DORmO mice develop progressive, HA-associated insulitis. DORmO mice developed autoimmune insulitis at approximately 4 weeks of age, with nearly all mice becoming diabetic (blood glucose >250 mg/dl) by 20 weeks of age (Supplemental Figure 1A; supplemental material available online with this article; doi:10.1172/JCI79271DS1). This pattern was observed irrespective of gender (Supplemental Figure 1B) or weight (Supplemental Figure 1C). The production of insulin by the islets progressively decreased over time (Figure 1, A–G, and Supplemental Figure 1D), while lymphocytic (CD3+) infiltrates increased (Figure 1, H–N).

Figure 1 Islet HA accumulates in tandem with progressive autoimmune insulitis in DORmO mice. Representative histologic staining of pancreatic tissue from BALB/c (control) and DORmO mice and average islet area positive over time for (A–G) insulin (INS), (H–N) CD3, and (O–U) HA. For G, N, and U, at least 25 islets were visualized per mouse, and staining and data are from n = 6–8 mice per condition. Original magnification, ×40. Data represent mean ± SEM; *P < 0.05 vs. control for each time point by unpaired t test.

Alongside these changes, the average islet area that stained positive for HA increased markedly during the progression to diabetes in DORmO mice (Figure 1, O–U). HA peaked at approximately 12 weeks of age (Figure 2A), around the time that these animals typically first become hyperglycemic (Supplemental Figure 1A). Our biochemical analyses indicated that at 15 weeks pancreatic HA content was nearly double that seen in control (BALB/c) mice (Figure 2B).

Figure 2 Islet HA deposition is temporally associated with insulitis and not hyperglycemia. (A) Percentage of islet area positive for HA staining of DORmO and BALB/c (control) mice over time (n = 6–15). (B) Pancreas and (C) plasma HA content in BALB/c and DORmO mice over time (n = 6). (D and E) Representative HA staining of pancreatic islet tissue from (D) a BALB/c mouse 1 week after STZ treatment and (E) a 12-week-old diabetic db/db mouse. Original magnification, ×40. Data represent mean ± SEM; *P < 0.05 vs. control for each time point by unpaired t test.

To discern whether the increase in islet HA was associated with increased synthesis or reduced catabolism of HA, we evaluated the expression of HA synthase or degradative (hyaluronidase) enzymes. Analyses of islet mRNA from 8-week-old mice showed that HA synthase 3 (Has3) was upregulated, while hyaluronidase 1 (Hyal1) was downregulated. However, hyaluronidase 2 (Hyal2) expression was increased, while that of Rhamm and layilin, two HA receptors that contribute to HA clearance, was increased (Supplemental Figure 2, A–G). This complexity may reflect the multiple cell types present within inflamed islets, with potentially disparate patterns of HA synthesis and catabolism.

We asked whether HA was systemically increased in DORmO mice. Levels of circulating HA were unchanged (Figure 2C), and we did not see heightened HA deposition in the heart, lung, or liver (data not shown).

We also considered whether islet HA deposition was driven by hyperglycemia. Arguing against this, islet HA was increased at 4 weeks (Figure 2A), well before the onset of hyperglycemia (Supplemental Figure 1A). In hyperglycemic mice in which the etiology of diabetes was not autoimmune, namely mice treated with the β cell toxin streptozotocin (STZ) (Figure 2D), and db/db mice (Figure 2E), less intraislet HA was seen, and this was not associated with insulitis (16). Consistent with this, we only observed substantial intraislet HA deposits in cadaveric islets from subjects with recent-onset T1D (6).

We next examined the relationship between HA and infiltrating lymphocytes during insulitis. In DORmO mice, we observed that HA was abundant in islets under autoimmune attack but was largely absent in neighboring islets without insulitis (Figure 3A). The same pattern was seen in islets in cadaveric human tissues (Figure 3B). In BALB/c control mice without insulitis, HA was concentrated around vascular tissues and in the interstitial spaces (Figure 3, C and E). In contrast, HA in DORmO mice was most abundant around lymphocytic infiltrates (Figure 3, D and F). This distribution again mirrored patterns of HA deposition in human cadaveric pancreatic tissues from subjects with recent-onset T1D (Figure 3, G and H).

Figure 3 HA deposits characterize sites of insulitis in DORmO and human T1D. (A and B) HA staining in pancreatic tissue isolated from (A) a DORmO mouse and (B) a human cadaveric donor with T1D. Infiltrated islets are circled in red; unaffected islets are circled in black. (C and D) HA staining of representative BALB/c and DORmO islets, demonstrating interstitial (orange arrowhead) and peri-islet (blue arrowheads) patterns of HA distribution. HA associated with lymphocytic infiltrates (red arrows) was only seen in DORmO mice. (E and F) Costaining of HA and DAPI, demonstrating HA accumulation in association with insulitis. (G and H) HA deposits in human insulitis (red arrows) from two cadaveric donors with T1D. Original magnification, ×40.

Together with our recently published work on human T1D (6), these data indicate that HA was temporally and anatomically associated with infiltrating lymphocytes in autoimmune diabetes.

It is notable that we observed some differences in the distribution of HA between mice and humans. In mice, a thin sheath of HA surrounded islets (Figure 3, C and E, and ref. 17), whereas this was less pronounced in humans (Figure 3, G and H). Furthermore, the progressive accumulation of HA during insulitis in DORmO mice was centripetal, spreading from the periphery of islets inward, while HA was more patchy in human T1D (Figure 3, G and H). These patterns mirror the known distribution and progression of insulitis in these two species (18).

Hyaladherins are altered during T1D progression. Along with changes in HA, we observed progressive alterations in both the amount and the distribution of hyaladherins in DORmO insulitis. Both TSG6 (Supplemental Figure 3, A–F, and S) and IαI (Supplemental Figure 3, G–L, and U) decreased in the DORmO pancreatic islets during disease progression. Tsg6 mRNA, however, was increased (Supplemental Figure 3T), while IαI-encoding mRNA expression decreased (Supplemental Figure 3V). Staining for versican was largely unchanged (Supplemental Figure 3, M–R, and W), but mRNA expression increased (Supplemental Figure 3X). These data indicate that insulitis was associated with extensive changes in the islet ECM.

4-MU treatment prevents progression to diabetes. While our data suggested that HA deposits are temporally and anatomically associated with insulitis, it was unclear whether HA contributed to disease pathogenesis. We therefore administered 4-MU to DORmO mice to test whether inhibition of HA synthesis could prevent autoimmune diabetes.

Adding 4-MU to chow significantly reduced HA content in DORmO islets (Figure 4, A–E). HA content was likewise reduced in isolated islets that were cultured overnight in 4-MU (Figure 4F). These data are consistent with the established role of 4-MU as an inhibitor of HA synthesis in other tissues (19, 20).

Figure 4 Inhibition of HA synthesis prevents progression to autoimmune diabetes. (A–D) Representative HA staining of pancreatic tissue from BALB/c (control) and DORmO mice fed 4-MU chow or control chow, beginning at 8 weeks of age, a month after the typical onset of insulitis in this model. (E) Average islet HA+ area in these same mice. 25 islets were visualized per mouse, and staining and data are for 6 mice per condition. (F) HA content in islets from BALB/c mice cultured with 50 μg/ml 4-MU or media alone. Data are for 100 islets, and experiments were done in triplicate. (G) Blood glucose of DORmO and BALB/c (control) mice fed 4-MU chow or control chow, beginning at 8 weeks of age, and maintained on 4-MU for 1 year (n = 12 mice per group). (H) Blood glucose of NOD mice fed with control chow or fed with control chow including a 1 week of 4-MU treatment from 5 to 6 weeks of age (n = 10 mice per group). Original magnification, ×40. Data represent mean ± SEM; *P < 0.05 vs. respective control and control for each time point by unpaired t test.

Together with this effect on HA, 4-MU treatment starting at 8 weeks of age prevented progression to hyperglycemia in DORmO mice (Figure 4G). Indeed, DORmO mice were normoglycemic during 4-MU treatment for up to a year, indicating that 4-MU–mediated prevention of diabetes was sustainable and efficacious in this model over long periods of time.

Further, treatment of NOD mice with 4-MU for 1 week prevented the subsequent progression to hyperglycemia in these animals (Figure 4H). Of note, insulitis is typically already well established in this model by the age at which mice were treated with 4-MU (5 weeks of age) (Supplemental Figure 4 and ref. 21). Moreover, HA deposition is also a feature of insulitis in NOD mice as well (16). These data indicate that 4-MU can prevent diabetes in multiple mouse models of T1D.

To evaluate whether 4-MU eliminated autoreactive T cells, we took DORmO mice made normoglycemic by 4-MU off of this drug. These mice rapidly became hyperglycemic (Figure 5A), suggesting that 4-MU suspends autoimmune destruction without eliminating the potential for autoimmunity.

Figure 5 4-MU treatment prevents progression of insulitis but does not cure established diabetes. (A) Blood glucose of DORmO and control mice started on 4-MU at 8 weeks of age, taken off 4-MU between 15 and 18 weeks of age, and restarted on 4-MU thereafter (n = 10). (B) Blood glucose of DORmO and BALB/c (control) mice fed 4-MU chow, beginning at 12 weeks of age (n = 10). (C) Blood glucose following IPGTT for control chow–fed BALB/c and DORmO mice at 10 weeks of age (n = 8 mice per group). (D) Blood glucose following IPGTT of DORmO mice from C treated for 2 weeks with 4-MU or control chow. (E) Blood glucose following IPGTT of BALB/c and DORmO mice at 10 weeks of age following 4-MU treatment for 2 weeks (n = 6). (F) Blood glucose following IPGTT for the same mice as in E, now made hyperglycemic by STZ treatment (n = 6). Data represent mean ± SEM.

To test whether 4-MU promoted regeneration of β cells, we initiated 4-MU treatment at 12 weeks of age, by which time DORmO mice are typically borderline hyperglycemic and have substantially diminished β cell mass. 4-MU treatment did not prevent the onset of diabetes in these animals (Figure 5B) or improve glycemic control, as measured by intraperitoneal glucose tolerance testing (IPGTT) (Figure 5, C and D). Further, we asked whether mice rendered diabetic via STZ treatment had improved glycemic control on 4-MU, and they did not (Figure 5, E and F). Thus, these data did not support the hypothesis that 4-MU promoted regeneration of β cells or improved metabolic control of hyperglycemia. Instead, these data suggested that 4-MU treatment suspended the progressive deterioration of insulin production otherwise observed in DORmO mice and that this effect was predicated upon the presence of viable β cells.

4-MU treatment establishes regulatory checkpoints in insulitis. By 15 weeks of age, insulin-producing β cells were characteristically lost in DORmO mice (Figure 6, A and B). However, in DORmO mice fed 4-MU, insulin staining was preserved (Figure 6, C–E), consistent with these mice remaining normoglycemic (Supplemental Figure 5A).

Figure 6 4-MU treatment promotes nondestructive insulitis. (A–D) Insulin staining of representative pancreatic tissue sections from DORmO and BALB/c mice fed either 4-MU or control chow. (E) Average insulin+ area of islets for these mice. 25 islets were visualized per mouse, and staining and data are for 6 mice per condition. (F and G) Representative images of insulin staining of pancreatic islets from 15-week-old DORmO mice treated with 4-MU for 7 weeks. Original magnification, ×40. Data represent mean ± SEM; *P < 0.05 vs. respective control by unpaired t test.

Robust insulitis was nonetheless still evident in 4-MU–treated DORmO mice (Figure 6, F and G). This effect was uniform across nearly all the islets that we examined. This nondestructive, “respectful” insulitis persisted while mice were maintained on 4-MU.

These effects were not associated with generalized immunosuppression. We observed a reduction in total splenocyte counts but unchanged lymphocyte counts in pancreatic lymph nodes (PLNs), mesenteric lymph nodes (MLNs), or inguinal lymph nodes (ILNs) (Supplemental Figure 5B). Proliferation of splenocytes in response to OVA peptide ex vivo was intact (Supplemental Figure 5C), and 4-MU–treated mice did not have reduced percentages of CD3+ T cells (Supplemental Figure 5D) or CD19+ B cells (Supplemental Figure 5E). Moreover, lymphocytes remained primed to destroy β cells, as evidenced by the persistent insulitis (Figure 6, F and G) and the rapid (<2 week) progression to diabetes that we observed after cessation of 4-MU treatment. Overall, the evidence was not consistent with generalized immune suppression.

We considered whether 4-MU treatment prevented autoimmunity through impaired leukocyte trafficking. However, the histologic data from 4-week-old mice (Figure 1P) indicated that lymphocytosis was typically already established prior to initiation of 4-MU at 8 weeks of age.

4-MU treatment promotes FOXP3 induction in vitro and in vivo. The reestablishment of the “respectful” insulitis that we observed upon histologic staining led us to wonder whether 4-MU treatment might promote peripheral immune tolerance. One major source of immune tolerance is FOXP3+ Tregs.

We indeed observed that the percentage of T cells expressing the Treg marker FOXP3 was increased in islets of DORmO mice after treatment with 4-MU (Figure 7). Along with this increase in Tregs in insulitis, we observed a nonsignificant increase in FOXP3+ Tregs in the spleens, ILNs, PLNs, and MLNs (Supplemental Figure 5F). Together, these data suggested that 4-MU treatment promoted an increase in Tregs in insulitis.

Figure 7 4-MU treatment increases islet FOXP3+ Tregs. (A–D) FOXP3 staining of representative pancreatic tissue sections from DORmO and BALB/c mice fed either 4-MU or control chow. (E) Average FOXP3+ islet area of these mice. At least 25 islets were visualized per mouse, and data are for 6 mice per condition. Original magnification, ×40. Data represent mean ± SEM; *P < 0.05 vs. respective control by unpaired t test.

To evaluate this possibility in the absence of hyperglycemia or other complicating factors, we examined Treg induction upon 4-MU treatment in a pair of in vivo mouse models. First, we assessed the impact of 4-MU on the percentage of GFP/FOXP3+ Tregs in total CD4+ T cells in BALB/c mice fed 4-MU or control chow for 2 weeks. We observed increased proportions of Tregs among CD4+ T cells in the spleens, ILNs, MLNs, and PLNs of the 4-MU–treated animals (Figure 8A) but no change in the total percentage of T cells (Supplemental Figure 6A). Similarly, the percentage of CD19+ B cells (Supplemental Figure 6B), CD86+CD19+MHC-II+ antigen-presenting cells (Supplemental Figure 6C), and activated CD44hiCD4+ T cells (Supplemental Figure 6D) was not altered.

Figure 8 4-MU treatment relieves CD44-mediated inhibition of FOXP3 induction. (A) Percentage of GFP/FOXP3+ Tregs of total CD4+ T cells in BALB/c mice fed 4-MU or control chow for 2 weeks (n = 5–6 mice per group). (B) In vivo induction of FOXP3+ Tregs assessed 4 days after transfer of GFP/FOXP3–CD4+ T cells into Rag–/– hosts given 4-MU or control chow (n = 3 Rag–/– recipient animals). Data are from the spleens of recipient animals. (C) CD25 and FOXP3 expression by CD4+GFP/FOXP3– T cells activated for 72 hours with or without plate-bound HA or anti-CD44 antibody. (D) Pooled data for 3 independent experimental replicates for the representative data in C. (E) FOXP3 induction using Cd44+/+, Cd44–/+, or Cd44–/– precursors. (F) Pooled data for 3 independent experimental replicates for the representative data in E. (G) In vivo induction of FOXP3 assessed using cotransfer of equivalent numbers of GFP/FOXP3–CD4+Cd44+/+CD45.1 and GFP/FOXP3–CD4+Cd44–/–CD45.2 T cells into Rag–/– hosts. After 4 days, the numbers of induced CD3+GFP/FOXP3+ Tregs in the spleens of recipient animals were assessed and the ratio of Cd44–/– Tregs versus Cd44+/+ Tregs was determined (n = 3 Rag–/– recipient animals). Data represent mean ± SEM; *P < 0.05 vs. respective control by unpaired t test.

To test whether 4-MU could alter the peripheral development of Tregs, we then examined the induction of Tregs upon transfer of purified CD4+GFP– (FOXP3–) T cells into RAG-deficient animals in the setting of 4-MU or control chow. We found that 4-MU treatment enhanced the fraction of GFP/FOXP3+ Tregs observed 4 days after transfer (Figure 8B). Together, these results indicated that 4-MU enhances the peripheral differentiation of Tregs.

HA and CD44 suppress FOXP3 induction. Because 4-MU, an inhibitor of HA synthesis, promotes FOXP3 levels, we asked whether HA and the HA receptor CD44 inhibit FOXP3 induction. Indeed, we observed that plate-coated HA and anti-CD44 antibody both diminished FOXP3 induction from CD4+GFP/FOXP3– precursors (Figure 8, C and D).

To better assess the contribution of CD44 to Treg induction, we then performed Treg induction using CD4+GFP/FOXP3– T cells isolated from Cd44+/+, Cd44+/–, or Cd44–/– mice as precursors. T cells from Cd44–/– mice had the greatest FOXP3 induction, with cells from heterozygous Cd44+/– mice and homozygous Cd44+/+ mice exhibiting less FOXP3 induction in inverse proportion to the number of CD44+ alleles they possessed (Figure 8, E and F).

To assess the impact of CD44 on Treg induction in vivo, we then performed a cotransfer of equivalent numbers of GFP/FOXP3–CD4+Cd44+/+CD45.1 and GFP/FOXP3–CD4+Cd44–/–CD45.2 T cells into Rag–/– hosts. After 4 days, the numbers of CD3+GFP/FOXP3+ cells and the ratio of Cd44–/– (CD45.2) vs. Cd44+/+ (CD45.1) Tregs were assessed. We found that Cd44–/– (CD45.2) Tregs represent the majority of these cells in vivo (Figure 8G).

In light of these data, we were initially surprised when we observed that Cd44–/– mice did not naturally have increased numbers of CD4+GFP/FOXP3+ Tregs (Supplemental Figure 7A). However, spleens from Cd44–/– mice had significantly greater total numbers of CD4+ T cells (Supplemental Figure 7B), and CD4+ T cells from these animals had a hyperproliferative response to anti-CD3/28 antibody activation (Supplemental Figure 7C). Both of these findings have been reported previously and attributed to defects in activation-induced cell death pathways (22). The absolute numbers of splenic CD4+GFP/FOXP3+ Tregs were actually higher on average in Cd44–/– mice, although this trend did not reach statistical significance (Supplemental Figure 7D). Together, these data are consistent with an increased generation of Tregs in Cd44–/– mice but suggest that this effect was obscured by heightened proliferation of effector T cells in these same animals.

Inhibition of ERK1/2 signaling partially overcomes CD44-mediated inhibition of FOXP3. We observed that the inhibitory effects of anti-CD44 antibody on FOXP3 induction enhanced anti-CD28 antibody-mediated FOXP3 inhibition (Figure 9, A and B), an effect known to proceed through AKT signaling (23). These data suggested that CD44 signaling might inhibit FOXP3 induction via pathways in addition to or other than AKT.

Figure 9 4-MU treatment promotes FOXP3 induction. (A) FOXP3 levels following induction from CD4+GFP/FOXP3– T cell precursors performed in the setting of anti-CD3, with or without anti-CD28 and/or anti-CD44 antibody costimulation. (B) Pooled data for 4 independent experimental replicates for the representative data in A. (C) Fold change in pERK1/2 MFI over time following CD44 crosslinking. The data shown incorporate 3 experimental replicates. (D) CD25 and GFP/FOXP3 levels following activation of CD4+GFP/FOXP3– T cells in the setting of TGF-β and IL-2, with or without CD44 costimulation and/or the ERK1/2 inhibitor SUO126. (E) Pooled data for 6 independent experimental replicates for the representative data in D. (F–K) CD44 staining of representative pancreatic tissue sections from BALB/c (control) or DORmO mice fed either 4-MU or control chow. (L) Average CD44+ area of islets for these mice. At least 25 islets were visualized per mouse (n = 6). Original magnification, ×40. Data represent mean ± SEM; *P < 0.05 vs. respective control and control for each time point by unpaired t test.

One signaling pathway known to inhibit FOXP3 induction is ERK1/2 (24). We previously reported that, when using human T cells, CD44 signaling promotes phosphorylation of ERK1/2 (pERK1/2) (25). In mouse CD4+ T cells we likewise found that CD44 crosslinking induced pERK1/2 (Figure 9C).

We then asked whether inhibition of pERK1/2 could restore the loss of FOXP3 induction we observed upon anti-CD44 antibody treatment. Using CD4+GFP/FOXP3– T cells activated for 72 hours in the setting of TGF-β and IL-2, we found that SUO126, an ERK1/2 inhibitor, could overcome the inhibition of FOXP3 induction with or without CD44 costimulation (Figure 9, D and E).

Finally, we asked whether infiltrating lymphocytes express CD44. We indeed observed an increase in CD44+ staining during insulitis progression (Figure 9, F–L), suggesting that cells present in insulitis may be responsive to local HA.

Together, these data implicate a role for CD44 and HA in inhibition of Treg differentiation and support the hypothesis that 4-MU treatment relieves this inhibition by reducing HA-mediated CD44 signaling.