportent stem cells (iPSCs) (Cheng et al, 2012) were serially expanded, in co-culture with pancreas mesenchyme or MEFs, respectively, and gave rise to glucose-responsive beta cells in vitro (Cheng et al, 2012) and glucose-sensing and insulin-secreting cells, when transplanted, in vivo (Sneddon et al, 2012). We have recently described a 3D culture system that allows long-term expansion of adult small intestine, stomach and liver cells without the need of a mesenchymal niche, while preserving the characteristics of the original adult epithelium (Sato et al, 2009; Barker et al, 2010; Huch et al, 2013). A crucial component of this culture medium is the Wnt agonist RSPO1 (Kim et al, 2005; Blaydon et al, 2006), the recently reported ligand of Lgr5 and its homologues (Carmon et al, 2011; de Lau et al, 2011). Here, we describe that Wnt signalling and Lgr5 are strongly upregulated in remodelling duct-like structures upon injury by PDL. We exploit the Wnt-Lgr5-Rspo signalling axis to generate culture conditions that allow long-term expansion of adult pancreatic duct cells, which maintain the ability to differentiate towards both duct and endocrine lineages when provided the proper signals. Results Wnt signalling and Lgr5 expression are upregulated during pancreas regeneration following PDL We first sought to document Wnt pathway activation in normal adult pancreas and following acute damage. We used the Axin2-LacZ allele as a general reporter for Wnt signalling (Leung et al, 2002; Lustig et al, 2002; Yu et al, 2005). In the head of a pancreas injured by PDL, where there is still healthy tissue, the reporter was inactive (Figure 1A), in agreement with the previous observations made with the TOPGAL Wnt reporter mice (DasGupta and Fuchs, 1999; Pasca di Magliano et al, 2007). However, after controlled injury by PDL (Watanabe et al, 1995; Xu et al, 2008), the Axin2 LacZ reporter was highly activated along the ductal tree of the ligated part of the pancreas (Figure 1B). Axin2 activation in the pancreas was already detectable at day 3 post injury, as assessed by qPCR (Figure 1C). Co-labelling with duct (pancytokeratin, CK) and endocrine (insulin, INS) markers revealed that the Axin2 upregulation was restricted to the duct compartment (Figure 1D). Thus, pancreas injury by PDL led to activation of Wnt target genes in the proliferative duct cell compartment (Scoggins et al, 2000) during the regenerative response. We have recently described that the Wnt target Lgr5 not only marks stem cells during physiological self-renewal (e.g., in the gut), but also marks a population of liver stem cells that is activated after liver damage (Huch et al, 2013). We utilized the Lgr5-LacZ knock-in allele (Barker, et al, 2007) to determine the expression of the Wnt target Lgr5 in the pancreas. Lgr5 is essentially undetectable in the head of a pancreas injured by PDL (non-ligated pancreas), in agreement with the absence of Wnt signalling in the tissue under homeostatic conditions (Figure 1E). However, in the tail of the pancreas upon PDL, we observed a significant Lgr5 LacZ reporter activity in the duct cells of the ligated pancreas, starting at day 3 and peaking at day 7 after PDL (Figure 1C and F). No background staining was detected in wild-type mice following pancreas injury (Supplementary Figure S1). The appearance of de novo expression of Lgr5 following pancreas regeneration by PDL suggested that pancreatic Lgr5 expression may herald de novo activation of regenerative stem/progenitor cells by Wnt upon injury. Pancreatic ducts self-renew in vitro Given the induction of Wnt and Lgr5 after injury, and the existence of pancreas progenitors in the ductal tree (Criscimanna et al, 2011; Furuyama et al, 2011), we reasoned that adult pancreas progenitors could be expanded from the duct cell compartment under our previously defined gut and stomach organoid culture conditions (Sato et al, 2009; Barker et al, 2010). Cultures of heterogeneous populations of pancreas cells have been previously established and typically include factors such as EGF, HGF and Nicotinamide (Bonner-Weir et al, 2000; Ramiya et al, 2000; Deutsch et al, 2001; Seaberg et al, 2004; Rovira et al, 2010; Cardinale et al, 2011; Smukler et al, 2011). Most of these approaches yield cell populations that undergo senescence over time unless the cells are transformed. To establish pancreas cultures, isolated pancreatic duct fragments from adult healthy mice (Figure 2A) were embedded in Matrigel containing the ‘generic' organoid culture factors EGF, RSPO1 and Noggin (Sato et al, 2009) to which FGF10 (Bhushan et al, 2001) and Nicotinamide were added. Under these conditions, small duct fragments formed closed structures within 24–48 h that expanded into budding cyst-like organoids (Figure 2B). The efficiency of cyst formation from isolated ducts and subsequent organoid formation was nearly 100%. Without EGF, RSPO1 or FGF10, the cultures deteriorated after 2–5 weeks (Supplementary Figure S2A). Noggin and Nicotinamide proved to be essential to maintain the cultures >2 months (∼passage 8) (Supplementary Figure S2A). The cultures maintained exponential growth with cell doubling times essentially unchanged during the culturing period (Figure 2C). Using these culture conditions, we have been able to expand the cultures by passaging at a 1:4–1:5 ratio weekly for over 10 months (Figure 2B). These culture conditions allowed the recovery of the cells after freezing and thawing. Of note, when transplanted into immunocompromised mice, the cultures did give rise only to ductal structures, and no tumour formation was detected in any of the mice analysed (n=5), confirming the non-transformed origin of the cultured cells (Supplementary Figure S2C and D). Also, the karyotype analysis revealed that chromosome numbers were essentially normal, even after >5 months in culture (Supplementary Figure S2B). Organoids generated from Axin2 LacZ and Lgr5 LacZ knock-in mice allowed localization of the Axin2- and Lgr5-positive cells. We observed XGAL staining in Axin2 LacZ pancreas organoids throughout the cysts, whereas XGAL staining in the Lgr5 LacZ -derived pancreas organoids was mainly restricted to small budding structures (Figure 2D). These results resembled the in vivo situation after pancreas injury by PDL, where only the ductal buds were Lgr5 +, whereas the Axin2 reporter showed a broader expression pattern (compare Figure 1B versus Figure 1F). Prospectively isolated single pancreatic duct cells but not endocrine or acinar cells self-renew long term in vitro We then prospectively isolated the different pancreatic epithelial cells (duct, acinar and endocrine lineages) and cultured the different populations in our defined 3D culture system. A prospective isolation procedure that allows isolation of single cells of the different pancreatic epithelial cell types and maintenance of their viability in culture has not been established yet. The epithelial cell-surface marker EpCAM and the high concentration of Zn2+ in secretory granules of endocrine cells, that allows binding of the fluorescent chelator TSQ (6-methoxy-8-p-toluenesulfonamido-quilone), were used as a basis for cell isolation. Pancreas tissue from both WT or transgenic mice that constitutively and ubiquitously express eGFP (Okabe et al, 1997) was dissociated into single cells. After depletion of non-epithelial (EpCAM−) and haematopoietic cells (CD45+, CD31+), the cell suspension was FACS sorted in order to separate the granulated endocrine fraction (EpCAM+TSQ+) from the non-endocrine component (EpCAM+TSQ-) with high purity (>99.6%) (Figure 3A–D; Supplementary Figure S3A and B). To rule out the possibility that endocrine cells might de-granulate during the isolation procedure and thus contaminate the non-endocrine fraction, we repeated the protocol on pancreas cells obtained from mouse insulin promoter (Mip)-RFP mice and found no RFP+ cells in the non-endocrine fraction (Supplementary Figure S4A). The separated fractions were then tested for their ability to survive, proliferate and give rise to organoids under the above-defined conditions. Only the EpCAM+TSQ− exocrine cells were able to generate duct-like structures that gave rise to larger organoids (1–1.5% organoid formation efficiency) and had to be split once a week (Figure 3E). As expected, the growth pattern of the single sorted cells followed an exponential curve (Supplementary Figure S3D). The duct-derived cell cultures were maintained for >5 months (Figure 3E). The EpCAM+TSQ+ endocrine cells did not proliferate, but survived for at least 30 days in culture (Figure 3F). Acino-ductal metaplasia can happen under conditions of stress or following injury (Means et al, 2005; Blaine et al, 2010). To confirm that duct rather than acinar cells are the long-term expanded cells isolated from the EpCAM+TSQ− fraction, we traced the progeny of isolated duct (Sox9 + ) or acinar (Ptf1a + ) cells in vitro. Transgenic mice with a Ptf1a CreER allele, that is exclusively expressed in the acinar compartment (Kopp et al, 2012; Pan et al, 2013), or mice carrying the Sox9 CreER allele, that is expressed predominantly (but is not absolutely restricted to) the duct cell compartment (Furuyama, et al, 2011; Kopp et al, 2012) were crossed with Rosa26R YFP mice and subcutaneously injected with tamoxifen as described in Supplementary Figure S5A. After the washout period, the pancreas was dissociated and single Sox9 YFP+ or Ptf1a YFP+ cells were FACS sorted and cultured in our defined pancreas culture medium (Supplementary Figure S5B–D). Only Sox9 YFP+ cells grew into budding organoids that expanded long term in culture, even when starting from a single cell (Supplementary Figure S5D, top panel). By contrast, the cultures derived from Ptf1a YFP+ cells gave rise to smaller duct-like structures that were able to proliferate only for 3–4 passages, after which they arrested proliferation (Supplementary Figure S5D, bottom panel). In conclusion, these data indicated that the long-term expanding pancreas organoid cultures derive from duct cells. Lgr5 cells sustain the growth of pancreas organoids that have a duct cell phenotype To test whether the Lgr5-expressing cells maintained the growth potential of the pancreas organoids, we sorted single Lgr5 LacZ+ cells from in vitro expanded organoids derived from Lgr5-LacZ knock-in mice (Barker et al, 2007). Indeed, the isolated Lgr5 + cells grew and formed organoids (Figure 4A–E) that were subsequently expanded for >4 months in culture by splitting the cultures weekly at a 1:6–1:8 ratio. The colony formation efficiency was ∼16%, similar to the colony formation of Lgr5 cells of small intestine and stomach (Barker et al, 2010; Sato et al, 2011) (Figure 4C). Of note, 1.6% of the Lgr5 neg -sorted population also grew into organoids (Figure 4C; Supplementary Figure S6A–D). These Lgr5 neg -derived clones rapidly re-expressed Lgr5 (Supplementary Figure S6B and C) and expanded at a similar ratio as their Lgr5 + counterparts (Supplementary Figure S6D). This result mirrors the efficiency of colony formation of the FACS-sorted EpCAM+TSQ− exocrine cells from healthy tissue (1.65%, Lgr5 neg versus 1–1.5%, exocrine cells). Overall, these results demonstrated that pancreas-derived Lgr5 + cells are capable of self-renewal and expansion in vitro, indicating that stem/progenitor cells can be activated both in organ-like structures and in secondary, single cell-derived organoids. Organoids derived from single, FACS-sorted, Sox9 + duct cells or from single isolated Lgr5 + cells (FACS sorted from Lgr5 LacZ cultures) allowed us to assess their lineage potential in vitro. Histologically, pancreas organoids displayed a duct-like phenotype characterized by a single-layered epithelium of cytokeratin-positive (CK) and MIC1-1C3-positive (Dorrell et al, 2008) cells (Figure 5A). Lgr5 + cells were readily detected in all organoids analysed (Figure 4E), similarly to what we had observed in the cultures derived from (non-clonal) duct fragments (Figure 2D). Ki67 and Edu staining demonstrated that only a subset of cells within the organoids proliferate (Figure 5A). Then, we performed comparative gene expression profiling of 1- to 2-month-old cultures and compared it with the gene expression profile of adult duct, acinar and islet cells. The overall gene expression profile of the organoid cultures clustered with the duct cell arrays, whereas it did not cluster with the gene profiles of acinar or endocrine cells (Figure 5B). Of note, among the genes whose expression pattern did cluster between the duct pancreatic cells and the organoids we found Sox9, Krt7, Krt19 and Spp1 (full list is provided in Supplementary Dataset 1). Comparison of the gene expression profile of the pancreas organoids and the pancreatic tissue (by in silico subtraction) confirmed the segregation of the non-ductal pancreatic markers (like Sst, Ins2, Gcg and Amy) and the ductal markers (like Krt7, Tcf2 and Sox9) (Figure 5C; Supplementary Dataset 2). Of note, the Wnt target genes Lgr5, Ccnd1 and Axin2 were also specifically highly expressed in the organoids (Figure 5C; Supplementary Dataset 2). As expected, gene set enrichment analysis (GSEA) confirmed that the organoid cultures are enriched in genes specifically expressed in adult Sox9+ pancreatic duct cells (Figure 5D and Supplementary Dataset 3). Interestingly, we also observed enrichment in genes previously reported in small intestinal and pancreas stem cells, that is, Lgr5, Prom1, Sox9 and Lrig1 (Figure 5D and Supplementary Dataset 4) (Barker et al, 2007; Snippert et al, 2009, Furuyama et al, 2011; Wong et al, 2012), while we found no significant enrichment in genes expressed in the developing pancreas at E14.5 or E17.5 (Figure 5E and Supplementary Datasets 5 and 6), confirming the adult nature of our pancreas progenitor cultures. To confirm this expression pattern, we performed qPCR analysis in cultures at early and late passages (Figure 5F). While some genes could be detected in pancreas organoids over time (Pdx1, Sox9 and Lgr5), no acinar (Amy2) or endocrine (Ins) markers were observed over several passages (Figure 5F). Immunofluorescent staining confirmed that the organoids were mainly formed by cells expressing Keratin19 (KRT19), SOX9, MUCIN-1 and PDX1 (Figure 5A) while negative for the endocrine marker Synaptophysin (SYP) (Supplementary Figure S4B). Overall, these results confirmed the pancreas progenitor and duct-like nature of the pancreas organoid cultures. Expanded organoids give rise to both pancreatic endocrine and duct cells in vivo The embryonic pancreas harbours all necessary factors and appropriate environmental cues to support the differentiation of bona fide pancreas progenitors to mature exocrine and endocrine cells in situ or when the embryonic pancreas is transplanted under the kidney capsule of an immunodeficient mouse (Zaret and Grompe, 2008). Therefore, to assess whether the organoid cells are capable of differentiating towards fully mature endocrine lineages (e.g., insulin producing cells) we developed a whole-organ morphogenetic assay based on the re-aggregation of dissociated cells from embryonic pancreas on one hand and organoids generated from adult pancreas on the other hand (Figure 6A; Supplementary Figure S7B). This type of morphogenetic assay has successfully been used to demonstrate fate potency of both skin and thymic epithelial stem cells after expansion in vitro (Bonfanti et al, 2010). When embryonic pancreas derived from either mouse (E13.5) or rat (E14) was isolated, dissociated, re-aggregated and then transplanted under the kidney capsule of an immune-deficient mouse, the embryonic tissue fully developed into the three mature pancreas lineages: duct, acinar and endocrine cells (Supplementary Figure S7A). Therefore, we isolated EpCAM+ TSQ− GFP+ epithelial cells from the pancreas of CAG eGFP adult mice (Okabe et al, 1997) as described above and expanded for at least 6 weeks (Supplementary Figure S3A–C), dissociated them into a single cell suspension and re-aggregated with embryonic E13 or E14 WT mouse or rat pancreas, respectively. The re-aggregates were kept overnight on a membrane and, the day after, were grafted under the kidney capsule of nude mice. After 2 or 3 weeks, mice were sacrificed and grafts harvested (Figure 6A; Supplementary Figure S7B). The transplanted re-aggregates did consistently grow pancreatic structures organized in both exocrine and endocrine areas, several of which contained eGFP+ integrated cells (Figure 6B and C; Supplementary Figure S7C and D). Immunohistochemical analysis of the re-aggregates revealed that eGFP+ cells mainly contributed to duct cells (Figure 6B and F). Of note, some eGFP+ cells, that located outside of the ducts, downregulated cytokeratin expression and contained high level of PDX1 protein, a feature of beta cells (Figure 6B). On the basis of their expression of synaptophysin, ∼5% of integrated eGFP+ cells were of endocrine nature, 50% of which were also insulin+ (Figure 6C and F). Quantification revealed that eGFP+ cells differentiated into duct cells at a frequency of 70% (Figure 6F; Supplementary Figure S7F). It is important to remark that these percentages roughly correspond to those found in the differentiating embryonic pancreas in vivo. More importantly, we obtained the same results using a different reporter mouse that expresses CFP under the control of E-Cadherin promoter (ECad CFP ) (Figure 6D and E; Supplementary Figures S7E and S8). We found that INS+ cells derived from cultivated organoids either from CAG eGFP or from ECad CFP reporter mice were functional and expressed C-peptide (Cppt) protein (Figure 6D and E; Supplementary Figure S7E). Cells with CFP membrane localization and cytoplasmic expression of both INS and mouse-specific C-peptide were readily detected throughout the grafted area, even when the organoid cells were engrafted in a rat pancreas microenvironment, where endogeneous INS+ cells were negative for the mouse/specific anti-Cppt antibody (Figure 6D and E; Supplementary Figure S8C). This last result excluded the possibility of fusion between mouse-cultivated eGFP+ or CFP+ cells and WT rat endocrine cells. The specificity of this antibody both in the ectopic rat pancreas and in the adult rat pancreas is shown in Figure 6D and Supplementary Figure S8B and C. Furthermore, the cultivated eGFP+ cells also gave rise to other endocrine lineages, such as Glucagon+ (GCG+) and Somatostatin+ (SST+) cells (Figure 7A and B). These cells were negative for INS, demonstrating that they had fully differentiated into mono-hormonal endocrine cells (Figure 7A and B). Then, we assessed whether a less permissive environment would also allow the adult expanded progenitor duct cells to achieve an endocrine cell fate. We directly transplanted ∼2-month-old adult duct pancreas cultures derived from both Bl6 WT mice and Ecad CFP mice into the kidney capsule of immunodeficient mice. We previously primed the 2-month-old cultures to express early endocrine markers by culturing them, 15 days prior to the transplantation, in a medium previously reported to allow ESC to acquire an endocrine fate (D'Amour et al, 2006; Kroon et al, 2008), with some modifications. We included the small molecule inhibitor ILV combined with FGF10, to induce Pdx1 expression (Bhushan et al, 2001; Chen et al, 2009), followed by DBZ treatment, to inhibit notch signalling (Milano et al, 2004) (Supplementary Figure S9A). This medium facilitated the expression of early endocrine progenitor markers (Neurogn3 and Chga) while retained the expression of the ductal marker Sox9, and suppressed Lgr5 (Supplementary Figure S9B). One month after transplantation, duct-like structures formed by Krt19+ cells were readily detectable throughout the graft (Supplementary Figure S9C). Also, albeit at much lower efficiency, Insulin+ and Cpeptide+ cells (Supplementary Figure S9C, D and E), as well as ChgA+ cells were detected (Supplementary Figure S9F). Overall, these results conclusively demonstrate that cultured organoids derived from either sorted adult duct cells (CAG eGFP or ECad CFP mice) or from freshly isolated ducts (Bl6 WT mice) are able to acquire both duct and endocrine fates, thus demonstrating their progenitor nature and bi-potency. Discussion The pancreas is a glandular organ that serves two important functions: the production of the digestive enzymes and the production of the hormones responsible of glucose homeostasis. This is mirrored in the wide range of pancreas diseases that vary from pancreatic cancer to disorders related to the glucose homeostasis, such as diabetes. While pancreas cancer is the result of the accumulation of oncogenic mutations in different epithelial cell types of the pancreas, diabetes is the result of severe reduction in functional beta-cell mass. The lack of primary culture systems capable of long-term expansion of primary tissue in vitro hampers the development of therapeutic strategies for pancreas diseases. The replacement of functional pancreatic beta cells may be envisioned as a potential definitive cure for diabetes. Unfortunately, human islet transplantation is hampered by the scarcity of donors and the need for immune suppression and also by graft failure (Lysy et al, 2012). Therefore, alternative sources for cell therapy replacement hold promise as a potential treatment for diabetes. ESCs and iPSCs can be differentiated towards beta cells in vitro (D'Amour et al, 2006; Zhang et al, 2009; Nostro et al, 2011; Cheng et al, 2012) and in vivo (Soria et al, 2000; Kroon et al, 2008; Sneddon et al, 2012), but the reproducibility of such procedures has been limited (Lysy et al, 2012). In addition, undifferentiated ESCs and iPSCs are prone to form teratomas upon transplantation in vivo, therefore any remaining undifferentiated cell must be completely removed prior to be used for transplantation. Adult pancreas progenitors able to expand long term in vitro while maintaining the potency to differentiate towards a duct or endocrine fate would potentially not encounter these limitations. We report here that damage of adult pancreas results in the upregulation of Wnt signalling and expression of the stem-cell marker Lgr5 in the neo-formed ducts. We exploit this Wnt-driven regenerative response to define a culture medium based on the Wnt activation (RSPO1) that allows the unlimited expansion of duct fragments or even single isolated cells in a defined medium without serum. Under these conditions, pancreatic duct cells upregulate the stem-cell marker Lgr5 (receptor for RSPO1), and self-renew while maintaining their genetic stability. Importantly, when the expanded adult progenitor cells receive the appropriate differentiation signals, as for instance the signals present in a developing embryonic pancreas, they are able to integrate into both exocrine and endocrine structures that express functional markers, demonstrating that they carry the hallmarks of bi-potent progenitors. Confirming the importance of the Wnt/Rspo signalling to facilitate the proliferation of pancreatic adult cells, Jin et al reported (while this study was under revision) that Rspo supplementation to a 3-week pancreatic culture facilitates the expansion of pancreas cells into heterogeneous cultures. The otherwise non-defined medium contains fetal bovine serum and ESC-derived conditioned medium (Jin et al, 2013). Thus, the conditions here described, based on the induction of the Wnt-Lgr5-Rspo axis, allow the long-term in vitro expansion of pancreas progenitors. The unlimited expansion potential of the adult progenitor cells may open avenues for building patient-derived disease models, as well as the development of regenerative strategies based on the expansion of adult, genetically non-modified, pancreas cells. Future optimization of the differentiation conditions may allow the generation of high numbers of specialized and functional pancreatic cells to be used for the treatment of pancreas diseases such as diabetes. Materials and methods Mice lines and injury models Generation and genotyping of the Lgr5 LacZ and ECad CFP mice is already described in Barker et al (2007) and Snippert et al (2010), respectively. Axin2 LacZ mice were obtained from EMMA (European Mouse Mutant Archive, Germany). C57BL/6-Tg(ACTB-EGFP)1Osb/J, Sox9 CreER and Ptf1a CreER mice were previously described (Okabe et al, 1997; Furuyama et al, 2011; Pan et al, 2013) and MipRFP mice were provided by Gérard Gradwohl (IGBMC, Strasbourg, France). Wild-type Sprague–Dawley (OFA) rats and OF1 mice were obtained from Janvier. Athymic (Swiss Nu−/−) were supplied by Charles River Breeding Laboratories NSG mice (Jackson Laboratory, Bar Harbor, MA, USA). All animal experiments were performed in accordance with the institutional review committee at the Hubrecht Institute and the VUB. Animals were maintained in a 12-h light cycle providing food and water ad libitum. To induce pancreas injury, 3- to 6-month-old mice were anaesthetized, with a mixture of fluanisone:fentanyl:midazolam injected intraperitoneally at a dosage of 3.3, 0.105 and 1.25 mg/kg, respectively. Following a median incision on the abdominal wall, the pancreas was exposed and, under a dissecting microscope, the pancreatic duct was ligated as described (Xu et al, 2008). For Sox9 and Ptf1a lineage labelling, tamoxifen (Sigma, T5648) was prepared at the concentration of 10 mg/ml in corn oil (Sigma, C8267). A total dose of 20 mg of tamoxifen was given subcutaneously in five doses of 4 mg over a 10-day period. A washout period of 14 days preceded pancreas harvesting and dissociation into single cells. Pancreas organoid cell culture Pancreatic ducts were isolated from the bulk of the pancreas of mice older than 8 weeks by collagenase dissociation (Collagenase type XI 0.012% (w/v) (Sigma), dispase 0.012% (w/v) (Gibco), FBS (Gibco) 1% in DMEM media (Gibco)) at 37°C. Isolated ducts were mixed with Matrigel (BD Bioscience) and seeded and cultured as we described previously (Sato et al, 2009; Barker et al, 2010). After Matrigel formed a gel, culture medium was added. Culture media was based on AdDMEM/F12 (Invitrogen) supplemented with B27 (Invitrogen), 1.25 mM N-Acetylcysteine (Sigma), 10 nM gastrin (Sigma) and the growth factors: 50 ng/ml EGF (Peprotech), 10% RSPO1-conditioned media (kindly provided by Calvin Kuo), 100 ng/ml Noggin (Peprotech) or 10% Noggin-conditioned media (in-house prepared), 100 ng/ml FGF10 (Peprotech) and 10 mM Nicotinamide (Sigma). One week after seeding, organoids were removed from the Matrigel, mechanically dissociated into small fragments, and transferred to fresh Matrigel. Passage was performed in a 1:4–1:8 split ratio once per week for at least 9 months. To prepare frozen stocks, organoid cultures were dissociated and mixed with Recovery cell culture freezing medium (Gibco) and froze following the standard procedures. When required, the cultures were thawed using standard thawing procedures, embedded in Matrigel and cultured as described above. For the first 3 days after thawing, the culture medium was supplemented with Y-27632 (10 μM, Sigma-Aldrich). Prospective isolation and pancreas organoid single cell (clonal) culture For clonogenic assays, whole pancreata were harvested from adult (8–12 weeks) mice and individually digested by collagenase type XI (0.3 mg/ml, Sigma) incubation at 37°C in a shaking incubator, and then dissociated into single cells by addition of trypsin (1 mg/ml, Sigma) and DNAse (0.4 mg/ml, Roche); cell suspension was filtered through a 70-μm cell strainer. Cell pellets were incubated with anti-mouse EpCAM/APC antibody (eBiosciences) for 30′ on ice. Cells were either processed directly for FACS sorting or were enriched for epithelial cells using magnetic beads (EasySepTM APC Positive selection kit or Epithelial enrichment kit; STEMCELL Technologies Inc.). Cells were re-suspended in a solution containing propidium iodide (PI, 1 mg/ml, Sigma), and N-(6-Methoxy-8-Quinolyl)-p-Toluenesulfonamide (TSQ, 1 mg/ml, Molecular Probes) and sorted on an FACSAria (Becton Dickinson). Clean separation between EpCAM+TSQ− and EpCAM+TSQ+ cell populations was confirmed by a second FACS analysis and immunocytochemistry. According to the mouse strain, an additional gate for eGFP or YFP signal was used for sorting cells. Pulse-width gating excluded cell doublets while dead cells were excluded by addition of PI and gating on the negative cells. For secondary clonal cultures, established cultures were dissociated into single cells and stained with the DetectaGene Green CMFDG LacZ Gene Expression Kit (Molecular Probes) according to the manufacturer's instructions. PI staining was used to label dead cells and FSC: pulse-width gating to exclude cell doublets. Sorted cells (EpCAM+TSQ−, EpCAM+TSQ+ or Lgr5 LacZ+ ) were embedded in Matrigel and seeded in 96-well plates at a ratio of 1 sorted cell/well. Cells were cultured in the pancreas media described above supplemented with Y-27632 (10 μM, Sigma-Aldrich) for the first 4 days. Passage was performed in split ratios of 1:4–1:5 once per week for at least 6 months. In vitro growth curves Expansion ratios were calculated from both sorted cells and duct fragments as follows: pancreas organoid cultures or 20 × 103 sorted cells were grown in our defined medium for 7 days. Then, the cultures were dissociated by incubation with TrypLE Express (Gibco) until single cells. Cell numbers were counted by trypan blue exclusion at the indicated time points. From the basic formula of the exponential curve y(t)=y 0 × e (growth rate × t) (y=cell numbers at final time point; y 0=cell numbers at initial time point; t=time) we derived the growth rate. Then, the doubling time was calculated as doubling time=ln(2)/growth rate for each time window analysed. Karyotyping Organoid cultures in exponential growing phase were incubated for 1–1.5 h with 0.05 μg/ml colcemid (Gibco). Then, cultures were dissociated into single cells using TrypLE express (Gibco) and processed as described (Huch et al, 2013). Chromosomes from 100 metaphase-arrested cells were counted. Pancreatic morphogenetic assay Pancreatic aggregates were obtained following a previously described protocol (Bonfanti et al, 2010), modified as follows. E13 mouse embryos (OF1) or E14 rat embryos (SD) were harvested from the uteri under sterile conditions, transferred in 100 mm Petri dishes containing HBSS supplemented with 10% FCS and stored on ice. Pancreatic tissue was removed from the embryonic abdomen and transferred into a solution containing collagenase type XI (1 mg/ml, Sigma) and DNAase (0.4 mg/ml, Roche Diagnostic) for about 5 min. A known number (from 75 × 103 to 105) of GFP-labelled single cells dissociated from in vitro expanded adult organoids were mixed with an ∼10-fold excess of unlabelled embryonic pancreatic cells. Aggregates were then transferred on a 0.8-μm Isopore membrane filter (Millipore) and incubated at 37°C for 24 h in RPMI medium supplemented with 10% FCS, before being grafted under the kidney capsule of nude mice as previously described (Bonfanti et al, 2010). Two to four weeks later, the grafts were harvested and processed for cryosection and immunohistochemistry. Pancreas organoid differentiation and kidney capsule transplantation Pancreas organoids derived from Bl6 isolated ducts or eGFP+- or CFP+-sorted cells were expanded in vitro for at least 2 months in our defined culture medium (EM) as described above. Then, the organoids were transferred into a differentiation medium (DM) to enhance their endocrine fate. To define the differentiation medium, we adapted the protocols already described by D'Amour et al (2006) and Chen et al (2009) as follows: organoids grown in Matrigel, in our defined expansion medium (EM), were removed from the Matrigel by using BD cell recovery solution (BD Biosciences), following the manufacturer's instructions, and transferred to suspension plates. The cells were maintained for 3 days in RPMI medium supplemented with 0.2% FBS and 100 ng/ml Activin A (Tocris BioScience). Then, the medium was changed to RPMI supplemented with 300 nM ILV (indolactam-V) (Tocris BioScience), 100 ng/ml FGF10 (Peprotech) and 2% FBS for 4–5 days. After, the medium was replaced by DMEM supplemented with 1% B27, Noggin (50 ng/ml), Retinoic Acid (2 μM) and KAAD-cyclopamine (0.25 μM) for the following 6 days. Finally, for the last 2–4 days prior to transplantation, the medium was changed to DMEM supplemented with 1% B27 and 10 μM DBZ (Tocris BioScience). During all the differentiation protocol, the cells were kept in suspension plates. After the last 2–4 days in DBZ supplemented medium, the organoids were collected and transplanted directly into the kidney capsule of nude mice using standard procedures. The grafts were allowed to grow for 1 month and then were harvested and processed for paraffin embedding and immunohistochemistry. To determine any potential transformation of the cells, pancreas organoids derived from Bl6 mice and cultured in our defined medium for at least 2 months were also directly transplanted into the kidney capsule of nude, SCID or NSG mice. The grafts were harvested 2 weeks and 3 months later and were processed for paraffin section and H&E staining using standard techniques. β-galactosidase (LacZ) staining, immunohistochemistry and immunoflorescence Tissues were fixed for 2 h in ice-cold fixative (1% Formaldehyde; 0.2% Glutaraldehyde; 0.02% NP-40 in PBS0) and incubated O/N at RT with 1–2 mg/ml of X-gal (bromo-chloro-indolyl-galactopyranoside) solution as we described in Barker et al (2010). The stained tissues were transferred to tissue cassettes and paraffin blocks were prepared using standard methods. Tissue sections (4 μM) were prepared and counterstained with neutral red. For immunohistochemistry, tissues and organoids were fixed using formalin 4%, and stained using standard histology techniques as described (Barker et al, 2010). The antibodies and dilutions used are listed in Supplementary Table SI. Stained tissues were counterstained with Mayer's Hematoxylin. Pictures were taken with a Nikon E600 camera and a Leica DFDC500 microscope (Leica). For whole-mount immunofluorescence staining, organoids were processed as described in Barker et al (2010). Tissue sections (4 μM) or cryosections from kidney capsule grafts were processed for immunofluorescent staining using standard procedures. For the paraffin-embedded kidney capsule grafts citrate retrieval was performed. Antibodies and dilutions are listed in Supplementary Table SI. Nuclei were stained with Hoechst33342 (Molecular Probes). Microarray For the expression analysis of pancreas cultures, total RNA was isolated from Sox9+ duct cells (isolated as described in Supplementary Figure S5), acinar and islets cells (prepared from whole pancreas after collagenase dissociation), whole adult pancreas and pancreas organoids cultured in our defined medium, using Qiagen RNAase kit following the manufacturer's instructions. Five hundred nanograms of total RNA were labelled with the low RNA Input Linear Amp kit (Agilent Technologies, Palo Alto, CA). Universal mouse Reference RNA (Agilent) was differentially labelled and hybridized to the tissue or cultured samples. A 4 × 44K Agilent Whole Mouse Genome dual colour Microarray (G4122F) was used. Labelling, hybridization and washing were performed according to Agilent guidelines. Microarray signal and background information were retrieved using the Feature Extraction software (V.9.5.3, Agilent Technologies). The hierarchical clustering analysis was performed in duct, acinar, islet and organoid arrays after in silico subtraction of the pancreas gene array. A cutoff of two-fold differentially expressed was used for the clustering analysis. GSEA was performed according to Subramanian et al (2005). The gene lists and gene sets used for the analysis are all provided in Supplementary Datasets 1–6. GEO accession number is GSE50103. RT-PCR and qPCR analysis RNA was extracted from cell cultures or freshly isolated tissue using the RNeasy Mini RNA Extraction Kit (Qiagen) or TRIzol (Invitrogen) respectively, and reverse transcribed using SuperScript II Reverse Transcriptase (Invitrogen). All targets were amplified (40 cycles) using gene-specific Taqman primers and probe sets (Applied Biosystems, London, UK). Data were analysed using the Sequence Detection Systems Software, Version 1.9.1 (Applied Biosystems). For Neurog3, cDNA was amplified in a thermal cycler (GeneAmp PCR System 9700; Applied Biosystems) as previously described (Huch et al, 2009). Primers used are listed in Supplementary Table SII. Image analysis Images of cultivated cells were acquired using either a Leica DMIL microscope and a DFC420C camera or a Nikon TE2000 inverted automated fluorescence microscope with motorized table and controlled by the NIS elements AR software. Immunofluorescence images were acquired using an upright Zeiss Axioplan2 fluorescence microscope with Hamamatsu C10600 ORKA-R2 camera or a confocal microscope (Leica, SP5) or a confocal microscope (Leica, SP8) or a confocal multiphoton Zeiss LSM710 NLO with the TiSa laser microscope. Images were analysed using the Leica LAS AF Lite software (Leica SP5 confocal) or Smartcapture 3 (version 3.0.8). Confocal images were processed using Improvision VolocityLE and Zeiss Zen softwares. Data analysis All values are represented as mean±standard error of the mean (s.e.m.). Mann–Whitney non-parametric test was used. P<0.05 was considered as statistically significant. In all cases, data from at least three independent experiments were used. All calculations were performed using the SPSS package. Supplementary Material Supplementary Information Supplementary Datasets Review Process File
Author and article information
Journal
Journal ID (nlm-journal-id): 0410462
Journal ID (pubmed-jr-id): 6011
Journal ID (nlm-ta): Nature
Journal ID (iso-abbrev): Nature
Title:
Nature
ISSN
(Print):
0028-0836
ISSN
(Electronic):
1476-4687
Publication date Nihms-submitted: 12
May
2017
Publication date
(Electronic):
03
May
2017
Publication date
(Print):
11
May
2017
Publication date PMC-release: 16
February
2018
Volume: 545
Issue: 7653
Pages: 234-237
Affiliations
[1
]Department of Molecular and Cellular Physiology, Howard Hughes Medical Institute,
and Department of Structural Biology, Stanford University School of Medicine, Stanford,
CA 94305, USA
[2
]Department of Biochemistry, Howard Hughes Medical Institute, and the Institute for
Protein Design, University of Washington, Seattle, WA 98195, USA
[3
]Division of Biophysics, Department of Biology, University of Osnabrück, 49076 Osnabrück,
Germany
[4
]Department of Medicine, Division of Hematology, Stanford University School of Medicine,
Stanford, CA, 94305, USA
[5
]Hubrecht Institute, Royal Netherlands Academy of Arts and Sciences, and University
Medical Center Utrecht, Uppsalalaan 8, 3584 CT, Utrecht, The Netherlands
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Manuscript ID: NIHMS863503DOI: 10.1038/nature22306
PMC ID: 5815871
PubMed ID: 28467818
SO-VID: 39db9af9-5aac-4879-9f8a-7fbf56c7e618
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