SUMMARY
Obese individuals exhibit an increase in pancreatic β-cell mass; conversely, scarce nutrition during pregnancy has been linked to β-cell insufficiency in the offspring (reviewed in [1, 2]). These phenomena are thought to be mediated mainly through effects on β-cell proliferation, since a nutrient sensitive β-cell progenitor population in the pancreas has not been identified. Here, we employed the FUCCI (Fluorescent Ubiquitination-based Cell Cycle Indicator) system to investigate β-cell replication in real-time, and found that high nutrient concentrations induce rapid β-cell proliferation. Importantly, we found that high nutrient concentrations also stimulate β-cell differentiation from progenitors in the intrapancreatic duct (IPD). Using a new zebrafish line where β-cells are constitutively ablated, we further show that β-cell loss and high nutrient intake synergistically activate these progenitors. At the cellular level, this activation process causes ductal cell reorganization as it stimulates their proliferation and differentiation. Notably, we link the nutrient-dependent activation of these progenitors to a down-regulation of Notch signaling specifically within the IPD. Furthermore, we show that the nutrient sensor mechanistic Target Of Rapamycin (mTOR) is required for endocrine differentiation from the IPD under physiological conditions as well as in the diabetic state. This study thus reveals critical insights into how cells modulate their plasticity in response to metabolic cues and identifies nutrient sensitive progenitors in the mature pancreas.
RESULTS AND DISCUSSION
β-cell mass increases in response to increased feeding
There is a tight correlation between nutrient intake and β-cell mass in nondiabetic obese individuals (reviewed in [1, 3]) and experimental models of over-nutrition [4, 5]. Whether nutritional cues impinge on the renewal and differentiation of β-cell progenitors remains to be investigated. In mouse, β-cell progenitors are found in the embryonic pancreatic ducts [6-8]. Analogously, in zebrafish, β-cells arise from epithelial cells lining the IPD [9, 10]. A unique advantage of the zebrafish model is the ability to visualize these ductal progenitors in vivo [9, 11]. To explore nutritional control of β-cell progenitors, we analyzed β-cell mass dynamics during two major metabolic transitions. First, by 5 days postfertilization (dpf) (Figure 1A), larvae deplete nutrients stored in the yolk, and transition into a feeding state. Second, between 15 and 16dpf, larvae are switched to a high-calorie diet and grow rapidly until late juvenile stages (45dpf) (Figure 1B) [12]. To characterize β-cell mass responses during these transitions, we examined Tg(TP1:H2BmCherry);Tg(ins:GFP) animals. Tg(TP1:H2BmCherry) drives H2BmCherry expression in Notch responsive cells (NRCs) in the IPD [9]. Since H2BmCherry has a long half-life, this transgenic combination allows the in vivo monitoring of NRC to β-cell differentiation (Figure 1C). This differentiation forms secondary islets (SIs) along the IPD [9, 11]. Intriguingly, we observed a dramatic increase in SI number and principal islet (PI) size after switching to a high-calorie diet at 15dpf (Figures 1D-1G). The new SIs were vascularized and individual β-cells appeared to establish contact with blood vessels (Figures S1A and S1B), suggesting that they contribute to the functional β-cell mass.
β-cells transition from quiescence to proliferation in response to nutrients
This rapid β-cell mass increase after switching to a high-calorie diet suggests that increased nutrient intake stimulates β-cell proliferation and/or differentiation. To determine the role of proliferation, we developed transgenics using the FUCCI system for real-time quantification of proliferation [13, 14]. We placed (zFucci-G1) and (zFucci-S/G2/M) under the insulin promoter for β-cell specific expression (Figure S1C). At 4.5dpf, Tg(ins:zFucci-G1) expression labeled the majority of β-cells in the PI, whereas Tg(ins:zFucci-S/G2/M) expression labeled rare cells (Figures 1H and S1C). To assess this system’s dynamics, we performed live imaging at 4.5dpf. Tg(ins:zFucci-S/G2/M) labeling disappeared shortly after mitosis (Figure 1H), indicating precise labeling of proliferating β-cells. To validate this system further, we performed 5-ethynyl-2’-deoxyuridine (EdU) analyses at 30dpf. Strikingly, the vast majority of Tg(ins:zFucci-S/G2/M) single positive cells (90±12%, n=9 animals) were also EdU+ whereas a minor fraction of Tg(ins:zFucci-S/G2/M);Tg(ins:zFucci-G1) double positive cells had incorporated low levels of EdU (Figures S1D-S1F). These double positive cells are likely in a very early stage of S-phase. For precision, we only scored Tg(ins:zFucci-S/G2/M) single positive cells as proliferating β-cells. Notably, after the switch to a high-calorie diet at 15dpf, β-cell proliferation increased throughout the day, peaking 10-12h after the first feeding (Figures 1I-1J). Furthermore, β-cells in SIs also exhibited high proliferation (Figure S1G). To test whether this β-cell proliferation was stimulated by nutrients, we deprived animals of food for 28h. Fasting dramatically reduced the number of proliferating β-cells (2.5±2.16 cells per PI, n=11 islets) (Figures 1L and 1M) compared to controls (21.4±4 cells, n=7 islets) (Figures 1K and 1M). The effects of fasting were reversible, as β-cells re-entered the cell cycle upon re-feeding (data not shown). This nutrient-driven β-cell proliferation was also observed at earlier time points after switching to a high-calorie diet (Figures S1H and S1I). Thus, β-cell mass expansion is dynamically regulated by nutritional intake.
Nutrients enhance β-cell differentiation from the IPD
Proliferation alone does not explain the increase in the number of SIs, and thus nutrients may also induce β-cell differentiation. We first analyzed whether β-cells in an SI originate from the differentiation of a single, rare IPD cell, followed by clonal expansion, or whether they derive from multiple IPD cells. This question is important because in mouse, IPD cells appear to lose progenitor potential around birth [6-8]. A tamoxifen inducible Cre recombinase was placed under the control of the Notch responsive element (TP1) - Tg(TP1:CreERT2). Using the ubiquitous reporter, Tg(ubi:Switch) [15], and 2F11 immunofluorescence, which marks IPD cells [16], we observed that 4-Hydroxytamoxifen (4-OHT) treatment at 14dpf mosaically labeled IPD cells by 17dpf (Figures 2A, S2A, and S2B). Next, we used Tg(TP1:CreERT2) in combination with the Tg(insulin:Switch) reporter [17]. In this combination, β-cells that originate from IPD cells containing Tg(TP1:CreERT2) activity exhibit H2BGFP expression instead of mCherry. In a single progenitor scenario and under limiting 4-OHT treatments, each SI would be composed of H2BGFP+ or mCherry+ cells. In a multiple progenitor scenario, SIs would be mosaic, containing both H2BGFP+ and mCherry+ cells (Figure 2B). Thus, we treated larvae with 4-OHT at 16dpf and analyzed pancreata at 35-40dpf. The lineage traced cells contributed to mosaic SIs (Figure 2C) with up to 3 discrete H2BGFP+ cells per SI (Figure 2D), indicating that multiple IPD cells contribute to an individual SI. We did not detect H2BGFP+ cells in the SIs two days after the 4-OHT pulse (Figure S2C), indicating that H2BGFP+ cells originated from neogenesis rather than from preexisting β-cells that had retained CreERT2 activity. Moreover, adding 4-OHT before SIs form (2.5dpf), also led to mosaic SI labeling at 32dpf (Figure S2D), supporting the multiclonality of β-cells in SIs, consistent with the polyclonality of mouse [18-20] and human [21] islets. Within the PI, we observed groups of H2BGFP+ cells (8±3.4 cells per group, n=7 groups) (Figure 2E), suggesting that individual NRCs differentiated into β-cells and then underwent several amplification rounds. We also detected H2BGFP+ cells in proximity to SIs (Figure 2F). Subsequent analyses using Tg(TP1:H2BmCherry);Tg(ins:GFP) revealed newly differentiated β-cells approaching an SI via directed migration (Figure 2G). To directly test the involvement of nutrients in β-cell differentiation, we compared the number of SIs in animals that were switched to a high-calorie diet versus siblings maintained on a low-calorie diet between 15 to 20dpf, (Figures 2H and 2I). The restricted diet significantly reduced the formation of new SIs (Figure 2J), indicating that high nutrients induce β-cell differentiation.
IPD cells exhibit a strong regenerative response to β-cell ablation under feeding
Whether IPD cells can increase their endocrine differentiation rate after a selective β-cell loss as well as the metabolic control of such a response, remain unknown. To address these questions, we employed a transgenic system in which β-cells express the cell-lethal Diptheria Toxin α-chain (DTA) [22] under the control of the insulin promoter, leading to complete ablation without a bystander effect (Hesselson et al., in preparation). Conditional β-cell ablation is achieved by Tg(ins:Cre) mediated excision of the BFP cassette from the floxed ins:loxp:BFPloxp:DTA transgene. In the absence of β-cells, the PI core was occupied by α-cells (Figures S3A and S3B).
We examined β-cell ablated animals under fasting metabolism (5 to 6dpf) [23] and during feeding (15 to 21dpf). At 5dpf, they are viable and exhibit a slight body length reduction compared to WT (WT=3.7±0.11mm; β-cell ablated animals=3.5±0.09mm, n=12 animals per group, p<0.0001). To monitor IPD endocrine lineage differentiation, we used Tg(TP1:H2BmCherry) in combination with the pan-endocrine marker Tg(neuroD:GFP). Quantification of the number of Tg(neuroD:GFP)+ SIs revealed no differences in endocrine differentiation in β-cell ablated animals by 6.5dpf (Figures 3A, 3B, and 3E).
Under external nutrition (16.5dpf), β-cell ablated animals feed actively, as indicated by the presence of food in their intestinal tract (Figure S3C), and do not exhibit lethality (1 out of 20) compared to controls (2 out of 20). However, they exhibit a significant growth retardation (Figures S3C and S3D), suggesting that insulin stimulates growth in zebrafish, as it does in humans [24, 25]. In addition, β-cell ablated animals exhibited higher free glucose levels (9.85±2.2 pmol/μg, n=6 animals) compared to unablated siblings (3.64±0.31 pmol/μg, n=3 animals) (p<0.001). Strikingly, by 15.5 and 16.5dpf, β-cell ablated animals presented excessive numbers of ectopic Tg(neuroD:GFP)+ cells in the pancreatic tail (Figures 3C,3D,3F, and S3E-S3G). A majority of Tg(neuroD:GFP)+ cells also exhibited Tg(TP1:H2BmCherry) expression, indicating differentiation from pancreatic NRCs (Figure S3G). By 21dpf, the ectopic Tg(neuroD:GFP)+ cells gave rise to hormone-producing endocrine cells, including Glucagon+ cells (Figures S3H and S3I). In addition, at 16.5dpf, we observed discrete β-cells which had clearly differentiated from NRCs (Figures 3H and S3G). This enhanced endocrine differentiation was accompanied by cell clustering and reduced branching of the IPD (Figure 3H), as well as a loss of duct cell markers, including 2F11 immunofluorescence (Figures 3G and 3H). Using EdU analysis at 20.5dpf, we also found an increase in NRC replication from 8.8% in WT (±2.86, n=9 animals) to 22.9% (±5.45, n=9 animals) in β-cell ablated animals exhibiting IPD clustering (Figures S3J-S3L). To test whether feeding at earlier stages could stimulate endocrine differentiation, we counted SIs after feeding from 5 to 6.5dpf. This short feeding did not significantly increase the number of SIs in β-cell ablated animals (1.4±1.13 SIs, n=13 animals) compared to WT (1.28±1.05 SIs, n=21 animals) (p>0.5). Thus, only under sustained feeding does the lack of β-cells trigger a strong regenerative response in β-cell progenitors, stimulating their proliferation and endocrine lineage differentiation.
IPD cells lose Notch signaling in the absence of β-cells
The IPD phenotypes in β-cell ablated animals, including endocrine cell differentiation and increased proliferation, resemble those of animals with impaired Notch signaling during early larval stages [9]. Therefore, we examined Notch signaling levels using the previously validated Tg(TP1:H2BmCherry);Tg(TP1:VenusPEST) transgenic system [9]. Cells with active Notch signaling are double-positive for H2BmCherry and VenusPEST, whereas cells that recently lost Notch signaling lack VenusPEST because of its short half-life. At 6.5dpf, a majority of IPD cells in WT exhibit active Notch signaling (Figure 3I). Similarly, under fasting conditions at 6.5dpf, the IPD cells in β-cell ablated animals exhibit active Notch signaling (Figure 3J), consistent with a lack of response of NRCs to β-cell ablation in the absence of feeding. In contrast, at 15.5dpf, β-cell ablated, fed animals exhibited strong downregulation of Notch signaling in the IPD (Figures 3K and 3L). This downregulation was IPD-specific, as other tissues, including the intrahepatic duct and brain, did not downregulate Tg(TP1:VenusPEST) expression (Figures S3M, S3N, and data not shown). We further examined direct effects of Notch signaling downregulation on the IPD during feeding stages. Treating WT animals with the γ-secretase inhibitor (GSI) LY411575 [26] from 15 to 18dpf increased β-cell number along the IPD (Figures S3O-S3Q), showing that after Notch signaling downregulation, some IPD cells can differentiate into mature endocrine cells, even at these late stages. GSI-treatment also caused clustering of NRCs, as observed in β-cell ablated animals (Figures S3R and S3S). Together, these data suggest that in β-cell ablated animals, nutrient intake triggers Notch signaling downregulation in the IPD, leading to progenitor activation. Interestingly, Notch signaling levels were reduced in the IPD of WT animals switched to a high-calorie diet compared to those that were maintained on a low-calorie diet between 15 and 20dpf (data not shown), indicating that nutrients also modulate Notch signaling under physiological conditions.
Nutrients and TOR signaling are required for the regenerative response of the IPD
We classified the phenotypes of β-cell ablated animals based on the severity of IPD disorganization and extent of endocrine differentiation (Figures 4A-4E). At 16.5dpf, 10 out of 34 β-cell ablated animals (29%) exhibited excessive clustering of the IPD, while also having the highest numbers of ectopic endocrine cells (Figures 4D-4F), whereas at 21.5dpf, 10 out of 18 animals (56%) exhibited this severe phenotype, suggesting increased penetrance under sustained β-cell demand. At 16.5dpf, we detected rare and weakly Insulin+ cells in the PI; however their numbers did not correlate with the severity of the IPD phenotype. More intriguingly, animals exhibiting the most severe phenotypes were slightly but significantly larger (5.37±0.4 mm, n=10 animals) compared to the rest of the β-cell ablated animals (4.73±0.5 mm, n = 21 animals) (p<0.01). Assuming that growth reflects nutrient intake, these data suggest that higher nutrition in some animals exacerbated the effects of β-cell loss. In agreement, dietary restriction, achieved by a 24h feeding/fasting regimen from 6 to 16dpf, suppressed the IPD phenotypes of β-cell ablated animals (Figures 4G-4I).
Next, we aimed to identify pathways linking β-cell deficiency and nutrient dependent endocrine differentiation. We first tested whether increased glucose could stimulate endocrine differentiation. WT larvae were incubated in D-glucose from 3.5 to 7dpf, a treatment that increases glucose levels during larval stages [27]. D-glucose treatments increased NRC differentiation into endocrine cells, doubling the number of SIs (Figures S4A, S4B, and S4E). L–glucose, which cannot be utilized as a nutrient, had no effect on endocrine differentiation (n=28 animals). In addition, glucocorticoid-treatment, which increases glucose levels [23], mildly increased SI numbers (Figures S4C-S4E). These data further reveal progenitor sensitivity to nutritional cues, including glucose. We also analyzed the role of mTOR, a key intracellular sensor that couples nutrient abundance with cell growth and division across all eukaryotes [28]. To assess its role, β-cell ablated animals were treated from 8 to 16dpf with a low but effective dose of the mTOR inhibitor Rapamycin (50nM) [29]. Significantly, this treatment strongly suppressed the β-cell ablation phenotypes (Figures 4J-4L). Furthermore, it inhibited the loss of Notch signaling (Figures 4M and 4N), indicating that activation of IPD cell plasticity in response to nutrient catabolism requires mTOR activity. To analyze mTOR’s role in endocrine differentiation under physiological conditions, we counted SIs in WT animals treated with Rapamycin or DMSO. Rapamycin treatment significantly reduced the number of newly differentiated SIs, revealing the sensitivity of endocrine progenitors to mTOR activity (Figures 4O-4Q). In addition, PI size was reduced by 26% (2962±364 μm2) compared to controls (3985±730 μm2, n=10 animals per group, p<0.01), consistent with mTOR’s role in endocrine cell growth/proliferation [30, 31]. Importantly, phosphorylated RPS6 (p-RPS6) immunoreactivity, a readout of mTOR activity, was high in the pancreata of fed animals (Figures S4F and S4H) but low after fasting (Figures S4G and S4I), confirming that pancreatic mTOR signaling is responsive to nutrients.
TOR is required for expanding the progenitor pool in the IPD
To further assess pancreatic progenitor sensitivity to mTOR activity, we examined their development in a zebrafish mTOR mutant (mtorxu015Gt), which develops relatively normally until 7dpf but exhibits lethality by 10dpf [32]. In WT, the pancreatic NRCs proliferate between 2.5 and 5dpf and expand posteriorly to form the branched IPD (Figure S4J). In contrast, mtorxu015Gt homozygous mutants display a strong defect in NRC expansion leading to a reduced pool of NRCs and IPD branching defects by 5dpf (Figure S4K). These data reveal a critical requirement for mTOR in establishing the endocrine progenitor pool during development.
Concluding Remarks
Whereas the effects of glucose on β-cell proliferation have previously been described [33-36], we now identify nutrient-sensitive endocrine progenitors in the IPD. Furthermore, we show that β-cell deficiency and nutrients cooperate to enhance IPD plasticity and differentiation (Figure 4R). It is likely that in obesity, impaired β-cell function and higher nutrition synergistically regulate progenitor differentiation as well [37]. Indeed, mature pancreatic cell types in mouse, including duct cells [38], acinar cells [39], and α-cells [40] exhibit enhanced plasticity after pancreatic injury. We also show that enhancement of duct cell plasticity in response to β-cell loss requires the nutrient sensor mTOR, and that Notch signaling, a critical regulator of endocrine differentiation in the embryo [41], is under metabolic control in the mature pancreas. Notably, Notch signaling in human exocrine cells prevents their reprogramming [42]. Furthermore, Notch1 was implicated as a tumor suppressor in a mouse model of pancreatic ductal adenocarcinoma [43], and increased duct cell replication has been reported in Type 2 diabetics and obese patients [44], observations consistent with our data in zebrafish. It will be important to identify the metabolic effectors of duct cell plasticity as they could provide pharmacological means to enhance β-cell differentiation.
Supplementary Material
HIGHLIGHTS.
β-cells transition from quiescence to proliferation in response to nutrient intake.
Nutrient intake induces β-cell differentiation from progenitors in the pancreatic duct.
Constitutive β-cell ablation triggers a regenerative response in the pancreatic duct.
High nutrient intake and mTOR signaling are required to trigger this regenerative response.
Acknowledgments
We thank Ana Ayala and Milagritos Alva for expert assistance with the fish, Oliver Stone and Alethia Villasenor for critical reading of the manuscript, all members of the Stainier group and Amnon Schlegel for helpful discussions. We acknowledge the generosity of Christian Mosimann for providing Tg(ubi:Switch) and technical advice, Atsushi Miyawaki and Asako Sakaue-Sawano for providing cDNAs encoding zFucci-S/G2/M and zFucci-G1, and Xiaolei Xu for mtorxu015Gt. This work was supported by Post-Doctoral Fellowships from the Canadian Diabetes Association (N.N), Larry L. Hillblom Foundation (D.H.), and Juvenile Diabetes Research Foundation (D.H.) as well as grants from the NIH (R01 DK075032; U01 DK089541) and the Packard Foundation to D.Y.R.S.
Footnotes
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