Lymphatic–stem cell crosstalk promotes tendon regeneration via Notch1–Srebp2–mediated cholesterol metabolism

Domain: biology
Published: 2025-12-26
Editorial outcome: major_revision

Abstract
Tendons, as bradytrophic tissues, exhibit limited healing capacity. The role of lymphatic vessels in tendon regeneration is largely overlooked. By comparing gene expression of the control and injured tendon cells from both neonatal and adult mice, we identify lymphatic vessels as key regenerative niche elements. Using the Transparent Embedding Solvent System for clearing in Prox1-CreERT2;tdTomato mice, we find that lymphatic signaling was activated following tendon injury, directing the fate of distinct tendon stem/progenitor cell (TSPC) subsets. Mechanistically, the glycoprotein Reelin secreted from lymphatic endothelial cells (LECs) triggers VLDLR-dependent phosphorylation of DAB1 in TSPCs. Activated DAB1 binds to NOTCH1 at the tyrosine Y-200 residue, initiating Srebp2-mediated cholesterol metabolism. Functionally, Reln−/− mice and Prox1-CreERT2;Relnfl/fl mice exhibit impaired tendon regeneration, underscoring the critical role of Reelin signaling. Furthermore, a slow-release Reelin delivery system established using mesoporous silica nanoparticles enhance tendon regeneration in mice and rabbits. These findings highlight the key role of lymphoangiocrine signaling in determining TSPC fate during tendon regeneration. Overall, this study provides a foundation for promoting tissue regeneration by targeting lymphatic signals.

Introduction
Tendon injuries, caused by overuse or age-related degeneration, account for nearly half of musculoskeletal disorders and are frequently linked to poor clinical outcomes1, creating a substantial burden on global healthcare systems2. The main challenge is fibrotic scarring in healed tendons, which disrupts matrix continuity and impairs function. Tendons are composed primarily of dense extracellular matrix (ECM), maintained by tenocytes and tendon stem/progenitor cells (TSPCs)3,4. Despite therapeutic advances, effective strategies to improve tendon healing remain scarce, underscoring the need for a deeper understanding of TSPC biology and regulatory mechanisms5,6. TSPCs exhibit clonogenicity, self-renewal, and multilineage differentiation, enabling limited tendon homeostasis under physiological conditions. Transcription factors and signaling pathways associated with TSPCs were first identified in tendon development studies, which established Scleraxis (Scx) as a tendon progenitor marker7,8. Advances in single-cell sequencing have revealed heterogeneity in TSPC populations, identifying different subtypes, such as Nestin+, and Tppp3+Pdgfrα+, Cd146+, and Cd9+Cd271+ subpopulations, which play critical roles in tendon development and regeneration9–11. Although progress has been made in defining early tendon formation, the cellular dynamics underlying differentiation, maturation, and scar formation during healing remain poorly understood.

Stem cell properties are regulated by specialized niches that provide signals essential for maintenance and differentiation. TSPCs reside within a unique niche enriched in ECM components, such as fibromodulin and biglycan, with low cellularity, which hinders repair after injury4. Although tendons are regarded as sparsely vascularized, they possess greater vascular complexity than previously believed. In particular, lymphatic vessels infiltrate repair tissue after injury, as shown in a rat tendon lesion model12,13. Lymphatic vessels have been extensively studied in other tissues, such as the intestine, for their roles in fluid transport, immune cell trafficking, and signaling14,15. However, their homeostatic and regenerative functions in tendons have not been fully examined. Reelin, a large extracellular glycoprotein traditionally associated with neuronal migration and cortical development16,17, has also been detected in lymphatic vessels. In peripheral tissues, Reelin regulates cardiac growth and repair, epithelial cell proliferation, and intestinal crypt stability18,19. It may also influence limb development and tissue regeneration; however, its mechanisms in tendons remain unclear20.

In this study, we identified lymphatic vessels as a central signaling hub for tendon regeneration and TSPC fate determination. Specifically, lymphoangiocrine Reelin was found to activate Notch1-Srebp2-mediated cholesterol metabolism, preserving TSPC stemness and enhancing tendon repair. Furthermore, a slow-release Reelin delivery system based on mesoporous silica nanoparticles (MSNs) was shown to markedly improve tendon regeneration. These findings reveal significant interaction between lymphatic vessels and TSPCs, offering a promising strategy for tendon regeneration.

Results
Lymphatic endothelial cells (LECs) are a key component of the tendon stem cell niche
Neonatal tendons (before postnatal day 7) acquire functional recovery after injury, while adult tendons typically heal with fibrovascular scarring and persistent impairment21. To identify the molecular drivers of tendon regeneration and maturation, we compared gene expression in cells from control and injured tendons of neonatal and adult mice using single-cell RNA sequencing (scRNA-seq, Fig. 1A). The tendon-derived cells were classified into nine major cell clusters based on gene expression profiles (Figs. 1B–E and S1A–D). Cluster 0 showed high expression of tenogenesis-associated genes, including Scx, tenomodulin (Tnmd), tenascin-C (Tnc), and fibromodulin (Fmod)22. To examine the mechanisms of tendon maturation, we isolated tenogenic subset cluster 0 for uniform manifold approximation and projection–based clustering. Clusters were annotated using marker gene expression and gene set scores derived from predefined canonical stem cell, progenitor cell, and mature tenocyte genes (Figs. 1F, G and S1E). The cluster expressed stem cell markers lymphocyte antigen 6 complex, locus A (Ly6a) and Thy-1 cell surface antigen, was annotated as tendon stem cells (TSCs). The cluster exhibited the highest progenitor cell-associated gene set score, leading to its classification as tendon progenitor cells (TPCs). Another cluster with high expression of the ECM genes Fmod and thrombospondin-4 (Thbs4) for mature tendon cells was designated tenocytes (Figs. 1G and S1E). The stemness scores calculated based on the stemness-related gene set showed decreasing stemness scores from TSCs to tenocytes, which aligned with the subpopulation definitions (Fig. 1H). Furthermore, Monocle3 and CytoTRACE analysis confirmed differentiation path from TSCs through TPCs to tenocytes (Fig. 1I, J), suggesting the three distinct TSPC subsets transited from neonatal to adult tissue. Immunofluorescence staining and gene expression on UMAP analysis demonstrated higher Ly6a levels in neonatal tendons, while higher Scx and proteoglycan 4 (Prg4) levels in adult ones (Fig. S1F, G).

The environment niche, where stem cells resident in, plays a crucial role in determining cell maintenance and fate23. The cell-cell communication analysis highlighted increased interactions between LECs and tenogenic cells after injury compared with control one (Fig. 1K). To define this interaction mechanisms, we isolated tenogenic related subsets and endothelial cells for UMAP-based clustering and annotation (Fig. S1H–J). We further examined the interaction between LECs with three subpopulations of tenogenic cells and the results revealed specifically increased interaction between LECs and TCs subsets after tendon injury (Fig. 1L). These findings indicate that lymphatics in tendons are key regulators of stem cell fate and tendon healing.

Lymphatic vessels expand after tendons injury
Tendons are composed of dense fibrous connective tissue, which render the lymphatic network difficult to visualize using conventional immunostaining13. To overcome this, we modified our previously reported Transparent Embedding Solvent System (TESOS)24 for efficient tendon clearing and immunolabeling, enabling rapid, high-resolution, and three-dimensional (3D) confocal imaging (Fig. S2A). To analyze the lymphatic vessels in and around tendons, we used Prox1-CreERT2;tdTomato mice and performed whole-mount imaging of the Achilles and Patellar tendons. This approach enabled visualization of Prox1-Tdtomato-positive lymphatic vessels in tendons (Fig. 2A, B; Supplementary Movies 1 and 2). To validate their identity, we performed immunolabeling using vascular endothelial and lymphatic vessel markers. Prox1-Tdtomato-positive lymphatic structures co-expressed the vascular endothelial marker CD31 and the LEC marker LYVE1 (Fig. S2B–D). Because vascular endothelial growth factor C (VEGF-C)/VEGF receptor 3 (VEGFR3) signaling is essential for lymphatic development and maintenance25,26, we also demonstrated VEGFR3 expression in Prox1-Tdtomato- positive lymphatic vessels via immunostaining (Fig. S2E). Moreover, to analyze the interstitial fluid drainage function of these structures, we injected indocyanine green (ICG) and Evans blue to trace lymphatic flow. Subcutaneous ICG injections into the footpad showed uptake in Achilles tendons within 30 min as well as time-dependent clearance (Fig. S2F). Evans blue accumulated in LYVE1 positive lymphatic vessels of tendon tissues, which was also cleared over time (Fig. S2G)27,28. These results indicate the functional lymphatic vessels distribution of tendon tissues. However, it was difficult to distinguish whether they distributed superficially on the tendon with in the tendon.

Next, to test LECs responses to tendon injury, we established an Achilles tendon transection model. Lymphatic vessels expanded after tendon injury in neonatal and adult mice, with increased Prox1-Tdtomato-positive and LYVE1-positive lymphatic vessels (Figs. 2C, D and S2H, I). Quantitative analysis also revealed elevated Prox1 and Lyve1 mRNA expression after tendon injury (Fig. S2J). As both the Achilles and patellar tendons are paratenon-sheathed tendons, comprising paratenon sheath and tendon substance. We therefore analyzed the precise distribution of expanded lymphatic vessels following injury. Our results demonstrated that in the control group, Prox1-Tdtomato and LYVE1-positive lymphatic vessels were mainly localized to the superficial tendon sheath. After injury, however, lymphatic vessels infiltrated to the tendon substance, as shown by the expression of COL1α1 and FMOD (Fig. S2K, L). Moreover, we treated injured tendons with VEGFR3-blocking antibody, which inhibited LYVE1-positive lymphatic vessel expansion (Fig. S2M). To model chronic tendinopathy, we induced Achilles tendon overuse. In this model, lymphatic expansion also occurred, as shown by Prox1-Tdtomato-positive vessels (Fig. S2N). Together, these results demonstrate the lymphatic vessels are mainly localized on tendon superficial sheath in homeostasis, while acute and chronic stress drive lymphatic expansion and growing into tendon.

Next, we examined how lymphatics contribute to tendon healing. CellChat analysis of TSPCs and LECs identified Reelin (Reln) signaling as the major pathway enriched after injury (Fig. 2E). Expression analysis also showed high Reln levels in LECs (Fig. 2F). Reelin protein has been reported to promote cardiac repair18. Immunostaining showed Reelin colocalization with the Prox1-tdTomato and LYVE1 in tendons. Reelin expression was also increased following tendon injury (Fig. 2G, H). Moreover, the ELISA assay verified the levels of Reelin was increased after tendon injury (Fig. S2O), indicating that lymphoangiocrine Reelin signaling is activated postinjury and may regulate tendon healing.

To assess the relevance of these findings in humans, we analyzed a published single-cell dataset of human tendons (Fig. S3)29. We compared gene expression in tendon cells from healthy samples with cells from pathologic samples at 3 days postinjury (dpi), 10 dpi, and 12 weeks post injury (wpi) (Fig. S3A–D). CellPhoneDB analysis revealed significant interactions between tenogenic cells and LECs (Fig. S3E). LECs count increased in pathologic samples at 10 dpi (proliferative phase) and 12 wpi (remodeling phase) (Fig. S3F). Interaction strength between tenogenic cells and LECs was higher at 10 dpi and 12 wpi compared with 3 dpi (Fig. S3G). Differential gene pathway enrichment analysis showed that ECM-receptor interaction signaling was upregulated in injured human tendons relative to healthy ones (Fig. S3H). These findings support increased lymphatic signaling involvement in tendon injury.

Reln signaling supports tendon regeneration
To examine the role of Reln signaling in tendon regeneration, we bred Reln−/− mice (Fig. S4A). Their Achilles, Patellar, and tail tendons appeared hypoplastic and more translucent compared with controls. Additionally, histological analysis showed reduced collagen density and fewer cells in Reln−/− mice tendons relative to controls (Figs. 3A and S4B). To evaluate Reln signaling in tendon healing, we created Achilles tendon transection defects in neonatal control and Reln−/− mice. By 4 weeks postoperatively, controls regained near-normal function, whereas Reln−/− mice showed limited regeneration, restricted mobility, and longer, narrower hind paw prints. Achilles functional index (AFI) values were lower in Reln−/− mice compared with controls (Fig. 3B). Newly formed tendons in control mice displayed well-organized, abundant collagen fibers, whereas Reln−/− mouse tissues contained sparse, fibrocartilage-like structures with fewer, disorganized collagen fibers, as shown via hematoxylin and eosin (H&E) and Masson’s trichrome staining (Fig. 3C). Polarized light microscopy confirmed thinner collagen fibers and reduced extracellular matrix deposition in newly formed Reln−/− tissues (Fig. S4C, D). Scanning electron microscopy (SEM) and transmission electron microscopy (TEM) revealed thinner, disorganized collagen fibers with decreased average diameter and density in Reln−/− mouse tendons compared with controls (Fig. 3D). Magnetic resonance imaging (MRI) with sagittal T2-weighted scans showed low-intensity, homogeneous signals with clear margins in control newly formed tendons but irregular and heterogeneous high-intensity signals in Reln−/− tendons (Fig. 3E). The inflammation-related M1 macrophage markers inducible nitric oxide synthase (iNOS) elevated in the Reln−/− repair group compared with the control repair group (Fig. S4E). Expression of M2 macrophage–related markers CD206 was lower in the Reln−/− repair group (Fig. S4F). Mechanical property analysis using an Instron tension/compression system30 revealed superior mechanical properties in control repaired tendons, which exhibited greater stress at failure and elastic modulus compared with Reln−/− tendons (Fig. 3F, G). Expression of the lymphatic markers LYVE-1 was also reduced in Reln−/− tendons compared with controls (Fig. S4G, H). Collectively, these results demonstrate that Reln signaling is critical for tendon regeneration.

Reelin secreted by LECs controls tendon regeneration
To specifically assess the role of LEC-derived Reelin in tendon regeneration, we selectively deleted Reln in LECs using Prox1-CreERT2;Relnfl/fl (Relncko) mice. We failed to detect the expression of Reln in LECs of Relncko mice (Fig. S5A). Under baseline conditions, Relncko and control Achilles tendons showed no significant morphological or histological differences (Fig. 4A–D). We then established Achilles tendon transection defects in Relnfl/fl and Relncko neonatal mice. At 4 weeks postoperatively, Relncko mice exhibited reduced mobility and function, with lower AFI values compared with the Relnfl/fl group (Fig. 4B). Newly formed tendons in Relncko mice showed decreased cellularity and disorganized collagen compared with Relnfl/fl tendons (Fig. 4C–F). SEM and TEM analyses confirmed thinner, more disordered fibers in the newly formed tissues of Relncko mice relative to control ones (Fig. 4G). MRI revealed hyperintensity signals in newly formed Relncko tendons (Fig. 4H). Functionally, newly formed Relncko tendons exhibited inferior mechanical properties (Fig. 4I, J). Additionally, the expression of the macrophage markers iNOS, and CD206 were elevated after tendon injury, while there was no significant difference between Relnfl/fl and Relncko groups (Fig. S5B, C). LYVE1 expression in Relncko tendons showed a slight but nonsignificant decrease (Fig. S5D). These findings demonstrate that LEC-derived Reelin is essential for tendon regeneration, promoting tissue organization and functional recovery.

Vldlr+ population harbors neonatal TSPCs
Next, we explored how Reelin signaling regulates TSPC fate. Analysis the expression of Reelin receptors in TSPC subpopulations showed that Vldlr but not Lrp8 (encodes ApoER2 protein) was highly expressed in TSCs (Fig. 5A). And the expression of Vldlr was upregulated in TSCs after injury (Fig. S6A). RT-qPCR and western blotting analyses confirmed higher Vldlr expression in neonatal TSPCs compared with adult ones (Fig. 5B, C), highlighting involvement of Vldlr in tendon maturation and healing.

Next, we generated Vldlr-CreERT2 mice using CRISPR/Cas9 and crossed them with R26R-tdTomato mice to generate Vldlr-CreERT2;tdTomato mice. Lineage tracing after tamoxifen induction at P3–P5 showed that Tomato+ cells contributed 4.55% ± 1.77% to 36.48% ± 7.54% of Col1α1+ tendon cells from 7 days to 3 months during tracing (Fig. 5D, E). In contrast, when tamoxifen was injected at 1 month and traced for 3 months, Tomato+ cell proportions in tendons did not change significantly (Fig. 5F, G). Following tendon injury in Achilles tendon, most Tomato+ cells infiltrated the lesion and integrated into a FMOD-rich matrix at 30 days after injury (Fig. 5H). These results indicate that Vldlr-Tomato+ cells contribute substantially to tendon development and injury repair.

To explore the role of Vldlr in tendon regeneration, we sorted Vldlr-Tomato+ and Vldlr-Tomato- tendon cells after 7 days of tamoxifen induction at P3–P5 (Fig. S6B). Vldlr-Tomato+ TSPCs displayed higher proliferation, as indicated by an increased in Ki67+ cell count (Fig. 5I). Vldlr-Tomato+ TSPCs also demonstrated a higher capacity for tenogenic differentiation, as shown by Sirius red staining and Masson trichrome staining (Fig. 5J). Transplantation of Vldlr-Tomato+ TSPCs into Achilles tendon transection model using ultraviolet-cured gelatin-methacryloyl (GelMA) as scaffolds revealed superior regenerative capacity. More organized fibers, abundant cells, and collagen deposition were detected in Vldlr-Tomato+ TSPCs transplantation group, as shown via H&E and Masson trichrome staining (Fig. 5K). Overall, Vldlr-Tomato+ TSPCs displayed a higher capacity for tenogenesis and injury repair.

Histological spatial analysis showed more VLDLR positive tendon cells located near lymphatic vessels, with this localized abundance increasing after tendon injury (Fig. S6C, D). The proliferation of VLDLR positive tendon cells was increased after injury compared with control ones. Additionally, compared with VLDLR positive tendon cells distant from lymphatic vessels, those adjacent ones showed higher proliferation index. Moreover, the non proliferating VLDLR positive cells were located relatively farther from lymphatic vessels than the proliferating cells (Fig. S6E–H). In Relncko mice, the proliferation of VLDLR positive cell was also reduced (Fig. 5L), implicating lymphatics as a signaling hub to maintain TSPC properties. To validate the effects of Reelin on TSPCs, TSPCs were treated with recombinant Reelin protein. The results showed that Reelin treatment promoted TSPC proliferation (Fig. S6I) and tenogenic differentiation (Figs. S6J, K). Following transplantation of GelMA scaffolds loaded with TSPCs or Reelin-treated TSPCs into nude mice, better-organized collagen fibers were observed in the Reelin-treated group (Fig. S6L). These findings demonstrate that Reelin enhances TSPC proliferation and tenogenesis, supporting tendon regeneration.

Reelin activates Notch1–Srebp2-mediated cholesterol metabolism
Next, we investigated how Reelin signaling regulates TSPC stemness. Expression and phosphorylation of disabled-1 protein (DAB1), a key downstream effector of Reelin, were increased after Reelin treatment (Fig. 6A). Reelin binds its receptors, recruiting DAB1 to the NPXY internalization motif of VLDLR and inducing tyrosine phosphorylation31. Consistent with this process, Vldlr levels in TSPCs increased in a time-dependent manner following Reelin treatment (Fig. S7A). Dab1 knockdown reversed Reelin-induced tenogenic differentiation, as shown via Masson trichrome and Sirius red staining (Figs. 6B, C and S7B). These results indicate that Reelin enhances tenogenesis through DAB1 phosphorylation.

Bulk RNA sequencing comparing Vldlr positive and Vldlr negative TSPCs revealed Notch1 pathway enrichment in Vldlr positive cells (Figs. 6D and S7C–E). Levels of Notch1 and its active form, the Notch1 intracellular domain (NICD), were substantially reduced in Vldlr− TSPCs (Fig. 6E). Reelin treatment elevated NICD levels and Notch1 nucleus translocation, which were suppressed by Dab1 knockdown, analyzed by western blotting and immunofluorescence (Fig. 6F–H). Coimmunoprecipitation analysis revealed a NICD–DAB1 interaction, which was enhanced by Reelin treatment, but was partially disrupted after Dab1 siRNA treatment (Fig. 6I). DAB1 is phosphorylated at multiple tyrosine (Y) residues, including Y185, Y198, Y200, Y220, and Y23232, which is critical for NICD protein stability33. The site-directed mutations of Y185F, Y198F, and Y200F, replacing tyrosine with a phenylalanine (F) to prevent phosphorylation at the tyrosine site, impaired DAB1’s ability to stabilize NICD. Additionally, Reelin-induced NICD binding was inhibited by these mutations (Fig. 6J). Notably, the Reelin-induced decrease in NICD ubiquitination was inhibited with the Y200F mutation, but not the Y185F and Y198F mutations (Fig. 6K). Thus, Reelin promotes DAB1 phosphorylation at Y200, activating Notch1 signaling. Functionally, Reelin-induced tenogenic differentiation of TSPCs were attenuated by the Notch1 inhibitor DAPT treatment (Fig. S7F, G).

VLDLR, together with LDLR and low-density lipoprotein receptor–related protein 1 (LRP1), is key endocytic receptor, recognizing apolipoprotein E (apoE)-containing lipoproteins and contributing to cholesterol metabolism and transport34. Notch1–Srebp2 was reported to regulate hematopoietic stem cell emergence35. Reelin treatment elevated total cholesterol levels in TSPCs (Fig. 6L). Mechanistically, Reelin promoted apoE lipoprotein–VLDLR binding (Fig. S8A) and increased expression of Srebp2, the master regulator of cholesterol biosynthesis34. This effect was partially blocked by DAPT treatment (Fig. 6M, N). Consistently, Reelin-induced cholesterol elevation was partially reversed by DAPT (Fig. 6O). Srebp2 target genes, including 3-hydroxy-3-methylglutaryl-CoA reductase (Hmgcr) and 3-hydroxy-3-methylglutaryl-CoA synthase (Hmgcs), were upregulated by Reelin treatment (Fig. S8B, C). Notably, Srebp2 knockdown using siRNA abrogated Reelin-induced cholesterol elevation (Fig. S8D, E). CUT&Tag-PCR assay revealed that Reelin enhanced NICD binding to the Srebp2 promoter (Fig. S8F). These findings indicate that Reelin regulates cholesterol uptake through apoE–VLDLR interaction and de novo cholesterol synthesis via Srebp2 activation. Moreover, Reelin-induced enhancements in TSPC tenogenic differentiation were partially attenuated by Srebp2 siRNA treatment (Fig. S8G, H). Thus, Reelin modulates cholesterol metabolism through Notch1–Srebp2 signaling, which preserves TSPC properties and support tenogenic differentiation.

Slow-released Reelin promotes tendon regeneration in mice and rabbits
To enhance Reelin delivery for tendon regeneration, we used MSNs to enable Reelin loading and sustained release36,37. TEM images showed spherical nanoparticles with a highly porous structure and uniform size (~190 nm). Nitrogen adsorption analysis, using the Barrett–Joyner–Halenda model, indicated average pore size of 33 nm based on the pore size distribution curve (Fig. S9A, B). Dynamic light scattering measurements suggested that MSNs had a hydrodynamic particle size of 200 nm and a zeta potential of −25 mV (Fig. S9C). Cone-shaped pores with large openings were visible on SEM images (Fig. 7A). With these pore features, MSNs achieved a Reelin loading concentration of 2 μg/mL and a loading efficiency of 40%. In protein release assays, ~60% of loaded Reelin was released over 72 h, confirming sustained delivery (Fig. 7B). MSNs also showed profound biocompatibility with TSPCs, which retained morphology, viability, and proliferation after nanoparticle exposure (Fig. 7C–E).

To investigate the regenerative effects of Reelin-loaded MSNs, they were embedded in a GelMA scaffold and applied to the Achilles tendon transection model. Reelin treatment partially improved tendon repair, and Reelin slow-release MSNs treatment resulted in superior tendon regeneration (Figs. 7F–H and S9D–F). Specifically, the Reelin slow-release MSNs showed improved mobility compared with Reelin MSNs alone, as measured via the AFI (Fig. S9D). Newly formed tendons contained well-organized, abundant collagen fibers with a larger average diameter (Fig. 7G). MRI sagittal T2-weighted imaging confirmed that regenerated tendons in the Reelin slow-release MSNs group exhibited low-intensity, homogeneous signals with clear and smooth contours (Fig. 7H). Biomechanically, tendons treated with Reelin slow-release MSNs showed higher stress at failure and modulus values (Fig. S9E, F). In rabbits, Reelin slow-release MSNs combined with TSPC transplantation enhanced functional tendon regeneration in Achilles tendon transection model (Fig. S9G–J). Collectively, these results show that Reelin-loaded MSNs promote tendon regeneration in mice and rabbits, underscoring their translational potential. Overall, this study identifies lymphatic signaling hub in tendons, where LEC-secreted Reelin promotes TSPC stemness through Notch1–Srebp2-mediated cholesterol metabolism. Functionally, we highlight the essential role of this lymphatic hub in tendon regeneration (Fig. S10).

Methods
Animals
All experimental procedures performed in this study were following animal welfare ethical guidelines and approved by the Animal Use and Care Committee of Peking University (#LA20221488) and Animal Use and Care Committee of Chinese Institute for Brain Research (#CIBR-IACUC-051), in accordance with the governmental regulations of China. Sex was not considered as a biological variable in this study. C57BL/6 (6–8 weeks) and BALB/c nude (8–10 weeks) mice were purchased from Charles River Laboratory (China). Vldlr-CreETR2 mouse line was generated at the Chinese Institute for Brain Research (CIBR) Genetic Manipulation Core. Reln−/− mice (T009126) and Relnfl/fl mice (T012926) were purchased from GemPharmatech (China), Prox1-CreERT2 mice (Jax#022075) and tdTomato mice (Jax#007914) were obtained from the Jackson Laboratory (USA). Prox1-CreERT2 mice were breeding with tdTomato mice and Relnfl/fl mice to get Prox1-CreERT2; tdTomato and Prox1-CreERT2; Relnfl/fl mice. Rabbits (4 months) were purchased from Beijing Long’an Experimental Animal Breeding Center.

Generation of Vldlr-CreETR2 mice
Vldlr-CreETR2 mice were generated by homologous recombination at the endogenous Vldlr locus using the CRISPR/Cas9 system. Briefly, we inserted P2A-CreERT2 before the stop codon TGA of the Vldlr gene. The sgRNA synthesis template was obtained by PCR and purified using the Qiagen MinElute kit with the T7 promoter. The sgRNA sequences were designed as 5′-GACTTGGAATCCCCTTTTGA-3′ and 5′-GGGGCTCAAGGGTTACAGAT-3′, and CreERT2 was amplified from addgene13777 by PCR. A mixture of donor DNA plasmid, sgRNA and Cas9 mRNA were microinjected into fertilized zygotes from C57BL/6J females. Injected zygotes were implanted into oviducts of pseudopregnant ICR female mice. The genotyping primers were 5′-GACTTGGAATCCCCTTTTGA-3′, 5′-GGGGCTCAAGGGTTACAGAT-3′, and 5′-ATCAGCATTCTCCCACCATC-3′. Any potential off-target effects were bred out by backcrossing to C57BL/6 mice for at least five generations before the mice were used for experiments.

Antibodies and reagents
The details for the reagents and antibodies used in this study were listed in Supplementary Table 1 and Table 2.

Plasmid construct and transfection
The pCMV6-NICD-HA plasmid was a gift from Peking Union Medical College, Beijing, China. The pCDN3.1-RELN plasmid was constructed by Ruibiotech (China). The pCMV6-DAB1-DDK plasmid and its site-directed mutants were constructed using Gibson assembly technology52. Plasmid transfection assay was carried out using the VIGOROUS Transfection System kit (#T001, Vigorous Biotechnology, Beijing, China). For siRNA transfection, cells were transfected with riboFECT CP Transfection Kit (#166T, RiboBio, China).

Single-cell RNA sequencing
For scRNA-seq sequencing, tendon tissues were harvested from the injury sites of Achilles and Patellar tendons in neonatal (P3) and adult (6-week-old) C57BL/6J mice after 7 days injury. The same age mice were used as control (n=10 per group, pooled for sequencing). After being washed twice with pre-cooled Phosphate-Buffered Saline (PBS), the collected tissues were cut into 0.5 mm pieces and then digested with 3 mg/mL type I collagenase (Merck, Germany) and 4 mg/mL dispase (Merck, Germany) for 30 min at 37°C with shaking. After removing erythrocytes with red blood cell (RBC) lysis buffer (Genview, China), the cells were harvested by centrifugation at 450 × g for 15 min at 4°C, and sequentially filtered through 70 μm cell strainers. The isolated cells were harvested for library preparation. The library was generated using 10× Genomics Single-Cell V2 Library Construction (Genenergy, Shanghai, China). High-throughput sequencing (Illumina HiSeq PE125/PE150) raw image files were processed by CASAVA base calling to convert into sequenced reads, which were stored in FASTQ format. Mapped RNA-seq reads to the mouse genome (GRCm38) by HISAT2. Illumina TruSeq SBS-Kit v2 (200 cycles). Illunina’s bcl2fastq v1.8.4 software (Illumina, San Diego, CA) was used to convert the resulting base calling (bcl) to FASTQ files. Mapping RNA-seq reads on the mouse genome were performed using DNASTAR Lasergene v15.0 (DNASTAR, Madison, WI) and reported using FPKM (fragments per kilobase of exon per pillion fragments mapped). Genes term with a P-value <0.05 and fold change >2 were considered statistically significant. The enrichment of gene clusters and pathways based on different genes between two groups were performed using Gene Set Enrichment Analysis (GSEA).

Analysis of scRNA-seq data
For the scRNA-seq data analysis, Seurat (version 3.0.2) was used to normalize the raw counts; Cells with less than 200 genes and mitochondrial gene percentages >15% were removed. After quality control, 25,423 genes from 20,326 neonate cells and 8159 adult cells were selected for further analysis. The filtered batches were integrated using Harmony method. And the scDblFinder function was used to remove doublets. Filtered cells were subjected to dimensionality reduction and clustering by performing RunPCA and RunUMAP functions with dims 1:30. Annotation of cell clusters was based on the most enriched genes and marker genes. For tenogenic subcluster analysis, predefined sets of marker genes identified from published literature were used to identify TSCs, TPCs, and Tenocytes subclusters. Geneset scores were calculated using the Seurat function AddModuleScore. The genesets were provided in the source data file. Then CytoTRACE was also used to compare cellular differentiation status of three tenogenic related cell subsets. To quantitatively evaluate the degree of differentiation across cell clusters, a predefined set of stemness-related genes was used to calculate the stemness. The stemness score was calculated by Seurat function AddModuleScore. The stemness-related geneset was provided in the source data file. Next, monocle3 was used for estimate a pseudotemporal ordering of tendon clusters. And the TSCs subcluster was assigned the root node of the trajectory. The differential genes and enriched clusters among different groups were analyzed by gene ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment analysis. For cell-cell communication analysis, CellChat and CellPhoneDB were employed.

For human tendon samples analysis, one healthy and three pathologic tendon samples at 3 days, 10 days and 12 weeks post-injury were obtained from published datasets (SRA: PRJNA975881, PRJNA976191). The comprehensive donor information was summarized as follows: all samples were obtained from donors aged between 28 and 49 years, with or without tendon injury. Detailed sample informations, including exact age, sex, disease status, and precise time points for sample collection, were presented in Fig. S3C. We performed a re-analysis using the Seurat package. Cells were filtered out with less than 200 expressed genes or mitochondria percentages greater than 20%. The variability across batches were corrected by the Harmony package. PCA was constructed based on the scaled data for the top 2000 high variable genes. And the top 10 principal components were used for UMAP constructions. The cell–cell communication analysis was performed by CellChat and CellPhoneDB.

Bulk RNA-seq analysis
For bulk RNA-seq analysis, Vldlr+ and Vldlr− tendon cells from mice were sorted by flow cytometry and the RNA were extracted using Trizol (#15596026CN, Invitrogen, USA). One microgram of total RNA were used to prepare the sequencing library. Then 2×100 paired-end sequencing in fast run mode was performed using the HiSeq 2500 and Illumina TruSeq SBS-Kit v2 (200 cycles). Illunina’s bcl2fastq v1.8.4 software (Illumina, San Diego, CA) was used to convert the resulting base calling (bcl) to FASTQ files. Mapping RNA-seq reads on the mouse genome were performed using DNASTAR Lasergene v15.0 (DNASTAR, Madison, WI) and reported using FPKM (fragments per kilobase of exon per pillion fragments mapped). Genes term with a P-value <0.05 and fold change >2 were considered statistically significant. The enrichment of gene clusters and pathways based on different genes between two groups were performed using Gene Set Enrichment Analysis (GSEA).

Tamoxifen treatment for genetic lineage tracing
Tamoxifen (#10540-29-1, Sigma-Aldrich, USA) was freshly prepared by first dissolving in 100% ethanol and then suspended in corn oil to a final concentration of 20 mg/mL. For tamoxifen induced genetic lineage tracing, Prox1-CreERT2 mouse lines were used. To induce Cre activity, tamoxifen was intraperitoneal injected at a dose of 7.5 μL/g body weight for three consecutive days. In all the experiments, tamoxifen injections were performed in both Cre+ and Cre− mice. At the indicated time, mice were euthanized by CO2 asphyxiation, and tissues were collected for analysis.

Tendon transection animal model
For mouse tendon transection model, postnatal 3 days (P3) and 6–8 weeks old C57BL/6 mice, P3 old Reln−/− mice and Prox1-CreERT2, Relnfl/fl mice were used in this study. Briefly, the Achilles tendon was resected in the middle part. For the regeneration groups, the UV photocuring GelMA with or without TSPCs, Reelin pretreated TSPCs, Reelin, MSNs and Reelin loaded MSNs were grafted into the defect area. Then the incision was suture and bandaged. The mice were provided with unrestricted access to a standard diet and water with free cage activity at constant temperature with a 12 h dark–light cycle. Four weeks postoperatively, the samples were harvested for further investigation (n=5).

To verify the lymphatic vessels expansion after injury, VEGFR3 neutralizing antibodies were used to determine whether this lymphatic vessel expansion was dependent on VEGFR3. Briefly, 100 μg VEGFR3 neutralizing antibody was injected into the footpads of Achilles tendon resection mouse mode every 2 days. The PBS injection ones were used as control. The samples were harvested 7 days postoperatively for further investigation (n=5).

For rabbit tendon regeneration, 4 months old rabbits were used. The Achilles tendon was resected in the middle part. For the regeneration groups, the UV photocuring GelMA combined 4×106 TSPCs with or without Reelin loading MSNs was grafted into the defect area. The rabbits were provided with unrestricted access to a standard diet and water with free cage activity at constant temperature with a 12 h dark–light cycle. The samples were harvested after 2 months for further investigation (n=5).

Tendon overuse mouse model
For tendon overuse model establishment, 6–8-week-old Prox1-CreERT2;tdTomato mice were underwent a totally 4-week exercise intervention. Briefly, the intervention start-speed was established at 15 m/min. Exercise intervention consisted of 1 h of running per day, 5 days per week. After 2 weeks exercise, 20 μl 1% type I collagenase was injected around the Achilles tendon. The Achilles tendons samples were collected two weeks later for the lymphatic distribution analysis.

Indocyanine green (ICG) near-infrared (NIR) lymphatic imaging
Local lymphatic drainage in the Achilles tendon was determined by ICG NIR lymphatic imaging. Briefly, 20 μl of ICG (#I2633, Sigma-Aldrich, USA) at 0.1 mg/mL was injected intradermally into the mouse footpad about 2 mm distal to the calcaneus. ICG imaging of mouse footpads and legs was collected by IVIS Spectrum Small Animal In Vivo Imaging System (PerkinElmer, USA). The signal intensity in the footpad was recorded immediately after ICG injection as the initial signal intensity. ICG imaging was collected again 0.5 h, 1 h, 2 h, 6 h, and 24 h after ICG injection. The images were analyzed using Living Image software. ICG clearance was quantified as the percentage of reduced ICG signal intensity in the footpad 24 h after injection relative to the initial.

The lymphatic drainage tracer Evans blue injection
To investigate the lymphatic drainage around Achilles tendon, 20 μL 0.5% Evans blue (G1810, MW 0.96 kDa, Beijing Solarbio Science & Technology Co., Ltd., China) was injected slowly into the mouse footpad about 2 mm distal to the calcaneus of Prox1-CreERT2;tdTomato mice. Then the needle was left in place for approximately 5 min to prevent backflow. The samples were collected for tissue section preparation at 0.5, 1, 2, 6 and 24 h after Evans blue injection.

ELISA assay
The tendon tissues protein was collected by using high-throughput homogenizer. The Reelin ELISA assay kit (#JL33224, Jonln, China) was used to detect the reelin concentration according to the manufacture’s instruction. The OD value was measured at a wavelength of 450 nm.

TSPCs subcutaneously transplantation in nude mice
TSPCs (1×106 cells) with or without 1 μg/mL Reelin pretreatment were seeded within GelMA scaffold, and then implanted subcutaneously into the dorsum of BLAB/c (8-week-old) nude mice. Four weeks later, the implants were harvested for further study.

Achilles functional index analysis
For Achilles functional index (AFI) analysis, a restrictive roadway (59.4 cm long and 42 cm wide) was set using papers as previous report53. Then the mice footprints using red inkpad were recorded and scanned (Epson Scanner, Japan). The following footprints’ parameters including print length (PL), toe spreading length (the distance between the first and fifth toes, TS), and intermediary toe spreading length (the distance between the second and fourth toes, IT) were measured. The footprints dimension factors including print length factor (PLF), toe spreading length factor (TSF), and intermediary toe spreading length factor (ITF) were acquired by using the following equations: PLF=(NPL−EPL)/EPL, TSF=(ETS−NTS)/NTS, ITF=(EIT−NIT)/NIT. Finally, AFI was calculated according to the equation: AFI=74(PLF)+161(TSF)+48(ITF)−5.

Histological staining
For H&E staining, the histological sections (5 μm) were prepared through a microtome and subsequently stained with hematoxylin and eosin following the manufacturer’s instruction (G1120, Beijing Solarbio Science & Technology Co., Ltd., China).

For Sirius Red staining, the section was stained with Iron Hematoxylin Staining Solution for 5 min. Then it was stained with Sirius Red Staining Solution for 30 min. Then the images were captured with fluorescence microscope (Olympus VS120, Japan). The collagen content was calculated as follows:
Collagen fraction cover area (%) = (Cover area of color fraction / Total area) × 100%

Modified Masson’s trichrome staining (G1346, Beijing Solarbio Science & Technology Co., Ltd., China) was performed to examine the maturity of the collagen fibers. Briefly, the section was incubated in Mordant Solution in 57–60°C incubator for 1 h. Then the section was stained with Celestite Blue Solution, Mayer Hematoxylin Solution, Ponceau-Acid Fuchsin Solution, Phosphmolybic Acid Solution and Aniline Blue Solution for 3–5 min sequentially. The images were acquired by Olympus fluorescence microscope.

Tenogenic differentiation induction
The tenogenic differentiation was induced by a tenogenic differentiation medium, which contained growth medium supplemented with 10 ng/mL TGF-β1 (Peprotech), 10 ng/mL GDF-5 (R&D System), and 0.05 mM l-ascorbic acid 2-phosphate. After 21-day induction, Sirius Red and Masson’s trichrome staining were applied to evaluate tenogenic differentiation capacity, respectively.

Immunofluorescence staining
For immunofluorescence staining, cells were fixed in 4% PFA for 15 min at room temperature. For tendon tissues, frozen histological sections (20 μm per slice) were prepared. The samples were permeabilized in 0.2% (v/v) Triton X-100 (9002-93-1, BioRuler, USA) for 15 min. Then the samples were incubated with 5% goat serum (ZSGB-BIO) for 1 h. Later the samples were incubated with primary antibody overnight at 4°C followed by the secondary antibody incubation at room temperature for 1 h. Nuclei were counterstained with DAPI (#C0065, Beijing Solarbio Science & Technology Co., Ltd., China). Confocal microscopic images were acquired with a laser-scanning microscope (Leica TCS SP8, Germany) and then processed with Leica Application Suite X.

The construction of GelMA scaffold
GelMA was purchased from Yongqinquan Intelligent Equipment Co., Ltd. (#EFL-GM-60, Engineering for Life, China). Briefly, GelMA was mixed with 5% photoinitiator in a 55°C water bath. Hydrogel solution was passed through a 0.22 μm filter, and mixed with cell pellet digested with trypsin–EDTA solution (#25200-072, Gibco, USA) and Reelin (#YBC775Mu01, YBio Biotechnology Co., Ltd, China). After curing under 405 nm light (#EFL-LS-1601-405, Engineering for Life, China) for 15 s. Cells or MSNs were uniformly encapsulated in GelMA. All hydrogels were completely crosslinked.

Scanning electron microscope (SEM)
The tissues were pre-fixed in 2.5% glutaraldehyde in PBS (pH 7.4) at 4°C for 12 h. Then the samples were dehydrated in a graded series of ethanol (50, 70, 80, 90, 95, and 100%), critical-point dried, and sputter-coated with gold for 2 min at 20 mA. The microstructure of GelMA and regenerated tendon tissues were examined using SEM (Hitachi SU8020, Japan) at 10 kV.

Transmission electron microscopy (TEM)
To detect collagen fibril diameters and alignment, the regenerated tendon samples were prepared following a standard procedure for TEM analysis. Briefly, the samples were fixed with 2.5% glutaraldehyde for 12 h at 4°C. Then they were fixed with 1% OsO4 for 1.5 h. Next, the samples were dehydrated by a graded series of acetone (30, 50, 70, 80, and 90%) for about 15 min at each step. Then they were treated with 100% acetone two times, 10 min per times. The specimens were placed in order into 1:1 mixture of absolute acetone and resin mixture for 3 h, and into absolute acetone-resin mixture overnight. After that, samples were placed into fresh resin and heated at 70°C at least for 9 h, and were sliced by LEICA EM UC7 ultramicrotome. Finally, samples were imaged by Hitachi Model H-7650 TEM. Counting and measurement of collagen diameter were processed by marking the fibers individually in ImageJ 1.52a software.

TESOS clearing, immunolabelling and three-dimensional image acquisition
For tissue clearing, we modified our previous reported protocol to better visualize lymphatic distribution in tendons24. Briefly, the mice were perfused with PBS and 4% PFA. Tendons were fixed in 4% PFA on the shaker overnight. Then they were decalcified in 20% EDTA-Na (pH 7.0) at 37°C on a shaker for 1 day. Then the samples were decolorized with the Quadrol decolorization solution, gradient tert-butanol (tB) delipidation solutions for 1–2 days, respectively.

For immunolabelling, the pretreated samples were blocked in 5% normal donkey serum (#017-000-121, Jackson ImmunoResearch) for 1 day at 4°C and incubated with the primary antibody at 4°C for 4 days. After primary antibody incubation, the samples were washed five times with PBS buffer for 4 h, and then incubated with secondary antibodies at 4°C for 3 days. Finally, the samples were washed five times over 6 h before clearing. Finally, samples were immersed in the BB-BED medium 47% (v/v) benzyl benzoate (BB), 48% (v/v) bisphenol-A ethoxylated diacrylate Mn 468 (BED) and 5% (v/v) Quadrol and then is supplemented with 2% (w/v) Irgacure 2959 as the UV initiator at room temperature for at least 1 day until transparency was achieved.

For three-dimensional image acquisition, the Leica SP8 confocal microscope equipped with resonant scanner (8000 Hz) and a piezo sample stage (Leica) were used. A magnetic kinematic base (KB25/M or SB1/M, Thorlabs) was used for mounting and transferring samples. Following alignment, embedded samples were mounted on the microtome and sectioned (5 μm/cut) to expose the sample’s top surface. After sectioning, the surface was dropped with a BB PEG immersion medium and imaged. The imaging depth of each imaging z-stack is determined mainly by the objective’s working distance.

Magnetic resonance imaging
The regenerated Achilles tendons were measured using an 8-channel surface coil at 3.0 T (Philips Ingenia, Philips Healthcare, Netherlands). Acquisition parameters for sagittal T2 were as follows: Turbo spin echo, repetition time = 3104 ms, echo time = 80 ms, field of view = 90 × 110 mm, matrix size = 384 × 384, slice thickness = 1 mm.

Mechanical property analysis
Mechanical property was conducted using an Instron tension/compression system with Fast-Track software (Model 5969, Instron, Canton, MA). The Achilles complexes harvested at 4 weeks postoperatively were wrapped in gauze soaked in saline. Before testing, the Achilles complexes were gradually thawed at room temperature, and all surrounding soft tissues were rigorously removed. Measurements of the tendon cross-sectional area were performed using aqueous rapid curing alginate dental impression paste. The complex was then fixed to custom-made clamps using high strength thread. After applying a preload of 0.1 N, each complex was cyclically elongated between 0 and 0.5 mm for 20 cycles at 5 mm/min in advance. The load–elongation behavior of the complex and failure modes was recorded. The structural properties of the complex were calculated by modulus (MPa) and stress at failure (MPa).

Stem cell isolation and culture
Primary TSPCs were isolated from P3, 6–8-week-old mice and 2–4 month-old rabbits according to previously established procedure. Briefly, the harvested Achilles tendon was cut into small pieces and digested in 3 mg/mL Type I collagenase (Merck) and 4 mg/mL dispase (Merck) at 37°C for 30 min to get the single cell suspension. After centrifugation at 400 × g for 5 min, cells were resuspended and cultured in low-glucose Dulbecco’s modified Eagle’s medium (DMEM, Gibco) supplemented with 20% fetal bovine serum (FBS, Gibco) and 100 U/mL penicillin/streptomycin (Hyclone). After growing to 80–90% confluence, cells were passaged with fresh medium. The passage 2–4 TSPCs were used for further experiments.

For Vldlr+ and Vldlr− TSPCs isolation, tendon cells were firstly isolated from Vldlr-CreERT2;tdTomato mice after 7 days of tamoxifen induction at P3–P5. Then the cells were stained with COL1α1 antibody, followed by the FITC-conjugated secondary antibody staining. Then the Vldlr+ and Vldlr− TSPCs among COL1α1 positive cells were sorted by BD Aria SORP (BD Biosciences, USA).

Cell viability staining
For cell viability analysis, the live/dead cells were stained according to the manufacturer’s protocol (Beijing Solarbio Science & Technology Co., Ltd., China). Briefly, cells were incubated 30 min with 2 μM calcein-AM and 4.5 μM Propidium iodide dissolved in PBS. Then the images were acquired with Olympus fluorescence microscope.

Western blotting
Total proteins from cell lysates were harvested by RIPA Buffer with Protease/Phosphatase Inhibitor Phenylmethylsulfonylfluoride (PMSF) (#P0100, Beijing Solarbio Science & Technology Co., Ltd., China). Nuclear and cytoplasmic proteins were extracted with Nuclear Protein Extraction Kit (#P0027, Beyotime, China). Proteins were separated by 4–15% SDS–polyacrylamide gel electrophoresis (SDS-PAGE), and then transferred to polyvinylidene difluoride membranes (PVDF) and blocked in 5% nonfat milk. The membranes were probed with corresponding primary antibodies overnight at 4°C. Then the secondary antibodies were incubated for 1 h at room temperature. After the membranes were enhanced with SuperSignal West Pico Chemiluminescent Substrate (#34077, Invitrogen, USA), The bands were detected by Imaging System (Bio-Rad, USA). GAPDH antibody was used to quantify the amount of loaded protein. The uncropped and unprocessed blots are provided in Source Data file.

Co-immunoprecipitation (Co-IP) analysis
For co-immunoprecipitation assay, cells were lysed with lysis buffer (#9803, Cell Signaling Technology, USA) for 30 min on ice. Then the cell lysate was centrifuged with 12,000 × g for 10 min and supernatant was incubated with antibodies overnight at 4°C, followed by Protein A/G agarose (#sc-2003, Santa Cruz Biotechnology, USA) incubation for 2 h at 4°C. The Anti-HA Rabbit mAb was used for NICD-HA protein immunoprecipitation. The Anti-SREBP2 Rabbit mAb, Anti-DAB1 Rabbit mAb, and Anti-VLDLR Rabbit mAb were used for SREBP2, DAB1 and VLDLR protein immunoprecipitation, respectively. The precipitated protein was collected by centrifugation (4000 × g for 5 min at 4°C). After washing, precipitated proteins were denatured (100°C for 5 min) and separated by SDS-PAGE, then detected by immunoblotting following the western blotting protocol.

RT-qPCR
For RT-qPCR analysis, tendon tissues or cultured cells were harvested using Trizol to obtain RNA. Then the 1 μg RNA was reverse-transcribed into cDNA with the ReverTra Ace® qPCR RT Master Mix Kit (#420600, Toyobo, Japan). Then 0.5 μg cDNA per sample were used to perform qPCR using gene-specific primers and SYBR Green Premix PRO Taq HS Quantitative PCR Kit II (AG11719, Accurate Biology, China) on 7900HT Fast Time PCR. The primers were listed in Supplementary Table 3.

SiRNA knockdown
To knock down Dab1 or Srebp2 in TSPCs, SiRNA transfection was performed according to the manufacturer’s instructions (C10511-05, RiboBio, China). Briefly, the riboFECT™ CP was added into the low-glucose Dulbecco’s modified eagle medium (L-DMEM) without penicillin-streptomycin solution. The siRNA oligonucleotide sequences used were listed in Supplementary Table 4. In 21-day tenogenic differentiation induction assay, the transfection complex was added every 3 days with the induction medium change. On day 2 and 21 post-transfection, Srebp2 and Dab1 mRNA levels were quantified by RT-qPCR to assess knockout efficiency.

Synthesis of surface modified mesoporous silica nanoparticles
Mesoporous silica nanoparticles (MSNs) were synthesized via the sol-gel method within an oil-water system36,37. In a typical synthesis, a water phase comprising 60 mL was prepared by blending 24 mL of cetyltrimethylammonium chloride (CTAC) (25% aqueous solution), 0.20 g of triethylamine (TEA), and 736 mL of MilliQ ultra-pure water at 60°C for 1 h. Subsequently, 40 mL of the oil phase, consisting of 37.5 mL of chlorobenzene and 2.5 mL of tetraethyl orthosilicate (TEOS), was gently introduced into the bottom of the water phase. The resulting mixture was stirred at 60°C with a stirring speed of 500 g for 12 h. The solid samples were then subjected to centrifugation at 18,522 × g for 15 min and washed thrice with ethanol. Finally, the MSNs products underwent calcination at 550°C for 5 h to eliminate the organic surfactant.

In the subsequent step, MSNs was functionalized with polyethyleneimine (PEI). Initially, 60 mg of MSNs was dispersed in 15 mL of MilliQ water, and the pH was adjusted to 10. Following this, 15 mL of N-(3-(trimethoxysilyl) propyl) ethylenediamine (THPMP) solution (56 mM) was added, and the mixture was stirred at 40°C for 2 h. The particles were then collected via centrifugation at 18,522 g for 15 min and washed thrice with water. Subsequently, the product was suspended in a PEI solution (10 mg/mL in 100 mM carbonate buffer, pH=9.6) and stirred for 4 h at room temperature. The final product underwent thorough washing with water and was air-dried at room temperature.

For the preparation of MSNs solution, nanoparticles were dispersed in PBS with a final concentration of 1 mg/mL with ultrasonic treatment. For Reelin protein loading in MSNs, 10 μL protein solution (200 μg/mL) was mixed with 10 μL silica nanoparticles (200 μg/mL) with gentle shaking for 24 h at 4°C.

Characterization of surface modified mesoporous silica nanoparticles
For MSNs characterization, the powder samples were dispersed in ethanol, dropped onto an aluminum foil piece, and affixed to a conductive carbon film. Following this, the samples were subjected to drying in a vacuum oven at 60°C for 12 h and subsequently cleaned with an Evactron 25/45 De-contaminator RF plasma cleaning system. Then the morphology and pores of MSNs were analyzed by SEM and TEM.

Nitrogen adsorption-desorption measurements were conducted at −196°C with a Micromeritcs Tristar ІІ system, before testing the samples were degassed at 100°C overnight on a vacuum line. The total pore volume was calculated from the amount adsorbed at a maximum relative pressure (P/P0) of 0.99. The Barrett–Joyner–Halenda (BJH) method was used to calculate the pore size of samples from the adsorption branches of the isotherms. The Brunauer-Emmett-Teller (BET) method was utilized to calculate the specific surface areas. Dynamic light scattering (DLS) was carried out at 25°C using a Zetasizer Advance Series-Pro from Malvern Panalytical Ltd.

Measurement of cholesterol
The cholesterol level was detected by Amplex Red Cholesterol and Cholesteryl Ester Assay Kit according to the manufacture’s instruction (#S0211M, Beyotime, China). Briefly, the cells were collected by centrifuged (500 × g for 5 min) and the supernatant was discarded. Then the buffer A was added to the sample and mixed well. The supernatants were collected by centrifugation (12,000 × g for 5 min at 4°C) for absorbance detection at 570 nm using multifunctional microplate reader (BioTek Synergy H1 Multimode Reader, Agilent, USA). The cholesterol level was quantified relative to the standard.

Cleavage under targets and tagmentation (CUT&Tag)
We performed chromatin immunoprecipitation with a NovoNGS CUT&Tag High-Sensitivity kit (N259-YH01, Novoprotein, China) following the manufacturer’s protocol. In brief, 100,000 cells were harvested, crosslinked with 1% formaldehyde, and permeabilized. The proteins DNA complex were then bound to ConA magnetic beads and incubated overnight at 4°C with anti-Notch1 antibody or IgG isotype control for immunoprecipitation. After subsequent incubation with secondary antibodies, the immunoprecipitated DNA was purified from beads and analyzed by qPCR. Primer sequences used can be found in Supplementary Table 5.

Statistical analysis
The quantitative data were presented using the mean ± standard deviation (SD). Statistical analysis was performed with GraphPad Prism 10. The comparison between two groups was analyzed by independent unpaired two tailed Student’s t tests, and comparisons among more than two groups were conducted using one-way analysis of variance (ANOVA) with Tukey’s multiple comparisons test. P<0.05 were statistically significant.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The authors perform a large number of studies that identify lymphatic endothelium as potentially important in tendon development and repair. Unfortunately studies are disjointed, many superficially performed and with lack of clarity. I would recommend strengthening the logic between individual experiments, retaking histologic images for clarity, rephrasing conclusions so as to more accurately represent the data, and focusing on the clear differences that exist between Reln KO and CKO tendons.\nSignificant gaps in logic exist in the analysis of Figure 1. Interactome analysis in Fig 1I shows high level of interaction between tendon cells and SMCs, Schwann cells, muscle cell and others. To focus on endothelial cells, and more specifically lymphatic endothelium, is not justified by this dataset. Moreover, this is a dataset comparing developing to juvenile intact tendons. To then conclude this has to do with tendon repair is essentially unfounded.\nCollagenase injury to tendons is an artificial methodology to induce injury. Recommend adding alternative tendon injury / overuse methods.\nMultiple injury models are used in later figures, or subfigures, but there is a real lack of clarity in terms of which models are used when. Moreover tendon transection differs widely from collagenase injury, making the combination of these two models difficult to interpret. All this makes review of the work challenging.\nExpression of Reln does not correlate well between single cell data in 2F and immunohistochemistry in oart H. Indeed a near absence of Reelin under control conditions is shown by IF staining.\nThe tendons are so different in the Relin KO mouse at baseline, that this appears to be a developmental defect and not a repair defect. Also it is unclear what \"Injury\" is in 3C, since it appears to be adipose tissue? Overall the histology is of poor quality and lacks clarity.\nSome histology is of such high magnification that is lacks value (e.g. Fig 4).", "reviewer_2": "NCOMMS-25-23311, Chen et al., Lymphatic - Stem Cell Crosstalk Controls Tendon Regeneration via Notch1-Srebp2 Mediated Cholesterol Metabolism\nLymphatic vessels drain tissue fluid and lipids and, also, traffic leukocytes. Recent publications have indicated contribution of the lymphatics in the intestinal, cardiomyocyte and hair follicle stem cell niche. In part, this paracrine function is taken care by lymphatic endothelial reelin (Liu et al. 2020, Nature).\nIn their manuscript, Chen et al. investigate Achilles and patellar tendon in postnatal and adult mice in homeostasis and upon injury. The authors use a variety of methods, such as scRNA sequencing, lineage tracing, cell type-specific gene deletions, and in vitro assays of cell differentiation and signalling. Based on these, Chen et al. claim that they have “identified lymphatic vessels as key components of tendon regenerative niche”. Further, they state that this effect is caused by lymphatic endothelial reelin-induced NOTCH-signalling in the tendon stem cells.\nThe study covers a huge area from identification of the essential cell types in tendon regeneration to molecular mechanisms of tendon stem cell niche and tendon regeneration. However, the provided data lacks many essential controls and the motivation for the sequence of experimentation is not always obvious to the reader. Especially, it is not clear whether there are lymphatic vessels in the tendons. My major comments are listed below.\n1. This manuscript and Tempfer et al (2015, Histochem. Cell Biol.) and Tong et al. (2023, J. Orthop. Translat.) describe the existence of lymphatic endothelial markers in the tendon tissue. However, these markers are also expressed in other cell types such as muscle cells, neurons and blood vessels.\nIn this manuscript, Chen et al. characterize these structures mostly with Prox1-TdTomato reporter mice. The provided images do not allow evaluation of whether the depicted structures are lymphatic vessels. For example, the structures shown in Fig. 2B-C resemble rather blood vessels or neurons than lymphatic vessels. Also, in figures 2H-I, lymphatic markers are displayed at the edge of the tissue, possibly reflecting a staining artifact, and do not show any vessel-like morphology. In some of the figures, the staining is dot-like, for example, Fig. 2A, 2H, S2G-I, and S5N. VE-cadherin staining, which should exclusively highlight lymphatic endothelial and endothelial junctions, shows a homogenous signal in the whole cell (Fig. S2D) and PROX1, which should be nuclear is, does not display nuclear localisation (Fig. 2H).\nTo solidify the existence of lymphatic vessels in the tendon and the basis for this manuscript the authors should:\n1.A. Provide high-resolution overview and Zoom in images to show the lymphatic vessel lumen (optical orthogonal sections or 3D stacks). Also, the localization of VE-cadherin and/or CD31 at cell junctions, and Prox1 in the nucleus of the same cells should be shown.\n1.B. Provide a functional draining assay (a tracer) to show that these structures drain the tissue fluid (the tracer should be seen inside the lymphatic vessels). Several time points should be included.\n1.C. To show that the growth of these structures in development and upon injury/inflammation is dependent on the VEGF-C to VEGFR3 signalling (the key growth factor signalling route of lymphatics), authors should use one of the established tools (blocking antibodies, VEGF-C ligand traps, or conditional mouse strains).\n2. Secreted reelin is expected to stay in the vicinity of the lymphatic vessels, suggesting that tendon stem cells should be located close to the lymphatic vessels. Authors should provide high-resolution double staining for the lymphatics and the tendon stem cells and quantify the distance of the stem cells from lymphatic in homeostasis and injury/inflammation (three independent experiments are required). Also, authors should add a proliferation marker to quantify, whether stem cells in the vicinity of the lymphatics have a higher proliferative index than the ones further away.\nThe authors state that the lymphatics expand upon injury/inflammation. Would this mean that only in these conditions do the lymphatics form the niche or, also, during homeostasis?\n3. Authors use Prox1CreERT2 to delete conditional allele of reelin. Authors should show that the reelin signal is lost in lymphatics upon tamoxifen induced reelin deletion.\n4. If reelin affects the growth and/or maintenance of lymphatics (Fig. S4G-H and S5N) it is difficult to judge whether the effect of reelin deletion on tendon stem cells is directly caused by the reduced reelin amount or by the absence of other lymphatic paracrine signals in vivo. Authors should discuss this in their manuscript.\n5. The authors use lymphadenectomy as described on rows 642-647. Here the authors execute “a combined popliteal and superficial inguinal lymph node excision”. Does this mean that the lymph nodes and the collector vessel in between are removed? Then the tendon lymphatics are probably still in place but the lymphatic drainage is disrupted. Authors should show, what happens to tendon lymphatics upon lymphadenectomy (are they still present in the tendon), does lymphadenectomy alter tendon drainage (see point # 1B) and would loss of drainage affect the paracrine function of the lymphatics (for example, acute reelin expression by the tendon lymphatics).\n6. Also, in other places, the methodology is not detailed enough and does not allow replication of the experiments. For example:\n6.A. In cell culture experiments authors silence Dab1 or Srebp2 with an siRNA approach (Fig. 6C and G, S7C, S8A-D) and validate the Dab1silencing with RT-PCR at 48 hours (Fig. S7B). Indeed transfected siRNAs are thought to cause downregulation of mRNAs for 2-3 days after which the target mRNA levels start to increase toward normal levels. In this context, it is difficult to understand why the silencing of Dab1 or Srebp2 has such a drastic effect on tenogenic differentiation in 21-day protocol. Authors should show the knockdown efficiency of Srebp2 mRNA at early and late time points and Dab1 mRNA levels at later time points as well. If the Dab1 or Srebp2 mRNA levels increase at a later time point, how do the authors explain the attenuation of differentiation?\n6.B. Authors should describe more carefully the protocol for the isolation of the tendon cells, for example, for scRNA sequencing.\n7. Reelin is a ligand of Vldlr. Authors, state “Reelin treatment could increase the total cholesterol level in TPSCs“ (rows 400-401 and Fig. 6L). It is not obvious, how reelin binding would increase cholesterol (apoE-lipoprotein) uptake since the increase in Vldlr mRNA levels upon reelin treatment is fairly moderate (Fig. S7A). Authors should investigate the molecular mechanism of increased cholesterol uptake. Further, does reelin compete with apoE lipoproteins on binding the Vldlr or are there synergistic effects (for example, co-binding)?\n8. The data on Notch1 signaling (NICD formation) consists largely of representative images. Authors should show quantifications of three independent experiments at least for Fig. 6E-G. Also, authors should use arrows in Fig. 6F to indicate which bands represent Notch1 and NICD. What are the bands in the lowest “row”? Authors should also include a control for the fractionation (nuclear and cytoplasmic marker for both fractions).\n9. The manuscript text needs proofreading and elaboration of some of the parts to allow easy digestion of the key messages. As an example:\n9.A. Rows 498-500: lineage tracing does not indicate the contribution of the Vldlr gene/protein but, rather, traces the Vldlr positive cells.\n9.B. Rows 487-491: What do the authors mean by “ Previous studies have identified various ligands in lymphatics, such as Ntn1, IL33 and VEGF-C (14, 37). Our study adds to this body of knowledge by identifying additional tendon lymphatic genes, including Reelin, and Rspo3, warranting further exploration into their roles in tendon development and regeneration.” VEGF-C is a ligand, produced by other cell types, which binds VEGFR3 on lymphatic endothelia, whereas reelin is a ligand produced by lymphatic endothelia.", "reviewer_3": "In the manuscript, a functional crosstalk between lymphatic vessels and tendon stem/progenitor cells (TSPCs) was found to play a critical role in tendon regeneration. Mechanistically, Reelin secreted by lymphatic endothelial cells (LECs) promoted phosphorylation of DAB1 through interaction with VLDLR in TSPCs. The activated DAB1 subsequently triggered a Notch1-SREBP2-mediated cholesterol metabolism pathway, which proved essential for maintaining TSPC stemness. This study reveals a novel lymphatic-tendon signaling axis and demonstrates its physiological relevance in tendon regeneration. The authors provided substantial experimental validation through rigorous in vitro and in vivo approaches. Although Reelin-DAB1-Notch1-SREBP2 signaling has been previously characterized in cardiac and intestinal tissues, this work represents the first elucidation of its mechanistic role in tendon biology. Given these findings, we recommend this manuscript for publication in Nature Communications, pending clarification of the following points:\n1. Discrepancies exist between the textual descriptions and annotations in Figure 5.\n2. Comprehensive documentation of human tendon samples is required, particularly regarding (a) donor demographics, (b) injury etiology, and (c) temporal parameters of sample collection post-injury.\n3. Methodological clarification: The collagenase-induced model (typically modeling chronic tendinopathy) and transection model (simulating acute injury) represent distinct pathological contexts. The authors should explicitly address how these models differentially inform their conclusions.\n4. Potential immune modulation: Given that (a) lymphatic signaling is intrinsically linked to immune regulation, and (b) observed alterations in macrophage markers between normal and injured tendons, the possible involvement of macrophage-mediated mechanisms in Reelin signaling warrants investigation."}, "author_response": "Point by point response\nEVIEWER COMMENTS\nReviewer #1 (Remarks to the Author):\n1. The authors perform a large number of studies that identify lymphatic\nendothelium as potentially important in tendon development and repair.\nUnfortunately studies are disjointed, many superficially performed and with lack\nof clarity. I would recommend strengthening the logic between individual\nexperiments, retaking histologic images for clarity, rephrasing conclusions so\nas to more accurately represent the data, and focusing on the clear differences\nthat exist between Reln KO and CKO tendons.\nResponse: We sincerely appreciate the reviewer’s valuable suggestions. To\nenhance the logical coherence between individual experiments, we have\ncarefully reorganized the sequence of figures and supplemented newly\ngenerated experimental data, thereby better elucidating the connections among\ndifferent experimental findings. Concerning the clarity of histological images,\nwe have replaced the images in Figure 3A, 3C, 4C, 5K, 7F and Figure S 9G\nwith higher-quality to ensure more clear visualization.\nFurthermore, we have refined the conclusions to more accurately reflect the\nexperimental data on Page 17: “In conclusion, this study establishes lymphatic\nsignaling as a critical mechanism in tendon regeneration. The\nlymphoangiocrine Reelin regulates TSPC stemness and tenogenesis through\nNotch1–Srebp2-mediated cholesterol metabolism. These findings not only\nprovide insights into tendon biology, but also introduce Reelin-loaded MSNs as\na promising therapeutic strategy. Future research should investigate other\nlymphangiogenic factors and metabolic pathways to optimize regenerative\ntherapies.”\n2. Significant gaps in logic exist in the analysis of Figure 1. Interactome analysis\nin Fig 1I shows high level of interaction between tendon cells and SMCs,\nSchwann cells, muscle cell and others. To focus on endothelial cells, and more\nspecifically lymphatic endothelium, is not justified by this dataset. Moreover, this is a dataset comparing developing to\njuvenile intact tendons. To then\nconclude this has to do with tendon repair is essentially unfounded.\nResponse: We greatly appreciate the reviewer’s valuable suggestion. We have\nsupplemented new experiments to compare the gene expression and signaling\ndifferences among neonate and adult tendons under both normal and injured\nconditions, using single-cell RNA sequencing analyses. The interactome\nanalysis revealed that the interactions between tendon cells and LCEs was\nsignificantly enhanced in injured tendons relative to control ones. Specifically,\nthe interaction between LECs and TSCs was strengthened post-injury, which\nindicates the crucial role of LECs in tendon repair. These bioinformatics\nanalysis hints provide a strong rationale for further investigating whether LECs\nare involved in tendon regeneration. The newly generated results and\ncorresponding revisions have been incorporated on Page 4-5: “To identify the\nmolecular drivers of tendon regeneration and maturation, we compared gene\nexpression in cells from control and injured tendons of neonates and adults\nusing single-cell RNA sequencing (scRNA-seq, Fig.1A). The tendon-derived\ncells were classified into nine major cell clusters based on gene expression\nprofiles (Fig. 1B-1E, and Fig. S1A–S1C). Cluster 0 showed high expression of\ntenogenesis-associated genes, including Scx, tenomodulin (Tnmd), tenascin-C\n(Tnc), and fibromodulin (Fmod)[22]. To examine the mechanisms of tendon\nmaturation, we isolated tenogenic subset cluster 0 for uniform manifold\napproximation and projection–based clustering. Clusters were annotated using\nmarker gene expression and gene set scores derived from predefined\ncanonical stem cell, progenitor cell, and mature tenocyte genes (Fig.1F,1G, and\nFig. S1D). The cluster expressed stem cell markers lymphocyte antigen 6\ncomplex, locus A (Ly6a) and Thy-1 cell surface antigen, was annotated as\ntendon stem cells (TSCs). The cluster exhibited the highest progenitor cell-\nassociated gene set score, leading to its classification as tendon progenitor\ncells (TPCs). Another cluster with high expression of the ECM genes Fmod and\nthrombospondin-4 (Thbs4) for mature tendon cells was designated tenocytes\n(Fig. 1G and Fig. S1D). The stemness scores calculated based on the\nstemness-related gene set showed decreasing stemness scores from TSCs to\ntenocytes, which aligned with the subpopulation definitions (Fig. 1H).\nFurthermore, Monocle3 and CytoTRACE analysis confirmed differentiation\npath from TSCs through TPCs to tenocytes (Fig. 1I and 1J), suggesting the\nthree distinct TSPC subsets transited from neonatal to adult tissue.\nImmunofluorescence staining and gene expression on UMAP analysis both\ndemonstrated higher Ly6a and tubulin polymerization-promoting protein family\nmember 3 (Tppp3) levels in neonatal tendons ， while higher Scx and\nproteoglycan 4 (Prg4) levels in adult ones (Fig.S1E and S1F).\nThe environment niche, where stem cells resident in, plays a crucial role in\ndetermining cell maintenance and fate[23]. The cell-cell communication\nanalysis highlighted increased interactions between lymphatic endothelial cells\n(LECs) and tenogenic cells after injury compared with control one (Fig. 1K). To\ndefine this interaction mechanisms, we isolated tenogenic related subsets and\nendothelial cells for UMAP-based clustering and annotation (Fig. S1G- S1I).\nWe further examined the interaction between LECs with three subpopulations\nof tenogenic cells and the results revealed specifically increased interaction\nbetween LECs and TSCs subsets after tendon injury (Fig. 1L). These findings\nindicate that lymphatics in tendons are key regulators of stem cell fate and\ntendon healing.”\n3. Collagenase injury to tendons is an artificial methodology to induce injury.\nRecommend adding alternative tendon injury / overuse methods.\nResponse: Thank you for your valuable suggestion. We have replaced the\ncollagenase injury model with Achilles tendon overuse model. The results were\nadded on Page 6-7: “To model chronic tendinopathy, we induced Achilles\ntendon overuse. In this model, lymphatic expansion also occurred, as shown\nby light-sheet imaging of Prox1-positive vessels (Fig. S2L). Together, these\nresults demonstrate that both acute and chronic stress drive lymphatic\nexpansion in tendons.”\n4. Multiple injury models are used in later figures, or subfigures, but there is a\nreal lack of clarity in terms of which models are used when. Moreover tendon\ntransection differs widely from collagenase injury, making the combination of\nthese two models difficult to interpret. All this makes review of the work\nchallenging.\nResponse: Thank you for your insightful comment. We have revised all the\nresults part related with the tendon injury model. For the single-cell RNA\nsequencing analyses and tendon regeneration, the tendon transection model\nwas used. Following your suggestions on Point #3, the tendon overuse\nmodel and tendon transection model were used for the lymphatic distribution\nanalysis. The detailed methods were revised on Page 24: “For mouse tendon\ntransection model, postnatal 3 days (P3) and 6-8 weeks old C57BL/6 mice, P3\nold Reln-/- mice and Prox1-CreERT2, Reln fl/fl mice were used in this study. Briefly,\nthe Achilles tendon was resected in the middle part.”, and “For tendon overuse\nmodel establishment, 6-8-week-old Prox1-CreERT2; tdTomato mice were\nunderwent a totally 4-week exercise intervention. Briefly, the intervention start-\nspeed was established at 15 m/min. Exercise intervention consisted of 1h of\nrunning per day, 5 day per week. After 2 weeks exercise, 20 μl 1% type I\ncollagenase was injected around the Achilles tendon. The Achilles tendons\nsamples were collected two weeks later for the lymphatic distribution analysis.”\nThe results were also revised to address the difference of these two models on\nPage 6-7: “Next, to test lymphatic responses to tendon injury, we established\nan Achilles tendon transection model. Lymphatic vessels expanded significantly\nafter 7 days tendon injury in neonatal and adult mice, with increased Prox1-\nTdtomato-positive and LYVE1-positive lymphatic vessels (Fig. 2C, 2D and Fig.\nS2H, S2I). Quantitative analysis confirmed significantly elevated Prox1 and\nLyve1 mRNA expression after tendon injury (Fig. S2J). To confirm the specificity,\nwe treated injured tendons with VEGFR3-blocking antibody, which inhibited\nLYVE1-positive lymphatic vessel expansion (Fig. S2K). To model chronic\ntendinopathy, we induced Achilles tendon overuse. In this model, lymphatic\nexpansion also occurred, as shown by light-sheet imaging of Prox1-positive\nvessels (Fig. S2L). Together, these results demonstrate that both acute and\nchronic stress drive lymphatic expansion in tendons.”\nMoreover, the Figure legends were revised accordingly to clarify the models\neach experiment used.\n5. Expression of Reln does not correlate well between single cell data in 2F and\nimmunohistochemistry in oart H. Indeed a near absence of Reelin under control\nconditions is shown by IF staining.\nResponse: We appreciate your suggestions greatly. To better demonstrate\nthe expression of Reelin in lymphatic endothelial, we have re-conducted the\nimmunofluorescence staining and replaced the images shown in Figure 2G\nand 2H. The results were consistent with the single cell sequencing analysis,\nwhich were added on Page 7: “Next, we examined how lymphatics contribute\nto tendon healing. CellChat analysis of TSPCs and LECs identified Reelin (Reln)\nsignaling as the major pathway enriched after injury (Fig. 2E). Expression\nanalysis also showed high Reln levels in LECs (Fig. 2F). Reelin protein has\nbeen reported to promote cardiac repair[18]. Immunostaining showed Reelin\ncolocalization with the Prox1-tdTomato and LYVE1 in tendons. Reelin\nexpression was higher in neonatal tendons than in adult ones, which also\nincreased following tendon injury (Fig. 2G, 2H).”\nIn addition, as immunofluorescence staining indicated the expression of\nReelin in a thin tissue layer, we performed the analysis of Reelin level in\ntendons from the control and injury groups using an ELISA assay, and the\nresults have been added on Page 7: “Moreover, the ELISA assay verified the\nlevels of Reelin was significantly increased after tendon injury (Fig. S2M),…”\n6. The tendons are so different in the Relin KO mouse at baseline, that this\nappears to be a developmental defect and not a repair defect. Also it is unclear\nwhat \"Injury\" is in 3C, since it appears to be adipose tissue? Overall the\nhistology is of poor quality and lacks clarity.\nResponse: We appreciate your suggestions. To clarify the role of Reelin on\ndevelopment, we repeated the histological analysis of Reln conventional\nknockout (Reln-/-) mice and Reln conditional knockout mice (Prox1-CreERT2;\nRelnfx/fl) under physical condition. The results demonstrated that Reln-/- mice\ntendons revealed a certain degree of hypoplasia, while no significantly\ndifference were detected between the control and Prox1-CreERT2; Relnfx/fl mice\nunder physical condition. The following results were revised and in-\ncooperated to the results part on Page 7: “Their Achilles, patellar, and tail\ntendons appeared hypoplastic and more translucent compared with controls.\nAdditionally, histology showed reduced collagen density and fewer cells in Reln-\n/- mice tendons relative to controls (Fig. 3A and Fig. S4B).”and Page 8-9：“To\nspecifically assess the role of LEC-derived Reelin in tendon regeneration, we\nselectively deleted Reln in LECs using Prox1-CreERT2; Relnfl/fl (RelnCKO) mice.\nThe expression of Reelin secreted by LECs were dramatically reduced in\ntendon of Prox1-CreERT2; RelnCKO mice (Fig. S5A). Under baseline conditions,\nRelnCKO and control Achilles tendons showed no significant morphological or\nhistological differences (Fig.4A–4D).”\nWe apologize for the confusion caused by including the injury group in Fig. 3C,\nwhich was intended to demonstrate the successful establishment of the Achilles\ntransection model. To eliminate this confusion, we have deleted this group.\nThe histological images in Figure 3A and 3C have been replaced with higher-\nquality versions at lower magnification to enhance clarity and improve\nvisualization of key details.\nMoreover, the following sentences were added in the Discussion part on\nPage 14-15: “In Reln-/- mice, newly formed tendons displayed reduced\ncellularity, disorganized collagen fibers, and impaired mechanical properties.\nConsistent with prior reports of Reln-/- mice exhibiting neural defects and weight\ngain failure[42], we also observed tendon developmental dysplasia, reduced\nlymphatic formation in Reln-/- mice. A previous study noted reduced smooth\nmuscle cell coverage and irregular diameters in lymphatic collecting vessels in\nReln-/- mice, despite normal lymphatic capillaries[43]. Collectively, these\nfindings suggest that the mechanisms controlling lymphatic vessel formation\nand tendon development may differ by tissue microenvironment and could be\ninfluenced by nervous system defects or growth impairments in Reln-/- mice. To\ntest the direct role of LEC-derived Reelin, we deleted Reln in lymphatics using\nProx1-CreERT2 mice. The resulting RelnCKO mice exhibited impaired tendon\nregeneration despite normal tendon development and lymphatic formation,\nconfirming that lymphatic-derived Reelin specifically regulated tendon\nregeneration. In vitro assays further verified the critical role of Reelin in\nmaintaining TSPC stemness.”\n7. Some histology is of such high magnification that is lacks value (e.g. Fig 4).\nResponse: We appreciate the reviewer’s suggestions. The histological\nimages in Figure 3A, 3C, 4C, 5K, 7F and Figure S 9G have been replaced with\nhigher-quality versions at lower magnification to enhance clarity and improve\nvisualization of key details.\nReviewer #2 (Remarks to the Author):\nNCOMMS-25-23311, Chen et al., Lymphatic - Stem Cell Crosstalk Controls\nTendon Regeneration via Notch1-Srebp2 Mediated Cholesterol Metabolism\nLymphatic vessels drain tissue fluid and lipids and, also, traffic leukocytes.\nRecent publications have indicated contribution of the lymphatics in the\nintestinal, cardiomyocyte and hair follicle stem cell niche. In part, this paracrine\nfunction is taken care by lymphatic endothelial reelin (Liu et al. 2020, Nature).\nIn their manuscript, Chen et al. investigate Achilles and patellar tendon in\npostnatal and adult mice in homeostasis and upon injury. The authors use a\nvariety of methods, such as scRNA sequencing, lineage tracing, cell type-\nspecific gene deletions, and in vitro assays of cell differentiation and signalling.\nBased on these, Chen et al. claim that they have “identified lymphatic vessels\nas key components of tendon regenerative niche”. Further, they state that this\neffect is caused by lymphatic endothelial reelin-induced NOTCH-signalling in\nthe tendon stem cells.\nThe study covers a huge area from identification of the essential cell types in\ntendon regeneration to molecular mechanisms of tendon stem cell niche and\ntendon regeneration. However, the provided data lacks many essential controls\nand the motivation for the sequence of experimentation is not always obvious\nto the reader. Especially, it is not clear whether there are lymphatic vessels in\nthe tendons. My major comments are listed below.\nResponse: We sincerely appreciate the reviewer’s positive comments and\nvaluable suggestions.\n1. This manuscript and Tempfer et al (2015, Histochem. Cell Biol.) and Tong et\nal. (2023, J. Orthop. Translat.) describe the existence of lymphatic endothelial\nmarkers in the tendon tissue. However, these markers are also expressed in\nother cell types such as muscle cells, neurons and blood vessels.\nIn this manuscript, Chen et al. characterize these structures mostly with Prox1-\nTdTomato reporter mice. The provided images do not allow evaluation of\nwhether the depicted structures are lymphatic vessels. For example, the\nstructures shown in Fig. 2B-C resemble rather blood vessels or neurons than\nlymphatic vessels. Also, in figures 2H-I, lymphatic markers are displayed at the\nedge of the tissue, possibly reflecting a staining artifact, and do not show any\nvessel-like morphology. In some of the figures, the staining is dot-like, for\nexample, Fig. 2A, 2H, S2G-I, and S5N. VE-cadherin staining, which should\nexclusively highlight lymphatic endothelial and endothelial junctions, shows a\nhomogenous signal in the whole cell (Fig. S2D) and PROX1, which should be\nnuclear is, does not display nuclear localisation (Fig. 2H).\nResponse: We appreciate the reviewer’s insightful suggestions. In response\nto the reviewer’s concern regarding the structure of lymphatic vessels and\nthe dot-like staining pattern of lymphatics observed using Prox1-CreERT2\ntdTomato reporter mice, we have conducted additional experiments.\nSpecifically, we performed immunolabeling with the vascular endothelial\nmarker CD31, alongside two distinct lymphatic vessel markers (LYVE1 and\nVEGFR3), to further characterize lymphatic vessel morphology. We also\nvalidated the structural integrity and functionality of these lymphatic vessels\nusing indocyanine green (ICG) and Evans blue tools that enable functional\nassessment of lymphatic drainage. Furthermore, to enhance clarity and\nsupport our observations, we have replaced the original figures with higher-\nquality images and revised the corresponding results section to incorporate\nthese new findings and improved visualizations on Page 6-7: “To test whether\ntendons contain lymphatic vessels, we used Prox1-CreERT2; tdTomato mice and\nperformed whole-mount imaging of the Achilles and Patellar tendons. This\napproach enabled visualization of Prox1-Tdtomato-positive lymphatic vessels\nin tendons (Fig.2A, 2B and Video1, 2). To validate their identity, we performed\nimmunolabeling using vascular endothelial and lymphatic vessel markers.\nProx1-Tdtomato-positive lymphatic structures co-expressed the vascular\nendothelial marker CD31 and the LECs marker LYVE1 (Fig. S2B-S2D).\nBecause vascular endothelial growth factor C (VEGF-C)/VEGF receptor 3\n(VEGFR3) signaling is essential for lymphatic development and\nmaintenance[25, 26], we also demonstrated VEGFR3 expression in Prox1-\nTdtomato- positive lymphatic vessels via immunostaining (Fig. S2E). Moreover,\nfor the interstitial fluid drainage function validation of these structures, we\ninjected indocyanine green (ICG) and Evans blue to trace lymphatic flow.\nSubcutaneous ICG injections into the footpad showed uptake in Achilles\ntendons within 30 min as well as time-dependent clearance (Fig. S2F). Evans\nblue accumulated in Prox1-Tdtomato-positive vessels and also cleared over\ntime, confirming the distribution of functional lymphatics in tendons (Fig.\nS2G)[27, 28].\nNext, to test lymphatic responses to tendon injury, we established an Achilles\ntendon transection model. Lymphatic vessels expanded significantly after 7\ndays injury in neonatal and adult mice, with increased Prox1-Tdtomato-positive\nand LYVE1-positive lymphatic vessels (Fig. 2C, 2D and Fig. S2H, S2I).\nQuantitative analysis confirmed significantly elevated Prox1 and Lyve1 mRNA\nexpression after tendon injury (Fig. S2J). To confirm the specificity, we treated\ninjured tendons with VEGFR3-blocking antibody, which inhibited LYVE1-\npositive lymphatic vessel expansion (Fig. S2K). To model chronic tendinopathy,\nwe induced Achilles tendon overuse. In this model, lymphatic expansion also\noccurred, as shown by confocal imaging of Prox1-positive vessels (Fig. S2L).\nTogether, these results demonstrate that both acute and chronic stress drive\nlymphatic expansion in tendons.”\nAs immunofluorescence staining of sections, it is technically challenging to\nvisualize entire lymphatic vessels in a single-layer view. We have replaced high\nquality of 3D confocal imaging in Fig 2A, Fig.2B, Fig 2C, Fig. S2G, Fig. S2H,\nand Fig. S2L.\nRegarding the PROX1 expression shown in Figure 2G: the sections were\nderived from Prox1-CreERT2; tdTomato mice, and the signals depicted\ncorrespond to tdTomato.\nTo solidify the existence of lymphatic vessels in the tendon and the basis for\nthis manuscript the authors should:\n1.A. Provide high-resolution overview and Zoom in images to show the\nlymphatic vessel lumen (optical orthogonal sections or 3D stacks). Also, the\nlocalization of VE-cadherin and/or CD31 at cell junctions, and Prox1 in the\nnucleus of the same cells should be shown.\nResponse: We greatly appreciate the reviewer’s valuable suggestion. We have\nreplaced Figure 2A with high-resolution overview to show lymphatic vessel\nusing Prox1-CreERT2; tdTomato mice. To better delineate lymphatic vessel\ndistribution, we stained tendon tissues isolated from Prox1-CreERT2; tdTomato\nmice with the vascular endothelial marker CD31 and lymphatic markers LYVE1\nand VEGFR3. Additionally, we conducted co-staining of LYVE1 (a lymphatic-\nspecific marker) with CD31 to further characterize lymphatic vessel identity and\nlocalization. The revised results incorporating these experiments have been\nupdated on Page 6: “To validate their identity, we performed immunolabeling\nusing vascular endothelial and lymphatic vessel markers. Prox1-Tdtomato-\npositive lymphatic structures co-expressed the vascular endothelial marker\nCD31 and the lymphatic endothelial marker LYVE1 (Fig. S2B-2D). Because\nvascular endothelial growth factor C (VEGF-C)/VEGF receptor 3 (VEGFR3)\nsignaling is essential for lymphatic development and maintenance[25, 26],\nwe also demonstrated VEGFR3 expression in Prox1-Tdtomato- positive lymphatic\nvessels via immunostaining (Fig. S2E).” and Page 6-7 “Next, to test lymphatic\nresponses to tendon injury, we established an Achilles tendon transection\nmodel. Lymphatic vessels expanded significantly after 7days tendon injury in\nneonatal and adult mice, with increased Prox1-Tdtomato-positive and LYVE1-\npositive lymphatic vessels (Fig. 2C, 2D and Fig. S2H, S2I).”\n1.B. Provide a functional draining assay (a tracer) to show that these structures\ndrain the tissue fluid (the tracer should be seen inside the lymphatic vessels).\nSeveral time points should be included.\nResponse: Thank you for the valuable suggestion. For the functional\nvalidation of lymphatic vessels in tendons, we employed indocyanine green\n(ICG) and Evans blue as tools to visualize and confirm the presence of\nfunctional lymphatic vessels. These newly generated experimental results\nhave been supplemented and incorporated on Page 6: “Moreover, for the\ninterstitial fluid drainage function validation of these structures, we injected\nindocyanine green (ICG) and Evans blue to trace lymphatic flow. Subcutaneous\nICG injections into the footpad showed uptake in Achilles tendons within 30 min\nas well as time-dependent clearance (Fig. S2F). Evans blue accumulated in\nProx1-Tdtomato-positive vessels and also cleared over time, confirming\nfunctional lymphatics in tendons (Fig. S2)[27, 28].”\n1.C. To show that the growth of these structures in development and upon\ninjury/inflammation is dependent on the VEGF-C to VEGFR3 signalling (the key\ngrowth factor signalling route of lymphatics), authors should use one of the\nestablished tools (blocking antibodies, VEGF-C ligand traps, or conditional\nmouse strains).\nResponse: We appreciate your suggestions. To determine whether lymphatic\nexpansion following injury is dependent on VEGFR3 signaling, we conducted\nanalyses using a VEGFR3-blocking antibody. The newly generated\nexperimental results were added on Page 6-7: “Next, to test lymphatic\nresponses to tendon injury, we established an Achilles tendon transection\nmodel. Lymphatic vessels expanded significantly after tendon injury in neonatal\nand adult mice, with increased Prox1-Tdtomato-positive and LYVE1-positive\nlymphatic vessels (Fig. 2C, 2D and Fig. S2H, S2I). Quantitative analysis\nconfirmed significantly elevated Prox1 and Lyve1 mRNA expression after tendon\ninjury (Fig. S2J). To confirm the specificity, we treated injured tendons with\nVEGFR3-blocking antibody, which inhibited LYVE1-positive lymphatic vessel\nexpansion (Fig. S2K).”\n2. Secreted reelin is expected to stay in the vicinity of the lymphatic vessels,\nsuggesting that tendon stem cells should be located close to the lymphatic\nvessels. Authors should provide high-resolution double staining for the\nlymphatics and the tendon stem cells and quantify the distance of the stem cells\nfrom lymphatic in homeostasis and injury/inflammation (three independent\nexperiments are required). Also, authors should add a proliferation marker to\nquantify, whether stem cells in the vicinity of the lymphatics have a higher\nproliferative index than the ones further away.\nThe authors state that the lymphatics expand upon injury/inflammation. Would\nthis mean that only in these conditions do the lymphatics form the niche or, also,\nduring homeostasis?\nResponse: We appreciate your suggestions. We conducted double staining\nfor the lymphatics and tendon stem cells to analyze the regulating effects of\nlymphatic vessels on tendon stem cells. The results were added on Page 10:\n“Histological spatial analysis showed more Vldlr+ TSPCs located near lymphatic\nvessels, with this localized abundance increasing after tendon injury.\nAdditionally, compared with Vldlr+ TSPCs distant from lymphatic vessels, those\nadjacent ones showed a significantly higher proliferation index. Moreover, the\nproliferation of Vldlr+ TSPCs was increased after injury compared with control\nones. (Fig. S6C-S6F). In RelnCKO mice, the proliferation of Vldlr+ cell was also\nreduced (Fig. 5L), implicating lymphatics as a signaling hub maintaining Vldlr+\nTSPC properties. To further validate Reelin's effects on TSPCs, TSPCs was\ntreated with recombinant Reelin protein ….”\nFurthermore, to state the role of lymphatics during physical and pathological\nconditions, the following sentences were added in the Discussion part on\nPage 14-15: “It’s reported that crypt lymphatics preserve intestinal stem cell\nstemness while inhibiting their premature differentiation[14], and skin\nlymphatics support hair follicle stem cells while contributing to skin regeneration\nand repair[40]. These observations indicates that lymphatics act as stem cell\nniches under physiological and pathological conditions[41]. However, the\nspecific function and underlying mechanism are tissue dependent and need\nfurther investigation. Our findings highlight Reelin from LECs as a key regulator\nof TSPC stemness and differentiation. Following injury, Reelin expression in\ntendon lymphatics increased, functioning as a lymphoangiocrine signal\nsupporting tendon repair.”\n3. Authors use Prox1CreERT2 to delete conditional allele of reelin. Authors\nshould show that the reelin signal is lost in lymphatics upon tamoxifen induced\nreelin deletion.\nResponse: Thanks for your suggestions. The deletion of allele of reelin in\nProx1CreERT2；Relnflox/flox was analyzed by immunofluorescence staining and\nthe results were added in Figure. S5A\n4. If reelin affects the growth and/or maintenance of lymphatics (Fig. S4G-H\nand S5N) it is difficult to judge whether the effect of reelin deletion on tendon\nstem cells is directly caused by the reduced reelin amount or by the absence of\nother lymphatic paracrine signals in vivo. Authors should discuss this in their\nmanuscript.\nResponse: We appreciate the reviewer’s suggestion. We repeated the staining\nfor the lymphatic markers LYVE1 and VEGFR3 in Reln-/- and Relncko tendons\nwith their corresponding controls and the results were revised on Page 8:\n“Expression of the lymphatic markers VEGFR3 or LYVE-1 was also reduced in\nReln-/- tendons compared with controls (Fig. S4G and S4H)” and Page 9:\n“VEGFR3 and LYVE1 expression in RelnCKO tendons showed a slight but\nnonsignificant decrease (Fig. S5D).”\nMoreover, the following sentences were added in the Discussion part on Page\n15: “Our findings further underscore the pivotal role of Reelin in regulating the\nstemness and differentiation of TSPCs. Following injury, Reelin expression in\ntendon lymphatics increased, functioning as a lymphoangiocrine signal\nsupporting tendon repair. In Reln-/- mice, newly formed tendons displayed\nreduced cellularity, disorganized collagen fibers, and impaired mechanical\nproperties. Consistent with prior reports of Reln-/- mice exhibiting neural\ndefects and weight gain failure[42], we also observed tendon developmental\ndysplasia, reduced lymphatic formation in Reln-/- mice. A previous study noted\nreduced smooth muscle cell coverage and irregular diameters in lymphatic\ncollecting vessels in Reln-/- mice, despite normal in lymphatic capillaries[43].\nCollectively, these findings suggest that the mechanisms controlling lymphatic\nvessel formation and tendon development may differ by tissue\nmicroenvironment and could be influenced by nervous system defects or\ngrowth impairments in Reln-/- mice. To test the direct role of LEC-derived\nReelin, we deleted Reln in lymphatics using Prox1-CreERT2 mice. The resulting\nRelnCKO mice exhibited impaired tendon regeneration despite normal tendon\ndevelopment and lymphatic formation, confirming that lymphatic-derived\nReelin specifically regulated tendon regeneration. In vitro assays further\nverified the critical role of Reelin in maintaining TSPC stemness.”\n5. The authors use lymphadenectomy as described on rows 642-647. Here\nthe authors execute “a combined popliteal and superficial inguinal lymph node\nexcision”. Does this mean that the lymph nodes and the collector vessel in\nbetween are removed? Then the tendon lymphatics are probably still in place\nbut the lymphatic drainage is disrupted. Authors should show, what happens\nto tendon lymphatics upon lymphadenectomy (are they still present in the\ntendon), does lymphadenectomy alter tendon drainage (see point # 1B) and\nwould loss of drainage affect the paracrine function of the lymphatics (for\nexample, acute reelin expression by the tendon lymphatics).\nResponse: We appreciate the reviewer’s valuable suggestions. The\nexperiments investigating the effects of lymphatics on tendon regeneration,\nusing a lymphadenectomy model, were conducted prior to our acquisition of\nRelncko mice. We acknowledge that the lymphadenectomy model has\nlimitations in clearly delineating the specific role of Reelin derived from LECs in\ntendon regeneration. To enhance the rigor and clarity of our work, we verified\nthe effect of Reelin using Relncko mice, which were revised on Figure 4. We also\nfurther analyzed lymphatic vessel distribution and lymphatic drainage function\nfollowing lymphadenectomy. To strengthen the logic of the manuscript, the\nresults using this model were presented in following attached Figure 1.\nTo investigate the effect of lymphatics on tendon healing, we performed\nlymphadenectomy prior to inducing tendon injury. The results showed that the\nlymphatic vessels were significantly decreased and distributed in an irregular\nscattered manner, with impairment of lymphatic vessel drainage function after\nlymphadenectomy (Attached Fig.1A-1C). The expression of Reelin were\nsignificantly decreased in lymphadenectomy group (Attached Fig.1D).\nFunctionally, the lymphadenectomy group showed impaired tendon healing\nwith lower hindlimb mobility, reduced and disorganized collagen fiber\narrangements diameters compared to the injury group, as assessed by AFI\nindex, HE and Masson trichrome staining (Attached Figure 1E, F). These\nresults indicated that the tendon regeneration capacity was impaired after\nlymphadenectomy. However, the precise mechanism needs further\ninvestigation.\nAttached Figure 1 The tendon regeneration capacity was impaired after\nlymphadenectomy (A) The lymphatic vessels and lymph node stained by Evans blue in control\nand the Lymphectomy in mice limbs. (B) The expression of LYVE-1 in control and lymph group,\nas assessed by immunofluorescence staining. (C) Representative ICG-NIR images of limbs\nimmediately (0 h),2h and 24 h and ICG 24h clearance after ICG injection in control and lymph\ngroup. (D) The expression and semi-quantification of LYVE-1 and Reelin in control and lymph\ngroup, as analyzed by immunofluorescence staining. (E) The footprints and AFI in the control,\ninjury and lymph groups. (F) HE staining, Masson’s trichrome staining and collagen content\nanalysis in the control, injury and lymph groups. (Control: the sham group, Repair: 4 weeks\nafter Achilles tendon resection in P3 mice; Lymph (lymphadenectomy): 4 weeks after Achilles\ntendon resection combined regional lymph ectomy in P3 mice; n=5 mice per group, data were\nrepresent with Mean±SD, *P < 0.05, **P < 0.01 and ***P < 0.001).\nFurthermore, the following sentences were added in the Discussion part on\nPage 15: “Lymphadenectomy mouse models, characterized by disrupted\nlymphatic vessel distribution and impaired lymphatic drainage, also\ndemonstrated compromised tendon regeneration (data not shown). However,\ndetermining whether the impaired tendon regeneration phenotype is mediated\nby lymphatic paracrine signals or defective lymphatic drainage using this model\nremains challenging. Other lymphatic-derived factors, such as netrin-1,\ninterleukin 33, and R-spondin 3[14, 44], may also contribute to tendon\ndevelopment and regeneration and warrant further investigation.”\n6. Also, in other places, the methodology is not detailed enough and does not\nallow replication of the experiments. For example:\nResponse: Thanks for the reviewer’s suggestions. We have revised the method\nparts thoroughly.\n6.A. In cell culture experiments authors silence Dab1 or Srebp2 with an siRNA\napproach (Fig. 6C and G, S7C, S8A-D) and validate the Dab1silencing with RT-\nPCR at 48 hours (Fig. S7B). Indeed transfected siRNAs are thought to cause\ndownregulation of mRNAs for 2-3 days after which the target mRNA levels start\nto increase toward normal levels. In this context, it is difficult to understand why\nthe silencing of Dab1 or Srebp2 has such a drastic effect on tenogenic\ndifferentiation in 21-day protocol. Authors should show the knockdown\nefficiency of Srebp2 mRNA at early and late time points and Dab1 mRNA levels\nat later time points as well. If the Dab1 or Srebp2 mRNA levels increase at a\nlater time point, how do the authors explain the attenuation of differentiation?\nResponse: Thanks for your suggestions. We have added the details for the\nsiRNA transfection for tenogenic differentiation on Page 32: “In 21-day\ntenogenic differentiation induction assay, the transfection complex was added\nevery 3 days with the induction medium change. On day 2 and 21 post-\ntransfection, Srebp2 and Dab1 mRNA levels were quantified by RT-qPCR to\nassess knockout efficiency.” We have added the RT-qPCR results to show the\nexpression of Srebp2 and Dab1 mRNA levels after 2 days and 21 days after\nsiRNA transfection to confirm the knockdown efficacy. The results have been\nadded to Figure S7B and Figure S8D.\n6.B. Authors should describe more carefully the protocol for the isolation of the\ntendon cells, for example, for scRNA sequencing.\nResponse: Thanks for your suggestions. We have revised the details of the\nprotocol for the isolation of the tendon cells in scRNA sequencing on Page 19-\n20: “For scRNA-seq sequencing, tendon tissues were harvested from the injury\nsites of Achilles and Patellar tendons in neonatal (P3) and adult (6-week-old)\nmale C57BL/6J mice after 7 days injury. The same age mice were used as\ncontrol (n=10 per group, pooled for sequencing). After being washed twice with\npre-cooled Phosphate-Buffered Saline (PBS), the collected tissues were cut\ninto 0.5 mm pieces and then digested with 3 mg/mL type I collagenase (Merck,\nGermany) and 4 mg/mL dispase (Merck, Germany) for 30 min at 37 °C with\nshaking. After removing erythrocytes with red blood cell (RBC) lysis buffer\n(Genview, China), the cells were harvested by centrifugation at 450 × g for 15\nminutes at 4 °C, and sequentially filtered through 70 μm cell strainers. The\nisolated cells were harvested for library preparation….”\n7. Reelin is a ligand of Vldlr. Authors, state “Reelin treatment could increase the\ntotal cholesterol level in TPSCs“ (rows 400-401 and Fig. 6L). It is not obvious,\nhow reelin binding would increase cholesterol (apoE-lipoprotein) uptake since\nthe increase in Vldlr mRNA levels upon reelin treatment is fairly moderate (Fig.\nS7A). Authors should investigate the molecular mechanism of increased\ncholesterol uptake. Further, does reelin compete with apoE lipoproteins on\nbinding the Vldlr or are there synergistic effects (for example, co-binding)?\nResponse: We thank the reviewer for the comment. To verify the role of Reelin\non Vldlr mRNA level, we analyzed Vldlr mRNA level at different time point after\nReelin treatment. The results showed that the expression of Vldlr was\nsignificantly increased in a time dependent manner. New experiments results\nwere added on Page 11: “Consistent with this process, Vldlr levels in TSPCs\nwas increased significantly in a time-dependent manner following Reelin\ntreatment (Fig. S7A).”\nTo elucidate the mechanism by which Reelin regulates cholesterol metabolism,\nwe analyzed their effects both on cholesterol uptake and de novo cholesterol\nsynthesis. The newly generated results were added on Page 12-13: “Reelin\ntreatment significantly elevated total cholesterol levels in TSPCs (Fig. 6L).\nMechanistically, Reelin promoted apoE lipoprotein–VLDLR binding (Fig. S8A)\nand increased expression of Srebp2, the master regulator of cholesterol\nbiosynthesis [34]. This effect was partially blocked by DAPT treatment (Fig. 6M,\n6N). Consistently, Reelin-induced cholesterol elevation was partially reversed\nby DAPT (Fig. 6O). Srebp2 target genes, including 3-hydroxy-3-methylglutaryl-\nCoA reductase (Hmgcr) and 3-hydroxy-3-methylglutaryl-CoA synthase (Hmgcs),\nwere significantly upregulated by Reelin treatment (Fig. S8B, S8C). Notably,\nSrebp2 knockdown using siRNA abrogated Reelin-induced cholesterol\nelevation (Fig. S8D, 8E). CUT&Tag-PCR assay confirmed that Reelin\nsignificantly enhanced NICD binding to the Srebp2 promoter (Fig. S8F). These\nfindings indicate that Reelin regulates cholesterol uptake through apoE–VLDLR\ninteraction and de novo cholesterol synthesis via Srebp2 activation. We also\nevaluated the functional relevance of this metabolic regulation, finding that\nReelin-induced enhancements in TSPC proliferation, and tenogenic\ndifferentiation were partially attenuated by Srebp2 siRNA treatment (Fig. S8G,\nS8H). Thus, Reelin modulates cholesterol metabolism through Vldlr–Notch1–\nSrebp2 signaling, which preserves TSPC properties and supports tenogenic\ndifferentiation.”\n8. The data on Notch1 signaling (NICD formation) consists largely of\nrepresentative images. Authors should show quantifications of three\nindependent experiments at least for Fig. 6E-G. Also, authors should use\narrow in Fig. 6F to indicate which bands represent Notch1 and NICD. What\nare the bands in the lowest “row”? Authors should also include a control for the\nfractionation (nuclear and cytoplasmic marker for both fractions).\nResponse: We appreciate the reviewer’s suggestions. The three repeats of\nwestern blot for Fig.6E-G were added in source data file. And the quantitative\nresults of western blot were added in Fig. S7F-S7H.\nThe labels were revised accordingly.\nThe controls for the nuclear and cytoplasmic fractionations were added.\n9. The manuscript text needs proofreading and elaboration of some of the parts\nto allow easy digestion of the key messages. As an example:\n9.A. Rows 498-500: lineage tracing does not indicate the contribution of the\nVldlr gene/protein but, rather, traces the Vldlr positive cells.\nResponse: we appreciate your detailed suggestions. The manuscript was\nrevised and proofread thoroughly. We have revised the manuscript accordingly\n9.B. Rows 487-491: What do the authors mean by “ Previous studies have\nidentified various ligands in lymphatics, such as Ntn1, IL33 and VEGF-C (14,\n37). Our study adds to this body of knowledge by identifying additional tendon\nlymphatic genes, including Reelin, and Rspo3, warranting further exploration\ninto their roles in tendon development and regeneration.” VEGF-C is a ligand,\nproduced by other cell types, which binds VEGFR3 on lymphatic endothelia,\nwhereas reelin is a ligand produced by lymphatic endothelia.\nResponse: We appreciate your valuable suggestions. The manuscript was\nrevised to make the description accurate: “However, determining whether the\nimpaired tendon regeneration phenotype is mediated by lymphatic paracrine\nsignals or defective lymphatic drainage using this model remains challenging.\nOther lymphatic-derived factors, such as netrin-1, interleukin 33, and R-spondin\n3 [14, 44], may also contribute to tendon development and regeneration and\nwarrant further investigation.”\nReviewer #3 (Remarks to the Author):\nIn the manuscript, a functional crosstalk between lymphatic vessels and tendon\nstem/progenitor cells (TSPCs) was found to play a critical role in tendon\nregeneration. Mechanistically, Reelin secreted by lymphatic endothelial cells\n(LECs) promoted phosphorylation of DAB1 through interaction with VLDLR in\nTSPCs. The activated DAB1 subsequently triggered a Notch1-SREBP2-\nmediated cholesterol metabolism pathway, which proved essential for\nmaintaining TSPC stemness. This study reveals a novel lymphatic-tendon\nsignaling axis and demonstrates its physiological relevance in tendon\nregeneration. The authors provided substantial experimental validation through\nrigorous in vitro and in vivo approaches. Although Reelin-DAB1-Notch1-\nSREBP2 signaling has been previously characterized in cardiac and intestinal\ntissues, this work represents the first elucidation of its mechanistic role in\ntendon biology. Given these findings, we recommend this manuscript for\npublication in Nature Communications, pending clarification of the following\npoints:\nResponse: We appreciate the reviewer’s positive comments.\n1. Discrepancies exist between the textual descriptions and annotations in\nFigure 5.\nResponse: Thank you very much for your valuable comment. The manuscript\nhas undergone thorough revision and rigorous proofreading to ensure the\ntextual descriptions and annotations in Figure 5 are consistent and accurate.\n2. Comprehensive documentation of human tendon samples is required,\nparticularly regarding (a) donor demographics, (b) injury etiology, and (c)\ntemporal parameters of sample collection post-injury.\nResponse: Thanks for your suggestions. We have added the detailed\ninformation to the Figure S3C. Moreover, the comprehensive details regarding\nthe human tendon samples used in this study were added in the Methods\nsection on Page 21: “For human tendon samples and three pathologic tendon\nsamples at 3, 10 days post-injury and 12 weeks post-injury, which were\nobtained from published datasets (SRA: PRJNA975881, PRJNA976191),\ncomprehensive donor information was summarized as follows: All samples\nwere derived from male donors aged between 28 and 49 years, with the specific\ninjury etiology being tendon injury. Detailed sample metadata, including exact\nage, sex, disease status, and precise time points of collection, were provided\nin the Figure S3C.”\n3. Methodological clarification: The collagenase-induced model (typically\nmodeling chronic tendinopathy) and transection model (simulating acute injury)\nrepresent distinct pathological contexts. The authors should explicitly address\nhow these models differentially inform their conclusions.\nResponse: Thanks for your suggestions. The tendon transection model was\nused as the acute tendon injury condition. Incorporating the suggestions from\nReviewer 1, we established the Achilles tendon overuse model to mimic\nTendinopathy, which is a chronic tendon disease in clinic and typically resulted\nfrom overuse. The new experimental data were added on Page 6-7: “Next, to\ntest lymphatic responses to tendon injury, we established an Achilles tendon\ntransection model. Lymphatic vessels expanded significantly after 7 days\ntendon injury in neonatal and adult mice, with increased Prox1-Tdtomato-\npositive and LYVE1-positive lymphatic vessels (Fig. 2C, 2D and Fig. S2H, S2I).\nQuantitative analysis confirmed significantly elevated Prox1 and Lyve1 mRNA\nexpression after tendon injury (Fig. S2J). To model chronic tendinopathy, we\ninduced Achilles tendon overuse. In this model, lymphatic expansion also\noccurred, as shown by light-sheet imaging of Prox1-positive vessels (Fig. S2L).\nTogether, these results demonstrate that both acute and chronic stress drive\nlymphatic expansion in tendons.”\n4. Potential immune modulation: Given that (a) lymphatic signaling is\nintrinsically linked to immune regulation, and (b) observed alterations in\nmacrophage markers between normal and injured tendons, the possible\ninvolvement of macrophage-mediated mechanisms in Reelin signaling\nwarrants investigation.\nResponse: We truly appreciate the reviewer's insightful suggestion. We\nanalyzed the macrophages in control and Reln−/− mice and the results were\nadded Page 8: “The inflammation-related M1 macrophage markers inducible\nnitric oxide synthase (iNOS) elevated in the Reln-/- repair group compared with\nthe control repair group (Fig. S4E). Expression of M2 macrophage–related\nmarkers CD206 was lower in the Reln-/- repair group (Fig. S4F).”\nMoreover, the macrophages infiltration in Relnfl/fl and Relncko mice were also\nanalyzed and added in Results part on Page 9: “Additionally, the expression\nof the macrophage markers iNOS, and CD206 was elevated after tendon injury,\nwhile there were no significantly difference between Relnfl/fl and RelnCKO groups\n(Fig. S5B, S5C).”\nFurthermore, the following sentences were added in the Discussion part on\nPage 15-16: “In the newly formed tendons of Reln-deficient mice, we also\ndetected a higher abundance of pro-inflammatory M1 macrophages and lower\nM2 macrophages. This observation aligns with a previous report demonstrating\nthat macrophages isolated from Reln-/- mice exhibit an increased propensity for\nM1 polarization[45]. Our study further revealed that Reelin treatment enhances\nthe functional properties of TSPCs. Notably, advancements in tissue\nregeneration may also be mediated by the role of stem cells or their secretomes,\nwhich have been shown to suppress pro-inflammatory responses and facilitate\nthe resolution of tendon pathologies. Specifically, mesenchymal stem cells are\nknown to secrete prostaglandin E2 (PGE2) and transforming growth factor-β\n(TGF-β), which could promote the activation of M2 macrophages[46].\nAdditionally, lymphatic vessels, which function to drain extravasated fluid,\nantigens, and inflammatory mediators, may themselves modulate macrophage\npolarization[47]. Collectively, whether Reelin regulates macrophages\npolarization directly or indirectly through modulating lymphatic function or the\nanti-inflammatory properties of stem cells，need further investigation to be fully\nelucidated.”"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have provided extensive new data. A few additional comments regarding revised Figure 1 are below.\nFig 1 -\n- 1B, comparison between neonate and adult could also be provided by UMAP\n- 1C, total cell numbers per cluster should be provided\n- 1E, As I understand, this is a 4 way analysis. Frequency data from each of the 4 groups should be shown. The only appearance of a 4 way analysis is in Supp Fig 1I, which is insufficient for interpretation.", "reviewer_2": "The authors have improved the manuscript, for example, the methodology section is improved (point #6 in the original statement), quantification of a western blot is added (#8 ) and interpretation of the lymphadectomy experiment has been modified (#5).\nHowever, the major concern still exists: are the lymphatics within the tendon, as stated by the authors, or superficial, i.e. growing on top of the tendon? This is a major concern and affects the interpretation of the results presented in this manuscript.\n1. The authors now show a draining assay (Fig. S2F-G) and state that “… showed uptake in Achilles tendons within 30 min as well as time-dependent clearance (Fig. S2F).” However, only approximate localization of the signal in the leg is shown. There are non-tendon lymphatic vessels in the leg that drain the footpad area. Thus, this assay does not show that there are functional/fluid-draining lymphatics within the tendon. The authors should show, for example, 3D imaging (a stack), showing lymphatic marker, a drainage tracer, and a tendon marker to show that draining lymphatics are within the tendon.\n2. Authors provide better quality images and videos of lymphatics (for example, Fig. 2A-C, and videos) but it is not possible to interpret whether the lymphatics are truly embedded within the tendon or just grow on top of it (superficial). Authors should use 3D stacks or cross sections to show that lymphatics are within the tendon. Here, “a tendon marker” would make the interpretation of the images easier. Authors should also always indicate in the figure legend whether the shown images are a flattened projection of a stack or images of a sectioned tendon.\n3. As expected, the authors show that the reelin signal is restricted to the direct vicinity of the stained lymphatic vessels (Fig. 2G-H). If lymphatic endothelial reelin directly controls tendon progenitor cells, then it would be expected that the progenitor cells reside next to lymphatic vessels. In response to point #2, authors show that upon injury, the distance of progenitor cells to lymphatic vessels is decreased (Fig. 6D), which is expected, since the density of the vessels is increased. The authors also show that within 100 micrometers from lymphatic vessels the KI67 positivity of VLDLR+ cells is higher. Authors should re-analyze their data and show a histogram of distances of KI67-positive VLDLR+ cells from lymphatic vessels and a histogram of distances of KI67-negative VLDLR+ cells from lymphatic vessels.\n4. Many of the requested data are shown only as representative images, for example, reelin loss (Fig. S5A) and anti-VEGFR3 effect on lymphatic markers (Fig. S2K). In the Figure legends, it should be mentioned how many independent experiments the data represents. The anti-VEGFR3 data should be quantified (3 independent experiments). Also, the used anti-VEGFR3 antibody should be mentioned in the methodology section (the reagent list).\n5. In response to point #4, now, the authors show that there is less lymphatic marker expression upon constitutive (Fig. S4G-H) and conditional loss of reelin (Fig. S5D) (a trend for the latter). It is possible that the tendon phenotype is caused, in part, by decreased amounts of lymphatic vessels and, thus, other lymphatic endothelial paracrine factors. This possibility should be clearly stated in the manuscript.\n6. In response to point #7, the authors state that reelin affects both VLDLR expression (Fig. S7A) and direct APOE binding to VLDLR (Fig. S8A). There is no description of the latter experiment neither in the Figure legend nor in the method section. Please elaborate on how the pull-down was done. The authors should also clarify which ones of the multiple bands represent VLDLR and APOE and provide quantification of 3 independent experiments (normalization of the pulldown protein levels to input levels).", "reviewer_3": "I have carefully reviewed the revised version of the manuscript entitled “Lymphatic–Stem Cell Crosstalk Promotes Tendon Regeneration via Notch1–Srebp2–Mediated Cholesterol Metabolism.”\nGeneral Assessment\nThe revised manuscript now presents a much clearer and coherent study demonstrating that lymphatic endothelial cell–derived Reelin orchestrates tendon stem/progenitor cell (TSPC) behavior through a VLDLR–DAB1–Notch1–SREBP2 axis, which is essential for tendon regeneration. The addition of new imaging, functional, and mechanistic data strongly supports the central hypothesis and improves the robustness of the conclusions. Overall, the quality of figures, histological evidence, and methodological descriptions has been markedly improved.\nRemaining Minor Points for Consideration\nQuantitative Data Presentation:\nWhile the main findings are supported, a few quantitative comparisons (e.g., Vldlr⁺ cell proliferation near lymphatic vessels, Fig. S6F) could benefit from clearer statistical annotation in the figure panels for readers’ clarity.\nDiscussion Section:\nThe discussion could be slightly condensed by merging overlapping statements on developmental versus regenerative mechanisms to improve focus and readability.\nTerminology Consistency:\nPlease ensure consistent use of “lymphatic endothelial cells (LECs)” rather than alternating with “lymphatic endothelium” or “lymphatics,” to maintain terminological precision across sections.\nSummary Recommendation\nThe manuscript now provides a well-structured, mechanistically compelling, and visually convincing demonstration of lymphatic–stem cell interaction in tendon regeneration. The revisions have adequately addressed all major concerns raised in the previous review."}, "author_response": "Response: We sincerely appreciate the reviewer’s valuable suggestions.\nFig 1 - 1B, comparison between neonate and adult could also be provided by UMAP\n- 1C, total cell numbers per cluster should be provided\n- 1E, As I understand, this is a 4 way analysis. Frequency data from each of the 4 groups should be shown. The only appearance of a 4 way analysis is in Supp Fig 1I, which is insufficient for interpretation.\nResponse: We sincerely appreciate the reviewer’s suggestions. We have\nrevised the figures accordingly based on the suggestions.\nIn Fig. 1B, we added the UMAP plot showing the clustering results comparing\nneonate and adult samples. The UMAP plot showing cell clusters for four\ngroups: control and injured tendons of neonatal and adult mice were also added\nin Supplementary Figure 1C.\nIn Fig. 1C, the total cell numbers per cluster were shown in the source data.\nIn Fig. 1E, the frequency data of different clusters for the 4 groups were added.\nReviewer #2 (Remarks to the Author):\nThe authors have improved the manuscript, for example, the methodology\nsection is improved (point #6 in the original statement), quantification of a\nwestern blot is added (#8 ) and interpretation of the lymphadectomy experiment\nhas been modified (#5).\nResponse: We sincerely appreciate the reviewers' thorough efforts and\nconstructive comments.\nHowever, the major concern still exists: are the lymphatics within the tendon,\nas stated by the authors, or superficial, i.e. growing on top of the tendon? This\nis a major concern and affects the interpretation of the results presented in this\nmanuscript.\n1. The authors now show a draining assay (Fig. S2F-G) and state that “…\nshowed uptake in Achilles tendons within 30 min as well as time-dependent\nclearance (Fig. S2F).” However, only approximate localization of the signal in\nthe leg is shown. There are non-tendon lymphatic vessels in the leg that drain\nthe footpad area. Thus, this assay does not show that there are functional/fluid-\ndraining lymphatics within the tendon. The authors should show, for example,\n3D imaging (a stack), showing lymphatic marker, a drainage tracer, and a\ntendon marker to show that draining lymphatics are within the tendon.\nResponse: We appreciate the reviewer’s valuable suggestion. To verify the\npresence of lymphatics within tendons, we analyzed the expression of tendon\nmarkers COL1α1 and FMOD in tendon tissues isolated from Prox1-CreERT2;\ntdTomato mice. The new generated results were added on Page 6-7: “As both\nthe Achilles and patellar tendons are paratenon-sheathed tendons, comprising\na paratenon sheath and tendon substance. We therefore analyzed the precise\ndistribution of expanded lymphatic vessels following injury. Our results\ndemonstrated that in the control group, Prox1-Tdtomato and LYVE1-positive\nlymphatic vessels were mainly localized to the tendon sheath. After injury,\nhowever, lymphatic vessels significantly infiltrated the tendon substance, as\nevidenced by the expression of COL1α1 and FMOD (Fig. S2K, 2L).”\nMoreover, we performed immunolabelling with the tendon marker COL1α1,\nalongside lymphatic vessel markers LYVE1 after a drainage tracer Evans blue\ninjection. The results were added to the results section on Page 6:“Moreover,\nfor the interstitial fluid drainage function validation of these structures, we\ninjected indocyanine green (ICG) and Evans blue to trace lymphatic flow.\nSubcutaneous ICG injections into the footpad showed uptake in Achilles\ntendons within 30 min as well as time-dependent clearance (Fig. S2F). Evans\nblue accumulated in LYVE1 positive lymphatic vessels of tendon tissues\nevidenced by COL1α1 staining, which was also cleared over time (Fig. S2G)”\n2. Authors provide better quality images and videos of lymphatics (for example,\nFig. 2A-C, and videos) but it is not possible to interpret whether the lymphatics\nare truly embedded within the tendon or just grow on top of it (superficial).\nAuthors should use 3D stacks or cross sections to show that lymphatics are\nwithin the tendon. Here, “a tendon marker” would make the interpretation of the\nimages easier. Authors should also always indicate in the figure legend whether\nthe shown images are a flattened projection of a stack or images of a sectioned\ntendon.\nResponse: Thank for your suggestions. To analyze the precise distribution of\nlymphatic vessels in tendon, we analyzed the expression of tendon markers\nCOL1α1 and FMOD in tendon tissues from Prox1-CreERT2; tdTomato mice.\nThe results were added on Page 6-7: “As both Achilles and patellar tendons\nare paratenon-sheathed tendons, comprising a paratenon sheath and tendon\nsubstance. We therefore analyzed the precise distribution of expanded\nlymphatic vessels following injury. Our results demonstrated that in the control\ngroup, Prox1-Tdtomato and LYVE1-positive lymphatic vessels were mainly\nlocalized to the tendon sheath. After injury, however, lymphatic vessels\nsignificantly infiltrated the tendon substance, as evidenced by the expression\nof COL1α1 and FMOD (Fig. S2K, 2L).”\nAdditionally, we have carefully revised the figure legends to clearly specify each\nimage acquisition strategy.\n3. As expected, the authors show that the reelin signal is restricted to the direct\nvicinity of the stained lymphatic vessels (Fig. 2G-H). If lymphatic endothelial\nreelin directly controls tendon progenitor cells, then it would be expected that\nthe progenitor cells reside next to lymphatic vessels. In response to point #2,\nauthors show that upon injury, the distance of progenitor cells to lymphatic\nvessels is decreased (Fig. 6D), which is expected, since the density of the\nvessels is increased. The authors also show that within 100 micrometers from\nlymphatic vessels the KI67 positivity of VLDLR+ cells is higher. Authors should\nre-analyze their data and show a histogram of distances of KI67-positive\nVLDLR+ cells from lymphatic vessels and a histogram of distances of KI67-\nnegative VLDLR+ cells from lymphatic vessels.\nResponse: We appreciate your suggestions. We reanalyzed the distances of\nKI67+ VLDLR+ cells and KI67- VLDLR+ cells from LYVE+ lymphatic vessels. The\nresults were added on Page 11: “Moreover, the proliferating VLDLR positive\ncells were located relatively farther from lymphatic vessels than the non-\nproliferating cells (Fig. S6E-S6H).”\n4. Many of the requested data are shown only as representative images, for\nexample, reelin loss (Fig. S5A) and anti-VEGFR3 effect on lymphatic markers\n(Fig. S2K). In the Figure legends, it should be mentioned how many\nindependent experiments the data represents. The anti-VEGFR3 data should\nbe quantified (3 independent experiments). Also, the used anti-VEGFR3\nantibody should be mentioned in the methodology section (the reagent list).\nResponse: Thank you for your thoughtful comment. The semi-quantifications\nfor Fig. S2K and Fig. S5A were added. The sample number (n=5) was added\nin the Figure legend. Similar findings were obtained in three independent\nexperiments.\nMoreover, the details for VEGFR3 blocking in tendon transection model were\nrevised in the method section on Page 22:“To verify the lymphatic vessels\nexpansion after injury，VEGFR3 neutralizing antibodies were used to determine\nwhether this lymphatic vessels expansion was dependent on VEGFR3 .Briefly,\n100μg VEGFR3 neutralizing antibody was injected into the footpads of Achilles\ntendon resection mouse model every 2 days. The PBS injection ones were\nused as control ones. The samples were harvested 7 days postoperatively for\nfurther investigation (n=5).”\nThe used anti-VEGFR3 antibody was also added to the reagent list in\nsupplementary table 2.\n5. In response to point #4, now, the authors show that there is less lymphatic\nmarker expression upon constitutive (Fig. S4G-H) and conditional loss of reelin\n(Fig. S5D) (a trend for the latter). It is possible that the tendon phenotype is\ncaused, in part, by decreased amounts of lymphatic vessels and, thus, other\nlymphatic endothelial paracrine factors. This possibility should be clearly stated\nin the manuscript.\nResponse: We appreciate the reviewer's comments The following sentences\nwere added to the Discussion part on Page 15:“These findings also suggest\nthat the impaired tendon regeneration, at least in Reln-/- mice, may be partially\nregulated by the decreased lymphatic vessels. Other lymphatic endothelial\nparacrine factors, such as netrin-1, interleukin 33, and R-spondin 3 [14, 44],\nmay also contribute to tendon regeneration and warrant further investigation.”\n6. In response to point #7, the authors state that reelin affects both VLDLR\nexpression (Fig. S7A) and direct APOE binding to VLDLR (Fig. S8A). There is\nno description of the latter experiment neither in the Figure legend nor in the\nmethod section. Please elaborate on how the pull-down was done. The authors\nshould also clarify which ones of the multiple bands represent VLDLR and\nAPOE and provide quantification of 3 independent experiments (normalization\nof the pulldown protein levels to input levels).\nResponse: We sincerely appreciate the reviewer's valuable suggestions. We\nhave revised the labels accordingly and provided the detailed protocol for\nprotein immunoprecipitation in the Methods section on page 30: “For co-\nimmunoprecipitation assay, cells were lysed with lysis buffer (#9803, Cell\nSignaling Technology, USA) for 30 min on ice. Then the cell lysate was\ncentrifuged with 12,000 g for 10 min and supernatant was incubated with\nantibodies overnight at 4 °C, followed by Protein A/G agarose (#sc-2003, Santa\nCruz Biotechnology, USA) incubation for 2 h at 4 °C. The Anti-HA Rabbit mAb\nwas used for NICD-HA protein immunoprecipitation. The Anti-SREBP2 Rabbit\nmAb, Anti-DAB1 Rabbit mAb, and Anti-VLDLR Rabbit mAb were used for\nSREBP2, DAB1 and VLDLR protein immunoprecipitation, respectively…..”\nFor multiple bands, we added a black arrow to indicate the position of VLDLR\nand APOE. Additionally, the triplicate western blot results for Fig. S8A have\nbeen included in the Source Data file, and the corresponding quantitative data\nwere added in Fig. S8A. We also added the quantitative method regarding\nVLDLR binding to APOE to the corresponding figure legend.\nReviewer #3 (Remarks to the Author):\nI have carefully reviewed the revised version of the manuscript entitled\n“Lymphatic–Stem Cell Crosstalk Promotes Tendon Regeneration via Notch1–\nSrebp2–Mediated Cholesterol Metabolism.”\nGeneral Assessment\nThe revised manuscript now presents a much clearer and coherent study\ndemonstrating that lymphatic endothelial cell–derived Reelin orchestrates\ntendon stem/progenitor cell (TSPC) behavior through a VLDLR–DAB1–\nNotch1–SREBP2 axis, which is essential for tendon regeneration. The addition\nof new imaging, functional, and mechanistic data strongly supports the central\nhypothesis and improves the robustness of the conclusions. Overall, the quality\nof figures, histological evidence, and methodological descriptions has been\nmarkedly improved.\nResponse: We sincerely appreciate the reviewers' positive comments.\nRemaining Minor Points for Consideration\n1. Quantitative Data Presentation:\nWhile the main findings are supported, a few quantitative comparisons (e.g.,\nVldlr⁺ cell proliferation near lymphatic vessels, Fig. S6F) could benefit from\nclearer statistical annotation in the figure panels for readers’ clarity.\nResponse: We sincerely appreciate your suggestions. We added the statistical\nannotation to Fig. S6F-6H. Moreover, incorporating the suggestions from\nReviewer 2, we reanalyzed the distances of KI67+ VLDLR+ cells and KI67- VLDLR+\ncells from LYVE+ lymphatic vessels. The results were added on Page\n11: “Moreover, the proliferating VLDLR positive cells were located relatively\nfarther from lymphatic vessels than the nonproliferating cells (Fig. S6E-S6H).”\n2. Discussion Section:\nThe discussion could be slightly condensed by merging overlapping statements\non developmental versus regenerative mechanisms to improve focus and\nreadability.\nResponse: We appreciate the reviewer’s valuable comments. We have\ncarefully revised the Discussion section to delete the overlapping statements\nand make the statement more focus and readability.\n3. Terminology Consistency:\nPlease ensure consistent use of “lymphatic endothelial cells (LECs)” rather than\nalternating with “lymphatic endothelium” or “lymphatics,” to maintain\nterminological precision across sections.\nResponse: Thank you for your suggestions. We have revised the manuscript to\nensure the terminological consistency and precision accordingly.\nSummary Recommendation\nThe manuscript now provides a well-structured, mechanistically compelling,\nand visually convincing demonstration of lymphatic–stem cell interaction in\ntendon regeneration. The revisions have adequately addressed all major\nconcerns raised in the previous review."}, {"round": 3, "reviewer_comments": {"reviewer_2": "The authors have improved the manuscript, for example, by adding quantifications (point #4 in the previous statement), improving the clarity of Western Blot panel (#6) and modifying the discussion and methods (#5 and #4). Major improvement is the co-staining of tendon and lymphatic markers (S2G and S2K-L, #1 and #2), which allows the evaluation of the major point: are the observed lymphatic vessels growing superficially on the tendon or are there lymphatic vessels in the tendon as stated by the authors (see the two previous statements). However, the interpretation of the results and the discussion should better reflect the provided data.\n1. Now, authors demonstrate by co-staining of tendon and lymphatic markers that most lymphatic vessels grow superficially on the tendon, whereas there is only some lymphatic marker staining in/within the tendon in homeostasis (Figure S2K-L). Also, the number of lymphatic vessels increases upon injury (Fig. S2K-L).\na. In relevant locations in the manuscript, the authors should state clearly that most of the lymphatics grow superficially on the tendon and that the performed functional experiments cannnot make a clear distinction between the role of superficial or “within the tendon” lymphatic vessels.\nb. The authors should clarify whether they have quantified in Fig. S2K-L only the LVs in/within the tendon or also the ones growing superficially on the tendon.\n2. Now, authors show Evans blue together with lymphatic and tendon marker staining. In the manuscript, the authors write (p. 6) “Evans blue accumulated in LYVE1 positive lymphatic vessels of tendon tissues evidenced by COL1α1 staining, which was also cleared over time (Fig. S2G)[27, 28]. These results verified the distribution of functional lymphatic vessels in tendons”. However, in Figure S2G it is shown that Evans blue is in the lymphatic vessels that grow superficially on the tendon (at the outer border of the tendon) but it is unclear whether there is any Evans blue within the tendon in the lymphatic vessels. Authors should clearly state that there are functional lymphatic vessels that are located superficially on the tendon, whereas it is unclear whether there are functional lymphatic vessels within/in the tendon.\n3. In the rebuttal letter (point #3) and manuscript (p. 11, rows 301-303) authors write “Moreover, the proliferating VLDLR positive cells were located relatively farther from lymphatic vessels than the non-proliferating cells (Fig. S6E-S6H).”, whereas they show in Figure S6H that the proliferating cells (Ki67+) are closer to lymphatics. Authors should clarify which one is correct.\nThe authors should, further, clarify whether they have quantified the VLDLR+Ki67+ distances only to the LVs in/within the tendon or also to the ones growing superficially on the tendon."}, "author_response": "REVIEWERS' COMMENTS\nReviewer #2 (Remarks to the Author):\nThe authors have improved the manuscript, for example, by adding\nquantifications (point #4 in the previous statement), improving the clarity of\nWestern Blot panel (#6) and modifying the discussion and methods (#5 and #4).\nMajor improvement is the co-staining of tendon and lymphatic markers (S2G\nand S2K-L, #1 and #2), which allows the evaluation of the major point: are the\nobserved lymphatic vessels growing superficially on the tendon or are there\nlymphatic vessels in the tendon as stated by the authors (see the two previous\nstatements). However, the interpretation of the results and the discussion\nshould better reflect the provided data.\n1. Now, authors demonstrate by co-staining of tendon and lymphatic markers\nthat most lymphatic vessels grow superficially on the tendon, whereas there is\nonly some lymphatic marker staining in/within the tendon in homeostasis\n(Figure S2K-L). Also, the number of lymphatic vessels increases upon injury\n(Fig. S2K-L).\na. In relevant locations in the manuscript, the authors should state clearly that\nmost of the lymphatics grow superficially on the tendon and that the performed\nfunctional experiments can not make a clear distinction between the role of\nsuperficial or “within the tendon” lymphatic vessels.\nb. The authors should clarify whether they have quantified in Fig. S2K-L only\nthe LVs in/within the tendon or also the ones growing superficially on the tendon.\nResponse: We sincerely appreciate the reviewer’s valuable suggestions. We\nhave added the discussion in page 6 “Together, these results demonstrate the\nlymphatic vessels are mainly localized on tendon superficial sheath in\nhomeostasis，while both acute and chronic stress drive lymphatic expansion\nand growing into tendon.”\nFurthermore, we have quantified the LVs in/within the tendon and the ones\ngrowing superficially on the tendon in Fig. S2K-L and revised in supplementary\nfigure 2 legend. “The lymphatics both within the tendon and the superficial\nsheath were quantified.”\n2. Now, authors show Evans blue together with lymphatic and tendon marker\nstaining. In the manuscript, the authors write (p. 6) “Evans blue accumulated in\nLYVE1 positive lymphatic vessels of tendon tissues evidenced by COL1α1\nstaining, which was also cleared over time (Fig. S2G)[27, 28]. These results\nverified the distribution of functional lymphatic vessels in tendons”. However, in\nFigure S2G it is shown that Evans blue is in the lymphatic vessels that grow\nsuperficially on the tendon (at the outer border of the tendon) but it is unclear\nwhether there is any Evans blue within the tendon in the lymphatic vessels.\nAuthors should clearly state that there are functional lymphatic vessels that are\nlocated superficially on the tendon, whereas it is unclear whether there are\nfunctional lymphatic vessels within/in the tendon.\nResponse: We sincerely appreciate the reviewer’s insightful suggestions. We\nhave stated that on page 6 “These results indicate the functional lymphatic\nvessels distribution of tendon tissues. However, it was difficult to distinguish\nwhether they distributed superficially on the tendon or within the tendon.”\n3. In the rebuttal letter (point #3) and manuscript (p. 11, rows 301-303) authors\nwrite “Moreover, the proliferating VLDLR positive cells were located relatively\nfarther from lymphatic vessels than the non-proliferating cells (Fig. S6E-S6H).”,\nwhereas they show in Figure S6H that the proliferating cells (Ki67+) are closer\nto lymphatics. Authors should clarify which one is correct.\nThe authors should, further, clarify whether they have quantified the\nVLDLR+Ki67+ distances only to the LVs in/within the tendon or also to the ones\ngrowing superficially on the tendon.\nResponse: Thank you for your thoughtful comment. We revised the statement\nin the manuscript on page 11 “Moreover, the nonproliferating VLDLR positive\ncells were located relatively farther from lymphatic vessels than the proliferating\ncells (Fig. S6E-S6H).”\nMoreover, we quantified the VLDLR+Ki67+ distances to the lymphatics both in\nthe tendon and the sheath. And we added figure legend in Fig. S6E-S6H “The\nVLDLR+Ki67+ distances to lymphatics both within the tendon and the superficial\nsheath were quantified.”"}]

Decision letter