Heparan sulfate fine-tuned interleukin-1 (IL-1) signaling inhibits insulin secretion of grafted pancreatic islets

Selina Wrublewsky; Sandra Rother; Franziska Pohlemann; Toni Radanovic; Fabian Junker; Anne S Boewe; Stefan Schunk; Leticia P Roma; Gloria Ruiz-Gómez; M Teresa Pisabarro; Patrick E MacDonald; Michael D Menger; Matthias W Laschke; Emmanuel Ampofo

doi:10.1126/sciadv.ady8566

. 2025 Aug 8;11(32):eady8566. doi: 10.1126/sciadv.ady8566

Heparan sulfate fine-tuned interleukin-1 (IL-1) signaling inhibits insulin secretion of grafted pancreatic islets

Selina Wrublewsky ¹, Sandra Rother ², Franziska Pohlemann ¹, Toni Radanovic ², Fabian Junker ², Anne S Boewe ¹, Stefan Schunk ³, Leticia P Roma ⁴, Gloria Ruiz-Gómez ⁵, M Teresa Pisabarro ⁵, Patrick E MacDonald ^6,⁷, Michael D Menger ¹, Matthias W Laschke ¹, Emmanuel Ampofo ^1,^*

^¹Institute for Clinical and Experimental Surgery, Saarland University, 66421 Homburg/Saar, Germany.

^²Center for Molecular Signaling, Präklinisches Zentrum für Molekulare Signalverarbeitung (PZMS), Saarland University, 66421 Homburg/Saar, Germany.

^³Department of Internal Medicine IV (Nephrology and Hypertension) and Translational Cardio-Renal Medicine, Saarland University, 66421 Homburg/Saar, Germany.

^⁴Biophysics Department, Center for Human and Molecular Biology, Saarland University, 66421 Homburg/Saar, Germany.

^⁵Structural Bioinformatics, BIOTEC TU Dresden, Tatzberg 47-51, 01307 Dresden, Germany.

^⁶Department of Pharmacology and Alberta Diabetes Institute, University of Alberta, Edmonton, Alberta, Canada.

^⁷Division of Metabolism, Endocrinology and Diabetes, University of Michigan, Ann Arbor, MI, USA.

Corresponding author. Email: emmanuel.ampofo@uks.eu

Roles

Selina Wrublewsky: Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Resources, Validation, Visualization, Writing - original draft, Writing - review & editing

Sandra Rother: Conceptualization, Investigation, Methodology, Visualization, Writing - review & editing

Franziska Pohlemann: Investigation

Toni Radanovic: Investigation

Fabian Junker: Investigation

Anne S Boewe: Investigation

Stefan Schunk: Conceptualization, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Validation, Writing - original draft, Writing - review & editing

Leticia P Roma: Conceptualization, Investigation, Resources, Supervision, Writing - review & editing

Gloria Ruiz-Gómez: Data curation, Formal analysis, Investigation, Methodology, Resources, Validation, Visualization, Writing - original draft, Writing - review & editing

M Teresa Pisabarro: Data curation, Formal analysis, Investigation, Methodology, Project administration, Resources, Supervision, Validation, Visualization, Writing - original draft, Writing - review & editing

Patrick E MacDonald: Conceptualization, Data curation, Resources, Writing - original draft

Michael D Menger: Conceptualization, Funding acquisition, Methodology, Project administration, Resources, Supervision, Writing - review & editing

Matthias W Laschke: Conceptualization, Methodology, Resources, Supervision, Visualization, Writing - original draft, Writing - review & editing

Emmanuel Ampofo: Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Software, Supervision, Validation, Visualization, Writing - original draft, Writing - review & editing

PMCID: PMC12333676 PMID: 40779638

Abstract

Islet-resident macrophages contribute to hypoxia-induced islet cell death during pancreatic islet transplantation. However, their specific role during this process remains elusive. Here, we report that interleukin-1α (IL-1α) and IL-1β are released by islet-resident macrophages, resulting in the suppression of insulin secretion. This may be due to a decreased inflammation-driven expression of pancreatic and duodenal homeobox 1 (PDX-1) and MafA in β cells. Islet-resident macrophages release significantly less IL-1α when compared to IL-1β. However, both cytokines inhibit insulin expression and secretion to a comparable extent. We identified heparan sulfate on the islet surface, which acts as a “molecular glue” potentiating the inhibitory action of IL-1α on insulin expression via specific binding to IL-1 receptor (IL-1R). In vivo analyses revealed that the loss of IL-1 signaling in isolated islets accelerates their revascularization and, thus, enhances their endocrine function. These findings indicate that heparan sulfate fine-tuned IL-1 signaling crucially determines the outcome of islet transplantation.

The binding of heparan sulfate to IL-1α potentiates the inhibitory action of IL-1α on insulin release.

INTRODUCTION

Islet transplantation represents a promising therapeutic approach to improve glycometabolic control in patients with type 1 diabetes mellitus (T1DM) (1). However, hypoxia-induced inflammatory graft failure is a major obstacle preventing long-term graft survival and function (2–7). To overcome this problem, various anti-inflammatory strategies have been developed in both experimental (5, 8) and clinical studies (9). In this context, we and others have introduced an anti-inflammatory approach that accelerates normoglycemia in diabetic animals after islet transplantation by inhibiting the nucleotide-binding oligomerization domain–like receptor protein 3 (NLRP3) inflammasome (10, 11).

The NLRP3 inflammasome and other inflammasome family members are highly expressed in circulating monocytes and tissue macrophages (12). Accordingly, intraislet inflammation is thought to be of immune cell origin. In line with this view, islets contain a small fraction of resident macrophages (13), which are in a persistent inflammatory state, however, without producing high levels of inflammatory cytokines (14–16). Recent studies revealed that islet-resident macrophage-derived proinflammatory cytokines are overproduced in pancreatic islets during obesity, aging, type 2 diabetes mellitus (T2DM), and T1DM (15–17). Hence, macrophage depletion or the use of cytokine-neutralizing antibodies has been shown to prevent the development of T1DM (18, 19).

Interleukin-1α (IL-1α) and IL-1β are potent inflammatory cytokines with immune response-amplifying effects (20). The two cytokines are mainly released by macrophages and trigger signaling transduction via binding to the IL-1 receptor (IL-1R) complex with a comparable affinity (21–23). However, despite these similarities, IL-1α and IL-1β differ in their biological relevance. IL-1β is only active in its extracellular and secreted form and is capable of recruiting macrophages, whereas IL-1α is a classic damage-associated molecular pattern, acts as an alarmin, and promotes neutrophil recruitment (24–26). Besides its role as a soluble cytokine, IL-1α has additional functions. For instance, nuclear IL-1α is capable of stimulating gene transcription by interacting with different transcription activators (27, 28). On the other hand, IL-1α can be found on the cell membrane, suggesting that it also plays an important role in cell-cell communication (29). Accordingly, we recently identified membrane-bound IL-1α on monocytes as a central regulator of inflammation in cardiorenal diseases (30). However, the mechanisms mediating the translocation of IL-1α to the cell surface and its binding to the plasma membrane are still under debate.

In this study, we identified islet-resident macrophages as the major source of IL-1α and IL-1β within pancreatic islets. Moreover, we found that IL-1–mediated signal transduction suppresses insulin secretion in a paracrine mode of action. By using different mouse models of islet transplantation, we could show that loss of IL-1α or IL-1β in islet-resident macrophages not only does accelerate the restoration of normoglycemia in diabetic mice but also promotes the revascularization of islet grafts in an insulin-dependent manner. In addition, we identified the selective binding of IL-1α, but not IL-1β, to heparan sulfate on the islets’ surface as a previously unknown mechanism potentiating the inhibitory action of the cytokine on insulin expression. Together, these findings highlight IL-1 signaling as a promising target to improve the future outcome of clinical islet transplantation.

RESULTS

Loss of IL-1α and IL-1β does not affect the viability and cellular composition of isolated islets

First, we proved by means of calcein/propidium iodide (PI) stainings (Fig. 1, A and B) and neutral red/trypan blue stainings (fig. S1) that isolated islets from wild-type (WT), IL-1α^−/−, and IL-1β^−/− mice exposed to hypoxia exhibit a comparable viability. We further confirmed by flow cytometry that the loss of IL-1–mediated signaling does not induce apoptosis or necrosis (Fig. 1, C and D). The analysis of the absolute islet cell number (fig. S2) and the cellular composition of isolated hypoxic WT, IL-1α^−/−, and IL-1β^−/− islets (Fig. 1, E and F) as well as of islets within the pancreas (fig. S3) demonstrated no difference between the groups. Additional analyses performed under normoxic conditions showed comparable results (fig. S4, A to C).

IL-1α and IL-1β suppress the gene expression and secretion of insulin

We first measured plasma insulin levels of fasted WT, IL-1α^−/−, and IL-1β^−/− mice by means of an intraperitoneal glucose tolerance test (IPGTT). We found increased plasma insulin levels in IL-1α^−/− and IL-1β^−/− mice when compared to those in WT controls (fig. S5). Next, we examined whether IL-1 signaling affects the insulin secretion of isolated islets exposed to hypoxia (1% O₂) for 16 hours. As expected, the loss of IL-1α and IL-1β ameliorated glucose-stimulated insulin secretion (GSIS) to a comparable extent, whereas the exposure of islets to recombinant IL-1α and IL-1β markedly reduced the endocrine function of WT islets (Fig. 2A). The exposure of IL-1α^−/− or IL-1β^−/− islets to IL-1α or IL-1β as well as of WT islets to IL-1R antagonist (IL-1RA) showed that the observed altered insulin secretion is mediated via the IL-1R complex (Fig. 2A). We additionally analyzed the effect of the two cytokines on insulin secretion from human islets. As expected, IL-1α or IL-1β also significantly repressed the islets’ C-peptide secretion (fig. S6). Of note, we did not observe any effects on murine islets under basal glucose conditions (fig. S7), indicating that a disturbed IL-1 signaling does not per se increase insulin secretion. Further, we could exclude that the reduced insulin secretion is caused by IL-1α or IL-1β–induced cell death (fig. S8).

Fig. 2. — (A) Quantitative analysis of insulin secretion (microunits per milliliter) from hypoxic WT, IL-1α^−/−, and IL-1β^−/− islets as well as hypoxic WT islets exposed to IL-1α, IL-1β, or IL-1RA, hypoxic IL-1α^−/− islets exposed to IL-1β and hypoxic IL-1β^−/− islets exposed to IL-1α (n = 4 each). Means ± SEM. (B) Representative Western blot of p65 and p-p65 from extracts of hypoxic WT and hypoxic WT islets exposed to IL-1α and IL-1β (bottom). Quantitative analysis of p-p65 expression (top). Data are expressed in % of WT (n = 3 each). Means ± SEM. (C and D) Quantitative analysis of pancreatic and duodenal homeobox 1 (*PDX-1*) (C) and *MafA* mRNA expression (D) in hypoxic WT and hypoxic WT islets exposed to IL-1α and IL-1β. Data are expressed as % of WT (n = 3 each). Means ± SEM. (E and F) Representative Western blot of PDX-1 (E) and MafA (F) from extracts of hypoxic WT and hypoxic WT islets exposed to IL-1α and IL-1β (bottom). Quantitative analysis of PDX-1 (E) and MafA expression (F) (top). β-Actin was used as loading control. Data are expressed in % of WT (n = 3 each). Means ± SEM. (G and H) Quantitative analysis of *Ins1* (G) and *Ins2* (H) mRNA expression in hypoxic WT and hypoxic WT islets exposed to IL-1α and IL-1β. Data are expressed as % of WT (n = 3 each). Means ± SEM. (I) Schematic illustration of the underlying mechanism. Hypoxia as well as the presence of IL-1α and IL-1β promotes the binding of p65 to the *PDX-1*- and *MafA*-promoter, which represses their gene expression. The low levels of PDX-1 and MafA, in turn, reduce *insulin* gene expression. Image provided by Servier Medical Art (https://smart.servier.com/), licensed under CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).

Cytokine-induced β cell dysfunction is associated with an impaired expression of β cell–specific genes, such as insulin (31, 32). Therefore, we next studied the effect of IL-1α and IL-1β on nuclear factor κ–light-chain enhancer of activated B cells (NF-κB)–mediated insulin expression. We detected higher levels of phosphorylated p65 (p-p65) in hypoxic islets exposed to IL-1α or IL-1β (Fig. 2B). Furthermore, both gene and protein expression of pancreatic and duodenal homeobox 1 (PDX-1) and MafA, the two major transcription factors for insulin, were diminished (Fig. 2, C to F). On the basis of on this, we also assumed a reduced insulin gene expression. We measured a lower Ins1/2 mRNA expression in IL-1α– and IL-1β–exposed islets (Fig. 2, G and H). Together, these findings indicate that IL-1 signaling may suppress insulin gene expression and secretion by reducing NF-κB activity and down-regulating PDX-1 and MafA expression in hypoxic islets (Fig. 2I).

IL-1α and IL-1β are released from islet-resident macrophages under hypoxic conditions

There is a controversial discussion, whether endocrine cells or resident macrophages within islets express and release IL-1α and IL-1β (17, 33–35). We herein found that mouse islets contain ~3 to 5% islet-resident macrophages, which do not express IL-1α or IL-1β under normoxia (Fig. 3A). However, the exposure of WT islets to hypoxia induced the expression of both cytokines in these cells (Fig. 3A). This was not the case in hypoxic IL-1α^−/− or IL-1β^−/− islets, which served as negative controls (Fig. 3A). Moreover, we found that islet-resident IL-1α–positive macrophages exhibit a lower mean fluorescence intensity (MFI) when compared to IL-1β–positive macrophages, indicating a decreased level of intracellular IL-1α in these cells (Fig. 3B). We additionally assessed the release of IL-1α or IL-1β from WT, IL-1α^−/−, and IL-1β^−/− islets (Fig. 3, C and D). In line with our flow cytometric MFI measurements, we assessed higher levels of IL-1β in the supernatants of WT and IL-1α^−/− islets when compared to the levels of IL-1α in the supernatants of WT and IL-1β^−/− islets. To verify these results, we performed control experiments with hypoxic U937-derived M1 macrophages, which also exhibited a significantly lower release of IL-1α when compared to IL-1β (Fig. 3E). On the basis of our findings showing identical effects of IL-1α and IL-1β on insulin secretion (Fig. 2A) despite a markedly lower release of IL-1α from hypoxic islets compared to IL-1β (Fig. 3, C and D), we speculated that the effect of IL-1α is potentiated by an extracellular mechanism (Fig. 3F).

Fig. 3. — (A) Representative flow cytometry scatterplots of dispersed islets from WT, IL-1α^−/−, and IL-1β^−/− mice cultivated under normoxia and hypoxia. The gating strategy shows the fractions of CD68-positive, CD68/IL-1α–positive, and CD68/IL-1β–positive cells. (B) Quantitative analysis of the MFI from IL-1α–positive cells (blue-bordered bars; left y axis) and IL-1β–positive cells (red-bordered bars; right y axis) from normoxic and hypoxic islets (n = 3 each). Means ± SEM. (C) Quantitative analysis of IL-1α secretion (picograms per milliliter) from normoxic and hypoxic WT, IL-1α^−/−, and IL-1β^−/− islets (blue-bordered bars; n = 3 each). Means ± SEM. n.d., not determined. (D) Quantitative analysis of IL-1β secretion (picograms per milliliter) from normoxic and hypoxic WT, IL-1α^−/−, and IL-1β^−/− islets (red-bordered bars; n = 3 each). Means ± SEM. (E) Quantitative analysis of IL-1α secretion (picograms per milliliter) from normoxic and hypoxic U937-M1 macrophages (blue-bordered bars; left y axis) and IL-1β secretion (picograms per milliliter) from normoxic and hypoxic U937-M1 macrophages (red-bordered bars; right y axis) (n = 3 each). Means ± SEM. (F) Schematic illustration of the proposed mechanism. Islet resident macrophages release markedly lower amounts of IL-1α when compared to IL-1β; however, both cytokines exhibit similar effects on insulin secretion. Therefore, we assume that an inter-islet effect potentiates the activity of IL-1α or attenuates that of IL-1β. Image provided by Servier Medical Art (https://smart.servier.com/), licensed under CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).

IL-1α binds to islet heparan sulfate

Heparan sulfate regulates the activity of cytokines through physical sequestration in the extracellular matrix and protection from enzymatic degradation (36). Moreover, heparan sulfate is required to ensure physiological insulin secretion (37). Therefore, we asked whether this sulfated glycosaminoglycan (GAG) increases the activity of IL-1α in hypoxic islets. To clarify this, we first performed a sandwich enzyme-linked immunosorbent assay (ELISA) to study the physical interaction of IL-1α and GAG (Fig. 4A). To mimic extracellular heparan sulfate, we used heparin (HEP), whereas hyaluronic acid (HA) served as nonsulfated control. Of interest, we found a concentration- and sulfate-dependent binding of IL-1α to GAG surfaces (Fig. 4A). Additional surface plasmon resonance (SPR) binding experiments revealed that HEP preferably binds to IL-1α, while only marginal binding responses were detected for HEP/IL-1β interaction (Fig. 4B). Furthermore, the formed HEP/IL-1α complexes showed a low dissociation over time (Fig. 4C).

On the basis of these findings, the molecular recognition of HEP by IL-1α was investigated through molecular modeling. For this purpose, the electrostatic potential of IL-1α was calculated to first visualize positively charged patches on the protein surface, which could potentially represent sites of recognition for HEP (Fig. 4D). Blind molecular docking was performed to predict binding sites and modes of HEP on IL-1α. For tetrameric HEP, binding poses were predicted in two main recognition sites (i.e., cluster1 and cluster2) surrounding the IL-1α N-terminal region (Fig. 4E), which exhibits a patch of positive electrostatic potential (Fig. 4F). For hexameric HEP, two more disperse clusters were predicted (Fig. 4F), which also coincided with the calculated positively charged patch on the protein surface. The binding poses predicted for tetra- and hexameric HEP were equivalent in one of the clusters (cluster2), whereas a small overlapping of poses was observed for the respective cluster1. Three representative HEP binding modes of each cluster were selected, and the corresponding HEP/protein complexes were energy minimized by molecular dynamics (MD). Calculated binding free energies indicated that binding poses in cluster1, for both tetra- and hexameric HEP, were slightly more favorable than poses in the two cluster2 (fig. S9). Per-residue binding free energy decomposition analysis revealed that, in the respective cluster1, Lys¹¹ and Lys¹⁰⁰ represent common residues of IL-1α relevant for recognizing tetra- and hexameric HEP (fig. S10, A and C). Furthermore, Arg¹⁶ may be relevant for recognition of the hexameric HEP. In the case of the respective cluster2, Asn⁹, Lys¹¹, and Lys⁶³ are the most contributing residues for recognition of tetra- and hexameric HEP (fig. S10, B and D). As IL-1α signals through IL-1R, we further tested whether the binding of IL-1α to IL-1RI and IL-1R–associated protein (IL-1RAcP) is affected by HEP using a competitive sandwich ELISA with IL-1RI and IL-1RAcP without HEP as controls. While HEP did not affect the binding of IL-1α to IL-1RAcP, the presence of the sulfated GAG markedly increased IL-1α binding to IL-1RI (Fig. 4G).

Together, these findings suggest that the interaction of IL-1α with heparan sulfate promotes its binding to IL-1R (Fig. 4H). This was next confirmed in pancreatic islets, which exhibit high levels of extracellular heparan sulfate (Fig. 4I). The loss of heparan sulfate by heparanase (HPSE) treatment significantly increased the amount of soluble IL-1α in the supernatant of hypoxic islets (Fig. 4J). This was not the case for IL-1β (Fig. 4J). To exclude that the observed effects are species specific, we additionally performed these experiments with human islets. Human islets within the pancreas also expressed high levels of heparan sulfate and HPSE treatment solely increased IL-1α and not IL-1β levels in isolated hypoxic human islets (Fig 4, K and L).

To assess the effect of heparan sulfate/IL-1α complex formation on the endocrine function of islets, we exposed hypoxic IL-1α^−/− and IL-1β^−/− islets, which did not differ in their total GAG amounts (fig. S11), to HPSE and subsequently measured GSIS (fig. S12A). In line with previous studies (38), we found that exposure of islets to HPSE reduces the secretion of insulin (fig. S12A). Of interest, this effect was less pronounced in HPSE-treated IL-1β^−/− islets when compared to WT and IL-1α^−/− islets (Fig. 4M). To further verify the binding of heparan sulfate to IL-1α, we used surfen, a small-molecule antagonist of heparan sulfate (39). The exposure of islets to surfen did not affect their viability (fig. S13). Surfen rather blocked the interaction of heparan sulfate with IL-1α, leading to higher insulin secretion in IL-1β^−/− islets when compared to IL-1α^−/− islets (Fig. 4N and fig. S12B). Of note, the inhibitory effect of HPSE and surfen on insulin secretion could also be found in human islets, as demonstrated by a reduced C-peptide secretion (fig. S14). This indicates that both HPSE and surfen may inhibit IL-1α signaling by reducing the interaction between IL-1α and heparan sulfate. Accordingly, insulin secretion is less suppressed in IL-1β^−/− islets. In contrast, the suppressive effect of IL-1 signaling on insulin secretion is still maintained in WT and IL-1α^−/− islets by IL-1β, which does not bind to heparan sulfate.

Loss of IL-1α and IL-1β improves the revascularization of transplanted islets

Insulin triggers angiogenic pathways in endothelial cells (40). Therefore, we assumed that the increased insulin secretion from isolated IL-1α^−/− and IL-1β^−/− islets promotes their revascularization after transplantation. Vice versa, the revascularization process is impaired by the pharmacological blockade of IL-1α/heparan sulfate interaction by means of surfen. To test this, we used a mouse dorsal skinfold chamber model in combination with intravital fluorescence microscopy (Fig. 5A). We detected a higher functional microvessel density and revascularized area throughout the entire 14-day observation period in grafted IL-1α^−/− and IL-1β^−/− islets when compared to WT controls (Fig. 5, B and C). The exposure of WT and IL-1α^−/− islets to surfen before their transplantation reduced their angiogenic activity and endocrine tissue perfusion over time (Fig. 5, B to D). Of note, this effect was less pronounced in IL-1β^−/− islets exposed to surfen, indicating that this small molecule blocks the IL-1α/heparan sulfate interaction and, thus, enhances the insulin-stimulated blood vessel formation (Fig. 5, B to D). Accordingly, loss of IL-1α or IL-1β improved the take rate of the grafts, i.e., the number of engrafted islets in relation to the overall number of transplants per group on day 14 (Fig. 5E). The additional assessment of microhemodynamic parameters showed that the loss of the two cytokines as well as the exposure to surfen do not affect the diameter, centerline red blood cell (RBC) velocity, and volumetric blood flow of microvessels within the grafted islets (fig. S15, A to C). These results demonstrate that IL-1 deficiency accelerates graft revascularization, which is partially mediated by heparan sulfate/IL-1α complex formation.

Fig. 5. — (A) Schematic illustration of the experimental setting. Dorsal skinfold chambers were implanted on day −2 followed by transplantation of WT, IL-1α^−/−, and IL-1β^−/− islets on day 0. Intravital fluorescence microscopy was performed on days 0, 3, 6, 10, and 14 after islet transplantation. On day 14, the tissue was harvested for immunohistochemical stainings. (B) Quantitative analysis of the functional microvessel density (centimeters per square centimeter) of WT, IL-1α^−/−, and IL-1β^−/− islets as well as WT, IL-1α^−/−, and IL-1β^−/− islets exposed to surfen for 24 hours prior transplantation (n = 8 each). Means ± SEM. (C) Quantitative analysis of the revascularized area (square millimeters) of islets as described in (B) (n = 8 each). Means ± SEM. (D) Quantitative analysis of the rhodamine 6G–positive area (% of islet size) within islets as described in (B) (n = 8 each). Means ± SEM. (E) Take rate of islets as described in (B) (% of transplanted islets; n = 8 each) on day 14 after islet transplantation onto the exposed striated muscle tissue. Means ± SEM.

The cellular composition of the grafts on day 14 after transplantation was analyzed by immunohistochemistry. We did not find any differences in the fractions of endocrine cells between IL-1α^−/−, IL-1β^−/− and WT islets (Fig. 6, A and B). However, a significantly higher fraction of intraislet CD31-positive endothelial cells could be detected in the group of IL-1α^−/− and IL-1β^−/− islets when compared to WT controls (Fig. 6, A and B). Additional analyses of the immune cell infiltration after transplantation revealed no differences in the fraction of infiltrated lymphocytes (Fig. 6, C and D). In contrast, we detected a lower fraction of CD68-positive macrophages and myeloperoxidase (MPO)–positive neutrophils within IL-1α^−/− and IL-1β^−/− islets when compared to WT controls (Fig. 6, C and D). To confirm that this was due to the loss of IL-1α and IL-1β and, thus, an impaired cytokine-mediated immune cell attraction during the hypoxic phase of islet transplantation, we further analyzed the fraction of lymphocytes, macrophages, and neutrophils within the pancreas as well as in isolated normoxic islets of WT, IL-1α^−/−, and IL-1β^−/− mice. As expected, we did not detect any differences between the groups (fig. S16, A to D).

Fig. 6. — (A) Representative immunofluorescence stainings of insulin and CD31 in WT, IL-1α^−/−, and IL-1β^−/− islets on day 14 after transplantation. Cell nuclei were stained with Hoechst 33342 (blue). (B) Quantitative analysis of insulin- (β cells), glucagon- (α cells), somatostatin- (δ cells), and CD31- (endothelial) positive cells in WT, IL-1α^−/−, and IL-1β^−/− islets in % of all islet cells (n = 20 each). Means ± SEM. (C) Representative immunohistochemical stainings of CD3-, CD68-, and MPO-positive cells (marked by arrowheads) in WT, IL-1α^−/−, and IL-1β^−/− islets on day 14 after transplantation. The border of the grafts is marked by broken lines. (D) Quantitative analysis of CD3- (B), CD68- (C), and MPO-positive cells (D) in WT, IL-1α^−/−, and IL-1β^−/− islets (n = 10 each). Means ± SEM. (E) Schematic illustration of pseudoislets + MVFs generation. Dispersed islet cells from WT, IL-1α^−/−, and IL-1β^−/− islets were fused with MVFs from WT mice by means of the liquid overlay technique and cultured for 5 days. On day 0, pseudoislets + MVFs were embedded into a collagen matrix and the sprouting activity was assessed on day 3. Representative image of pseudoislets + MVFs on day 3. (F) Quantitative analysis of sprouting areas of the indicated pseudoislets + MVFs. Data are expressed in % of initial size (day 0) (n = 12). Means ± SEM.

It is well-known that IL-1 signaling promotes angiogenesis (41, 42). Therefore, it was an unexpected finding that IL-1α^−/− and IL-1β^−/− islets exhibit a markedly improved revascularization after transplantation. We assumed that this is due to the proangiogenic effect of insulin in these islets. To confirm this, we performed a modified spheroid sprouting assay by the fusion of islet cells and small blood vessel segments, also known as microvascular fragments (MVFs), to pseudoislets (Fig. 6E) (43). In line with our hypothesis, we found that pseudoislets consisting of IL-1α^−/− and IL-1β^−/− islet cells and MVFs from WT mice exhibit a higher angiogenic activity when compared to pseudoislets consisting of WT islet cells and MVFs (Fig. 6F). This proangiogenic effect was abolished by the exposure of pseudoislets consisting of IL-1α^−/− and IL-1β^−/− islets to the insulin receptor (IR) antagonist linsitinib (Fig. 6F).

Loss of IL-1α and IL-1β in transplanted islets accelerates the restoration of normoglycemia in diabetic mice

Last, we transplanted WT, IL-1α^−/−, and IL-1β^−/− islets as well as WT, IL-1α^−/−, and IL-1β^−/− islets exposed to surfen under the kidney capsule of streptozotocin (STZ)–induced diabetic mice and determined their blood glucose levels and body weights over 28 days (Fig. 7A). Nondiabetic animals served as negative controls. The body weights of the mice did not differ between the groups throughout the experiments (fig. S17). However, we measured significantly lower blood glucose levels of mice transplanted with IL-1α^−/− and IL-1β^−/− islets after transplantation (Fig. 7B and fig. S18A). By performing an IPGTT on day 28 after islet transplantation, we measured lower blood glucose levels and a higher insulin content in mice transplanted with IL-1α^−/− and IL-1β^−/− islets (Fig. 7, C and D, and fig. S18B).

Fig. 7. — (A) Schematic illustration of the experimental setting: A diabetic phenotype was induced by a single injection of STZ 8 days before islet transplantation. On day 0, 300 islets were transplanted under the left kidney capsule of diabetic mice. Blood glucose levels and body weights were measured at the indicated time points. On day 28, IPGTT were performed and grafts were explanted for the determination of total insulin content. (B) Blood glucose levels (milligrams per milliliter) of diabetic mice transplanted with WT, IL-1α^−/−, and IL-1β^−/− islets and with WT, IL-1α^−/−, and IL-1β^−/− islets exposed to surfen at the indicated time points. Nondiabetic animals served as negative control (n = 8 each). Means ± SEM. (C) Quantitative analysis of blood glucose levels (milligrams per deciliter) according to the IPGTT of diabetic mice transplanted with WT, IL-1α^−/−, and IL-1β^−/− islets and with WT, IL-1α^−/−, and IL-1β^−/− islets exposed to surfen. Nondiabetic animals served as negative control (n = 8 each). Means ± SEM. (D) Insulin content (microunits per milliliter) of the removed grafts from diabetic mice transplanted with WT, IL-1α^−/−, and IL-1β^−/− islets and with WT, IL-1α^−/−, and IL-1β^−/− islets exposed to surfen (n = 4 each). Means ± SEM. (E) The fraction of mice of the different groups that achieved normoglycemia after transplantation (n = 8 each).

On the basis of our previous results showing a reduced surfen-dependent insulin secretion of isolated islets and revascularization of transplanted islets, we expected a deteriorated restoration of normoglycemia in diabetic mice receiving surfen-exposed islets when compared to diabetic mice receiving islets not exposed to surfen. However, this was not the case. We even detected a tendency toward lower blood glucose levels in mice transplanted with surfen-exposed islets (Fig. 7, B and C). This may be explained by the fact that chronic hyperglycemia, as found in patients with T1DM and STZ-treated mice, is characterized by high systemic levels of inflammatory cytokines (44–46), which, in turn, represses GSIS (47, 48). Moreover, surfen has been reported to reduce the expression of these inflammatory cytokines (49). Hence, surfen may improve the restoration of normoglycemia in diabetic mice by counteracting the cytokine-mediated lower GSIS. In line with this view, we could additionally demonstrate that the treatment of hypoxic- and cytokine-exposed islets with surfen significantly increases GSIS (fig. S19).

Together, we could demonstrate that the transplantation of IL-1–deficient islets accelerates the restoration of normoglycemia when compared to WT islets (Fig. 7E). The accelerated restoration of normoglycemia by surfen in this experimental setting is most probably caused by the abovementioned off-target effects of this heparan sulfate antagonist.

DISCUSSION

Previous studies have shown that IL-1α and IL-1β promote islet cell death (17, 34, 35, 50). Accordingly, the down-regulation of their expression as well as the inhibition of IL-1–driven signaling in islets represents a promising approach to improve the success of clinical islet transplantation. In this context, it should be considered that the mode of action of IL-1α and IL-1β signaling is highly complex, because both cytokines are not only processed in different ways but also exert distinct regulatory effects (21–24, 26). In the present study, we found that islet-resident macrophages release IL-1α and IL-1β under hypoxic conditions. In this context, we observed a reduced insulin gene expression, which may be driven by a diminished NF-κB activity and decreased PDX-1 and MafA expression. We additionally discovered the binding of IL-1α to heparan sulfate as a previously unknown mechanism potentiating IL-1α–mediated signaling transduction in mouse and human islet cells. Last, we could demonstrate that the loss of IL-1 signaling in transplanted islets accelerates their revascularization, reduces their immune cell infiltration, and improves their endocrine function.

Multiple islet stress events induce cytokine production during islet transplantation (51). Particularly oxygen and glucose deprivation combined with physical stress during the initial posttransplant phase promote the expression and release of proinflammatory cytokines, which, in turn, results in massive islet cell death (5, 43, 52). IL-1 family cytokines, including IL-1α and IL-1β, are expressed by various cells as cytosolic proforms that require cleavage for their activation and cellular release (53). It has already been reported that IL-1β promotes apoptosis in β cell lines (54, 55). Hence, it is conceivable that the loss or the exposure of hypoxic islets to IL-1α and IL-1β may protect or trigger cell death, respectively. However, we did not detect any effect of the two cytokines on islet cell viability. This indicates that the herein observed effects on islet endocrine function are not due to IL-1–induced cell death.

The effect of IL-1β on insulin secretion is still a matter of debate (50, 56, 57). We have recently shown that the transplantation of Nlrp3^−/− islets restores normoglycemia in diabetic animals (11). Of note, the NLRP3 inflammasome is crucially involved in the processing of IL-1β (58), indicating that reduced levels of IL-1β may promote insulin secretion. We herein found that the loss of IL-1α or IL-1β increases insulin secretion, whereas exposure to the two cytokines reduces insulin secretion from hypoxic islets. To elucidate the underlying mechanisms, we investigated the NF-κB pathway, which has been shown to be involved in inflammatory-driven insulin expression (50, 59). On the basis of our analyses, it may be assumed that the expression of insulin is inhibited by IL-1R–induced NF-κB activation, which down-regulates the two major transcription factors of insulin, PDX-1 and MafA, in pancreatic islets.

During isolation and culture, islets release high levels of cytokines, including IL-1β (60–62), that are suggested to be produced from resident immune cells (10, 17). In contrast, it is still unclear, which islet cell type is the source of IL-1α (17, 35). Our results show that only resident macrophages within hypoxic islets express IL-1α and IL-1β. Of note, we found that these immune cells release markedly less IL-1α when compared to IL-1β. Therefore, we assumed that the activity of IL-1α is potentiated by an extracellular mechanism. IL-1α has originally been identified as a membrane-bound cytokine (63), which was later on confirmed by many studies (30, 64–66). So far, it has been speculated that IL-1α is glycosylated and anchored onto the membrane by a lectin-like interaction. In the present study, we now identified heparan sulfate as a crucial binding partner for IL-1α on the surface of pancreatic islets, facilitating the binding of this cytokine to IL-1R.

Heparan sulfate is a structurally highly variable polysaccharide found on all cell surfaces and in extracellular matrices (67, 68). Its negatively charged chains are conjugated to defined core proteins forming proteoglycans, which interact mainly ionically with the positively charged lysine and arginine residues of proteins. The specific arrangement of more or less sulfated domains within the heparan sulfate chains regulates the activity of many binding proteins, such as growth factors, chemokines, and cytokines. For example, heparan sulfate can act either as presenting protein at defined locations or as scaffold for protein-protein interactions (69). Our in vitro results showed that mainly IL-1α and not IL-1β binds to HEP, which is a higher sulfated form of heparan sulfate. The binding of IL-1 to IL-1RI is not sufficient for the activation of intracellular pathways. It is known that a second chain, the accessory protein IL-1RAcP, is required for IL-1–dependent signaling transduction (21). The structure of IL-1RAcP is very similar to IL-1RI, but it interacts with the IL-1/IL-1RI complex without binding to IL-1 (22, 23). Our results showed that heparan sulfate only improves the binding of IL-1α to IL-1RI. Heparan sulfate acts as a kind of “molecular glue” potentiating the inhibitory action of IL-1α on insulin expression. In this context, Koehler et al. (70) demonstrated that sulfated hyaluronan binds to vascular endothelial growth factor (VEGF) and, thus, affects angiogenesis. In addition, HEP is capable of promoting fibroblast growth factor (FGF) signaling transduction via binding to FGF within the FGF receptor recognition site (71, 72).

IL-1 signaling promotes angiogenesis by up-regulating the expression of angiogenic factors, such as VEGF (41, 42). Accordingly, transplanted IL-1α^−/− and IL-1β^−/− islets should exhibit an attenuated revascularization when compared to WT controls. However, we detected an improved revascularization of these islets, which may be explained by their improved endocrine function, as indicated by a higher expression and secretion of insulin. We and others previously reported that insulin promotes blood vessel formation via binding to endothelial IR/insulin growth factor receptor (IGFR) (43, 73–75). In line with these findings, the blockade of endothelial IR/IGFR abolished the proangiogenic effect of insulin in IL-1α^−/− and IL-1β^−/− islets in the present study.

Together, we found that the loss of IL-1α and IL-1β markedly ameliorates islet transplantation by increasing insulin expression and, thus, accelerating insulin-driven graft revascularization. During the last decades, IL-1 has been extensively studied in a wide range of medical fields, and several clinical trials have been conducted highlighting the role of different IL-1 inhibitors as potential anti-inflammatory drugs (24, 53, 76), including bermekimab (anti–IL-1α antibody) (77), canakinumab (anti–IL-1β antibody) (78), and anakinra (recombinant IL-1R1 antagonist) (79). The latter has already been reported to protect cultured human islets from IL-1–mediated islet death and, hence, may be a promising approach to improve islet survival and function (80). Anakinra enhances the outcome of islet transplantation in preclinical and clinical studies when used in combination with etanercept, a tumor necrosis factor blocker, when compared to regimens without anti-inflammatory drugs (9, 81–84). Accordingly, our results not only strengthen these findings showing that blockade of IL-1 signaling is highly beneficial to improve the outcome of islet transplantation but also reveal the importance of heparan sulfate in fine-tuning IL-1 signaling.

MATERIALS AND METHODS

Material

RPMI 1640 medium and recombinant IL-1β (PMC0814) were purchased from Thermo Fisher Scientific (Karlsruhe, Germany). IL-1α, IL-1RAcP, and IL-1RI were purchased from Bio-Techne (Minneapolis, USA). Fluorescein isothiocyanate (FITC)–labeled dextran 150,000, glycerine gelatin, Hoechst 33342, neutral red solution, penicillin, rhodamine 6G, STZ, IL-1RA, HA, HEP, surfen, and Tween 20 were purchased from Sigma-Aldrich (Taufkirchen, Germany). Bovine serum albumin (BSA) was purchased from Santa Cruz Biotechnology (Heidelberg, Germany). Cell lysis reagent QIAzol was purchased from QIAGEN (Hilden, Germany). The qScriber cDNA Synthesis Kit and ORA SEE qPCR Green ROX L Mix were purchased from HighQu (Kraichtal, Germany). Collagenase NB 4G was purchased from SERVA GmbH (Heidelberg, Germany). Collagenase NB 8 Broad Range was purchased from Nordmark Biochemicals (Uetersen, Germany). HepatoQuick and Annexin-V-FLUOS Staining Kit were purchased from Roche (Basel, Switzerland). Matrigel was purchased from Corning (Wiesbaden, Germany). Polyvinylidene difluoride (PVDF) membrane was purchased from Bio-Rad (Feldkirchen, Germany). Accutase was purchased from BioLegend (Koblenz, Germany). PI was purchased from BD Biosciences (Heidelberg, Germany). Calcein was purchased from Molecular Probes (Eugene, OR, USA). Hematoxylin was purchased from Morphisto (Offenbach am Main, Germany). Linsitinib (OSI-906) was purchased from Selleckchem (München, Germany).

Antibodies

The anti-CD31 antibody (DIA310) was purchased from Dianova (Hamburg, Germany). The anti-MafA (sc-390491) and anti-PDX-1 (sc-25403) antibodies were purchased from Santa Cruz Biotechnology (Heidelberg, Germany). The anti-insulin (ab181547), anti-somatostatin (ab30788), anti-glucagon (ab92587), anti-MPO (ab9535), anti-CD3 (ab16669), and anti-CD68 (ab125212) antibodies were purchased from Abcam (Cambridge, UK). The anti-p65 (80979) and anti–β-actin (HRP-66009) antibodies were purchased from Proteintech (Rosemont, USA). The anti–p-p65 antibody (3033) was purchased from Cell Signaling Technology (Leiden, The Netherlands). The anti–IL-1β (17-7114-80), anti-rabbit immunoglobulin G (IgG) Alexa Fluor 555 (A-21429), anti-mouse IgG Alexa Fluor 488 (A-11001), anti-rat IgG Alexa Fluor 488 (A-21434), and anti-guinea pig IgG Alexa Fluor 488 (A-11073) antibodies were purchased from Thermo Fisher Scientific (Karlsruhe, Germany). The anti–IL-α and CD68 (137008) antibodies were purchased from BD Biosciences. The anti–heparan sulfate antibody (370255) was purchased from Amsbio (Alkmaar, The Netherlands). The peroxidase-labeled anti-rabbit (NIF 824) and peroxidase-labeled anti-mouse (NIF 825) antibodies were purchased from GE Healthcare (Freiburg, Germany).

Cell culture

U937 cells were cultivated in RPMI 1640 [10% (v/v) fetal calf serum (FCS), penicillin (100 U/ml), and streptomycin (0.1 mg/ml)] at 37°C under a humidified 95 to 5% (v/v) mixture of air and CO₂. Cells were passaged at a split ratio of 1:3. For differentiation, U937 were stimulated with phorbol 12-myristate 13-acetate (25 ng/ml) for 3 hours and incubation for 21 hours with lipopolysaccharide (100 ng/ml) and interferon-γ (20 ng/ml). The M1 status of the macrophages was assessed by flow cytometric detection of CD86, CD68, and CD11c.

U937-derived macrophages, isolated islets or isolated islets exposed to recombinant IL-1α (200 pg/ml), IL-1β (200 pg/ml), HPSE (5 mU/ml), surfen (20 μM), or IL-1RA (1 μg/ml) were cultivated in RPMI 1640 [penicillin (100 U/ml) and streptomycin (0.1 mg/ml)] under hypoxic conditions (95% N₂, 5% CO₂, and 1% O₂) for 16 hours. Afterward, the cells and islets were harvested for further experiments.

Western blot analysis

Whole-cell extracts from hypoxic WT islets exposed to IL-1α or IL-1β were generated, separated through a 12.5% SDS–polyacrylamide gel electrophoresis and transferred onto a PVDF membrane. The membrane was incubated in 5% dry milk in phosphate-buffered saline (PBS) (0.1% Tween 20) for 1 hour and exposed to the indicated primary antibodies, which were diluted (1:500) in PBS (0.1% Tween 20) containing 1% dry milk. After incubation of the membrane with a peroxidase-coupled secondary antibody [anti-rabbit (1:2000) or anti-mouse (1:2000)] for 1 hour, the protein expression was visualized by the incubation of the membrane with enhanced chemoluminescence Western blotting substrate (GE Healthcare) in a Chemocam device (Intas, Göttingen, Germany). The intensity of the measured signals was quantified using ImageJ software and normalized by the corresponding housekeeping protein.

Quantitative real time-polymerase chain reaction

Total RNA from hypoxic islets exposed to IL-1α or IL-1β was isolated using QIAzol lysis reagent (QIAGEN). The corresponding cDNA was synthesized from 1 μg of total RNA by QuantiNova Reverse Transcription Kit (QIAGEN) according to the manufacturer’s instructions. ORA qPCR Green ROX L Mix (highQu) was used for quantitative real time-polymerase chain reaction (qRT-PCR). Data analysis was performed by the MiniOpticon Real-Time PCR System (Bio-Rad). Murine glyceraldehyde-3-phosphate dehydrogenase (GAPDH) served as internal control for mRNA detection.

Forward and reverse primers were used in a concentration of 700 nM solved in ribonuclease/deoxyribonuclease-free H₂O. Primer sequences for qPCR were coded as follows: mouse Ins-1, (forward) 5′-AACAACTGGAGCTGGGAGGAAG-3′ and (reverse) 5′-GGTGCAGCACTGATCCACAATG-3′; mouse Ins-2, (forward) 5′-GCAGCACCTTTGTGGTTCC-3′ and (reverse) 5′-CTTGTGGGTCCTCCACTTC-3′; mouse PDX-1, (forward) 5′- GAAATCCACCAAAGCTCACG-3′ and (reverse) 5′- GAATTCCTTCTCCAGCTCCAG-3′; mouse MafA, (forward) 5′- GGTCATCCGACTGAAACAGA-3′ and (reverse) 5′- CACTTCTCGCTCTCCAGAAT-3′; and mouse GAPDH, (forward) 5′-CGGTGCTGAGTATGTC-3′ and (reverse) 5′-TTTGGCTCCACCCTTC-3′.

Calcein/PI staining

Isolated islets and isolated islets cultivated for 24 hours in 0.2% H₂O₂ (positive control) were incubated for 20 min at 37°C with calcein (1 μg/ml) and PI (5 μg/ml). Cell nuclei were stained for 10 min at 37°C with Hoechst 33342 (2 μg/ml). The islets were washed with PBS, resuspended in glycerin gelatin, and covered on slides. The cellular stainings were visualized by using fluorescence microscopy [BX60F fluorescence microscope (Olympus)] 15 min after mounting.

Neutral red/trypan blue staining

Isolated islets and isolated islets incubated for 24 hours in 0.2% H₂O₂ (positive control) were incubated for 2 min at room temperature with neutral red (1:100) or trypan blue (1:100) and washed with PBS. The cellular stainings were visualized by bright field images using a 20× objective of a BX60F microscope (Olympus).

Flow cytometry

Normoxic and hypoxic isolated islets were dispersed into single cells by Accutase (1:20), or U937-derived macrophages were detached from culture plates by scratching. The cells were washed and incubated with the indicated phycoerythrin (PE)–labeled primary antibodies as well as the corresponding PE-labeled control antibodies for 30 min at room temperature. The cells were washed in PBS, and the number of positive cells and the MFI for the indicated antibodies were determined by a FACSLyric flow cytometry system (BD Biosciences).

For PI/annexin V staining, isolated islets were dispersed into single cells by Accutase (1:20). Subsequently, the cells were washed in PBS, resuspended in incubation buffer, and stained for 15 min with PI and annexin V (100 μg/ml), according to the manufacturer’s protocol (Roche). The stained cells were analyzed by flow cytometry using a FACSLyric flow cytometry system (BD Biosciences), and the fractions of vital, apoptotic, necrotic, and necroptotic cells were given in % of all measured cells.

Enzyme-linked immunosorbent assay

The amount of secreted insulin was measured by an insulin or C-peptide ELISA kit. For this purpose, 10 isolated mouse or human islets were washed with Krebs Ringer Buffer (KRB) (115 mM NaCl, 4.7 mM KCl, 1.28 mM CaCl₂, 1.2 mM MgSO₄, and 0.1% BSA) and incubated for 1 hour at 37°C and 5% CO₂. The supernatants were discarded, and the islets were incubated for 30 min in KRB containing 16.5 mM glucose. The supernatants were collected and the amount of secreted insulin or C-peptide was determined by using an insulin or C-peptide ELISA kit (Invitrogen, USA) according to the manufacturer’s protocol.

The amount of released IL-1α or IL-1β was measured by an IL-1α or IL-1β ELISA according to the manufacturer’s protocol (Invitrogen, USA). For this purpose, 100 isolated islets or 1 × 10⁵ U937-derived macrophages (per well of a 24-well plate) were cultivated for 16 hours under hypoxia or normoxia. The supernatants were collected and the amount of released IL-1α or IL-1β was determined.

The binding of IL-1α to GAG surfaces was analyzed by ELISA. HA and HEP solutions in 25 mM citrate-phosphate buffer were immobilized onto high-binding ELISA plates, as previously described (85). Nonspecific binding was evaluated using BSA-coated wells. GAG surfaces were incubated with IL-1α dissolved in 1% BSA/PBS at 4°C for 16 hours. Afterward, the supernatants containing the nonbound IL-1α were analyzed using a sandwich ELISA.

Competitive IL-1α ELISAs were performed with the Duo-Set IL-1α ELISA kit (Invitrogen, USA). High-binding ELISA plates were coated with IL-1α capture antibody overnight. Afterward, IL-1α (80 ng/ml) was added together with increasing concentrations of IL-1RI or IL-1RAcP in the absence or presence of 500 μM HEP related to its disaccharide units. After 60 min of incubation at room temperature followed by extensive washing, the IL-1α detection antibody was added and IL-1α was quantified.

SPR analysis

Interactions of HA and HEP with human IL-1α and human IL-1β were analyzed using a Biacore T200 instrument (Cytiva). IL-1α or IL-1β was immobilized onto a Series S Sensor Chips CM5 (Cytiva) via amine coupling at 25°C according to Cytiva. After surface activation, IL-1α/β (8 μg/ml) in sodium acetate buffer (pH 4.5) was injected at a flow rate of 10 μl/min until an immobilization level of ~800 RU was achieved. Unreacted groups were saturated by injecting 1 M ethanolamine-HCl, pH 8.5 (7 min, 10 μl/min). A nonprotein-immobilized surface served as control.

Binding analysis was performed at 37°C with a flow rate of 30 μl/min. Each analyte was diluted in Hepes-buffered saline–EDTA and polysorbate (EP). GAGs were injected at a concentration of 500 μM related to their disaccharide units. Binding levels were detected 10 s before the end of sample injection. After a 1000-s dissociation phase, the sensor chip surface was regenerated for 60 s with 5 M NaCl followed by a 1000-s stabilization time. Binding parameters were evaluated with Biacore T200 evaluation software.

Electrostatic potential

The Adaptive Poisson-Boltzmann Solver (86) plugin in PyMOL (The PyMOL Molecular Graphics System, version 2.4.1 Schrödinger LLC) was used to calculate the electrostatic potential of IL-1α [Protein Data Bank (PDB) ID 2KKI] (87).

Molecular docking

The lowest energy structure of the NMR ensemble of IL-1α obtained from the PDB (PDB ID 2KKI) (87) was used for blind docking calculations to HEP with Autodock 3. The structure of HEP was modeled as tetramer and hexamer [(i.e., tetrasaccharide (dp4) and hexasaccharide (dp6), respectively], as previously described (88).

HEP was treated completely flexible, whereas IL-1α was considered rigid. Autogrid3 was used to compute the atomic potential of IL-1α covering the full protein surface with a grid box and spacing grid of 126 Å by 126 Å by 126 Å and 0.4 Å, respectively. Calculations and clustering analysis were performed as previously described (88). Three representative HEP/IL-1α complexes for each of the obtained docking clusters were selected for further refinement.

MD simulations

The selected representative complexes were further refined by MD simulations in AMBER 2019 (89). Charges and parameters for sulfate groups, HEP, and protein were taken from the literature (90), GLYCAM-06j (91), and ff14SB (89) force fields, respectively. Each HEP/protein complex was solvated in a truncated octahedral box of TIP3P water molecules and neutralized with Na⁺ counterions. Three independent 100-ns MD simulations were carried out following the same protocol as previously described (88). MD trajectories were recorded every 10 ps. HEP pyranose rings were harmonically restrained. Trajectories were visualized with Visual Molecular Dynamics (VMD) (92). Energy decomposition per residue as well as binding free energy post-processing analysis of 200 frames distributed along the last 50 ns of the MD production runs were performed in implicit solvent with MM-GBSA (93, 94). Data analysis was carried out with Origin2019b (Origin, version 2019b, OriginLab Corporation, Northampton, MA, USA). Figures were created with PyMOL (The PyMOL Molecular Graphics System).

Human islets

Islets were isolated from donor organs obtained with informed written consent and research ethics approval at the University of Alberta and Saarland University. The characteristics of human pancreas donors and preparations are summarized in table S1. Experiments conformed to the principles set out in the WMA Declaration of Helsinki and the Department of Health and Human Services Belmont Report. Human islets were cultured in low-glucose (5.5 mM) Dulbecco’s modified Eagle’s medium with l-glutamine, sodium pyruvate (110 mg/liter), 10% FCS, and penicillin (100 U/ml) and streptomycin (0.1 mg/ml).

Human islets for research were provided by the Alberta Diabetes Institute IsletCore at the University of Alberta in Edmonton (www.bcell.org/adi-isletcore.html) with the assistance of the Human Organ Procurement and Exchange program, Trillium Gift of Life Network, and other Canadian organ procurement organizations. Islet isolation was approved by the Human Research Ethics Board at the University of Alberta (Pro00013094). All donors’ families gave informed consent for the use of pancreatic tissue in research.

Animals

Animals were maintained on a standard 12/12-hour day/night cycle. Water and standard pellet chow (Altromin, Lage, Germany) were provided ad libitum. C57BL/6J WT as well as IL-1a^−/− (Il1a^tm1Yiw) and IL-1b^−/− (Il1b^tm1Yiw) mice [provided by T. Speer (30) and originally described by Horai et al. (95)] with a body weight of 25 to 30 g (age of 3 to 12 months) served as donors for islet and pancreas isolation. Male C57BL/6J WT mice with a body weight of 30 to 35 g (age of 3 to 6 months) served as donors for MVFs isolation. C57BL/6J WT mice with a body weight of 22 to 27 g (age of 2 to 3 months) were used for the dorsal skinfold chamber model. Diabetes was induced in male C57BL/6J WT mice with a body weight of 24 to 28 g (age of 2 to 3 months).

All animal experiments were performed in compliance with the National Institutes of Health (NIH) Guidelines on the Care and Use of Laboratory Animals (NIH publication no. 85-23 Rev. 1985) and the European legislation on the protection of animals (Directive 2010/63/EU). They were approved by the local authorities (permission numbers 45/2018, 06/2020, and 08/2024; State Office for Consumer Protection, Saarbrücken, Germany).

Isolation of pancreatic islets

Mice were anesthetized by intraperitoneal injection of ketamine (100 mg/kg body weight) and xylazine (12 mg/kg body weight). Following cervical dislocation and midline laparotomy, the pancreatic duct was injected with collagenase NB 8 (1 mg/ml) containing neutral red solution (25 μl/ml), and pancreatic islets were isolated as described previously in detail (96). Isolated islets were cultivated in RPMI 1640 [supplemented with 10% (v/v) FCS, penicillin (100 U/ml), and streptomycin (0.1 mg/ml)] for 24 hours at 37°C and 5% CO₂ for further experiments.

GAG quantification

For quantification of sulfated GAGs, tissues from C57BL/6J WT, IL-1a^−/−, and IL-1b^−/− mice were washed twice in Dulbecco’s balanced salt solution (DPBS) to remove residual blood. The tissues were lyophilized until dry, powderized with a spatula, and resuspended in DPBS with 10 mM CaCl₂, 0.1% (v/v) Triton X-100, and pronase (2 mg/ml). The tissues were digested at 37°C overnight under continuous shaking. The tissue homogenates were centrifuged at 20,000g for 20 min. The supernatant was used for quantification of sulfated GAGs by dimethylmethylene blue (DMMB) assay. For the DMMB assay, a dilution series of chondroitin sulfate C was used for standard curve. The DMMB assay was performed in 96 well plates with 20 μl of standard or sample. After addition of 200 μl of DMMB reagent, the absorbance at 525 nm was measured with a plate reader.

Generation of pseudoislets and sprouting assay

Mice were anesthetized by an intraperitoneal injection of ketamine (100 mg/kg body weight) and xylazine (12 mg/kg body weight) and euthanized by cervical dislocation. Subsequently, MVFs were isolated by mechanic and enzymatic digestion (collagenase NB 4G) of epididymal fat pads of mice, as described previously in detail (97). After isolation, MVFs were fused with dispersed islet cells to pseudoislets by means of the liquid overlay technique in a 96-well plate covered with 1% agarose (43). After 5 days, the pseudoislets were harvested, and their angiogenic activity was determined by a sprouting assay, as previously described in detail (43). The sprouted pseudoislets were visualized by a BX60F microscope (Olympus). The sprouting area was assessed by means of the Fiji software (NIH). Data are given in % of the initial pseudoislet area.

Preparation of the dorsal skinfold chamber and islet transplantation

Mice were anesthetized by an intraperitoneal injection of ketamine (100 mg/kg body weight) and xylazine (12 mg/kg body weight), and the dorsal skinfold chamber was implanted, as described previously in detail (98). Briefly, two symmetrical titanium frames were prepared on the extended dorsal skinfold of anesthetized mice, resulting in the doubling of the skin in two layers. One layer, including skin, subcutis, and the retractor muscle, was completely removed in a circular area of 15 mm in diameter. This area was then covered by a removable cover slip and a snap ring providing direct microscopic access to the microcirculation of the chamber. After the procedure, the animals were allowed to recover for 48 hours.

After recovery, the mice were again anesthetized by an intraperitoneal injection of ketamine (100 mg/kg body weight) and xylazine (12 mg/kg body weight), the cover glass was removed, and the tissue was washed with saline. Subsequently, eight isolated islets (cultivated for 24 hours prior their transplantation) were transplanted onto the exposed striated muscle tissue. Last, the chamber was sealed with a new cover slip for repeated intravital fluorescence microscopic analyses.

Intravital fluorescence microscopy

Anesthetized dorsal skinfold chamber–equipped mice received a retrobulbary intravenous injection of 0.05 ml of FITC-labeled dextran (5%) for plasma staining and 0.05 ml of rhodamine 6G (2%) for the visualization of microvessel fenestration (99) on day 0 as well as days 3, 6, 10, and 14 after islet transplantation. Thereafter, the dorsal skinfold chamber was positioned under a fluorescence microscope (Zeiss), and the microscopic images were recorded for offline evaluation by the computer-assisted image analysis system CapImage (Zeintl, Heidelberg, Germany). The revascularized area (square millimeters), the functional microvessel density (centimeters per square centimeter), and the rhodamine 6G–positive area (calculated by the ratio of the rhodamine 6G–positive area and the revascularized area on that day in %) of islets were assessed as previously described (100, 101). In addition, we measured the diameter (micrometers), centerline RBC velocity (micrometers per second), and volumetric blood flow (picoliters per second) of four to eight individual microvessels within the grafts (100, 101). Moreover, the take rate (%), i.e., the number of engrafted islets on day 14 in relation to the number of transplanted islets on day 0, was determined.

Diabetes induction and islet transplantation under the kidney capsule

Diabetic phenotypes were induced by a single intraperitoneal injection of STZ (180 mg/kg) 8 days before islet transplantation. Body weights and nonfasting blood glucose levels of STZ-injected mice were measured twice a week during the entire observation period of 28 days. Blood samples were taken from the tail vein and analyzed by a portable blood glucose monitoring system (GL50; Breuer). Mice with a nonfasting blood glucose level ≥ 350 mg/dl served as recipients for islet transplantation. Three hundred isolated islets were injected under the left kidney capsule of diabetic mice using a 10-μl Hamilton syringe. Normoglycemia was defined by blood glucose levels below 200 mg/dl.

IPGTT and insulin content

The IPGTT was performed on day 28 after islet transplantation under the kidney capsule of diabetic mice. After 16 hours of fasting, the mice were intraperitoneal injected with a 10% glucose solution. The blood glucose levels were determined 0, 15, 30, 45, 60, 120, and 180 min after glucose injection from the tail vein and analyzed by a portable blood glucose monitoring system (GL50; Breuer).

To determine the total insulin content of the grafts, the islet transplants underneath the kidney capsule were dissected and lysed in 1 ml radioimmunoprecipitation assay lysis buffer, and the intracellular insulin content was determined by an insulin ELISA kit according to the manufacturer’s protocol.

Plasma insulin levels were determined from fasted mice. After 7 hours of fasting, the mice were intraperitoneal injected with a 10% glucose solution, and the blood samples were collected from the tail vein. The blood plasma was separated by centrifugation and stored at −80°C. The plasma insulin levels were analyzed by means of an insulin ELISA kit according to the manufacturer’s protocol.

Immunohistochemistry

For the preparation of histological sections, dorsal skinfold chamber–equipped mice were anesthetized by an intraperitoneal injection of ketamine (100 mg/kg body weight) and xylazine (12 mg/kg body weight) and euthanized by cervical dislocation. The dorsal skinfold chamber tissue was excised and fixed for 24 hours in 4% paraformaldehyde (PFA). Specimens from pancreatic tissue of mice and human were excised and fixed for 24 hours in 4% PFA. In addition, isolated islets were incubated for 45 min at 37°C in 100 μl of HepatoQuick, 50 μl of human citrate plasma, and 10 μl of 10% CaCl₂ solution. The resulting clot was also fixed for 24 hours in 4% PFA. The PFA-fixed specimens were embedded in paraffin and 3-μm-thick sections were cut.

The sections were stained with the indicated primary antibodies and visualized by their corresponding secondary antibodies. Cell nuclei were stained with Hoechst 33342 for fluorescence microscopy and with hematoxylin for bright field microscopy. The sections were analyzed by means of fluorescence microscopy [BX60F fluorescence microscope (Olympus)]. The quantification of positively stained cells was done by Fiji software (NIH) and is given in % of all islet cells.

Statistical analysis

All in vitro experiments were reproduced at least three times. The in vivo experiments were performed with eight animals per group. After testing the data for normal distribution and equal variance, differences between the groups were assessed by the one-way analysis of variance (ANOVA) following by the Tukey post hoc test. Differences between the groups over time were assessed by a mixed-effects model or repeated measures ANOVA followed by the Tukey post hoc test. The statistical analysis was performed by means of Prism software 10.2.3 (GraphPad, USA). The results were expressed as means ± SEM. P values of < 0.05 indicated statistical significance.

Acknowledgments

We thank Servier Medical Art for providing access to designed medical elements (https://smart.servier.com/), supporting the generation of graphical items in this publication. We are grateful for the excellent assistance of C. Bickelmann and R. M. Nickels (Institute for Clinical and Experimental Surgery). We thank T. Speer for providing the IL-1a^−/− and IL-1b^−/− mice.

Funding: This work is supported by the Deutsche Forschungsgemeinschaft (DFG) (AM 640/2-1).

Author contributions: Conceptualization: L.P.R., S.W., M.D.M., E.A., P.E.M., M.W.L., S.R., and S.S. Methodology: S.W., M.D.M., G.R.-G., M.T.P., E.A., M.W.L., S.R., and S.S. Investigation: L.P.R., S.W., T.R., G.R.-G., M.T.P., E.A., F.J., S.R., F.P., A.S.B., and S.S. Visualization: S.W., G.R.-G., M.T.P., E.A., M.W.L., and S.R. Funding acquisition: M.D.M., E.A., and S.S. Project administration: M.D.M., M.T.P., E.A., S.S. Supervision: L.P.R., M.D.M., M.T.P., E.A., M.W.L., and S.S. Writing—original draft: S.W., G.R.-G., M.T.P., E.A., P.E.M., M.W.L., and S.S. Writing—review and editing: L.P.R., S.W., M.D.M., G.R.-G., M.T.P., E.A., M.W.L., S.R., and S.S. Resources: L.P.R., S.W., M.D.M., G.R.-G., M.T.P., E.A., P.E.M., M.W.L., and S.S. Data curation: S.W., G.R.-G., M.T.P., E.A., and P.E.M. Validation: S.W., G.R.-G., M.T.P., E.A., and S.S. Formal analysis: S.W., G.R.-G., M.T.P., E.A., and S.S. Software: E.A.

Competing interests: The authors declare that they have no competing interests.

Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials. The human islets can be provided by Alberta Diabetes Institute IsletCore pending scientific review and a completed material transfer agreement. Requests for the human isltes should be submitted to Alberta Diabetes Institute IsletCore at the University of Alberta in Edmonton (www.bcell.org/adi-isletcore.html).

Supplementary Materials

This PDF file includes:

Figs. S1 to S19

Table S1

sciadv.ady8566_sm.pdf^{(2.7MB, pdf)}

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figs. S1 to S19

Table S1

sciadv.ady8566_sm.pdf^{(2.7MB, pdf)}

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PERMALINK

Heparan sulfate fine-tuned interleukin-1 (IL-1) signaling inhibits insulin secretion of grafted pancreatic islets

Selina Wrublewsky

Sandra Rother

Franziska Pohlemann

Toni Radanovic

Fabian Junker

Anne S Boewe

Stefan Schunk

Leticia P Roma

Gloria Ruiz-Gómez

M Teresa Pisabarro

Patrick E MacDonald

Michael D Menger

Matthias W Laschke

Emmanuel Ampofo

Roles

Abstract

INTRODUCTION

RESULTS

Loss of IL-1α and IL-1β does not affect the viability and cellular composition of isolated islets

Fig. 1. Loss of IL-1α and IL-1β does not affect the viability and cellular composition of isolated islets.

IL-1α and IL-1β suppress the gene expression and secretion of insulin

Fig. 2. IL-1α and IL-1β suppress insulin gene expression.

IL-1α and IL-1β are released from islet-resident macrophages under hypoxic conditions

Fig. 3. IL-1α and IL-1β are expressed and released from islet resident macrophages under hypoxic conditions.

IL-1α binds to islet heparan sulfate

Fig. 4. IL-1α binds to islet heparan sulfate.

Loss of IL-1α and IL-1β improves the revascularization of transplanted islets

Fig. 5. Loss of IL-1α and IL-1β improves the revascularization of transplanted islets.

Fig. 6. Cellular composition of transplanted islets and angiogenic activity of PI + MVFs.

Loss of IL-1α and IL-1β in transplanted islets accelerates the restoration of normoglycemia in diabetic mice

Fig. 7. In vivo endocrine function of transplanted islets.

DISCUSSION

MATERIALS AND METHODS

Material

Antibodies

Cell culture

Western blot analysis

Quantitative real time-polymerase chain reaction

Calcein/PI staining

Neutral red/trypan blue staining

Flow cytometry

Enzyme-linked immunosorbent assay

SPR analysis

Electrostatic potential

Molecular docking

MD simulations

Human islets

Animals

Isolation of pancreatic islets

GAG quantification

Generation of pseudoislets and sprouting assay

Preparation of the dorsal skinfold chamber and islet transplantation

Intravital fluorescence microscopy

Diabetes induction and islet transplantation under the kidney capsule

IPGTT and insulin content

Immunohistochemistry

Statistical analysis

Acknowledgments

Supplementary Materials

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REFERENCES AND NOTES

Associated Data

Supplementary Materials

ACTIONS

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