Weaning from maternal milk induces the switch from mTORC1 to AMPK signaling in β cells.

To investigate what regulates the reciprocal relationship between mTORC1 and AMPK signaling in maturing β cells, we analyzed both adult human and adult mouse islets that were treated for 12 hours with either everolimus (an mTORC1 inhibitor) or 5-aminoimiazole-4-carboxamide-1-β-D-ribofuranoside (AICAR, an AMPK activator) (19). As expected, everolimus treatment inhibited mTORC1 activity in β cells, as revealed by a dramatic reduction in specific rpS6 phosphorylation, but hardly altered AMPK phosphorylation. By contrast, AICAR treatment was sufficient to both activate AMPK and completely abrogate mTORC1 phosphorylation in both mouse and human islets (Figure 2, A and B, with associated quantifications in Supplemental Figure 2, A and B). Taken together, these findings indicate that inhibiting mTORC1 activity is not sufficient on its own to activate AMPK in β cells, but that activating AMPK is sufficient to repress mTORC1. This unidirectional relationship supports the concept that AMPK, as in other systems, is a key repressor of mTORC1 in β cells. Indeed, its activation is responsible for shutting off mTORC1 signaling, triggering a key step in β cell maturation.

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Figure 2

Weaning from maternal milk induces the switch from mTORC1 to AMPK signaling in β cells.

(A and B) Representative immunoblots showing p-AMPK, p-rpS6, total AMPK, total rpS6, and β-tubulin in (A) adult human islets and (B) adult mouse islets after treatment with 1 mM AICAR or 40 μM everolimus for the indicated time (n = 3 for both human and mouse islets except only for the 3-hour AICAR treatment condition, where n = 2). (C) Experimental paradigm for the milk fat–supplemented diet (MFD), and immunofluorescence (IF) staining for insulin (green) and p-rpS6 (red) in representative pancreatic sections from P40 mice otherwise fed either standard chow (control) or the MFD. Nuclei were counterstained with DAPI (blue). Original magnification, ×64. (D) Experimental paradigm for the MFD and associated representative Western blots (WB), showing p-AMPK, p-rpS6, total S6, and total AMPK in control and MFD mice. β-Tubulin was used as loading control. Quantification of p-AMPK/AMPK and AMPK/β-tubulin is shown below. **P = 0.0062 (unpaired t test corrected for multiple comparisons using the Holm-Sidak method). (E) Intraperitoneal glucose tolerance test in control and MFD mice (n = 7). *P < 0.05 (unpaired t test corrected for multiple comparisons using the Holm-Sidak method). (F) Percentage of Ki67⁺ and insulin⁺ cells in control and MFD islets (n = 3–5). ***P = 0.0002 (2-tailed unpaired t test).

The process of weaning, which in mice starts in the third week of life, is associated with decreased reliance on the consumption of fat-rich maternal milk in favor of an adult-type diet with a more carbohydrate-rich macronutrient composition (33). Given the role of mTORC1 as a nutrient-sensing kinase, we hypothesized that postnatal β cell maturation may represent an adaptation to the cessation of milk consumption, and that mTORC1 repression through AMPK activation may act as a physiological mediator of this process.

To test this hypothesis, we enabled mice to continue to consume a diet highly enriched for milk fat by giving them access to a milk fat–rich diet and supplementing this with daily milk fat gavage, beginning at P11–P13 and continuing through adulthood (P40). This allowed us to maintain high milk fat consumption in the mice during the period over which weaning normally occurs. Remarkably, allowing mice to continue assimilating milk fat throughout their entry into adulthood, a period during which this is usually declining, was sufficient to allow β cells to maintain neonatal levels of mTORC1 activity, which was otherwise completely repressed in the control mice (Figure 2C).

We next assessed the expression and phosphorylation of β cell AMPK and rpS6 in the islets of these mice. Unlike the AMPK activation seen in the islets of adult mice weaned onto standard chow, islets from mice that continued milk fat consumption past weaning displayed a marked suppression of AMPK phosphorylation, although not all the way to neonatal levels, despite the entry into adulthood (Figure 2D). Similarly, extending milk fat consumption past weaning tended to produce higher levels of p-rpS6 within the islets of adult mice, though this trend was not significant (Figure 2D and Supplemental Figure 2C).

We also examined whether continuing milk fat consumption from the neonatal period into adulthood in mice impacts β cell function. Mice provided with milk fat by gavage from P11 into adulthood had lower serum glucose levels than did vehicle-treated control mice, both at baseline and during a glucose tolerance test, indicative of constitutively enhanced glucose clearance (Figure 2E and Supplemental Figure 2D). Supporting this, β cell (INS⁺) area tended to be higher in milk fat-supplemented mice than in controls, though this trend was not statistically significant (Supplemental Figure 2E). Moreover, the percentage of proliferating (Ki67⁺) β cells was higher in the mice provided milk fat than it was in control mice (Figure 2F). We also found that mRNA levels for the maturity markers Ins1, Ins2, and Mafa were lower in the islets of the mice provided milk fat (Supplemental Figure 2F). Islet mRNA levels of Ucn3, by contrast, were not affected, although this is not surprising given that approximately 93% of β cells already express Ucn3 by P13, which is prior to weaning (1).

These data together highlight that continuing milk fat consumption from the neonatal period into adulthood in mice is sufficient to maintain an immature-cell phenotype, and strongly support the concept that shifting from fat to carbohydrate consumption during weaning is a previously unrecognized mechanistic driver of mTORC1-to-AMPK switching and consequent functional β cell maturation.

mTORC1 maintains the immature phenotype of pancreatic β cells.

To definitively test whether mTORC1 is responsible for maintaining the immature phenotype of neonatal pancreatic β cells, we generated Rip-Cre Tsc1^fl/fl (βTSC1-KO) mice, where TSC1, the upstream negative regulator of mTORC1, is deleted, leading to a constitutive activation of mTORC1 in β cells (Figure 3A). βTSC1-KO mice also carry a YFP allele, having been crossed onto a ROSA26-stop-EYFP line, with the latter serving as a lineage tracing tool to assess Cre expression and genomic recombination. All subsequent experiments were performed using βTSC1-KO mice or littermate (Tsc1^fl/fl; or Tsc1^+/+, Cre⁺) controls. We confirmed that Cre expression in βTSC1-KO mice is restricted to pancreatic β cells by demonstrating colocalization of insulin and YFP staining (data not shown). Transcriptional analysis confirmed that Tsc1 mRNA is significantly reduced in islets from βTSC1-KO mice versus those from control mice (Figure 3B). Confirming the direct consequence of deleting TSC1 specifically in β cells, approximately 85% of pancreatic β cells in βTSC1-KO mice had increased specific p-rpS6 staining intensity, indicative of upregulated β cell mTORC1 activity (Supplemental Figure 3, A and B). Measuring pancreatic INS⁺ area histologically showed that constitutive activation of mTORC1 in adult βTSC1-KO mice increases β cell mass (Figure 3C), mirroring what was seen in response to milk fat gavage.

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Figure 3

mTORC1 maintains the immature phenotype of pancreatic β cells.

(A) Genetic model for constitutive activation of mTORC1. (B) mRNA levels of Tsc1 in islets from control and βTSC1-KO mice (n = 7–9). ***P = 0.0002 (2-tailed unpaired t test). (C) Relative β cell areas in pancreatic sections from control and βTSC1-KO mice, measured as the ratio of insulin⁺ area/insulin⁺ cell number (n = 37–38 islets/group). ****P < 0.0001 (2-tailed unpaired t test). (D) GSEA, showing enrichment of disallowed genes in RNAseq data from βTSC1-KO islets (P < 0.05). (E) Heatmap, showing the transcriptional profile of β cell maturity markers in βTSC1-KO and control mice (P < 0.05). (F) Insulin secretion (percentage of insulin content) in a perifusion assay on βTSC1-KO and control islets exposed to 2.8 mM and 16 mM glucose (n = 4–5). *P < 0.05 (unpaired t test corrected for multiple comparisons using the Holm-Sidak method). (G) Model for inducing mTORC1 activation in adult mice. (H) Plasma insulin levels after glucose challenge in iTSC1-KO and control mice (n = 8–9). *P = 0.017, **P = 0.0012 (unpaired t test corrected for multiple comparisons using the Holm-Sidak method). (I) Intraperitoneal glucose tolerance test in iTSC1-KO and control mice (n = 10–11). *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 (unpaired t test corrected for multiple comparisons using the Holm-Sidak method).

To more fully determine whether constitutively activating mTORC1 recapitulates an immature islet phenotype in adult mice, we performed RNAseq on islets from adult βTSC1-KO and control mice. We first explored a list of β cell disallowed genes that are known to be upregulated in neonatal islets (2, 34). GSEA showed that adult βTSC1-KO islets were enriched for the expression of such genes (Figure 3D). Moreover, expression of key maturation genes (Ins1, Ins2, Unc3, G6pc2, Erol1b, Syt4, Pcsk1, and Mafa) was concomitantly reduced in adult βTSC1-KO islets when compared with control islets (Figure 3E). Assessment of MafA by immunostaining mirrored the transcriptional data, as MafA expression was reduced in the islets of βTSC1-KO versus control mice (Supplemental Figure 3C). Together, these data indicate that forcing β cell–specific mTORC1 activation in mice results in the expression of a transcriptional pattern, including critical genes, that is typical of neonatal islets.

In addition to sharing a transcriptional signature with neonatal β cells, βTSC1-KO islets were also functionally immature. For example, serum insulin levels in βTSC1-KO mice were higher, both after fasting and after glucose stimulation when compared with those of control mice (Supplemental Figure 3D). Correspondingly, βTSC1-KO mice had relatively lower fasting blood glucose levels (mean value of 83 mg/dL in βTSC1 KO vs. 109 mg/dL in control) and cleared injected glucose faster (Supplemental Figure 3, E and F). Furthermore, analysis of dynamic insulin secretion ex vivo showed that βTSC1-KO islets have increased basal insulin secretion at low glucose concentrations but fail to significantly increase this secretion in response to glucose challenge, a behavior seen in immature β cells (Figure 3F).

To further confirm the ability of mTORC1 to elicit an immature phenotype in adult pancreatic β cells, we also crossed Tsc1^fl/fl mice with those expressing the inducible MIP1-CreER driver in order to generate iTSC1-KO mice (Figure 3G). We first noted that treating adult iTSC1-KO mice with tamoxifen was sufficient to markedly induce p-rpS6 staining among β cells in pancreatic islets (57% stained for p-rpS6), indicative of strong mTORC1 activation (Supplemental Figure 3G). In assessing β cell function we noted that, similar to what was seen in both βTSC1-KO mice and WT mice maintained on dietary milk fat during their entry into adulthood, fasting blood glucose levels were lower after 3 weeks of tamoxifen administration in iTSC1-KO mice than in Tsc1^fl/fl (no Cre) littermates (Supplemental Figure 3H). Moreover, serum insulin levels were higher, both after fasting and after 30 minutes of glucose stimulation, and injected glucose was correspondingly cleared more rapidly in iTSC1-KO mice versus controls (Figure 3, H and I). Together, these data show that inducing mTORC1 activation in adult β cells is sufficient to spontaneously revert them to an immature, neonatal-type functional state.

AMPK activation triggers a switch to oxidative metabolism in mature islets.

In line with data from Yoshihara et al. (3), analysis of our RNAseq data from neonatal and adult pancreatic β cells (Supplemental Figure 1C) revealed an induction of TCA cycle and electron transport chain pathway genes during the transition to adulthood. Intriguingly, 9 out of the total 13 genes encoded by mitochondrial DNA (mtDNA) were upregulated in adult pancreatic β cells during this transition. This included the genes encoding cytochrome c oxidase subunits I, II, and III; the NADH dehydrogenases 1, 2, 4, 5, and 6; and the ATP synthase F₀ subunit 6. The expression of other genes, including Uqcrh, Uqcrq, Cox5b, and Cox6a2, which are part of the mitochondrial respiratory chain, was also increased in adult versus neonatal pancreatic β cells (Figure 4A). Together, these data suggest that the maturation of pancreatic β cells is orchestrated in conjunction with increased mitochondrial function and oxidative metabolism.

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Figure 4

AMPK activation triggers a switch to oxidative metabolism in mature islets.

(A) Heatmap, profiling mitochondrial genes and genes related to ox-phos metabolism in P6 and P45 β cells (P < 0.05). (B) qPCR, showing the ratio of mtDNA to nuclear DNA in P6 and adult islets (n = 4–5). ****P < 0.0001 (2-tailed unpaired t test). (C) Relative Ppargc1a and Ppargc1b mRNA levels in P6 and adult islets (n = 4). *P = 0.016, **P = 0.0029 (unpaired t test corrected for multiple comparisons using the Holm-Sidak method). (D) Oxygen consumption rates for cultured neonatal and adult islets (Seahorse XF24), with data presented relative to baseline (n = 7–8). *P < 0.01, **P < 0.001, ***P < 0.0001 (unpaired t test corrected for multiple comparisons using the Holm-Sidak method). Oligo, oligomycin; Rot, rotenone; AA, antimycin A. (E) Heatmap, showing the transcriptional profile of mitochondrial genes those related to ox-phos metabolism in AMPKdKO control islets (P < 0.05). (F and G) Morphometric analysis of mitochondria in AMPKdKO and control β cells, showing (F) the number of mitochondria per 70 μm² (n = 32–36), and (G) the volume density (mL%). n = 29–33 β cells per group. ****P < 0.0001 (2-tailed unpaired t test). (H) qPCR, showing the ratio of mtDNA to nuclear DNA in WT adult islets treated with 10 μM compound C (CC) (n = 3). ***P = 0.0003 (2-tailed unpaired t test). (I) Ppargc1a and Ppargc1b mRNA levels in the islets of control mice vs. those receiving milk fat throughout their development into adulthood (n = 3). MFD, milk fat–supplemented diet. **P = 0.0013 (unpaired t test corrected for multiple comparisons using the Holm-Sidak method).

To address the mechanism(s) leading to increased expression of mitochondrial genes, we assessed the ratio of mtDNA copy number per genomic DNA copy number by qPCR. mtDNA copy number in adult islets (mean of 444/cell) was approximately2 times higher than in neonatal ones (mean of 218/cell) (Figure 4B), suggesting that β cell maturation is tied to increased mitochondrial mass. Consistent with this concept, mRNA levels of 2 regulators of mitochondrial biogenesis, Ppargc1a (PGC1α) and Ppargc1b (PGC1β), were also higher in adult versus neonatal islets, suggesting that mitochondrial biogenesis is important for β cell maturation (Figure 4C).

Given that the abundance of mtDNA is tightly correlated with cellular respiratory activity (35), we sought to directly assess mitochondrial function by measuring oxygen consumption in neonatal and adult islets using a bioenergetic analyzer (Seahorse XF24). The glucose-stimulated increase in oxygen consumption was markedly higher in isolated adult versus neonatal islets, as was maximal oxygen consumption, which can be measured after treating islets with the mitochondrial uncoupler carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone (FCCP) (Figure 4D). These data confirm that the increase in β cell mitochondrial mass seen during the transition from neonatal to adult life is commensurate with a pronounced induction of mitochondrial activity.

AMPK activation is known to promote oxidative metabolism (36). We therefore hypothesized that AMPK activation might be responsible for the increase in β cell mitochondrial number and function that we saw during islet maturation. To investigate this possibility, we simultaneously eliminated both AMPK1 and AMPK2 (AMPKdKO) expression selectively in pancreatic β cells by crossing Prkaa1^fl/fl/Prkaa2^fl/fl mice with Ins-Cre (26) or Rip-Cre (37) mice. RNAseq of islets from adult AMPKdKO (Ins-Cre) and floxed (no Cre) control mice showed that the transcriptomic signatures of AMPKdKO islets were similar to those of WT neonatal islets, importantly including downregulation of the same set of mtDNA-encoded and respiratory pathway genes induced during normal adult β cell maturation (Figure 4E). Monitoring mitochondrial numbers by electron microscopy (EM) confirmed that adult AMPKdKO (Rip-Cre) islets have fewer and smaller mitochondria than those of controls (Figure 4, F and G). We also treated adult islets from WT mice with compound C, an AMPK inhibitor (38) to confirm our findings in AMPKdKO mice. Remarkably, compound C treatment for 24 hours reduced the number of mitochondria in adult islets, recapitulating the knockout phenotype (Figure 4H).

The importance of AMPK regulation during β cell maturation was also evident in our studies with milk fat supplementation. For example, WT mice that continued to consume milk fat throughout the period when weaning normally occurs had lower islet mRNA levels of Ppargc1a (no difference for Ppargc1b) than did mice weaned onto regular chow in a normal manner (Figure 4I). Taken together, our findings highlight the important role of cellular AMPK activation as a switch that is flipped in response to shifting patterns of macronutrient consumption and that turns on β cell oxidative metabolism, a hallmark of their adult phenotype.

Metabolic and physiological analyses.

Fasting blood glucose concentrations in mice were determined using a handheld glucometer (Freestyle Lite) after 6 hours of fasting. For in vivo glucose challenges, mice were fasted for 6 hours, after which serum insulin and blood glucose were collected by submandibular bleeding prior to and 30 minutes following an i.p. injection of glucose (2 g/kg); serum was collected by centrifugation and insulin levels measured by ELISA (Mercodia). For glucose tolerance tests, blood glucose levels were measured before, and 15, 30, 60, and 120 minutes following an i.p. injection of glucose (2 g/kg).

An automated perifusion system (BioRep Technologies) was used to measure dynamic insulin secretion by islets. Twenty to 30 size-matched islets per mouse were immobilized in Bio-Gel P-4 Gel (Bio-Rad) that was maintained at 37°C in a temperature-controlled chamber. Islets were perifused at 100 μL/min with Krebs-Ringer bicarbonate buffer supplemented with 0.25% BSA (KRB buffer) and containing 2.8 mM glucose for a preincubation period of 30 minutes. Islets were next perifused in sequence with KRB buffer containing 2.8 mM glucose for 20 minutes, following by KRB buffer containing 16.7 mM glucose for 30 minutes, and finally KRB buffer containing 30 mM KCl for 10 minutes. Flow-through was collected every minute during the 3 perifusion steps of the assay. Islets were next recovered from the chambers and insulin content was extracted by overnight incubation in acid/ethanol solution. For static measurements of GSIS by db/db islets, the newly isolated islets were first incubated overnight in medium with 11 mM glucose, after which they were preincubated in KRB buffer containing 2.8 mM glucose with or without 1 mM AICAR or 40 μM everolimus for 1 hour. The islets were next incubated in KRB buffer containing 2.8 mM glucose for 30 minutes, 16.7 mM glucose for another 30 minutes, and then KCl as above, with respective AICAR or everolimus treatment maintained during steps. The supernatant was collected after each incubation. Total insulin content was then extracted from islets by overnight incubation with acid/ethanol solution. Insulin in the supernatants and islet extracts was then measured by ELISA.

Flow cytometry and β cell sorting.

Islets from neonatal (P6) or postweaning (P45) mice were dissociated, fixed, permeabilized, and stained for p-rpS6 and insulin (5018 or 9008, respectively, Cell Signaling Technology) for FACS analysis using a BD LSRFortessa X20 Dual machine. FACS data were analyzed with FlowJo software.

Islets from MIP-GFP mice were digested into a single-cell suspension and sorted for GFP by FACS to 85%–95% average purity. Sorted cells were then processed for RNA extraction.

Human islet samples were obtained from both healthy adult organ donors and adult donors with T2D. Islets were dissociated using Accumax (STEMCELL Technologies) for 7–8 minutes and washed 3 times with PBS containing 2% BSA and 2.8 mM glucose. Cells were next incubated for 30 minutes at 4°C in the presence of the pancreatic marker antibodies HPi2 (77-HPI2-PU-0.1, clone HIC1-2B4, ALPCO) and HPa3 (provided by Markus Grompe, Oregon Health Sciences University, Portland, Oregon, USA). After washing, samples were incubated at room temperature for an additional 30 minutes in the presence of fluorescent-coupled secondary antibodies. Cells were next washed and resuspended in PBS containing 2% BSA and 2.8 mM glucose. Before sorting, DAPI stain was added to distinguish live versus dead cells. The details of the gating strategy are shown in Supplemental Figure 4A.

Transmission EM analysis.

Pancreatic islets (~30) were fixed in 2.5% (w/v) glutaraldehyde in PBS at 4°C. After being washed in PBS, islets were postfixed in 0.1% (w/v) osmium tetroxide in PBS and dehydrated in a graded series of ethanol. The islets were then rapidly transferred to propylene oxide and embedded in PolyBed 812 (Polysciences, Inc).

Ultrathin sections were cut with a diamond knife, stained with uranyl acetate and lead citrate, and observed under a Zeiss 902 transmission electron microscope. β Cells were identified under EM by their distinct morphology including dense core granules (60). β Cells were considered dead based on a loss of plasma membrane integrity, fragmentation into discrete bodies, or engulfment of the cell corpse or its fragments by an adjacent cell (61). The presence of marked chromatin condensation and/or of blebs was considered signs of apoptosis, whereas massive vacuole accumulation and absence of chromatin condensation were considered signs of autophagy (62).

For morphometric studies, volume density values were derived from the evaluation of 3–5 different islets for each pancreas. Twelve photomicrographs were taken of each islet at ×10⁵ magnification. A graticule (11 × 11 cm) composed of 169 points was placed on the micrographs, and the number of points intersecting the autophagic vacuoles and mitochondria was counted. Volume density of intracellular organelles was calculated according to the formula VD = Pi/Pt, where Pi is the number of points within the subcellular component and Pt is the total number of points within the cells, and expressed in mL/100 mL of tissue (mL%) (63).

Assessments of cellular metabolism and mitochondrial number.

The OCR of β cells was measured in real time using a Seahorse XF24 analyzer (Seahorse Bioscience). Batches of 30 islets were loaded into islet-capture microplates and covered with a mesh to prevent movement of the islets during the assay. This was followed by incubation in a non-CO₂ incubator at 37°C for 1 hour. Three to 5 measurements in 2.8 mM glucose were taken, followed by replacement with 16.7 mM glucose and sequential addition of 5 μM oligomycin (Cell Signaling Technology, 9996L), 1 μM FCCP (C2920-10mg), 5 μM rotenone (R8875-1G), and 5 μM antimycin A (A8674-50MG) (all 3 from Sigma-Aldrich). OCR was normalized to the average baseline measurement in 2.8 mM glucose, and expressed as a percentage change from this baseline during the assay. After the experiment, the islets underwent DNA extraction, and the number of mitochondria was determined by qPCR as a ratio of mtDNA copy number per genomic DNA copy number using the following primers. For mtDNA: forward, GAGCATCTTATCCACGCTTCC; reverse, GGTGGTACTCCCGCTGTAAA. For genomic DNA: forward, CTTCCCCACTGGCCTCAAG; reverse, CCAAAACCCAGTGATCCAGC.

Immunofluorescence staining and morphometric analysis of islets.

A standard immunofluorescence protocol was performed to stain paraffin sections of fixed human and mouse pancreata using the following primary antibodies: anti–p-Ser240/244-rpS6 (5364, Cell Signaling Technology), anti–p-Thr172-AMPK (2535S, Cell Signaling Technology,) anti-insulin (A056401-2, Agilent), anti-Pdx1 (F109-D12, Developmental Studies Hybridoma Bank), and anti-MafA (NBP1-00121, Novus Biologicals). Briefly, pancreatic tissue was flushed with PBS, dissected, fixed in buffered zinc formalin fixative (Z-Fix, Anatech Ltd) overnight, dehydrated in grades of ethanol, and processed for paraffin embedding. Sections (5 μm) were heated, de-waxed, rehydrated through a series of ethanol dilutions, and then placed in distilled H₂O. Heat-induced epitope retrieval was performed by immersion in antigen unmasking solution (H-3300, Vector) and microwave heating. After blocking with antibody diluent (DAKO), sections were incubated with a primary antibody and then with appropriate Alexa Fluor 488– or 555–coupled secondary antibodies (Jackson ImmunoResearch). Nuclei were counterstained using DAPI. To stain with anti–p-AMPK (Thr172), PBS-flushed pancreata were perfused in vivo with 4% paraformaldehyde (PFA) via intracardiac catheter prior to dissection in order to fix the pancreas as quickly as possible. The pancreata were then dissected and incubated for 2 hours in 4% PFA and then conserved in OCT at –80°C until sectioning. Cryosections were then washed with PBS followed by the antigen retrieval and blocking steps as described above.

RNA isolation, RT-PCR, and real-time PCR.

RNA was isolated using Direct-zol (Zymo Research) and digested with RNase-free DNase to eliminate DNA. RNA (50–200 ng) was used for preparation of cDNA using Superscript IV Reverse Transcriptase (Invitrogen) by the oligo(dT) priming method. qPCR was performed using the 1× Fast SYBR Green Mix (Applied Biosystems) in an ABI7900HT system (Applied Biosystems). The expression levels of each transcript were normalized to the housekeeping gene RPLP19. The primer sequences used are listed in Supplemental Table 7.

RNAseq library generation.

For neonatal versus adult, βTSC1 KO versus control, and db/db versus control RNAseq analyses, total RNA was isolated from the appropriate preparation using Direct-zol RNA MicroPrep plus (Zymo Research), following the manufacturer’s instructions. Sequencing libraries were prepared from 10–25 ng of total RNA using the SMARTer Stranded RNAseq Kit (Clontech) according to the manufacturer’s protocol. Briefly, after rRNA depletion, remaining RNA was used for first-strand cDNA synthesis and purification, after which the samples were ligated to unique adapters and PCR amplified. Libraries were then validated using a 2100 BioAnalyzer (Agilent), normalized, and pooled for sequencing using the UCSF Institute for Human Genetics core service.

To process RNAseq libraries, adaptor sequences were trimmed using Cutadapt version 1.13 (64), requiring a length greater than 20 nt after trimming, and quality filtered by requiring all bases to have a minimum score of 20 (-m 20 -q 20). Only reads that passed the quality or length threshold on both strands were considered for mapping. Reads were mapped with the spliced-read mapper GSNAP (version 2016-08-10) (65) using default parameters. Reads were aligned against the NCBIM37 (mm9) genome build release 67 downloaded from Ensemble. Reads mapping to the genes of interest were counted using HTseq (v.0.7.1) (66) in the union mode. Differential expression analysis was performed in R using DESeq2 (v.1.16.0) (67) with the default parameters, including the Cook’s distance treatment to remove outliers. AMPKdKO RNAseq data were obtained as previously described (26).

RNAseq data that support the findings of this study have been deposited in the NCBI’s Gene Expression Omnibus (GEO) under accession code GSE132261. From Kone et al., data for RNAseq are available at the European Molecular Biology Laboratory–European Bioinformatics Institute (EMBL-EBI) ArrayExpress website under accession number E-MTAB-2791.

Immunoblotting.

Cells were lysed in ice-cold RIPA buffer containing a cocktail of protease and phosphatase inhibitors. Lysates were kept on ice for 5 minutes and cleared by centrifugation. Protein concentrations were determined using the Pierce 660nm Protein Assay, and immunoblotting was performed using the Bolt mini-gel and iBlot2 system. Membranes were next blocked with 5% BSA and incubated overnight with primary antibodies anti–p-rpS6 (Ser240/244, 1:1000, 5364), anti–p-AMPK (Ser172, 1:1000, 2535S), anti-rpS6 (2217S, 1:1000), anti-AMPK (5831S, 1:1000) (all from Cell Signaling Technology), and anti–β-tubulin (ab11307, 1:5000) and anti-GAPDH (ab8245, 1:5000) from Abcam. Primary incubation was followed by a 1-hour incubation with the secondary antibody at room temperature, after which the blots were revealed using the ECL system. We routinely used both GAPDH and tubulin as loading controls. Rarely, as was the case for the data presented in Figure 5G, we were unable to effectively blot for β-tubulin. In this case, GAPDH served as the sole loading control.

Statistics.

Statistical analyses performed for specific data sets are described in the figure legends. All statistical tests were performed using GraphPad Prism software v8. Data are presented as mean ± SEM and were subjected to 2-tailed unpaired Student’s t test. For multiple comparisons, unpaired t tests were corrected using the Holm-Sidak method. Significance was assessed using P-value cutoffs indicated as follows: *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001.

GAR was supported by a Wellcome Trust Senior Investigator (WT098424AIA) and Investigator Award (212625/Z/18/Z), by MRC Programme grants (MR/R022259/1, MR/J0003042/1, MR/L020149/1, and MR/R022259/1), an Experimental Challenge Grant (DIVA, MR/L02036X/1), and Project grants from the MRC (MR/N00275X/1), and Diabetes UK (BDA/11/0004210, BDA/15/0005275, and BDA 16/0005485). YD was supported by grants from the Human Islet Research Network (HIRN) and the DON foundation. This project was also supported by NIH/NIDDK grants, including DK108666 to MH, DK103175 and DK112304 to SKK, a Diabetes Research Center P30 grant (DK063720), and a Nutrition and Obesity Research Center P30 grant (DK098722).

Version 2. 08/26/2019

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