Reduced calcium levels and accumulation of abnormal insulin granules in stem cell models of HNF1A deficiency

González, Bryan J.; Zhao, Haoquan; Niu, Jacqueline; Williams, Damian J.; Lee, Jaeyop; Goulbourne, Chris N.; Xing, Yuan; Wang, Yong; Oberholzer, Jose; Blumenkrantz, Maria H.; Chen, Xiaojuan; LeDuc, Charles A.; Chung, Wendy K.; Colecraft, Henry M.; Gromada, Jesper; Shen, Yufeng; Goland, Robin S.; Leibel, Rudolph L.; Egli, Dieter

doi:10.1038/s42003-022-03696-z

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Published: 02 August 2022

Reduced calcium levels and accumulation of abnormal insulin granules in stem cell models of HNF1A deficiency

Communications Biology volume 5, Article number: 779 (2022) Cite this article

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Abstract

Mutations in HNF1A cause Maturity Onset Diabetes of the Young (HNF1A-MODY). To understand mechanisms of β-cell dysfunction, we generated stem cell-derived pancreatic endocrine cells with hypomorphic mutations in HNF1A. HNF1A-deficient β-cells display impaired basal and glucose stimulated-insulin secretion, reduced intracellular calcium levels in association with a reduction in CACNA1A expression, and accumulation of abnormal insulin granules in association with SYT13 down-regulation. Knockout of CACNA1A and SYT13 reproduce the relevant phenotypes. In HNF1A deficient β-cells, glibenclamide, a sulfonylurea drug used in the treatment of HNF1A-MODY patients, increases intracellular calcium, and restores insulin secretion. While insulin secretion defects are constitutive in β-cells null for HNF1A, β-cells heterozygous for hypomorphic HNF1A (R200Q) mutations lose the ability to secrete insulin gradually; this phenotype is prevented by correction of the mutation. Our studies illuminate the molecular basis for the efficacy of treatment of HNF1A-MODY with sulfonylureas, and suggest promise for the use of cell therapies.

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Introduction

Maturity onset diabetes of the young (MODY) is an autosomal dominant form of diabetes with onset typically before the age of 25 years accounting for 1–5% of diabetes incidence^1,2. There are at least 11 genes with mutations causing MODY, due to derangements in β-cell development or function. HNF1A-MODY is caused by mutations in the transcription factor HNF1A^3,4 and is among the most commonly diagnosed instances of MODY⁵. HNF1A-MODY patients have normal glucose tolerance during childhood and early adult life but show a progressive reduction of insulin secretion in response to glucose^6,7,8,9. Glycemia typically increases over time, resulting in the need for treatment with insulin secretory sulfonylurea drugs. Eventually, 30–40% of patients become insulin-dependent due to progressive deterioration of β-cell function.

HNF1A is a 631-amino acid transcription factor with three major domains: dimerization, DNA-binding, and transactivation. Over 200 HNF1A diabetes-related mutations have been identified in all major ethnic groups¹⁰. Understanding the role of the HNF1A gene and the pathophysiology of HNF1A-MODY has been difficult because of the limited access to islets of affected individuals. Mouse models of HNF1A deficiency do not accurately mimic patient phenotypes¹¹.

Stem cell-derived β-cells provide a useful model system, and have been used to study β-cell development in humans^12,13 and to recapitulate disease phenotypes^14,15. Differentiation of pluripotent stem cells to pancreatic endocrine cells can be achieved by a multistep protocol resulting in islet-like clusters containing all endocrine cell types^16,17,18. Transplanting these islet-like clusters into mice enables maturation and functional testing of stem cell-derived β-cells (scβ-cells) in vivo^19,20. Here we show that human stem cell-based models of HNF1A deficiency display islet developmental bias towards α-cells, altered insulin granule morphology and the stoichiometry of insulin:C-peptide secretion in vitro and in vivo. We used these models to identify disordered calcium homeostasis and accumulation of abnormal insulin granules as key mechanisms accounting for the secretory defects observed. This study was designed using two cell-based models: Human embryonic stem cell (hESC) lines rendered null for HNF1A in Figs. 1–5 and S1–S10; and induced pluripotent stem cell (iPSC) lines with HNF1A-MODY patient-specific mutations (R200Q) in Figs. 6 and S11–S13. Figure S1 provides a schematic overview of the study.

**Fig. 1: HNF1A deficiency impairs a network of genes required for calcium signaling, glucose-stimulated insulin secretion, and β-cell fate in vitro.**

**Fig. 2: HNF1A deficiency causes a developmental bias toward the α-cell fate.**

**Fig. 3: HNF1A deficiency affects insulin secretion due to reduced intracellular calcium levels in association with CACNA1A and SYT13 down-regulation in vitro.**

**Fig. 4: HNF1A deficiency alters the stoichiometry of insulin to C-peptide secretion in vivo.**

**Fig. 5: HNF1A deficiency causes abnormal insulin granule structure and fails to maintain glucose homeostasis in diabetic mice.**

**Fig. 6: HNF1A haploinsufficiency gradually impairs scβ-cell function in vivo.**

Results

Isogenic cell lines with HNF1A mutations in hESCs and HNF1A-MODY iPSCs

To elucidate the cellular functions of HNF1A, we used CRISPR/Cas9 to generate hESC lines (Mel1) harboring non-naturally occurring heterozygous and homozygous null mutations. The Mel1 hESC line has a INS^GFP/wt and GAPDH^{Luciferase/wt} dual-reporter, enabling imaging and isolation of INS-GFP⁺ cells²¹. GFP expression does not alter β-cell function in mice²² and does not cause ER stress in human scβ-like-cells²³. Short guide RNAs (sgRNAs) were used to target exon 3 of the HNF1A gene (Fig. S2a and Supplementary Data 1) because the DNA-binding domain in exon3 has the highest frequency of mutation in HNF1A-MODY patients⁵. Transfection of hESC lines with Cas9-GFP and sgRNAs #12 or #14, followed by sorting for GFP (Fig. S2b), achieved 17.3% and 20.4% indel efficiency as shown by surveyor assay (Fig. S2c). Gene editing resulted in compound heterozygous knockouts and heterozygous mutations (Fig. S2d) with no off-target mutations detected (Supplementary Data 2). Heterozygous (hESC HNF1A Het) and compound heterozygous-null mutant cell lines (hESC HNF1A KO1 and KO2) with premature protein termination were chosen for further studies (Fig. S2e). hESC HNF1A Het, KO1, and KO2 lines retained the HNF1A dimerization domain (truncated HNF1A), while the hESC HNF1A KO3 line was mutated at the start codon, deleting the entire HNF1A protein (Fig. S2e).

To understand the consequences of HNF1A-MODY patient-specific heterozygous mutations, we also generated iPSC lines from HNF1A-MODY patient fibroblasts containing heterozygous mutations in the transactivation domain (HNF1A-MODY iPSC Het: +/460_461insCGGCATCCAGCACCTGC, ID1056); and DNA-binding domain (HNF1A-MODY iPSC Het: +/R200Q, ID1075 and ID1076). The R200Q variant has been previously associated with HNF1A-MODY²⁴; the effect of this missense mutation is unknown, but likely pathogenic²⁴. We corrected the R200Q mutation from HNF1A-MODY iPSC Het (Fig. S2f) with an efficiency of 7.9% (5 clones out of 63) to generate isogenic wild type cells. We also generated iPSCs compound heterozygous knockout lines with an efficiency of 42.9% (27 clones out of 63) (Fig. S2d). An iPSC line with a deletion of the WT allele, and the patient’s mutation (HNF1A-MODY iPSC: R200Q/−) was chosen for further studies (Fig. S2g). We also introduced the same R200Q mutation into both alleles in the hESC line (hESC Hom: R200Q/R200Q) (Fig. S2h) with an efficiency of 20.8% (5 clones out of 24) (Fig. S2i). All genetically manipulated cell lines (Fig. S2j) resulted in modified versions of HNF1A protein (Fig. S2k, l), had a normal karyotype (Fig. S2m, n) and no mutations at potential off-target sites (Supplementary Data 2).

HNF1A is not required to generate pancreatic endocrine cells

To determine the functional consequences of HNF1A deficiency, hESC cells were differentiated into the pancreatic lineage²⁰ (Fig. S3a). Differentiation of wild type stem cells consistently generated 82% PDX1⁺/NKX6.1⁺ cells at the pancreatic progenitor stage (day 11). At the endocrine stage (day 27), we obtained 45% PDX1⁺/CPEP⁺ cells with 60% of them co-expressing NKX6.1 (scβ-like cells), 30% GCG⁺ cells (scα-like cells), and 10% SST⁺ cells (scδ-like cells) (Fig. S3b).

To determine the timing of HNF1A expression, we performed qPCR at different stages of differentiation. Insulin mRNA was detected at the endocrine stage (day 20) (Fig. S3c), whereas HNF1A mRNA is first detected at the primitive gut tube stage (day 5), increased at the pancreatic progenitor stage (day 11) and increased further at the endocrine stage (day 27) (Fig. S3d). HNF1A knockout resulted in a significant reduction of HNF1A transcript at the endocrine stage (Fig. S3e); HNF1A protein was detected in hESC WT, but only a faint signal was detected in hESC HNF1A KO-derived endocrine cells by western blot (Fig. S3f).

Mutations in HNF1A did not affect hESC lines in generating definitive endoderm cells (SOX17⁺) at day 3 or pancreatic progenitor cells (PDX1⁺/NKX6.1⁺) at day 11 of differentiation (Fig. S3g). At the endocrine stage (day 27), organoids were morphologically indistinguishable by the HNF1A genotype and showed no differences in INS-GFP fluorescence (Fig. S3h). HNF1A KO cells were capable of differentiation to all islet endocrine cell types, including PDX1⁺/NKX6.1⁺/SYP⁺/CPEP⁺ cells (scβ-like cells), GCG⁺ cells (scα-like cells), and SST⁺ cells (scδ-like cells) (Figs. 1a and S3i–m), indicating that HNF1A is not required to generate pancreatic endocrine cells. No differences were found for PDX1 and NKX6.1 in hESC HNF1A WT and HNF1A KO endocrine cells (day 27) by immunohistochemistry (Fig. S3j, k).

HNF1A deficiency impairs a network of genes required for calcium signaling, glucose-stimulated insulin secretion, and β-cell fate

To determine the transcriptional consequences of HNF1A deficiency in scβ-cells, bulk and single-cell RNA sequencing was performed in INS-GFP sorted β-like cells derived from hESC HNF1A KO and WT lines in vitro. Volcano plot analysis of bulk RNA sequencing identified 30 up-regulated genes and 148 down-regulated genes in HNF1A KO β-like cells (Fig. 1b and Supplementary Data 3). Single-cell RNA sequencing showed that HNF1A KO β-like cells segregated from HNF1A WT β-like cells based on their gene expression profile by principal component analysis (Fig. 1c), and volcano plot analysis (Fig. 1d) revealed previously undescribed HNF1A target genes involved in intracellular calcium signaling (CACNA1A, S100A6, and TMEM37), calcium-mediated insulin secretion (KCNH6) and glucose-stimulated insulin secretion (RFX6, DPEP1, DPP4, and IAPP) (Fig. 1e) as commonly downregulated with bulk RNA sequencing from HNF1A KO scβ-like cells. Among the up-regulated genes in HNF1A KO β-like cells, we found genes involved in the synaptic vesicle cycle (AP2A2, SH3KBP1, and HSPA1B) (Fig. 1e and Supplementary Data 4).

To understand the transcriptional consequences of HNF1A mutation in other endocrine cell types, we analyzed all single cells within the islet-like clusters. In addition to HNF1A KO lines, an hESC line homozygous for HNF1A R200Q (R200Q/R200Q) was included. Using clustering analysis from those cells, we grouped cells into 13 different populations (Fig. S4a, b). To cluster different endocrine cells, we used the expression of endocrine cell markers SYP, INS, GCG, and SST (Fig. S4c) and pancreatic progenitor markers NKX6.1, PDX1, MAFA, and HNF1A (Fig. S4d). This allowed the identification of insulin-expressing cells (cluster 0) and glucagon-expressing cells (cluster 2) (Fig. 1f). From the population of insulin-expressing cells (cluster 0), cells defined by high insulin and low glucagon expression were considered to be scβ-like cells and expression was compared between genotypes (Fig. S5a).

In scβ-like cells, a total of 73 genes were differentially expressed between HNF1A WT, homozygous R200Q, and KO lines (Fig. 1g and Supplementary Data 5). Pathway analysis of differentially expressed genes in scβ-like cells revealed genes involved in MODY, endocrine cell development, calcium signaling/sensing, insulin secretion and synaptic vesicle cycle (Fig. S5b). Furthermore, several MODY genes (RFX6, PAX4 and NEUROD1) were down-regulated (Fig. 1g). These transcription factors are important for the identity of adult pancreatic β-cells^25,26. We also identified down-regulated genes important for intracellular calcium signaling (S100A6 and TMEM37), exocytosis-regulated insulin secretion (SYT13), and glucose-stimulated insulin secretion (IGF2 and LLGL2) in HNF1A-mutated β-like cells (Fig. 1g and Supplementary Data 5)^25,27,28. Among the up-regulated genes, we found genes involved in the synaptic vesicle cycle (RAB3A and VAMP2) (Fig. 1g and Supplementary Data 5). RFX6, KCNH6, S100A6, A1CF, DPEP1, and DPP4 were the most consistently downregulated genes in HNF1A mutated β-like cells. In addition to those genes, SYT13, CACNA1A, and PAX4 were down-regulated in the scβ-like-cell subpopulation (Fig. 1h). A recent case study of cadaveric human islets of an HNF1A-MODY donor (+/T260M) also found down-regulation of RFX6, CACNA1H, IAPP, and TMEM37 in β-cells, and down-regulation of CACNA1A, RFX6, PCBD1, and PPP1R1A in α-cells²⁹. These genes are required for calcium signaling, glucose-stimulated insulin secretion, and determination of endocrine cell fate.

To determine the molecular consequences of HNF1A deficiency in other endocrine cell types, we analyzed mono-hormonal scα-like cells (cluster 2a), characterized by high glucagon and low insulin expression (Fig. S5c, d). We identified up-regulated genes involved in glucagon signaling pathways (PKM, CALM1, and CALM2) and down-regulated genes involved in insulin secretion (ADCY1, KCNH6, RFX6, IGF2, TM4SF4 and ATF4), calcium-mediated exocytosis (SYT7) and β-cell dedifferentiation (GC)³⁰ (Fig. S5e, f). Furthermore, in bi-hormonal cells (cluster 2b) expressing both insulin and glucagon, PYY was down-regulated, while GCG was up-regulated (Fig. S5g, h) in HNF1A mutated lines.

A recent publication by Cardenas-Diaz et al. ³¹ identifies LINC01139 as an HNF1A target implicated in β-cell respiration and mitochondrial function. We found no significant difference in the expression of this long non-coding RNA between WT and HNF1A mutated (hESC Het, hESC Hom R200Q/R200Q, hESC KO or iPSC HNF1A-MODY Het +/460ins, ID1056) β-like (Fig. S5i–k) and α-like cells (hESC Hom R200Q/R200Q and hESC KO) (Fig. S5l). No difference was found in cadaveric human β-cells from non-diabetic donors as compared to one HNF1A-MODY donor (+/T260M) (Fig. S5m).

In summary, HNF1A orchestrates a network of genes promoting endocrine differentiation to β-cells, and in differentiated β-cells regulates genes involved in calcium signaling, hormone exocytosis, and glucose-stimulated insulin secretion.

HNF1A deficiency causes a developmental bias towards the α-cell fate

Downregulation of genes involved in adult β-cell identity (PAX4 and RFX6) with upregulation of glucagon in HNF1A KO cells point to a developmental role of HNF1A. Both hESC HNF1A KO and WT cells gave rise to insulin-positive cells (Fig. S6a, b), with increased glucagon-positive cells in the hESC HNF1A KO line (Figs. 2a and S6b). This finding was confirmed by an increase in GCG⁺/CPEP⁺ cell type ratio by immunohistochemistry (Fig. 2b) and glucagon content by ELISA (Fig. 2c). The increase in scα-like cells in HNF1A KO cells was driven by an ~2-fold increase in the number of bi-hormonal scβ/α-like cells as shown by flow cytometry (Figs. 2d and S3m), immunohistochemistry (Figs. 2b and S6c), and RNA sequencing (Fig. S6d). Transcriptional analysis by single-cell RNA sequencing of glucagon and insulin double hormone-positive cells showed an increase in glucagon gene expression and a decrease in insulin gene expression in HNF1A KO cells compared to WT cells (Fig. 2e). These findings are consistent with PAX4 repressing pancreatic glucagon gene expression³².

To assess the developmental requirements of HNF1A in endocrine cells in vivo, we transplanted pancreatic clusters derived from hESC HNF1A WT and hESC HNF1A KO lines with GAPDH^{Luciferase/wt} and INS^GFP/wt dual-reporter²¹. Mice received ~180 clusters of stem cell-derived islet-like cells (Fig. S7a) containing a similar amount of CPEP/PDX1 positive cells across genotypes (Fig. S7b, c). Cell transplantation was done by injection into the ventral and medial muscle groups of the left quadriceps in NOD SCID gamma immunodeficient mice (NSG mice) (Fig. 2f). Skeletal muscle has been used for other endocrine transplants, including for parathyroid auto-transplantation in patients undergoing parathyroidectomy with a 93% success rate³³. Analysis of engraftment efficiency was evaluated by bioluminescence intensity (BLI). Mice with successful engraftment showed a two-fold increase in BLI 4–6 weeks post-transplantation, while those with failed engraftment showed a decrease (Fig. S7d). Transplantation was successful in 79% (31/39) of mice, independent of the HNF1A genotype (Fig. S7e). 92.5% (49/53) of mice transplanted remained teratoma-free (Fig. S7f). In those mice, the graft explant (30 weeks post-transplantation) (Fig. S7g) was a vascularized tissue of about ~220 mg (Fig. S7h). According to luminescence intensity, there was a gradual increase in BLI from week 0 to 30 post-transplantation without significant differences by HNF1A genotype (Fig. 2g). Grafts remained localized for up to 50 weeks post-transplantation, and in no case (0/39) was luminescence detected in an ectopic location. These results show that skeletal muscle is a stable and suitable transplantation site for SC-derived islet-like cells.

Thirty weeks post-transplantation, we isolated grafts from normoglycemic animals. Quantification of scβ-mass as determined by GFP fluorescence did not differ by HNF1A genotype (Figs. 2h, i and S7i, j). The presence of exclusively mono-hormonal endocrine cells, including scβ-cells (CPEP⁺/PDX1⁺/NKX6.1⁺), as well as glucagon- and somatostatin-positive cells were apparent in both hESC HNF1A WT and hESC HNF1A KO grafts (Fig. 2j). No double hormone-positive cells were identified in either genotype. Consistent with the in vitro studies, the percentage of scα-cells in HNF1A KO compared to HNF1A WT islet-like structures was increased by 24% (Figs. 2k and S7k), leading to a significant increase in GCG⁺/CPEP⁺ cell type ratio (Fig. 2l); comparable to cadaveric human islets of an HNF1A-MODY donor (+/T260M)²⁹ (Fig. 2k, l). Ex vivo analysis of isolated HNF1A KO grafts showed an increase in glucagon protein content by ELISA (Figs. 2m and S7l, m). Glucagon secretion was higher in HNF1A KO grafts when stimulated with glibenclamide, a second-generation sulfonylurea drug compared to HNF1A WT grafts (Fig. 2n). Similarly, elevated glucagon secretion upon KCl stimulation was detected in vitro from HNF1A KO-derived endocrine cells (Fig. S7n). These results point to both a gain of α-cell number and enhanced α-cell function due to HNF1A deficiency. This is consistent with the up-regulation of genes (PKM, CALM1, and CALM2) involved in glucagon signaling pathways in scα-like cells.

HNF1A deficiency affects glucose-mediated secretion of insulin

To determine the consequences of HNF1A deficiency on β-cell function, we measured glucose-stimulated insulin secretion. HNF1A KO scβ-like cells had reduced basal insulin secretion (Fig. S8a) and impaired glucose-stimulated insulin secretion compared to WT scβ-like cells and compared to human pancreatic islets in vitro (Fig. 3a). These differences in hormone secretion between HNF1A mutant and HNF1A WT scβ-like cells were not due to a reduction in insulin content (Fig. S8b). In contrast to glucose, treatment with the insulin secretagogues, tolbutamide, or potassium chloride resulted in insulin secretion in HNF1A KO cells comparable to HNF1A WT β-like cells (Fig. 3b). Thus, membrane depolarization by sulfonylurea causes secretion of insulin from HNF1A mutant β-cells that do not respond to glucose alone, consistent with the clinical efficacy of sulfonylurea drugs. These differences in response to insulin secretagogues are likely intrinsic to the β-cell and not due to augmented glucagon³⁴.

As calcium signaling is a mechanism for the release of insulin, we measured intracellular calcium levels in dispersed scβ-like cells in vitro. Both genotypes had glucose-stimulated calcium responses (Fig. 3c). However, hESC HNF1A KO β-like cells had significantly lower calcium levels relative to hESC HNF1A WT scβ-like cells as measured by Fura-2 fluorescence (Fig. 3c). This finding was further supported by measurements of absolute intracellular calcium concentrations by Fura-2 fluorescence (Fig. 3d). Among the HNF1A target genes, reduced expression of CACNA1A is potentially involved in reducing intracellular calcium levels, and SYT13 may be required for efficient exocytosis-mediated insulin secretion. CACNA1A encodes a voltage-dependent calcium channel mediating the entry of calcium and is involved in calcium-dependent insulin secretion and type 2 diabetes³⁵. Synaptotagmins are calcium sensors localized in the β-cell insulin granules and are required for vesicle fusion and glucose-stimulated insulin release^36,37,38. SYT13 is a member of the synaptotagmin family and is predicted to be involved in calcium-regulated exocytosis. SYT13 is down-regulated in human T2D islets and silencing of SYT13 impairs insulin secretion in INS1-832/13 cells³⁹. To determine whether the down-regulation of CACNA1A and SYT13 seen in HNF1A mutant cells (Fig. 1h) reproduced the reduced intracellular calcium levels and reduced glucose-stimulated insulin secretion, we generated hESC CACNA1A KO and SYT13 KO cell lines using CRISPR/Cas9. Sanger sequencing revealed homozygous mutations resulting in frame shifts (Fig. S8c). Differentiation of CACNA1A KO and SYT13 KO lines generated INS-GFP organoids (Fig. 3e) comprised of ~51% PDX1/CPEP and ~5% GCG positive cells (Figs. 3f and S8d) with no differences compared to isogenic WT cells. Dispersed CACNA1A KO β-like cells (Fig. S8e) had significantly reduced intracellular calcium levels compared to WT cells (Figs. 3g and S8f). The reduction of intracellular calcium in CACNA1A KO was intermediate to HNF1A KO cells, indicating that CACNA1A is not the only gene affecting intracellular calcium levels in HNF1A KO cells. Intracellular calcium levels in SYT13 KO were unchanged compared to WT cells (Figs. 3g and S8f). Reduced basal and stimulated insulin secretion were observed in both CACNA1A KO and SYT13 KO scβ-like cells at intermediate levels to HNF1A KO scβ-like cells (Fig. 3h). Treatment with tolbutamide or KCl stimulated insulin secretion in all KO lines comparable to the corresponding WT lines. Impairments in insulin release due to reduced levels of these molecules can be overcome by elevating calcium beyond physiological levels using tolbutamide or KCl. The milder phenotypes of cells deficient for single target genes compared to HNF1A KO cells suggests a combined effect on several HNF1A target genes on insulin secretion.

HNF1A deficiency alters the stoichiometry of insulin to C-peptide secretion

To interrogate the function of HNF1A KO β-cells in vivo, we monitored circulating human insulin and C-peptide concentrations in euglycemic mice transplanted with sc-islet-like clusters. By 4 weeks post-transplantation, plasma human C-peptide concentrations were significantly lower in mice transplanted with HNF1A KO β-cells compared to HNF1A WT β-cells (Fig. 4a). Circulating human C-peptide concentrations in mice transplanted with HNF1A WT β-cells increased gradually, reaching human physiological levels (652 ± 146 pM) 24–30 weeks post-transplantation. In mice transplanted with HNF1A KO β-cells, plasma human C-peptide concentrations increased to a maximum of 352 ± 142 pM at 30 weeks post-transplantation (Fig. 4a). In mice transplanted with HNF1A WT β-cells, human insulin was detected as early as 4 weeks post-transplantation and concentrations increased over time, reaching human physiological levels (12.74 ± 2.1 mU/L) 24–30 weeks post-transplantation. However, mice transplanted with hESC HNF1A KO β-cells had virtually undetectable plasma human insulin concentrations for 30 weeks (Fig. 4b) despite human C-peptide levels of >300 pM. These differences in hormone secretion between HNF1A mutant and WT scβ-cells were not due to reduced scβ-cell mass in vivo (Fig. 2i). These findings led us to quantify the secretion ratio of hormones in scβ-cells in vitro. For equivalent secretion of human C-peptide at basal glucose condition (Fig. S8g), secretion of human insulin was significantly reduced in hESC HNF1A KO lines (Fig. S8h), leading to decreased insulin:C-peptide secretion ratios in vitro (Fig. S8i). Thus, the absence of insulin in the plasma was not a limitation of the sensitivity of the assay but a decrease in the stoichiometry of insulin to C-peptide in circulation. While plasma of mice grafted with HNF1A WT β-cells consistently showed an insulin:C-peptide molar ratio of 0.22 ± 0.11 from week 4 to 30 post-transplantation, mice grafted with HNF1A KO β-cells had an 18-fold lower ratio (Fig. 4c). This decrease in circulating insulin:C-peptide ratio was reciprocal to a 3-fold increase in the insulin:C-peptide ratio of intracellular content from HNF1A KO isolated grafts (Fig. 4d), indicating a complementary imbalance of insulin to C-peptide. The altered insulin:C-peptide ratio is not attributable to differences in insulin processing because insulin:proinsulin ratios in HNF1A KO and HNF1A WT grafts were identical in vivo (Fig. 4e) and in vitro (Fig. S8j–l), and no differences were found in the transcript levels of processing genes (PC1/PC3) (Supplementary Data 3–5). Impaired stoichiometry of circulating insulin to C-peptide was also observed in mice transplanted with two additional homozygous-mutant hESC lines (Fig. S8m). Therefore, HNF1A deficiency not only impairs insulin secretion, but also the stoichiometry of circulating insulin to C-peptide ratios.

To evaluate glucose-stimulated insulin secretion, we performed intraperitoneal glucose tolerance tests (iPGTT). During fasting and following an iPGTT, mice transplanted with HNF1A WT β-cells displayed normal human insulin and C-peptide secretion profiles (Fig. 4f–h). In contrast, HNF1A KO β-cells were non-responsive to glucose: plasma human C-peptide was not decreased by fasting and showed no significant increase after glucose injection (Fig. 4g). Circulating human insulin remained undetectable in animals engrafted with HNF1A KO β-cells (Fig. 4h). Therefore, HNF1A deficiency affects glucose-stimulated insulin secretion, and C-peptide is released independent of metabolic state, resulting in an altered ratio of insulin to C-peptide.

Glibenclamide restores insulin secretion in HNF1A-deficient scβ-cells

In contrast to glucose, intra-peritoneal injection of the sulfonylurea drug, glibenclamide, in mice transplanted with hESC HNF1A KO β-cells resulted in a significant increase in plasma C-peptide after 10 min (Fig. 4i). In parallel, plasma human insulin concentrations in mice transplanted with HNF1A KO β-cells increased from undetectable levels to 14 ± 21 mU/L, reaching levels comparable to the control group (Fig. 4j). Insulin:C-peptide ratios in plasma of animals transplanted with HNF1A KO β-cells were increased 10-fold by glibenclamide, equal to the ratios in mice transplanted with HNF1A WT β-cells (Fig. 4k). Clearance of insulin from the circulation occurred with the same kinetics in mice grafted with HNF1A WT and HNF1A KO β-cells 20 min after glibenclamide administration (Fig. 4k), excluding insulin stability or increased clearance in plasma as the cause for the low insulin concentrations in mice transplanted with HNF1A mutant scβ-cells. To further test whether reduced insulin:C-peptide secretion ratios from HNF1A KO β-cells were due to secretory defects and not insulin stability or clearance, we isolated the HNF1A KO grafts from mice and identified a significant decrease in insulin:C-peptide secretion ratios compared to HNF1A WT grafts ex vivo. This ratio was restored after exposure of the grafts to glibenclamide (Fig. 4l). Thus, membrane depolarization initiated by the closure of the ATP-sensitive K⁺ (K_ATP) channels (target of glibenclamide) and high intracellular calcium levels cause the secretion of insulin that is abnormally retained in HNF1A-mutant β-cells.

HNF1A deficiency causes abnormal insulin granule structure

To identify the basis of the insulin secretory defect in HNF1A KO β-cells we examined insulin granule morphology by electron microscopy from 30-week explants of normoglycemic mice. HNF1A WT grafts contained scβ-cells with granules of a high-electron density core separated from limiting membranes by a “halo” (Figs. 5a and S9a), similar to human β-cells. In contrast, HNF1A KO islets-like scβ-cells showed a majority (>90%) of insulin granules that were abnormally enlarged (Fig. 5a–c) with a diffuse electron-light core (Figs. 5d, e and S9a), characteristic of immature granules. Enlarged insulin granules were further confirmed by immunogold staining for human C-peptide (Fig. 5f). Similar results were observed in vitro, where HNF1A-mutated cells had increased insulin granules diameters with a diffuse light core (Fig. S9b, c). For scα-cells, there were no differences in morphology, structure, or granule size between HNF1A genotypes (Fig. S9d). These results show that HNF1A is required for normal dense-core insulin granule formation.

HNF1A deficient scβ-cells are unable to maintain glucose homeostasis in diabetic mice

To test the ability of HNF1A KO β-cells to maintain in vivo normoglycemia, endogenous murine β-cells were ablated by streptozotocin (STZ) 22 weeks post-transplantation. By virtue of the species-related differences in glucose transporters, STZ specifically targets murine β-cells without affecting human β-cells⁴⁰. We first confirmed that STZ treatment did not alter the secretion of c-peptide in sc-β-like cells. There was no change in C-peptide secretion (Fig. S10a) between genotypes in vitro. In vivo, circulating plasma human C-peptide from mice transplanted with HNF1A WT or HNF1A KO cells was unchanged or increased post STZ treatment (Fig. 5g). Mouse C-peptide was undetectable by ELISA (Fig. S10b) and immunohistochemistry of the pancreas (Fig. S10c) demonstrating successful ablation of mouse β-cells. Mice transplanted with the HNF1A WT β-cells were normoglycemic (~100 mg/dl and HbA1C ~5%) for at least 10 weeks post-STZ injection (Figs. 5h and S10d) and had normal human insulin and C-peptide secretion profiles during an iPGTT (Fig. S10e–g). In contrast, mice transplanted with HNF1A KO β-cells did not increase human C-peptide secretion (Fig. 5g) after becoming severely diabetic (blood glucose >500 mg/dl) within one week following STZ injection (Fig. 5h). HNF1A KO β-cells were unresponsive to blood glucose even at high levels, failing to maintain systemic glucose homeostasis in a mouse model of β-cell deficient diabetes.

HNF1A R200Q homozygous mutation is pathogenic and causes a developmental bias towards the α-cell fate in vitro

HNF1A-MODY patients are heterozygous for the causal HNF1A alleles. To more directly characterize the consequences of heterozygous patient-specific mutations in HNF1A, we differentiated two heterozygous HNF1A-MODY iPSC lines (+/460_461insCGGCATCCAGCACCTGC, ID1056 and +/R200Q), isogenic mutation-corrected HNF1A-MODY iPSC lines R200Q -corrected WT(+/+) and a hESC HNF1A Het line (Fig. S2J, C30) into endocrine cells. Heterozygous mutations in HNF1A did not affect the generation of definitive endoderm cells (SOX17⁺) or pancreatic progenitor cells (PDX1⁺/NKX6.1⁺) compared to WT lines as determined by immunohistochemistry (Fig. S11a). At the endocrine stage (day 27), organoids were morphologically indistinguishable and all heterozygous cell lines differentiated efficiently to all pancreatic endocrine cells (scα-, scβ- and scδ-like cells); scβ-like cells co-expressed PDX1/NKX6.1/HNF1A (Fig. S11b). HNF1A protein was detected in WT, HNF1A-MODY R200Q Het (+/R200Q), and HNF1A-MODY R200Q-corrected WT (+/+) iPSC-derived endocrine cells by Western blot (Fig. S11c). In contrast to HNF1A truncated lines, the heterozygous point mutation R200Q of HNF1A-MODY iPSC line did not affect the total amount of HNF1A protein, suggesting that the mutant R200Q protein was produced (Fig. S11c). No significant differences were found in the proportion of endocrine cell types (scα-, scβ- and scδ-like cells) between HNF1A WT and heterozygous cell lines at day 27 of differentiation by immunohistochemistry (Fig. S11d).

To determine HNF1A-MODY patient lines for the bias towards α-cell fate, we measured the percentage of glucagon cells co-expressing C-peptide in two HNF1A-MODY iPSC Het (+/460ins and +/R200Q) lines in vitro. We found no significant differences between HNF1A-MODY iPSC lines and control iPSC R200Q-corrected WT lines (Fig. S11e). To determine the pathogenicity of the HNF1A R200Q mutation, we knocked out the HNF1A wild type allele in the HNF1A-MODY iPSC Het (+/R200Q) line to make an HNF1A-MODY iPSC Hom (R200Q/−) line, which resulted in a significant ~70% increase in GCG and CPEP co-expressing cells compared to the R200Q-corrected WT control lines (Fig. S11e, f). We also detected a significant increase in the percentage of GCG and CPEP co-expressing cells in hESC HNF1A R200Q homozygous (R200Q/R200Q) and hESC HNF1A Het (+/−) lines compared to hESC WT (+/+) lines (Figs. S11g and S11h). Thus, while both heterozygous HNF1A-MODY patient cells (+/460ins and +/R200Q) show no developmental bias towards the α-cell fate, complete loss of HNF1A WT allele in HNF1A-MODY iPSC R200Q homozygous (R200Q/−) and hESC HNF1A R200Q homozygous (R200Q/R200Q) results in a developmental bias towards α-cell fate, indicating the pathogenicity of the R200Q mutation. Interestingly, the hESC heterozygous line harboring a frameshift mutation (Fig. S2J, clone C30) in the DNA-binding domain and causing a premature stop codon presented a strong bias towards α-cell fate (Fig. S11h). Previous studies have shown that frameshift mutations in the DNA-binding domain can cause dominant-negative effects⁴¹.

HNF1A haploinsufficiency gradually impairs scβ-cell functional capacity in the context of metabolic stress

To assess the developmental potential of HNF1A-MODY patient-specific mutations in vivo, we transplanted pancreatic islet-like clusters derived from iPSCs. While 87.9% (51/58) of mice transplanted with isogenic HNF1A-MODY iPSC Het (+/R200Q) and R200Q-corrected WT (+/+) lines were teratoma-free, the majority (61.5%, 8/13) of the HNF1A-MODY iPSC Het (+/460ins) and control iPSC WT lines showed teratoma formation (Fig. S11i). Because of variable teratoma formation among different iPSC lines²⁰, only teratoma-free mice were used for further analysis. Teratoma-free transplanted mice had 67.3% (35/52) engraftment efficiency. Thirty weeks post-transplantation, we isolated grafts from normoglycemic animals (Fig. S11j) and found no differences in glucagon-to-insulin content ratios or endocrine cell types between HNF1A-MODY iPSC Het (+/R200Q) and HNF1A-MODY iPSC R200Q-corrected WT (+/+) control grafts (Fig. S11k, l). A recent study of cadaveric HNF1A-MODY human islets with a T260M heterozygous mutation in the DNA-binding domain²⁹, showed a bias of endocrine cells toward the α-cell fate (Fig. 2k). A bias was also observed in hESC HNF1A heterozygous line with a frameshift mutation in the DNA-binding domain (Fig. S11h). Mutations affecting the DNA-binding domain or truncating mutations are associated with the earlier onset of diabetes in HNF1A-MODY patients⁴². Thus, different mutations affect HNF1A function to different degrees, resulting in phenotypic differences among patients as well as characteristic cellular phenotypes in a stem cell system.

To understand the function of HNF1A-MODY iPSC Het β-cells in a physiological context, we monitored circulating human insulin and C-peptide in transplanted euglycemic mice over 30 weeks. Mice transplanted with iPSC-derived cells from heterozygous HNF1A-MODY patients (+/460_461insCGGCATCCAGCACCTGC) had low, but detectable circulating human C-peptide 2, 4, 8 (Fig. S12a) and 30 weeks post-transplantation (Fig. S12b) with undetectable human insulin (Fig. S12c). Since human insulin levels are under the minimal threshold limit for detection by ELISA, this deficiency can be attributed to the lower ability of these mutant iPSCs to generate functional insulin-producing cells in vivo. In mice transplanted with iPSC-derived cells from HNF1A-MODY patient (+/R200Q) and R200Q-corrected WT (+/+) isogenic lines, circulating human C-peptide (Fig. S12b) and human insulin (Fig. S12c) concentrations reached comparable levels 30 weeks post-transplantation. Transplantation of mice with hESC HNF1A Het (+/−), thought to be a dominant negative allele, and hESC HNF1A R200Q homozygous (R200Q/R200Q) line resulted in low circulating concentrations of human C-peptide over 30 weeks post-transplantation (Fig. S12a, b) accompanied by low to undetectable human insulin (Fig. S12c). These results show that HNF1A heterozygous frameshift mutations in the DNA-binding domain (hESC +/−) and HNF1A R200Q homozygosity (hESC R200Q/R200Q) are pathogenic and that heterozygous patient mutations (iPSC +/R200Q) are less detrimental to β-cell function.

To further interrogate the consequence of the R200Q mutation in β-cells, we performed intraperitoneal glucose tolerance tests (iPGTT) in mice transplanted with the HNF1A-MODY iPSC Het (+/R200Q) and R200Q-corrected WT (+/+) isogenic lines, as these iPSC lines allowed the reliable generation of transplanted mice with high C-peptide and insulin levels over 30 weeks post-transplantation (Fig. S12d, e). During an iPGTT, mice transplanted with both isogenic lines showed comparable changes in blood glucose (Fig. S12f) and plasma human C-peptide (Fig. 6a); plasma human insulin concentrations dropped below detection during fasting and increased upon glucose injection (Fig. 6b), demonstrating homeostatic glucose responsiveness. Glibenclamide injection increased C-peptide and insulin concentrations in both genotypes (Fig. S12g, h) at equivalent ratios (Fig. S12i). No significant differences in insulin secretion (Fig. S12j) or endocrine hormone content were found in vitro or in vivo between genotypes (Fig. S12k, l).

HNF1A-MODY patients generally display normal glucose tolerance during early childhood and exhibit symptomatic diabetes in their late teens or early adulthood⁹ depending on the type and position of the HNF1A mutation⁴². To determine whether there are functional differences between HNF1A-MODY heterozygous (+/R200Q) and HNF1A-MODY R200Q-corrected WT (+/+) scβ-cells, we treated mice with STZ, thereby exposing transplanted scβ-cells to higher insulin demand due to ablation of mouse endogenous β-cells. Two weeks post STZ, animals transplanted with HNF1A-MODY iPSC R200Q Het (+/R200Q) cells showed rapid progression to hyperglycemia (Fig. 6c). The increase in blood glucose (mice n = 9) was accompanied by a failure to increase circulating human C-peptide (Fig. 6d) and a reduction of human insulin concentrations (Fig. 6e) over 5–6 weeks after STZ treatment. In contrast, in mice (n = 6) transplanted with isogenic HNF1A-MODY iPSC R200Q-corrected WT cells (+/+), concentrations of human plasma C-peptide (Fig. 6d) and insulin increased during this period (Fig. 6e).

Five to six weeks post STZ treatment, we found that the ratio of circulating insulin to C-peptide fell by 55% in mice transplanted with HNF1A-MODY iPSC R200Q Het (+/R200Q) cells. In contrast, insulin:C-peptide ratios remained constant in mice transplanted with HNF1A-MODY R200Q-corrected WT (+/+) cells (Fig. 6f). The decrease in circulating insulin:C-peptide ratios were reciprocal to a significant increase by 74% in the insulin:C-peptide ratios of intracellular content from isolated HNF1A-mutated grafts compared to isogenic controls (Fig. 6g), demonstrating complementary imbalance of insulin to C-peptide secreted. These results show that HNF1A R200Q haploinsufficiency gradually impairs the stoichiometry of circulating insulin:C-peptide, and that gene correction of the R200Q mutation protects HNF1A-MODY iPSC β-cells from acquiring this imbalance.

In sequential iPGTTs 2–8 weeks post-STZ administration, HNF1A-MODY iPSC R200Q Het (+/R200Q) scβ-cells showed progressive impairment of glucose-stimulated insulin secretion: mild at two weeks post-STZ (Fig. S13a, b) and completely unresponsive to glucose challenges four weeks post-STZ (Fig. S13c, d). Mice transplanted with HNF1A-MODY scβ-cells fail to clear glucose from the circulation during a glucose tolerance test at 6–8 weeks post STZ treatment (Figs. 6h and S13e). HNF1A R200Q heterozygous (+/R200Q) scβ-cells showed no glucose-stimulated insulin secretion, whereas HNF1A-MODY R200Q-corrected WT (+/+) scβ-cells were sensitive to fasting, remained glucose responsive 6–8 weeks post STZ treatment, and cleared glucose from circulation (Figs. 6h, i and S13e, f). Mice transplanted with HNF1A-MODY iPSC R200Q Het (+/R200Q) scβ-cells became diabetic as reflected by elevated blood glucose (Fig. S13g) and HbA1c levels (Fig. 6j) compared to control mice. These results show that HNF1A-MODY iPSC (+/R200Q) β-cells fail to compensate for higher metabolic insulin demands and that hyperglycemia is due to the gradual development of insulin secretory defects. Correction of R200Q mutations protects HNF1A-MODY iPSC β-cells from acquiring abnormal insulin secretion profiles.

Discussion

Here we report the use of stem cell-derived islets containing HNF1A mutations to elucidate the molecular basis for β-cell dysregulation of insulin secretion in HNF1A hESC null and HNF1A-MODY hypomorphic (R200Q) patient lines. In HNF1A knockout stem cell-derived β-cells, glucose-stimulated insulin secretion is compromised in association with a reduction in CACNA1A and intracellular calcium levels, and down-regulation of SYT13, a gene involved in calcium-regulated granule exocytosis. Mutations in these genes reproduce the insulin secretion phenotypes found on the stem cell model of HNF1A deficiency. These differences in gene expression were seen in both knockout and hESC-derived cells rendered homozygous for R200Q (R200Q/R200Q), confirming the pathogenicity of the R200Q allele. The deficiency in insulin secretion can be overcome through the application of sulfonylureas, by increasing the low calcium levels found in HNF1A mutant cells. Thereby we demonstrate the molecular basis for the efficacy of sulfonylureas in the treatment of HNF1A-MODY. However, HNF1A deficiency has pleiotropic effects, which will need to be further elucidated for effective treatment long term. The triggering of insulin release through sulfonylureas often becomes insufficient at later stages of the disease.

In particular, HNF1A also has developmental functions; hypomorphic mutations in HNF1A perturb the expression of genes important in β-cell identity such as PAX4 and lead to increases in glucagon gene expression, α-cell number, and the α- to β-cell ratio in mature pancreatic islets. This α-cell bias phenotype was observed in homozygous hESC HNF1A knockouts as well as homozygous hESC and iPSC for R200Q (iPSC R200Q/− and in hESC R200Q/R200Q) but was not seen in HNF1A-MODY patient iPSC-derived heterozygous cells (+/460ins and +/R200Q). A recent case study of cadaveric human islets from a 33-year-old HNF1A-MODY donor with a heterozygous mutation (+/T260M) in the HNF1A DNA-binding domain and with a 17 years history of diabetes also showed an increase in the ratio of α- to β-cells²⁹.

The accumulation of abnormal insulin secretory granules in HNF1A knockout β-cells was associated with a deviation of the 1:1 stoichiometric release of C-peptide and insulin from β-cells, characterized by the constitutive release of C-peptide and intracellular retention of insulin. Their equimolar secretion was restored by depolarization of the cells with sulfonylurea. Previous studies have identified a “constitutive-like” secretory pathway that is characterized by the accumulation of immature secretory granules that secrete newly synthesized C-peptide in molar excess of insulin^43,44. Our data are consistent with this form of secretion, but a detailed study of insulin secretory dynamics in HNF1A-MODY patients is needed to fully elucidate the clinical contributions of the secretory defects described here. We cannot formally exclude the possibility that insulin secreted from mutant cells is more efficiently removed from the circulation than insulin secreted from wild type cells.

The insulin secretory phenotype of heterozygous HNF1A-MODY iPSC-derived β-cells differed from complete HNF1A deficiency. Heterozygous β-cells (+/R200Q) initially functioned normally in transplanted mice, consistent with normal glucose tolerance of heterozygous carriers during early childhood, followed by symptomatic diabetes within the first three decades of life (Supplementary Data 6 for the age of onset)⁹. In response to increasing the requirements for human insulin following ablation of the mouse’s intrinsic β-cells, iPSC-derived β-cells from HNF1A-MODY patients were unable to fully compensate for the increased insulin requirement, and gradually developed phenotypes resembling those of HNF1A knockout β-cells, including a failure to secrete insulin in response to glucose, and reduced insulin to C-peptide secretion ratios. Correction of the R200Q mutation in HNF1A-MODY iPSC-derived β-cells protected cells from acquiring abnormal insulin secretion profiles.

Our study illustrates both congruences as well as differences with other stem cell models of HNF1A deficiency. Our data and another study identify a bias in endocrine differentiation to glucagon-positive cells in HNF1A knockout cells⁴⁵. Cardenas-Diaz also documents this bias in heterozygous cells, to an extent comparable to biallelic knockout cells. Mutations in HNF1A introduced by Cardenas-Diaz are located adjacent to the start codon, excluding possible dominant-negative effects of a truncated protein. Such dominant-negative truncating mutations were made by another group and shown to affect pancreatic differentiation from pluripotent stem cells by interfering with HNF1B function⁴⁶. Both our study and Cardenas-Diaz et al. identify a deficiency of glucose-stimulated insulin secretion in HNF1A biallelic knockout cells⁴⁵. Surprisingly, Cardenas-Diaz and colleagues found that heterozygous and homozygous knockout cells were equally impaired in insulin secretion. The finding that haploinsuffiency is as detrimental as a complete knockout or a dominant negative mutation across different phenotypic aspects, is surprising and was not observed here. Cardenas-Diaz et al. linked insulin secretion phenotypes to altered mitochondrial function decreased expression of LINC01139 and increased expression of LDHA involved in anaerobic glycolysis, reporting similar changes in expression of these genes in both heterozygous and homozygous mutant cells. An increase of LDHA due to HNF1A deficiency was not observed in our study, neither in bulk-RNAseq nor in single-cell RNAseq, and neither in HNF1A knockout or heterozygous cells. Neither did Haliyur et al. see these changes in the pancreatic islets of an HNF1A-MODY heterozygous (+/T260M) donor²⁹. LDHA is highly expressed in undifferentiated pluripotent stem cells; its expression may thus be affected by the maturity of stem cell-derived pancreatic cell clusters. An intriguing concordance of phenotypes of a stem cell model (Cardenas-Diaz et al.) and islets from an HNF1A-MODY patient (Haliyur et al.) is the increased basal secretion rate of insulin. However, such phenotypic concordances among studies are not necessarily due to concordant mechanisms. Other factors, including increased LDHA expression and rates of glycolysis, may also affect basal insulin secretion in stem cell-derived insulin-producing cells. Interestingly, we find that HNF1A knockout cells show reduced expression of KCNH6, mutations which are associated with elevated insulin secretion⁴⁷. Another study has found downregulation of glucose transporter GLUT2 expression and reduced ATP generation in HNF1A-MODY patient iPS-derived β-cells with heterozygous mutation (+/H126D) in comparison to control wild type lines of a different genetic background⁴⁸. In summary, these stem cell-based studies on HNF1A deficiency reveal pleiotropic effects of HNF1A deficiency at several levels of β-cell biology and function, consistent with the diversity of its transcriptional targets.

Among the clinical implications of this study are that stem cell-derived human islet cell clusters capable of supporting glucose homeostasis in mice. In MODY patients, β-cell autoimmunity is not mechanistically involved, so islets created from somatic cells of such patients should be immunologically tolerated by the patient. As shown here, the mutant alleles of HNF1A-MODY patients could be corrected to produce autologous stem-cell-derived islets with a normal HNF1A allele. While sulfonylureas can be used to treat patients with HNF1A-MODY effectively, insulin dependence is common after years of treatment. For these patients, cell therapy might be considered.

Methods

Human subject and cell lines

Nine HNF1A-MODY subjects (Pt1–Pt9), three MODY2 subjects (1068, 1133 and 1144), and four control subjects (1023, 1098, 1136 and 1015) were recruited at the Naomi Berrie Diabetes Center and monitored over 2–5 years. Samples were coded to protect subjects’ identities (Fig. S2j). Biopsies from HNF1A-MODY subjects (Pt1, Pt2 and Pt3) and one control subject (1023) were cut into small pieces (approximately 5 × 5 mm in size). 2–3 pieces of minced skin were placed next to a droplet of silicon in a well of the six-well dish. A glass coverslip (22 × 22 mm) was placed over the biopsy pieces and silicon droplet. 5 ml of biopsy plating media was added and incubated for 5 days at 37 °C. Biopsy pieces were then grown in a culture medium for 3–4 weeks. Biopsy plating medium is composed of DMEM (Gibco, 10569), 10% FBS (GE Healthcare, SH30088.03HI), 1% GlutaMAX (Gibco, 35050061), 1% Anti–Anti (Gibco, 15240062), NEAA (Gibco, 11140-050), 0.1% 2-Mercaptoethanol (Gibco, 21985023) and nucleosides (Millipore ES-008-D). Culture medium contains DMEM (Gibco, 10569), 10% FBS (GE Healthcare, SH30088.03HI), 1% GlutaMAX (Gibco, 35050061) and 1% Pen-Strep (Gibco, 15070063). The hESCs (Mel1)²¹ used in this manuscript is an NIH-approved line. All research involving human subjects was approved by the Institutional Review Board (IRB) of Columbia University Medical Center, and all participants provided written informed consent.

Generation of iPSCs and cytogenic analysis of stem cells

Primary fibroblasts were reprogrammed into pluripotent stem cells using CytoTune™-iPS Sendai Reprogramming Kit (Invitrogen). 50,000 fibroblast cells (between passages 2 and 5) were seeded in a well of a six-well dish and allowed to recover overnight. The next day, cells were infected by the Sendai virus expressing human transcription factors Oct4, Sox2, Klf4, and C-Myc mixed in fibroblast medium according to the manufacturer’s instructions. Two days later, the medium was exchanged for human ES medium supplemented with the ALK5 inhibitor SB431542 (2 µM; Stemgent), the MEK inhibitor PD0325901 (0.5 µM; Stemgent), and thiazovivin (0.5 µM; Stemgent). Human ES medium contained KO-DMEM (Gibco 10829), 15% KnockOut Serum Replacement (Gibco, 10828), 1% GlutMAX, 0.1% 2-Mercaptoethanol, 1% NEAA, 1% PenStrep and 0.1 µg/mL bFGF (all from Gibco). On days 7–10 post-infection, cells were detached using TrypLE™ Express (Gibco, 12605036) and passaged onto mouse embryonic fibroblast feeder cells (GlobalStem CF-1 MEF IRR). Individual colonies of induced pluripotent stem cells were manually picked between day 21 and 28 post-infection and each iPSC line was expanded from a single colony. All stem cell lines were cultured on a human ES medium. Cytogenic analysis was performed on 20 G-banded metaphase cells from each line by Cell Line Genetics Inc (Fig. S2m).

Stem cell culture

hESC and iPSCs were grown on plates coated with primary mouse embryonic fibroblasts or MEFs (GlobalStem, CF-1 MEF IRR) and dissociated every 4–5 days using TrypLE™ Express (Life Technology, 12605036) for passaging. After dissociation, cells were suspended in a human ES medium containing 10 µM ROCK inhibitor Y27632 (Selleckchem, S1049).

In silico gRNA design

sgRNAs (Supplementary Data 1) were designed using an online CRISPR design tool (crispr.mit.edu) and cloned into a gRNA cloning vector (Addgene, 41824) following option B from the gRNA Synthesis Protocol⁴⁹. The resulting vector was transformed into competent bacteria using Gibson Assembly^® chemical transformation protocol (E5510). Single clones were picked and grown on 3 ml of LB broth for 16–18 h at 37 °C in a shaker (250 rpm); DNA is extracted and Sanger sequenced.

CRISPR/Cas9 nucleofection and mutagenesis

Stem cells were cultured with human ES media containing ROCK inhibitor Y27632 (Selleckchem, S1049) 3 h prior to nucleofection, dissociated, and filtered through a 70 µm cell strainer (Thermo Fisher Scientific, 8-771-2). Approximately 2 × 10⁶ cells were nucleofected (Lonza nucleofector, program A23) with 5 µg of Cas9-GFP plasmid (Addgene, 44719), 5 µg of sgRNA and 5 µg of ssDNA donor template (Supplementary Data 1) using Human Stem Cell Nucleofector^TM kit 1 (Lonza, VVPH-5012) according to the manufactures protocol and cells replated. Cells were dissociated 48 h later for GFP sorting using BD FACS Aria II cell sorter. As a quality control step, some unsorted cells were used to test the sgRNAs mutation efficiency using the Transgenomic SURVERYOR^® mutation detection kit according to the manufacturer's protocol. Sorted cells were plated in a 10 cm dish and single clones were picked 7–10 days post sorting, clones were further expanded and DNA was extracted for PCR using HNF1A primers (Supplementary Data 1). Amplicons were sent for Sanger sequencing to GENEWIZ and clones with indels were further validated by TOPO^® TA cloning (Thermo Scientific, 450641) (at least 6 clones were picked) followed by Sanger sequencing. For the hESC WT line, five different clonal lines were used for analysis throughout the study.

Differentiation into pancreatic endocrine cells

Cells were grown to 80–90% confluency, dissociated, and suspended in mTeSR™ medium (STEMCELL Technology, 05850) with 10 µM ROCK inhibitor Y27632 (Selleckchem, S1049) and plated in a 1:1 ratio into Matrigel-coated (Fisher Scientific, 354277) wells for differentiation. Differentiation was performed using a published protocol²⁰. The initial stages of differentiation were conducted in planar culture (d0–d11). For the definitive endoderm stage (d1–d3) cells were cultured using STEMdiffTM Definitive Endoderm Differentiation Kit (Stemcell Technologies, 05110). For primitive gut stage (d4–d6), cells were cultured in RPMI containing GlutaMAX (Life Technology, 61870-127), 1% (v/v) Penicillin–Streptomycin (PS) (Thermo Fisher Scientific, 15070-063), 1% (v/v) B27 Serum-Free Supplement (50×) (Life Technology, 17504044) and 50 ng/ml FGF7 (R&D System, 251-KG). For posterior foregut stage (d7 and d8), cells were cultured in DMEM -containing GlutaMax, 1% (v/v) PS, 1% (v/v) B27, 0.25 μM KAAD-Cyclopamine (Stemgent, 04-0028), 2 μM Retinoic acid (Stemgent, 04-0021) and 0.25 μM LDN193189 (Stemgent, 04-0074). For pancreatic progenitor stage (d9–d11), cells were cultured in DMEM containing GlutaMax, 1% (v/v) PS, 1% (v/v) B27, and 50 ng/ml EGF (R&D System, 236-EG). Cells were then dissociated using TrypLE™ Express (Life Technology, 12605036) and seeded into low-attachment 96 well-plates (Corning, 7007) (1 well of six-well-plate to 60 wells of 96-well-plate) for clustering step to form aggregates or clusters of endocrine cells in DMEM containing GlutaMax, 1% (v/v) PS, 1% (v/v) B27, 0.25 μM Cyclopamine, 1 μM thyroid hormone (T3) (Sigma, T6397), 10 μM Alk5i, 10 μM Zinc sulfate (Sigma-Aldrich, Z4750) and 10 μg/ml Heparin (Sigma-Aldrich, H3149) for 2 days (d12–d13). For pancreatic endocrine stage (d14–d20) cells were cultured using DMEM containing GlutaMax, 1% (v/v) PS, 1% (v/v) B27, 100 nM LDN, 1 μM T3, 10 μM Alk5i, 10 μM Zinc sulfate, 10 μg/ml Heparin and 100 nM gamma-secretase inhibitor (DBZ) (EMD Millipore, 565789). For mature pancreatic endocrine stage (d21–d27) cells were cultured using DMEM-containing GlutaMax, 1% (v/v) PS, 1% (v/v) B27, 1 μM T3, 10 μM Alk5i, 10 μM Zinc sulfate, 10 μg/ml Heparin, 1 mM N-acetyl cysteine (N-Cys) (Sigma-Aldrich, A9165-5G), 10 μM Trolox (EMD Millipore, 648471-500MG) and 2 μM R428 (Tyrosine kinase receptor AXL inhibitor) (ApexBio, A8329). From d1 to d11 media was changed every day and from d12 to d27 media was changed every other day. All differentiations were done for 27 to 30 days.

In vitro insulin secretion and content

Static insulin secretion assay was performed on days 27–30 of differentiation; 10 islet-like clusters of cells were used per experiment. Islet-like clusters were pre-incubated for 1 h in Kreb’s Ringer Buffer (128 mM NaCl, 5 mM KCl, 2.7 mM CaCl₂, 1.2 mM MgSO₄, 1 mM NaHPO₄, 1.2 mM KH₂PO₄, 5 mM NaHCO₃, 10 mM HEPES, 0.1% Bovine Serum Albumin, pH = 7.4) containing 3.3 mM glucose, washed and incubated for another hour in 3.3 mM glucose and the medium was collected. Subsequently, 200 µl of buffer containing 16.7 mM glucose or 400 µM tolbutamide (abcam, ab120278) or 30.5 mM KCl was used to treat cells for 1 h, after which the medium was collected. Insulin content was measured by acid ethanol extraction; cells were resuspended in 50 µl of water and sonicated for 15 s. The sonicate is mixed with acid ethanol (0.18 M HCl in 96% ethanol (vol/vol)), in a 1:3 ratio of sonicate to acid ethanol. The mixed solution is incubated at 4 °C for 12 h. Human C-peptide, human insulin, and human proinsulin secretion and content were measured using Ultrasensitive C-peptide ELISA (Mercodia, 10-1141-01), Insulin ELISA (Mercodia, 10-1113-01), and Proinsulin ELISA (Mercodia, 10-1118-01) kit according to the manufacturer’s protocol. Mouse insulin was measured using an Ultrasensitive Insulin ELISA (Mercodia, 10-1249-01) kit according to the manufacturer's protocol. All samples were handled the same way.

Dynamic insulin secretion

Perifusion was performed as follows and according to ref. ⁵⁰. Twenty randomly chosen clusters of cells were examined using a Biorep Technologies (Miami, FL) perifusion system. Clusters were perifused with Kreb’s buffer [115 mM NaCl, 5 mM KCl, 24 mM NaHCO₃, 1 mM MgCl₂, 2.2 mM CaCl₂ at pH 7.4] supplemented with 0.17% bovine serum albumin and 3.3 mM glucose (26 min), followed by 16.7 mM glucose (35 min), 3.3 mM glucose (15 min), 400 µM tolbutamide in 3.3 mM glucose (35 min), 20 mM KCl plus 3.3 mM glucose (10 min) and finally 3.3 mM glucose (15 min). The medium was collected at a flow rate of 100 µl/min to assess insulin secretion. Insulin concentration was measured using an insulin ELISA kit (Alpco, 80-INSHU-E01.1). Clusters of cells were collected at the end of the study and placed in acidified ethanol overnight to determine total insulin levels. All samples were handled the same way.

Calcium Imaging

Stem cell-derived β-cells were dissociated from clusters and plated onto 35 mm glass-bottom dishes coated with 5% Matrigel (Fisher Scientific, 354277). Cells were washed twice with basal KRBH solution composed of (mM): 128 NaCl, 5 KCl, 2.7 CaCl₂, 1.2 MgSO₄, 1 NaHPO₄, 1.2 KH₂PO₄, 5 NaHCO₃, 10 HEPES, 0.1% Bovine Serum Albumin, 3.3 glucose, pH to 7.4. Cells were then incubated in the same solution containing 1 µM fura-2, AM (Thermo Fischer, F1221) with 0.05% pluronic F-127 in DMSO (Thermo Fischer, P3000MP) for 15 min at 37 °C, 5% CO₂. Cells were washed twice with basal KRBH solution and then imaged on an inverted Nikon Ti-eclipse microscope with a Nikon Plan fluor ×20 objective (0.45 NA). Fura-2 measurements were collected at excitation wavelengths of 340 and 380 nm using EasyRatioPro (HORIBA Scientific). Stimulation solutions included either 16.7 mM glucose, 600 µM tolbutamide, or 30.5 mM KCl, with NaCl concentrations adjusted accordingly to balance osmolarity with KRBH solution.

Calcium concentrations were calculated as follows:

$$\left[{{\rm {{Ca}}}}^{2+}\right]={K}_{\rm {{d}}}\cdot \frac{R-{R}_{{\min }}}{{R}_{{\max }}-R}\cdot \frac{{S}_{{\rm {f}}2}}{{S}_{{\rm {b}}2}}$$

(1)

where K_d, the apparent Ca²⁺ binding affinity to Fura-2, was assumed to be 225 nM and R_max, R_min, S_f2, and S_b2 values were obtained using 10 μm ionomycin with either no Ca²⁺ or 2 mM Ca²⁺, values^51,52.

Immunohistochemistry

Clusters of cells and tissue (graft or pancreas) were fixed with 4% paraformaldehyde for 20 min at room temperature or 24 h at 4 °C respectively, washed in PBS, then dehydrated with 30% sucrose for 24 h. Cells and tissues were washed in PBS, then cryopreserved in frozen OCT and sectioned at 7 µm on microscope slides. Slides were then incubated with a blocking solution containing 3% donkey serum with 0.1% triton X-100 diluted in PBS for 1 h at room temperature. Primary antibodies (Supplementary Data 7) were added in block solution overnight at 4 °C, washed three times for 5 min in PBST (PBS with 0.1% Tween-20) and incubated with secondary antibodies (Supplementary Data 7) diluted in block solution for 1 h at room temperature, washed three times in PBST containing 10 µg/ml DAPI for 5 min and mounted with a coverslip in fluorescent mounting media (Dako, S3023). All images were taken using a Zeiss LSM 710 confocal microscope and quantified manually using ImageJ software. Quantification of cell type at the definitive and pancreatic progenitor stage (planar culture) was done by randomly choosing 4 sections in the well and averaging the percentage of cells from all sections. For clusters of cells (3D culture) and tissue (graft or pancreas), they were cut in 3–6 sections spaced 20 and 100 µm, respectively, and cell type was quantified in each section as a percentage. The final quantification result is the average of all sections. The ratio of hormone-positive cells for each insulin and glucagon over the total number of cells identified by DAPI nuclear staining was quantified using ImageJ and reported as the fraction of total pancreatic parenchyma for each hormone (i.e. hormone-positive area). Each cluster of cells is comprised of ~10,000 cells.

Flow cytometry

Clusters of endocrine cells were dissociated into single cells using TrypLE™ Express (Life Technology, 12605036). Cells were then fixed with 4% paraformaldehyde for 20 min at room temperature followed by 10 min permeabilization with cold methanol at −20 °C. Cells were washed with 3% donkey serum diluted in PBS and primary antibodies diluted (Supplementary Data 7) in a blocking solution containing 3% donkey serum with 0.1% triton X-100 diluted in PBS overnight at 4 °C. For tunel assay (Biotium, 30074) cells were incubated with fluorescent biotinylated nucleotide conjugate according to the manufacturer’s instructions. Cells were washed in PBST (PBS with 0.1% Tween-20) and incubated with secondary antibodies diluted (Supplementary Data 7) in block solution for 1 h at room temperature. Cells were washed and filtered with a BD Falcon 12 × 75 mm tube with a cell strainer cap (BD Biosciences, 352235) and analyzed by flow cytometer BD Fortessa. Data were analyzed using FlowJo software. Gating for flow cytometry was determined by incubating the cells only with secondary antibodies (negative control) and was consistent across experiments (shown in magenta in the figures).

Western Blot

Clusters of endocrine cells were dissociated into single cells using TrypLE™ Express (Life Technology, 12605036). Cells were lysed using RIPA buffer (NP40 1%, NaCl 150 mM, EDTA 1 mM, Tris–HCl pH 7.5 50 nM, SDS 0.1%, sodium deoxychol 0.5%, NaF 10 mM, protease inhibitor tablet) and whole-cell lysates obtained by subsequent centrifugation. Immunoblots were incubated with primary antibodies against HNF1A (abcam, ab96777, 1:500) and β-Tubulin-III (Sigma, T2200, 1:500) (Supplementary Data 7). Raw images are provided in Fig. S14.

Bulk RNA sequencing

Around 200 clusters of cells were dissociated and sorted for GFP using BD FACS Aria II cell sorter. Sorted cells were pelleted by centrifugation and RNA was extracted using total the RNA purification micro (Norgen Biotek, 35300) kit according to the manufacturer's protocol. RNA quality and concentration are determined by an Agilent bioanalyzer and RNAseq performed by the Columbia University Genome Center. A poly-A pull-down was used to enrich mRNAs from total RNA samples (200 ng–1 µg per sample, RIN > 8 required) and proceed with library preparation by using the Illumina TruSeq RNA prep kit. Libraries were then sequenced using an Illumina HiSeq2000. RTA (Illumina) was used for base calling and bcl2fastq (version 1.8.4) for converting BCL to fastq format, coupled with adaptor trimming. The reads were mapped to a reference genome (Human: NCBI/build37.2; Mouse: UCSC/mm9) using Tophat⁵³ (version 2.1.0) with 4 mismatches (–read-mismatches = 4) and 10 maximum multiple hits (–max-multihits = 10). To tackle the mapping issue of reads that are from exon-exon junctions, Tophat infers novel exon-exon junctions ab initio and combines them with junctions from known mRNA sequences (refgenes) as the reference annotation. The relative abundance (aka expression level) of genes and splice isoforms is estimated using cufflinks⁵⁴ (version 2.0.2) with default settings. Differentially expressed genes were tested under various conditions using DEseq⁵⁵. An R package based on a negative binomial distribution that models the number reads from RNA-seq experiments to test for differential expression.

Single-cell RNA sequencing

Around 100 clusters of cells per genotype at day 27 of differentiation were dissociated and sorted for GFP using BD FACS Aria II cell sorter. Single-cell RNA-Seq data from a total of 271 hESC-derived cells (113 for WT and 158 for KO) were obtained. Raw reads were aligned to the hg19 reference genome using STAR (2.5.2b) and gene quantification was performed using FeatureCounts (1.5.2). Quality control was conducted as follows: cells with low mapped reads (<1,000,000), low exon count (<100,000), low exon mapped rate (<0.2), high intergenic mapped rate (>0.3), low detected gene count (<1000) and with more than 10% Mitochondria genes’ count were removed. In addition, genes with <10 non-zero counts across all the cells were eliminated. All the downstream analysis was done within R-3.4.1. Filtered raw count data were normalized and scaled by the Seurat package and only variable genes generated by Seurat were used to do PCA and t-SNE. n = 3 for each genotype.

10X genomics single-cell RNA sequencing

Around 100 clusters of cells per genotype at day 27 of differentiation were dissociated. Single cells were suspended in PBS + 0.04% BSA. Cellular suspensions (~6000 cells) were loaded on a Chromium Single Cell Instrument (10X Genomics) to generate single cell GEMs. Single-cell RNAseq libraries were prepared using Chromium Single cell 3¢ Library, Gel beads & Multiplex kit (10X Genomics). Sequencing was performed on Illumina NextSeq500 using the following read length: 59-bp Read1 for transcript read, 14-bp I7 Index for Cell Barcode read, 8-bp I5 Index for sample index read, and 10-bp Read2 for UMI read. The samples were sequenced in ten different libraries (batches). Among all these samples, Batch 980379 and Batch 993091 (2 libraries) are HNF1A mutants (R200Q/R200Q), Batch 980380 and Batch 993090 (2 libraries) are HNF1A knockouts (KO) while Batch 989516 and Batch 993089 (2 libraries) are HNF1A wild type cells (WT). In addition, to increase the sequencing depth, each batch was sequenced from 7 flowcells. The raw fastq files were processed via 10X Cell Ranger pipeline (cellranger-2.1.1). To note that GRCh38 reference genome was applied for the whole analysis. “cellranger count” with “–expect-cells = 3000” was run for each library, respectively, and “cellranger aggr” with default parameters was run to aggregate results generated from multiple libraries. A normalized umi-count matrix for all the cells was then generated. As for the quality control, cell and gene filtering process, cells with umi-count <5000 and umi-count >40,000 and cells with <2000 detected genes were removed. Also, genes with <~1% (25) non-zero counts across all the cells were eliminated from the analysis. To get rid of the Mitochondria cells, cells with more than 10% Mitochondria genes’ count were also removed. All the downstream analysis was done with Seurat package in R-3.4.1. Every batch is an independent differentiation (biological replicates). n = 2 for each genotype.

Cell transplantation and in vivo assays

After 27–30 days of differentiation, 50 µg of stem cell-derived pancreatic cell clusters (~180 islet-like clusters of 300–350 µm diameter; each cluster comprised ~10,000 cells in total) including all islet endocrine cell types were suspended in 30 µl of ice-cold Matrigel (Fisher Scientific, 354277) and loaded into ice-cold 1 ml syringe with a 21 G needle for transplantation. 6–10 weeks old male immunodeficient NSG (NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ, Stock No. 005557, The Jackson Laboratories) mice were transplanted in the ventral and medial muscles (medial thigh and posterior lower) of the left thigh. Every 2 weeks post-transplantation, human C-peptide and human insulin were determined in the plasma of recipient mice at a fed state (morning) for 30 weeks. An intraperitoneal glucose tolerance test was performed 18–24 weeks (before STZ injection) and 28–30 weeks (after STZ injection) post-transplantation by injecting 2 g/kg body weight of 20% d-glucose (Sigma, G8270) in PBS overnight fasting (16–18 h). For glibenclamide (Sigma, G0639) administration, 1 mg/kg body weight of the drug diluted in PBS was i.p. injected 24–28 weeks (before STZ injection) post-transplantation, all mice were at fed state but the food was retrieved during the experiment. Blood was collected by tail bleeding in heparinized Eppendorf tubes at time points; plasma was isolated by centrifugation for 15 min at 2000 × g at 4 °C. Human C-peptide was measured using Mercodia Ultrasensitive C-peptide ELISA (10-1141-01) and human insulin using Mercodia Insulin ELISA (10-1113-01) kit according to the manufacturer's protocol. All samples were handled the same way. The insulin detection limit is 1 mU/L as determined by the methodology described in the kit. Blood glucose levels were measured using a glucometer (FreeStyle Lite) and HbA1C using Siemens DCA 2000 Vantage Reagent kit (Siemens, 5035C). To induce diabetes, mice were i.p. injected for 5 consecutive days with 40 mg/kg body weight of Streptozotocin (STZ) (Sigma-Aldrich, S0130-1G) in PBS 24–28 weeks post-transplantation. All experimental procedures on mice were performed according to Columbia University-approved IACUC protocols.

Bioluminescence and fluorescence imaging

NSG mice transplanted with GAPDH^{Luciferase/wt} and INS^GFP/wt double reporter hESCs lines were i.p. injected with 150 mg/kg body weight of d-luciferin potassium salt (Gold Biotechnology, luck-2G) in PBS 15 min before imaging on a IVIS spectrum optical imaging system (PerkinElmer). Signals were acquired with 1-min exposure and analyzed using the Living image analysis software (Xenogen Corp.). Circular regions of interest (ROI) of the same size for all experiments were drawn around the signal in the left thigh and photons emitted over the time of exposure within the ROI measured. Bioluminescence was measured every 2 weeks post-transplantation for 30 weeks and after isolation of the graft at that time point. Luminescence was measured as described for bioluminescence after isolation of the graft. Background signals were subtracted from a nearby region and were consistent over time.

Electron microscopy

Fixed cells/tissues were incubated with 2.5% glutaraldehyde and 2% paraformaldehyde in 100 mM sodium cacodylate buffer (pH 7.4) overnight. Samples were then treated with 1% osmium tetroxide in 100 mM sodium cacodylate buffer for 1 h, washed in distilled water four times (10 min/wash), and then treated with 2% aqueous uranyl acetate overnight at 4 °C in the dark. Samples were washed and sequentially dehydrated with increasing concentrations of acetone (20%, 30%, 50%, 70%, 90%, and 100%) for 30 min each, followed by three additional treatments with 100% acetone for 20 min each. Samples were then infiltrated with increasing concentrations of Spurr’s resin (25% for 1 h, 50% for 1 h, 75% for 1 h, 100% for 1 h, 100% overnight at room temperature), and then incubated overnight at 70 °C in a resin mold. Sections of 50–90 nm were cut on a Leica ultramicrotome with a diamond knife. Imaging was performed on an FEI Talos L120C operating at 120 kV.

Immunogold electron microscopy

70 nm-thick sections from the embedded samples were placed onto carbon formvar 75 mesh nickel grids and etched using 4% sodium metaperidotate for 10 min before being washed twice in distilled water and then blocked for 1 h. Grids were incubated with the primary C-peptide antibody (Supplementary Data 7) (1 in 10 dilutions) for 1 h at 4 C overnight. The next day grids underwent seven washes in 1xPBS and then incubated in anti-rat 6 nm gold secondary (1 in 50 dilutions) for 1 h (Supplementary Data 7). After this, the grid was washed seven times in 1xPBS and twice in distilled water. Samples were then imaged on a Thermo Fisher Talos L120C operating at 120 kV.

Statistics and reproducibility

All statistics were performed using Prism GraphPad software (La Jolla, CA). Normality of data set distributions was assessed by Shapiro–Wilk test and D’Agostino and Pearson omnibus tests. Normally distributed data were analyzed by unpaired two-tailed t-test. For data not normally distributed we used a two-tailed Mann–Whitney test. Data were expressed as mean ± standard error of the mean (SEM). p < 0.05 was considered statistically significant. *p < 0.05, **p < 0.01, ***p < 0.001. n.s. indicates a non-significant difference.

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability

Single-cell RNA sequencing and bulk RNA-sequencing data were deposited in NCBI’s Gene Expression Omnibus (GEO) database and the accession number is GSE128331 and GSE129653. Additional data from Haliyur et al. ²⁹ GSE116559 were used. Unprocessed blots can be found in Fig. S14 and source data is available in Supplementary Data 8.

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Acknowledgements

We would like to thank Giacomo Diedenhofen for help with western blots and Qian Du for help with differentiation and flow cytometry. This research was supported by the American Diabetes Association (grant #1-16-ICTS-029) and the NYSTEM IDEA award # C029552, Leona and Harry Helmsley Charitable Trust, Helmsley Trust Diabetes Cell Repository, the NYSCF-Robertson award from the New York Stem Cell Foundation, the Sanofi iAwards, and the Naomi Berrie Foundation program for Cellular Therapies of Diabetes. These studies used the resources of the Herbert Irving Comprehensive Cancer Center Flow Cytometry Shared Resources funded in part through Center Grant P30CA013696 and the Diabetes and Endocrinology Research Center Flow Core Facility funded in part through DRC Center Grant (5P30DK063608) and the MBMG Core in the New York Nutrition and Obesity Research Center (5P30DK026687).

Author information

Jesper Gromada
Present address: Vertex Cell and Genetic Therapies, Watertown, MA, 02472, USA

Authors and Affiliations

Naomi Berrie Diabetes Center & Departments of Pediatrics and Medicine, Vagelos College of Physicians and Surgeons, Columbia University, New York, NY, 10032, USA
Bryan J. González, Maria H. Blumenkrantz, Charles A. LeDuc, Wendy K. Chung, Robin S. Goland, Rudolph L. Leibel & Dieter Egli
Institute of Human Nutrition, Columbia University Medical Center, New York, NY, 10032, USA
Bryan J. González
Department of Systems Biology, Columbia University Medical Center, New York, NY, 10032, USA
Haoquan Zhao, Jaeyop Lee & Yufeng Shen
Department of Physiology and Cellular Biophysics, College of Physicians and Surgeons, Columbia University, New York, NY, 10032, USA
Jacqueline Niu & Henry M. Colecraft
Stem Cell Core Facility, Department of Rehabilitation and Regenerative Medicine, Columbia University, New York, NY, 10032, USA
Damian J. Williams
Center for Dementia Research, Nathan S. Kline Institute, Orangeburg, NY, 10962, USA
Chris N. Goulbourne
Department of Surgery, University of Virginia, Charlottesville, VA, 22908, USA
Yuan Xing, Yong Wang & Jose Oberholzer
Columbia Center for Translational Immunology, Department of Surgery, Columbia University Medical Center, New York, NY, 10032, USA
Xiaojuan Chen
Regeneron Pharmaceuticals, Tarrytown, NY, 10591, USA
Jesper Gromada

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Henry M. Colecraft
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Jesper Gromada
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Yufeng Shen
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Robin S. Goland
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Dieter Egli
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Contributions

B.J.G. designed the studies, performed differentiation, transplantation of all cell lines and downstream experiments, gathered the data, analyzed the data, and wrote the paper with input from authors. D.E. designed the studies, discussed results, reviewed and edited the manuscript. D.E. performed skin biopsy stem cell derivation. J.G. performed single-cell RNA sequencing and B.J.G., H.Z., J.L., and Y.S. designed, analyzed, and compiled the single-cell RNA-sequencing data. B.J.G. and C.N.G. performed EMC imaging. W.K.C. provided genetic sequence information of HNF1A-MODY subjects. Y.X., Y.W., and J.O. performed microfluidic perfusion and generated data. M.H.B. performed flow cytometry of HNF1A-WT and HNF1A-KO. J.N., D.J.W., and H.M.C. performed calcium imaging, measurements, and data interpretation. X.C. contributed to human islets. R.S.G. performed oversight of human subject research and provided de-identified clinical information. D.E., C.A.L., and R.L. provided guidance in the design and interpretation of studies.

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Correspondence to Dieter Egli.

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González, B.J., Zhao, H., Niu, J. et al. Reduced calcium levels and accumulation of abnormal insulin granules in stem cell models of HNF1A deficiency. Commun Biol 5, 779 (2022). https://doi.org/10.1038/s42003-022-03696-z

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Received: 20 May 2021
Accepted: 11 July 2022
Published: 02 August 2022
DOI: https://doi.org/10.1038/s42003-022-03696-z

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