Abstract
Epithelial organs undergo steady-state turnover throughout adult life, with old cells being continually replaced by the progeny of stem cell divisions1. To avoid hyperplasia or atrophy, organ turnover demands strict equilibration of cell production and loss2–4. However, the mechanistic basis of this equilibrium is unknown. Using the adult Drosophila intestine5, we find that robustly precise turnover arises through a coupling mechanism in which enterocyte apoptosis breaks feedback inhibition of stem cell divisions. Healthy enterocytes inhibit stem cell division through E-cadherin, which prevents secretion of mitogenic EGFs by repressing transcription of the EGF maturation factor rhomboid. Individual apoptotic enterocytes promote divisions by loss of E-cadherin, which releases cadherin-associated β-catenin/Armadillo and p120-catenin to induce rhomboid. Induction of rhomboid in the dying enterocyte triggers EGFR activation in stem cells within a discrete radius. When we block apoptosis, E-cadherin-controlled feedback suppresses divisions, and the organ retains the same number of cells. When we disrupt feedback, apoptosis and divisions are uncoupled, and the organ develops either hyperplasia or atrophy. Altogether, our work demonstrates that robust cellular balance hinges on the obligate coupling of divisions to apoptosis, which limits the proliferative potential of a stem cell to the precise time and place that a replacement cell is needed. In this manner, localized cell-cell communication gives rise to tissue-level homeostatic equilibrium and constant organ size.
Main Text
Throughout an animal’s lifetime, mature organs undergo continuous cell turnover yet can maintain the same approximate size. This remarkable ability implies the existence of robust mechanisms to ensure that turnover is zero-sum, with cell production and loss held precisely equal1,2,4. In most organs, production of new cells ultimately depends on divisions of resident stem cells. Although much is understood about how excessive or insufficient divisions lead to disease, little is known about how equal rates of division and loss are sustained during the steady-state turnover of healthy tissues.
We investigated the regulation of turnover in the midgut epithelium of adult Drosophila5 (Extended Data Fig. 1a–e). To establish whether production of new cells equals loss of old cells, we measured kinetics of cell addition and loss in the R4ab region using escargot>GFP flp-out labeling (esgF/OGFP)6,7 (Fig. 1a–e, Extended Data Fig. 1f–g). Newly-added, GFP+ cells increased linearly over time and, after 4 days, comprised all cells in R4ab. Concomitantly, total cell number remained near-constant. We conclude that production of new cells quantitatively equals loss of old cells.
To probe the relationship between cell production and loss, we devised a system to manipulate mature enterocytes and simultaneously track divisions of stem cells by combining enterocyte-specific mexGAL4;GAL80ts (mexts) with split-nlsLacZ clonal labeling8 (Fig. 1f; Extended Data Fig. 2). Using this two-pronged system, we expressed the apoptotic inhibitor p35 in enterocytes and assessed the impact on stem cell divisions. Blocking enterocyte apoptosis resulted in fewer divisions, as indicated by smaller clones (Fig. 1g–i). Apoptotic inhibition also impeded S phase progression (Fig. 1j), consistent with a prior report9. Reduced divisions could be a compensatory means to keep a constant number of total cells. Indeed, total cell number, as well as physical size and morphology, of apoptosis-inhibited midguts remained normal (Fig. 1k, Extended Data Figs. 3a and 4a, b, d, e). These findings imply that enterocyte apoptosis and stem cell division are homeostatically coupled to maintain constant cell number and organ size.
How is coupling mediated? The cell-cell adhesion protein E-cadherin (E-cad) drew our attention because in the mouse intestine, enterocyte E-cad represses stem cell divisions10, and because in other epithelia, E-cad is degraded by caspases during apoptosis11. In the Drosophila midgut, we found that E-cad::mTomato was largely eliminated from the interfaces of dying, Sytox+ enterocytes (Fig. 2a), indicating that apoptotic enterocytes lose junctional E-cad.
To investigate whether E-cad couples divisions to apoptosis, we depleted E-cad in apoptosis-inhibited enterocytes and assessed stem cell divisions by measuring clones. Depletion of enterocyte E-cad did not disrupt epithelial architecture or integrity (Extended Data Fig. 4a, c, d, f–j). It did, however, prevent the reduction in stem cell divisions that would otherwise have occurred following apoptotic inhibition (Fig. 2b–e). Consequently, total cell number increased by 70%, and organs became markedly hyperplastic (Fig. 2h–j, Extended Data Fig 3a). These effects were specific to E-cad as depletion of another cell-cell adhesion protein, echinoid, did not affect cell number (Fig. 2h). Without apoptotic inhibition, E-cad depletion caused excess divisions but not hyperplasia (Fig. 2b), likely due to other, tissue-level effects (Extended Data Fig 5); conversely, E-cad overexpression suppressed divisions (Fig. 2f). These findings show that in apoptosis-inhibited midguts, homeostatic suppression of stem cell divisions requires enterocyte E-cad.
Because E-cad functions as an intercellular homodimer, we considered whether enterocyte E-cad acts by dimerizing with stem cell E-cad12,13. Surprisingly, we found that manipulation of E-cad in stem and enteroblast cells did not alter the rate of stem cell divisions, at least as measured by 4-day clones (Fig. 2k). Thus, enterocyte E-cad acts not through stem cell E-cad, but through a distinct intermediary.
We sought to identify this intermediary. Prime candidates included four E-cad-associated pathways: Wingless/Wnt, Hippo, JAK-STAT, and EGFR. To assess whether these pathways act downstream of enterocyte E-cad, we asked whether E-cad knockdown induced pathway-specific target genes and reporters (Extended Data Fig. 6a–j). E-cad knockdown did not induce Wingless or Hippo targets, JAK-STAT pathway components, or STAT signaling in stem cells, although STAT activity in enterocytes was mildly elevated. Strikingly by contrast, E-cad knockdown induced significant activation of EGFR targets in diploid cells, likely stem cells.
We therefore probed the functional relationship between enterocyte E-cad, stem cell EGFR, and organ size control. In the Drosophila midgut, a specific readout of EGFR activation is diphosphorylation of the effector kinase ERK (dpERK); endogenous diphosphorylation of ERK requires EGFR and occurs primarily in stem cells14–17 (Fig. 3d, e; Extended Data Figs. 3b and 6k). We found that enterocyte E-cad knockdown caused dpERK+ stem cells to increase, whereas overexpression caused them to decrease (Fig. 3a–c, Extended Data Fig. 3b). Furthermore, EGFR was necessary for the excess divisions and organ hyperplasia induced by knockdown of E-cad in apoptosis-inhibited enterocytes (Fig. 3f–g; Extended Data Fig. 3a). Thus, enterocyte E-cad inhibits stem cell EGFR to mediate homeostatic control of cell number and organ size.
How does E-cad on enterocytes control EGFR on stem cells? One possible mechanism involves direct, E-cad-EGFR interaction18; another involves a dispersed signal. To examine these possibilities, we generated isolated, GFP-marked enterocytes that were depleted of E-cad. Measuring the spatial distribution of dpERK+ stem cells surrounding each mutant enterocyte (Fig. 3h–j), we found a zone of strong EGFR activation within 25 µm and a second zone of weak activation from 25–50 µm (Fig. 3k). The spatial extent of these zones suggests the involvement of a dispersed signal.
Consistent with a dispersed signal, two enterocyte-derived EGFs, spitz (spi) and keren (krn), were necessary for organ hyperplasia caused by loss of E-cad in apoptosis-inhibited enterocytes (Fig. 3g, Extended Data Fig. 3a). Surprisingly, however, spi and krn were not induced by E-cad knockdown (Fig. 4a). The visceral muscle EGF vein, modulators of EGF star and argos, and egfr itself were also unaffected. In contrast, the obligate EGF protease rhomboid (rho) was markedly induced by E-cad knockdown and repressed by E-cad overexpression.
Rho operates within EGF-producing cells, where it cleaves EGF precursors for secretion19. Consistent with this function, we found that E-cad repressed rho-lacZ specifically in enterocytes (Extended Data Fig. 7c, d, f). Overexpression of rho in enterocytes caused dpERK+ stem cells to increase; conversely, rho depletion caused dpERK+ cells to virtually disappear (Extended Data Figs. 3b, 7m–o, s). Furthermore, combined depletion of rho and E-cad precluded the hyperactivation of ERK caused by depletion of E-cad alone (Fig. 4b, c). Altogether, these results imply that enterocyte E-cad inhibits stem cell EGFR by preventing EGF secretion through repression of rho.
Does this E-cad-Rho-EGFR relay couple stem cell divisions to enterocyte apoptosis? If so, then: (1) apoptotic enterocytes, which lose E-cad, should upregulate rho; (2) loss of E-cad in apoptotic enterocytes should underlie stem cell EGFR activation; and (3) exogenous manipulation of rho should alter organ size. We investigated each prediction. First, rho-lacZ predominantly marked apoptotic enterocytes during normal turnover (Fig. 4d–f). By comparison, the cardinal injury signal upd320,21 rarely marked apoptotic enterocytes; upd3 was also dispensable for EGFR activation (Extended Data Figs. 6l, 7i; Supplemental Discussion). Thus, rho is silenced in healthy enterocytes but upregulated in enterocytes undergoing physiological apoptosis.
To test the second prediction, we blocked enterocyte apoptosis and examined stem cell EGFR activation. ERK-activated stem cells were virtually absent following apoptotic inhibition but were restored, in a rho-dependent manner, by additional depletion of enterocyte E-cad (Fig. 4g–j, Extended Data Fig. 3b). These results demonstrate that the loss of E-cad in apoptotic enterocytes underlies EGFR activation in stem cells.
To investigate the third prediction, we manipulated enterocyte rho and measured cell number and organ size. Overexpression of rho in apoptosis-inhibited enterocytes led to organ hyperplasia (Fig. 4k, Extended Data Fig. 3a). Conversely, loss of rho in apoptosis-competent enterocytes resulted in organ atrophy (Extended Data Fig. 8). Moreover, combined loss of both rho and E-cad in apoptosis-inhibited enterocytes thwarted the hyperplasia that would have resulted from loss of E-cad alone (Fig. 4k, Extended Data Fig. 3a). These results show that downstream of E-cad, rho is the pivot point that balances division and death to sustain cellular equilibrium.
Finally, how does E-cad control rho expression? To address this, we examined three transcription factors whose nuclear localization is precluded by binding to junctional E-cad: β-catenin/Armadillo (Arm), p120-catenin (p120), and YAP/Yorkie (Yki)22. arm and p120, but not yki, were required in E-cad knockdown enterocytes for induction of rho and hyperactivation of stem cell EGFR (Extended Data Figs. 3b, 7a, g–l). In addition, arm and p120 were required for organ hyperplasia following combined E-cad knockdown and apoptotic inhibition (Fig 4k, Extended Data Fig. 3a). Conversely, overexpression of p120, but not activated armS10, was sufficient for rho induction, EGFR hyperactivation, and organ hyperplasia (Extended Data Figs. 3b, 7b–f, p–r). Thus, E-cad controls rho by inhibiting p120 and Arm, likely through physical sequestration at cell junctions.
Our results demonstrate that steady-state turnover is not driven by constitutive cycling of stem cells. Rather, healthy enterocytes enforce a default state of stem cell quiescence, while sporadic, apoptotic enterocytes trigger replacement divisions (Fig 4l). Because divisions are coupled to apoptosis, turnover remains zero-sum over time.
The molecular mechanism of coupling suggests a simple model for how, during continuous turnover, total cell number is held constant with such robust precision. Apoptotic enterocytes trigger stem cell divisions through loss of E-cad, which induces rho to permit secretion of mitogenic EGFs. Crucially, a single enterocyte can efficiently activate EGFR on stem cells within a ~25 µm radius (Fig. 3k). We propose that this local zone of activation enables organ size homeostasis. If, by chance, stem cells produce excess enterocytes, the stem cells’ physical spacing would increase; consequently, fewer stem cells would reside in the activation zone of the next dying enterocyte, and fewer divisions would result. Similarly, insufficient production of enterocytes would place more stem cells in the activation zone, and more divisions would result. We propose that the radii of individual activation zones, when integrated over the entire epithelium, sets total cell number and organ size. In this manner, localized cell-cell communication can give rise to tissue-level homeostatic equilibrium.
Methods
Drosophila Husbandry
Crosses utilizing the GAL4/GAL80ts system were performed at 18°C. Upon eclosion, adult animals remained at 18°C for 4 days, unless otherwise indicated. On adult day 4, animals were temperature shifted to 29°C to inactivate GAL80ts and induce GAL4-mediated expression. Midguts were harvested for immunostaining 4 days after induction, unless specified otherwise in the figure caption. All other crosses were performed at 25°C; refer to figure legends for individual timepoint information. Adult female flies were used in all experiments.
Fly Stocks
The following stocks were obtained from the Bloomington Stock Center: y, w; shg[mTomato]23, UAS-E-cadherinRNAi (TRiP.HMS00693, TRiP. JF02769, and TRiP.GL00646), UAS-echinoidRNAi (TRiP.GL00648), UAS-rhomboidRNAi (TRiP.JF03106), UAS-spitzRNAi (TRiP.HMS01120), UAS-armadilloRNAi (TRiP.JF01251), UAS-p120ctnRNAi (TRiP.HMC03276), UAS-yorkieRNAi (TRiP.JF03119), UAS-unpaired3RNAi (TRiP.HM05061), UAS-hisH2A:RFP, UAS-p35, UAS-diap1, UAS-rhomboid, UAS-armadilloS10, UAS-p120ctn, Egfrf24/T(2;3)TSTL, Egfrtsla/T(2;3)TSTL, cycE-lacZ, and 10xSTAT-GFP. UAS-kerenRNAi (KK104299) and UAS-armadilloRNAi (KK107344) were obtained from the Vienna Drosophila Resource Center. The following stocks were generous gifts: esg flp-out line6 (a gift from Bruce Edgar), mexGAL424 (a gift from Carl Thummel), UAS-E-cadherinDEFL (a gift from Margaret Fuller), rhoX81 (rho-lacZ) and Upd3.1-lacZ (gifts from Huaqi Jiang), UAS-groucho (a gift from Amir Orian). Other stocks used: esgGAL4, y w hsflp; X-15-29 w+ (‘split-lacZ’)8, y w; y+ X-15-33 (‘split-lacZ’)8, w UAS-CD8:GFP hsflp; tubGAL4; FRT82 tubGAL80, w; FRT82 (used in our prior study3). Detailed information on Drosophila genes and stocks is available from FlyBase (http://flybase.org/).
Immunohistochemistry and Microscopy
Samples were fixed, immunostained, and mounted as previously described3. Primary antibodies: mouse anti-β-galactosidase (1:400, Promega Z3781), mouse anti-Armadillo (1:100, DSHB N27A1), rabbit anti-cleaved caspase 3 (1:200, Cell Signaling, generous gift from D. Bilder3) rabbit anti-diphospho-ERK (1:400, Cell Signaling 4370P), goat anti-HRP-Cy3 (Cappel, 1:100) which stains stem cells and enteroblasts3, mouse anti-Coracle (1:50, DSHB C615.16), mouse anti-Discs large C615.16 (1:50, DSHB 4F3), and rabbit anti-phospho-histone H3, Ser 10 (1:1000, EMD Millipore). Secondary antibodies: Alexa Fluor 488-, 555- or 647-conjugated donkey anti-rabbit or anti-mouse IgGs (1:800, LifeTechnologies A31570, A11001, and A21244). Nuclei were stained with DAPI (LifeTechnologies, 1:1000). Actin was stained with SiR-Actin (Spiro-chrome, 1:500) or Alexa 647-conjugated phalloidin (1:100, LifeTechnologies). Samples were mounted in ProLong (LifeTechnologies). Imaging of samples was performed on a Leica SP8 confocal microscope, with serial optical sections taken at 3.5 µm intervals through the entirety of whole-mounted, immunostained midguts. Representative images are shown in all panels.
Regionalization of the Adult Midgut; Cell Counts and Size Measurements of the R4ab (P1–2) compartment
The Drosophila midgut is compartmentalized along its proximal-distal axis. Each compartment exhibits a characteristic digestive physiology, gene expression pattern, and stem cell division rate7,25. In general, stem cell clones do not cross compartment boundaries25. Our study focused specifically on two adjacent compartments, known alternatively as R4ab or P1–2, which comprise the major region of nutrient absorption. We observed that R4ab consistently exhibited complete cellular turnover between adult days 4–8, as indicated by esgF/O labeling (Fig. 1a–e, Extended Data Fig. 1f–g). Other midgut compartments exhibited variable, incomplete turnover during the same time period, consistent with prior reports; they were not analyzed in this study.
To perform total cell counts of R4ab, this region was first identified in confocal image stacks using morphological landmarks7 (Extended Data Fig 1b–e, g) and digitally isolated in Fiji. Bitplane Imaris software algorithms were applied to generate three-dimensional organ reconstructions and comprehensively count individual cell nuclei by mapping DAPI signals to Imaris surface objects. For analysis of esgF/O midguts, GFP+ cells were additionally counted by mapping DAPI/GFP colocalization signals to Imaris surface objects. R4ab lengths were measured by a spline through the center of individual midguts in Fiji.
Split-lacZ Clone Induction and Analysis
Animals were raised at 18°C and shifted to 29°C four days post-eclosion. Split-lacZ clone induction8 was performed by subjecting animals to two 30-min, 38.5°C heat shocks separated by a 5-min chill on ice. Four days after clone induction, midguts were immunostained and clones in the R4ab region were identified and analyzed by visual examination of serial confocal sections. Clones in regions outside R4ab were excluded from analysis. Clone size was measured as the number of contiguous cells in one discrete clone, as previously described3. No labeled cells were observed in the absence of 38.5°C heat shock.
To ensure that clone counts comprised exclusively stem cell clones and excluded any non-stem cell (transient) clones that were directly labeled by the heat shock, our split-lacZ clonal analyses incorporated two, redundant safeguards. First, a 4-day chase period was included between heat-shock induction and subsequent clonal analysis. Enteroblasts/enterocytes that were directly labeled by the heat shock would have been lost during the succeeding chase period. Confirming that transient clones were nearly absent, only 1–3 single labeled enterocytes were observed per midgut R4ab region after the 4-day chase. As a second safeguard, all single, labeled enterocytes were excluded from our clone counts. This induction protocol resulted in an average of 6–8 clones per midgut R4ab gut region, depending on experimental genotype.
EdU Labeling
Animals were fed yeast paste prepared with 1mg/mL EdU (5-ethynyl-2’-deoxyuridine, Invitrogen) dissolved in water. After 2 days, tissues were fixed as described above and stained for EdU using the Click-iT EdU kit (Invitrogen) based on manufacturer’s protocol.
Sytox Staining
Sytox Green (ThermoFisher, 5mM in DMSO) or Sytox Orange (ThermoFisher, 5mM in DMSO) was diluted 1:5,000 in 5% sucrose. Sytox solution was fed to animals in an empty vial for 5–6 hours26, after which midguts were dissected and mounted in ProLong (LifeTechnologies). Because Sytox is incompatible with fixation, live organs were imaged immediately after mounting.
MARCM Clone Induction
MARCM clone inductions27 were performed by subjecting animals to two 30-min, 38.5°C heat shocks separated by a 5-min chill on ice. For single-enterocyte MARCM clones, animals were dissected 5 days post-induction and terminal clones consisting of one GFP+ enterocyte (identified by its polyploid nucleus) were selected for analysis. GFP+ enterocytes were excluded from analysis if another GFP+ clone was present within an 80 µm radius. Fiji was used to measure the distance between the plasma membrane of the nearest GFP+ enterocyte and the center of dpERK+ stem cells within a 60 µm radius. For mosaic analyses of multicell MARCM clones, animals were fed Sytox three days post-induction and dissected. The proportion of labeled clone cells (GFP+) that were also Sytox+ was quantified.
AG1478 Drug Treatment
Stocks of AG1478 (Sigma) were dissolved in EtOH and subsequently diluted in dH2O to reach a working concentration of 100 µM AG1478 (in 0.02% EtOH). This 100 µM stock solution was used to prepare yeast paste, which was fed to animals as a supplement to their standard cornmeal-molasses diet for the duration of induced gene expression.
Smurf Assay
Smurf assays were conducted by feeding adult animals yeast paste containing 2.5% Brilliant Blue FCF (Sigma) and scoring animals for leakage of dye into the abdomen. Animals were scored as ‘non-Smurf’ if the blue dye was confined to the GI tract and ‘Smurf’ if blue dye leaked outside the GI tract. As a positive control, animals were fed dye in conjunction with 1% SDS.
qRT-PCR
mRNA was extracted from midguts (5 animals/experiment) followed by cDNA synthesis with Invitrogen SuperStrand III First Script Super Mix (Invitrogen). Real-time PCR was performed using the relative standard curve method with SYBR GreenER Supermix (Invitrogen) on a StepOnePlus ABI machine. Expression levels were normalized to mexGAL4ts>CD4-GFP midguts; mef2 transcripts were used as a reference3.
Statistical Analysis
All statistical analyses were performed using Graphpad Prism 6. For comparisons of clone size distributions, unpaired two-tailed Mann-Whitney tests were used to assess statistical significance. (Clone size distributions are non-normal, independent, and derived from a simple random sample.) For comparisons of cell numbers and gut length, unpaired two-tailed t-tests were used to assess statistical significance. (Organ cell number and size distributions are normal, independent, and derived from a simple random sample.) For comparisons of rho gene expression, unpaired two-tailed t-tests were used to assess statistical significance.
Study Design
Sample sizes were chosen based on our previous study3, which also characterized changes in organ cell number and clone sizes. In split-lacZ experiments, single enterocyte clones were excluded from analysis. No other exclusion criteria were applied. No sample randomization or blinding was performed, although automated, Imaris-based computer algorithms were used to analyze and quantify most data in this study.
Extended Data
Supplementary Material
Acknowledgments
J.L. was supported by NSF GRFP DGE-114747 and NIH T32GM007276. This work was supported by NIH R03DK104027 and R01GM116000-01A1 to L.E.O. Confocal microscopy was performed at the Stanford Beckman Cell Sciences Imaging Facility (NIH 1S10OD01058001A1). We thank D. Bilder for the gift of cCas-3 antibody; the Developmental Studies Hybridoma Bank for other antibodies; D. Bilder, B. Edgar, M. Fuller, H. Jiang, B. Ohlstein, C. Thummel, the Bloomington Drosophila Stock Center (NIH P40OD018537), the TRiP at Harvard Medical School (NIH/NIGMS R01-GM084947), and the Vienna Drosophila Resource Center for fly stocks; J. Axelrod, M. Goodman, M. Fuller, W.J. Nelson, R. Nusse, M. Krasnow, T. Nystul, and D. Fox for comments on the manuscript; and M. Mirvis, B. Benham-Pyle, N. Pierce, and D. Gordon for helpful discussions.
Footnotes
The authors declare no competing interests.
Author contributions: J.L. and L.E.O. designed the experiments and wrote the manuscript. J.L. and S.N. prepared microscopy specimens. J.L., S.B., and S.N. performed confocal microscopy. J.L. performed all other experiments, genetic crosses, data analysis, and statistical analysis.
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