Deletion of Carboxypeptidase E in β-Cells Disrupts Proinsulin Processing but Does Not Lead to Spontaneous Development of Diabetes in Mice

Proinsulin is processed into mature insulin and C-peptide by prohormone convertase (PC)1/3 and PC2 and carboxypeptidase E (CPE) (1–3) prior to secretion from pancreatic β-cells. Failure of this process leads to insufficient mature insulin release and onset of hyperglycemia (4–6) and has been observed in diabetes pathogenesis (7,8).

Prohormone processing enzymes are highly expressed in neuroendocrine cells, and subjects with mutations in these genes often display cognitive impairments and obesity. A CPE truncating mutation (c.76_98del) causes morbid obesity and severe hyperglycemia (9), a CPE nonconservative missense mutation (c.847C>T) reduces enzymatic activity associated with early-onset type 2 diabetes (10), and homozygous nonsense CPE mutations (c.405C>A) cause obesity and hypogonadotropic hypogonadism (11). Similarly, Cpe whole-body knockout mice develop spontaneous obesity and behavioral abnormalities (12), and Cpe mutant mice (Cpe^fat/fat) are obese and infertile (13).

To understand the role of Cpe in β-cell function and glucose homeostasis, we generated pancreatic β-cell–specific Cpe knockout mice. We performed biochemical and top-down proteomic analysis to evaluate hormone processing patterns in β-cells. We also analyzed β-cell transcriptomic profiles and performed live-cell imaging analysis to understand whether deficiency of Cpe, and the increased compensatory production of proinsulin, leads to islet dysfunction and dysglycemia. Finally, we tested whether the lack of Cpe in β-cells increases susceptibility to diet- or secretory stress–induced hyperglycemia in mice. Our model provides a useful tool to understand the role of reduced prohormone processing efficiency and increased (pro)insulin translation in β-cells during diabetes development and sheds light on whether impaired prohormone processing in β-cells is a cause of diabetes and obesity in subjects with CPE mutations.

Paraffin-embedded pancreatic tissue sections were obtained from the Network for Pancreatic Organ donors with Diabetes (nPOD) and Alberta Diabetes Institute IsletCore (Supplementary Table 1).

β-Cell–specific Cpe knockout (βCpeKO) and inducible β-cell–specific Cpe knockout (iβCpeKO) mice were generated through crossing the offspring of C57BL/6N-Cpe^{tm1a(EUCOMM)Hmgu}/Ieg mice and Tg(CAG-flpo)1Afst mice with B6(Cg)-Ins1^{tm1.1(cre)Thor}/J or Tg(Pdx1-cre/Esr1) mice. In addition, Gt(ROSA)26Sor^{tm4(ACTB-tdTomato,-EGFP)Luo} reporter mice were bred with βCpeKO mice for β-cell sorting and RNA sequencing. The control mice used for biochemical, imaging, and metabolic experiments are as follows: Ins1^Cre/+;Cpe^fl/+ (βCpeHet) as well as Ins1^+/+;Cpe^fl/fl or Ins1^+/+;Cpe^fl/+ (Wt). For diet studies, 8-week-old mice received either a low-fat diet (LFD) (10% fat), or a high-fat diet (HFD) (45% fat; Research Diets). For secretory stress studies, 10-week-old male mice received saline or streptozotocin (STZ): multiple injections of low-dose STZ (MLD-STZ) (35 mg/kg body wt i.p. daily for 5 days; Sigma-Aldrich). Metabolic assays (such as intraperitoneal glucose tolerance test (IPGTT), insulin tolerance test (ITT), and body mass composition analysis) were performed in a blinded fashion and have previously been described (14). For in vivo β-cell proliferation studies, after tamoxifen-induced Cpe deletion, a 60% fat diet (Research Diets) was given for 48 h (15) in combination with 5-ethynyl-2′-deoxyuridine (40 mg/kg body wt i.p. injection twice daily, EdU; Toronto Research Chemicals) (16). All studies were approved by the Animal Care and Use Committee at the University of British Columbia.

Mouse islets were isolated and cultured as previously described (14). For electron micrograph studies, freshly isolated islets were fixed in 2% glutaraldehyde (pH 7.4) at room temperature, shipped, processed, and imaged by the Electron Microscopy Facility at McMaster University Health Science Centre. PC1/3 and PC2 enzyme activity assays were performed as previously described (17) with a SpectraMax M3 plate reader (Molecular Devices). For respirometry studies, mouse islets were dispersed and analyzed by Seahorse XFe96 Analyzer (Agilent Technologies). For analysis of insulin secretion dynamics, islets were incubated in a perifusion system (Biorep Technologies) with 1.67 mmol/L glucose, 1.67 mmol/L glucose, and 16.7 mmol/L glucose plus 30 mmol/L KCl Krebs-Ringer buffer sequentially. Insulin concentrations in perifusates were analyzed with rodent insulin (ALPCO) and proinsulin (Mercodia) ELISAs. For measurement of glucose uptake, islets were precultured in glucose-free media, dispersed with Accutase (Innovative Cell Technologies), and treated with 2-NBDG (2-deoxy-2-[(7-nitro-2,1,3-benzoxadiazol-4-yl)amino]-d-glucose; Invitrogen) for 5 min, prior to flow cytometry analysis.

For exocytosis studies, dispersed islet β-cells were patch clamped in whole-cell voltage-clamp configuration with use of a HEKA EPC 10 amplifier and PATCHMASTER software (HEKA Electronik, Lambrecht, Germany) as previously described (18). Exocytosis was monitored as increases in cell capacitance, elicited by either a series of 500-ms membrane depolarizations from −70 to 0 mV or increase in the duration of membrane depolarizations. For FACS-sorted β-cell bulk-RNA sequencing experiments, freshly isolated islets were dispersed and GFP⁺ live cells were collected with BD FACSAria Cell Sorter for RNA isolation via an RNeasy Plus Micro Kit (QIAGEN). After quality control analysis with an Agilent 2100 Bioanalyzer, an RNA library was prepared with use of the NeoPrep Library Prep system with TruSeq Stranded mRNA kit (Illumina); RNA sequencing was performed with Illumina NextSeq 500; reads were aligned with TopHat to the reference genome of UCSC Genome Browser mm10, assembled by Cufflinks; and a list of differentially expressed genes was generated via DESeq2. Gene set enrichment analysis, network visualization, and volcano plot were generated via Gene Set Enrichment Analysis (GSEA v4.2.3), Cytoscape (v3.9.1), and EnhancedVolcano (v1.14.0) in RStudio (v1.4.1717).

Islet pellets were homogenized in 8 mol/L urea lysis buffer, reduced, alkylated, quenched, and clarified with tris(2-carboxyethyl)phosphine, iodoacetamide, and dithiothreitol, before 3 kDa molecular weight cutoff filtration. Samples were analyzed with a Waters nanoACQUITY UPLC system with mobile phases consisting of 0.2% formic acid in H2O and 0.2% formic acid in acetonitrile. For tandem mass spectrometry analysis of proteins, the nanoACQUITY UPLC system was coupled to a Thermo Scientific Orbitrap Fusion Lumos mass spectrometer equipped with the FAIMS Pro interface (19). Proteoform identification was performed with TopPIC (v1.4). Downstream data analysis and quantification were performed with use of MSstats (v4.0.1) and TopPICR (v0.0.3) R packages.

Mouse islets were lysed in an NP-40–based buffer and analyzed through reducing or nonreducing Tricine–urea–SDS-PAGE (20), and blotted with use of antibodies listed in Supplementary Table 2, on a LI-COR Biosciences Odyssey Imaging System. For analysis of insulin biosynthesis, islets were preincubated in methionine-free RPMI medium for 90 min and then treated with l-azidohomoalaine (Invitrogen) and 5 or 25 mmol/L glucose Krebs-Ringer buffer for 90 min. Islets were lysed, click labeled with biotin-alkyne (Invitrogen), and immunoprecipitated and the eluted proteins were analyzed on a Tricine–urea–SDS-PAGE system.

Dispersed mouse islet cells were seeded on chamber slides (ibidi or Thermo Fisher Scientific) overnight and cultured in 5 or 25 mmol/L glucose RPMI media for the indicated times. For cell proliferation experiments, EdU was added during the last 24 h of treatment, followed by click labeling and staining (Invitrogen). TUNEL staining was performed according to the manufacturer’s manual (Roche). For live-cell imaging experiments, cells were labeled with CellROX, MitoSOX, tetramethylrhodamine methyl ester (TMRM) (Thermo Fisher Scientific), and MitoTracker Green (MTG) (New England Biolabs) and imaged with use of an SP5II laser scanning confocal microscope (Leica Microsystems). Cpd and proinsulin costained β-cells were imaged on an SP8 X STED (STimulated Emission Depletion) white light laser confocal imaging system. β-Cell area was analyzed with immunohistochemistry staining against insulin with a BX61 microscope (Olympus). All antibodies used are listed in Supplementary Table 2. Image analyses were performed with ImageJ (21), QuPath, ilastik, and CellProfiler pipelines.

Islet mRNA and DNA were isolated with PureLink RNA Micro (Invitrogen) and QIAamp DNA Micro (QIAGEN) kits, and cDNA was synthesized with a SuperScript VILO kit (Invitrogen). mRNA and DNA levels were analyzed with SYBR Green–based quantitative real-time PCR (ViiA 7 Real-Time PCR System; Applied Biosystems). Primer sequences are listed in Supplementary Table 2.

Statistical analyses were performed through GraphPad Prism 9 or R. After normality tests, data sets with normal distribution or with small sample numbers were analyzed using Student t test or ANOVA followed by post hoc analysis. Data with nonnormal distribution were analyzed with the Wilcoxon rank sum test. Statistical significance is indicated in the figures as follows: *P < 0.05. All data are presented as mean ± SEM.

Data and reagents generated in the current study are available from the corresponding author on reasonable request.

CPE is highly expressed in human and mouse islet endocrine cells (Fig. 1A and B). To study the roles of Cpe in β-cells, we generated βCpeKO mice by crossing Ins1^Cre/+ and Cpe^fl/fl mice (Fig. 1C–E). The deletion of Cpe in β-cells leads to near-total loss of mature insulin granules (Fig. 1F) and significantly elevated fasting plasma proinsulin-like immunoreactivity (Fig. 1G and H). As most immunoassays likely cross-react with target peptide with various C- and N-terminal extensions, we decided to analyze the propeptide repertoire with biochemical and proteomic approaches.

Proinsulin is first processed by PC1/3 to form split-32,33 proinsulin with overhanging basic residues. Cpe then removes these basic residues to yield the des-31,32 proinsulin intermediate, which is cleaved by PC2 (or PC1/3) to produce mature insulin following trimming of the remaining basic residues by Cpe (1–3). To study the impact of β-cell Cpe deficiency on proinsulin processing, we analyzed proinsulin forms using a nonreducing SDS-PAGE system. We found that higher-molecular-weight proinsulin forms were increased in βCpeKO islets. The lower bands are likely the combination of mature insulin and insulin with basic residue extensions, as their molecular weights are similar and may not be separated by electrophoresis. (Fig. 2A). Similar to proinsulin, islet amyloid polypeptide (IAPP) is also synthesized as a larger precursor, proIAPP, and is processed by PC1/3, PC2, Cpe, and Pam to form amidated IAPP (14,22–24). Nonamidated IAPP and intermediate proIAPP (proIAPP_1–48) forms are increased in βCpeKO islets, although amidated IAPP levels appear comparable between Wt and βCpeKO islets (Fig. 2B). Proteomic assessment confirmed that intact proinsulin levels are increased in βCpeKO islets, while levels of the mature insulin are reduced (Fig. 2C and D). Full-length proIAPP levels are also increased, while levels of amidated mature IAPP level are not reduced, in βCpeKO islets (Fig. 2E and F).

To determine whether Cpe deletion creates feedback inhibition of peptide hormone maturation, we analyzed prohormone processing enzyme transcript and protein levels. Expression of insulin, IAPP, and processing enzyme transcripts are comparable between βCpeKO and Wt islets (Fig. 2G). ProPC1/3 protein (87 kDa) levels are elevated, as are PC2 and proSAAS (Fig. 2H and I); however, total islet PC1/3-specific activity is not changed (Fig. 2J). PC2-specific activity is reduced in βCpeKO mice (Fig. 2K), which may occur through increased 7B2-mediated inhibition of PC2 activity (25). We also found that Cpe is not the only carboxypeptidase capable of processing peptide hormones in β-cells. Despite near-complete recombination and deletion of Cpe (Fig. 1C–E), mature insulin remains detectable, and mature IAPP is expressed at levels similar to those of Wt islets (Fig. 2B). Carboxypeptidase D (CPD) has been detected in the Golgi network of rodent β-cells (26) and may not be removed from immature insulin granules in conditions such as cargo protein CCDC186 deficiency (27). CPD is expressed in human islet β-cells (Fig. 2L). Increased presence of Cpd in proinsulin-containing organelles such as the Golgi network or immature insulin granules may aid the processing of prohormones in the absence of Cpe, as the colocalization of Cpd and proinsulin is significantly higher in Cpe-deficient β-cells (Fig. 2M and N).

Unlike Cpe whole-body knockout mice or Cpe mutant mice (12,13), 8 week-old βCpeKO mice do not develop early-onset obesity and diabetes (males [Fig. 3A–C] and females [Fig. 3D–F]). Islets from βCpeKO mice displayed insulin secretion dynamics comparable with those of Wt islets (Fig. 3G), speaking against a role for Cpe as a granule-sorting receptor in β-cells. Interestingly, proinsulin was released upon high glucose and KCl stimulation (Fig. 3H), suggesting that the proinsulin-containing granules are likely equipped with appropriate granule contents that allow for efficient granule release. Rates of exocytosis were also comparable between Wt and Cpe-deficient β-cells (Fig. 3I), although exocytosis events proximal to the plasma membrane, measured by a time-train depolarization experiment, were slightly reduced in β-cells from βCpeKO mice (Fig. 3J).

To promote the development of obesity and insulin resistance, we placed 8-week-old βCpeKO mice on a control LFD (10% fat) or HFD (45% fat) for a duration of 6 months. In the LFD-treated group, weight gain of βCpeKO mice was similar to that of their Wt littermates (males [Fig. 4A] and females [Fig. 4F]), suggesting that the lack of Cpe in β-cells does not lead to spontaneous development of obesity. At 20 weeks post-LFD, male βCpeKO mice displayed modestly increased fasting blood glucose levels (Fig. 4B); however, their glucose tolerance, insulin tolerance, and percent fat mass, measured at 16 weeks and 20 weeks post-LFD, remained comparable with those of Wt littermates (Fig. 4C–E). Female βCpeKO mice have slightly elevated fasting blood glucose levels at 4 and 22 weeks post-LFD, yet displayed glucose tolerance, insulin tolerance, and body fat mass similar to those of littermates (Fig. 4G–J). Inappropriate proinsulin processing associated with β-cell Cpe deficiency does not contribute to accelerated development of HFD-induced obesity (males [Fig. 4K] and females [Fig. 4P]). Male and female βCpeKO mice showed fasting blood glucose levels, glucose tolerance, insulin tolerance, and percent fat mass similar to those of their littermates (Fig. 4L–O and Q–T).

Despite comparable glucose tolerance, β-cell area in LFD-treated βCpeKO male and female mice was elevated (Fig. 5A and B). Although HFD promoted compensatory β-cell expansion in Wt mice, βCpeKO mice did not have an increase in β-cell area (Fig. 5C and D). To study the cause of increased β-cell area in βCpeKO mice on LFD, we first examined the β-cell proliferation rate by analyzing the frequency of Ki67⁺ β-cells. However, the number of proliferating β-cells in mice fed LFD for 6 months was too low to allow for appropriate comparison between βCpeKO and Wt mice (data not shown). We therefore isolated islets from 10-week-old mice, cultured them in 5 or 20 mmol/L glucose media for 72 h, and analyzed EdU incorporation in insulin⁺ β-cells. β-Cell proliferation was significantly elevated in islets from βCpeKO mice (Fig. 5E). We also generated iβCpeKO mice by crossing Pdx1-Cre^ER mice with Cpe^flox/flox mice (Fig. 5F). Shortly after oral tamoxifen administration, male, but not female, iβCpeKO mice become mildly glucose intolerant (Fig. 5G and H). To induce β-cell proliferation, we treated iβCpeKO and their Wt littermates with a 60% fat diet for 2 days and analyzed EdU incorporation in the mouse pancreas. We found that the β-cell proliferation rate was significantly elevated in iβCpeKO female mice (Fig. 5I [males] and Fig. 5J [females]).

To identify the underlying molecular mechanisms contributing to increased β-cell proliferation in βCpeKO mice, we performed transcriptomic analysis in sorted β-cells from Wt and βCpeKO mice (Fig. 6A). As expected, Cpe was drastically reduced in β-cells from βCpeKO mice. Expression of many Hif1α-regulated genes (including Ldha, Hmox1, P4ha1, Pgk1, Mif, Ak4, Bnip3, Pfkp, P4ha2, Slc2a1, and Pfkfb3) was increased. Gene ontology analysis showed that expression of genes related to glycolysis and hypoxia are enriched, while hallmarks of pancreatic β-cells are reduced, in Cpe-deficient mouse β-cells (Fig. 6B). We found that the rate of glucose uptake into islet cells is comparable in βCpeKO and Wt mice (Fig. 6C), and glycolytic flux analysis showed that βCpeKO islet cells have an oxygen consumption rate similar to that of Wt islet cells (Fig. 6D). Interestingly, despite elevated (pro)insulin production (Fig. 6E) and increased accumulation of proinsulin oligomers (28) (Fig. 6F), (pro)insulin protein stability was not significantly reduced in islets from βCpeKO mice (Supplementary Fig. 1), and canonical unfolded protein response elements were not elevated in freshly isolated βCpeKO islets (Supplementary Fig. 2A–E).

Because (pro)insulin production is elevated in βCpeKO islets, we asked whether the morphology or function of the fuel-providing mitochondria is altered in Cpe-deficient β-cells. Electron micrograph analysis of mitochondrial images showed that βCpeKO β-cells have similar area, yet reduced size (Fig. 7A–C). Additional confocal image analysis showed mitochondrial number, area, perimeter, and branch number were all reduced in βCpeKO β-cells (Fig. 7D–H), while mtDNA content was not reduced (Fig. 7I). This suggests that mitochondria in Cpe-deficient β-cells work to accommodate an increased demand for (pro)insulin synthesis. Nevertheless, after prolonged glucose treatment, β-cells from βCpeKO mice had reduced mitochondrial membrane potential upon high glucose stimulation (Fig. 7J) and displayed elevated levels of mitochondrial and cellular reactive oxygen species (ROS) (Fig. 7K and L). β-Cells from βCpeKO mice have no increase in mitochondria biogenesis upon high glucose culture, as Pgc1a transcript levels are not significantly elevated (Fig. 7M). Islets from βCpeKO mice failed to display elevated MafA transcript level upon high glucose treatment (Fig. 7N). Rather, Aldh1a3 transcript levels were significantly elevated, suggesting loss of β-cell identity in βCpeKO islets (Fig. 7O). Glucose metabolism was likely altered, as transcript levels of Pfkp were significantly increased in βCpeKO islets (Fig. 7P). Of note, treatment of islets with high glucose led to increased expression of endoplasmic reticulum (ER) stress markers such as spliced Xbp1 (Xbp1s), but βCpeKO islets showed no increase in Xbp1s (Fig. 7Q), inferring that elevated proinsulin biosynthesis does not contribute to increased ER stress in Cpe-deficient β-cells.

We administered βCpeKO and littermate mice with MLD-STZ to induce cell dysfunction in a small portion of β-cells and to create secretory stress in the remaining cells (Fig. 8A). βCpeKO mice showed higher blood glucose levels at 10 days after the last STZ treatment, compared with Cpe heterozygous (βCpeHet) or Wt mice (Fig. 8B–D). βCpeKO mice did not display increased STZ-induced β-cell death: β-cell area (analyzed at 10 days post-STZ) and number of TUNEL⁺ β-cells (analyzed at 3 days post-STZ) were similar in βCpeKO and Wt mice (Fig. 8E and F). Building on our finding of increased β-cell area in βCpeKO mice (Fig. 5A), we found that buffer-treated βCpeKO mice had a higher percentage of β-cells in their islets (Fig. 8G). Although MLD-STZ led to a modest increase in the percentage of β-cells in Wt islets, it reduced the percentage of β-cells in βCpeKO islets (Fig. 8G). In agreement with previous reports, we showed that the portion of Glut2⁺ β-cells was reduced after MLD-STZ (Fig. 8H); however, the number of Glut2⁺ β-cells remained comparable in Wt and βCpeKO mice. We also found an increased percentage of Aldh1a3⁺ β-cells and increased ER stress markers in the islet cells upon MLD-STZ treatment (Fig. 8I and Supplementary Fig. 2F). However, the deficiency of Cpe in β-cells did not cause further elevation of ER stress, as immunofluorescence intensity of phosphorylated eIF2α, Atf4, and Bip was comparable between MLD-STZ Wt and βCpeKO mice (Supplementary Fig. 2). As the insulin antibody used for immunostaining recognizes both proinsulin and mature insulin, the increased mean fluorescence intensity observed in β-cells from buffer-treated βCpeKO versus Wt mice likely reflects the significantly elevated proinsulin protein expression in βCpeKO islets (Fig. 8J). Upon STZ treatment, insulin and Glut2 expression levels were significantly reduced in βCpeKO, but not Wt, mice (Fig. 8J and K), suggesting that β-cells in βCpeKO are more susceptible to secretory stress–induced degranulation and dysfunction. β-Cell maturity remained comparable, as islet Aldh1a3 expression levels were not further increased in STZ-treated βCpeKO mice (Fig. 8L).

Cpe mutations in mice and humans lead to obesity and hyperglycemia; however, the underlying cellular and physiological mechanisms remain unknown. We hypothesized that lack of Cpe in pancreatic β-cells is the main contributor to such clinical phenotypes because 1) Cpe is required for proper proinsulin processing (3), 2) reduced expression of Cpe is associated with β-cell dysfunction in multiple experimental models of diabetes (29–31), and 3) Cpe may play a protective role in preventing β-cell death (32). To address this hypothesis, we generated β-cell–specific Cpe knockout mice. Our data indicate that while Cpe is important in normal proinsulin processing, Cpe deficiency alone does not contribute to obesity or cause marked dysglycemia.

Islets from βCpeKO mice contain markedly more proinsulin peptides, yet have detectable mature insulin peptide, suggesting that another carboxypeptidase, likely Cpd, is compensating. In agreement with an in vitro study suggesting that Cpe is essential for PC2-mediated peptide processing (25), we showed that loss of Cpe results in reduced PC2 enzyme activity and increased N-terminally extended proIAPP (which is normally processed by PC2 into mature IAPP [23]). Although total islet PC1/3 enzyme activity was not changed in βCpeKO mice, PC1/3 protein levels were elevated, suggesting that on a per-enzyme basis, PC1/3 activity is likely reduced in βCpeKO β-cells. To our surprise, even with markedly impaired proinsulin processing and diminished output of mature insulin, βCpeKO mice failed to develop obesity and hyperglycemia spontaneously or when challenged with an HFD, contrary to a recent report that Pdx-Cre^ERT-mediated Pcsk1 deletion and elevated proinsulin promote the development of obesity in mice (33). It is plausible that elevated proinsulin, possessing 5% activity compared with insulin (34,35), is sufficient to maintain glucose homeostasis. Another possibility is that obesity and overt hyperglycemia observed in Cpe mutations are driven by insufficient Cpe and defective neuropeptide processing in other tissues, such as in the hypothalamus, although mice with Cpe deletion in proopiomelanocortin (POMC)-expressing neurons do not become obese (36). Whether Cpe controls body weight and metabolic homeostasis in non-POMC-expressing neuroendocrine cells remains to be tested.

β-Cells are able to adapt to increased insulin demand by increasing production, secretion, and mass prior to hyperglycemia onset (37–39). βCpeKO mice have increased proinsulin production and elevated β-cell area but remain normoglycemic. Because protein overproduction may change intrinsic metabolic pathways and alter β-cell fate (40), it is plausible that increased demand caused by increased insulin production (41) contributes to metabolic pathway rewiring and concomitant β-cell proliferation in βCpeKO mice. In a recent large-scale small molecule screen a compound was identified that promotes protein synthesis and β-cell regeneration. The authors showed that the increased β-cell regeneration is associated with hypo-translation of mRNAs that are integral to mitochondrial-related processes (42). In support of this idea, we observed altered glycolytic gene signatures and changes in mitochondrial morphology and membrane potential in βCpeKO β-cells. Oxygen consumption rate was not reduced in islets from βCpeKO mice, hinting that additional pathological stimuli are likely needed to disrupt oxidative phosphorylation. Alternatively, metabolic flux analysis may offer more quantitative insights into carbon metabolism and energy flow in islets with inherently elevated proinsulin biosynthesis. It is also possible that the increased proinsulin oxidative folding burden may create ER redox imbalances (43), leading to increased mitochondrial and cellular ROS levels. Future live-cell imaging experiments with ROS biosensors could illuminate the cellular sequence of events. Both mild ER stress and ROS have been reported to facilitate β-cell proliferation (16,44). Despite an accumulation of proinsulin oligomers (28), we failed to detect significant changes in transcripts encoding ER chaperons or unfolded protein response proteins. Of note, β-cell de-differentiation markers Serpina7 (45) and Ldha (46) are reduced in sorted β-cells from normoglycemic βCpeKO mice, suggesting that the slight loss of β-cell identity may occur during early-stage β-cell compensation prior to the development of hyperglycemia. Whether insulin biosynthesis impacts insulin granule secretion requires further investigation. We speculated that increased proinsulin biosynthesis burden and altered glucose metabolism or redox handling capacity may contribute to defects in coupling or translocation of granules to the site of Ca²⁺ channels, which results in reduced exocytotic response of βCpeKO β-cells to a train of membrane depolarizations. Alternatively, increased insulin production may affect the composition of the secretory granule membrane (47), which alters its interaction with plasma membrane Ca²⁺ channels or the fusion with plasma membrane (48). It is also worth mentioning that we have not analyzed possible changes in paracrine signaling in βCpeKO islets.

βCpeKO mice adapted to chronic dietary stress weight gain and glucose tolerance similar to those of their littermates. We administered βCpeKO mice and their littermates with MLD-STZ to induce acute insulin secretory stress without extensive β-cell death or loss of β-cell mass. MLD-STZ led to loss of β-cell identity in both Wt and βCpeKO mice, evidenced by an increased percentage of Aldh1a3⁺ cells and reduced Glut2⁺ cells in islets. After MLD-STZ, βCpeKO mice also had accelerated development of hyperglycemia and displayed reduced (pro)insulin and Glut2 expression levels in β-cells. These findings were mirrored in vitro, as we observed no induction of MafA, and increased Aldh1a3, in high glucose–treated βCpeKO islets. We speculate that suboptimal mitochondrial function, or altered cellular redox homeostasis, resulting from the combination of secretory stress and Cpe deficiency, contributes to β-cell dysfunction in MLD-STZ Cpe-deficient islets (49–51). It has been reported that islets from Cpe mutant mice are more susceptible to palmitic acid–induced β-cell apoptosis (32). We were unable to observe detectable differences in TUNEL⁺ β-cells in MLD-STZ–treated βCpeKO 3 days after the last STZ injection, when β-cell apoptosis rates are at their highest (52). Instead, β-cells from βCpeKO mice have reduced Glut2 expression, which may present an adaptive mechanism to attenuate glucose uptake and metabolic stress–induced β-cell death (53).

In summary, we demonstrated that loss of Cpe in pancreatic β-cells does not contribute to spontaneous development of obesity and hyperglycemia in mice. However, elevated proinsulin output likely reshapes β-cell glucose metabolism and increases its susceptibility to secretory stress–induced dysfunction and diabetes. Our model may shed light on β-cell translational adaptation, which likely occurs early during the development of diabetes. Additional studies in other prediabetes models and human islets are needed for a better understanding of these early adaptive events and will aid discovery of new therapeutic targets to preserve β-cell function prior to the onset of diabetes.

Acknowledgments. The authors thank Dr. Paul Orban (University of British Columbia) for helpful comments on the project, Dr. Iris Lindberg (University of Maryland Medical Center) for providing the enzyme activity assay protocol and reagents, Dr. Rohit Sharma (University of Massachusetts Amherst) and Dr. Aaron Cox (Baylor College of Medicine) for providing suggestions on β-cell proliferation experiments, Mr. Daniel Pausula (University of British Columbia) for providing protocol for live cell imaging experiments, Drs. Lei Dei and Galina Soukhatcheva (University of British Columbia) for technical assistance, and Dr. Elizabeth Simpson (University of British Columbia) for help with mouse rederivation.

Funding. This work is supported by JDRF (advanced postdoctoral fellowship 3-APF-2022-1141-A-N to Y.-C.C.), Canadian Institutes of Health Research (grant PJT-153156 to C.B.V.), BC Children’s Hospital Foundation and BC Children’s Hospital Research Institute (Canucks for Kids Fund Childhood Diabetes Laboratories Summer Studentship to K.L.C.W.), Natural Sciences and Engineering Research Council of Canada (NSERC) grant RGPIN-2020-05390946 and Michael Smith Health Research BC Scholar Award to R.I.K.-G., and National Institutes of Health grants R01DK122160 and U01DK124020 to W.-J.Q. Mass spectrometry proteomics experiments were performed in the Environmental Molecular Sciences Laboratory, Pacific Northwest National Laboratory, a national scientific user facility sponsored by the Department of Energy under contract DE-AC05-76RL0 1830.

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. Y.-C.C. contributed to study conceptualization, investigation, formal analysis, and visualization and wrote the manuscript. A.J.T. contributed to study conceptualization and investigation and edited the manuscript. J.M.F., X.-Q.D., and M.K. contributed to study methodology, investigation, and formal analysis. K.L.C.W. contributed to study investigation and formal analysis and edited the manuscript. K.F. and A.E.P. contributed to study investigation. A.C.S. contributed to study investigation and methodology and edited the manuscript. R.I.K.-G. contributed to study methodology and formal analysis. P.E.M. and W.-J.Q. contributed to study investigation and methodology. C.B.V. contributed to study conceptualization and investigation and reviewed and edited the manuscript. All authors approved the final version of the manuscript. C.B.V. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.

Prior Presentation. Parts of this study were presented in abstract form at the 55th Annual Meeting of the European Association for the Study of Diabetes, Barcelona, Spain, 16–20 September 2019; the 58th Annual Meeting of the European Association for the Study of Diabetes, 19–23 September 2022, Stockholm, Sweden; and the Gordon Research Conference on Protein Processing, Trafficking and Secretion, New London, NH, 17–22 July 2022.