Therapeutic potential of lymphatic endothelial progenitor cells in secondary lymphedema: a preclinical murine study

Therapeutic potential of lymphatic endothelial progenitor cells in secondary lymphedema: a preclinical murine study | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article Therapeutic potential of lymphatic endothelial progenitor cells in secondary lymphedema: a preclinical murine study Ibon Jaunarena, María-Teresa Iglesias-Gaspar, Inazio Arriola-Alvarez, and 4 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-8524204/v1 This work is licensed under a CC BY 4.0 License Status: Posted Version 1 posted You are reading this latest preprint version Abstract Background Secondary lymphedema is a chronic and disabling condition with scarce disease-modifying treatments, particularly after oncologic surgery or radiotherapy. Current management remains largely palliative rather than restorative. Regenerative medicine strategies are emerging as promising alternatives, and lymphatic endothelial progenitor cells (LEPCs) may support reconstruction of damaged lymphatic networks. Methods Secondary lymphedema was induced in C57BL/6J mice using standardized tail skin excision with interruption of superficial lymphatics. Animals received intradermal phosphate-buffered saline (PBS), mesenchymal stem cells (MSCs), or adipose-derived LEPCs on days 1 and 7 post-surgery. Lymphatic function was monitored longitudinally by IVIS-based near-infrared indocyanine green (ICG) imaging and serial tail circumference measurements. Lymphatic vessels were assessed by LYVE-1 immunofluorescence, and fibrosis by Picrosirius Red staining. Results LEPC treatment markedly improved functional lymphatic recovery, as evidenced by faster indocyanine green (ICG) transit on near-infrared imaging, and significantly reduced tail swelling compared with both the MSC and PBS groups. Immunofluorescence analysis revealed a higher density of LYVE-1⁺ lymphatic vessels in LEPC-treated tissue, supporting enhanced lymphatic network reconstruction. Conclusions LEPC administration significantly enhanced lymphatic repair and functional recovery in a murine model of secondary lymphedema. These findings reinforce LEPCs as a biologically targeted and potentially disease-modifying therapeutic option that deserves further translational evaluation. Secondary lymphedema regenerative medicine lymphatic endothelial progenitor cells mouse tail lymphedema model Lymphangiogenesis Figures Figure 1 Figure 2 Figure 3 Background Secondary lymphedema is a chronic consequence of lymphatic system failure, where sustained impairment of fluid transport drives progressive swelling, tissue fibrosis and loss of function. Among cancer survivors, particularly those treated for gynecologic and breast malignancies, it represents a frequent and long-lasting complication that substantially affects quality of life [ 1 – 4 ]. The pathophysiology involves sustained lymphatic injury combined with chronic inflammation and maladaptive tissue remodelling. The accumulation of interstitial fluid promotes an IL-4/IL-13–driven fibrotic and adipogenic response that compromises lymphatic vessel repair and valve integrity [ 5 ]. Within this environment, lymphatic endothelial progenitor cells (LEPCs) play an essential role in vascular regeneration, although their reparative capacity is hindered by the profibrotic milieu. Recent evidence also suggests that mesenchymal progenitors may acquire a lymphatic endothelial phenotype through PROX1-dependent pathways without requiring a venous intermediate stage [ 6 ], indicating considerable developmental plasticity within lymphangiogenic precursor populations. Current management strategies remain symptomatic rather than curative. Complex decongestive therapy—which includes manual lymphatic drainage, compression therapy, exercise, and skin care—continues to be the cornerstone of care, despite requiring lifelong adherence and offering variable clinical benefits [ 4 , 7 ]. Microsurgical procedures such as lymphovenous anastomosis (LVA) and vascularized lymph node transfer (VLNT) can improve limb volume and function but remain technically demanding and do not reliably restore physiologic lymphatic flow [ 8 – 11 ]. These limitations highlight the need for therapies that target the biological mechanisms underlying lymphatic regeneration. Regenerative medicine has therefore gained attention as a potential means to rebuild damaged lymphatic networks. Preclinical and early clinical studies indicate that cell-based approaches may promote lymphangiogenesis, modulate inflammation, and reduce fibrosis, thereby improving lymphatic drainage [ 12 , 13 ]. Mesenchymal stem cells (MSCs) and adipose-derived stem cells (ADSCs) have been the most widely studied, acting mainly through paracrine mechanisms that support host tissue repair [ 14 ]. Early-phase trials have demonstrated the feasibility and safety of ADSC-based interventions in treating cancer-related lymphedema [ 15 , 16 ]. Nonetheless, clinical outcomes remain inconsistent, underscoring the need for more targeted, mechanism-driven strategies. Recent advances have expanded the therapeutic potential of adipose-derived cells. Modulation of the Hippo–YAP pathway in ADSCs has been shown to increase lymphangiogenesis and reduce fibrosis in a mouse model of tail lymphedema [ 17 ]. In parallel, LEPCs have emerged as strong candidates for lymphatic regeneration because of their ability to differentiate into mature lymphatic endothelial cells and contribute directly to new vessel formation [ 5 , 6 , 18 , 19 ]. However, despite these promising findings, studies specifically evaluating LEPC-based therapy in secondary lymphedema remain scarce. Experimental work has confirmed their capacity to integrate into lymphatic structures, but their performance in clinically relevant models and their potential advantages over other regenerative approaches have not been fully defined. This gap emphasizes the importance of rigorous preclinical studies focused on mechanistic and functional outcomes. On the basis of these considerations, this study investigated the therapeutic effect of adipose-derived LEPCs in a validated mouse model of secondary lymphedema. By integrating IVIS–ICG functional imaging, quantitative tail measurements, and detailed histological analyses of lymphangiogenesis and fibrosis, we provide a comprehensive assessment of LEPC-mediated tissue regeneration. This work addresses a critical unmet need in the field and lays the foundation for future translational applications of LEPC-based therapy [ 14 , 15 , 17 ]. Materials and methods Study Design This preclinical experimental study was designed to investigate the therapeutic potential of adipose-derived LEPCs for secondary lymphedema. We employed a validated mouse tail model that reliably induces a chronic lymphatic drainage defect and reproduces the main pathological features of human lymphedema, including fluid stasis, inflammatory activation, and progressive fibrotic remodelling [ 20 – 22 ]. To generate this condition, standardized surgical injury consisting of full-thickness circumferential skin excision combined with disruption of the superficial lymphatic collectors was performed, following established methodologies for inducing stable lymphatic dysfunction in vivo [ 20 – 22 ]. 1 day post-surgery, the animals were randomly allocated to receive phosphate-buffered saline (PBS), MSCs, or adipose-derived LEPCs. The therapeutic response was evaluated through a multimodal approach combining near-infrared indocyanine green (ICG) imaging via the IVIS SpectrumCT system, serial measurements of tail circumference, and detailed histological assessment of lymphatic vessel density, cell engraftment, and extracellular matrix remodelling. This integrative design enabled a comprehensive characterization of functional and structural regeneration following LEPC treatment [ 21 , 23 ]. Animal Model A total of 17 C57BL/6J mice (male and female, 8 weeks old, 20–25 g) were obtained from Janvier Labs (France). The animals were acclimatized for at least 7 days before experimentation and housed under specific pathogen-free (SPF) conditions in groups of 3–5 per cage. The environment was controlled for temperature (22 ± 2°C), humidity (45–65%), and a 12:12 h light/dark cycle, with ad libitum access to food and water. All the cages were provided with nesting material and shelters to increase their welfare. All procedures complied with Spanish Royal Decree 53/2013 and the European Directive 2010/63/EU. The study protocol was approved by the Ethics Committee for Animal Experimentation of IIS Biogipuzkoa and the competent authority of Gipuzkoa (OH-23-25). The animals were checked daily during the first postoperative week and at least three times weekly thereafter. Humane endpoints included > 15% body weight loss, persistent infection or necrosis, impaired mobility, or signs of sustained distress. Euthanasia was performed under deep isoflurane anaesthesia followed by cervical dislocation, following established welfare recommendations [ 24 ]. The study design, conduct and reporting followed the ARRIVE 2.0 recommendations for in vivo research [ 25 ]. Mouse tail lymphedema model Secondary lymphedema was induced via a standardized and stereomicroscope-assisted surgical protocol. A narrow full-thickness circumferential strip of dorsal tail skin (approximately 2 mm wide) was carefully removed 1–2 cm distal to the tail base to interrupt the superficial lymphatic collectors. To facilitate the identification of lymphatic vessels, 25 µL of 1% Evans blue dye (Sigma‒Aldrich) was injected intradermally into the distal tail, allowing selective visualization and subsequent ligation of marked lymphatic channels via fine Vannas microscissors. This technique ensures reproducible interruption of superficial lymphatics while preserving major blood vessels, thereby minimizing ischemic artifacts. Anaesthesia was induced with 4–5% isoflurane and maintained at 1–2% in oxygen (0.5–1 L/min). Perioperative analgesia was provided with subcutaneous meloxicam (0.2 mg/kg). No postoperative bandaging was applied to avoid irritation or self-mutilation behaviours. Lymphedema development was confirmed by a persistent increase in tail circumference measured 1 cm distal to the excision site, following established criteria. This protocol reliably induces a chronic defect in lymphatic drainage and results in progressive interstitial fluid accumulation, inflammatory activation, fibroadipose remodelling, and delayed endogenous lymphangiogenesis—closely recapitulating key pathological features of human secondary lymphedema [ 20 – 22 ]. Recent ultrastructural studies further confirmed that this model reproduces the intussusceptive and sprouting lymphangiogenesis patterns observed in human disease and in high-resolution murine regeneration models [ 26 ]. For these reasons, the mouse tail lymphedema model remains the most widely adopted and technically robust platform for evaluating lymphatic dysfunction and testing regenerative therapies in vivo. Cell isolation and characterization Mesenchymal stem cells (MSCs). All MSCs used in this study were murine (mouse) adipose-derived cells; no human-derived cells or human material were used. Adipose-derived MSCs were obtained from the stromal vascular fraction of murine adipose tissue following enzymatic and mechanical dissociation. Briefly, excised adipose tissue was finely minced and subjected to collagenase type XI digestion (1 mg/mL, 60 min, 37°C) with gentle agitation to release stromal cells. The digested material was filtered through a 70-µm cell strainer to remove debris and centrifuged to isolate the stromal vascular pellet [ 27 ]. The resulting cell fraction was resuspended and expanded directly onto the plastic plates with complete Mesencult medium (05513, StemCell) and cultured in a humidified incubator with a gas mixture of 5% CO2 at 37ºC. Cultures were maintained until they reached approximately 70–80% confluence and were used at passage 3. Cell viability (> 90%) was confirmed prior to administration. Lymphatic endothelial progenitor cells (LEPCs). At passage 3, MSCs were transferred to new dishes using 0.25% Trypsin-EDTA, seeded at 20,000 cells/cm² with complete mouse endothelial cell medium (ECGM, M1168, Cell Biologics) and supplemented with 50 ng/mL of VEGF-C (752-VC, RD Systems). Cells were cultured under standard conditions (37°C with 5% CO₂) for 7 days. Subsequently, the Pod⁺ cells were sorted, briefly expanded (≤ passage 3) and characterized by immunofluorescence and flow cytometry for canonical lymphatic markers (podoplanin and LYVE-1). Only cell preparations with viability > 90% were used for in vivo administration. Experimental Groups, Randomization, and Blinding The animals were allocated into three treatment groups: Control group (n = 4): intradermal PBS MSC group (n = 7): 1 × 10⁶ MSCs in PBS LEPC group (n = 6): 1 × 10⁶ LEPCs in PBS Treatments were administered on postoperative days 1 and 7. A total volume of 100 µL (cell suspension or PBS) was delivered through five intradermal microinjections evenly distributed around the excision site via a 30-gauge needle to ensure uniform coverage of the lymphatic injury zone. Randomization was performed by an investigator not involved in the follow-up assessments. Tail measurements, IVIS/ICG imaging, and histological analysis were conducted by blinded examiners in accordance with best practices for minimizing experimental bias in animal research [ 25 , 28 ]. Functional assessment Tail circumference measurements Tail circumference was recorded before treatment (baseline) and weekly for 36 days via a digital micrometer. Measurements were consistently obtained 1 cm distal to the surgical site to ensure reproducibility. This approach represents a quantitative and widely implemented method for monitoring lymphedema progression and resolution in validated murine tail models [ 22 ]. Lymphatic function assessment Indocyanine green (ICG) near-infrared fluorescence imaging (IVIS) The quantitative assessment of lymphatic transport was performed via near-infrared ICG imaging. The mice received 25 µL of ICG (1 mg/mL in PBS; Sigma‒Aldrich) intradermally at the tail tip, followed by imaging at days 18 and 36 posttreatment via the IVIS SpectrumCT system (PerkinElmer, USA). Fluorescence was captured using 745/800 nm excitation/emission settings. Regions of interest (ROIs) were placed immediately distal and proximal to the surgical gap via Living Image software, and fluorescence intensity ratios (proximal/distal) were calculated to quantify tracer passage across the area of lymphatic disruption. This setup allows objective and reproducible quantification of lymphatic transport over time in the same animal and has recently been validated as a sensitive readout of lymphatic recovery in preclinical lymphedema models [ 23 ]. Histological analysis On day 36, the mice were euthanized, and tail tissues encompassing the injury site were harvested for structural analysis. The samples were fixed in 4% paraformaldehyde (24 h at 4°C), decalcified, dehydrated, embedded in paraffin, and sectioned at 5 µm. The samples were stained with hematoxylin and eosin to help identify different types of cells and tissues and to obtain important information about the cellular structure of the tissue. Fibrosis Collagen deposition was assessed by Picrosirius Red staining (Sigma‒Aldrich) and visualized under polarized light. Fibrotic area quantification was performed via ImageJ software (NIH, USA) with standardized color threshold algorithms. This technique provides sensitive detection of extracellular matrix remodelling and has been validated in recent preclinical lymphedema studies [ 21 ]. Lymphatic vessel density and cell engraftment Immunofluorescence staining was performed using antibodies against LYVE-1 (Abcam, UK), followed by Alexa Fluor–labelled secondary antibodies and DAPI counterstaining. Images were acquired with a Leica SP8 confocal microscope. Lymphatic vessel density was quantified in five randomly selected high-power fields per sample via ImageJ [ 29 ]. Statistical analysis Statistical analyses were conducted via GraphPad Prism v10.0 (GraphPad Software, San Diego, CA, USA). The data are reported as the means ± standard deviations (SDs). Normality was assessed with the Shapiro–Wilk test [ 30 ]. For comparisons between two groups, unpaired Student’s t tests were applied to normally distributed data, and the Mann–Whitney U test was used for nonparametric distributions. Comparisons across three groups were performed with one-way ANOVA followed by Tukey’s post hoc test [ 31 ] or, when appropriate, with the Kruskal–Wallis test and Dunn’s multiple comparisons [ 32 ]. The outliers were identified via Grubbs’ test [ 33 ]. Sample size selection was guided by prior studies using the murine tail lymphedema model [ 20 , 21 ] and aligned with recommendations for rigorous preclinical research [ 25 , 28 ]. Statistical significance was set at p < 0.05. Results Tail swelling over time (tail diameter) Tail diameter was measured weekly for 36 days following lymphedema induction. All animals developed increased tail thickness after surgery, with group-dependent differences in the magnitude and evolution of swelling. Baseline diameters were comparable among groups: control (3.13 ± 0.08 mm), MSC (3.18 ± 0.10 mm), and LEPC (3.19 ± 0.11 mm). Peak swelling occurred on Day 18 in the control group (6.09 ± 0.55 mm), whereas MSC-treated animals reached a lower maximum diameter (5.33 ± 0.36 mm). LEPC-treated mice exhibited the mildest peak swelling (5.25 ± 0.23 mm). From Day 18 onwards, all groups showed a partial reduction in tail diameter, with the LEPC group exhibiting the lowest final diameter on Day 36 (4.61 ± 0.47 mm), followed by the MSC (5.12 ± 0.59 mm) and control (5.59 ± 0.55 mm) groups (Fig. 1). Figure 1. Tail diameter over time across treatment groups. Compared with the MSC and control groups, the LEPC-treated group showed a milder and more transient increase in tail swelling. To better characterize treatment effects, we analysed the difference between each animal’s peak tail diameter and its baseline value (ΔMax–Day 0). The control group showed the greatest increase (3.33 ± 0.46 mm), followed by the MSC (2.68 ± 0.47 mm) and LEPC (2.45 ± 0.51 mm) groups. The difference between the control and LEPC groups was statistically significant (p = 0.032), suggesting reduced swelling progression in LEPC-treated mice. Histological assessment of tissue remodeling (fibrosis and non-tissue/void spaces) Interstitial tissue composition was quantified using the WSI Sirius Red Manual workflow. Within each region of interest (ROI), Sirius Red–positive area and non-tissue/void area were segmented and expressed as percentage of the total ROI. Quantitative analysis showed a significantly lower non-tissue/void area in LEPC-treated mice (7.37 ± 2.87%) compared with MSC-treated (15.87 ± 2.64%) and control animals (19.62 ± 9.31%). No significant differences were observed between the MSC and control groups (p = 0.548). The non-tissue/void area was significantly reduced in the LEPC group compared with both control (p = 0.048) and MSC groups (p = 0.002). Representative histological images are shown in Fig. 2. These histological results were consistent with the reduced tail swelling observed in LEPC-treated animals during longitudinal follow-up. Figure 2. Histological assessment of tissue remodelling. a) Representative macroscopic view of the mouse tail on day 36 post-surgery. b) Hematoxylin and eosin–stained section of the injury area. c) Picrosirius Red–stained section highlighting collagen deposition and non-tissue/void spaces. Scale bar, 200 µm. Lymphatic function assessment Near-infrared ICG imaging provides a dynamic evaluation of lymphatic transport. On Day 18, none of the groups demonstrated tracer passage across the surgical site. By Day 36, LEPC-treated mice exhibited clear proximal-to-distal tracer migration (100% of the animals), as reflected by significantly higher fluorescence intensity ratios than those of the MSC-treated (43% of the animals, 3 of 7) and PBS-treated controls (0% of the animals) ( p < 0.05). Representative IVIS images on Day 36 illustrate these differences (Fig. 3a), with LEPC-treated animals showing visible tracer progression, whereas MSC-treated mice displayed minimal signal and control mice showing no transit. Figure 3. Lymphatic function and structural regeneration. a) Representative IVIS–ICG near-infrared fluorescence images on day 36 post-surgery showing tracer transit across the excision site in LEPC-treated mice compared with MSC-treated and PBS control animals. Arrows indicate the excision site (region of lymphatic disruption) in all groups b) Representative confocal images of LYVE-1 immunostaining in the excision region on day 36. LEPC-treated tissues exhibit a higher density of organized LYVE-1⁺ lymphatic capillaries compared with MSC-treated and PBS control tissues. Arrows indicate lymphatic vessels. Scale bar, 200 µm. Lymphatic Vessel Regeneration Immunostaining of tissue sections confirmed the increased density of LYVE-1⁺ capillaries and the presence of organized lymphatic networks in the LEPC-treated group (Fig. 3b). In contrast, MSC and control tissues displayed sparse and disorganized lymphatic structures. These findings indicate that LEPC treatment enhances both structural regeneration and functional restoration of lymphatic drainage. Discussion In this study, we show that adipose-derived lymphatic endothelial progenitor cells (LEPCs) enhance both structural and functional recovery in a validated murine model of secondary lymphedema. Across independent readouts, LEPC-treated animals showed a consistent improvement in tail thickness, ICG tracer transport, and lymphatic vessel density compared with both PBS and MSC-treated groups. Overall, LEPC treatment produced concordant functional recovery (IVIS–ICG transit and tail thickness) together with increased LYVE-1⁺ vessel density, supporting a true regenerative effect in this model. The concordance between functional imaging, morphometric follow-up, and histological findings supports a true biological treatment effect rather than a measurement-specific artifact. Methodological considerations supporting translational relevance An important methodological aspect of this study is the use of IVIS-based near-infrared ICG imaging as the main functional readout. Conventional assessments, such as Evans blue transit, provide limited sensitivity and cannot be performed longitudinally. In contrast, IVIS enables dynamic, repeated, and quantitative evaluation of lymphatic flow within the same animal, thereby improving reproducibility and translational value. Recent studies support the use of longitudinal near-infrared ICG imaging as a sensitive functional endpoint in preclinical lymphatic models [ 23 ]. In our dataset, IVIS clearly separated the functional effects of LEPCs from those of MSCs and controls, highlighting its utility for preclinical therapeutic screening. The murine tail model employed in this study remains one of the most widely used and reproducible systems to evaluate lymphatic dysfunction and regeneration [ 20 – 22 ]. Its defined injury pattern, predictable edema kinetics, and compatibility with high-resolution imaging provide a stable platform for testing regenerative interventions. The integration of IVIS-ICG imaging into this model further enhances sensitivity and allows comprehensive evaluation of lymphatic recovery. Regenerative advantages of LEPCs over MSC- and ADSC-based therapies Previous studies using MSCs or adipose-derived regenerative cell populations have demonstrated beneficial paracrine effects on lymphangiogenesis and tissue remodelling in experimental lymphedema [ 14 , 15 ]. Enhanced approaches such as Hippo–YAP modulation have further amplified the lymphangiogenic potential of adipose-derived stromal cells in murine tail lymphedema [ 17 ]. Our results expand this landscape by showing that LEPCs can induce substantial lymphatic regeneration without requiring gene engineering. The podoplanin-positive progenitor population used here was associated with improved lymphatic repair, as shown by reduced edema and reconstitution of LYVE-1⁺ and VEGFR-3⁺ lymphatic networks. These findings support the hypothesis that LEPC-mediated recovery may involve paracrine activity and/or direct endothelial incorporation, although the relative contribution of each mechanism cannot be determined in the absence of dedicated cell-tracking approaches. The pattern of structural remodelling observed is consistent with intussusceptive lymphangiogenesis, as recently described in high-resolution imaging studies [ 26 ]. Microenvironmental barriers and potential combinatorial strategies Secondary lymphedema is characterized by a fibrotic and inflammatory milieu that can hinder tissue repair and limit long-term recovery [ 5 , 34 ]. In our study, LEPC-treated tissues presented a reduced Sirius Red–positive area at day 36, suggesting that improving lymphatic transport may help attenuate fibrotic remodelling within the injury zone. In parallel, the lower non-tissue/void area observed histologically is consistent with reduced interstitial expansion and tissue disruption in the treated group. Emerging evidence suggests that targeting extracellular matrix remodelling may potentiate regenerative strategies in secondary lymphedema. Inhibition of uPARAP has been reported to improve lymphatic architecture and reduce fibrosis in preclinical models [ 35 ]; mechanotransduction pathways such as Hippo–YAP/TAZ may also influence lymphatic endothelial behaviour [ 17 ]. In our study, the reduced fibrotic signal in LEPC-treated tissues supports the relevance of these microenvironmental constraints. Paracrine cooperation and integration with next-generation therapeutics Reported synergy between MSCs and LEPCs suggests that paracrine support may enhance progenitor survival and lymphangiogenic activity; for example, MSC-derived IGF-1 has been linked to improved LEPC function via PI3K/Akt/mTOR signalling and superior outcomes with cotherapy compared with single-cell approaches [ 36 ]. Other “next-generation” combination strategies—such as Apelin + VEGF-C delivery via mRNA [ 37 ] and platelet-derived extracellular vesicles [ 38 ]—further support a multimodal regenerative framework. Together, these data raise the possibility that integrating LEPCs with paracrine-active or microenvironment-modulating interventions could improve durability beyond the functional benefit observed with LEPCs alone in our model, although this was not tested here. Positioning of LEPC therapy within the evolving lymphatic regeneration landscape LEPCs represent a mechanistically rational strategy for secondary lymphedema given their lymphatic-lineage orientation. In our study, LEPC treatment was associated with concordant functional improvement (ICG transport and tail thickness) together with increased LYVE-1⁺ lymphatic vessel density, supporting further translational evaluation. Key next steps include standardizing LEPC isolation and characterization workflows [ 39 ], testing whether combination approaches improve performance in fibrotic tissue, and extending follow-up to assess durability, biodistribution, and safety. Limitations Several limitations of this study must be acknowledged. First, the sample size, although consistent with established murine tail lymphedema studies [ 20 – 22 ], remains modest and may limit the statistical power for detecting small or late therapeutic effects. Increasing the group size, especially for MSCs, could help confirm the differences observed in this work. Second, the tail model—despite being one of the most reproducible and widely used platforms for lymphatic research—does not fully recapitulate the anatomical and biomechanical complexity of human limb lymphedema. The absence of muscle compartments, fascia, and weight-bearing forces limits the extrapolation of certain mechanotransductive phenomena and stromal interactions. Third, although IVIS-ICG provides sensitive and longitudinal functional assessment, it reflects tracer transit rather than true lymphatic pumping or valve competence. Additional techniques, such as near-infrared dynamic contractility imaging or lymphoscintigraphy, could strengthen functional characterization. Fourth, the identification of LEPCs relies on classical lymphatic markers (LYVE-1, VEGFR-3, and PROX1). While consistent with current standards, the heterogeneity of lymphatic progenitor populations is increasingly recognized [ 6 , 39 ]. Single-cell profiling or lineage-tracing approaches would more precisely define the subpopulations responsible for the regenerative effect. Fifth, LEPC–host interactions within the fibrotic microenvironment have not been investigated mechanistically. The contributions of macrophages, fibroblasts, and Th2-skewed inflammation to graft success or failure remain largely unexplored in this setting. Finally, long-term durability and potential off-target effects could not be assessed because of the 36-day follow-up. Chronic models—beyond 8–12 weeks—are required to capture lymphatic remodelling, stabilize fibrosis, and maintain the persistence of engrafted cells. Future directions Future work should prioritize mechanistic validation and translational strengthening. Key next steps include cell-tracking approaches to define the relative contribution of paracrine effects versus endothelial incorporation, extending follow-up to assess durability and safety, and testing clinically aligned models (e.g., hindlimb and/or irradiation-associated lymphedema). In parallel, standardization of LEPC isolation/characterization will be essential to improve reproducibility and support future translation [ 39 ]. Conclusions Our findings show that LEPC transplantation improves multiple hallmarks of secondary lymphedema, outperforming MSCs and PBS controls across functional, morphometric, and histological endpoints. The integration of IVIS–ICG imaging with quantitative tail measurements and lymphatic marker analysis provides convergent evidence of treatment-associated lymphatic recovery in this preclinical model. Together, these results support further investigation of LEPC-based interventions as a regenerative strategy for secondary lymphedema, including studies designed to define mechanisms, durability, and translational feasibility. Abbreviations ADSCs: Adipose-derived stem cells ARRIVE: Animal Research: Reporting of In Vivo Experiments ECGM: Endothelial cell growth medium H&E: Hematoxylin and eosin ICG: Indocyanine green IL: Interleukin IVIS: In vivo imaging system LEPCs: Lymphatic endothelial progenitor cells LVA: Lymphaticovenular anastomosis LYVE-1: Lymphatic vessel endothelial hyaluronan receptor 1 MSCs: Mesenchymal stem cells PBS: Phosphate-buffered saline PROX1: Prospero homeobox 1 ROI: Region of interest SPF: Specific pathogen-free UPV/EHU: University of the Basque Country VEGFR-3: Vascular endothelial growth factor receptor 3 VLNT: Vascularized lymph node transfer Declarations Ethics approval and consent to participate Ethics approval and consent to participate: All animal procedures were authorized by the Diputación Foral de Gipuzkoa ( Provincial Council of Gipuzkoa) under the approved project “Capacidad regenerativa de precursores linfangiogénicos para el tratamiento de linfedema secundario en modelo murino (OH-23-25)” (administrative identification code PRO-AE-SS-311; approval date 13 December 2023). All procedures complied with Spanish Royal Decree 53/2013 and the European Directive 2010/63/EU. Use of artificial intelligence The authors declare that artificial intelligence–based tools were used exclusively for language editing, grammar correction, and improvement of clarity and structure in selected sections of the manuscript. No artificial intelligence tools were used for data generation, data analysis, figure creation, or interpretation of results. All scientific content, analyses, and conclusions were verified and approved by the authors, who take full responsibility for the integrity and accuracy of the work. Consent for publication Not applicable. Availability of data and materials The datasets generated and/or analysed during the current study are available from the corresponding author on reasonable request. Competing interests The authors declare that they have no competing interests. Funding This work received no specific funding. Authors’ contributions IJ and HL conceived and designed the study, performed the experiments, acquired the data (including tail measurements and imaging), analysed and interpreted the results, and wrote the manuscript. MTI performed the biostatistical analysis of the tail measurement dataset and critically revised the manuscript. IA established and performed the LEPC isolation workflow and contributed to cell preparation and characterization. AI, ID and AL contributed to manuscript revision and approved the final version. All authors read and approved the final manuscript. Acknowledgements The authors thank the Animal Facility and Imaging Core of IIS Biogipuzkoa for technical support and the Histology Service of CIMA (Centro de Investigación Médica Aplicada, Universidad de Navarra, Pamplona, Spain) for assistance with tissue processing and immunofluorescence imaging. 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Adipose-derived regenerative cells and lipotransfer in alleviating breast cancer-related lymphedema: An open-label phase I trial with 4 years of follow-up. Stem Cells Transl Med. 2021;10(6):844–854. doi:10.1002/sctm.20-0394. Hu L, Zhang N, Zhao C, Pan J. Engineering ADSCs by manipulating YAP for lymphedema treatment in a mouse tail model. Exp Biol Med (Maywood). 2024 Nov 20;249:10295. doi:10.3389/ebm.2024.10295. Lee JY, Park C, Cho YP, Lee E, Kim H, Kim P, Yun SH, Yoon YS. Podoplanin-expressing cells derived from bone marrow play a crucial role in postnatal lymphatic neovascularization. Circulation. 2010 Oct 5;122(14):1413–1425. doi:10.1161/CIRCULATIONAHA.110.941468. Dai T, Jiang Z, Cui C, Sun Y, Lu B, Li H, Cao W, Chen B, Li S, Guo L. The Roles of Podoplanin-Positive/Podoplanin-Negative Cells from Adipose-Derived Stem Cells in Lymphatic Regeneration. Plast Reconstr Surg. 2020 Feb;145(2):420–431. doi:10.1097/PRS.0000000000006474. Weiler MJ, Cribb MT, Nepiyushchikh Z, Nelson TS, Dixon JB. A novel mouse tail lymphedema model for observing lymphatic pump failure during lymphedema development. Sci Rep. 2019 Jul 18;9(1):10405. doi:10.1038/s41598-019-46797-2. Zhou C, Su W, Han H, Li N, Ma G, Cui L. Mouse tail models of secondary lymphedema: fibrosis gradually worsens and is irreversible. Int J Clin Exp Pathol. 2020 Jan 1;13(1):54–64. PMCID: PMC7013376. Hassanein AH, Sinha M, Neumann CR, Mohan G, Khan I, Sen CK. A Murine Tail Lymphedema Model. J Vis Exp. 2021 Feb 10;(168):e61848. doi:10.3791/61848. Yamaji Y, Akita S, Akita H, Miura N, Gomi M, Manabe I, Kubota Y, Mitsukawa N. Development of a mouse model for the visual and quantitative assessment of lymphatic trafficking and function by in vivo imaging. Sci Rep. 2018 Apr 12;8(1):5921. doi:10.1038/s41598-018-23693-9. American Veterinary Medical Association. AVMA Guidelines for the Euthanasia of Animals: 2020 Edition. Schaumburg (IL): American Veterinary Medical Association; 2020. Percie du Sert N, Hurst V, Ahluwalia A, Alam S, Avey MT, Baker M, Browne WJ, Clark A, Cuthill IC, Dirnagl U, Emerson M, Garner P, Holgate ST, Howells DW, Karp NA, Lazic SE, Lidster K, MacCallum CJ, Macleod M, Petersen OH, Rawle F, Reynolds P, Rooney K, Sena ES, Silberberg SD, Steckler T, Würbel H. The ARRIVE guidelines 2.0: updated guidelines for reporting animal research. PLoS Biol. 2020 Jul;18(7):e3000410. doi:10.1371/journal.pbio.3000410. Hayashida K, Ogino R, Suda S, Yamakawa S. Scanning electron microscopy analysis of lymphatic regeneration in a secondary lymphedema mouse model: a preliminary study. Lymphatics. 2023;1(3):237–243. doi:10.3390/lymphatics1030014. Bourin P, Bunnell BA, Casteilla L, Dominici M, Katz AJ, March KL, Redl H, Rubin JP, Yoshimura K, Gimble JM. Stromal cells from the adipose tissue-derived stromal vascular fraction and culture expanded adipose tissue-derived stromal/stem cells: a joint statement of the IFATS and the ISCT. Cytotherapy. 2013;15(6):641–648. doi:10.1016/j.jcyt.2013.02.006. Festing MFW, Altman DG. Guidelines for the design and statistical analysis of experiments using laboratory animals. ILAR J. 2002;43(4):244–258. doi:10.1093/ilar.43.4.244. Mohan G, Khan I, Diaz SM, Kamocka MM, Hulsman LA, Ahmed S, Neumann CR, Jorge MD, Gordillo GM, Sen CK, Sinha M, Hassanein AH. Quantification of lymphangiogenesis in the murine lymphedema tail model using intravital microscopy. Lymphat Res Biol. 2024 Jun;22(3):195–202. doi:10.1089/lrb.2023.0048. Shapiro SS, Wilk MB. An analysis of variance test for normality (complete samples). Biometrika. 1965;52(3–4):591–611. doi:10.1093/biomet/52.3-4.591. Tukey JW. Comparing individual means in the analysis of variance. Biometrics. 1949;5(2):99–114. Dunn OJ. Multiple comparisons using rank sums. Technometrics. 1964;6(3):241–252. doi:10.1080/00401706.1964.10490181. Grubbs FE. Procedures for detecting outlying observations in samples. Technometrics. 1969;11(1):1–21. doi:10.1080/00401706.1969.10490657. Hossain L, Gomes KP, Safarpour S, Gibson SB. The microenvironment of secondary lymphedema. The key to finding effective treatments? Biochim Biophys Acta Mol Basis Dis. 2025 Mar;1871(3):167677. doi:10.1016/j.bbadis.2025.167677. Gucciardo F, Lebeau A, Pirson S, Buntinx F, Ivanova E, Blacher S, Brouillard P, Deroye J, Baudin L, Pirnay A, Morfoisse F, Villette C, Nizet C, Lallemand F, Munaut C, Alitalo K, Geris L, Vikkula M, Gautier-Isola M, Noel A. Targeting uPARAP modifies lymphatic vessel architecture and attenuates lymphedema. Circulation. 2025 May 13;151(19):1412–1429. doi:10.1161/CIRCULATIONAHA.124.072093. Xue Z, Yang D, Jin Z, Li Y, Yu Y, Zhao X, Huang Y, Jia S, Zhang T, Huang G, Hou J. IGF-1 secreted by mesenchymal stem cells affects the function of lymphatic endothelial progenitor cells: a potential strategy for the treatment of lymphedema. Front Genet. 2025;16:1584095. doi:10.3389/fgene.2025.1584095. Creff J, Lamaa A, Benuzzi E, Balzan E, Pujol F, Draia-Nicolau T, Bonnevie T, Boisset N, Lemaigre F, Chuvin N, Clavel L, Carpentier G, Pratx S, Duhamel A, Bidault G, Jude B, Soncin F. Apelin-VEGF-C mRNA delivery as therapeutic for the treatment of secondary lymphedema. EMBO Mol Med. 2024 Feb;16(2):386–415. doi:10.1038/s44321-023-00017-7. Vachon L, Jean G, Milasan A, Ganivet A, Monge-Rojas J, Rolland-Yasuhara M, Cartier R, Martel C. Platelet extracellular vesicles preserve lymphatic endothelial cell integrity and enhance lymphatic vessel function. Commun Biol. 2024 Aug 11;7(1):975. doi:10.1038/s42003-024-06675-8. Saha S, Graham F, Knopp J, Patzke C, Hanjaya-Putra D. Robust differentiation of human pluripotent stem cells into lymphatic endothelial cells using transcription factors. Cells Tissues Organs. 2024;213(6):464–474. doi:10.1159/000539699. Additional Declarations No competing interests reported. Cite Share Download PDF Status: Posted Version 1 posted You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. Our growing team is made up of researchers and industry professionals working together to solve the most critical problems facing scientific publishing. Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-8524204","acceptedTermsAndConditions":true,"allowDirectSubmit":true,"archivedVersions":[],"articleType":"Research Article","associatedPublications":[],"authors":[{"id":587844673,"identity":"ff50625e-f493-4181-a4ed-03333266e0b8","order_by":0,"name":"Ibon Jaunarena","email":"data:image/png;base64,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","orcid":"","institution":"Biogipuzkoa Health Research Institute","correspondingAuthor":true,"prefix":"","firstName":"Ibon","middleName":"","lastName":"Jaunarena","suffix":""},{"id":587844675,"identity":"fb213857-d228-4361-98d6-bfaa69bd9169","order_by":1,"name":"María-Teresa Iglesias-Gaspar","email":"","orcid":"","institution":"Donostia University Hospital","correspondingAuthor":false,"prefix":"","firstName":"María-Teresa","middleName":"","lastName":"Iglesias-Gaspar","suffix":""},{"id":587844676,"identity":"304fd004-c44b-456d-af32-d8ae25816884","order_by":2,"name":"Inazio Arriola-Alvarez","email":"","orcid":"","institution":"Biogipuzkoa Health Research Institute","correspondingAuthor":false,"prefix":"","firstName":"Inazio","middleName":"","lastName":"Arriola-Alvarez","suffix":""},{"id":587844678,"identity":"d5325c0b-da13-46ff-98ca-f4bfe60cd851","order_by":3,"name":"Ander Izeta","email":"","orcid":"","institution":"Biogipuzkoa Health Research Institute","correspondingAuthor":false,"prefix":"","firstName":"Ander","middleName":"","lastName":"Izeta","suffix":""},{"id":587844679,"identity":"46fc0030-b001-4b29-b8ab-7711749a9802","order_by":4,"name":"Irene Diez-Itza","email":"","orcid":"","institution":"Donostia University Hospital","correspondingAuthor":false,"prefix":"","firstName":"Irene","middleName":"","lastName":"Diez-Itza","suffix":""},{"id":587844681,"identity":"ae250228-d524-4d50-8729-8e449c132e21","order_by":5,"name":"Arantza Lekuona","email":"","orcid":"","institution":"Donostia University Hospital","correspondingAuthor":false,"prefix":"","firstName":"Arantza","middleName":"","lastName":"Lekuona","suffix":""},{"id":587844682,"identity":"89a21303-6b33-4831-b6d5-b93cf61069b8","order_by":6,"name":"Héctor Lafuente","email":"","orcid":"","institution":"Biogipuzkoa Health Research Institute","correspondingAuthor":false,"prefix":"","firstName":"Héctor","middleName":"","lastName":"Lafuente","suffix":""}],"badges":[],"createdAt":"2026-01-05 18:38:42","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-8524204/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-8524204/v1","draftVersion":[],"editorialEvents":[],"editorialNote":"","failedWorkflow":false,"files":[{"id":102750640,"identity":"87151f80-89f3-4426-b098-162f4b6ae033","added_by":"auto","created_at":"2026-02-16 09:21:08","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":36859,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eTail diameter over time across treatment groups. Compared with the MSC and control groups, the LEPC-treated group showed a milder and more transient increase in tail swelling.\u003c/strong\u003e\u003c/p\u003e","description":"","filename":"1.png","url":"https://assets-eu.researchsquare.com/files/rs-8524204/v1/fc4c9fbbae230357e199d1c3.png"},{"id":102751742,"identity":"a37138b0-b3fc-40b4-9fb7-66e26d842850","added_by":"auto","created_at":"2026-02-16 09:27:20","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":1114376,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eHistological assessment of tissue remodelling.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003ea) Representative macroscopic view of the mouse tail on day 36 post-surgery.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eb) Hematoxylin and eosin–stained section of the injury area.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003ec) Picrosirius Red–stained section highlighting collagen deposition and non-tissue/void spaces. Scale bar, 200 µm.\u003c/strong\u003e\u003c/p\u003e","description":"","filename":"2.png","url":"https://assets-eu.researchsquare.com/files/rs-8524204/v1/bda233bcda3521f29fe2d624.png"},{"id":102751717,"identity":"e36763a2-2991-4634-8f7c-83306f56abc0","added_by":"auto","created_at":"2026-02-16 09:27:15","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":430509,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eLymphatic function and structural regeneration.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003ea) Representative IVIS–ICG near-infrared fluorescence images on day 36 post-surgery showing tracer transit across the excision site in LEPC-treated mice compared with MSC-treated and PBS control animals. Arrows indicate the excision site (region of lymphatic disruption) in all groups\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eb) Representative confocal images of LYVE-1 immunostaining in the excision region on day 36. LEPC-treated tissues exhibit a higher density of organized LYVE-1⁺ lymphatic capillaries compared with MSC-treated and PBS control tissues. Arrows indicate lymphatic vessels. Scale bar, 200 µm.\u003c/strong\u003e\u003c/p\u003e","description":"","filename":"3.png","url":"https://assets-eu.researchsquare.com/files/rs-8524204/v1/cc4abaae332b4eb2ac1e3aad.png"},{"id":103056504,"identity":"dc74102d-97ae-4903-a026-4a235597c17e","added_by":"auto","created_at":"2026-02-20 09:12:25","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":3592295,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-8524204/v1/1c2e07ca-4849-4703-8700-052ebd098b3f.pdf"}],"financialInterests":"No competing interests reported.","formattedTitle":"Therapeutic potential of lymphatic endothelial progenitor cells in secondary lymphedema: a preclinical murine study","fulltext":[{"header":"Background","content":"\u003cp\u003eSecondary lymphedema is a chronic consequence of lymphatic system failure, where sustained impairment of fluid transport drives progressive swelling, tissue fibrosis and loss of function.\u003c/p\u003e \u003cp\u003eAmong cancer survivors, particularly those treated for gynecologic and breast malignancies, it represents a frequent and long-lasting complication that substantially affects quality of life [\u003cspan additionalcitationids=\"CR2 CR3\" citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eThe pathophysiology involves sustained lymphatic injury combined with chronic inflammation and maladaptive tissue remodelling. The accumulation of interstitial fluid promotes an IL-4/IL-13\u0026ndash;driven fibrotic and adipogenic response that compromises lymphatic vessel repair and valve integrity [\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e]. Within this environment, lymphatic endothelial progenitor cells (LEPCs) play an essential role in vascular regeneration, although their reparative capacity is hindered by the profibrotic milieu. Recent evidence also suggests that mesenchymal progenitors may acquire a lymphatic endothelial phenotype through PROX1-dependent pathways without requiring a venous intermediate stage [\u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e], indicating considerable developmental plasticity within lymphangiogenic precursor populations.\u003c/p\u003e \u003cp\u003eCurrent management strategies remain symptomatic rather than curative. Complex decongestive therapy\u0026mdash;which includes manual lymphatic drainage, compression therapy, exercise, and skin care\u0026mdash;continues to be the cornerstone of care, despite requiring lifelong adherence and offering variable clinical benefits [\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e, \u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e]. Microsurgical procedures such as lymphovenous anastomosis (LVA) and vascularized lymph node transfer (VLNT) can improve limb volume and function but remain technically demanding and do not reliably restore physiologic lymphatic flow [\u003cspan additionalcitationids=\"CR9 CR10\" citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e]. These limitations highlight the need for therapies that target the biological mechanisms underlying lymphatic regeneration.\u003c/p\u003e \u003cp\u003eRegenerative medicine has therefore gained attention as a potential means to rebuild damaged lymphatic networks. Preclinical and early clinical studies indicate that cell-based approaches may promote lymphangiogenesis, modulate inflammation, and reduce fibrosis, thereby improving lymphatic drainage [\u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e, \u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e]. Mesenchymal stem cells (MSCs) and adipose-derived stem cells (ADSCs) have been the most widely studied, acting mainly through paracrine mechanisms that support host tissue repair [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e]. Early-phase trials have demonstrated the feasibility and safety of ADSC-based interventions in treating cancer-related lymphedema [\u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e, \u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e]. Nonetheless, clinical outcomes remain inconsistent, underscoring the need for more targeted, mechanism-driven strategies.\u003c/p\u003e \u003cp\u003eRecent advances have expanded the therapeutic potential of adipose-derived cells. Modulation of the Hippo\u0026ndash;YAP pathway in ADSCs has been shown to increase lymphangiogenesis and reduce fibrosis in a mouse model of tail lymphedema [\u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e]. In parallel, LEPCs have emerged as strong candidates for lymphatic regeneration because of their ability to differentiate into mature lymphatic endothelial cells and contribute directly to new vessel formation [\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e, \u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e, \u003cspan citationid=\"CR18\" class=\"CitationRef\"\u003e18\u003c/span\u003e, \u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eHowever, despite these promising findings, studies specifically evaluating LEPC-based therapy in secondary lymphedema remain scarce. Experimental work has confirmed their capacity to integrate into lymphatic structures, but their performance in clinically relevant models and their potential advantages over other regenerative approaches have not been fully defined. This gap emphasizes the importance of rigorous preclinical studies focused on mechanistic and functional outcomes.\u003c/p\u003e \u003cp\u003eOn the basis of these considerations, this study investigated the therapeutic effect of adipose-derived LEPCs in a validated mouse model of secondary lymphedema. By integrating IVIS\u0026ndash;ICG functional imaging, quantitative tail measurements, and detailed histological analyses of lymphangiogenesis and fibrosis, we provide a comprehensive assessment of LEPC-mediated tissue regeneration. This work addresses a critical unmet need in the field and lays the foundation for future translational applications of LEPC-based therapy [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e, \u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e, \u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e].\u003c/p\u003e"},{"header":"Materials and methods","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003eStudy Design\u003c/h2\u003e \u003cp\u003eThis preclinical experimental study was designed to investigate the therapeutic potential of adipose-derived LEPCs for secondary lymphedema. We employed a validated mouse tail model that reliably induces a chronic lymphatic drainage defect and reproduces the main pathological features of human lymphedema, including fluid stasis, inflammatory activation, and progressive fibrotic remodelling [\u003cspan additionalcitationids=\"CR21\" citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eTo generate this condition, standardized surgical injury consisting of full-thickness circumferential skin excision combined with disruption of the superficial lymphatic collectors was performed, following established methodologies for inducing stable lymphatic dysfunction in vivo [\u003cspan additionalcitationids=\"CR21\" citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e]. 1 day post-surgery, the animals were randomly allocated to receive phosphate-buffered saline (PBS), MSCs, or adipose-derived LEPCs.\u003c/p\u003e \u003cp\u003eThe therapeutic response was evaluated through a multimodal approach combining near-infrared indocyanine green (ICG) imaging via the IVIS SpectrumCT system, serial measurements of tail circumference, and detailed histological assessment of lymphatic vessel density, cell engraftment, and extracellular matrix remodelling. This integrative design enabled a comprehensive characterization of functional and structural regeneration following LEPC treatment [\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e, \u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e].\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eAnimal Model\u003c/h3\u003e\n\u003cp\u003eA total of 17 C57BL/6J mice (male and female, 8 weeks old, 20\u0026ndash;25 g) were obtained from Janvier Labs (France). The animals were acclimatized for at least 7 days before experimentation and housed under specific pathogen-free (SPF) conditions in groups of 3\u0026ndash;5 per cage. The environment was controlled for temperature (22\u0026thinsp;\u0026plusmn;\u0026thinsp;2\u0026deg;C), humidity (45\u0026ndash;65%), and a 12:12 h light/dark cycle, with ad libitum access to food and water. All the cages were provided with nesting material and shelters to increase their welfare.\u003c/p\u003e \u003cp\u003eAll procedures complied with Spanish Royal Decree 53/2013 and the European Directive 2010/63/EU. The study protocol was approved by the Ethics Committee for Animal Experimentation of IIS Biogipuzkoa and the competent authority of Gipuzkoa (OH-23-25). The animals were checked daily during the first postoperative week and at least three times weekly thereafter. Humane endpoints included\u0026thinsp;\u0026gt;\u0026thinsp;15% body weight loss, persistent infection or necrosis, impaired mobility, or signs of sustained distress. Euthanasia was performed under deep isoflurane anaesthesia followed by cervical dislocation, following established welfare recommendations [\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e]. The study design, conduct and reporting followed the ARRIVE 2.0 recommendations for in vivo research [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e].\u003c/p\u003e\n\u003ch3\u003eMouse tail lymphedema model\u003c/h3\u003e\n\u003cp\u003eSecondary lymphedema was induced via a standardized and stereomicroscope-assisted surgical protocol. A narrow full-thickness circumferential strip of dorsal tail skin (approximately 2 mm wide) was carefully removed 1\u0026ndash;2 cm distal to the tail base to interrupt the superficial lymphatic collectors. To facilitate the identification of lymphatic vessels, 25 \u0026micro;L of 1% Evans blue dye (Sigma‒Aldrich) was injected intradermally into the distal tail, allowing selective visualization and subsequent ligation of marked lymphatic channels via fine Vannas microscissors. This technique ensures reproducible interruption of superficial lymphatics while preserving major blood vessels, thereby minimizing ischemic artifacts.\u003c/p\u003e \u003cp\u003eAnaesthesia was induced with 4\u0026ndash;5% isoflurane and maintained at 1\u0026ndash;2% in oxygen (0.5\u0026ndash;1 L/min). Perioperative analgesia was provided with subcutaneous meloxicam (0.2 mg/kg). No postoperative bandaging was applied to avoid irritation or self-mutilation behaviours. Lymphedema development was confirmed by a persistent increase in tail circumference measured 1 cm distal to the excision site, following established criteria.\u003c/p\u003e \u003cp\u003eThis protocol reliably induces a chronic defect in lymphatic drainage and results in progressive interstitial fluid accumulation, inflammatory activation, fibroadipose remodelling, and delayed endogenous lymphangiogenesis\u0026mdash;closely recapitulating key pathological features of human secondary lymphedema [\u003cspan additionalcitationids=\"CR21\" citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e]. Recent ultrastructural studies further confirmed that this model reproduces the intussusceptive and sprouting lymphangiogenesis patterns observed in human disease and in high-resolution murine regeneration models [\u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e]. For these reasons, the mouse tail lymphedema model remains the most widely adopted and technically robust platform for evaluating lymphatic dysfunction and testing regenerative therapies in vivo.\u003c/p\u003e\n\u003ch3\u003eCell isolation and characterization\u003c/h3\u003e\n\u003cp\u003e \u003cdiv class=\"BlockQuote\"\u003e \u003cp\u003e \u003cb\u003eMesenchymal stem cells (MSCs).\u003c/b\u003e \u003c/p\u003e \u003c/div\u003e \u003c/p\u003e \u003cp\u003eAll MSCs used in this study were murine (mouse) adipose-derived cells; no human-derived cells or human material were used.\u003c/p\u003e \u003cp\u003eAdipose-derived MSCs were obtained from the stromal vascular fraction of murine adipose tissue following enzymatic and mechanical dissociation. Briefly, excised adipose tissue was finely minced and subjected to collagenase type XI digestion (1 mg/mL, 60 min, 37\u0026deg;C) with gentle agitation to release stromal cells. The digested material was filtered through a 70-\u0026micro;m cell strainer to remove debris and centrifuged to isolate the stromal vascular pellet [\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eThe resulting cell fraction was resuspended and expanded directly onto the plastic plates with complete Mesencult medium (05513, StemCell) and cultured in a humidified incubator with a gas mixture of 5% CO2 at 37\u0026ordm;C. Cultures were maintained until they reached approximately 70\u0026ndash;80% confluence and were used at passage 3. Cell viability (\u0026gt;\u0026thinsp;90%) was confirmed prior to administration.\u003c/p\u003e \u003cp\u003e \u003cb\u003eLymphatic endothelial progenitor cells (LEPCs).\u003c/b\u003e \u003c/p\u003e \u003cp\u003eAt passage 3, MSCs were transferred to new dishes using 0.25% Trypsin-EDTA, seeded at 20,000 cells/cm\u0026sup2; with complete mouse endothelial cell medium (ECGM, M1168, Cell Biologics) and supplemented with 50 ng/mL of VEGF-C (752-VC, RD Systems). Cells were cultured under standard conditions (37\u0026deg;C with 5% CO₂) for 7 days.\u003c/p\u003e \u003cp\u003eSubsequently, the Pod⁺ cells were sorted, briefly expanded (\u0026le;\u0026thinsp;passage 3) and characterized by immunofluorescence and flow cytometry for canonical lymphatic markers (podoplanin and LYVE-1). Only cell preparations with viability\u0026thinsp;\u0026gt;\u0026thinsp;90% were used for in vivo administration.\u003c/p\u003e\n\u003ch3\u003eExperimental Groups, Randomization, and Blinding\u003c/h3\u003e\n\u003cp\u003eThe animals were allocated into three treatment groups:\u003c/p\u003e \u003cp\u003e \u003col\u003e \u003cspan\u003e \u003cli\u003e \u003cp\u003eControl group (n\u0026thinsp;=\u0026thinsp;4): intradermal PBS\u003c/p\u003e \u003c/li\u003e \u003c/span\u003e \u003cspan\u003e \u003cli\u003e \u003cp\u003eMSC group (n\u0026thinsp;=\u0026thinsp;7): 1 \u0026times; 10⁶ MSCs in PBS\u003c/p\u003e \u003c/li\u003e \u003c/span\u003e \u003cspan\u003e \u003cli\u003e \u003cp\u003eLEPC group (n\u0026thinsp;=\u0026thinsp;6): 1 \u0026times; 10⁶ LEPCs in PBS\u003c/p\u003e \u003c/li\u003e \u003c/span\u003e \u003c/ol\u003e \u003c/p\u003e \u003cp\u003eTreatments were administered on postoperative days 1 and 7. A total volume of 100 \u0026micro;L (cell suspension or PBS) was delivered through five intradermal microinjections evenly distributed around the excision site via a 30-gauge needle to ensure uniform coverage of the lymphatic injury zone.\u003c/p\u003e \u003cp\u003eRandomization was performed by an investigator not involved in the follow-up assessments. Tail measurements, IVIS/ICG imaging, and histological analysis were conducted by blinded examiners in accordance with best practices for minimizing experimental bias in animal research [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e, \u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e].\u003c/p\u003e \u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003ch2\u003eFunctional assessment\u003c/h2\u003e \u003cdiv id=\"Sec9\" class=\"Section3\"\u003e \u003ch2\u003eTail circumference measurements\u003c/h2\u003e \u003cp\u003eTail circumference was recorded before treatment (baseline) and weekly for 36 days via a digital micrometer. Measurements were consistently obtained 1 cm distal to the surgical site to ensure reproducibility. This approach represents a quantitative and widely implemented method for monitoring lymphedema progression and resolution in validated murine tail models [\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e].\u003c/p\u003e \u003c/div\u003e \u003c/div\u003e\n\u003ch3\u003eLymphatic function assessment\u003c/h3\u003e\n\u003cdiv id=\"Sec11\" class=\"Section2\"\u003e \u003ch2\u003eIndocyanine green (ICG) near-infrared fluorescence imaging (IVIS)\u003c/h2\u003e \u003cp\u003eThe quantitative assessment of lymphatic transport was performed via near-infrared ICG imaging. The mice received 25 \u0026micro;L of ICG (1 mg/mL in PBS; Sigma‒Aldrich) intradermally at the tail tip, followed by imaging at days 18 and 36 posttreatment via the IVIS SpectrumCT system (PerkinElmer, USA). Fluorescence was captured using 745/800 nm excitation/emission settings. Regions of interest (ROIs) were placed immediately distal and proximal to the surgical gap via Living Image software, and fluorescence intensity ratios (proximal/distal) were calculated to quantify tracer passage across the area of lymphatic disruption.\u003c/p\u003e \u003cp\u003eThis setup allows objective and reproducible quantification of lymphatic transport over time in the same animal and has recently been validated as a sensitive readout of lymphatic recovery in preclinical lymphedema models [\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e].\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec12\" class=\"Section2\"\u003e \u003ch2\u003eHistological analysis\u003c/h2\u003e \u003cp\u003eOn day 36, the mice were euthanized, and tail tissues encompassing the injury site were harvested for structural analysis. The samples were fixed in 4% paraformaldehyde (24 h at 4\u0026deg;C), decalcified, dehydrated, embedded in paraffin, and sectioned at 5 \u0026micro;m. The samples were stained with hematoxylin and eosin to help identify different types of cells and tissues and to obtain important information about the cellular structure of the tissue.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec13\" class=\"Section2\"\u003e \u003ch2\u003eFibrosis\u003c/h2\u003e \u003cp\u003eCollagen deposition was assessed by Picrosirius Red staining (Sigma‒Aldrich) and visualized under polarized light. Fibrotic area quantification was performed via ImageJ software (NIH, USA) with standardized color threshold algorithms. This technique provides sensitive detection of extracellular matrix remodelling and has been validated in recent preclinical lymphedema studies [\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e].\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003ch2\u003eLymphatic vessel density and cell engraftment\u003c/h2\u003e \u003cp\u003eImmunofluorescence staining was performed using antibodies against LYVE-1 (Abcam, UK), followed by Alexa Fluor\u0026ndash;labelled secondary antibodies and DAPI counterstaining. Images were acquired with a Leica SP8 confocal microscope. Lymphatic vessel density was quantified in five randomly selected high-power fields per sample via ImageJ [\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e].\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec15\" class=\"Section2\"\u003e \u003ch2\u003eStatistical analysis\u003c/h2\u003e \u003cp\u003eStatistical analyses were conducted via GraphPad Prism v10.0 (GraphPad Software, San Diego, CA, USA). The data are reported as the means\u0026thinsp;\u0026plusmn;\u0026thinsp;standard deviations (SDs). Normality was assessed with the Shapiro\u0026ndash;Wilk test [\u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e]. For comparisons between two groups, unpaired Student\u0026rsquo;s t tests were applied to normally distributed data, and the Mann\u0026ndash;Whitney U test was used for nonparametric distributions. Comparisons across three groups were performed with one-way ANOVA followed by Tukey\u0026rsquo;s post hoc test [\u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e31\u003c/span\u003e] or, when appropriate, with the Kruskal\u0026ndash;Wallis test and Dunn\u0026rsquo;s multiple comparisons [\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e]. The outliers were identified via Grubbs\u0026rsquo; test [\u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e]. Sample size selection was guided by prior studies using the murine tail lymphedema model [\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e, \u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e] and aligned with recommendations for rigorous preclinical research [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e, \u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e]. Statistical significance was set at p\u0026thinsp;\u0026lt;\u0026thinsp;0.05.\u003c/p\u003e \u003c/div\u003e"},{"header":"Results","content":"\u003cdiv id=\"Sec17\" class=\"Section2\"\u003e\n \u003ch2\u003eTail swelling over time (tail diameter)\u003c/h2\u003e\n \u003cp\u003eTail diameter was measured weekly for 36 days following lymphedema induction. All animals developed increased tail thickness after surgery, with group-dependent differences in the magnitude and evolution of swelling. Baseline diameters were comparable among groups: control (3.13\u0026thinsp;\u0026plusmn;\u0026thinsp;0.08 mm), MSC (3.18\u0026thinsp;\u0026plusmn;\u0026thinsp;0.10 mm), and LEPC (3.19\u0026thinsp;\u0026plusmn;\u0026thinsp;0.11 mm). Peak swelling occurred on Day 18 in the control group (6.09\u0026thinsp;\u0026plusmn;\u0026thinsp;0.55 mm), whereas MSC-treated animals reached a lower maximum diameter (5.33\u0026thinsp;\u0026plusmn;\u0026thinsp;0.36 mm). LEPC-treated mice exhibited the mildest peak swelling (5.25\u0026thinsp;\u0026plusmn;\u0026thinsp;0.23 mm). From Day 18 onwards, all groups showed a partial reduction in tail diameter, with the LEPC group exhibiting the lowest final diameter on Day 36 (4.61\u0026thinsp;\u0026plusmn;\u0026thinsp;0.47 mm), followed by the MSC (5.12\u0026thinsp;\u0026plusmn;\u0026thinsp;0.59 mm) and control (5.59\u0026thinsp;\u0026plusmn;\u0026thinsp;0.55 mm) groups (Fig.\u0026nbsp;1).\u003c/p\u003e\n \u003cp\u003e\u003cstrong\u003eFigure 1. Tail diameter over time across treatment groups. Compared with the MSC and control groups, the LEPC-treated group showed a milder and more transient increase in tail swelling.\u003c/strong\u003e\u003c/p\u003e\n \u003cp\u003eTo better characterize treatment effects, we analysed the difference between each animal\u0026rsquo;s peak tail diameter and its baseline value (\u0026Delta;Max\u0026ndash;Day 0). The control group showed the greatest increase (3.33\u0026thinsp;\u0026plusmn;\u0026thinsp;0.46 mm), followed by the MSC (2.68\u0026thinsp;\u0026plusmn;\u0026thinsp;0.47 mm) and LEPC (2.45\u0026thinsp;\u0026plusmn;\u0026thinsp;0.51 mm) groups. The difference between the control and LEPC groups was statistically significant (p\u0026thinsp;=\u0026thinsp;0.032), suggesting reduced swelling progression in LEPC-treated mice.\u003c/p\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec18\" class=\"Section2\"\u003e\n \u003ch2\u003eHistological assessment of tissue remodeling (fibrosis and non-tissue/void spaces)\u003c/h2\u003e\n \u003cp\u003eInterstitial tissue composition was quantified using the WSI Sirius Red Manual workflow. Within each region of interest (ROI), Sirius Red\u0026ndash;positive area and non-tissue/void area were segmented and expressed as percentage of the total ROI.\u003c/p\u003e\n \u003cp\u003eQuantitative analysis showed a significantly lower non-tissue/void area in LEPC-treated mice (7.37\u0026thinsp;\u0026plusmn;\u0026thinsp;2.87%) compared with MSC-treated (15.87\u0026thinsp;\u0026plusmn;\u0026thinsp;2.64%) and control animals (19.62\u0026thinsp;\u0026plusmn;\u0026thinsp;9.31%). No significant differences were observed between the MSC and control groups (p\u0026thinsp;=\u0026thinsp;0.548). The non-tissue/void area was significantly reduced in the LEPC group compared with both control (p\u0026thinsp;=\u0026thinsp;0.048) and MSC groups (p\u0026thinsp;=\u0026thinsp;0.002). Representative histological images are shown in Fig.\u0026nbsp;2. These histological results were consistent with the reduced tail swelling observed in LEPC-treated animals during longitudinal follow-up.\u003c/p\u003e\n \u003cp\u003e\u003cstrong\u003eFigure 2. Histological assessment of tissue remodelling.\u003c/strong\u003e\u003c/p\u003e\u003cspan\u003e\n \u003cp\u003e\u003cstrong\u003ea) Representative macroscopic view of the mouse tail on day 36 post-surgery.\u003c/strong\u003e\u003c/p\u003e\n \u003c/span\u003e \u003cspan\u003e\n \u003cp\u003e\u003cstrong\u003eb) Hematoxylin and eosin\u0026ndash;stained section of the injury area.\u003c/strong\u003e\u003c/p\u003e\n \u003c/span\u003e \u003cspan\u003e\n \u003cp\u003e\u003cstrong\u003ec) Picrosirius Red\u0026ndash;stained section highlighting collagen deposition and non-tissue/void spaces. Scale bar, 200 \u0026micro;m.\u003c/strong\u003e\u003c/p\u003e\n \u003c/span\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec19\" class=\"Section2\"\u003e\n \u003ch2\u003eLymphatic function assessment\u003c/h2\u003e\n \u003cp\u003eNear-infrared ICG imaging provides a dynamic evaluation of lymphatic transport. On Day 18, none of the groups demonstrated tracer passage across the surgical site. By Day 36, LEPC-treated mice exhibited clear proximal-to-distal tracer migration (100% of the animals), as reflected by significantly higher fluorescence intensity ratios than those of the MSC-treated (43% of the animals, 3 of 7) and PBS-treated controls (0% of the animals) (\u003cem\u003ep\u003c/em\u003e\u0026thinsp;\u0026lt;\u0026thinsp;0.05).\u003c/p\u003e\n \u003cp\u003eRepresentative IVIS images on Day 36 illustrate these differences (Fig.\u0026nbsp;3a), with LEPC-treated animals showing visible tracer progression, whereas MSC-treated mice displayed minimal signal and control mice showing no transit.\u003c/p\u003e\n \u003cp\u003e\u003cstrong\u003eFigure 3. Lymphatic function and structural regeneration.\u003c/strong\u003e\u003c/p\u003e\u003cspan\u003e\n \u003cp\u003e\u003cstrong\u003ea) Representative IVIS\u0026ndash;ICG near-infrared fluorescence images on day 36 post-surgery showing tracer transit across the excision site in LEPC-treated mice compared with MSC-treated and PBS control animals. Arrows indicate the excision site (region of lymphatic disruption) in all groups\u003c/strong\u003e\u003c/p\u003e\n \u003c/span\u003e \u003cspan\u003e\n \u003cp\u003e\u003cstrong\u003eb) Representative confocal images of LYVE-1 immunostaining in the excision region on day 36. LEPC-treated tissues exhibit a higher density of organized LYVE-1⁺ lymphatic capillaries compared with MSC-treated and PBS control tissues. Arrows indicate lymphatic vessels. Scale bar, 200 \u0026micro;m.\u003c/strong\u003e\u003c/p\u003e\n \u003c/span\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec20\" class=\"Section2\"\u003e\n \u003ch2\u003eLymphatic Vessel Regeneration\u003c/h2\u003e\n \u003cp\u003eImmunostaining of tissue sections confirmed the increased density of LYVE-1⁺ capillaries and the presence of organized lymphatic networks in the LEPC-treated group (Fig.\u0026nbsp;3b). In contrast, MSC and control tissues displayed sparse and disorganized lymphatic structures. These findings indicate that LEPC treatment enhances both structural regeneration and functional restoration of lymphatic drainage.\u003c/p\u003e\n\u003c/div\u003e"},{"header":"Discussion","content":"\u003cp\u003eIn this study, we show that adipose-derived lymphatic endothelial progenitor cells (LEPCs) enhance both structural and functional recovery in a validated murine model of secondary lymphedema. Across independent readouts, LEPC-treated animals showed a consistent improvement in tail thickness, ICG tracer transport, and lymphatic vessel density compared with both PBS and MSC-treated groups. Overall, LEPC treatment produced concordant functional recovery (IVIS\u0026ndash;ICG transit and tail thickness) together with increased LYVE-1⁺ vessel density, supporting a true regenerative effect in this model. The concordance between functional imaging, morphometric follow-up, and histological findings supports a true biological treatment effect rather than a measurement-specific artifact.\u003c/p\u003e \u003cdiv id=\"Sec22\" class=\"Section2\"\u003e \u003ch2\u003eMethodological considerations supporting translational relevance\u003c/h2\u003e \u003cp\u003eAn important methodological aspect of this study is the use of IVIS-based near-infrared ICG imaging as the main functional readout. Conventional assessments, such as Evans blue transit, provide limited sensitivity and cannot be performed longitudinally. In contrast, IVIS enables dynamic, repeated, and quantitative evaluation of lymphatic flow within the same animal, thereby improving reproducibility and translational value. Recent studies support the use of longitudinal near-infrared ICG imaging as a sensitive functional endpoint in preclinical lymphatic models [\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e]. In our dataset, IVIS clearly separated the functional effects of LEPCs from those of MSCs and controls, highlighting its utility for preclinical therapeutic screening.\u003c/p\u003e \u003cp\u003eThe murine tail model employed in this study remains one of the most widely used and reproducible systems to evaluate lymphatic dysfunction and regeneration [\u003cspan additionalcitationids=\"CR21\" citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e]. Its defined injury pattern, predictable edema kinetics, and compatibility with high-resolution imaging provide a stable platform for testing regenerative interventions. The integration of IVIS-ICG imaging into this model further enhances sensitivity and allows comprehensive evaluation of lymphatic recovery.\u003c/p\u003e \u003cdiv id=\"Sec23\" class=\"Section3\"\u003e \u003ch2\u003eRegenerative advantages of LEPCs over MSC- and ADSC-based therapies\u003c/h2\u003e \u003cp\u003ePrevious studies using MSCs or adipose-derived regenerative cell populations have demonstrated beneficial paracrine effects on lymphangiogenesis and tissue remodelling in experimental lymphedema [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e, \u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e]. Enhanced approaches such as Hippo\u0026ndash;YAP modulation have further amplified the lymphangiogenic potential of adipose-derived stromal cells in murine tail lymphedema [\u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e]. Our results expand this landscape by showing that LEPCs can induce substantial lymphatic regeneration without requiring gene engineering.\u003c/p\u003e \u003cp\u003eThe podoplanin-positive progenitor population used here was associated with improved lymphatic repair, as shown by reduced edema and reconstitution of LYVE-1⁺ and VEGFR-3⁺ lymphatic networks. These findings support the hypothesis that LEPC-mediated recovery may involve paracrine activity and/or direct endothelial incorporation, although the relative contribution of each mechanism cannot be determined in the absence of dedicated cell-tracking approaches. The pattern of structural remodelling observed is consistent with intussusceptive lymphangiogenesis, as recently described in high-resolution imaging studies [\u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e].\u003c/p\u003e \u003c/div\u003e \u003c/div\u003e \u003cdiv id=\"Sec24\" class=\"Section2\"\u003e \u003ch2\u003eMicroenvironmental barriers and potential combinatorial strategies\u003c/h2\u003e \u003cp\u003eSecondary lymphedema is characterized by a fibrotic and inflammatory milieu that can hinder tissue repair and limit long-term recovery [\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e, \u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e]. In our study, LEPC-treated tissues presented a reduced Sirius Red\u0026ndash;positive area at day 36, suggesting that improving lymphatic transport may help attenuate fibrotic remodelling within the injury zone. In parallel, the lower non-tissue/void area observed histologically is consistent with reduced interstitial expansion and tissue disruption in the treated group.\u003c/p\u003e \u003cp\u003eEmerging evidence suggests that targeting extracellular matrix remodelling may potentiate regenerative strategies in secondary lymphedema. Inhibition of uPARAP has been reported to improve lymphatic architecture and reduce fibrosis in preclinical models [\u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e]; mechanotransduction pathways such as Hippo\u0026ndash;YAP/TAZ may also influence lymphatic endothelial behaviour [\u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e]. In our study, the reduced fibrotic signal in LEPC-treated tissues supports the relevance of these microenvironmental constraints.\u003c/p\u003e \u003cdiv id=\"Sec25\" class=\"Section3\"\u003e \u003ch2\u003eParacrine cooperation and integration with next-generation therapeutics\u003c/h2\u003e \u003cp\u003eReported synergy between MSCs and LEPCs suggests that paracrine support may enhance progenitor survival and lymphangiogenic activity; for example, MSC-derived IGF-1 has been linked to improved LEPC function via PI3K/Akt/mTOR signalling and superior outcomes with cotherapy compared with single-cell approaches [\u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e]. Other \u0026ldquo;next-generation\u0026rdquo; combination strategies\u0026mdash;such as Apelin\u0026thinsp;+\u0026thinsp;VEGF-C delivery via mRNA [\u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e37\u003c/span\u003e] and platelet-derived extracellular vesicles [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e]\u0026mdash;further support a multimodal regenerative framework. Together, these data raise the possibility that integrating LEPCs with paracrine-active or microenvironment-modulating interventions could improve durability beyond the functional benefit observed with LEPCs alone in our model, although this was not tested here.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec26\" class=\"Section3\"\u003e \u003ch2\u003ePositioning of LEPC therapy within the evolving lymphatic regeneration landscape\u003c/h2\u003e \u003cp\u003eLEPCs represent a mechanistically rational strategy for secondary lymphedema given their lymphatic-lineage orientation. In our study, LEPC treatment was associated with concordant functional improvement (ICG transport and tail thickness) together with increased LYVE-1⁺ lymphatic vessel density, supporting further translational evaluation. Key next steps include standardizing LEPC isolation and characterization workflows [\u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e], testing whether combination approaches improve performance in fibrotic tissue, and extending follow-up to assess durability, biodistribution, and safety.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec27\" class=\"Section3\"\u003e \u003ch2\u003eLimitations\u003c/h2\u003e \u003cp\u003eSeveral limitations of this study must be acknowledged. First, the sample size, although consistent with established murine tail lymphedema studies [\u003cspan additionalcitationids=\"CR21\" citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e], remains modest and may limit the statistical power for detecting small or late therapeutic effects. Increasing the group size, especially for MSCs, could help confirm the differences observed in this work.\u003c/p\u003e \u003cp\u003eSecond, the tail model\u0026mdash;despite being one of the most reproducible and widely used platforms for lymphatic research\u0026mdash;does not fully recapitulate the anatomical and biomechanical complexity of human limb lymphedema. The absence of muscle compartments, fascia, and weight-bearing forces limits the extrapolation of certain mechanotransductive phenomena and stromal interactions.\u003c/p\u003e \u003cp\u003eThird, although IVIS-ICG provides sensitive and longitudinal functional assessment, it reflects tracer transit rather than true lymphatic pumping or valve competence. Additional techniques, such as near-infrared dynamic contractility imaging or lymphoscintigraphy, could strengthen functional characterization.\u003c/p\u003e \u003cp\u003eFourth, the identification of LEPCs relies on classical lymphatic markers (LYVE-1, VEGFR-3, and PROX1). While consistent with current standards, the heterogeneity of lymphatic progenitor populations is increasingly recognized [\u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e, \u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e]. Single-cell profiling or lineage-tracing approaches would more precisely define the subpopulations responsible for the regenerative effect.\u003c/p\u003e \u003cp\u003eFifth, LEPC\u0026ndash;host interactions within the fibrotic microenvironment have not been investigated mechanistically. The contributions of macrophages, fibroblasts, and Th2-skewed inflammation to graft success or failure remain largely unexplored in this setting.\u003c/p\u003e \u003cp\u003eFinally, long-term durability and potential off-target effects could not be assessed because of the 36-day follow-up. Chronic models\u0026mdash;beyond 8\u0026ndash;12 weeks\u0026mdash;are required to capture lymphatic remodelling, stabilize fibrosis, and maintain the persistence of engrafted cells.\u003c/p\u003e \u003c/div\u003e \u003c/div\u003e \u003cdiv id=\"Sec28\" class=\"Section2\"\u003e \u003ch2\u003eFuture directions\u003c/h2\u003e \u003cp\u003eFuture work should prioritize mechanistic validation and translational strengthening. Key next steps include cell-tracking approaches to define the relative contribution of paracrine effects versus endothelial incorporation, extending follow-up to assess durability and safety, and testing clinically aligned models (e.g., hindlimb and/or irradiation-associated lymphedema). In parallel, standardization of LEPC isolation/characterization will be essential to improve reproducibility and support future translation [\u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e].\u003c/p\u003e \u003c/div\u003e"},{"header":"Conclusions","content":"\u003cp\u003eOur findings show that LEPC transplantation improves multiple hallmarks of secondary lymphedema, outperforming MSCs and PBS controls across functional, morphometric, and histological endpoints. The integration of IVIS\u0026ndash;ICG imaging with quantitative tail measurements and lymphatic marker analysis provides convergent evidence of treatment-associated lymphatic recovery in this preclinical model.\u003c/p\u003e \u003cp\u003eTogether, these results support further investigation of LEPC-based interventions as a regenerative strategy for secondary lymphedema, including studies designed to define mechanisms, durability, and translational feasibility.\u003c/p\u003e"},{"header":"Abbreviations","content":"\u003cp\u003eADSCs: Adipose-derived stem cells\u003c/p\u003e\n\u003cp\u003eARRIVE: Animal Research: Reporting of In Vivo Experiments\u003c/p\u003e\n\u003cp\u003eECGM: Endothelial cell growth medium\u003c/p\u003e\n\u003cp\u003eH\u0026amp;E: Hematoxylin and eosin\u003c/p\u003e\n\u003cp\u003eICG: Indocyanine green\u003c/p\u003e\n\u003cp\u003eIL: Interleukin\u003c/p\u003e\n\u003cp\u003eIVIS: In vivo imaging system\u003c/p\u003e\n\u003cp\u003eLEPCs: Lymphatic endothelial progenitor cells\u003c/p\u003e\n\u003cp\u003eLVA: Lymphaticovenular anastomosis\u003c/p\u003e\n\u003cp\u003eLYVE-1: Lymphatic vessel endothelial hyaluronan receptor 1\u003c/p\u003e\n\u003cp\u003eMSCs: Mesenchymal stem cells\u003c/p\u003e\n\u003cp\u003ePBS: Phosphate-buffered saline\u003c/p\u003e\n\u003cp\u003ePROX1: Prospero homeobox 1\u003c/p\u003e\n\u003cp\u003eROI: Region of interest\u003c/p\u003e\n\u003cp\u003eSPF: Specific pathogen-free\u003c/p\u003e\n\u003cp\u003eUPV/EHU: University of the Basque Country\u003c/p\u003e\n\u003cp\u003eVEGFR-3: Vascular endothelial growth factor receptor 3\u003c/p\u003e\n\u003cp\u003eVLNT: Vascularized lymph node transfer\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eEthics approval and consent to participate\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eEthics approval and consent to participate: All animal procedures were authorized by the Diputaci\u0026oacute;n Foral de Gipuzkoa ( Provincial Council of Gipuzkoa) under the approved project \u0026ldquo;Capacidad regenerativa de precursores linfangiog\u0026eacute;nicos para el tratamiento de linfedema secundario en modelo murino (OH-23-25)\u0026rdquo; (administrative identification code PRO-AE-SS-311; approval date 13 December 2023). All procedures complied with Spanish Royal Decree 53/2013 and the European Directive 2010/63/EU.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eUse of artificial intelligence\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare that artificial intelligence\u0026ndash;based tools were used exclusively for language editing, grammar correction, and improvement of clarity and structure in selected sections of the manuscript. No artificial intelligence tools were used for data generation, data analysis, figure creation, or interpretation of results. All scientific content, analyses, and conclusions were verified and approved by the authors, who take full responsibility for the integrity and accuracy of the work.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eConsent for publication\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eNot applicable.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAvailability of data and materials\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe datasets generated and/or analysed during the current study are available from the corresponding author on reasonable request.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eCompeting interests\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare that they have no competing interests.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunding\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis work received no specific funding.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthors\u0026rsquo; contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eIJ and HL conceived and designed the study, performed the experiments, acquired the data (including tail measurements and imaging), analysed and interpreted the results, and wrote the manuscript. MTI performed the biostatistical analysis of the tail measurement dataset and critically revised the manuscript. IA established and performed the LEPC isolation workflow and contributed to cell preparation and characterization. AI, ID and AL contributed to manuscript revision and approved the final version. All authors read and approved the final manuscript.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAcknowledgements\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors thank the Animal Facility and Imaging Core of IIS Biogipuzkoa for technical support and the Histology Service of CIMA (Centro de Investigaci\u0026oacute;n M\u0026eacute;dica Aplicada, Universidad de Navarra, Pamplona, Spain) for assistance with tissue processing and immunofluorescence imaging. The authors critically reviewed and approved the final manuscript and take full responsibility for the integrity and accuracy of the work.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n \u003cli\u003eDessources K, Aviki E, Leitao MM Jr. Lower extremity lymphedema in patients with gynecologic malignancies. Int J Gynecol Cancer. 2020 Feb;30(2):252\u0026ndash;260. doi:10.1136/ijgc-2019-001032.\u003c/li\u003e\n \u003cli\u003eCormier JN, Askew RL, Mungovan KS, Xing Y, Ross MI, Armer JM. Lymphedema beyond breast cancer: a systematic review and meta-analysis of cancer-related secondary lymphedema. Cancer. 2010;116(22):5138\u0026ndash;5149. doi:10.1002/cncr.25458.\u003c/li\u003e\n \u003cli\u003eDiSipio T, Rye S, Newman B, Hayes S. Incidence of unilateral arm lymphoedema after breast cancer: a systematic review and meta-analysis. 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Front Genet. 2025;16:1584095. doi:10.3389/fgene.2025.1584095.\u003c/li\u003e\n \u003cli\u003eCreff J, Lamaa A, Benuzzi E, Balzan E, Pujol F, Draia-Nicolau T, Bonnevie T, Boisset N, Lemaigre F, Chuvin N, Clavel L, Carpentier G, Pratx S, Duhamel A, Bidault G, Jude B, Soncin F. Apelin-VEGF-C mRNA delivery as therapeutic for the treatment of secondary lymphedema. EMBO Mol Med. 2024 Feb;16(2):386\u0026ndash;415. doi:10.1038/s44321-023-00017-7.\u003c/li\u003e\n \u003cli\u003eVachon L, Jean G, Milasan A, Ganivet A, Monge-Rojas J, Rolland-Yasuhara M, Cartier R, Martel C. Platelet extracellular vesicles preserve lymphatic endothelial cell integrity and enhance lymphatic vessel function. Commun Biol. 2024 Aug 11;7(1):975. doi:10.1038/s42003-024-06675-8.\u003c/li\u003e\n \u003cli\u003eSaha S, Graham F, Knopp J, Patzke C, Hanjaya-Putra D. Robust differentiation of human pluripotent stem cells into lymphatic endothelial cells using transcription factors. Cells Tissues Organs. 2024;213(6):464\u0026ndash;474. doi:10.1159/000539699.\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":true,"highlight":"","institution":"","isAcceptedByJournal":false,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true},"keywords":"Secondary lymphedema, regenerative medicine, lymphatic endothelial progenitor cells, mouse tail lymphedema model, Lymphangiogenesis","lastPublishedDoi":"10.21203/rs.3.rs-8524204/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-8524204/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003ch2\u003eBackground\u003c/h2\u003e \u003cp\u003eSecondary lymphedema is a chronic and disabling condition with scarce disease-modifying treatments, particularly after oncologic surgery or radiotherapy. Current management remains largely palliative rather than restorative. Regenerative medicine strategies are emerging as promising alternatives, and lymphatic endothelial progenitor cells (LEPCs) may support reconstruction of damaged lymphatic networks.\u003c/p\u003e\u003ch2\u003eMethods\u003c/h2\u003e \u003cp\u003eSecondary lymphedema was induced in C57BL/6J mice using standardized tail skin excision with interruption of superficial lymphatics. Animals received intradermal phosphate-buffered saline (PBS), mesenchymal stem cells (MSCs), or adipose-derived LEPCs on days 1 and 7 post-surgery. Lymphatic function was monitored longitudinally by IVIS-based near-infrared indocyanine green (ICG) imaging and serial tail circumference measurements. Lymphatic vessels were assessed by LYVE-1 immunofluorescence, and fibrosis by Picrosirius Red staining.\u003c/p\u003e\u003ch2\u003eResults\u003c/h2\u003e \u003cp\u003eLEPC treatment markedly improved functional lymphatic recovery, as evidenced by faster indocyanine green (ICG) transit on near-infrared imaging, and significantly reduced tail swelling compared with both the MSC and PBS groups. Immunofluorescence analysis revealed a higher density of LYVE-1⁺ lymphatic vessels in LEPC-treated tissue, supporting enhanced lymphatic network reconstruction.\u003c/p\u003e\u003ch2\u003eConclusions\u003c/h2\u003e \u003cp\u003eLEPC administration significantly enhanced lymphatic repair and functional recovery in a murine model of secondary lymphedema. These findings reinforce LEPCs as a biologically targeted and potentially disease-modifying therapeutic option that deserves further translational evaluation.\u003c/p\u003e","manuscriptTitle":"Therapeutic potential of lymphatic endothelial progenitor cells in secondary lymphedema: a preclinical murine study","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2026-02-11 16:46:14","doi":"10.21203/rs.3.rs-8524204/v1","editorialEvents":[{"type":"communityComments","content":0}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"208208d3-a8f4-4822-9f7c-adcab968c5a8","owner":[],"postedDate":"February 11th, 2026","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"posted","subjectAreas":[],"tags":[],"updatedAt":"2026-02-17T16:25:48+00:00","versionOfRecord":[],"versionCreatedAt":"2026-02-11 16:46:14","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-8524204","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-8524204","identity":"rs-8524204","version":["v1"]},"buildId":"XKTyCvWXoU3ODBz1xrDgd","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}