SUMMARY
Intestinal epithelial homeostasis is maintained by adult intestinal stem cells, which, alongside Paneth cells, appear after birth. It is unclear how neonatal intestinal epithelial development is regulated. We found that Lysine-specific demethylase 1A (Kdm1a/Lsd1) is required for the postnatal maturation of crypts, including Paneth cell differentiation. Lsd1-deficient epithelium retains a neonatal state into adulthood, which is beneficial for repair after irradiation injury. Lsd1-deficient crypts, devoid of Paneth cells, are still able to form and maintain organoids. Mechanistically, we find a spatiotemporal expression of LSD1 during crypt formation, and LSD1 represses genes that are normally expressed in fetal and neonatal epithelium. Surprisingly, we could not link the transcriptional control by LSD1 functionally to fetal- or neonatal specific H3K4me1 sites. Nevertheless, enzymatic inhibition of LSD1 also leads to a loss of Paneth cells and increase in LGR5-expressing cells in human organoids. In summary, we found an important regulator of neonatal intestinal development and identified a druggable target to reprogram intestinal epithelium towards a reparative state.
INTRODUCTION
The intestinal epithelium undergoes a dramatic change during the neonatal period. Crypt formation occurs after birth together with the appearance of Paneth cells (PCs) and the development of adult intestinal stem cells (ISCs). Adult ISCs rely on niche factors such as Wnt ligands. In vivo, mesenchymal cells are important sources of Wnt to support ISC maintenance (Degirmenci et al., 2018; Greicius et al., 2018; Shoshkes-Carmel et al., 2018), whereas in vitro, it is PCs that are required to supply the necessary Wnt (Durand et al., 2012; Farin et al., 2012; Kim et al., 2012; Sato et al., 2011). In contrast, Wnt ligands or PCs are dispensable for fetal organoid homeostasis (Fordham et al., 2013; Mustata et al., 2013). Thus, ISCs undergo a fetal-to-adult transition that includes a change in Wnt dependency.
Recently, the existence of bona fide fetal ISCs has been challenged by the finding that any fetal epithelial cell can be or become an adult ISC as long as the appropriate environment is supplied (Guiu et al., 2019). This model fits nicely with studies showing that after injury, the intestinal epithelium is temporarily reprogrammed into a fetal-like state that is needed for proper repair (Gregorieff et al., 2015; Nusse et al., 2018; Yui et al., 2018). This, in turn, complements work specifying that adult intestinal epithelial lineages can dedifferentiate to give rise to new ISCs to rebuild the epithelium after injury (Buczacki et al., 2013; Tetteh et al., 2016; van Es et al., 2012; S. Yu et al., 2018). In hindsight, these high levels of cell-fate reversion make sense because the intestine is a common site for chemical and mechanical challenges as well as the host for many putative pathogens. Nonetheless, it is not yet fully understood how the fetal-to-adult ISC transition, or its reversal upon injury, is mediated, and whether epithelial reprogramming can be targeted therapeutically.
Adult ISCs give rise to all intestinal epithelial subtypes. Unlike other stem cell systems, both ISCs and differentiating intestinal epithelial cells have a similar chromatin state at lineage defining genes, which allows for Notch-mediated lateral inhibition in ISC differentiation (Kim et al., 2014). However, the same group subsequently identified differences in open chromatin distinguishing ISCs from secretory precursors, which was reversed upon irradiation when these secretory precursors dedifferentiate into ISCs (Jadhav et al., 2017). Three groups separately identified a crucial role for the polycomb repressive complex 2 (PRC2) in maintaining crypt physiology (Chiacchiera et al., 2016; Jadhav et al., 2019; Koppens et al., 2016). Although some epigenetic modifiers, such as the PRC2 complex, are critical, others, such as DOT1L, play a minimal role in maintaining intestinal epithelial physiology (Ho et al., 2013). In summary, although it is clear there is an important role for certain epigenetic modifiers in intestinal epithelial biology, the role and importance of others remain undefined.
Here we exploit the availability of chemical probes targeting epigenetic modifiers (Scheer et al., 2018), and combine it with intestinal organoid cultures (Sato et al., 2009), to identify the demethylase LSD1 as a critical regulator of crypt maturation including PC differentiation. LSD1 is a demethylase of H3K4me1/2 that is known to ‘decommission’ enhancers during embryonic development (Agarwal et al., 2017; Whyte et al., 2012). We observe that LSD1 itself is tightly regulated and its levels are reduced during crypt formation, both in development and after injury, yet, our data suggests that LSD1 is not required for the decommissioning of H3K4me1 sites that occurs during intestinal epithelial development. Nevertheless, mice lacking Lsd1 in the intestinal epithelium have crypts resembling newly formed crypts that lack PCs and are filled with ISC-like cells.
RESULTS
Identification of LSD1 as a regulator of Paneth cell differentiation
The intestinal epithelium undergoes a dramatic change during the neonatal period including the appearance of Paneth cells (PCs) two weeks after birth. To study PC differentiation in organoids, we developed a differentiation protocol (adapted from (Yin et al., 2013)) using CHIR (GSK3 inhibitor) and DAPT (γ-secretase inhibitor) to activate Wnt and block Notch signalling, respectively (Fig. S1A). CHIR-DAPT treatment led to a robust enrichment of Lysozyme+ PCs by confocal microscopy, mRNA, and protein expression compared to standard EGF, Noggin, and R-spondin 1 (ENR) organoid growing conditions (Fig. 1A, S1B, S1C). Next, we tested chemical probes targeting 18 methyltransferases and demethylases and identified the LSD1 inhibitor GSK-LSD1 to consistently repress PC differentiation (Fig. 1B, 1C, S1D-G, Supplementary table 1). In support, GSK-LSD1 similarly affected PC differentiation in organoids grown in ENR conditions (Fig. 1D, S1H). Use of a different LSD1 inhibitor led to a similar near loss of PCs (Fig. S1I). Consistent with the irreversible binding nature of GSK-LSD1 to its target (Mohammad et al., 2015), we found that upon withdrawal of GSK-LSD1, PCs re-appeared after 2 organoid splitting events (Fig. 1E, S1J). To test if only a selection or all PC genes are downregulated upon LSD1 inhibition, we performed RNA-seq on organoids treated with GSK-LSD1 for 24 h, between day 1 and 2 after splitting, which we anticipate is right when PCs develop in organoids. We found a robust downregulation of all PC specific genes (gene set from (Haber et al., 2017)) (Fig. 1F). In contrast, we did not observe change in expression of genes recently identified to be important for breaking of symmetry (Serra et al., 2019)(Fig. 1G). This could explain why we still observe budding and crypt formation, even in the absence of new PC formation.
Inhibition of LSD1 renders Lgr5+ cells independent of niche-providing PCs in vitro
PCs are crucial for adult small intestinal organoid growth as they supply niche factors to retain a stem cell population, and normally, PC-devoid organoids only sustain growth upon Wnt supplementation (Durand et al., 2012; Farin et al., 2012; Kim et al., 2012; Sato et al., 2011). To test the role of LSD1 in intestinal stem cells, we used Lgr5-EGFP derived organoids and treated them with GSK-LSD1. We found that GSK-LSD1 treatment resulted in a 2-3 fold increase in percentage of Lgr5+ cells (Fig. 1H, 1I, S1K), and co-treatment of GSK-LSD1 with CHIR or valproic acid had an additive effect on the Lgr5+ population (Fig. S1K). Thus, GSK-LSD1 treatment expands the Lgr5+ population and renders these cells independent of PC-derived niche factors such as Wnt3 (Fig. S1L). To test if GSK-LSD1 is able to act like Wnt ligands we quantified clonal organoid formation from single sorted Lgr5+ cells. However, unlike Wnt3A, GSK-LSD1 is not able to increase organoid formation from single cells (Fig. 1J). In addition, in contrast to the rapid loss of PC differentiation, 24 h GSK-LSD1 treatment did not lead to an expansion of the Lgr5+ population (Fig. 1K). Thus, this feature does not precede PC loss. Nevertheless, using a recently described culture condition that allows PC differentiation in human organoids (Fujii et al., 2018), we found that GSK-LSD1 also blocks PC differentiation in human organoids while simultaneously expanding the LGR5+ population (Fig. 1L, 1M). Together, we identify that inhibition of LSD1 leads to a loss of PCs and an increase of Lgr5+ cells in organoids, and our data suggests this occurs separately from Wnt or Notch pathway activation.
Spatiotemporal expression of LSD1 during postnatal development
In adult mouse epithelium, we observed a clear enrichment of LSD1 expression in crypts compared to villi (Fig. S2A, S2B). However, we did observe LSD1-negative cells at the bottom of crypts, which is where ISCs and PCs reside. Therefore, we examined the expression of LSD1 using intestinal tissue and organoids derived from Lgr5-EGFP mice to mark ISCs (Barker et al., 2007). We found that LSD1 is markedly absent from PCs both in vivo and in vitro, whereas LSD1 was detected in nuclei of all other crypt cells including Lgr5+ ISCs (Fig. 2A, 2B). A major part of intestinal epithelial development occurs after birth, including crypt formation, PC differentiation, and appearance of adult Lgr5+ ISCs. We observed a change in the LSD1 expression pattern when we compared P7 and P21 developmental stages (Fig. 2C). Specifically, at P7 we see robust expression of LSD1 in villus regions with reduced LSD1 expression in the zone where newly forming crypts appear (Fig. 2C). In contrast, at P21 the LSD1 pattern resembles adulthood with an enrichment in the crypt region compared to the villi (Fig. 2C, S2A).
LSD1 is required for PCs but not goblet or enteroendocrine cells in vivo
To test the role of LSD1 in vivo, we crossed Lsd1f/f (Kerenyi et al., 2013) with Villin-Cre mice to delete Lsd1 in intestinal epithelial cells specifically (KO mice). Although these mice appear normal, we found that KO mice lack PCs throughout the small intestine (Fig. 2D, S2C). We did observe that KO mice had ‘escaper’ crypts still expressing LSD1, and LSD1+ crypts were positive for Lysozyme (Fig. S2D). Thus, these mice do not completely lack PCs. Currently, two genes are known to be absolutely required for PC differentiation in vivo; Sox9 and Atoh1 (Bastide et al., 2007; Yang et al., 2001). We did not observe differences in Sox9 expression (Fig. 2D), and, although we found fewer Atoh1+ cells in KO epithelium (Fig. 2D), reduction of Atoh1+ cells unlikely causes a complete lack of PCs. We reasoned that perhaps KO crypts are filled with PC precursors expressing Wnt3, however, similar to GSK-LSD1 treated organoids, Wnt3 was markedly reduced in crypts of KO mice except for Lsd1+ escaper crypts (Fig. 2D, S2E).
Next, we examined other intestinal secretory lineages and found a reduction of goblet cells, but equal numbers of enteroendocrine cells, comparing adult WT and KO littermates (Fig. 2E). When we examined fetal and postnatal intestines of WT and KO littermates, we found that the reduction in goblet cells emerges after developmental stage P7, similar to when PCs develop (Fig. 2F, S2F, S2G). These results suggest that LSD1 KO epithelium maintains neonatal characteristics into adulthood, including the absence of PCs and fewer goblet cells.
Mice lacking LSD1 sustain crypt-bottom ISCs independent of PCs, and KO organoids grow independent of endogenous Wnt
To test the role of LSD1 in ISCs in vivo, we used in situ hybridization (ISH) and antibody-based detection of the ISC marker Olfm4 in tissues of WT and KO mice. We found Olfm4+ cells completely filling the bottom of crypts in KO mice, compared to the standard PC/ISC pattern observed in WT crypts (Fig. 2G, S2H). In addition, all crypt-base cells in KO mice are Ki67+, suggesting that these Olfm4+ cells are proliferating (Fig. S2H). Atoh1-/- mice lack PCs, and Atoh1-/- crypts do not sustain organoid growth without Wnt supplementation (Durand et al., 2012). In contrast, Lsd1 WT and KO crypts were equally able to form organoids, even in the absence of PCs in KO organoids (Fig. 2H). This led us to hypothesize that KO organoids do not rely on endogenous Wnt. Indeed, blockage of Wnt signaling by the porcupine inhibitor IWP-2, showed that treated KO organoids sustained growth unlike WT organoids (Fig. 2I, 2J). Of note, IWP-2 distinctively reduced growth rate in KO organoids, which makes long-term expansion unfeasible, yet, after splitting there were still surviving KO organoids under continuous IWP-2 treatment, and, LSD1 inhibitor treatment greatly increased splitting efficiency (Fig. S2I, S2J). In contrast, both KO and WT organoids could not sustain growth in medium lacking R-spondin 1 (Fig. 2I, 2J).
LSD1 represses a fetal-like, reparative gene program that allows PC differentiation independent of YAP/TAZ
Next, we sought to find the mechanism by which LSD1 controls intestinal epithelial biology. We performed RNA-seq on FACS-sorted Epcam+ small intestinal crypt cells from WT and KO mice. We found 2564 up and 1522 down regulated genes (p-adj<0.1) in KO cells compared to WT cells (Fig. 3A). In support of our findings that there are Atoh1+ cells in KO crypts and the differential ability of Atoh1-KO and Lsd1-KO crypts to form organoids, we found no shift towards an Atoh1-/- gene signature in the KO transcriptional profile (Fig. S3A) (Kim et al., 2014). To verify our gene expression profile, we tested a PC-specific gene signature (Haber et al., 2017), and, expectedly, found that this is suppressed in KO crypts (Fig. 3B). However, an Lgr5-ISC signature gene set (Muñoz et al., 2012) was not enriched in our KO transcript profile (Fig. 3C). Over the years, there have been various different intestinal stem cell and stem-cell like populations described, either using genetic markers such as Bmi1, or by techniques such as label-retaining capacity and scRNA-seq (Ayyaz et al., 2019; Buczacki et al., 2013; van Es et al., 2012; Yan et al., 2017). To test if there was enrichment for a certain stem cell type, we analyzed expression of defining genes for each population (Fig. S3B), however, we did not find enrichment for any stem cell subtype.
Recently, two groups elegantly identified and characterized a cellular repair state in the intestinal epithelium (Nusse et al., 2018; Yui et al., 2018). Indeed, two repair gene signatures from these studies were enriched in the KO transcriptional profile (Fig. 3D). These reparative profiles are fetal-like, and, in support, we also find enrichment of two fetal gene sets in the KO transcriptional profile (Fig. 3E) (Mustata et al., 2013; Yui et al., 2018), with clear enrichment of defining markers specifically identified in these studies (Fig. 3F). Yui et al. revealed that the reparative state was mediated by YAP/TAZ (Yui et al., 2018). Indeed, a separate YAP gene signature was enriched in our KO transcriptional profile (Fig. 3G) (Gregorieff et al., 2015). To test if YAP/TAZ is required for LSD1-mediated PC differentiation, we treated organoids derived from mice lacking both Yap and the gene encoding for TAZ Wwtr1 (Vil-Cre;Yapf/f;Wwtr1f/f) with GSK-LSD1 and found that PC differentiation was equally impaired in WT and mutant organoids (Fig. S3C). However, we noted that approximately half of these organoids contained Yap and Wwtr1 based on qPCR (Fig. S3D). We thus tested a second model, deleting YAP/TAZ in an inducible manner (Vil- CreERT2; Yapf/f;Wwtr1f/f organoids). Indeed, tamoxifen treatment led to near undetectable levels of both Yap and Wwtr1 (Fig. S3E), impaired survival and led to an increase in PCs (Fig. 3H, 3I), in accordance with previous results (Azzolin et al., 2014; Gregorieff et al., 2015). Strikingly, GSK-LSD1 treatment completely abrogated PC differentiation independently of YAP/TAZ (Fig. 3H, 3I).
LSD1 represses GFI1 mediated genes
We observed a rapid and broad reduction of Paneth cell genes upon GSK-LSD1 treatment (Fig. 1F), however, we did not observe an increase of Lgr5+ cells within that same timeline (Fig. 1K). In support, we also do not observe enrichment of both fetal gene programs after 24h of GSK-LSD1 treatment (Fig. 3J), suggesting this requires sustained inhibition or deletion of LSD1. Nevertheless, a study recently reported that an LSD1-GFI1 complex was rapidly disturbed by LSD1 inhibitors (Maiques-Diaz et al., 2018). Notably, Gfi1-/- mice have a near loss of Paneth cells, reduced goblet cell numbers, but more enteroendocrine cells (Shroyer et al., 2005). When we examined genes known to be repressed by GFI1, we found increase of Neurog3 and Neurod1, but not that of mature enteroendocrine cell markers Chga and Chgb (Fig. 3K, 3L). This is in accordance with equal enteroendocrine cells in vivo in WT and KO mice (Fig. 2E, S2F). Together, this suggests that inhibition of LSD1 rapidly leads to de-repression of GFI1-targeted enteroendocrine-progenitor markers, that in turn block PC differentiation, but fail to become fully mature enteroendocrine cells. Additionally, a broad, yet undefined, lack of epithelial maturation would impair PC and enteroendocrine differentiation, which together leads to a complete loss of PCs and equal ChgA+ cells in adult intestinal epithelium of KO mice.
LSD1 controls H3K4me1 levels of genes associated with a fetal-like profile
LSD1 controls embryonic development by repressing enhancers to allow for embryonic stem cell differentiation (Agarwal et al., 2017; Whyte et al., 2012). We did not observe major differences in global H3K4me1/2 levels by immuno-histochemistry comparing escaper LSD1+ crypts in KO tissue (Fig. S4A). To assess if LSD1 mediates H3K4 demethylation and/or chromatin accessibility, we performed ChIP-seq for H3K4me1 and H3K4me2 as well as ATAC-seq comparing sorted WT and KO crypt cells. Analysis of ChIP-seq for H3K4me1 identified 2059 sites with associated altered methylation levels, of which the majority (1622) were enriched in KO crypts (Fig. 4A). ChIP-seq of H3K4me2 revealed a very similar pattern and the large majority of genes affected by LSD1 with gain of H3K4me2 were also significant in the analysis for H3K4me1 (Fig. S4B, S4C). In support, analysis of ATAC-seq of sites with differential H3K4me1 levels showed a modest increase in open chromatin (Fig. 4A). In addition, most of these peaks are not in close proximity to TSS sites (Fig. S4D). The top 300 genes associated with increased H3K4me1 levels in the KO, are overall enriched in the KO transcriptional profile (Fig. 4B). These include established regenerative markers such as Ly6a, Ly6e, and Anxa6 (Fig. S4E). Analysis of ATAC-seq data identified 864 sites to have more open chromatin of which 608 sites had increased ATAC levels in KO crypt cells and were associated with higher H3K4me1 levels (Fig. 4C). In addition, the majority of genes associated with higher ATAC peaks were upregulated in KO crypts (Fig. S4F). Furthermore, using a high stringency (adj. p<0.01), we combined our RNA-seq, ChIP-seq, and ATAC-seq data to find a decent overlap of the genes associated by ChIP, ATAC, or both (Fig. 4D). Because LSD1 demethylates H3K4me1, we assume by combining our H3K4me1 ChIP-seq and RNA-seq data, we establish a core list of 228 genes putatively directly affected by Lsd1 loss (Fig. 4F, Table S2). Importantly, 84% of the increased H3K4me1 peaks associated with these 228 genes are located outside the 2 kb surrounding the TSS. This suggests that LSD1 would drive enhancer-mediated regulation of these genes, which fits with a role generally associated with LSD1 (Agarwal et al., 2017; Whyte et al., 2012). In addition, the LSD1 core signature is enriched in a transcriptional profile comparing fetal with adult organoids (Fig. 4E (Yui et al., 2018)).
LSD1 controls genes separately from PRC2
Several groups have shown that EED, an essential component of the Polycomb Repressive Complex 2 (PRC2), is essential for maintaining adult ISCs and crypts, likely by repressing fetal and embryonic genes (Chiacchiera et al., 2016; Jadhav et al., 2019; Koppens et al., 2016). Comparing the LSD1 core with the EED core (genes up in EEDKO crypts and associated with H3K27me3 peaks (Koppens et al., 2016)) revealed strikingly little overlap between regulated genes (Fig. 4F). Further analysis confirmed that also the majority of putative LSD1-controlled H3K4me1 genes are not ‘co-repressed’ by the PRC2-mediated H3K27me3 (Fig. S4G). Thus, this suggests that both LSD1 and PRC2 control fetal-like genes but in an unrelated fashion.
Defining fetal and early life gene programs in developing intestinal epithelium
So far, we have found that LSD1 controls postnatal cell lineage differentiation (Fig. 1, 2), which correlates with a fetal-like or reparative gene program (Fig. 3), that in turn may be mediated by the H3K4me1 demethylation role of LSD1 and/or its co-repressive role with other regulators such as GFI1 (Fig. 3, 4). To distinguish fetal from postnatal intestinal epithelial development we performed RNA-seq on embryonic (E) day 18.5, as well as postnatal (P) days 7 and P21 (Fig. S5A, S5B). Figure 5A shows how established cell lineage markers behave during development. As expected, we observed a stepwise increase in ISC markers, an abrupt appearance of PC genes at P21, and goblet and enteroendocrine gene expression at E18.5 and P7 stages that only increases slightly at P21 (Fig. 5A). This indeed supports our hypothesis that KO mice have intestinal epithelium that is ‘stuck’ at a P7 stage, lacking PCs, with reduced goblet cell and immature enteroendocrine cells, but with crypts containing ISC-like cells. When we compared genes upregulated in KO crypts to different developmental stages we see overlap with both E18.5 and P7 stages compared to P21 (Fig. 5B), and, a very similar pattern is noted in EED KO crypts (Fig. 5C). In a separate test, we found enrichment by GSEA for our own fetal (E18vsP7) and neonatal (P7vsP21) gene sets as well as E18 and P7 unique genes (Fig. 5D, S5C). Together, this suggests that both EED and LSD1 control different gene sets throughout intestinal epithelial development in a similar fashion, which results in a striking phenotypical difference: EED KO epithelium returns to a fetal and even embryonic state and mice become moribund (Jadhav et al., 2019), whereas LSD1 KO mice retain an early-life postnatal stage and appear normal up to at least 1 year.
Developmental profiles suggest no determining role for LSD1 in H3K4me1 demethylation for fetal or postnatal specific sites
To support our transcriptomic analysis, we also performed H3K4me1 ChIP-seq on FACS sorted epithelium at different developmental stages. As an example, we find a robust stepwise increase of H3K4me1 levels near the ISC marker gene Olfm4, whereas another ISC marker Lgr5 has mostly unchanged H3K4me1 levels (Fig. 5E, S5D). When we systemically assess differential sites, we find that there is an abrupt loss of H3K4me1 at P21 from sites that are significantly up in E18.5 and P7 (Fig. 5F, 5G). In contrast, ChIP levels that are low in E18.5 or P7 display a stepwise increase that culminates at P21 (Fig. 5F, 5G). Notably, there are fewer peak changes from E18.5 vs. P7, compared to P7 vs. P21 or E18.5 vs. P21 (3700, 15522, 26942 significant sites respectively) (Fig. 5F, 5G, S5E). This suggest that the major H3K4me1 transition occurs after P7, which would fit very well with our hypothesis that LSD1 controls this transition specifically. However, when we take these sites and display the H3K4me1 ChIP-seq data of WT vs. KO crypts, we find a complete lack of regulation of sites lost at P21 (Fig. 5F, 5G, S5E), nor did we observe increased ATAC levels (data not shown).
This suggests that LSD1 is not involved at all in the observed reduction of H3K4me1 sites at P21. In contrast, we observe a modest increase of H3K4me1 levels in KO crypts of the sites that normally culminate at P21 (Fig. 5F, 5G, S5E). In support, if we do the reverse, by taking the significant KO sites from Fig. 4A and display the different developmental stages, we see a stepwise increase of those peaks in time (Fig. S5F). This surprising finding suggests that LSD1 broadly modulates H3K4me1 levels of peaks that appear during development, but that LSD1 is not involved in H3K4me1 sites that are lost during the different developmental stages in intestinal epithelium.
To test if LSD1 controls other transcriptional regulators, we selected these from the genes upregulated in KO crypts (Egolf et al., 2019; Lambert et al., 2018). We found 73 significantly upregulated transcriptional regulators (Padj<0.01), and many are specifically fetal and/or neonatally expressed, including Prdm1 (Blimp1) (Fig. 5H, S5G). Prdm1KO mice have previously been shown to have increased maturation of the intestinal epithelium, with PCs already present at P7 (Muncan et al., 2011). In addition, many other candidates such as Arid3a and several Hoxa family members could be involved in regulating postnatal intestinal epithelial development. Future studies will be directed to test whether any of these have a dominant role.
LSD1 expression is downregulated during repair
Based on our data we hypothesize a model in which LSD1 is actively controlled during postnatal epithelial development where first, de novo crypt formation is allowed in the absence of LSD1, and afterwards, epithelial maturation (including PC differentiation) requires LSD1. In mice, crypt formation occurs in the first week after birth during development, but also after damage such as upon radiation injury. Therefore, we studied LSD1 expression after whole body irradiation (10 gy) and found that LSD1 is much lower expressed in regenerating epithelium 3 days post irradiation, compared to naïve mice (Fig. 6A). Of note, this is near opposite of the Hippo-transducer YAP expression pattern (Fig. S6A). In support, we assessed Lsd1 expression in a recently described single cell RNA-seq experiment comparing crypt cells during homeostasis and during active repair (Ayyaz et al., 2019), and found a clear reduction in number of crypt cells expressing Lsd1 (Fig. 6B). In summary, Lsd1 is actively downregulated during crypt formation both in developing as well as repairing intestines.
Lsd1-deficient epithelium has superior reparative capacity
We found that genetic loss of Lsd1 in vivo or LSD1 inhibition in vitro leads to intestinal epithelium with neonatal and/or repairing features. This prompted us to test whether a ‘reparative’ state during homeostasis would be beneficial after injury. We irradiated WT and KO mice with 10 Gy and we did not observe pathological differences 6 days post treatment (Fig. 6C, S6B). However, we did find an increase in crypt length 3 days after injury, as measured by Ki67+ (Fig. 6D, 6E). We found no evidence of appearing PCs after injury in KO tissue (Fig. 6F). We did find that the Olfm4+ ISC zone similarly expanded as the Ki67+ crypts (Fig. 6D, 6F). We next irradiated mice with 16 Gy, when WT mice are unable to recover by day 6, in contrast, KO epithelium regained crypt-villus structures and had lower pathology scores compared to WT epithelium (Fig. 6H, 6I, S6C, S6D). Thus, KO epithelial tissue that have a pre-existing repairing profile enhances repair in vivo after radiation injury.
Finally, we performed a gene expression array of GSK-LSD1 treated human organoids, and found that the LSD1 core and a fetal signature were enriched in GSK-LSD1 treated human organoids (Fig. 6J).
In summary, we provide evidence that inhibition of LSD1 may be a viable target for the reprogramming of intestinal epithelium into a reparative state that is beneficial after injury, such as inflicted by radiation therapy.
DISCUSSION
In this study, we show that spatiotemporal regulation of LSD1 occurs in the developing intestine and during repair, and that LSD1 is important for crypt maturation, which includes the appearance of PCs. Notably, while LSD1 KO mice fail to develop any PCs, organoids derived from KO epithelium are formed and normally maintained, even in the absence of Wnt. Although these resemble organoids derived from fetal epithelium that also lack PCs (Fordham et al., 2013; Mustata et al., 2013), there are some key distinctions. Unlike fetal organoids, KO organoids cannot grow without R-spondin 1 (Fig. 3F). In addition, KO organoids form crypts, likely because Notch-ligand expressing cells required for ‘symmetry breaking’ are still present (Serra et al., 2019), such as enteroendocrine progenitors (van Es et al., 2019). In contrast, fetal organoids remain spheroid, and only form crypts after a ‘maturation’ trigger, at which point they also lose the ability to grow without R-spondin 1. Our data suggests that KO organoids would be in between fetal and adult, having features of both; not relying on Wnt yet unable to grow without R-spondin 1.
Intestinal epithelial repair is crucial to prevent chronic disease, and YAP/TAZ are established initiators and critical regulators of this process (Cai et al., 2010; Gregorieff et al., 2015; Yui et al., 2018). YAP/TAZ are sensors of mechanical signals (Dupont et al., 2011), so it is plausible that upon damage, and subsequent ECM cues, YAP/TAZ are activated by control of their protein levels (Cai et al., 2010; Yui et al., 2018). We found a clear reduction of LSD1 levels after irradiation, which coincides with an increase in YAP levels. Although it is unclear how LSD1 levels are controlled after regeneration, in glioblastoma tumor cells GSK3β stabilizes LSD1 by direct phosphorylation (Zhou et al., 2016). Thus, activation of the Wnt pathway, which occurs during the regenerating phase, could lead to fast reduction of LSD1 levels by Wnt-controlled inhibition of GSK3. We further found that LSD1 represses a reparative gene expression profile, similar to that activated by YAP. However, GSK-LSD1-mediated depletion of PCs was completely independent of YAP/TAZ (Fig. 3H, 3I). In combination, after damage, reducing levels of the repressor LSD1, and inducing levels of activators YAP/TAZ, can together mediate a robust reparative response.
Maintenance of chromatin accessibility is required for intestinal physiology, for example by DNA methylation (D.-H. Yu et al., 2015). In addition, three groups have independently identified a crucial role for the PRC2 complex in intestinal epithelial biology (Chiacchiera et al., 2016; Jadhav et al., 2019; 2016; Koppens et al., 2016). In 2019, Jadhav et al. proposes a model in which, upon deletion of PRC2 member EED, first fetal and subsequent embryonic genes are re-activated in adult epithelium (Jadhav et al., 2019). This initially leads to loss of ISCs and aberrant differentiation, and ultimately leads to loss of structure and moribund mice 14 days after deletion of EED (Jadhav et al., 2019). When we compared EED core genes (genes up in EED KO crypts combined with H3K27me3 associated genes) with LSD1 core genes (up in LSD1 KO, and increase H3K4me1 levels in LSD1 KO) we found very little overlap, suggesting that there are not many genes co-repressed by PRC2 and LSD1 complexes, which would explain the difference in severity of their KO phenotypes. Surprisingly, LSD1 does not control H3K4me1 sites that are specific for E18 or P7 developmental stages (Fig. 5F, 5G). This is in stark contrast to PRC2 controlled sites that are primarily fetal and embryonic (Jadhav et al., 2019). Nonetheless, putative LSD1-controlled genes include those associated with altered H3K4me1 levels in KO crypts, as well as genes that may be regulated independently of LSD1 demethylase activity, such as those being co-repressed by GFI1 (Fig. 3K, 3L) (Maiques-Diaz et al., 2018). In addition, we present a list of transcriptional regulators with increased expression in LSD1 KO crypts, including previously identified regulator of intestinal epithelial development PRDM1 (BLIMP-1) (Muncan et al., 2011). Together, we conclude that LSD1 controls fetal and neonatal genes but in a rapid reversible fashion where cells retain their ability to mature, which happens either between P7-P21 or after injury, making LSD1 an attractive targeting candidate for stimulating intestinal repair in patients. Future studies will be directed towards identifying the specific regulatory mechanisms driving the repression of fetal and neonatal genes in adult intestinal epithelium by LSD1.
Author contributions
R.T.Z., H.T.L., M.F., M.T.P., Y.O., A.D.S, M.M.A., J.O., M.M., N.P., E.K., R.R.S., and M.J.O. performed experiments. K.N. and C.A. provided essential reagents. M.R., F.D., C.A., J.A.D., K.B.J., T.S., and M.J.O. supervised and/or provided conceptual insight. M.J.O wrote a first draft with subsequent input from all authors.
Competing interests
T.S. is an inventor on several patents related to organoid culture. Correspondence and request for materials should be addressed to M.J.O.
METHODS
Animal work
Lgr5-EGFP-IRES-CreERT2 (stock no: 008875) mouse strain was obtained from Jackson Laboratories. Villin-Cre and Villin-CreERT2 (Marjou et al., 2004) were a kind gift from Sylvie Robine, Lsd1f/f mice were a kind gift from Dr. Stuart Orkin (Kerenyi et al., 2013). Mice were housed and maintained at the Comparative Medicine Core Facility (CoMed), and experiments were ethically approved by the Norwegian Food Safety Authority (FOTS ID: 11842). Mice were lethally irradiated (10 Gy or 16 Gy) and small intestinal repair was assessed 3 and 6 days post irradiation. Yapf/f;Wwtr1f/f animals (Azzolin et al., 2014) were a kind gift from Stefano Piccolo and were crossed with Villin-Cre and Villin-CreERT2 at University of Copenhagen under the approval of the National Animal Ethics Committee in Denmark.
Crypt, IEC, and ISC isolation, mouse organoid cultures
Adult crypt isolation
Crypt, IEC, and ISC isolation, as well as organoid culture, were essentially done as described (Sato and Clevers, 2013). For adult crypt isolation duodenum tissue was rinsed with ice-cold PBS, cut open longitudinally, and villi were scraped off. Tissue was cut in ∼2 mm pieces and washed 5 times with ice-cold PBS. Tissue pieces were incubated in 2 mM EDTA in ice-cold PBS for 30-60 minutes. Crypts were isolated from up to 10 fractions after pipetting up and down 5 times with PBS. To isolate single cells for sorting, crypts were incubated in TrypLE (ThermoFisher) at 37 °C for 20-45 minutes. Single cells were stained and sorted, DAPI-negative and Epcam-positive cells were used for RNA-seq and ChIP-seq experiments. ISCs for clonal organoid outgrowth experiments were isolated by sorting DAPI-negative, GFP-high (top 5%) cells from Lgr5-EGFP mice.
E18.5/P7/P21 IEC isolation
Whole (E18.5) or proximal 10 cm (P7 and P21) small intestines were isolated and flushed with ice cold PBS when possible (P21). Small intestines were opened longitudinally (P21 and P7) and cut into small pieces that were washed with ice-cold PBS, and incubated with 2 mM EDTA in ice-cold PBS for 30 min. Whole epithelium was isolated by collecting all fractions, which was used directly for RNA isolation, for ChIP fractions were made single cell using TrypLE (ThermoFisher) and sorted as described.
Organoid cultures
Organoids were grown and maintained in ‘basal crypt medium’ (Advanced DMEM/F12 medium supplemented with penicillin/streptomycin, 10 mM HEPES, 2 mM Glutamax, N2 (Thermo Fisher, 17502048), B-27 (Thermo Fisher, 17504044)) supplemented with N-acetyl-L-cysteine (Sigma, A7250), 50 ng/ml murine EGF [ThermoFisher, PMG8041], 20% R-spondin 1s conditioned medium (CM) (kind gift from Dr. Calvin Kuo), and 10% Noggin-CM (kind gift from Dr. Hans Clevers). For ISC clonal experiments, in the first 48 h after seeding, the medium was supplemented with Rock inhibitor (Y-27632) and Jagged-1 peptide (amino-acid sequence CDDYYYGFGCNKFCRPR, made in house, peptide synthesized as described (Bolscher et al., 2011)), 33% Wnt3-CM (kind gift from Dr. Hans Clevers) served as control. Medium was renewed every other day. For passaging, organoids cultures were washed, and matrigel and organoids were disrupted mechanically by strong pipetting, centrifuged at 200g, 5 min at 4 °C and resuspended in Matrigel to re-plate. Imaging of live organoids was done using an EVOS FL Auto 2. Structural Genomics Consortium supplied the inhibitors for the screen (www.thesgc.org), all of which are commercially available, concentrations are listed in Supplementary Table 1. Additionally, CHIR99021 (3 µM), IWP-2 (2 µM), VPA (1 mM) and DAPT (10 µM) were used.
Vil-Cre; Yapf/f/Wwtr1f/f and Vil-CreERT2; Yapf/f;Wwtr1f/f organoids were cultured in Basal Medium supplemented with NAC, B27, 50ng/ml human EGF (Peprotech, AF-100-15) and 100 ng/ml murine Noggin (Peprotech, 250-38) and either 500 ng/ml mouse RSPO1 (R&D systems, 3474-RS) or 10% RSPO1-conditioned medium. Established Vil-CreERT2; Yapf/f;Wwtr1f/f organoids were cultured in the presence of 1 uM 4-OH-Tamoxifen (Sigma-Aldrich) for 72h prior to plating in the absence or presence of GSK-LSD1.
Human organoids. Culture and staining
Human small intestine samples were obtained from patients undergoing elective surgery at Tokyo University Hospital with written informed consent. This was approved by the ethical committee (No. G3553-(7)). Crypt isolation and optimized organoid culture conditions that allow PC differentiation was done essentially as described5,6. In short, stroma was physically removed and the remaining epithelium was cut into 1-mm3 pieces, washed at least 5 times in ice cold PBS, and incubated in 2.5 mM EDTA in ice cold PBS for 1 hour. Isolated crypts were then suspended in Matrigel and seeded in 48-well plates. Domes of polymerized Matrigel were given the refined medium consisting of ‘basal crypt medium’ (see above) supplemented with 10 nM gastrin I (Sigma-Aldrich), 1 mM N-acetylcysteine (Sigma-Aldrich), 100 ng/ml recombinant mouse Noggin (PeproTech), 50 ng/ml recombinant mouse EGF (Thermo Fisher Scientific), 100 ng/ml recombinant human IGF-1 (BioLegend), 50 ng/ml recombinant human FGF-basic (FGF-2) (Peprotech), 1 mg/ml recombinant human R-spondin1 (R&D), 500 nM A83-01 (Tocris) and 50% Afamin-Wnt-3A serum-free conditioned medium. LGR5-iCaspase9-tdTomato organoids were made previously(Fujii et al., 2018). For staining, organoids were isolated from Matrigel using Cell Recovery Solution (Corning) and fixed in 4% paraformaldehyde for 20 min at room temperature. Next, organoids were washed with PBS, and permeabilized with 0.2% Triton X-100 in PBS for 20 min at room temperature. Blocking was done using Power Block Universal Blocking Reagent (BioGenex) for 20 min at room temperature, and rabbit anti-Lysozyme antibody (A0099, DAKO 1:1000 (Fig. 1h), GTX72913, GeneTex, 1:200 (Fig. 2f)) and anti-RFP (600-401-379, Rockland 1:500) was incubated overnight at 4 °C. Organoids were washed 3 times with PBS, and secondary antibody incubation was done for 30 min at room temperature. Nuclear counterstaining was done simultaneously with secondary antibody incubation using Hoechst 33342 (Thermo Fisher Scientific). Stained organoids were suspended in 1 drop of ProLong Diamond Antifade Mountant (Thermo Fisher Scientific) and mounted onto a 35-mm glass bottom dish. Images were captured using a confocal microscope (SP8, Leica).
Immunohistochemical staining of intestinal tissue
For immunohistochemical staining and imaging, tissues were harvested and fixed in swiss rolls. After fixation in formalin, tissues were embedded in paraffin and cut in 4 µm sections. Paraffin sections were rehydrated and peroxidase activity was blocked in 3% hydrogen peroxide. Antigen retrieval was performed in citrate buffer pH6. Sections were stained overnight with primary antibodies against Ki67 (1:500, Thermo Scientific MA5-14520), Lysozyme (1:750, Dako A0099), Sox9 (1:200, Millipore), LSD1 (1:200, Cell Signalling 2184S), H3K4me1 (1:100, Cell Signaling 9723) and H3K4me2 (1:1500, Cell Signalling 9725). The sections were washed in TBS and Tween-20 and stained for 1 hour with HRP-labelled secondary antibody (Dako K4003). The staining was developed with diaminobenzidine (DAB) chromogenic substrate (Dako K5007) and mounted with Glycergel mounting medium (Dako C056330). Tissues were imaged using a Nikon eclips Ci-L microscope.
Immunofluorescence staining of intestinal tissue and organoids
For immunofluorescence labeling and imaging, tissues (first 5 cm of duodenum) were harvested and fixed in swiss rolls. After fixation in formalin, tissues were paraffin embedded and cut in 4 µm sections. Briefly, paraffin sections were treated as before for IHC, and after antigen retrieval were blocked and permeabilized in PBS with TX-100 0.2%, Normal Goat Serum (NGS) 2%, BSA 1% and Tween-20 0.05%. Sections were then stained overnight in the same blocking buffer with primary antibodies against GFP (1:2000 Abcam 13970), YAP (1:200 Cell Signaling Technologies 14074S), OLFM4 (1:200 Cell Signalling 39141S) or LSD1 (1:200 Cell Signaling Technologies 2184S). Tissues were then incubated with the corresponding secondary antibodies for 3 hours (1:500 Alexa Fluor), Rhodamine-labeled UEA1 (5 µg/ml Thermo Fisher Scientific NC9290135) and Hoechst 33342 (1:10,000). Washes were performed with PBS + Tween-20 0.1%.
For organoid staining, organoids were grown in Matrigel on eight-chamber μ-slides (Ibidi 80826) and fixed after exposition to the specific treatments in PBS containing 4% paraformaldehyde (pH 7.4) and 2% sucrose for 20-30 min, permeabilized (PBS, 0.2% Triton X-100) and blocked (PBS-Triton X-100 0.2%, 2% NGS, 1% BSA). Primary antibodies against the following antigens were used, diluted in the same blocking buffer: Lysozyme (1:500, Dako A0099), GFP (1:2000 Abcam 13970) and LSD1 (1:400 Cell Signaling Technologies 2184S) overnight at 4°C with slow agitation. Rhodamine-labeled UEA1 (5 µg/ml Thermo Fisher Scientific NC9290135) and Hoechst 33342 (1:10,000) were used to stain secretory cells and nuclei respectively together with the corresponding secondary antibodies (1:500 Alexa Fluor) and incubated overnight in PBS with 0.2% Triton X-100, 1% NGS and 0.5% BSA at 4°C. Tissue sections and organoids were both mounted using Fluoromount G (ThermoFisher Scientific, 00-4958-02) and imaged with a Zeiss 510 Meta Live or a Zeiss LSM880 confocal microscope, using 20x and 40x objective lens.
In situ hybridization
In situ hybridization was performed on FFPE tissues using RNAscope® 2.5 HD BROWN reagent kit (Advanced Cell Diagnostics (ACD) 322371). Tissue sections (4µm) were deparaffinised with Neoclear and 100% ethanol. The slides were pretreated with hydrogen peroxide for 10 minutes, target antigen retrieval reagent for 15 minutes, and protease plus reagent for 30 minutes (ACD 322300 and 322000). The sections were hybridized with probes for Mm-Wnt3 (ACD 312241), Mm-Olfm4 (ACD 311831), Mm-Atoh1 (ACD 408791), positive control Mm-Ppib (ACD 313911) and negative control Mm-DapB (ACD 310043). For amplification and chromogenic detection the 2.5 HD Detection Reagents BROWN kit (ACD 322310) was used. The slides were counterstained with hematoxylin, dehydrated and mounted with Neomount (Merck 109016). Tissues were imaged using a Nikon eclips Ci-L microscope.
Flow cytometry analysis of organoids
Organoids were mechanically disrupted, centrifuged at 2000 rpm and incubated with TripLE (ThermoFisher) at 37 degrees Celsius for 50 minutes for dissociation into single cells. Cells were incubated with DAPI and analyzed using a flow cytometer (FACSCanto II; BD). Stem cell populations were gated as DAPI negative and GFP-high (top 5%) and analyzed using FlowJo software.
Western Blot
Organoids were harvested in lysis buffer (1% NP-40, 0.02% SDS in 1X TBS) on ice for 30 minutes. Debris was pelleted by spinning down at 14 000 RPM for 30 min. Supernatant was diluted in 4x NuPage sample buffer with 100 mM DTT, and samples were run using precast 4-12% gels using the NuPage system, and blotted using iblot 2 (all ThermoFisher). Membranes were incubated with antibodies against Lysozyme (Dako A0099), Tubulin (Abcam Ab6046), Yap/Taz (Santa Cruz, SC-101199). Secondary antibodies (HRP linked) were swine anti–rabbit (P039901-2) and goat anti–mouse (P044701-2) (DAKO). Imaging was done using SuperSignal West Femto (ThermoFisher) on a Lycor machine. Bands were quantified using Image Studio software.
qPCR
RNA from organoids was isolated using either an RNeasy kit (Qiagen) or Quick-RNA kit (Zymo). Reverse transcription was carried out by using the High-Capacity RNA-to-cDNA Kit (ThermoFisher). Quantitive PCR was performed using the QuantiFast SYBR Green PCR Kit (Qiagen) using primers for Hprt (fwd: cctcctcagaccgcttttt, rev: aacctggttcatcatcgctaa), Actb (fwd: actaatggcaacgagcggttc, rev: ggatgccagaggattccatacc), Lyz1 (fwd: ggcaaaaccccaagatctaa, rev: tctctcaccaccctctttgc), Lyz1 (fwd: gccaaggtctacaatcgttgtgagttg, rev: cagtcagccagcttgacaccacg), Defa, fwd: aatcctcctctctgccctcg, rev: accagatctctcaatgattcctct), Yap1 (fwd: tggccaagacatcttctggt, rev: caggaacgttcagttgcgaa), Wwtr1 (fwd: tggggttagggtgctacagt, rev: ggattgacggtcatgggtgt), Gapdh (fwd: tgttcctacccccaatgtgt, rev: tgtgagggagatgctcagtg), Olfm4 (fwd: ggatcctgaacttttggtgct, rev: acgccaccatgactacagc), and Wnt3 (fwd: ctcgctggctacccaattt, rev: gaggccagagatgtgtactgc). Samples were commonly analyzed in duplicate and RNA expression was calculated either normalized to reference gene, or additionally normalized to control conditions.
RNAseq preparation
RNA for WT and LSD1KO crypts was isolated by sorting IECs (DAPI-, Epcam+) in 2x RNA shield buffer (Zymo) and RNA isolation using the Quick-RNA Micro prep kit (Zymo). Library preparation was done using the Illumina TruSeq Stranded protocol, and samples were sequenced at 75X2 bp PE reads on an Illumina NS500 MO flow-cell. Sequencing was performed by the Genomics core facility (GCF, NTNU). Crypts from E18.5, P7 and E18.5 was directly dissolved in RNA isolation buffer and RNA was isolated using the Quick-RNA micro prep kit (Zymo). Library preparation was done using the NEB Next® Ultra™ RNA Library Prep Kit. Sequencing was performed by Novogene (UK) Co.
RNAseq analysis
Sequenced reads were aligned with STAR to the Mus musculus genome build mm10 (Dobin et al., 2013; Frankish et al., 2019). The number of reads that uniquely aligned to the exon region of each gene in GENCODE annotation M18 of the mouse genome was then counted using featureCounts (Liao et al., 2014). Genes that had a total count less than 10 were filtered out. Differential expression was then determined with DESeq2 using default settings (Love et al., 2014). Interesting differential genes were plotted with a volcano plot using the R package EnhancedVolcano. Heatmaps where generated using the R-package pheatmap. Count values for each gene were transformed to rates per bp by dividing the count for a gene by the length of the total exon region for that gene. Rates per bp where then converted to Transcripts Per Million (TPM) by dividing the rate per bp for each gene by the sum of rates per bp for all the genes in that sample and multiplying with one million. PCA analysis was performed using the function sklearn.decomposition.PCA in scikit-learn. Gene set enrichment analysis (GSEA) was done by sorting the output from DESeq2 by log2 fold change and with the log2 fold change as weights. GSEA was run with the R package clusterProfiler using 10000 permutations and otherwise default settings. Gene sets were generated from published datasets and can be found in supplementary table 3.
Microarray analysis
Gene expression in human small intestinal organoids was analyzed using the PrimeView Human Gene Expression Array. Raw expression data were normalized with the rma function in the R/Bioconductor package affy and the normalized values were used to calculate log fold change(Gautier et al., 2004). For each gene, the probe with the highest absolute log fold change was used. GSEA was run on this list of genes as described for the RNAseq analysis.
ChIP-seq
DAPI-negative, Epcam-positive IECs were sorted in PBS containing 20 mM Sodium Butyrate, and cross-linked by incubation in 1% formaldehyde for 8 minutes. Glycine was added to a final concentration of 125 mM and incubated for 5 minutes at room temperature. Using a swing-out rotor cells were washed 3 times in ice-cold PBS with 20 mM Sodium Butyrate. After washing, cells were snap frozen in liquid nitrogen and stored in −80 °C. The ChIP-seq was carried out similarly to previously described protocols(Dahl and Collas, 2008) Binding of antibodies to paramagnetic beads. The stock of paramagnetic Dynabeads Protein A was vortexed thoroughly to ensure a homogenous suspension before pipetting. Dynabeads stock solution (5 μL per IP) was transferred into a 1.5-ml tube, which was placed on a magnetic rack and the beads captured on the tube wall. The buffer was discarded, the beads washed twice with 200 μL standard RIPA buffer (10 mM Tris-HCl pH 8.0, 140 mM NaCl, 1 mM EDTA, 0.5 mM EGTA, 1% Triton X-100, 0.1% SDS, 0.1% Na-deoxycholate) and resuspended in standard RIPA buffer to a final volume of 100 uL per IP. 99 μL of this was aliquoted into each 0.6 mL tube on ice, and antibody (1.2 ug of anti-H3K4me1: Diagenode, C15410194, Lot A1862D or 4 uL anti-H3K4me2: Cell Signaling #9725, Lot 9) was added per 0.6 mL-tube. Tubes were then incubated at 40 r.p.m. on a ‘head-over-tail’ tube rotator for at least 16h at 4 °C.
Chromatin preparation, Lsd1 cre+ / cre-
Crosslinked cell pellets containing 335 000-500 000 cells were thawed on ice. The 6-10μL pellets were added Lysis buffer (50 mM Tris– HCl pH 8.0, 10 mM EDTA pH 8.0, 1% SDS, 20 mM sodium butyrate, 1 mM PMSF and protease inhibitor cocktail) to a total of 160 μL and incubated on ice for 10 min. The samples were sonicated for 8 x 30 s using a UP100H Ultrasonic Processor (Hielscher) fitted with a 2-mm probe. We allowed for 30 s pauses on ice between each 30 s session, using pulse settings with 0.5 s cycles and 27% power. After the final sonication, 340 μL standard RIPA (with 20 mM sodium butyrate, 1 mM PMSF and protease inhibitor cocktail) was added to the tube while washing the probe, followed by thorough mixing by pipetting. 20 μL was removed as input, and the remaining solution was diluted further with 1mL standard RIPA buffer (with 20 mM sodium butyrate, 1 mM PMSF and protease inhibitor cocktail). The samples and inputs were centrifuged at 12,000 g in a swinging-bucket rotor for 10 min at 4 °C and the supernatants were transferred to a 1.5-ml tube on ice. 66 000 – 100 000 cells were used per IP.
Chromatin preparation, E18.5/P7/P21
The H3K4me1 ChIP-seq for the different developmental stages was slightly modified at the chromatin preparation step. Crosslinked cell pellets containing 50 000-300 000 cells were thawed on ice. The 10 μL pellets were added a modified Lysis buffer (50 mM Tris–HCl pH 8.0, 10 mM EDTA pH 8.0, 0.8% SDS, 20 mM sodium butyrate, 1 mM PMSF and protease inhibitor cocktail) to a total of 120μL and incubated on ice for 10 min, followed by addition of 30 uL PBS with 20 mM sodium butyrate. The samples were sonicated as described above. After the final sonication, 360 uL RIPA without SDS (with 20 mM sodium butyrate, 1 mM PMSF and protease inhibitor cocktail) was added to the tube while washing the probe, followed by thorough mixing by pipetting. The sample and input were centrifuged at 12,000g in a swinging-bucket rotor for 10 min at 4 °C and the supernatants were transferred to a 1.5 ml-tube on ice. Chromatin corresponding to 1 000 - 2 000 cells was removed from each sample to be decross-linked and sequenced as inputs. For the ChIPs, chromatin corresponding to 50 000 cells was used for E18.5 Replicate A, while chromatin from 100 000 cells was used for the remaining samples Immunoprecipitation and washes. Pre-incubated antibody–bead complexes were washed twice in 200 μl standard RIPA buffer by vortexing roughly. The tubes were centrifuged in a mini-centrifuge to bring down any solution trapped in the lid and antibody–bead complexes were captured in a magnetic rack. After removal of RIPA, 177-500 μl of chromatin (equivalent of 50,000-100,000 cells per ChIP) was added to each tube, then incubated at 4 °C, 40 r.p.m. on a ‘head-over-tail’ rotator for at least 16 h. For H3K4me1 ChIPs, the chromatin– antibody–bead complexes were washed three times in 100 μl ice-cold standard RIPA buffer. For H3K4me2 ChIPs, the reactions were washed once in standard RIPA, once in RIPA with increased salt and SDS (300 mM NaCl and 0.20% SDS), once in RIPA with increased salt and SDS (300 mM NaCl and 0.23% SDS), once in standard RIPA. All washing steps were performed with 100uL ice cold buffer supplemented with 20 mM sodium butyrate, 1 mM PMSF and protease inhibitor cocktail. Each wash involved rough vortexing at full speed, repeated twice with pauses on ice in between. Next, a wash in 100μl TE and tube shift was carried out.
DNA isolation and purification
TE was removed and 150μl ChIP elution buffer was added (20 mM Tris-HCl pH 7.5, 50 mM NaCl, 5 mM EDTA. 1% SDS, 30 μg RNase A) and incubated at 37 °C, 1 h at 1,200 r.p.m. on a Thermomixer. The input samples were added ChIP elution buffer up to 150uL and incubated similarly. 1μl of Proteinase K (20 mg/ml stock) was added to each ChIP or input tube and incubated at 68 °C, 4 h at 1,250 r.p.m. The ChIP eluates were transferred to a 1.5-ml tube. Then, a second elution with 150μl was performed for 5 min and pooled with the first supernatant. The ChIP and input DNA was purified by phenol-chloroform isoamylalcohol extraction, ethanol-precipitated with 11μl acrylamide carrier and dissolved in 10-15μl EB (10 mM Tris-HCl).
Library preparation and sequencing
ChIP and input library preparations were performed according to the QIAseq Ultralow Input Library Kit procedure. Sequencing procedures were carried out according to Illumina protocols, on a NextSeq 500 instrument, with 75bp single end reads using high output reagents. The sequencing service was provided by the Norwegian Sequencing Centre (www.sequencing.uio.no).
ATAC-seq
Cells were harvested and frozen in culture media containing FBS and 10% DMSO. Cryopreserved cells were sent to Active Motif to perform the ATAC-seq assay. The cells were then thawed in a 37°C water bath, pelleted, washed with cold PBS, and tagmented as previously described (Buenrostro et al., 2013), with some modifications based on (Corces et al., 2017). Briefly, cell pellets were resuspended in lysis buffer, pelleted, and tagmented using the enzyme and buffer provided in the Nextera Library Prep Kit (Illumina). Tagmented DNA was then purified using the MinElute PCR purification kit (Qiagen), amplified with 10 cycles of PCR, and purified using Agencourt AMPure SPRI beads (Beckman Coulter). Resulting material was quantified using the KAPA Library Quantification Kit for Illumina platforms (KAPA Biosystems), and sequenced with PE42 sequencing on the NextSeq 500 sequencer (Illumina).
Analysis of ATAC-seq data was very similar to the analysis of ChIP-Seq data. Reads were aligned using the BWA algorithm (mem mode; default settings). Duplicate reads were removed, only reads mapping as matched pairs and only uniquely mapped reads (mapping quality >= 1) were used for further analysis. Alignments were extended in silico at their 3’-ends to a length of 200 bp and assigned to 32-nt bins along the genome. The resulting histograms (genomic “signal maps”) were stored in bigWig files. Peaks were identified using the MACS 2.1.0 algorithm at a cutoff of p-value 1e-7, without control file, and with the – nomodel option. Peaks that were on the ENCODE blacklist of known false ChIP-Seq peaks were removed. Signal maps and peak locations were used as input data to Active Motifs proprietary analysis program, which creates Excel tables containing detailed information on sample comparison, peak metrics, peak locations and gene annotations.
ChIP-seq and ATAC-seq analysis
Sequence reads were deduplicated with BBMaps clumpify tool and then aligned with STAR to the Mus musculus genome build mm10 (Dobin et al., 2013; Frankish et al., 2019) (Bushnell, B. BBMap. SourceForge Available at: https://sourceforge.net/projects/bbmap Accessed: 12th February 2019). Peaks were identified using Model-Based analysis of ChIP-seq 2 (MACS2) with peak type set to broad and genome size 2652783500 (Zhang et al., 2008). Input files where supplied for H3K4me1/2 but not for ATAC seq. Peaks from all samples that were compared in differential expression were merged with BEDTools to create a union set and featureCounts was used to count the number of reads, including multi-mappers, for each sample in the union set of peaks (Zhang et al., 2008). Differential peaks was determined from the counts with DESeq2 (Love et al., 2014) using default settings and. deepTools2 was used to create heatmaps (Ramírez et al., 2016). Peak locations were associated with the gene that has the closest transcriptional start site (TSS) with the closest command in bedtools (Quinlan and Hall, 2010). Ties were resolved by only reporting the first hit. TSS sites were downloaded from biomart for GRCm38.p6. The peaks were grouped on the distance to the TSS and the size of each group was plotted. The list of differential peaks with associated genes was grouped by gene and sorted on the differential peak that had the smallest p-value for each gene. Each gene was determined to be either up or down in signal based on whether the total change was above or below zero, where total change is defined as the average of log fold change multiplied by peak length of all peaks associated with that gene. Venn diagrams where created with the R package Euler (Larsson2018, n.d.). Bigwig files describing the score across the genome were created with deepTools2 and scaled to the count of the sample with the least aligned reads for each group (e.g. H3K4me1, H3K4me2 and ATAC) (Ramírez et al., 2016). Heatmaps of regions of interest was created with deepTools2. Chipseq profiles were created in integrative genomics viewer (IGV) (Robinson et al., 2011).
Fetal RNAseq (from Yui et al)
Published microarray raw data was downloaded from ArrayExpress under the accession number “E-MTAB-5246”, normalized with “neqc” function in the R package limma and then log2fc was calculated from the normalized expression values (Ritchie et al., 2015; Yui et al., 2018). GSEA was performed as described in the RNAseq methods.
scRNAseq analysis
Preprocessed and normalized scRNAseq data where downloaded from GSE117783 (Ayyaz et al., 2019). The control treated cells where randomly sub sampled so the two groups had equal number of cells and the density of LSD1 expressing cells where plotted in base R. Y-axis is shortened to show distribution of cells that has an expression larger than zero.
Statistical analysis
Statistical significance was determined either using Student’s t test or 1-way ANOVA with Tukey’s post hoc test, or, when n<10 non-parametric testing (Mann Whitney test) was done. Significance levels are indicated in figure legends.
Data availability statement
All raw data is available through ArrayExpress. WT/KO Crypt RNAseq: E-MTAB-7862, WT/KO Crypt ChIP seq: E-MTAB-7871, WT/KO Crypt ATAC: E-MTAB-8718, E18/P7/P21 RNA-seq E-MTAB-8713, E18/P7/P21 H3K4me1 ChIP-seq: E-MTAB-8710 and Human microarray: E-MTAB-7871.
ACKNOWLEDGEMENTS
We kindly thank Colby Zaph for the initial support of this study and critical reading of the manuscript. We thank Unni Nonstad for assistance with cell sorting. We are indebted to Anne Marthinsen for performing the irradiation after working hours, and the Department of Radiology and Nuclear Medicine (St. Olavs Hospital) for allowing the use of their instruments. We thank the imaging (CMIC) and animal care (CoMed) core facilities that assisted in this work (NTNU). The WT KO crypt RNA-seq was done by the Genomics Core Facility at NTNU, which receives funding from the Faculty of Medicine and Health Sciences and Central Norway Regional Health Authority. The ChIP sequencing was done at the Norwegian Sequencing Centre (www.sequencing.uio.no), a national technology platform hosted by the University of Oslo and supported by the “Functional Genomics” and “Infrastructure” programs of the Research Council of Norway and the Southeastern Regional Health Authorities. Funding of this work was provided by the Norwegian Research Council (Centre of Excellence grant 223255/F50, and ‘Young Research Talent’ 274760 to MJO) and the Norwegian Cancer Society (182767 to MJO). MMA is the recipient of a Marie Skłodowska-Curie IF (DLV-794391). This work was also supported by the South-Eastern Norway Regional Health Authority, Early Career Grant 2016058, and the Research Council of Norway “Young Research Talent” grant to JAD. MF is supported by the Norwegian Research Council (grant no. 275286). The SGC is a registered charity (number 1097737) that receives funds from AbbVie, Bayer Pharma AG, Boehringer Ingelheim, Canada Foundation for Innovation, Eshelman Institute for Innovation, Genome Canada through Ontario Genomics Institute [OGI-055], Innovative Medicines Initiative (EU/EFPIA) [ULTRA-DD grant no. 115766], Janssen, Merck KGaA, Darmstadt, Germany, MSD, Novartis Pharma AG, Ontario Ministry of Research, Innovation and Science (MRIS), Pfizer, São Paulo Research Foundation-FAPESP, Takeda, and Wellcome. This project also received funding from the European Union’s Horizon 2020 research and innovation programme (grant agreements INTENS 668294, MTP, KBJ). The Novo Nordisk Foundation Center for Stem Cell Biology is supported by a Novo Nordisk Foundation grant number NNF17CC0027852 (KBJ).