Research ArticleOsteoarthritis

Inhibition of TET1 prevents the development of osteoarthritis and reveals the 5hmC landscape that orchestrates pathogenesis

See allHide authors and affiliations

Science Translational Medicine  15 Apr 2020:
Vol. 12, Issue 539, eaax2332
DOI: 10.1126/scitranslmed.aax2332

Preventing progression

The etiology and progression of osteoarthritis, a disease of joint degeneration, remains incompletely known. Studying epigenetics in a mouse model of osteoarthritis, Smeriglio et al. found that ten-eleven-translocation enzyme 1 (TET1) regulated activation of WNT signaling, metalloproteinases, and mTOR via cytosine hydroxymethylation. Tet1−/− mice were protected from osteoarthritis progression, and similar protection was seen with 2-hydroxyglutarate treatment in chondrocytes from patients with osteoarthritis and a mouse model of surgically induced osteoarthritis. Results suggest that inhibition of TET1 activity could be therapeutic for osteoarthritis.

Abstract

Osteoarthritis (OA) is a degenerative disease of the joint, which results in pain, loss of mobility, and, eventually, joint replacement. Currently, no disease-modifying drugs exist, partly because of the multiple levels at which cartilage homeostasis is disrupted. Recent studies have highlighted the importance of epigenetic dysregulation in OA, sparking interest in the epigenetic modulation for this disease. In our previous work, we characterized a fivefold increase in cytosine hydroxymethylation (5hmC), an oxidized derivative of cytosine methylation (5mC) associated with gene activation, accumulating at OA-associated genes. To test the role of 5hmC in OA, here, we used a mouse model of surgically induced OA and found that OA onset was accompanied by a gain of ~40,000 differentially hydroxymethylated sites before the notable histological appearance of disease. We demonstrated that ten-eleven-translocation enzyme 1 (TET1) mediates the 5hmC deposition because 98% of sites enriched for 5hmC in OA were lost in Tet1−/− mice. Loss of TET1-mediated 5hmC protected the Tet1−/− mice from OA development, including degeneration of the cartilage surface and osteophyte formation, by directly preventing the activation of multiple OA pathways. Loss of TET1 in human OA chondrocytes reduced the expression of the matrix metalloproteinases MMP3 and MMP13 and multiple inflammatory cytokines. Intra-articular injections of a dioxygenases inhibitor, 2-hydroxyglutarate, on mice after surgical induction of OA stalled disease progression. Treatment of human OA chondrocytes with the same inhibitor also phenocopied TET1 loss. Collectively, these data demonstrate that TET1-mediated 5hmC deposition regulates multiple OA pathways and can be modulated for therapeutic intervention.

INTRODUCTION

Aging and trauma to articular cartilage are associated with osteoarthritis (OA), a chronic disease of the joints that is increasing in incidence and is a leading cause of disability (1). A clear understanding of OA etiology and progression has remained elusive (2), and no disease-modifying OA drug is available. Various mutations and polymorphisms have been linked to OA; however, these genes identified by genome-wide association studies are not considered causative of OA because the low frequency is not sufficient to represent a risk for the general population (3). Beyond genetic susceptibility, multiple factors including age, sex, metabolic status, obesity, and joint trauma contribute to OA (4). Development of OA is therefore multifactorial and is considered a disease of the whole joint, affecting cartilage, synovium, and bone (5).

Recent evidence has demonstrated epigenetic dysregulation of multiple genes involved in the pathogenesis of OA (615), with many of these genes undergoing DNA demethylation (1619). Besides 5-methylcytosine (5mC), 5-hydroxymethylcytosine (5hmC) has been found to be stably present in adult tissues and aberrantly distributed in various diseases. This distribution is tissue-specific and mediated by the activity of the ten-eleven-translocation (TET) enzymes, TET1, TET2, and TET3, which convert 5mC to 5hmC and its oxidative derivatives formylcytosine (5fC) and carboxylcytosine (5caC) in a Fe(II)-dependent and α-ketoglutarate (α-KG)–dependent manner (20). TET1 is critical for chondroprogenitor differentiation (21), and both TET1 and TET2 are required for the maintenance of bone marrow mesenchymal stem cells and bone homeostasis (22). Our previous findings that 5hmC accumulates on OA-associated genes in human osteoarthritic cartilage (23, 24) highlighted a previously unknown role for 5hmC in OA pathogenesis and motivated us to investigate 5hmC dynamics during OA initiation and progression. Toward this goal, we have used an established mouse model of posttraumatic OA (PTOA), wherein the destabilization of the medial meniscus (DMM) (2527) leads to surgical induction of OA. DMM mimics human OA progression, including changes to gait, size and location of lesions, and formation of osteophytes (2830). Through global 5hmC mapping and RNA sequencing (RNA-seq) analyses, we identified several pathways in the pathogenesis of OA that are regulated by changes in the 5hmC epigenome. The direct TET1 targets that gain 5hmC and are activated during OA onset regulate multiple disease-associated networks, including WNT signaling, expression of metalloproteinases, and mammalian target of rapamycin (mTOR) signaling. Impairment of the 5hmC deposition in Tet1−/− mice protected them against OA development. Last, we demonstrate that modulation of TET activity using a broad small-molecule inhibitor, 2-hydroxyglutarate (2-HG), dampens the OA phenotype in vivo and in human OA chondrocytes, providing an attractive therapeutic strategy.

RESULTS

5hmC increases in joint cartilage after PTOA induction

To identify changes in the 5hmC landscape in OA, we used the DMM model of PTOA, in mice 4 to 6 months of age (Fig. 1, A and B) (25, 26). In this model, surgical intervention causes a destabilization of the knee joint, mimicking many of the known features of human disease and producing more reliable disease initiation than spontaneous age-associated OA (29). Only male mice were used for these studies (31) because sex hormones are known to cause lower disease severity with altered dynamics in female mice. Immunofluorescence on the knee joint showed that DMM mice had twofold (P = 0.01) and fivefold (P = 0.0028) increase in global 5hmC at 6 and 8 weeks after surgery (Fig. 1, C and D, and fig. S1A). These changes are mainly attributable to DMM-induced OA because no increase in global 5hmC was observed in mice when aged in the absence of surgical induction (fig. S1B). To determine the sites of 5hmC accumulation genome-wide, we performed reduced representation hydroxymethylation profiling (RRHP) (32) to assess individual CCGG sites. Globally, we identified 43,267 differentially hydroxymethylated CCGG (dhCCGG) sites 6 weeks after surgery, with 93% of these sites gaining 5hmC during OA induction (Fig. 1E). Upon exploring the distribution of these dhCCGG sites, we noted that they were predominantly gained in gene bodies (46%), where 5hmC accumulation has been associated with gene activation, and in intergenic regions (45%) (Fig. 1F) (23). Only a small fraction of the dhCCGG sites (8%) were found in promoter regions (Fig. 1F), which we broadly defined as ±5000 base pairs from the transcription start site (TSS). It was difficult to discern the precise 5hmC changes in enhancer regions, which can be found near TSS or in intergenic regions, because no datasets have defined enhancers in mouse articular chondrocytes.

Fig. 1 5hmC increases in joint cartilage after PTOA induction.

(A) Schematic of the time course for DMM. Mice were evaluated at 6 and 12 weeks after surgery. (B) Graphic representation of the DMM surgical procedure with DMM indicated by the red arrows. PCL, posterior cruciate ligament; ACL, anterior cruciate ligament. (C) Representative immunostaining images of 5hmC (red) and 4′,6-diamidino-2-phenylindole (DAPI) (blue) of sham and DMM knee cartilage 6 weeks after surgery. Fem, femur condyle; Tibia, tibial plateau. Scale bars, 30 μm. (D) Quantification of 5hmC staining 6 and 8 weeks after surgery and respective shams. Log2 mean fluorescence intensity (MFI) ± SD (n = 3; 6-month-old mice); *P = 0.05 and **P = 0.0028, unpaired samples t test (two-tailed). (E) Percentage of up- and down-regulated hydroxymethylated CCGG sites in Tet1+/+ knee cartilage 6 weeks after surgery compared to sham controls (n = 3 for each group; 6-month-old mice). (F) Overall genomic distribution of the up-regulated dhCCGGs in intragenic (promoter and gene body) and intergenic locations. (G) Overlap between number of genes that gained 5hmC in either the gene body or the promoter regions and transcriptionally up-regulated genes in mouse knee cartilage 6 weeks after DMM induction. (H) Top six pathways identified by analysis of genes that were transcriptionally up-regulated and gained 5hmC in the gene body in mouse cartilage 6 weeks after DMM induction (n = 3; 6-month-old mice). fMLP, N-formyl-Met-Leu-Phe. (I) Fold change in 5hmC at CCGG sites in representative genes whose 5hmC deposition increases in DMM mice compared to sham controls (each dot indicates a different CCGG along the gene) (n = 3; 6-month-old mice). (J) Overlap between the dhCCGGs gained after 6 weeks of mouse DMM induction and DhMRs (differentially hydroxymethylated regions) accumulated in human OA.

To monitor the effects of this 5hmC deposition on gene expression, we performed RNA-seq on cartilage microdissected from sham and DMM mouse joints 6 and 12 weeks after surgery. Eight hundred ten genes were found to be up-regulated 6 weeks after DMM surgery. Forty-six percent of these up-regulated genes showed a gain in 5hmC. Three hundred forty-seven of the activated genes gained 5hmC in gene bodies. On the other hand, only 60 genes gained 5hmC in the promoter regions, and 32 of these genes also gained 5hmC in gene bodies (Fig. 1G). In the core set of 375 activated genes, we observed an enrichment for pathways related to WNT signaling, protein kinase A activity, and inositol metabolism (Fig. 1H). The 347 activated genes that gained 5hmC in gene bodies included several extracellular matrix (ECM) remodeling enzymes (Fig. 1I).

A total of 911 genes were up-regulated 12 weeks after DMM as compared to the sham control, and 106 of these genes were already up-regulated 6 weeks after DMM, revealing the early-stage genes that persist toward the later stages of PTOA. In addition, 43% of the genes up-regulated at 12 weeks in the DMM joints had gained 5hmC at 6 weeks (fig. S1C), suggesting that 5hmC also marks genes that are important in later stages of OA. This is consistent with our studies of human OA cartilage (23, 24). Upon cross-referencing the datasets, we found that 52% of the genes that gain 5hmC in the DMM model accumulated 5hmC in the end-stage human OA chondrocytes as well (Fig. 1J). Our findings in both mouse and human samples therefore corroborate that 5hmC gain is a hallmark of OA pathogenesis.

TET1 mediates OA-related 5hmC deposition

Oxidation of 5mC to 5hmC, 5fC, or 5caC can be catalyzed by any of the three members of the TET family (20). The short form of TET1 (TET1s) is expressed in somatic tissues and has been shown to participate in targeted binding and oxidation events (3335). Unlike TET2 and TET3, TET1’s catalytic domain is not processive; rather, it releases 5hmC intermediates (36), putatively explaining how stable 5hmC can build up in many genomes (37), including that of differentiating chondrocytes (21). Therefore, conceptually, TET1 is the ideal candidate responsible for 5hmC deposition during OA induction despite being the lowest expressed of the three TET enzymes in the knee cartilage (fig. S2A). Immunohistochemistry of 4-month-old mouse joints confirmed the expression of TET1 in a subpopulation of cells in joint cartilage (fig. S2B), consistent with human data (24, 38).

Recent work has posited that OA is a disease of the whole joint, with multiple tissues involved, including bone and synovium. Therefore, to test whether TET1 was necessary for the 5hmC deposition during OA development, we performed DMM on a Tet1−/− mouse model that lacked TET1 in the whole joint (39). Unexpectedly, Tet1−/− mice showed no deterioration of the cartilage architecture 6 weeks after surgery (Fig. 2A). Grading of the DMM joints using Osteoarthritis Research Society International (OARSI) scoring (27) exhibited statistically significant lower (P < 0.0001) maximum and summit scores and decreased cartilage loss in the Tet1−/− mice compared to the Tet1+/+ mice (Fig. 2, B and C). Accordingly, Tet1−/− mice had lower matrix metalloproteinase-13 (MMP-13) expression in articular chondrocytes of the joint compared to Tet1+/+ control mice (Fig. 2, D and E). Tet1−/− DMM mice lost 5hmC at 65% of CCGG sites compared to Tet1+/+ DMM mice (Fig. 2F) and failed to gain 5hmC at 98% of the dhCCGGs that showed increased 5hmC in Tet1+/+ DMM mice (Fig. 2G). Collectively, these data demonstrate that TET1 is the major TET protein responsible for the 5hmC gain upon DMM. We did not observe any significant change (P > 0.05) in Tet1 expression upon DMM induction in the RNA-seq data at 6 or 12 weeks (fig. S2C), suggesting that the increase in 5hmC deposition was due to either increased activity or increased targeting.

Fig. 2 TET1 mediates OA-related 5hmC deposition.

(A) Representative images of Safranin O (SafO)/fast green staining of Tet1+/+ and Tet1−/− knee joints 6 weeks after DMM induction. Black arrows indicate sites of cartilage loss in Tet1+/+ samples (6-month-old mice). Scale bars, 100 μm. Boxed regions are shown at higher magnification below. (B) Maximum and summit scores of the knee joint in Tet1+/+ and Tet1−/− mice based on the OARSI scoring system. Means ± SD (n = 7; 6-month-old mice); ***P < 0.0001, one-way ANOVA with Tukey’s multiple comparisons test. (C) Percentage loss of cartilage in Tet1+/+ and Tet1−/− mice. Mean staining intensity ± SD (n = 7; 6-month-old mice); *P = 0.015, unpaired samples t test (two-tailed). (D) Representative images of MMP-13 immunostaining (brown) of Tet1+/+ and Tet1−/− knee joints 6 weeks after DMM induction. Boxed regions are shown at higher magnification below. Scale bars, 100 μm (top) and 30 μm (bottom). (E) Quantification of MMP-13 immunostaining in Tet1+/+ and Tet1−/− knee joints. Mean staining intensity ± SD (n = 3; 6-month-old mice); *P = 0.0378, unpaired samples t test (two-tailed). (F) Percentage of CCGG sites that are up- or down-regulated in Tet1−/− DMM knee joints compared to Tet1+/+ DMM. (G) Percentage of CCGG sites lost or maintained in Tet1−/− DMM cartilage among the 93% CCGGs gained after Tet1+/+ DMM induction. (H) Three-dimensional (3D) reconstructions of phase-contrast micro-CT imaging data in Tet1+/+ and Tet1−/− sham and 12-week-old DMM knee joints. Z1, Z2, Z3, and Z4 arrowheads and panels indicate frontal view for sites of osteophyte formation in the Tet1+/+, which are missing in the Tet1−/−; Z1′ and Z3′ arrowheads and panels indicate lateral view of Z1 and Z3. (I) Representative 2D micro-CT images showing osteophytes in red arrows and zoomed red boxes.

We next wanted to determine whether the protection from OA in the Tet1−/− DMM mice would persist into later disease stages. Upon assessing joints from mice 12 weeks after surgery, which mimic late-stage human disease (29, 30), we observed that the Tet1−/− DMM mice maintained intact cartilage and proteoglycan staining (fig. S2D), had limited loss of cartilage thickness (fig. S2E), and reduced MMP-13 production and secretion (fig. S2F). In addition, micro–computed tomography (micro-CT) analyses of the DMM joints showed the formation of osteophytes and ectopic bone in the Tet1+/+ but not Tet1−/−, DMM mice (Fig. 2, H and I). To complement these data, we also studied the joints in uninjured, 18-month-old Tet1+/+ and Tet1−/− mice to assess any spontaneous development of OA. A variable onset of OA was observed in wild-type mice, and although not statistically significant (P > 0.05), the Tet1−/− mice trended toward being protected from OA (fig. S2, G and H). Together, these data show that TET1 is the hydroxymethylase predominantly responsible for the accumulation of pathogenic 5hmC in OA and that its loss impairs both OA initiation and progression.

TET1-mediated 5hmC controls activation of key OA pathways including WNT, metalloproteinases, and signal transducer and activator of transcription 3 signaling

We next sought to understand how the TET1-mediated 5hmC landscape in chondrocytes drives the molecular events of OA. RNA-seq analyses of the microdissected cartilage of DMM mice at 6 weeks revealed that multiple OA-related pathways, including retinoic acid receptor (RAR), WNT, and interleukin-1 (IL-1) signaling networks, failed to activate in Tet1−/− DMM mice compared to Tet1+/+ DMM mice (Fig. 3, A to C). To determine all ECM regulators involved, we overlapped the differentially expressed gene lists of Tet1+/+ and Tet1−/− with the murine matrisome database (40). We observed 16 significant (P < 0.05) gene expression changes, including metalloproteinases Adams and Adamts (Fig. 3D). Comparison of the 5hmC maps between the Tet1+/+ and Tet1−/− DMM mice showed a loss of 5hmC at OA-associated genes involved in key pathways like HIF-1A (hypoxia-inducible factor 1A), JAK (Janus kinase)/STAT (signal transducer and activator of transcription), and NF-κB (nuclear factor κB) (Fig. 3, E and F).

Fig. 3 TET1-mediated 5hmC controls activation of many key OA pathways including WNT, metalloproteinases, and STAT3 signaling.

(A) Heat maps with hierarchical clustering of differentially expressed genes in Tet1+/+ and Tet1−/− knee cartilage 6 weeks after DMM induction (n = 3; 6-month-old mice). (B) Top seven pathways identified by pathway analysis of genes that are differentially expressed in Tet1−/− knee cartilage compared to Tet1+/+. BMP, bone morphogenetic protein. (C) Changes in expression of OA-associated genes in Tet1+/+ and Tet1−/− cartilage 6 weeks after surgery. Log2 transcripts per million (TPM) ± SD (n = 3; 6-month-old mice). (D) Changes in expression of ECM regulators in Tet1+/+ and Tet1−/− knee cartilage 6 weeks after surgery compared to their respective sham. Log2(TPM) fold change ± SD (n = 3; 6-month-old mice). All represented values are significant; *P < 0.05, multiple t test with Bonferroni-Dunn correction (α = 0.049). (E) Top six pathways identified by canonical pathway analysis of dhCCGGs in DMM Tet1−/− versus DMM Tet1+/+ controls. JNK, c-Jun N-terminal kinase; GPCRs, G protein–coupled receptors; IFN, interferon. (F) Fold change in 5hmC at CCGG sites in DMM Tet1−/− versus DMM Tet1+/+ control cartilage for genes involved in pathways reported in (E). Multiple dhCCGG sites are represented for each gene (each dot indicates a different CCGG along the gene). (G) Overlap between number of genes that lost 5hmC and transcriptionally down-regulated genes in Tet1−/− versus Tet1+/+ DMM-induced cartilage (n = 3 for each group). (H) STRING analysis of protein interaction networks identified from the 313 “direct target” genes in (G) that lost both 5hmC accumulation and gene expression. GnRH, gonadotropin-releasing hormone; AMPK, adenosine monophosphate–activated protein kinase. (I) Changes in expression of Mapk1 and Mapk14 in Tet1+/+ and Tet1−/− knee cartilage 6 weeks after surgery. Mean TPM ± SD (n = 3; 6-month-old mice); *P = 0.02 for Mapk1 and *P = 0.01 for Mapk14, unpaired samples t test (two-tailed). (J) Enrichr analysis to identify TFs controlling the 313 genes resulting from the overlap in (G). (K) Schematic representation of pathways and genes that are modulated in Tet1−/− cartilage and may contribute to the protection from OA pathogenesis. SASP, senescence-associated secretory phenotype.

To determine which of these genes were direct targets of TET1, we overlapped genes losing both 5hmC and gene expression in the Tet1−/− DMM joints (Fig. 3G), resulting in 313 targets. Of these, 59% gained 5hmC during Tet1+/+ OA induction. Upon assembling a protein interaction network using STRING (41), these direct TET1 target genes were found to be responsible for controlling several important nodes in the homeostasis of cartilage, including mTOR signaling, WNT signaling, apoptosis, and chemokine signaling (Fig. 3H). Some members of the mitogen-activated protein kinase (MAPK) family, including Mapk1 (extracellular signal–regulated kinase 2) and Mapk14 (p38), were found to be down-regulated in Tet1−/− DMM cartilage (Fig. 3I). In addition, several other kinases, including Oxsr1, which activates pathways related to oxidative stress, tumor necrosis factor (TNF) signaling, and apoptosis, were direct target genes (table S1).

Among the 313 direct target genes, several were transcription factors (TFs) whose down-regulation might explain the TET1-dependent gene expression changes that are independent of 5hmC. These included Cxxc5, Gata2, Klf3, Kl9, Tcf3, and Yy1 (table S1). Among these, YY1 has been associated with rheumatoid arthritis and is known to up-regulate the expression of IL-6 and inhibit the expression of IL-8 (42, 43). In addition, it acts as an activator for Gdf5 (44) and inhibitor of miR-10a, whose inhibition leads to increased NF-κB activation (45). Several chromatin regulators were also down-regulated, including Setd8, Kdm2b, and Prdm16 (table S1).

To determine which TFs might be sensitive to TET1-mediated 5hmC deposition, we used Enrichr (46) to analyze expression data from knockout studies of known TFs. In the top 30 enriched sets, we found TFs with known roles in OA, such as Creb1, Gata1, Stat3, and Yap1 (Fig. 3J), that can act upstream of the 313 direct TET1 targets. Previous work on the interaction between TET1 and STAT3 suggests a model in which TET1-mediated 5hmC deposition allows for TF binding at its target sites and subsequent target activation (47). These TFs may also represent the factors that facilitate the binding of TET1 to their target genes during OA initiation.

TET1 is also able to interact with the polycomb repressive complex 2 and act as a repressor (48). To determine whether the loss of this repressor activity led to the up-regulation of any chondroprotective genes in the Tet1−/− mice in a 5hmC-independent manner, we focused on targets that were up-regulated in the Tet1−/− sham compared to Tet1+/+ control sham mice. Pathway analyses on the 263 up-regulated genes showed an enrichment for negative regulation of Toll-like receptor 4 signaling and protein catabolic processes. Sham Tet1−/− mice also had a 20-fold up-regulation of Dot1l (P = 0.0004), which was previously shown to be chondroprotective in OA by repressing sirtuin 1 (SIRT1)–induced expression of the WNT pathway (49). DNA methyltransferase 3b (Dnmt3b) was also 20-fold up-regulated (P = 0.0004), whose gain of function has been shown to be chondroprotective through altering metabolism and mTOR signaling (9), further underscoring the broad effects of TET1 on multiple aspects of cartilage regulation. The genes up-regulated in Tet1−/− sham mice were not enriched for any ECM components, as tested using a curated matrisome list (40). After DMM induction, however, we observed that Tet1−/− mice up-regulated pathways related to N-glycan biosynthesis (Dolpp1) and sialic acid biosynthesis that are involved in the modification of ECM proteins, along with chondroprotective genes like Smad4, Pparg, and Timp1 (Fig. 3K). Therefore, although the Tet1−/− mice do not up-regulate ECM genes directly, they might preserve cartilage integrity by decreasing metalloproteinase activity and through glycan metabolism.

Down-regulation of inflammatory pathways, such as PPARγ (peroxisome proliferator–activated receptor γ) and IL-12 signaling, was maintained in the 12-week-old Tet1−/− DMM mice (fig. S3, A and B). Other target genes related to ECM degradation, cell cycle, and cytoskeletal rearrangement (Adamtsl4, Nek1, and Wipf1) were also persistently down-regulated in the Tet1−/− joints 12 weeks after DMM induction. Together, these data collectively highlight the multiple TET1-regulated pathways (Fig. 3K), demonstrating that TET1 is a key epigenetic regulator of OA.

Loss of Tet1 protects cartilage from OA without impairing the systemic response to inflammation

Our analyses of 5hmC deposition and gene expression provided an overview of the cartilage intrinsic mechanisms of protection in the Tet1−/− mice. Next, we asked whether Tet1 loss predominantly affected cartilage or also caused a systemic dampening of low-grade inflammation (50). We first assessed synovial inflammation by (i) the extent of immune filtration in the synovium and (ii) the amount of inflammatory cytokines in the serum of the DMM mice. DMM joints were immunostained for F4/80, a marker of monocyte and macrophage immune cell lineages, 6 weeks after surgery. We noted that immune cell infiltration in the synovial lining was not affected because we observed twofold more infiltration of F4/80-positive immune cells in the Tet1−/− joints than in the Tet1+/+ DMM joints (Fig. 4, A and B). We also observed increased expression of Cxcr4 and Lcp2, two markers of B cells, in Tet1−/− joints in the RNA-seq dataset. Thus, we ruled out a potentially lowered immune infiltration as the mechanism of protection from OA in the in Tet1−/− mice.

Fig. 4 Loss of Tet1 protects cartilage from OA without impairing the systemic response to inflammation.

(A) Representative images of F4/80 immunostaining (brown) of Tet1+/+ and Tet1−/− synovial membrane 6 weeks after DMM induction. Scale bars, 30 μm. (B) Severity of synovitis score for Tet1+/+ and Tet1−/−. Mean scores ± SD (n = 3; 6-month-old mice); *P < 0.05, unpaired samples t test (two-tailed) with Welch’s correction. (C) Heat map of a 39-plex mouse inflammatory cytokine panel on Tet1+/+ and Tet1−/− serum 6 weeks after surgery. Significant changes in LEPTIN, CCL7, and EOTAXIN are plotted on the right. MFI ± SD (n = 3 for Tet1+/+ and n = 4 for Tet1−/−; 6-month-old mice); *P < 0.05, ***P < 0.001, and ****P < 0.0001, one-way ANOVA with Tukey’s multiple comparisons test. (D) Heat map of a 39-plex mouse inflammatory cytokine panel on Tet1+/+ and Tet1−/− serum 4 hours after systemic LPS administration and in unstimulated controls (n = 3 for each group; 6-month-old mice). (E) Schematic of the time course for the in vivo injection of IL-1β in Tet1+/+ and Tet1−/− knee joints. Collection of tissues for analysis was performed at day 6. (F) Representative images of Safranin O/fast green staining of Tet1+/+ and Tet1−/− knee joints after three IL-1β intra-articular injections (n = 3 for each group; 6-month-old mice). Black boxes indicate area of higher magnification. Scale bars, 100 μm. (G) Representative images of MMP-13 immunostaining (brown) of Tet1+/+ and Tet1−/− knee joints after three IL-1β or control saline intra-articular injections. Brown arrowheads indicate MMP-13–positive signal. Scale bars, 100 μm. (H) Percentage loss of Safranin O staining in Tet1+/+ and Tet1−/− mice after IL-1β injection compared to saline injection. Mean staining intensity ± SD (n = 3; 6-month-old mice); *P = 0.03, unpaired samples t test (two-tailed). (I) Quantification of MMP-13 immunostaining in (G). Mean staining intensity ± SD (n = 3; 6-month-old mice); *P = 0.04, unpaired samples t test (two-tailed).

To assess the low-grade inflammation typically observed in osteoarthritic mice, we characterized serum from mice 6 and 12 weeks after surgery using a 39-plex Luminex cytokine panel. In Tet1+/+ DMM mice, we detected an up-regulation of LEPTIN, CCL7 (C-C motif chemokine ligand 7), and EOTAXIN over the course of disease progression. When compared to their own baseline, Tet1−/− mice also had increased amount of LEPTIN (at 6 and 12 weeks), CCL7 (at 12 weeks), and EOTAXIN (at 6 and 12 weeks); however, the absolute quantity was lower than Tet1+/+ controls (Fig. 4C). These data ruled out a potentially heightened inflammatory microenvironment due to increased infiltration of F4/80+ immune cells in the synovium. However, the decrease in OA-associated cytokines LEPTIN, CCL7, and EOTAXIN (5153) suggested that loss of TET1 partially modulated inflammation in the joint microenvironment. To exclude any systemic immune defects in the Tet1−/− mice, as observed in the Tet2−/− mice (54), we challenged mice with an acute infection by intraperitoneal lipopolysaccharide (LPS) injection. Both Tet1−/− and Tet1+/+ mice mounted a comparable, robust response against LPS (Fig. 4D and fig. S4A). In particular, CCL7 and EOTAXIN were up-regulated to a similar extent in both Tet1−/− and Tet1+/+ mice.

Last, we challenged Tet1−/− and Tet1+/+ mice with IL-1β injections to test the cartilage response to direct cytokine stimulation in the joint. Mouse knee joints were injected every other day with IL-1β over a course of 6 days (Fig. 4E). As previously described (55, 56), wild-type animals exhibited loss of proteoglycan content in cartilage after injections as observed by Safranin O staining (Fig. 4, F and H). In contrast, proteoglycan staining in Tet1−/− cartilage was still high (Fig. 4, F and H). In line with these findings, MMP-13 secretion, a target of IL-1β (57), was ~25% lower in Tet1−/− joints compared to wild type (Fig. 4, G and I). These data were corroborated by in vitro studies using differentiated ATDC5 cells, a mouse chondrogenic cell line (58). We observed that ~75% Tet1 knockdown through short hairpin RNA (shRNA) inhibited the up-regulation of Mmp3, Mmp13, and IL-6 in response to IL-1β stimulation (fig. S4B). Collectively, these data demonstrate that Tet1−/− cartilage is intrinsically protected from the effects of the proinflammatory cytokine milieu in the joint.

As a corollary, we wanted to test whether TET1 was sufficient to activate cytokine-induced gene expression changes in healthy chondrocytes. For this, we used chondrocytes isolated from three normal human donors with no history of OA. We found no difference in the amount of MMP3 or CCL2 in Tet1-overexpressing chondrocytes (fig. S5). Likewise, both control and Tet1-overexpressing cells had the same magnitude of MMP3 and CCL2 up-regulation after treatment with IL-1β or with a combination of recombinant TNFα and oncostatin M (fig. S5). This suggests that although Tet1 is necessary for the expression of these OA-associated genes, it is not sufficient.

Pharmacological inhibition of TET activity prevents OA

Next, we wanted to directly test the therapeutic potential of TET1 inhibition in OA. As a proof-of-principle experiment, we performed shRNA-mediated Tet1 knockdown on three samples from patients with OA, obtaining a 40 to 70% knockdown and a subsequent 80 to 90% reduction in 5hmC (fig. S6A and Fig. 5A). Consistent with the mouse data, we found that the silencing of TET1 decreased MMP3 and MMP13 expression (fig. S6B). To assess any low-grade inflammation in the OA cells, we used a cytokine array and observed that Tet1 knockdown significantly (P < 0.001) decreased concentration of IL-6, IL-8, vascular endothelial growth factor (VEGF), vascular cell adhesion molecule 1 (VCAM1), plasminogen activator inhibitor 1 (PAI1), and CCL2 (Fig. 5, B and C). The plurality of pathways altered by the loss of Tet1 validated our strategy in targeting an upstream epigenetic regulator rather than perturbing each pathway separately.

Fig. 5 Inhibition of TET activity prevents the OA phenotype.

(A) Fold change in 5hmC quantified by antibody-based enzyme-linked immunosorbent assay (ELISA) for three human OA chondrocyte samples after nontarget (NT) or TET1 (T1-Sh) knockdown. Mean absorbance ± SD (n = 3 patients with OA and n = 2 shRNA for each patient); ***P < 0.001, unpaired samples t test (two-tailed). (B) Sankey diagram of a 62-plex cytokine panel in OA chondrocytes after nontarget or TET1 knockdown. Cytokines are grouped by detection quantity (n = 3 patients with OA and n = 2 ShRNA for each patient). (C) Log2(MFI) of representative cytokines that change upon TET1 knockdown. Log2(MFI) ± SD (n = 3 patients with OA and n = 2 shRNA for each patient); ***P < 0.001, multiple t test comparison (two-tailed). (D) Fold change in 5hmC quantified by antibody-based ELISA for human OA chondrocyte samples upon inhibition of TETs with 2-hydroxyglutarate (2-HG). Mean absorbance ± SD (n = 3 patients with OA, each treated twice). (E) Sankey diagram of a 68-plex cytokine panel before and after TET inhibition with 2-HG treatment (n = 5 patients with OA). Cytokines are grouped by detection quantity. (F) Log2(MFI) of representative cytokines that change upon TET inhibition. Log2(MFI) ± SD (n = 5 patients with OA); ***P < 0.001, multiple t test comparison (two-tailed). (G) Schematic of the time course for the in vivo injection of 2-HG inhibitor in surgically induced knee joints of Tet1+/+ and Tet1−/− mice. Collection of tissues for further analysis was performed 6 weeks after DMM induction. (H) Representative images of Safranin O/fast green staining of Tet1+/+ knee joints injected with 2-HG inhibitor (100 mM) or saline controls and analyzed 6 weeks after DMM induction. Scale bars, 100 μm. (I) Summit and maximum scores based on the OARSI scoring system of Tet1+/+ knee joints injected with 2-HG inhibitor (100 mM) or saline controls and analyzed 6 weeks after DMM induction. Mean scores ± SD (n = 3 controls and n = 5 inhibitors; 6-month-old mice); *P = 0.01 for summit score and *P = 0.03 for maximum score, unpaired samples t test (two-tailed). (J and K) Representative images of MMP-13 (J) and 5hmC (K) immunostaining (brown) of Tet1+/+ DMM knee cartilage upon saline control or inhibitor injection. Scale bars, 100 μm. Boxed regions shown at higher magnification. (L) Quantification of 5hmC immunostaining in (K). Mean staining intensity ± SD (n = 3); **P = 0.001, unpaired samples t test (two-tailed). (M) Fold expression change in Oxsr1, Igfbp7, and Adamtsl4 in Tet1+/+ cartilage injected with 2-HG inhibitor (100 mM) or saline controls and analyzed 6 weeks after DMM induction. Mean DDCT ± SD (n = 3); **P = 0.001 (n = 3 for each group), unpaired samples t test (two-tailed) with Welch’s correction.

Next, we tested the feasibility of modulating TET1 activity pharmacologically using a known TET inhibitor, 2-HG (59). Treatment of chondrocytes from three patients with OA with 2-HG resulted in a 25% decrease in 5hmC (Fig. 5D). Although 2-HG is a broad inhibitor affecting other demethylases besides TET (60), we observed effects similar to Tet1 knockdown on MMP3 and MMP13 expression and proinflammatory cytokine secretion (Fig. 5, E and F, and fig. S6C) despite some expected variability between patients.

Last, we asked whether early intervention with the 2-HG inhibitor could prevent the onset of OA in vivo. Tet1+/+ control mice were injected with 2-HG twice a week for 5 weeks after DMM surgery (Fig. 5G). Periodic administration of the inhibitor in the joints was sufficient to reduce cartilage degradation (Fig. 5, H and I), along with decreasing the secretion of MMP-13 (Fig. 5J) and reducing 5hmC (Fig. 5, K and L), akin to the genetic loss of TET1. This demonstrates that 2-HG is active within the joint environment and can alter the pathological expression of 5hmC observed during OA onset. Furthermore, we tested the expression of three direct TET1 targets (Oxsr1, Igfbp7, and Adamtsl4) identified in Tet1−/− mice in inhibitor-treated joints by quantitative polymerase chain reaction and found their expression down-regulated (Fig. 5M). Collectively, these data show that pharmacological inhibition of TET mirrors the genetic loss of TET1, further confirming our hypothesis that TET inhibition would be therapeutic for OA.

DISCUSSION

Previous work from our laboratory established the accumulation of 5hmC in OA cartilage and its association with genes linked to OA pathogenesis (23, 24). To study the dynamics of 5hmC and the TET enzymes in OA, we used a mouse model of PTOA. After surgical destabilization of the knee joint, we observed a global increase in the amount of 5hmC in joint cartilage. In agreement with our previous findings in human OA, 5hmC is predominantly gained in gene bodies and intergenic regions, with 46% of these genes becoming activated during OA induction. This indicates a functional role for 5hmC in the modulation of gene expression during OA pathogenesis. Corroborating this idea, the gain of 5hmC was associated specifically with the induction of PTOA in mice because uninjured mice aged up to 1 year did not show any increase in 5hmC in joint cartilage. In contrast to gene bodies, only 60 genes gained 5hmC in the promoter regions. This predominance of gene body 5hmC gain is consistent with data from other tissues, where it has been correlated with gene activation (6163). Chromatin immunoprecipitation sequencing in multiple cell types has demonstrated that although TET1 binding is correlated to where 5hmC is gained, TET1 is not always found bound at the sites of 5hmC gain (63). Therefore, TET1 binding upon injury in the DMM model is not expected to correspond completely with the 5hmC gain because a transient binding and release of TET1 could be sufficient for 5hmC deposition.

Despite the smaller size of the mutant mice observed in the early postnatal life (39), adult Tet1−/− joint cartilage appears phenotypically normal and comparable to wild-type littermates. A similar observation has also been recently described for the bone tissue (22). Upon DMM induction, Tet1−/− mice did not develop the classical signs of OA, including cartilage degradation, and failed to accumulate ~98% of the dhCCGGs gained in the wild-type DMM. Protection from OA was maintained 12 weeks after DMM induction in the Tet1−/− mice. These data revealed TET1 to be the major family member responsible for the 5hmC gain in OA.

Loss of TET1-regulated genes perturbed multiple pathways known to be crucial in OA pathogenesis, including Wnt, MAPK, mTOR, and chemokine signaling. Moreover, analyses of the TFs upstream of the down-regulated genes identified GATA1 (GATA binding protein 1), TCF3 (transcription factor 3), and STAT3, demonstrating that TET1 activity controls several key nodes in cartilage homeostasis and OA progression (fig. S7). WNT signaling and reactive oxygen species (ROS)/oxidative stress pathways were identified for aberrant 5hmC gain in our studies of end-stage human OA chondrocytes as well (23). Another notable change observed was an alteration in ECM dynamics in the Tet1−/− cartilage as compared to that of the wild-type mice, showing suppression of several serine proteases including Mmp and Adam families of genes (fig. S7).

Another major effect of TET1 loss was that inflammatory pathways were down-regulated in the transcriptome of Tet1−/− cartilage. Because this effect could be cartilage-specific or due to an overall dampening of inflammation, we investigated whether TET1 controlled inflammation at a systemic level. However, neither synovial tissue infiltration by F4/80-positive monocytes after DMM nor the serum proinflammatory cytokines released upon LPS stimulation were adversely affected by the loss of TET1. This confirmed that, unlike the Tet2−/− mice (54), there was no broad immune impairment in the Tet1−/− mice. It is noted, however, that in these experiments, we tested only a small immune population (F4/80-positive cells). Similarly, the LPS stimulation assessed innate immune function but did not rule out any potential changes in the adaptive immune function upon TET1 loss. The overall amount of LEPTIN, CCL7, and EOTAXIN, which are associated with increased severity of OA, was lower in the serum of Tet1−/− mice compared to the Tet1+/+ mice after DMM. To independently test the effect of a cartilage-specific loss, TET1 was knocked down in chondrocytes differentiated from a mouse chondrogenic cell line, resulting in an impaired response to IL-1β and greatly dampened up-regulation of Mmp3, Mmp13, and Il-6. Similarly, shRNA-mediated knockdown of Tet1 in late-stage human OA chondrocytes was sufficient for the suppression of inflammatory response. Together, these data support that loss of TET1 caused a cartilage-specific failure to respond to inflammation and modulation of the overall inflammatory microenvironment, likely because of muted cross-talk between the cartilage and the inflamed synovium (5). The observation that there was an about twofold increase in infiltrated F4/80+ immune cells in the Tet1−/− mouse after DMM could be a result of a delayed resolution of inflammation in the synovium due to this impaired cross-talk. Because a global TET1 knockout mouse was used, it is possible that the other TET1-deficient cells in the joint also contribute to the overall dampening of inflammation. Future studies should investigate the cartilage-specific loss of Tet1 as well as the effect of Tet2 and Tet3 inhibition because these two members are more highly expressed than Tet1 in the joint cartilage.

An independent study found that cartilage-specific loss of Dnmt3b resulted in changes to the gene expression of pathways involved in inflammation, chondrogenesis, and energy metabolism and accelerated OA in mice (9). Although 5mC or 5hmC patterns were not interrogated during OA progression in this study, it has been shown in other tissues that TET and DNMT proteins are responsible for maintaining the 5mC-5hmC homeostasis. Complementary and competitive binding of TET1 and DNMT3A has been demonstrated in embryonic stem cells wherein TET1 limits the binding of DNMT3A and the subsequent DNA methylation (64). Loss of TET1 in this scenario would therefore be similar to DNMT3A gain of function. This model can nicely explain the results of OA induction in the genetic models of TET1 loss and DNMT3B gain because both led to protection from OA. Furthermore, we observe that in our sham baseline Tet1−/− mice, Dnmt3b and Dot1l, which also have a known chondroprotective effect, are up-regulated. Future studies will unravel the cross-talk between TET1, DNMT3B, and DOT1L because they are all important epigenetic regulators of OA.

Our mouse and human data suggested that an intervention based on inhibition of TET1 activity, with its involvement in multiple nodes of OA pathways, would make for a stronger and more efficacious therapy than previous approaches targeting single pathways. To test this hypothesis, we pharmacologically inhibited TET activity using 2-HG, which inhibits multiple α-KG–dependent dioxygenases including the TET family (59, 60). Intra-articular injections of 2-HG in animals after DMM surgery prevented both 5hmC deposition and cartilage degeneration, recapitulating the knockout mouse phenotype. Furthermore, the use of the same inhibitor treatment dampened a broad range of inflammatory cytokines and decreased 5hmC quantity in OA patient-derived chondrocytes, akin to TET1 knockdown. These data provide a proof of principle that modulating TET activity can be an effective early-stage intervention for OA through the modulation of the 5hmC epigenome and consequent changes in gene transcription. Future experiments using the mouse PTOA model can test the time course of TET1 inhibition and determine whether a late-stage intervention, once the disease is established, will also be effective. Recently, a TET1-specific peptide inhibitor with an IC50 (half maximal inhibitory concentration) of 1.1 μM has been developed (65), further opening the possibility of translating these benefits into the clinic. Future studies should test these TET1-specific inhibitors on preclinical large animal models that more accurately represent the load-bearing human joints.

MATERIALS AND METHODS

Study design

The primary objective of this research study was to unravel the role of TET1-mediated 5hmC deposition in the pathogenesis of OA. The study is based on controlled experimental analyses of Tet1 knockout mice and wild-type controls, along with TET1 loss induced via either shRNA or 2-HG inhibitor treatment in human chondrocytes from patients with OA. A surgical model was primarily used to induce OA in the knee joints of Tet1−/− and Tet1+/+ control littermates. Several measurements were used to assess the phenotype of mouse cartilage tissue and human chondrocytes including bioinformatic analyses of RNA-seq and CCGG sequencing, histological evaluation, and quantitative bone assessment by micro-CT imaging and multiplex cytokine analyses in the serum of mice in vivo and in the supernatant of human chondrocyte cultures. To confirm the protection of Tet1−/− cartilage to tissue damage, we used an additional in vivo injury method via intra-articular injection of recombinant IL-1β. Cartilage histology was performed by Safranin O staining and immunohistochemistry or fluorescence staining. All experiments were performed thrice, and the experimental groups were randomly assigned. Blinded methods were used for both study design and outcome assessment. In particular, histological evaluation of the DMM model was performed in a blinded way by three independent investigators, and human chondrocyte experiments were blindly executed by two independent investigators. We estimated the sample size on the basis of known variability of assays, and no outliers were excluded. Sample size for the DMM mice cohort was established by prospective power analysis before the study initiation, and no outliers were excluded from the study at any point. Primary data are reported in data file S1.

Surgically induced mouse OA model

All animal procedures were approved by the Stanford University Administrative Panel on Laboratory Animal Care. Tet1−/− mice (Tet1tm1.1Jae, the Jackson laboratory) were generated from Tet1+/− heterozygous mice. DMM was performed on male mice 4 to 6 months old. After anesthesia with 3% isoflurane, hindlimbs were shaved, and the knee joint was exposed after a medial capsular incision. Medial meniscotibial ligament was transected with microscissors. The joint capsule was closed with a 7-0 taper Vicryl suture, and the skin incision was closed with a 5-0 taper Vicryl. On weeks 6, 8, or 12, mice were euthanized, and the joints were collected for histological assessment through blinded grading with the OARSI scoring system (27). Briefly, we stained sections of mouse joints with Safranin O, and we evaluated cartilage degeneration by assigning numbers 0 to 4 for increasing depth of cartilage degeneration. For the summit scores, six regions were summed.

Reduced representation hydroxymethylation sequencing

RRHP libraries were made using the RRHP 5-hmC Library Prep Kit from Zymo Research (D5450). One microgram of DNA from the mouse joint was used as input. Libraries were made according to the manufacturer’s specifications. Library size selection was done using Agencourt AMPure Beads (A63881) at a 1.8× ratio. Libraries were indexed using NEBNext Multiplex Oligos Ultra index primers (set 1; E7600S) and amplified for 15 cycles. Individual libraries were analyzed on the Bioanalyzer and pooled to a final concentration of 8 to 12 pM. A negative control library was generated by including a sample not treated with β-glucosyltransferase. Pooled libraries were sequenced on the Illumina HiSeq 4000 as paired-end 101–base pair reads. On average, 40 million paired reads were obtained per sample.

Multiplex autoantibody assay

This assay was performed in the Human Immune Monitoring Center at Stanford University. Human 62-plex or mouse 39-plex kits (eBioscience/Affymetrix) were used according to the manufacturer’s recommendations. Briefly, beads were added to a 96-well plate and washed in a BioTek ELx405 washer. Samples were added to the plate containing the mixed antibody-linked beads and incubated at room temperature for 1 hour, followed by overnight incubation at 4°C with shaking at 500 to 600 rpm. Plates were then washed in a BioTek ELx405 washer, and biotinylated detection antibody was added for 75 min at room temperature with shaking. Plate was washed, and streptavidin-phycoerythrin was added. After 30-min incubation and washes, reading buffer was added to the wells. Plates were read using a Luminex 200 instrument with a lower bound of 50 beads per sample per cytokine. Custom assay control beads by Radix Biosolutions are added to all wells.

TET1 inhibitor treatment

Inhibition of TET1 activity was obtained by treating human OA chondrocytes with 10 mM R-ɑ–2-HG (L-2-HG; 907990, Sigma Aldrich) for 8 to 10 hours. In vivo inhibition was performed by intra-articular injection of 2-HG (100 mM diluted in sterile H2O in 5-μl volume) twice a week for 5 weeks starting at 7 days after surgical DMM.

Statistical analysis

Planned comparisons were performed with the GraphPad software. We used (i) one-way analysis of variance (ANOVA), followed by Tukey’s post hoc test to identify specific differences between genotypes, drug treatment groups, selected patients with OA group, or across the time point tested, and (ii) nonparametric, two-tailed Welch’s t test for comparisons between only two groups. P values were corrected for multiple-hypothesis testing, such that the familywise error was capped at 0.05, using the Bonferroni correction method. The exact method and specific P values for significant comparisons are stated in the appropriate Results section.

SUPPLEMENTARY MATERIALS

stm.sciencemag.org/cgi/content/full/12/539/eaax2332/DC1

Materials and Methods

Fig. S1. TET1-mediated 5hmC deposition does not increase during aging.

Fig. S2. Protection from OA in Tet1−/− mice persists in later stages of the disease and is also observed in spontaneous age-induced OA.

Fig. S3. Down-regulation of inflammatory pathways is maintained in Tet1−/− mice 12 weeks after DMM.

Fig. S4. Inflammatory response of chondrocytes to IL-1β is dampened in the absence of Tet1, whereas systemic response to LPS is not altered.

Fig. S5. TET1 is necessary but not sufficient for the expression of genes associated with OA.

Fig. S6. Both TET1 knockdown and pharmacological inhibition reduced the expression of OA markers.

Fig. S7. Loss of TET1 protects the cartilage from OA development at multiple stages.

Table S1. Direct TET1 targets in DMM.

Data file S1. Primary data.

REFERENCES AND NOTES

Acknowledgments: We would like to thank G. Yang and Z. Wang for helpful setup of the animal model, Y. Rosenberg-Hasson at the Stanford Human Immune Profiling Center for help with the Luminex analysis, and T. Doyle for the expert advice on the micro-CT analyses. Funding: P.S. and N.B. are supported by NIH/NIAMS grant R01 AR070865-01. F.C.G. is supported by an NSF GRFP award. This work used the Genome Sequencing Service Center by the Stanford Center for Genomics and Personalized Medicine Sequencing Center, supported by grant award NIH S10OD020141 and the Stanford Center for Innovation in In Vivo Imaging (SCI3) supported by NIH S10 Shared Instrumentation Grant 1S10OD02349701. Author contributions: P.S., F.C.G., and N.B. designed the overall study, selected the methods, and determined the outcome analyses. P.S. performed in vivo and in vitro experiments. F.C.G. performed in vitro experiments and bioinformatic analyses. S.D. and V.M. performed blinded OARSI scoring. P.F.I. and S.B.G. provided the human samples from patients with OA. P.S. and F.C.G. analyzed data. P.S., F.C.G., and N.B. wrote the manuscript. Competing interests: No financial support or other benefits have been obtained from any commercial sources for this study. The authors declare that they have no competing financial interests. Data and materials availability: All data associated with this study are present in the paper or the Supplementary Materials. Raw data for RNA-seq and RRHP datasets are deposited in GEO (accession number: GSE143576).

Stay Connected to Science Translational Medicine

Navigate This Article