Dysfunctional polycomb transcriptional repression contributes to lamin A/C–dependent muscular dystrophy

Lamin A is a component of the inner nuclear membrane that, together with epigenetic factors, organizes the genome in higher order structures required for transcriptional control. Mutations in the lamin A/C gene cause several diseases belonging to the class of laminopathies, including muscular dystrophies. Nevertheless, molecular mechanisms involved in the pathogenesis of lamin A–dependent dystrophies are still largely unknown. The polycomb group (PcG) of proteins are epigenetic repressors and lamin A interactors, primarily involved in the maintenance of cell identity. Using a murine model of Emery-Dreifuss muscular dystrophy (EDMD), we show here that lamin A loss deregulated PcG positioning in muscle satellite stem cells, leading to derepression of non–muscle-specific genes and p16^INK4a, a senescence driver encoded in the Cdkn2a locus. This aberrant transcriptional program caused impairment in self-renewal, loss of cell identity, and premature exhaustion of the quiescent satellite cell pool. Genetic ablation of the Cdkn2a locus restored muscle stem cell properties in lamin A/C–null dystrophic mice. Our findings establish a direct link between lamin A and PcG epigenetic silencing and indicate that lamin A–dependent muscular dystrophy can be ascribed to intrinsic epigenetic dysfunctions of muscle stem cells.

Our recent results showed a lamin A/C–PcG crosstalk during in vitro myogenesis (10). We thus wondered if the altered MuSC balance observed in LmnaΔ8–11^–/– muscles might be ascribed to aberrant PcG functions. We first performed immunostaining of Ezh2, the catalytic subunit of polycomb repressive complex 2 (PRC2) (Supplemental Figure 3A) in d19 MuSCs. We fixed MuSCs before FACS isolation to preserve the nuclear architecture of lamin A–deficient cells (see Methods). We found a general intranuclear diffusion of Ezh2 in LmnaΔ8–11^–/– MuSCs, ascertained by measuring PcG body parameters (22) (Supplemental Figure 3, A–C). We also measured Ezh2 expression both in MuSCs and whole muscles (Supplemental Figure 3D) and we analyzed Ezh2 protein levels in whole muscles (Supplemental Figure 3, E and F; see complete unedited blots in the supplemental material). We found no major differences between LmnaΔ8–11^+/+ and LmnaΔ8–11^–/– mice. To further analyze the Ezh2 intranuclear distribution in QSCs, we performed triple immunostaining on muscle cryosections (Supplemental Figure 3, G and H). Ezh2 levels, assessed measuring fluorescence intensity, were similar in LmnaΔ8–11^+/+ and LmnaΔ8–11^–/– MuSCs in PAX7⁺Ki67^– and PAX7⁺Ki67⁺ cells. Because Ezh2 is hardly detectable in adult quiescent satellite cells (23–25), this result suggests that during postnatal growth developmental signals might instead contribute to maintaining Ezh2 expression in nonproliferating MuSCs.

On the other hand, evaluation of the number of PcG bodies on the same sections highlighted a decrease in the number of Ezh2 bodies in the mutant (Supplemental Figure 3, I and J), leading us to conclude that the absence of lamin A/C does not affect Ezh2 protein levels but influences its nuclear distribution. To gain further insight into possible PcG-dependent transcriptional defects, we performed RNA sequencing (RNA-seq) on freshly isolated MuSCs at d19, finding 1424 upregulated genes and 1842 downregulated genes in the LmnaΔ8–11^–/– MuSCs compared with WT (Supplemental Figure 4A). Interestingly, performing a gene set enrichment analysis (GSEA) based on differential expression generated after conditional ablation of Ezh2 in MuSCs (24) and LmnaΔ8–11^–/– upregulated genes, we found a significant association between the 2 data sets, suggesting that lamin A absence impairs Ezh2 function (Supplemental Figure 4B). We also followed the deposition of the Ezh2-dependent H3K27me3 histone mark in LmnaΔ8–11 mice by quantitative spike-in ChIP-seq (26) (see Supplemental Methods and Supplemental Figure 4, C and D). Integrative analysis of RNA-seq and ChIP-seq revealed that upregulated genes in the LmnaΔ8–11^–/– condition are significantly enriched for H3K27me3 targets (identified in the WT condition) (Figure 3A). Indeed, analysis of H3K27me3 distribution around the transcription start sites (TSSs) and along the body of genes indicated a decrease in this repressive mark in LmnaΔ8–11^–/– MuSCs compared with WT (Figure 3, B and C), which was not accompanied by a statistically significant decrease in global H3K27me3 levels in MuSCs (Supplemental Figure 5, A and B; see complete unedited blots in the supplemental material) and in whole muscle (Supplemental Figure 5, C and D; see complete unedited blots in the supplemental material). In contrast, a deep analysis of H3K27me3 ChIP-seq read coverage in the intergenic genomic regions between the known H3K27me3 enrichment peaks interestingly showed a higher average coverage in the LmnaΔ8–11^–/– MuSCs compared with WT counterparts (Figure 3D). These results are compatible with a diffusion of PcG proteins along the chromatin fibers rather than a complete PcG displacement. To identify the PcG targets mostly affected by lamin A deficiency, genes were grouped according to their transcription level in WT MuSCs. We thus defined 4 equally sized groups of genes based on expression level quartiles (Figure 3E). For each expression category, we reanalyzed the H3K27me3 distribution along the body of genes and at the TSS and the percentage of upregulated genes in the LmnaΔ8–11^–/– MuSCs (Figure 3, E and F, and Supplemental Figure 5E). In quartile I we found only a small percentage (0.65%) of upregulated genes in LmnaΔ8–11^–/–, suggesting that the H3K27me3 decrease/redistribution is not sufficient to activate transcription in highly repressed genes (Figure 3F, Supplemental Figure 5E, and Supplemental Figure 6A). In contrast, quartiles II, III, and IV are more affected by the diminished H3K27me3 levels in LmnaΔ8–11^–/– (Figure 3F, Supplemental Figure 5E, and Supplemental Figure 6B), showing a percentage of upregulated genes between 5% and 9%. Specifically, we noticed that in WT MuSCs, H3K27me3 ChIP-seq signal enrichment around the TSS and within the body of genes is progressively lower in quartiles of higher expression, as expected (Figure 3F, quartiles III and IV, and Supplemental Figure 5E). However, for LmnaΔ8–11^–/– mice the decrease in H3K27me3 signal inside the gene body is relatively less marked than in WT mice; in fact, the average enrichment is slightly higher. We quantified and confirmed this observation by considering the ratio of H3K27me3 ChIP-seq enrichment signal at the TSS and 2.5 kb downstream of the TSS, for each gene, in WT and LmnaΔ8–11^–/– mice (Supplemental Figure 7A), showing that this ratio is significantly different for higher expression quartiles (Supplemental Figure 7B).

Figure 3

LmnaΔ8–11^–/– dystrophic MuSCs display PcG displacement. (A) Heatmap reporting log₂ ratios of observed over expected (colored bar) number of genes in the intersections between H3K27me3 targets identified in LmnaΔ8–11^+/+ mice and the upregulated genes in LmnaΔ8–11^–/– mice. Fisher’s exact test P = 2.38 × 10^–5. (B–D) Average profile of H3K27me3 ChIP-seq signal calculated as the IP/input ratio over annotated mouse genes. (B) Average profile of H3K27me3 signal around the TSS. (C) Average profile of H3K27me3 signal along the gene body. TES, annotated transcript end. (D) Average profile of H3K27me3 signal in regions outside H3K27me3 peaks and outside annotated genes. (E) Normalized expression distribution of genes stratified using WT expression level in the 3 biological replicates (see Supplemental Methods). Distribution of average log₂ transcripts per million (TPM + 0.1) values is plotted for WT and hom. Data in the boxes extend from the 25th to the 75th percentiles with the median indicated. The upper whisker extends from the hinge to the highest value that is within 1.5 × IQR of the hinge, where IQR is the interquartile range, or distance between the first and third quartiles. The lower whisker extends from the hinge to the lowest value within 1.5 × IQR of the hinge. Data beyond the end of the whiskers are outliers and plotted as points. (F) Average profile of H3K27me3 signal (IP/input) along the gene body using gene categories as in E. WT = LmnaΔ8–11^+/+; hom = LmnaΔ8–11^–/–.

The altered PcG binding observed in LmnaΔ8–11^–/– MuSCs prompted us to examine in greater detail the bivalent genes, a subgroup of PcG targets whose expression is more susceptible to variations in PcG occupancy (27). Bivalent genes are characterized by the concurrent presence of both H3K27me3 and H3K4me3 marks around TSSs and have an intermediate gene expression state (28). We first performed H3K27me3 and H3K4me3 ChIP-seq in WT MuSCs (Supplemental Figure 8, A and B) and we defined bivalent genes using the parameters described in Bernstein et al. (17) for the H3K4me3 window at the TSS (Supplemental Figure 8C). Then, we tested the association between bivalent and upregulated genes in the LmnaΔ8–11^–/– MuSCs by means of Fisher’s exact test. We observed a significant overrepresentation of bivalent genes among upregulated ones in the LmnaΔ8–11^–/– MuSCs (Figure 4A). To gain more insight into the biological relevance of deregulated genes in the mutant mice, we performed semantic similarity analysis of all Gene Ontology (GO) terms associated with upregulated genes (Figure 4B) together with GSEA (Supplemental Figure 8, D and E). These analyses showed a negative correlation with muscle specification (Supplemental Figure 8D) together with an acquisition of markers related to lipid metabolic processes (Figure 4B and Supplemental Figure 8E). Notably, Fisher’s exact test analysis highlighted a significant overlap between genes with bivalent promoters and the LmnaΔ8–11^–/– MuSC upregulated genes involved in adipogenesis (Figure 4C), suggesting that lamin A is involved in preserving MuSC identity by ensuring the correct PcG-mediated transcriptional repression of non-muscle genes.

Figure 4

Lamin A–PcG–mediated transcriptional repression preserves MuSC identity. (A) Heatmap reporting log₂ ratios of observed over expected number of genes (colored bar) in the intersections between bivalent promoters identified in WT satellite cells and the upregulated genes in LmnaΔ8–11^–/– mice. Fisher’s exact test P = 4.57 × 10^–7. (B) Semantic similarity analysis of GO terms enriched in upregulated genes in hom versus WT comparison (FDR < 0.05) with macrocategories identified using the REVIGO web tool (http://revigo.irb.hr/). (C) Heatmap reporting log₂ ratios of observed over expected number of genes (colored bar) in the intersections between upregulated genes in LmnaΔ8–11^–/– mice in Pparg-related GO terms and the bivalent genes identified as above. Fisher’s exact test P = 6.73 × 10^–6. WT = LmnaΔ8–11^+/+; hom = LmnaΔ8–11^–/–.

Given this strong association between bivalent gene reactivation and adipogenesis markers (Figure 4C), we analyzed different lipid-related GO categories, finding among the top GO terms peroxisome proliferator–activated receptor γ (Pparg) (Supplemental Table 1). This master transcription factor for adipose cell differentiation (29, 30) was found to be significantly upregulated in LmnaΔ8–11^–/– MuSCs (FDR < 0.05). Moreover, the Pparg gene is a polycomb target and has a bivalent signature in WT MuSCs (Supplemental Table 1 and Supplemental Figure 9A). These observations prompted us to analyze Pparg transcriptional deregulation. We stained muscles for PAX7 and PPARγ to directly test if MuSCs displayed aberrant expression of PPARγ in the absence of lamin A. Strikingly, we found approximately 10% of LmnaΔ8–11^–/– MuSCs that simultaneously express both muscular and adipogenesis markers, being PAX7⁺PPARγ⁺ (Figure 5, A and B). Accordingly, the genomic region of the Pparg gene showed a decrease in H3K27me3 enrichment around the TSS in the LmnaΔ8–11^–/– MuSCs, accompanied by transcriptional upregulation (Supplemental Figure 9B). To evaluate if the aberrant expression of adipogenic genes in MuSCs of mutant mice culminates with fatty infiltration we performed immunofluorescence staining for perilipin 1, a protein present on the surface of lipid droplets (ref. 31 and Figure 5, C and D), on cryosections of muscles derived from d19 LmnaΔ8–11 mice. We found large areas of adipose accumulation between myofibers of lamin A/C^+/– and lamin A/C^–/– muscles, which were instead undetectable in the WT mice. Considering the key role of PcG proteins in mediating the formation of chromatin loop structures (32, 33), we reasoned that the loss of H3K27me3 and transcriptional upregulation of the Pparg locus could be related to the alteration of chromatin 3D structure. The genomic 3D architecture is organized in structurally separated topologically associated domains (TADs), chromosomal structures that favor intradomain looping interactions (34). TADs can be identified by genome-wide chromosome conformation capture (Hi-C) and are largely conserved across different cell types. We verified that the Pparg locus is included in a TAD encompassing a region extending also upstream of the Pparg locus itself, using high-resolution Hi-C data on mouse embryonic stem cells (35) and the 3D Genome Browser (ref. 36 and Supplemental Figure 10). Then, we performed 3D multicolor DNA FISH analysis on prefixed MuSCs using 1 BAC probe overlapping the Pparg promoter and a second probe annealing at the other TAD border. We observed an overlap of the signals from the 2 regions in the WT LmnaΔ8–11^+/+ MuSCs that indicates the presence of a DNA looping in cis (Figure 5, E and F). By contrast, in LmnaΔ8–11^–/– MuSCs the distance between the signals was higher, definitely suggesting the lack of DNA/DNA interactions. Indeed, from the analysis of H3K27me3 ChIP-seq tracks we noticed in the LmnaΔ8–11^–/– MuSCs a reduction in H3K27me3 peaks upstream of the Pparg locus (Supplemental Figure 10). FISH analysis also highlighted that in WT the entire genomic region is close to the nuclear periphery (Figure 5, E and G), whereas in LmnaΔ8–11^–/– it is relocated in the nuclear interior, suggesting that lamin A absence interferes with chromatin anchoring to the nuclear lamina and PcG-dependent DNA conformation.

Figure 5

Deregulation of Pparg locus in LmnaΔ8–11^–/– MuSCs. (A) Immunohistochemical staining of PAX7 and PPARγ markers in LmnaΔ8–11 mice at 19 days of postnatal growth (d19). Scale bars: 50 μm. (B) Quantification of PPARγ⁺ MuSCs in A; n = 4–5 animals per genotype. Data are the mean ± SEM. (C) Immunohistochemical staining of perilipin (PLIN) in LmnaΔ8–11 muscles at d19. Nuclei of muscle fibers were stained with DAPI. Scale bars: 25 μm. (D) Quantification of perilipin staining in C; n = 5 animals per genotype. (E) Representative image of FISH analysis of fixed and sorted MuSCs from LmnaΔ8–11 mice at d19 with probes indicated in Supplemental Figure 10. Scale bars: 2 μm. (F) Quantification of FISH probe (represented in Supplemental Figure 10) distances (x axis) versus cumulative frequency distributions (y axis). Only probes with distances of 0.5 μm or less are reported. n = 1–2 animals per genotype. (G) Quantification of FISH probe position with respect to the nuclear envelope. In the box-and-whisker plots in D and G, horizontal lines within the boxes represent the medians, upper and lower bounds of the boxes represent quartiles Q3 (75th percentile) and Q1 (25th percentile), respectively, and whiskers min to max. *P < 0.05; ***P < 0.001 by 2-way ANOVA with multiple comparisons (B and D) or Kolmogorov-Smirnov test (F and G). WT = LmnaΔ8–11^+/+; het = LmnaΔ8–11^+/–; hom = LmnaΔ8–11^–/–.

Taken together, these results clearly point toward a role of lamin A in mediating PcG-transcriptional repression in MuSCs to safeguard their identity and regenerative capacity. This lack of cell identity and the impairment of self-renewal displayed by LmnaΔ8–11^–/– MuSCs are all features reminiscent of the phenotype described for Ezh2-null MuSCs (24). Moreover, the impairment in self-renewal and the progressive decline of the MuSC pool are also typical traits of aged MuSCs (37) in which both lamin A/C and PcG proteins play a key role (38, 39). A major cellular mechanism that ensures self-renewal and hence the maintenance of the MuSC pool is asymmetric division (40). At the molecular level, in aged mice, the accumulation of the activated form of p38 (phospho-p38 [p-p38]) and its symmetric distribution in MuSC doublets heavily compromise the self-renewal capacity, leading to MuSC functional decline (41, 42). To test whether premature exhaustion of quiescent LmnaΔ8–11^–/– MuSCs could be ascribed to defective asymmetric division, we stained myofiber-associated MuSCs for p-p38 after 48 hours of culture, a timing at which d19 myofibers formed MuSC-derived doublets (Figure 6A). In contrast to WT, LmnaΔ8–11^–/– MuSC doublets showed a preferential symmetric distribution of p-p38, quantified by relative fluorescence intensity (Figure 6, A and B), often accompanied by a planar orientation with respect to myofibers (see Supplemental Methods and Figure 6C). This highlights problems in asymmetric division, which should be instead characterized by apico-basal orientation (43). In line with this result, in LmnaΔ8–11^–/– muscle sections we found higher amounts of p-p38⁺ (Supplemental Figure 11, A and B) MuSCs and signs of genomic instability, as measured by increased γH2AX DNA repair signal foci (Supplemental Figure 11, C and D), not accompanied by apoptosis or necrosis as evidenced by annexin staining (Supplemental Figure 12A). To test if defective asymmetric division is associated with premature senescence we then analyzed RNA-seq to determine if LmnaΔ8–11^–/– MuSCs share the same transcriptional signature of MuSCs isolated from aged mice. We performed a GSEA using 2 different RNA data sets from MuSCs of 24-month-old mice (25, 44) and LmnaΔ8–11^–/– upregulated genes. In line with our hypothesis, we found that d19 LmnaΔ8–11^–/– MuSCs present an upregulated transcriptome similar to 20- to 24-month-old MuSCs (Figure 6D and Supplemental Figure 12B), but different from geriatric 28- to 32-month-old MuSCs (Supplemental Figure 12C). At the molecular level, the senescence program is supported by upregulation of some PcG-regulated cyclin-dependent kinase inhibitors (CDKIs) (45) such as p21, which is involved in cellular senescence and in cell cycle arrest. p21 maintains the viability of DNA damage–induced senescent cells (46) and aberrant expression of p21 has been observed in EDMD-derived human myoblasts (47). ChIP-seq and RNA-seq analyses of the Cdkn1a/p21 locus showed a displacement of Ezh2 from the promoter accompanied by an upregulation of p21 transcripts in LmnaΔ8–11^–/– MuSCs (Figure 6E).

Figure 6

LmnaΔ8–11^–/– MuSCs acquire senescence transcriptional traits. (A) Representative image of myofiber-derived MuSCs from LmnaΔ8–11 mice at d19 immunostained for p-p38 and PAX7 after 48 hours of culture. Scale bars: 25 μm. (B) Quantification of asymmetric and symmetric divisions assessed by p-p38 distribution as shown in A. (C) Quantification of asymmetric apico-basal division versus symmetric planar divisions. n = 46 ± 6 doublets of MuSCs per genotype, n = 7–9 mice per group. Data in B and C are the mean ± SEM. (D) GSEA of expression data from old and young mouse quiescent satellite cells (25). Upregulated (log[fold change] > 1) genes in hom versus WT comparison added to Biocarta mouse pathways from the gskb R package were used as gene sets (NES = 4.70, FDR < 1 × 10^–4). (E) ChIP-seq of H3K27me3 mark and RNA-seq signal tracks on the Cdkn1a/p21 locus. Promoter regions are highlighted by light blue rectangles. Statistics by 2-way ANOVA with multiple comparisons. **P < 0.01; ***P < 0.001. WT = LmnaΔ8–11 ^/+; het = LmnaΔ8–11^+/–; hom = LmnaΔ8–11^–/–.

To further corroborate our findings, we also analyzed the Cdkn2a locus, a PcG target primarily involved in muscular senescence (ref. 44 and Supplemental Figure 13A). Two transcripts, p16^INK4a and p19^ARF, originate from the Cdkn2a locus (48). Interestingly, it was recently reported that p16^INK4a expression is a second event, subsequent to p21 upregulation, in cellular senescence progression (49). In line with these observations, p16^INK4a expression is specifically induced in geriatric 28- to 32-month-old MuSCs (but not in 24-month-old MuSCs) (44). Moreover, depletion of p16^INK4a is sufficient to reduce senescence-associated gene expression in geriatric MuSCs. RNA-seq and quantitative reverse transcription PCR (qRT-PCR) on d19 LmnaΔ8–11^–/– MuSCs did not reveal any transcription of p16^INK4a in WT or LmnaΔ8–11^–/– (Figure 7A). However, qRT-PCR analysis performed in older mice (d26) revealed higher levels of p16^INK4a transcripts in LmnaΔ8–11^–/– MuSCs and whole muscles compared with the WT counterpart (Figure 7A), suggesting a transition during dystrophy progression toward a geriatric condition. We thus decided to test whether genetic ablation of the Cdkn2a locus could reverse LmnaΔ8–11^–/– MuSC premature aging, by crossing LmnaΔ8–11^+/– with Cdkn2a^–/– mice (50). Analysis of the LmnaΔ8–11^+/+ background showed no differences in the percentages of QSCs and ASCs, nor in CSA, suggesting that Cdkn2a is dispensable for postnatal muscle development (Figure 7, B–D; LMNA^+/+). On the other hand, Cdkn2a^–/– LmnaΔ8–11^–/– mice partially rescued the quiescent MuSC pool and CSA defects observed in the absence of lamin A (Figure 7, B–D; LMNA^–/–; Supplemental Figure 13, B and C), emphasizing that lamin A–dependent muscular dystrophy might be due to progressive MuSC functional decline caused by acquisition of premature aging features.

Figure 7

Cdkn2a genetic ablation restores regenerative capacity of LmnaΔ8–11^–/– dystrophic mice. (A) Transcriptional analysis of p16^INK4 and p19^ARF at the Cdkn2a locus in LmnaΔ8–11 mouse MuSCs (left graph) at d19 and d26 and whole muscles (right graph) at d26. Values were normalized to Gapdh and compared with the average of WT amplification. nd, not detected. n = 3–10 animals per genotype. (B) Immunohistochemical staining of PAX7 and MYOD markers in Cdkn2a/LmnaΔ8–11 mice at d19. Basement membrane of muscle fibers was stained with anti-laminin. Activated, ASC (PAX7⁺/MYOD⁺) and self-renewing, QSC (PAX7⁺MYOD^–) MuSCs are shown. Scale bars: 50 μm. (C) Quantification of MuSC pool composition in B. n = 4–5 animals per genotype. Data in A and C are the mean ± SEM. (D) Quantification of myofiber size, evaluated by the cross-sectional area (CSA). n = 600 muscle fibers. n = 4–5 animals per genotype. The horizontal lines within the boxes represent the medians, upper and lower bounds of the boxes represent quartiles Q3 (75th percentile) and Q1 (25th percentile), respectively, and whiskers min to max. *P < 0.05; **P < 0.01; ***P < 0.001 by unpaired t test (A) or by 1-way (D) or 2-way (C) ANOVA with multiple comparisons. WT = LmnaΔ8–11^+/+; hom = LmnaΔ8–11^–/–.

CL conceived the project and designed experiments. AB and CM performed experiments and analyzed data. FL and DP performed and interpreted ChIP-seq experiments. SV performed bioinformatic analysis. GP, VR, RR, and CB performed experiments in mice. AB, AC, and BB performed and interpreted FISH experiments. LA, FG, and GO quantified immunofluorescence and FISH images. MDB performed FACS. FF and CP analyzed ChIP-seq data. CL supervised the project and analyzed data. CL, AB, and CM wrote the manuscript. All authors edited and approved the manuscript.

View Supplemental data

We thank Maria Vivo, Gisèle Bonne, Federica Marasca, Pierluigi Manti, Davide Gabellini, Lorenzo Puri, Daniela Palacios, Giovanna Lattanzi, the Italian network of laminopathies, and members of the laboratory for stimulating discussions and constructive criticisms. We are grateful to Chiara Cordiglieri and INGM Imaging Facility for assistance during image acquisition and to Mariacristina Crosti, Monica Moro, and the INGM FACS Facility for assistance in cell sorting. This work was supported by grants from the Italian Minister of Health no. GR-2013-02355413 to CL, My First AIRC Grant (MFAG) no. 18535 to CL, AFM-Telethon no. 21030 to CL and FF, and Cariplo 2017-0649 to CL and FF.

Address correspondence to: Lanzuolo Chiara. Via Fratelli Cervi, 93, 20090 Segrate (MI), Italy. Phone: 39.02.00660358; Email: lanzuolo@ingm.org. MC’s present address is: CNR Institute of Molecular Biology and Pathology (IBPM), Rome, Italy. VS’s present address is: San Raffaele Telethon Institute for Gene Therapy (SR-Tiget), IRCCS San Raffaele Scientific Institute, Milan, Italy.

Conflict of interest: The authors have declared that no conflict of interest exists.

Copyright: © 2020, American Society for Clinical Investigation.

Reference information: J Clin Invest. 2020;130(5):2408–2421.https://doi.org/10.1172/JCI128161.

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