Research ArticleISLET TRANSPLANTATION

Human umbilical cord perivascular cells improve human pancreatic islet transplant function by increasing vascularization

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Science Translational Medicine  15 Jan 2020:
Vol. 12, Issue 526, eaan5907
DOI: 10.1126/scitranslmed.aan5907

Improving islet transplantation

Pancreatic islet transplantation can stabilize blood glucose in individuals with type 1 diabetes; however, transplant function decreases occur over time, and suitable donor islet availability remains limited. Forbes et al. investigated the supportive effects of cotransplanting islets and perivascular mesenchymal stromal cells isolated from human umbilical cords (HUCPVCs). The HUCPVCs inhibited T cells and expressed proregenerative and immunoregulatory markers in vitro and increased islet vascularization in vivo. Cotransplanting human islets and HUCPVCs under the kidney capsule or via the hepatic portal vein improved control of blood glucose in diabetic immunodeficient and immunocompetent mouse models for up to 16 weeks. Results suggest that HUCPVCs could help improve long-term islet transplant function.

Abstract

Islet transplantation is an efficacious therapy for type 1 diabetes; however, islets from multiple donor pancreata are required, and a gradual attrition in transplant function is seen. Here, we manufactured human umbilical cord perivascular mesenchymal stromal cells (HUCPVCs) to Good Manufacturing Practice (GMP) standards. HUCPVCs showed a stable phenotype while undergoing rapid ex vivo expansion at passage 2 (p2) to passage 4 (p4) and produced proregenerative factors, strongly suppressing T cell responses in the resting state and in response to inflammation. Transplanting an islet equivalent (IEQ):HUCPVC ratio of 1:30 under the kidney capsule in diabetic NSG mice demonstrated the fastest return to normoglycemia by 3 days after transplant: Superior glycemic control was seen at both early (2.7 weeks) and later stages (7, 12, and 16 weeks) versus ratios of 1:0, 1:10, and 1:50, respectively. Syngeneic islet transplantation in immunocompetent mice using the clinically relevant hepatic portal route with a marginal islet mass showed that mice transplanted with an IEQ:HUCPVC ratio of 1:150 had superior glycemic control versus ratios of 1:0, 1:90, and 1:210 up to 6 weeks after transplant. Immunodeficient mice transplanted with human islets (IEQ:HUCPVC ratio of 1:150) exhibited better glycemic control for 7 weeks after transplant versus islet transplant alone, and islets transplanted via the hepatic portal vein in an allogeneic mouse model using a curative islet mass demonstrated delayed rejection of islets when cotransplanted with HUCPVCs (IEQ:HUCPVC ratio of 1:150). The immunosuppressive and proregenerative properties of HUCPVCs demonstrated long-term positive effects on graft function in vivo, indicating that they may improve long-term human islet allotransplantation outcomes.

INTRODUCTION

Islet transplantation is effective for stabilizing glycemic control and restoring awareness of hypoglycemia in people with type 1 diabetes (19), and there is evidence of diminished progression of microvascular and macrovascular complications (10). However, transplant is suboptimal, and individuals typically require two to three islet transplants to improve their glycemic control (4). Inflammatory and immunological rejection mechanisms lead to the early loss of islets (11, 12) and contribute to the attrition in graft function seen over time (4). Repeat immune sensitization events may arise due to the necessity for multiple transplants. Furthermore, the number of people that may receive transplants is limited by the availability of suitable donor pancreata (13). Therapeutic strategies to improve graft function are urgently needed. Ideally, a therapeutic strategy would modulate the inflammatory and immune response and enhance engraftment of islets through a variety of mechanisms, reducing the need for multiple transplants and high doses of immunosuppression. This would allow more patients to be treated and would enable the use of pancreata with marginal numbers of isolated islets to be used.

Mesenchymal stromal cells (MSCs) are multipotent cells found in most tissues, which are capable of self-renewal and have potential for trilineage differentiation: osteogenic, adipogenic, and chondrogenic (14). They have been shown to promote regeneration of tissues by supporting blood vessel formation through production of a broad spectrum of growth factors and extracellular matrix building molecules (15). MSCs suppress the host immune response, both innate and adaptive pathways, by activating anti-inflammatory factors, suppressing B and T cell proliferation, and increasing production of regulatory T cells and polarized M2 macrophages. Such functions are highly relevant to the therapy of the chronic inflammatory autoimmune disease type 1 diabetes (16).

Bone marrow–derived MSCs (bmMSCs) have been the gold standard in clinical practice, enhancing donor bone marrow cell engraftment and chimerism (17), and have been cotransplanted with islets in animal models of diabetes with favorable effects on graft function and longevity (18). However, bmMSCs are present at low frequency in the marrow compartment, cell harvesting is painful and invasive, and bmMSCs also exhibit a reduced ex vivo expansion capacity as donor age increases (19). MSCs from other sites, including umbilical cord, have been used in animal models of diabetes, although not in animal models of islet transplantation (20). Cellular preparations derived from umbilical cord can be obtained in clinically relevant quantities with procurement advantages: Cell harvesting is noninvasive, and cells are highly expandable ex vivo. Preparations from umbilical cord, which include MSC-type cells and other cell types (epithelial), can undergo freeze-thaw cycles for off-the-shelf use (21) and have been used in clinical trials (20, 21). However, before such MSCs can be used in clinical islet allotransplantation, they must be fully characterized, demonstrate greater efficacy on cotransplantation with islets versus islet transplantation alone, and must be compliant with GMP.

There have been no studies in islet transplantation in man using allogeneic MSCs. MSC studies in type 1 diabetes are heterogeneous and difficult to compare because the numbers and types of MSCs used have differed, the time interval between diagnosis of diabetes and infusion of stem cells is highly variable, and some studies use autologous stem cells, whereas other studies use allogeneic stem cells (22, 23). No studies in man have developed and/or characterized GMP-compliant umbilical cord MSCs for cotransplant with islets or have systematically examined the islet:MSC ratio, which has the greatest impact on short- and long-term glycemic control.

Here, we have adapted existing methods for production of well-characterized human umbilical cord perivascular mesenchymal stromal cell (HUCPVC) (2427) to a fully GMP-compliant, xeno-free culture system. Our aims were to repeatedly derive and characterize GMP-grade HUCPVCs with respect to immunoregulatory and regenerative properties in vitro as well as their metabolic efficacy in cotransplantation experiments with islets in vivo [including infusion via the hepatic portal vein (HPV) route used in human transplantation] and to quantify islet graft vascularization using rodent models of islet transplantation.

RESULTS

Generation of GMP-grade HUCPVCs with stable MSC phenotype

Six donations of umbilical cord were processed into HUCPVC lines, frozen after the initial seed culture was established, and then thawed and expanded up to p4 using three GMP-compliant media: StemMACS (SM), SM + group AB donor plasma (SMAB), and SM + platelet lysate (SMPL). HUCPVCs exhibited the characteristic MSC phenotype [International Society for Cellular Therapy criteria (28)]: expressing the markers CD73, CD90, and CD105 but negative for expression of CD34, CD45, CD14, and CD11b (Fig. 1A). HUCPVCs also consistently expressed CD10, which is definitively associated with perivascular cells of the Wharton’s jelly (29, 30). SMPL gave the shortest population doubling times; however, differences were not significant, and all three media gave satisfactory results (Fig. 1B). The HUCPVC phenotype was stable throughout serial passage of the cells, and HUCPVCs maintained similar stemness potential in all media, assessed by colony-forming unit–fibroblast (CFU-F) assay. Lipoaspirate-derived MSCs, derived from abdominal subcutaneous tissue (used as a comparator material, see characterization in fig. S1), thawed, and grown in identical medium from p2 to p4, demonstrated significantly lower CFU-F activity in the same assay (Fig. 1C). HUCPVCs exhibited the ability to differentiate into adipocytes, osteocytes, and chondrocytes in vitro (Fig. 1D). One donation from an individual umbilical cord could be manufactured into a cell product containing a minimum of 1.04 × 108 cells [Dulbecco’s modified Eagle medium (DMEM) + PL (DMPL) at initiation: 1.04 × 108 to 2.97 × 108; n = 3] or minimum of 4.35 × 108 cells (SMPL at initiation: 4.25 × 108 to 24.5 × 108; n = 3) by p2. HUCPVCs constitutively expressed the T cell inhibitory surface molecules programmed death-ligand 1 (PD-L1) (CD274) and PD-L2 (CD273) (Fig. 1E). “Licensing” (exposure to inflammatory cytokines) of HUCPVCs with interferon-γ (IFN-γ) increased the relative fluorescence intensity of CD273 and CD274 (Fig. 1E).

Fig. 1 HUCPVCs grown in different GMP-compliant media.

(A) Representative phenotypic analysis of HUCPVCs grown in GMP-compliant medium (SMAB). HUCPVCs p3, baseline fluorescence set using matched isotype controls. Right: Percentage of HUCPVCs expressing a CD14CD45CD73+CD90+CD105+ phenotype across three serial passages (p2 to p4). n = 3 individual donations. (B) HUCPVCs grown in different GMP media across three passages (p2 to p4) from n = 5 donated samples of umbilical cord. Mean doubling time ± SEM (days) using different GMP-compliant media. No significant differences between media were used to expand cell lines with two-way ANOVA with Tukey’s multiple comparison test. P ranges from 0.22 to 0.99. (C) Colony-forming unit scores for HUCPVCs and lipoaspirate MSCs grown in different GMP-compliant media from three different tissue donations across three passages (p2 to p4) with numbers of colonies obtained from plating 10 cells/cm2 in CFU-F medium; quantified mean ± SEM is shown; CFU-F is significantly higher in HUCPVC than in lipoaspirate cells (P < 0.01), paired t test in all media tested. (D) Differentiation of HUCPVCs to adipocytes, osteocytes, and chondrocytes. Samples were stained with Oil Red O (Adipose), Alizarin Red (Osteo), or Safranin O (Chondro) as described (63). Bright-field microscopy magnification, ×40. (E) Analysis of PD-L2 (CD273) and PD-L1 (CD274) expression on HUCPVCs with and without licensing. Light gray peak/green outline, isotype control; gray peak/blue outline, unlicensed; white peak, licensed. SM, StemMACS; SMAB, SM + AB plasma; SMPL, SM + platelet lysate; CFU-F, colony-forming unit–fibroblasts.

GMP HUCPVCs are potent T cell inhibitors that do not require a priori licensing by IFN-γ

HUCPVCs exhibited robust inhibition of T cell proliferation. Addition of these MSCs to mitogen-stimulated peripheral blood mononuclear cells (PBMCs) in a ratio of 2:1 PBMC:HUCPVC inhibited T cell division (Fig. 2A), with an effect still measurable at 8:1 (Fig. 2A). T cell inhibition by HUCPVCs was achieved without licensing. Addition of IFN-γ as a single licensing cytokine did not significantly increase the ability of the HUCPVCs to suppress T cell proliferation in this assay (Fig. 2B). Licensing of HUCPVCs with multiple cytokines—IFN-γ, interleukin-1β (IL-1β), and tumor necrosis factor–α (TNF-α)—did not increase the ability of HUCPVCs to suppress T cell proliferation compared to unlicensed cells (fig. S2). Unlicensed HUCPVCs grown in all three GMP-compliant media types efficiently suppressed T cell proliferation irrespective of whether the complete SM medium was supplemented with AB plasma or PL.

Fig. 2 Inhibition of T cell proliferation and requirement for IFN-γ licensing for HUCPVCs to mediate this effect.

(A) Representative dye dilution results measuring inhibition of T cell proliferation by HUCPVCs. HUCPVCs were grown in SMAB, assayed at p4, and cultured with Ef670-stained PBMCs; ratios are PBMC:HUCPVC at outset of culture. PHA, phytohemagglutinin. (B) Inhibition of T cell proliferation with and without IFN-γ licensing. Comparative measurement of inhibitory effects was made by comparing the percentage of T cells remaining in the undivided peak after activation with or without titrated amounts of MSCs. Mean ± SD of three different HUCPVC lines grown in SMPL ratios are PBMC:HUCPVC (no significant differences between licensed and unlicensed groups at each concentration, paired t test).

GMP HUCPVCs express multiple immunoregulatory and proregenerative factors

HUCPVCs were assessed in the unlicensed and licensed state for the expression of a variety of anti-inflammatory and proregenerative factors, compared to lipoaspirate-derived MSCs. Both MSC types expressed steady amounts of transcript for transforming growth factor–β (TGF-β) with and without licensing (raw CT range, 21 to 24; Fig. 3A). Licensing of MSCs strongly increased the expression of TNF-α–stimulated gene 6 (TSG-6) and indoleamine-2,3-dioxygenase (IDO) (raw CT range: unstimulated, 20 to 23; stimulated, 14 to 16) (Fig. 3, B and C). CD274 expression was also significantly increased after licensing in HUCPVCs compared to lipoaspirate-derived MSCs (Fig. 3D). HUCPVCs significantly up-regulated production of CXCL8 and vascular endothelial growth factor (VEGF) protein after licensing (Fig. 3, E and F). Lipoaspirate-derived MSCs also produced increased amounts of CXCL8 and VEGF in response to licensing, although this did not reach significance (Fig. 3, E and F). Both factors are strongly associated with blood vessel formation; VEGF is also a strong T cell activation inhibitor. Measurement of MSC-derived TGF-β was not possible in the SM-based medium because this medium contains TGF-β as a supplement.

Fig. 3 Gene expression and protein production by resting and licensed MSCs.

(A) TGF-β, (B) CD274, (C) IDO, and (D) TSG-6 gene expression in resting and licensed MSCs from lipoaspirate (blue) and HUCPVCs (red), measured by quantitative reverse transcription polymerase chain reaction (qRT-PCR) and expressed as 2(−ΔCT) against expression of RPLP0. Means ± SEM (n = 3 donors), MSC at p3. Where plain bars are absent, gene expression had lower limit of detection and was not included in measurement of significance. (E) CXCL8 and (F) VEGF production by resting (plain bars) and licensed (hatched bars) MSCs from lipoaspirate (blue) and HUCPVCs (red) measured by Luminex 100 (in pg/ml). Means ± SEM (n = 3 donors), HUCPVCs and MSCs at p3. Protein measurements showed net of background measurement of analyte in culture medium alone.

Comparative expression of multiple chemokines in the resting and licensed state of HUCPVCs and adipose tissue MSCs

HUCPVCs and MSCs derived from adipose tissue were assessed in the unlicensed and licensed state with respect to changes in chemokine expression. Chemokine expression profiles associated with an inflammatory state, an anti-inflammatory state, a proangiogenic state (chemokines containing the ELR motif), and a potentially angiostatic state (chemokines lacking the ELR motif) were assessed using four separate analyses. The ELR motif consists of three amino acids (Glu, Leu, and Arg) present in a family of chemokines, which are potent inducers of angiogenic activity (31).

The heat map in Fig. 4 demonstrates major differences between the MSC types, showing anti-inflammatory and proangiogenic properties of HUCPVCs and proinflammatory and potentially angiostatic activity of the lipoaspirate-derived MSCs. The HUCPVCs do not express inflammatory-associated chemokines in either the steady state or after cytokine challenge, whereas lipoaspirate-derived MSCs express a suite of potentially inflammatory chemokines at rest and then switch to a different inflammatory state after cytokine challenge.

Fig. 4 Chemokine expression by resting and licensed lipo-MSCs and HUCPVC.

The expression of inflammatory, ELR+ve, and ELR−ve chemokines by resting (−) and licensed (+) MSCs from lipo-MSC (adipose) and HUCPVCs (umbilical cord) were measured by RT2 Profiler PCR Arrays. The expression of anti-inflammatory genes was measured using qRT-PCR as indicated. All samples were run at the same time on the same plate. In each case, the average 2(−ΔCT) is plotted, and heat maps were generated using Heatmapper software (www2.heatmapper.ca/). Each group of genes was analyzed separately. Genes are clustered using the single linkage method. n > 3 < 6 donors, HUCPVC/MSCs at p3 (expression of CD274, TSG-6, IDO, and TGF-β same samples as Fig. 3A).

Kidney capsule transplantation in immunodeficient mice

To begin to evaluate the therapeutic potential of GMP HUCPVCs, experiments were set up to first examine islet grafts: For this, the kidney capsule route was used. All immunodeficient mice were transplanted with a marginal mass of human islets [3000 islet equivalents (IEQs) per mouse] within 4 weeks of administration of streptozotocin (STZ). Before transplant, there were no differences in glucose concentrations or weights among groups [glucose, 30.1 ± 1.0 mM versus 32.6 ± 1.0 mM (mean ± SEM); weight, 26.4 ± 0.5 g versus 25.6 ± 0.6 g; islet transplant group versus islet + HUCPVCs group, respectively (P > 0.05)]. In the pilot experiments transplanting human islets (4000 IEQs per mouse) within 4 weeks of administration of STZ, there were no differences in glucose concentrations or weights among groups [glucose, 22.7 ± 1.6 mM versus 20.8 ± 0.9 mM versus 24.1 ± 1.4 mM (mean ± SEM); weight, 33.7 ± 0.5 g versus 29.8 ± 1.5 g versus 31.9 ± 0.8 g; islet transplant versus islet + HUCPVCs versus islet + lipo-MSCs (all P > 0.05)]. Tables S1 and S2 show the clinical information for the human islet preparations used in these kidney capsule transplant studies, including donor characteristics and in vitro static glucose-stimulated insulin release (GSIS) and oxygen consumption rate (OCR) normalized to DNA.

IEQ:HUCPVCs (1:30) in a marginal islet transplant model in immunodeficient mice produced optimal long-term metabolic results

In the initial pilot experiments, 4000 IEQs were transplanted in nonobese diabetic (NOD) severe combined immunodeficient (SCID) γ (NSG) mice with or without 1 × 105 HUCPVCs or lipoaspirate-MSCs. Lipoaspirate-MSCs did not enhance islet function (fig. S3A). In mice receiving islets alone or islets plus HUCPVCs, glycemic control was tighter at 6 weeks after transplant (fig. S3B and tables S1 and S2, islet preparation 1). Stimulated blood glucose concentrations were below counterpart control NSG mice that had not received STZ (with normal glucose tolerance; fig. S3B). Therefore, a marginal islet mass approach was adopted, and subsequent experiments were performed using 3000 IEQs per mouse via the kidney capsule route, and only HUCPVCs were tested further for their ability to optimize islet transplant (tables S1 and S2, islet preparation 2).

Mice receiving 3000 IEQs plus 90,000 HUCPVCs (1:30 IEQ:HUCPVCs) reached “cured” blood glucose concentrations after transplant (defined as nonfasted plasma glucose <11 mM after STZ and transplant) faster compared to mice receiving grafts of islets alone or lower numbers of HUCPVCs (post hoc tests, P < 0.001; Fig. 5A). The average number of days to cure was 16.0 ± 1.7, 12.0 ± 2.6, 3.0 ± 0.4, and 9.0 ± 3.6 for mice receiving grafts of 1:0, 1:10, 1:30, and 1:50 IEQ:HUCPVCs, respectively.

Fig. 5 Time to cure diabetes with IPGTTs in NSG mice receiving 3000 IEQs and HUCPVCs.

(A) All mice were transplanted with 3000 IEQs plus different ratios of IEQ:HUCPVCs (1:0, 1:10, 1:30, and 1:50). Time to normoglycemia (nonfasted glucose <11.0 mM) was determined. Tail vein sampling for glucose concentrations (mM) during IPGTT (2 g/kg) was done at 0, 15, 30, 60, 90, and 120 min after IPGTT. Stimulated C-peptide concentrations (pM) were sampled at 60 min after IPGTT. Glucose concentrations at (B) 2.8 weeks, (C) 7 weeks, (D) 12 weeks, and (E) 16 weeks after islet transplant. Stimulated C-peptide concentration (pM) divided by glucose concentration (mM) at (F) 12 weeks and (G) 16 weeks. n = 6 to 8 mice per group. Data presented as mean ± SEM. See Table 1 for additional information. Black bar, 3000 IEQs alone; red bar, 1:10 ratio of IEQ:HUCPVCs; red horizontal lined bar, 1:30 ratio of IEQ:HUCPVCs; red vertical lined bar, 1:50 ratio of IEQ:HUCPVCs. Post hoc analyses for significant differences between ratios of IEQ:HUCPVCs are denoted: a, 1:0 versus 1:10; b, 1:0 versus 1:30; c, 1:0 versus 1:50; d, 1:10 versus 1:30; e, 1:10 versus 1:50; f, 1:30 versus 1:50.

At 2.8 weeks after transplant, glucose control as determined in the intraperitoneal glucose tolerance test (IPGTT) was superior in mice transplanted with islets plus HUCPVCs, with no statistical significance between the islet plus HUCPVCs groups (Fig. 5B and Table 1). At 7 weeks after transplant, glucose control determined by IPGTT was superior with 3000 islets plus 90,000 HUCPVCs (ratio of 1:30 IEQ:HUCPVCs), with no significant difference between grafts with islets alone and other ratios of HUCPVCs (Fig. 5C and Table 1). At 12 weeks after transplant, the tightest glycemic control was seen in mice receiving 1:30 and 1:50 IEQ:HUCPVCs, with no difference in those receiving islets alone and 1:10 IEQ:HUCPVCs (Fig. 5D). By 16 weeks, the best glycemic control was seen in mice transplanted with 1:30 IEQ:HUCPVCs (Fig. 5E). On two-way analysis of variance (ANOVA), there was an effect of time after transplant on the integrated glucose response, with an interaction between time after transplant and numbers of HUCPVCs received. However, post hoc analyses examining the effect of time (from 2.8 to 16 weeks) on graft function found no deterioration with the 1:30 IEQ:HUCPVCs grafts.

Table 1 AUC of glucose concentrations after IPGTT.

Data presented as mean ± SEM. The integrated area under the curve of glucose concentrations was calculated during the 120-min IPGTT (mM × min2) at 2.8, 7, 12, and 16 weeks after islet transplant; n = 6 to 8 mice per group. One-way ANOVA analyses comparing effect of integrated glucose response with ratio of islets: HUCPVCs transplanted under the kidney capsule were compared at separate time intervals. Post hoc analyses for significant differences between ratios of islets to HUCPVCs are denoted: a, 1:0 versus 1:10; b, 1:0 versus 1:30; c, 1:0 versus 1:50; d, 1:10 versus 1:30; e, 1:10 versus 1:50; f, 1:30 versus 1:50.

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The ratios of circulating C-peptide to glucose concentration at 60 min concurred with these observations and were elevated in mice transplanted with 1:30 IEQ:HUCPVCs at 12 and 16 weeks (Fig. 5, F and G). Control experiments were performed in STZ-treated mice receiving Matrigel alone versus HUCPVCs in Matrigel. Glucose concentrations did not fully normalize in either group; however, glucose concentrations were lower in mice receiving HUCPVCs in Matrigel versus Matrigel alone (fig. S4).

Confirmatory studies demonstrate hyperglycemia after removal of islet grafts

After 16 weeks, mice with transplants (n = 5) underwent left nephrectomies to remove islet grafts; these mice reverted to diabetes, and blood glucose concentrations increased from 5.0 ± 0.1 mM on the day of surgery to hyperglycemia (18.8 ± 1 mM) 3 days after surgery. These results confirmed that the normoglycemic state was a consequence of islet transplant rather than the regeneration of mouse islets in the pancreas (fig. S5).

Vessel density in islet graft was increased with islets co-transplanted with HUCPVCs

To determine whether vascularization was increased in the grafts where islets were cotransplanted with HUCPVCs, vessel density was quantified within the islet grafts. Vessel density was increased in islet grafts cotransplanted with 3000 islets plus HUCPVCs, in ratios of 1:10 and 1:50 (Fig. 6, A and B), as compared to grafts of islets alone. Results were confirmed using immunofluorescence analysis with a further endothelial marker to ETS-related gene (ERG) (fig. S6). Grafts with 4000 islets and 1 × 105 HUCPVCs were also examined and demonstrated nonsignificantly greater vessel density in these grafts with HUCPVCs (P = 0.07, fig. S7).

Fig. 6 Vessel density in islet grafts implanted under kidney capsule at 16 weeks after transplant.

Diabetic NSG mice were transplanted with 3000 IEQs ± HUCPVCs (IEQ:HUCPVC ratios of 1:0, 1:10, 1:30, and 1:50). Serial sections (5 μm) in n = 5 mice per group were cut through the entire islet graft below the kidney capsule, and vessel density associated with the islet graft was quantified. (A) Vessel density at 16 weeks after transplant; data presented as mean ± SEM. Black bar, 3000 IEQs alone; red bar, 1:10 ratio of IEQ:HUCPVCs; red horizontal lined bar, 1:30 ratio of IEQ:HUCPVCs; red vertical lined bar, 1:50 ratio of IEQ:HUCPVCs. Post hoc analyses for significant differences between ratios of IEQ:HUCPVCs are denoted: a, 1:0 versus 1:10; b, 1:0 versus 1:30; c, 1:0 versus 1:50; d, 1:10 versus 1:30; e, 1:10 versus 1:50; f, 1:30 versus 1:50. (B) Immunofluorescence image of a representative section showing vessel quantification in an islet graft. Endothelial cells (isolectin B4) and mature vessels (anti–smooth muscle α-actin antibody) were identified by immunostaining. DAPI, 4′,6-diamidino-2-phenylindole.

In vivo syngeneic, allogeneic, and human islet transplants in mice via HPV. To examine the metabolic function of islets with HUCPVCs, transplant experiments were done via the HPV into the liver, emulating the clinical route used in man. Islet transplantation in man involves transplantation of islets from a donor with a different tissue and blood type to the recipient, and islets may be rejected in part secondary to alloimmune mechanisms. To model this allogeneic transplant in mice, immunocompetent C57Bl/6 mice may be transplanted with islets from BALB/c mice, eliciting a brisk rejection of transplanted BALB/c islets. A marginal BALB/c islet mass (500 IEQs per mouse) was used, and IEQ:HUCPVC ratios of 1:0, 1:90, 1:150, and 1:210 were explored. In a second experiment, a curative mass was used (900 IEQs per mouse), comparing IEQ:HUCPVC ratios of 1:0 versus 1:150. To investigate syngeneic transplant, immunocompetent C57Bl/6 mice were transplanted with islets from C57Bl/6 mice within 9 days of administration of STZ. Last, immunodeficient NSG mice were transplanted with human islets from a single islet isolation (donor 3; tables S1 and S2).

Before transplant, all groups of mice had established diabetes with blood glucose readings >17 mM, and there were no differences in glucose concentrations or weights between treatment groups (P > 0.05; table S3). There was no evidence of cellular-mediated rejection in the liver after transplantation of 150,000 HUCPVCs with mouse islets via the HPV of immunocompetent C57Bl/6 mice at days 1 and 8 after transplantation (fig. S8).

IEQ:HUCPVCs (1:90 and 1:150) via the HPV produced optimal metabolic results in short-term allogeneic experiments. Using a marginal islet mass approach, transplant of 500 IEQs per mouse ± HUCPVCs was tested to assess optimized islet function up to 10 days after transplant. Islets from a BALB/c mouse were transplanted into a C57Bl/6 mouse via the HPV at IEQ:HUCPVC ratios of 1:0, 1:30, 1:90, and 1:150. Blood glucose concentrations decreased most in the mice transplanted with 1:90 and 1:150 IEQ:HUCPVCs (Fig. 7A), although this decrease did not reach statistical significance. Blood glucose concentrations did not change in mice transplanted with islets alone. Blood glucose concentrations reached a nadir at days 2 and 3 after transplant, but glycemic control deteriorated in all groups by day 8 after transplant, reflecting rejection of the islet grafts.

Fig. 7 Blood glucose concentrations after islet transplant ± HUCPVCs in mismatch (allogeneic) and syngeneic transplants.

(A) Immunocompetent C57Bl/6 mice (n = 4 to 6 per group) were transplanted with allogeneic BALB/c islets (n = 500 IEQs) and HUCPVCs (ratios, 1:0, 1:30, 1:90, and 1:150). Mean values of glucose concentration are shown (P > 0.05). (B) Glucose concentration over time, comparing islet-only grafts (n = 900 IEQs) to IEQ:HUCPVCs (1:150) in immunocompetent C57Bl/6 mice transplanted with BALB/c islets (n = 8 mice per group). (C) Immunocompetent C57Bl/6 mice (n = 8 per group) were transplanted with syngeneic C57Bl/6 mouse islets and HUCPVCs (ratios, 1:0, 1:90, 1:150, and 1:210). Mean values of glucose concentration are shown. Tx, transplant. (D) Stimulated (60 min) insulin concentration (pM) after IPGTT divided by glucose concentration (mM) at 6 weeks in immunocompetent syngeneic islet transplants. n = 8 mice per group. Data are presented as mean ± SEM.

The experiments were repeated with 900 IEQs per mouse ± HUCPVCs in a ratio of 1:150 versus 900 islets alone in an attempt to cure the mouse of diabetes and to ascertain whether HUCPVCs ameliorated or delayed graft rejection. Five of six mice receiving HUCPVCs were cured of diabetes at day 1 versus three of seven mice receiving islets alone. There was a slower increment in blood glucose concentrations in all mice receiving islets plus HUCPVCs, consistent with delayed graft rejection (Fig. 7B). Frank hyperglycemia (plasma glucose ≥13 mM) was evident by day 5 in all mice receiving islets alone, whereas hyperglycemia in the mice receiving islets plus HUCPVCs was delayed until day 8 (P = 0.04).

IEQ:HUCPVC (1:150) transplanted via the HPV in an immunocompetent syngeneic model produced optimal metabolic results in long-term experiments. Using a marginal islet mass approach, 500 IEQs were implanted per mouse with or without HUCPVCs to assess optimized islet function up to 6 weeks after transplant. Islets from C57Bl/6 mice were transplanted into C57Bl/6 recipient mice via the HPV in the following IEQ:HUCPVC ratios: 1:0, 1:90, 1:150, and 1:210.

Blood glucose concentrations decreased in the mice transplanted with 1:150 IEQ:HUCPVCs, and these mice maintained the tightest glycemic control over the 6-week duration of the experiment (Fig. 7C). The mice receiving an IEQ:HUCPVC ratio of 1:150 had the greatest insulin:glucose concentrations after IPGTT 6 weeks after transplant (Fig. 7D). All mice (n = 8) transplanted with 1:150 IEQ:HUCPVCs were cured from their diabetes versus n = 3 to 4 mice in all other groups (χ2 = 7.5; P = 0.02). Liver function tests at cull were normal in both groups, with no significant differences between the groups (fig. S9).

1:150 human IEQ:HUCPVC transplanted via the HPV in an immunodeficient mouse model cured diabetes and led to long-term cure rates. Using a marginal islet mass approach, 700 human IEQs per mouse with or without HUCPVCs in a ratio of 1:150 IEQ:HUCPVCs were transplanted to determine ability to cure diabetes and maintain cure for 7 weeks after islet transplantation. Blood glucose concentrations decreased significantly in the mice transplanted with 1:150 IEQ:HUCPVCs and diabetes in mice was cured, maintaining normoglycemia for the 7-week duration of the experiment (Fig. 8). At the end of the experiment, the average glucose concentration in mice receiving islet-only transplants versus islet ± HUCPVCs was 34.7 ± 7.2 mM versus 11.2 ± 2.6 mM (mean ± SEM, P < 0.01). Six of eight mice transplanted with 1:150 IEQ:HUCPVCs were cured from their diabetes versus n = 0 mice in the group transplanted with islets without HUCPVCs (χ2 = 6; P < 0.01).

Fig. 8 Blood glucose measurement after human islet ± HUCPVC transplant in immunodeficient mice.

Immunodeficient NSG mice (n = 8 per group) were transplanted with 700 human islets ± HUCPVCs in a ratio of 1:150 IEQ:HUCPVCs. Mean ± SEM values are shown; difference between groups, P < 0.01.

DISCUSSION

Here, we demonstrated that GMP-compliant HUCPVCs produced in xeno-free conditions are capable of rapid post-thaw in vitro expansion at p2 to p4, with stability of phenotype throughout passages. HUCPVCs exhibit immunosuppressive properties in the nonlicensed state. We also demonstrate that these specific MSCs are distinct from those derived from lipoaspirate, with anti-inflammatory and proregenerative properties in the licensed state. In vivo, using a kidney capsule route of administration, the ratio of HUCPVCs to human islets, which demonstrated optimal metabolic function (32), was 30:1. Transplanting grafts of this ratio led to rapid control of glucose by 3 days after transplant and superior graft function at both early (2.7 weeks after transplant) and later stages (7, 12, and 16 weeks after transplant). After 100 days, defined as long-term in such models (33), there was no evidence of graft dysfunction, and there was increased vascular density in grafts containing HUCPVCs versus grafts of islets alone.

In immunocompetent mice, we demonstrated that these human umbilical cord MSCs did not elicit an immune response, consistent with other studies (34), and provided proof of concept of using this model in syngeneic islet transplant experiments in the absence of immunosuppression. Transplanting mouse islets with HUCPVCs (1:150 IEQ:HUCPVCs) by the HPV route in immunocompetent mouse models produced optimal glycemic results in the short and longer term (up to 6 weeks after transplant). Previous experiments had not systematically examined the dose of MSCs relative to islets for efficacy regarding glycemic control. We further confirmed that transplantation of human islets with HUCPVCs, transplanted via the HPV route, in immunodeficient mice led to a cure of diabetes in these mice, which was maintained for the 7-week duration of the experiment without evidence of long-term adverse effects on the liver.

The ease of procurement of umbilical cord tissue over other tissue sources, ability to rapidly expand this MSC population, and stable phenotype make this an ideal MSC source for off-the-shelf banking and use (35). MSCs used in these experiments were assayed after a cryopreservation step at p1, thawing cells in xeno-free GMP medium before use, with demonstrable in vivo function. The functional utility of thawed HUCPVCs used immediately after cryopreservation has also recently been demonstrated (36), contrasting with previous reports suggesting that it is necessary to culture bone marrow mesenchymal cells after cryopreservation to restore their immunoregulatory properties (37, 38). In contrast to our findings, some groups have demonstrated that MSCs derived from adipose tissue have a beneficial effect on islet transplantation (39); the source of fat and other donor factors may be critical.

The ability to suppress T cell activation and express TGF-β in the nonstimulated state has not previously been described for MSCs (40, 41) and is beneficial, along with the ability to up-regulate proregenerative factors when induced by inflammation. TSG-6 expression has been recently shown to play a central role in MSC suppression of inflammation in vivo (42), and the HUCPVCs used here highly up-regulated this factor in response to inflammation. At 24 to 48 hours after islet transplantation, there is marked inflammation in the liver (11, 43, 44), which compromises islet survival and engraftment. MSCs may exert protective effects by promoting angiogenesis and immunomodulation (45, 46). With time, as inflammation becomes quiescent, vascularization of islets takes place and is largely complete within 4 to 6 weeks.

Autologous bmMSC trials are underway in different disease areas including patients with type 1 diabetes (47). Furthermore, in certain diseases including graft-versus-host disease (4853), critical limb ischemia (54, 55), and acute myocardial infarction (56), allogeneic bmMSCs have been administered with considerable success, although no phase 3 trials have been reported. No center has performed randomized controlled clinical trials using allogeneic MSCs in patients with type 1 diabetes undergoing islet transplantation. One open-label, randomized pilot study in humans transplanting autologous bone marrow mononuclear cells (bmMNCs) with a small proportion of Wharton’s jelly–derived cells that had not been phenotyped demonstrated evidence of preservation of graft function (23). No conclusions with respect to the effect of the mesenchymal cell population can be reached from these bmMNC studies, although such studies illustrate the clinical utility of MSCs for transplantation in type 1 diabetes.

There are several small studies that have examined autologous bmMSCs in type 1 diabetes and in animal models of islet transplantation. Many studies used an IEQ:MSC ratio of about 1:10 or lower (5759), showing modest effects on graft function but pronounced effects on graft vascularity. Although dosage experiments are difficult to perform and interpret in clinical practice, the optimum ratio that we demonstrate is in excess of previously reported ratios and in concord with MSC numbers used in primate studies (18). In our experiments, an IEQ:MSC dose of 1:10 did not yield sustained effects on graft function. It is of interest that the largest quantity of MSCs in our study did not demonstrate the greatest benefit in terms of graft function, indicating the possibility of a plateau effect with MSCs that may be tissue source dependent. This further emphasizes the importance of our dose response studies. The experimental route of human islet transplantation under the kidney capsule was selected for experiments in immunodeficient mice because islets remain at the site of delivery, enabling quantification of vasculogenesis; we confirmed vascular density using two different methodologies. Islet delivery via the hepatic portal route more closely reflects the clinical scenario; however, the islets tend to disseminate throughout the liver, as well as the systemic circulation, and it is therefore not possible to quantify vasculogenesis. Our HPV studies in immunocompetent mouse models using mouse islets, as well as in immunodeficient mouse models using human islets, inform the optimal ratio of islets to HUCPVCs to use via the hepatic portal route of delivery in man, and we further confirmed longevity of graft function to 7 weeks in these models using a separate isolation of islets.

The fate of MSCs after transplant is largely unknown; tracking cells to determine whether they engraft or differentiate is necessary but remains a limitation of the present work. We presented extensive data regarding expression of inflammatory response modifying factors by the MSCs, which associated with improved transplant outcome. However, the effects of transplanted MSCs on recruitment and modification of other host cell populations in the transplant setting is not yet determined. Immunomodulation and angiogenic factor expression are strongly up-regulated in vitro after licensing with cytokines, and repeating our observations with licensed MSCs could aid interpretation of the therapeutic relevance of the many factors expressed by the MSC. Here, we examined both grafts formulated in Matrigel and transplanted into the kidney, and grafts were introduced to the liver via the HPV, the route used in humans. We cannot robustly compare the numbers of HUCPVCs used successfully in this study to numbers of bmMSCs used in other published studies due to differences in the experimental route of transplant. Other limitations include the HUCPVC studies that were not carried out with immunosuppression in immunocompetent recipients and the experiments that were confined to small animal models using only male animals. Dose escalation studies are required to fully evaluate the effect of HUCPVCs with immunosuppression in man.

An important aspect of these studies is that human GMP-grade HUCPVCs at p3 and human islets were used in immunodeficient mouse models; furthermore, a single large islet donation was used for the main transplant experiments via the kidney capsule and the HPV route in the mice, facilitating interpretation of the findings. It is well recognized that there is considerable donor heterogeneity in islet samples. Thus, such studies are difficult to perform in animal models with respect to timing.

The findings of this study have major clinical implications. At present, subjects with type 1 diabetes need to receive islets from at least two donor pancreata to improve glycemic control; there is a shortage of suitable donor pancreata, and attrition in graft function is seen over time, likely secondary to loss of β cells through autoimmune, alloimmune, and inflammatory mechanisms. Furthermore, more than 60% of islets are lost in the first 24 to 48 hours after transplant (11, 43, 44). Our study demonstrates the mechanism of action of GMP-grade HUCPVCs in vitro and positive effects in vivo on graft function in vivo. These results lay the foundation for future safety studies in animal models, which should include the tracking of MSCs by state-of-the-art molecular techniques and an assessment of long-term safety and tumorigenesis. With such data, it will be possible to proceed with randomized controlled clinical trials in man.

MATERIALS AND METHODS

Study design

The purpose of this study was to explore whether MSCs improved islet transplantation outcomes and to elucidate the mechanism of action. We carried out islet transplantation experiments using the kidney capsule route to examine the islet graft in its entirety, as well as transplantation via the HPV to emulate the transplant route used clinically. MSCs manufactured using GMP-compliant methods from human umbilical cord and adipose tissue were extensively characterized at a phenotypic and molecular level, including expression of inflammation and angiogenesis-modulating chemokines and cytokines.

Human islets were transplanted into immunodeficient mouse models, and mouse islets were transplanted in immunocompetent mouse models. The optimal ratio of islets to HUCPVCs was determined. Sample sizes were determined on the basis of previous experience with similar studies and pilot experiments. All mice used were males to eliminate any potential confounding influences of differences in engraftment secondary to gender. All mice were randomly assigned to treatment groups, and all analyses were performed blinded to treatment condition. No surviving animals were excluded from analysis, and no outliers were excluded. The number of biologic replicates is specified in the figure legends.

Materials

All human tissue (donated material) was fully consented for use in research, and ethical committee approval was obtained {pancreatic islets ethics approval [Scottish National Blood Transfusion Service (SNBTS)] REC reference: 13/NW/0105 Integrated Research Application System project ID: 124650 and sample governance 14-29v2 and University of Alberta Research Ethics Board, Pro00013094}. The use of umbilical cord material was governed under ethics approval [Edinburgh Reproductive Tissue Biobank (ERTBB)–097 and National Health Service (NHS) Greater Glasgow and Clyde Project REC reference GN14NN406]. All blood samples used in immune response assays were governed under the SNBTS sample governance committee approval process permission 14-02.

Generation of GMP-grade HUCPVC

HUCPVCs were generated for use in this study under a collaborative research agreement with Tissue Regeneration Therapeutics Inc. Methodology was based on published methods for establishment of HUCPVC from Wharton’s jelly (25) with modifications to use commercially available xeno-free medium with or without clinical-grade AB plasma or PL supplements. Briefly, umbilical cords were aseptically dissected as described (26), and Wharton’s jelly was placed in culture. The cells are described as HUCPVCs, as Wharton’s jelly was harvested from the distinguished tunica media of the three vessels of the cord [as discussed by Davies et al. (30)]. The cells are thus perivascular stromal cells but not pericytes. Umbilical cord (6 to 10 cm) yielded about 2 ml of dissected material that was used to seed one 175-cm2 flask. Explants were cultured in DMEM (Gibco), heparin (Leo Labs) at a final concentration of 2 IU/ml, and 1× nonessential amino acids (Gibco) supplemented with 25% human PL in Corning CellBIND flasks with no additional substrate. Once the cells had reached 80 to 90% confluence, the adherent MSCs were recovered with a 10-min incubation at 37°C with 10 ml of 1× TrypLE (Gibco). To remove cell debris, the material was passed through a 100-μm cell strainer (Falcon). The cells were counted using a hemocytometer and designated as p1 (for standardization purposes; the growth and time in culture for each cell line were recorded according to doubling rate). These MSCs were then cryopreserved at 1 × 106 per vial in CryoStor (CS10, Sigma Ltd.). Later in the study, further three-cell isolate cells were grown to p1 in SM medium supplemented with 25% PL instead of the DMEM-based medium. Thawed cells were washed once in SM medium and recultured at 3000 cells/cm2 in fresh flasks for up to four passages. Cells were cultivated from p2 to p4 in three GMP-compliant xeno-free media—StemMACS MSC XF (SM) (Miltenyi Biotec), StemMACS MSC XF supplemented with 5% human PL (SM PL), or StemMACS MSC XF supplemented with 10% human AB plasma (SM AB).

Preparation of mouse models before islet transplantation experiments

All animal procedures complied with Home Office Guidance on the operation of the Animals (Scientific Procedures) Act 1986 and conformed to local ethical requirements of the University of Edinburgh. Male NSG (NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ /NSG) mice were used to establish the human islet transplant models, and male C57Bl/6 mice were used to establish syngeneic and allogeneic islet transplant models. Animals (n = 8 to 10 per group) were rendered hyperglycemic at 16 to 17 weeks old by administration of STZ (160 to 180 mg/kg intraperitoneally; Sigma-Aldrich) in ice-cold acetate buffer (pH 4.5) after a 4-hour fast. Daily tail blood glucose measurements were taken using a glucometer (One Touch Verio, LifeScan). Mice were classed as hyperglycemic if their nonfasted glucose concentrations were ≥20 mM for two consecutive days. Once diabetes was confirmed, daily subcutaneous injections (0.5 to 2.5 U) of insulin glargine (Lantus, Sanofi) were administered until the day before islet transplantation. All islet transplantations took place within 4 weeks of administration of STZ.

Procurement and preparation of human islets and HUCPVCs

Three human islet preparations were isolated from donor pancreata by the islet isolation laboratories using the Edmonton protocol (2). In the clinical islet isolation laboratory (SNBTS), those islet preparations that did not meet clinical release criteria because of numbers (<200,000 IEQs) and that were consented for research purposes were used in the laboratory. Because of the inherent heterogeneity of donor islet preparations, one large islet preparation was used in the longitudinal NSG mouse experiments with 3000 IEQs transplanted via the kidney capsule route as described (tables S1 and S2). Before this one, preparation had been used in pilot experiments that used 4000 IEQs transplanted via the kidney capsule route. Similarly in the HPV transplants in NSG mice, one large islet preparation from a single islet isolation (donor preparation 3; Alberta Diabetes Institute, Islet Core Facility) was transplanted. Upon receipt of islets by the research laboratory, the islets were cultured in CMRL-1066 media (Mediatech), supplemented with 10% (v/v) fetal calf serum (Scientific Laboratory Supplies), heparin (10 U/ml; Leo Pharma), penicillin-streptomycin (100 U/ml and 100 μg/ml, respectively; Gibco), and 2 mM l-glutamine (Gibco), at 22°C/5% CO2. All HUCPVCs for in vivo use were thawed from the p1 stock and grown to p3 for use. Cells were cultured in SM (Miltenyi Biotec) and 5% human PL.

Islet transplantation in immunodeficient NSG mouse model via kidney capsule route

Islet transplantation was carried out at 4000 IEQs initially ±1 × 105 lipoaspirate MSC or HUCPVC with relevant control groups in the initial studies (fig. S1). Subsequently, a marginal mass model using 3000 IEQs was transplanted with three ranges of HUCPVCs (ratios: 1:10, 1:30, and 1:50) with parallel control groups including with islets alone, Matrigel, and HUCPVCs plus Matrigel (six groups).

In all cotransplantation experiments of MSC plus islets, both islets and HUCPVCs were placed into 0.5-ml Eppendorf tubes in coculture for an hour at room temperature before transplant. Excess medium was drawn off to leave an islet pellet, to which 10 to 20 μl of ice-cold Matrigel (reduced growth factor, Corning) were added. The islet-Matrigel solution was drawn up into PE50 tubing just before transplant.

All mice were 20 to 21 weeks old at the time of transplant. Mice were anesthetized using isoflurane (1.5 to 3%), the left kidney was exposed under aseptic conditions, and the Matrigel (±islets ± HUCPVCs) was transplanted under the left kidney capsule. The wound was sutured and clipped before recovery. Control experiments were performed at the same time in the same cohorts of mice.

In vivo assessment of efficacy of HUCPVCs in islet engraftment and function in an immunodeficient NSG mouse model and immunocompetent C57Bl/6 mouse model

The function of the islet plus MSC grafts was assessed with an IPGTT at 2.7, 7, 12, and 16 weeks after transplantation under the kidney capsule. NSG mice were fasted overnight and, after a basal tail blood glucose reading, received 2 g of glucose per kilogram of fasted body mass by intraperitoneal injection. Glucose measurements were then taken at 15, 30, 60, 90, and 120 min after glucose injection. A blood sample was taken at the 60-min time point (into EDTA) for stimulated plasma C-peptide analysis. At the 16-week IPGTT, further blood samples were also taken at 0- and 120-min time points. IPGTTs were similarly carried out after transplantation of C57Bl/6 mouse islets via the HPV route into C57Bl/6 mice. At 6 weeks after transplant, a blood sample was taken while fasting and at the 60-min time point (into EDTA) for stimulated plasma insulin analysis.

Unilateral left nephrectomy as confirmation of insulin-secreting function of islets ± HUCPVCs engrafted kidney

The left kidneys of half the mice (receiving 1:30 or 1:50 IEQ:HUCPVCs at transplant) were removed via laparotomy at day 101, defined as the point of long-term engraftment (60). These mice were followed for a further 5 days to confirm the return of hyperglycemia. All mice were culled after the IPGTT at 16 weeks, and histology, including assessment of vascular density, was performed.

Mouse islet isolation and culture

Pancreatic islets were isolated from C57BL/6, BALB/c mice (9 weeks old, 20 to 22 g) by a collagenase digestion method (61). The islets (250 islets/ml) were cultured free floating (37°C, 5% CO2) in RPMI 1640 (BioWhittaker, Walkersville, MD) supplemented with l-glutamine (Sigma), penicillin-streptomycin (1000 U/ml and 10 mg/ml, respectively; Sigma), and 10% (v/v) fetal calf serum (HyClone, Celbio, Logan, UT) for up to 12 hours before the transplant. Islet purity was ≥90%.

Transplantation experiments via the HPV route in immunocompetent mice

All mice were 12 to 14 weeks old at the time of transplant. Animals were anesthetized using isoflurane (1.5 to 3%), and HPV transplant was performed after a midline laparotomy exposing the HPV with subsequent injection of islets into the vein using a 27-gauge needle followed by control of bleeding and resuturing of the skin wound.

Islets ± HUCPVC transplant experiments in mice via the HPV route

HUCPVCs (n = 150,000) plus islets versus control experiments with islets alone (500 IEQs per mouse) were injected into C57Bl/6 mice via the HPV route. Mice were sacrificed at days 3 and 8 (n = 8 per group). The livers were removed, step-sectioned (5 μm) and hematoxylin and eosin–stained for an immune infiltrate, and quantified using a Nikon Eclipse TE2000-S microscope.

Allogeneic islet ± HUCPVC transplants from BALB/c into C57Bl/6 mice

Allogeneic experiments used islets from BALB/c mice transplanted into C57Bl/6 mice via the HPV route with IEQ:HUCPVC ratios of 1:0, 1:30, 1:90, and 1:150 (n = 6 per group; n = 500 IEQs per mouse). Mice were sacrificed at day 10 after transplant.

In a second series of experiments, mice were transplanted with IEQ:HUCPVC ratios of 1:0 and 1:150 (n = 8 per group; 900 IEQs per mouse). Mice were sacrificed at day 21 after transplant.

Syngeneic islet ± HUCPVC transplants from C57Bl/6 donor and recipients

Syngeneic experiments used C57Bl/6 mice as donors and recipients with IEQ:HUCPVC ratios of 1:0, 1:90, 1:150, and 1:210 (n = 7 to 9 per group). Mice were sacrificed at 6 weeks after transplant. At-cull bloods were analyzed for liver function tests (aspartate aminotransferase, alanine aminotransferase, and albumin).

Statistical analysis

Data were tested for normality, and transformations were applied as appropriate. The integrated area under the curve (AUC) for glucose was calculated by the trapezoid rule (62). Glucose AUC measurements, time to cure diabetes with IEQ ± HUCPVCs, 60-min C-peptide:glucose ratios, and graft vessel densities were compared by ANOVA with post hoc testing between groups. Gene expression and protein concentrations were examined by ANOVA or, in the case of two datasets, unpaired t tests or Mann-Whitney U test as specified. Statistical significance was taken as P < 0.05. Group sizes of n = 6 for the mouse studies (59) demonstrated 80% power to detect a 20% difference in response at the 95% level of significance.

SUPPLEMENTARY MATERIALS

stm.sciencemag.org/cgi/content/full/12/526/eaan5907/DC1

Materials and Methods

Fig. S1. Characterization of lipoaspirate-derived MSCs in vitro.

Fig. S2. Inhibition of T cell proliferation by HUCPVC.

Fig. S3. Characterization of lipoaspirate-derived MSCs in vivo.

Fig. S4. AUC glucose 6 weeks after transplant.

Fig. S5. Glucose concentrations after removal of kidney islet graft by nephrectomy.

Fig. S6. Immunofluorescence analysis with endothelial marker ERG.

Fig. S7. Vessel density in islet grafts with and without HUCPVCs.

Fig. S8. Histological assessment of cell infiltrate into liver after syngeneic islet grafts ± HUCPVC.

Fig. S9. Liver function 6 weeks after transplant.

Table S1. Human islet preparation and associated clinical characteristics.

Table S2. GSIS and OCR of islet preparations.

Table S3. Glucose concentrations and weights before surgery.

Data file S1. Primary data.

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REFERENCES AND NOTES

Acknowledgments: We thank the Scottish Clinical Islet Transplant Laboratory and the Alberta Diabetes Institute, Islet Core Facility for provision of human islets; the Scottish National Blood Transfusion Service for manufacturing and immune testing of MSCs; and the Edinburgh Reproductive Tissue Biobank for provision of umbilical cords. We acknowledge the technical support of J. Henderson and the Centre for Biological Services. Funding: Chief Scientist Office (ETM/325 and TCS/17/31) to S.F. and J.D.M.C., Diabetes UK (BDA 13/0004682) to S.F., Wellcome Trust–University of Edinburgh Institutional Strategic Support Fund to S.F. and J.D.M.C., and the Edinburgh and Lothians Health Foundation Award to S.F. R.N.C. and N.M.M. were funded by a Wellcome Trust Investigator Award (100981/Z/13/Z to N.M.M.). We acknowledge NHS Scotland, the UK Islet Transplant Consortium, which has received funding from Diabetes UK (BDA 06/0003362), Diabetes Research and Wellness Foundation, and the Juvenile Diabetes Research Foundation. We acknowledge the research infrastructure support of the British Heart Foundation Centre of Research Excellence, Edinburgh. We are grateful to the Society for Endocrinology for supporting S.F.’s laboratory visit to the Islet Transplant Programme Edmonton, Alberta, Canada. We also gratefully acknowledge the support and advice from Tissue Regeneration Therapeutics (Toronto, Canada) as part of a research collaboration agreement to generate HUCPVC in this study. Author contributions: S.F.: principal investigator for all rodent in vivo metabolic and in vitro experiments and histological analyses; all rodent experiments were performed in S.F.’s laboratory. J.D.M.C.: principal investigator for MSCs and immunology experiments; S.F., J.D.M.C., M.L.T., J.C.M., and D.C. contributed to the concept and design of the experiments. S.F. and J.D.M.C. drafted and revised manuscript and performed statistical analyses. A.R.B., J.N., G.B., P.B., and K.S. performed animal experiments. J.N., G.B., and A.R.B. performed histological analyses. P.S.L. assisted with immunohistochemical techniques and analysis. D.C., K.S., J.C.M., P.B., and K.L.T. manufactured, maintained, and characterized MSCs including T cell assays. K.L.T. and G.J.G. performed all gene analysis. K.L.T., K.S., N.W.A.M., and A.R.F. performed cytokine analyses. A.R.B., J.N., P.B., R.N.C., and N.M.M. assisted and advised with metabolic assays. A.R.B. helped draft methods. All authors commented and reviewed the manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data associated with this study are present in the paper or the Supplementary Materials. Islets from the Alberta Diabetes Institute, Islet Core Facility were obtained under a material transfer agreement.

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