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. Author manuscript; available in PMC: 2026 Feb 18.
Published in final edited form as: Am J Physiol Endocrinol Metab. 2025 Dec 15;330(2):E152–E160. doi: 10.1152/ajpendo.00342.2025

Ppid is Necessary for Overnutrition-Induced β-cell Loss

Brittney A Covington 1, Zihan Tang 1, Lisette Maddison 1, Bingyuan Yang 1, Wenbiao Chen 1
PMCID: PMC12911484  NIHMSID: NIHMS2130868  PMID: 41397247

Abstract

Type 2 diabetes (T2D) involves progressive loss of functional β-cell mass. In a zebrafish insulin-resistant model (zMIR), overnutrition triggers islet inflammation and nocturnal β-cell death. The cell death is prevented by the cyclophilin D (Ppid) inhibitor, cyclosporin A (CsA). Reducing mitochondrial ROS with mito-TEMPO or mitochondrial calcium with Ru360 protects β cells, further implicating the mitochondrial permeability transition pore (mPTP) in β-cell loss. The timing of β-cell death coincides with lower mitochondrial antioxidant gene expression, indicating nocturnal mitochondrial vulnerability. Global ppid−/− preserves β-cell mass without altering islet inflammation or macrophage recruitment. Conversely, β-cell-specific PPID re-expression restores—and exacerbates—β-cell loss, which remains CsA-sensitive. These findings identify Ppid as a β-cell-intrinsic mediator of overnutrition-induced β-cell loss.

New & Noteworthy:

This study provides the first evidence that Ppid-mediated opening of the mPTP is necessary for overnutrition-induced β-cell death in zebrafish. Pharmacological inhibition of mPTP opening protected β cells. Global knockout of ppid prevented β-cell loss, which was reversed by transgenic PPID re-expression in the β cells. The time of β-cell death coincides with lower antioxidant gene expression at night. Thus, relieving mitochondrial stress at night may preserve β cells.

Graphical Abstract

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Introduction

Type 2 diabetes (T2D) affects over 500 million people worldwide (1). Alarmingly, T2D diagnosis in adolescents has significantly increased over the past two decades, driven in part by obesity and sedentary behavior (2,3). Unfortunately, T2D in adolescents is associated with a higher risk and earlier onset of chronic diseases, including hypertension, nephropathy, cardiovascular disease, and non-alcoholic fatty liver disease (4). Elucidating the mechanisms leading to T2D pathogenesis in adolescents is essential to prevent or delay this debilitating chronic disease for future generations.

A hallmark of T2D pathogenesis is the progressive loss of functional β-cell mass, which impairs insulin secretion and glucose homeostasis (5,6). Insulin resistance from obesity and puberty increases β-cell workload, driving compensatory insulin secretion (7–9). The heightened demand triggers stress in the endoplasmic reticulum (ER), where immature insulin undergoes a series of post-translational modifications (10,11). The amplified workload also elevates oxidative stress within β cells (12,13). When the adaptive capacity of β cells exceeded, dysfunction and death ensue, leading to diabetes (14,15).

To investigate mechanisms of T2D pathogenesis, we developed a zebrafish muscle-specific insulin resistance (zMIR) model (16). Insulin resistance arises from transgenic expression of a dominant-negative IGF-1 receptor in skeletal muscle (17). Under normal feeding, zMIR fish display increased β-cell mass in the first 6 months, followed by a gradual mass decline and impaired glucose tolerance (16). However When challenged with a high-fat diet, zMIR larvae undergo rapid compensation and decompensation, with significant β-cell loss after 3 days of overnutrition (18). This accelerated disease progression in a short developmental window may model aspects of adolescent T2D, where puberty-induced insulin resistance accelerates β-cell failure (2,3,16). Although the model aligns with features of adolescent metabolic stress, further comparative studies are needed to fully establish its translational relevance.

We previously discovered a sequential inflammatory cascade preceding β-cell loss in this model (18,19): 1) ER stress activates Ripk3; 2) Ripk3-mediated induction of il1b; 3) Il1b-driven recruitment of macrophages and Tnfa secretion; 4) Tnfa-dependent cxcl8a induction in β cells; 5) β-cell loss (18,19). Disruption of any step prevents β-cell death, underscoring the cascade’s essential role.

Several mechanisms contribute to β-cell loss in T2D, including dedifferentiation, loss of identity, senescence, and cell death (5,20,21). As T2D is heterogeneous, the contribution of each likely varies across individuals (22). Because dying β cells are quickly removed, its contribution may be underestimated (23,24). Nevertheless, increased β-cell death has been reported in diabetic animal models and organ donors (5,25). However, the molecular pathways of β-cell death in vivo have not been fully characterized. Identifying the pathway(s) driving cell death is critical for developing interventions aimed at preserving functional β-cell mass.

Mitochondria are central to β-cell function, coupling glucose metabolism to insulin secretion via ATP production (6). Conditions of metabolic stress can lead to mitochondrial dysfunction(26,27), and subsequent ROS generation, calcium overload, and disrupted ATP production which contribute to β-cell dysfunction and death (28). A key mediator of mitochondrial-induced cell death is the mitochondrial permeability transition pore (mPTP). Its prolonged opening, regulated by Cyclophilin D (PPID gene) and triggered by ROS and Ca2+ overload, causes membrane depolarization and loss of bioenergetic function (27,29). While mPTP-dependent cell death is well-characterized in other tissues, its role in overnutrition-induced β-cell death T2D remains incompletely defined.

In the zMIR model, we have excluded apoptosis and canonical necroptosis, as neither pan-caspase (zVAD) nor MLKL (NSA) inhibition prevented β-cell loss (18). Here, we identified the mPTP as a necessary mediator of overnutrition-induced β-cell death. Reducing mitochondrial ROS or Ca2+ and pharmacological or genetic inactivation of Ppid all diminish β-cell death. Conversely, transgenic expression of PPID in ppid-deficient fish exacerbates β-cell death. Thus, β-cell loss results from mPTP-mediated cell death.

Materials and Methods

Zebrafish Husbandry and Lines:

Zebrafish were maintained on a 14:10 h light/dark cycle at 28.5°C. Embryos were kept under the same conditions. β cells were visualized using Tg(ins:H2B-mCherry), and macrophages using Tg(mpeg1:EGFP). β-cell–specific PPID rescue lines were generated using Tol2 transgenesis, and ppid mutants were created by CRISPR–Cas9 as described (31).

Overnutrition model:

Overnutrition was induced by culturing larvae in 5% chicken egg yolk emulsion for 8 h daily, followed by 16 h in 0.3× Danieau buffer, as previously described (32,33).

Drug Treatments:

Compounds were added directly to 0.3× Danieau buffer. Larvae were treated from 56–72 hpo with: MitoTEMPO (10 μM), Ru360 (500 nM), Cyclosporin A (CsA; 10 μM)

Islet Isolation:

Larvae expressing Tg(ins:H2B-mCherry) were euthanized in ice water and digested in HBSS containing 1.5 mg/mL collagenase. Digests were quenched with HBSS + 30% FCS, and individual islets were manually isolated using a fluorescent stereomicroscope.

Glucose Tolerance and Whole-Body Glucose:

Larvae were exposed to 30 mM glucose for 20 min at 28°C, rinsed, and collected at 0, 30, 60, and 120 minutes thereafter. Whole-larval glucose was quantified using the Amplex Red Glucose/Glucose Oxidase Assay Kit (Thermo Fisher). Pools of 10–15 larvae were homogenized (10 μL buffer per fish), centrifuged, and 50 μL supernatant was used per reaction.

qRT-PCR:

RNA from pooled larvae or isolated islets was extracted with TRIzol and reverse-transcribed using the PrimeScript RT kit (Takara). qPCR was performed with SYBR Green (Bio-Rad or MCE) on a Bio-Rad CFX96. Expression was analyzed by the ΔΔCt method and normalized to β-actin or ef1a.

Live Imaging and β-Cell Quantification:

Larvae were anesthetized in tricaine and mounted in 1.5% low-melting agarose. Imaging was performed on a Zeiss LSM 880 confocal microscope using 20×–63× objectives. β-cell nuclei (Tg(ins:H2B-mCherry)) and macrophages (Tg(mpeg1:EGFP)) were imaged using 561-nm and 488-nm lasers, respectively. β-cell number in the principal islet was manually counted. Images were processed using Zen Blue and Imaris.

Statistics:

Unpaired two-tailed t-tests were used for two-group comparisons; one-way ANOVA with Tukey’s post hoc test was used for multi-group analyses. Data are presented as mean ± SEM. Significance was set at P ≤ 0.05. Analyses were performed using GraphPad Prism.

Study approval:

All animal studies were approved by the Vanderbilt Institutional Animal Care and Use Committee.

Results

Pharmacologic inhibition of Ppid prevents β-cell loss in a zebrafish model of overnutrition-induced stress

In the diabetes-prone zebrafish model, zMIR, larval fish exhibit β-cell loss following a 3-day overnutrition challenge in a 5% egg yolk solution for 8 hours per day (Fig. 1A). This loss is specific to zMIR fish and is not observed in wild-type controls. β-cell loss occurs between 56 and 72 hours post overnutrition (hpo), during the nocturnal phase of the circadian cycle (Fig. 1B). In this model, β cells are labeled with H2B-mCherry, a long-lived nuclear fluorescent protein that is used for lineage tracing (14,35), confirming that observed cell loss reflects true depletion of cells rather than dedifferentiation or identity switching (Fig. 1C).

Figure 1: Pharmacologic inhibition of ppid prevents β-cell loss in an overnutrition model.

Figure 1:

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(A) Schematic of the overnutrition paradigm (5% egg yolk, 8 h/day for 3 days starting at 6 dpf).(B) β-cell number in zMIR and WT larvae at 48, 56, and 72 hpo; Data represent ± SEM (≥ 6 per group) (C) Representative images of β cells in zMIR and WT larvae at 56 and 72 hpo (Tg(ins:H2B-mCherry)). (D–E) β-cell number at 72 hpo in zMIR larvae treated with DMSO, CsA, Ru360, or mitoTEMPO. Data represent ± SEM (≥ 7 per group) (F–H) Whole-body expression of antioxidant genes (sod2, txn2, gpx1a) across ZT4–ZT22 in 6 dpf zMIR larvae. Data represent ± SEM (≥ 5 per group) (I–K) Islet expression of sod2, txn2, and gpx1a at ZT4 and ZT16 during overnutrition in 9 dpf larvae. Data represent ± SEM (≥ 3 per group). (L) ppid expression in adult islets at ZT4 and ZT16. Data represent ± SEM (≥ 3 per group). Statistics: Mean ± SEM; two-way ANOVA with Tukey’s test for multiple groups or unpaired Student’s t-test for comparison of 2 groups; ****p < 0.0001, ***p < 0.001, **p < 0.01, *p < 0.05, ns p > 0.05.

To test the role of mPTP-dependent cell death, we treated larvae with cyclosporin A (CsA), an inhibitor of the mPTP regulator cyclophilin d. CsA significantly preserved β-cell number at 72 hpo (Fig. 1D). CsA can also inhibit calcineurin via cyclophilin A (37). Although calcineurin inhibition can enhance endocrine progenitor proliferation and promote β-cell regeneration 7 days after injury (38), CsA treatment for 16 hours did not increase β-cell number in wild-type controls under overnutrition conditions (Fig. 1D).

Ppid is an essential regulator of mitochondrial permeability transition pore (mPTP) opening, which is triggered by mitochondrial calcium overload and elevated reactive oxygen species (ROS) (39,40). While transient mPTP opening regulates mitochondrial activity physiologically, prolonged opening is a well-characterized mechanism of regulated necrosis (27). To validate a role for mPTP in β-cell loss, we treated fish with 10 μM of mito-TEMPO (Fig. 1E), a mitochondrially-targeted ROS scavenger, or 500 nM Ru360 an inhibitor of the mitochondrial calcium uniporter (MCU), and observed protection against β-cell loss at 72 hpo, supporting a role for mPTP activation in β-cell loss in this model.

As overnutrition increases β cell metabolic load and is expected to elevate mitochondrial ROS (41), we assessed expression of the mitochondrial antioxidant genes, gpx1a, sod2 and txn2 in 6 days post-fertilization (dpf) zebrafish across ZT4, ZT10, ZT16 and ZT22 (12pm, 6pm, 12am, and 6am). Sod2 serves as the first line of defense against mitochondria-derived ROS, catalyzing the dismutation of superoxide into hydrogen peroxide (42). Gpx1a reduces hydrogen peroxide to water using glutathione (GSH) (43). Txn2 reduces disulfide bonds and detoxifies peroxides through the mitochondrial thioredoxin system (44). These antioxidant enzymes play a critical role in mitigating mitochondrial ROS and maintaining redox balance (45). Whole-body expression levels of sod2 and txn2 was elevated during the day and decreased at night (Fig. 1F,G), while gpx1a expression trended lower at ZT16 (12am) but was not significantly decreased (Fig 1.H).

Because whole-body gene expression may mask minute islet-specific changes, we examined antioxidant gene expression in isolated islets during overnutrition. Larval islet gpx1a again did not have a significant difference in expression from ZT4-ZT16 (12 pm to 12 am) when exposed to overnutrition feeding (Fig 1K). Notably, both sod2 and txn2 were significantly lower at 12 am (64 hpo), suggesting a potential nocturnal window of diminished islet antioxidant capacity coinciding with β-cell loss (Fig. 1I, J). Interestingly, ppid mRNA levels were more than tenfold higher at ZT16 (12 a.m.) than ZT4 in adult zMIR islets (Fig. 1L).

Genetic knockout of ppid prevents overnutrition-induced β-cell loss.

To confirm the role of Ppid in β-cell loss in overnutrition-induced β-cell loss, we generated a ppid mutation deleting 17 bp using CRISPR/Cas9. This frameshift mutation significantly reduced ppid mRNA as measured by RT-qPCR (Fig. 2A), likely due to nonsense-mediated mRNA decay. The zMIR, ppid−/− fish were viable, exhibiting no differences in survival percentage, β-cell number or whole-body larval glucose at 6 dpf compared to control larvae (Fig. 2B–D).

Figure 2: Genetic knockout of ppid results in normal development and prevents overnutrition-induced β-cell loss.

Figure 2:

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(A) ppid mRNA expression at ZT0 in zMIR and zMIR ppid−/− larvae. Data represent ± SEM (≥ 4 per group). (B–D) Percentage survival, β-cell number and relative glucose at 6 dpf in zMIR or zMIR ppid −/− larvae. Data represent ± SEM (≥ 4 per group). (E-G) yolk area in foregut, IgY levels, and body length at 56 hpo in zMIR or zMIR ppid −/− larvae. ± SEM (≥ 3 per group). (H) Representative whole-larva images at 48 and 56 hpo. Scale bar 500 microns (I) β-cell number at 56 and 72 hpo; zMIR ppid−/− larvae show protection at 72 hpo. Data represent ± SEM (≥ 9 per group) (J) Representative β-cell images Tg(ins:H2B-mCherry). Scale bar 10 microns. Statistics: Mean ± SEM; two-way ANOVA with Tukey’s test for multiple groups or unpaired Student’s t-test for comparison of 2 groups; ***p < 0.001, *p < 0.05, ns p > 0.05.

Feeding behavior also appeared normal in zMIR, ppid−/− larvae. Foregut yolk accumulation could be visualized through the translucent body wall of post-mortem zebrafish from 48 hpo (fasted) to 56 hpo (fed) (Fig. 2H) and did not differ between zMIR, ppid−/− and zMIR controls at 56 hpo (Fig. 2E). To quantify egg yolk ingestion whole-larval IgY levels were measured by ELISA. IgY is a maternal immunoglobulin that is specifically deposited in chicken egg yolk to provide passive immunity to the developing embryo. There were no differences in IgY accumulation between zMIR, ppid−/− and zMIR control groups (Fig. 2G). Length was also consistent between genotypes at 56 hpo (Fig. 2G), supporting a normal feeding behavior in ppid −/− fish (Fig. 2H). Loss of ppid in zMIR larvae prevented overnutrition-induced β-cell loss (Fig. 2I, J).

β-cell ppid is essential for overnutrition-induced β-cell loss in zMIR fish.

Because ppid is ubiquitously expressed and prior pharmacological and genetic manipulations inhibit Ppid function globally, we generated a β cell-specific rescue line expressing human PPID under the insulin promoter using Tol2 transgenesis. This line, referred to as zMIR, ppid−/− βR showed human PPID expression exclusively in the islets and not in the skeletal muscle nor was PPID observed in ppid −/− controls (Fig. 3A). These zMIR, ppid−/− βR larvae develop normally, with no difference in β-cell number or glucose levels at 6 dpf (Fig. 3B,C).

Figure 3: Β-cell ppid is essential for overnutrition-induced β-cell loss in zMIR fish.

Figure 3:

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(A) PPID expression in zMIR ppid−/− and zMIR, ppid βR larvae. Data represent ± SEM (≥ 2 per group). (B–C) β-cell number and relative glucose/larvae at 6 dpf in zMIR and zMIR, ppid−/− βR larvae. Data represent ± SEM (≥ 4 per group). (D) β-cell number at 56 and 72 hpo in zMIR and zMIR, ppid−/− βR larvae. Data represent ± SEM (≥ 9 per group). (E) Representative β-cell images at 56 and 72 hpo. β cells labeled with Tg(ins:H2B-mCherry). Scale bar 20 μm (F) CsA treatment in zMIR ppid−/− βR larvae. Data represent ± SEM (≥ 9 per group). Statistics: Mean ± SEM; two-way ANOVA with Tukey’s test for multiple groups or unpaired Student’s t-test for comparison of 2 groups; ****p < 0.0001, **p < 0.01, *p < 0.05, ns p > 0.05.

β-cell-specific rescue of PPID function restored overnutrition-induced β-cell loss (Fig. 3D, E). Interestingly, the loss was significantly higher than in the zMIR fish (Fig 3D), likely due to over-expression of PPID from the insulin promoter. Notably, treatment with 10 μM CsA, prevented β-cell loss in the zMIR, ppid−/− βR fish (Fig. 3F). These results support a cell-autonomous role for ppid in nutrient-induced β-cell loss.

Loss of ppid does not alter islet inflammation or macrophage recruitment to the islet

β-cell loss is initiated by ER stress-triggered islet inflammation in our zMIR overnutrition model (18). We performed RT-qPCR on isolated islets from zMIR and zMIR, ppid −/− larvae. Expression levels of the ER stress markers chop and edem were not significantly different in the zMIR, ppid−/− compared to the control zMIR larvae (Fig. 4A). Likewise, the pro-inflammatory cytokines il1b, il8a, il6, and tnfa remained unchanged between groups (Fig. 4B).

Figure 4: Loss of ppid does not alter islet inflammation or macrophage recruitment to the islet.

Figure 4:

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(A–B) Islet expression of ER-stress markers (chop, edem) and inflammatory genes (il1b, il6, cxcl8a, tnfa) at 64 hpo. Data represent ± SEM (≥ 3 per group). (C) Intra-islet macrophage number at 65–66 hpo in zMIR and zMIR ppid−/− larvae. (D) Representative macrophage, Tg(mpeg:EGFP), and β cell, Tg(insulin: H2B-mCherry), images (E) Glucose tolerance at 9 dpf after overnutrition in zMIR and zMIR ppid−/− larvae. Statistics: Mean ± SEM; two-way ANOVA with Tukey’s test for multiple groups or unpaired Student’s t-test for comparison of 2 groups, **p < 0.01, *p < 0.05, ns p > 0.05.

Macrophage recruitment to the islet is essential for β-cell loss (19). Live imaging at 64 hpo showed no differences in macrophage recruitment to the islet in zMIR, ppid−/− versus controls, suggesting that ppid does not alter macrophage infiltration into the islet (Fig. 4C, D).

To determine β-cell function in the ppid−/− background, we performed a glucose tolerance test at 72 hpo. zMIR, ppid−/− larvae had lower baseline glucose than zMIR controls (Fig. 4E). Although glucose levels were similar at 30 minutes post exposure, the zMIR, ppid−/− larvae displayed significantly lower glucose at 90 minutes, indicating faster glucose disposal. At 120 minutes post exposure, both groups had come back down to baseline measurements (Fig. 4F). These results suggest that zMIR, ppid−/− larvae have improved glucose tolerance and enhanced glucose disposal capacity.

Discussion

Here, we identify cyclophilin D (Ppid) as a critical mediator of β-cell loss in a zebrafish model of overnutrition-induced diabetes. Inhibition of mPTP opening, either pharmacologically with CsA or by reducing mitochondrial ROS and calcium, preserved β-cell number. Consistently, global knockout of ppid prevented β-cell loss. β-cell-specific re-expression of human PPID in global ppid−/− fish restored and even exacerbated β-cell loss, confirming that Ppid acts cell-autonomously in β cells to promote mPTP-mediated death.

Despite its role in β-cell death, ppid deficiency did not alter islet inflammation. Macrophage recruitment and expression of inflammatory cytokines (il1b, cxcl8a, tnfa) and ER stress markers (chop, edem) were unchanged, indicating Ppid functions downstream of inflammatory signaling as an executioner of β-cell death rather than a modulator of immune activity.

Metabolically, zMIR ppid−/− larvae showed improved glucose tolerance with lower baseline glucose and improved glucose disposal, consistent with preserved β-cell function. Notably, β-cell death occurs preferentially at night, when ppid expression is higher and antioxidant gene expression is reduced, suggesting β cells are more susceptible to mitochondrial stress at night.

Several limitations remain. Although zebrafish offer advantages for in vivo imaging and rapid genetic manipulation, validation in mammalian models is required to establish translational relevance. Although our data strongly support mPTP-mediated necrosis, we did not directly visualize pore opening or cell rupture. Additionally, we were unable to directly visualize mitochondrial ROS accumulation in zMIR larvae. Fluorescent dye CellROX has been used to detect increased ROS from metronidazole metabolism in β cells in 4-day-old larval zebrafish (46). We attempted several approaches to administer CellROX and MitoSOX Red but were unable to obtain a detectable signal within the islet. This was likely due to limited dye penetration into the islet, presumably due to the older age of the larvae (9-day-old) or a less severe oxidative stress, or both. As a result, our conclusions regarding mitochondrial oxidative stress are based on indirect evidence, such as antioxidant gene expression and pharmacological rescue with MitoTEMPO, rather than direct imaging.

Furthermore, antioxidant gene expression was measured during overnutrition, meaning metabolic state (fed at ZT4 vs fasted ZT16) could represent a confounding variable, as feeding and fasting are known to influence mitochondrial redox status and antioxidant gene expression (47–49). Additionally, expression measurements were only taken at two time points. Therefore, we cannot determine whether these differences reflect circadian oscillations, feeding-driven effects, or an interaction of the two.

Finally, these experiments were performed in larval zebrafish, whose β cells differ metabolically and developmentally from adult β cells. In some models, immature β cells rely on glycolytic ATP production, have reduced mitochondrial maturity, and altered stress-response pathways, all of which may influence baseline antioxidant gene expression and responses to metabolic stress (50–52).

Together, these findings identify Ppid as a β-cell-intrinsic effector of nutrient-induced β-cell loss, acting downstream of macrophage infiltration. Targeting Ppid or upstream mitochondrial activators may represent a therapeutic strategy to preserve β-cell mass in metabolic disease, particularly if aligned with windows of vulnerability. Given the alarming rise in adolescent T2D and its aggressive disease progression, insights into β-cell death mechanisms—especially those linked to nutrient stress and insulin resistance—is of critical importance.

Acknowledgements:

We thank other members of the Chen lab for constructive discussions, Corey Guthier for expert fish care. Cell imaging was performed in part through the use of the Vanderbilt Cell Imaging Shared Resource (supported by NIH grants CA68485, DK20593, DK58404, DK59637, and EY08126). National Institutes of Health grant R01DK117147 supported this study to WC and 5T32 DK07563 to BAC.

Footnotes

Conflict of Interest: The authors have no competing financial interests in relation to the work described.

Availability of Data and Materials:

All data needed to evaluate the conclusions in the paper are presented in the paper.

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