A novel approach for reliable differentiation of lymphatic endothelial progenitor cells in vitro

A novel approach for reliable differentiation of lymphatic endothelial progenitor cells in vitro | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (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];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results A novel approach for reliable differentiation of lymphatic endothelial progenitor cells in vitro View ORCID Profile Inazio Arriola-Alvarez , View ORCID Profile Maria Rodriguez-Hidalgo , View ORCID Profile Hector Lafuente , View ORCID Profile Sonia Alonso-Martin , View ORCID Profile Ander Izeta doi: https://doi.org/10.1101/2025.09.23.677051 Inazio Arriola-Alvarez 1 Biogipuzkoa Health Research Institute, Stem Cells & Aging Group , Donostia/San Sebastián, 20014, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Inazio Arriola-Alvarez Maria Rodriguez-Hidalgo 1 Biogipuzkoa Health Research Institute, Stem Cells & Aging Group , Donostia/San Sebastián, 20014, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Maria Rodriguez-Hidalgo Hector Lafuente 1 Biogipuzkoa Health Research Institute, Stem Cells & Aging Group , Donostia/San Sebastián, 20014, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Hector Lafuente Sonia Alonso-Martin 1 Biogipuzkoa Health Research Institute, Stem Cells & Aging Group , Donostia/San Sebastián, 20014, Spain 2 CIBERNED, ISCIII (CIBER, Carlos III Institute, Spanish Ministry of Sciences and Innovation) , 28031, Madrid, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Sonia Alonso-Martin Ander Izeta 1 Biogipuzkoa Health Research Institute, Stem Cells & Aging Group , Donostia/San Sebastián, 20014, Spain 3 Donostia University Hospital , Advanced Therapies Unit, Donostia/San Sebastian, 20014, Spain 4 Tecnun-University of Navarra, School of Engineering , Donostia/San Sebastian, 20009, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Ander Izeta For correspondence: ANDER.IZETAPERMISAN{at}osakidetza.eus Abstract Full Text Info/History Metrics Preview PDF Abstract The lymphatic system plays critical roles in fluid homeostasis, immune regulation, and lipid transport, making its dysfunction a contributor to numerous pathological conditions. Lymphatic tissue engineering aims to develop therapeutic strategies for lymphatic regeneration and repair, depending on the availability of suitable cell sources for lymphatic endothelial cell (LEC) generation. Key cell sources explored for lymphatic tissue engineering include the dermis, bone marrow and stromal vascular fraction. To our knowledge, no existing differentiation protocols can accurately generate lymphatic endothelial progenitor cells (LEPCs). This study presents a novel in vitro differentiation protocol for generating LEPCs from stromal vascular fraction, dermis and bone marrow-derived mesenchymal stem cells (MSCs). The feasibility of using them as tissue sources was first evaluated leading to dermal cells exclusion due to low yield post-isolation, and highlighting the limited differentiation efficiency of bone marrow MSCs. However, the stromal vascular fraction emerged as the optimal source, exhibiting robust LEPCs differentiation and superior scalability. In vitro characterization confirmed that LEPCs maintain a lymphatic endothelial phenotype, exhibiting high migratory ability, robust angiogenic structure formation, and consistent expression of key lymphatic markers in both 2D and 3D cultures, as validated by RNA-Seq analysis. The protocol provides a consistent platform for studying lymphatic endothelial biology and potential applications in regenerative medicine and therapeutic angiogenesis. Introduction The lymphatic system plays a critical role in maintaining tissue fluid homeostasis, immune response regulation, and lipid metabolism. 1 Dysfunction in this system is implicated in a range of pathological conditions, including lymphedema, cancer metastasis, obesity, and neurodegenerative disorders. 2 , 3 LECs form the structural foundation of the lymphatic system, constructing the vessel walls throughout the entire network. The system’s strongly hierarchical structure originates during embryogenesis, when a small subset of endothelial cells (ECs) in the cardinal vein differentiate into lymphatic lineage and begin expressing crucial markers like PDPN, LYVE1, and VEGFR3 around embryonic day 9.5 (E9.5). 4 These cells, primarily guided by VEGF-C signaling via the VEGF-C/VEGFR3 pathway, develop into lymph sacs, which—along with non – venous origin LECs—form the mature lymphatic system. 5 At this stage, additional factors such as IGF2, angiopoietins, HGF and FGF2 are also required for proper lymphatic development and maintenance. 6 – 8 PDPN, a transmembrane glycoprotein, is critical for lymphatic vessel formation and function, playing roles in cell migration and maintaining lymphatic endothelial integrity. 9 , 10 LYVE1 is involved in hyaluronan transport and serves as a key marker for LECs, indicating proper differentiation and functional potential of the cells. 11 VEGFR3 is a primary receptor mediating VEGF-C signalling as described above, essential for lymphangiogenesis and the proliferation, migration, and survival of LECs. 12 , 13 Recently, the limited but growing evidence supporting the importance of the lymphatic system in maintaining the tissue homeostasis and its implication on the diversity of diseases has gathered some interest, resulting in the development of few therapeutic approaches to stimulate lymphangiogenesis, i.e., the formation of new lymphatic vessels from preexisting ones. 14 – 16 One of the strategies followed for this task is cell therapy. In this context, stem cells have gained significant interest for their potential in treating various diseases. These cells differ in potency, or their ability to differentiate into multiple cell types, with totipotent cells found only in embryos and adult stem cells, with lower potency –tissue specific–, residing in tissue niches to aid in repair and regeneration. 17 Research has focused on harnessing adult stem cells for regenerative medicine, particularly MSCs. These are distinguished by their ability to form colonies, express specific surface markers, and maintain self-renewal while retaining differentiation potential, making them key to tissue engineering and therapeutic applications. 18 – 20 Among these, bone marrow-derived (BM-MSCs), dermis-derived (D-MSCs), adipose tissue-derived (AD-MSCs) and umbilical cord-derived MSCs (UC-MSCs) have shown potential for lymphatic tissue engineering, each with distinct advantages. 21 BM-MSCs, traditionally the most studied MSC type, have been explored for their ability to support lymphangiogenesis through paracrine signaling and direct differentiation. 22 – 25 However, their invasive collection process and lower yield limit their practicality for large-scale applications. 26 D-MSCs are a very invasive option but exhibit promising lymphangiogenic properties. 27 , 28 Their role in supporting lymphatic endothelial cell differentiation has been noted, particularly through interactions with extracellular matrix components that resemble the native lymphatic niche. 29 AD-MSCs offer a highly accessible and abundant cell source, with minimal donor morbidity. 30 – 32 They have gained increasing attention for their potential in lymphatic tissue engineering due to their robust proliferation capacity, differentiation potential, and ability to secrete pro-lymphangiogenic factors, such as VEGF-C. 12 , 33 , 34 UC-MSCs provide benefits, including easier and less invasive collection, enhanced proliferation capacity, and greater potency. 35 , 36 Cord-blood endothelial cell colony-forming cells (ECFCs), have been recognized for their role in promoting lymphangiogenesis through the contribution of the lymphatic ECFC subpopulation they contain. 37 , 38 Despite these promising attributes, tissue engineering has not put hardly any interest in their use for lymphatic tissue engineering. 21 Overall, lymphatic tissue engineering lacks of adequate approaches to reliably differentiate MSCs into functional LECs in vitro . Therefore, this study presents a novel differentiation protocol for deriving LEPCs from MSCs, focusing on dermis, bone marrow (BM) and adipose tissue-stromal vascular fraction (SVF) as the main cell sources. By leveraging key growth factors and signaling pathways involved in lymphatic embryogenesis and postnatal lymphangiogenesis, we have established a robust and reproducible method for generating functional LEPCs. Furthermore, in this work, we have proven that SVF-derived LEPCs display the best differentiation potential, and coupled with their ease of access and proliferation capacity, pose as a great alternative for LEPC differentiation in the clinical settings. This work not only provides a valuable tool for advancing lymphatic disease modeling and therapeutic applications but also highlights the broader potential of MSCs in lymphatic tissue engineering. Materials and methods Isolation of SVF, BM and dermis SVF and BM were extracted from C57BL/6 mice (8 week old male and female). Briefly, adipose tissue was scraped off the inner part of the back skin. Then, it was minced into approximately 1 mm 2 pieces, and incubated under 180 rpm agitation in a collagenase type IA solution (1.000 u/mL, C2674, Sigma) for approximately 25 minutes. Later, the digested adipose tissue solution was inactivated and centrifuged at 1.500 rpm for 5 minutes to separate into layers. The top two layers were removed and the bottom layer (containing the SVF’s cells) was washed twice with 2% fetal bovine serum (FBS, 10270-106, Gibco) in D-PBS (14190-094, Gibco), filtered at 100 µm and plated for culture. Besides, the femur, the tibia and the humerus were extracted from the mice, and the epiphysis of the bones were cut leaving an open wound. Afterwards, the bones were placed in a 0.5 mL tube with a small hole in the bottom. Then, these two were placed inside a 1.5 mL tube and centrifuged at 10.000 rpm for 10 seconds. The BM was flushed out into the 1.5 mL whilst the empty bones were retained in the smaller tubes. The BM pellet was washed twice again with 2% FBS and plated for culture. Skin specimens clean of blood vessels and adipose tissue were minced into 1-2 mm² fragments. Digestion was performed in 1 mg/ml collagenase XI at 37°C for ∼60 min until tissue fragments became pale and easily dispersible. Digestion was stopped with cold wash medium. After centrifugation (300g, 7 min, 4 °C), pellets were mechanically dissociated in wash medium via pipette grinding (2-3 min/repetition) until complete dissociation (milky suspension). Cells were filtered (40 μm), centrifuged, and resuspended in proliferation medium. Viability was assessed by trypan blue exclusion. Cell culture Isolated SVF-derived MSCs were directly seeded onto the plastic plates with complete Mesencult medium (05513, StemCell) and cultured in a humidified incubator with a gas mixture of 5% CO 2 at 37°C. Bone marrow MSCs were also cultured in complete Mesencult medium in a humidified incubator with a gas mixture of 1% O 2 , 5% CO 2 at 37°C. MSCs were seeded at 10.000 cell/cm 2 and cultured until passage 3. Once cells had reached 80% confluence, they were transferred to new dishes using 0.25% Trypsin-EDTA (25200-072, Gibco). Human dermal lymphatic endothelial cells (HDLEC, C-14021, PromoCell) were cultured according to the providers’ guidelines. Adipogenesis MSCs isolated from SVF and BM were expanded to passage 2 using standard subculture techniques. At 90% confluence, cells were trypsinized and seeded at 10,000 cells/cm² in 12-well plates. Differentiation was induced using the MesenCult Adipogenic Differentiation Kit (05507, StemCell) under hypoxia (1% O₂, 5% CO₂, N₂-balanced) in a tri-gas incubator. The medium was refreshed every 3–4 days, with differentiation periods of 14 days (SVF-derived MSCs) or 7 days (BM-derived MSCs), as per the manufacturer’s guidelines. Post-differentiation, cells were fixed with Histofix (256462, Panreac AppliChem) for 10 min and stored in dPBS for subsequent staining. Osteogenesis MSCs at passage 2 were seeded at 10,000 cells/cm² in 12-well plates and differentiated using the MesenCult Osteogenic Stimulatory Kit (05504, StemCell) upon reaching 90% confluence. Cells were cultured under hypoxia (1% O₂, 5% CO₂) with medium changes every 3-4 days for 21 days (SVF-derived MSCs) or 7 days (BM-derived MSCs), as per manufacturer’s guidelines. Post-differentiation, cells were fixed in Histofix for 10 min and stored in dPBS for subsequent staining. Histological analysis To confirm adipogenesis, cells were stained with Oil Red O (O0625, Sigma). A stock solution (300 mg Oil Red O in 99 mL 99% isopropanol) was diluted 3:2 with deionized water to prepare the working solution, filtered (0.22 μm), and used within 2 h. Fixed cells were washed with dPBS, treated with 60% isopropanol (5 min) (59310, Sigma) then stained with Oil Red O (5 min). After rinsing, nuclei were counterstained with hematoxylin (1 min) and imaged using AxioObserver 7 microscope (Zeiss). To confirm osteogenesis, a 40 mM Alizarin Red S solution (A5533, Sigma) was prepared by dissolving 6.846 g in 400 mL deionized water, adjusting pH to 4.2 with NaOH (131687.1211, Panreac), and bringing to 500 mL final volume before 0.22 μm filtration. Fixed samples were washed with dPBS, air-dried, then stained with 1 mL/well Alizarin Red solution (15 min, RT). After gentle aspiration, samples were washed 3x with tap water (1 mL/well) while preserving mineralized deposits. Imaging was performed using an AxioObserver 7 microscope (Zeiss). Differentiation of isolated MSCs toward LEPCs At passage 3, MSCs were transferred to new dishes using 0.25% Trypsin-EDTA, and seeded at 20.000 cells/cm 2 with complete mouse endothelial cell medium (ECGM, M1168, Cell Biologics) supplemented with 50 ng/mL of VEGF-C (752-VC, RD Systems), CCBE1 (H00147372-P01, Abnova), angiopoietin-2 (7186-AN, Angiopoietin-2), IGF2 (792-MG, RD Systems), FGF2 (3339-FB, RD Systems) and angiopoietin-1 (9936-AN, RD Systems). The differentiation protocol was split into 3 stages: induction, differentiation and maturation. Induction was carried for 5 days and cells were cultured with ECGM supplemented with VEGF-C. Differentiation followed the previous and lasted 7 days cultured in ECGM supplemented with VEGF-C, CCBE1 and angiopoietin-2. Maturation comprised an additional 7 days of culture with ECGM supplemented with VEGF-C, IGF2, FGF2 and angiopoietin-1. Cells were maintained under standard culture conditions (37 °C with 5% CO 2 ). Immunofluorescence staining Cultured LEPCs and MSCs were fixed with Hystofix for 15 minutes diluted 1:1 in D-PBS, blocked and permeabilized with 10% FBS and 0.3% Triton X-100 (T8787, Sigma) in D-PBS and immunostained. Briefly, cells were immunostained with: anti-mouse PDPN (127402, BioLegend), anti-mouse LYVE1 (103-PA50AG, Reliatech) and anti-mouse VEGFR3 (AF743, RD Systems) as the primary antibodies; and Alexa Fluor 647-conjugated anti-hamster (A21451, Invitrogen), Alexa Fluor 555-conjugated anti-rabbit (A31572, Molecular Probes) and Alexa Fluor 488-conjugated anti-goat (A11055, Molecular Probes) as secondary antibodies. Alexa Fluor 488-conjugated phalloidin (A12379, Invitrogen) was used to selectively stain f-actin. 4′, 6-Diamidino-2-Phenylindole (DAPI, D3571, Invitrogen) was used to stain nuclei (please refer to table S1 for antibody details). Immunofluorescence images were taken using Zeiss AxioObserver 7 and LSM900 confocal microscope (Zeiss). View this table: View inline View popup Download powerpoint Table S1. Antibodies and other stainings used for immunofluorescence staining. Flow cytometry analysis The expression of PDPN, SCA-1, CD90.2, CD105, CD44, CD31 (PECAM1) and CD34 of the isolated and differentiated cells was detected using the APC-conjugated anti-mouse PDPN (127410, BioLegend), PE-Cy7-conjugated anti-mouse SCA-1 (558162, BioLegend), anti-mouse CD90.2 (553009, BD Biosciences), anti-mouse CD105 (14-1051-82, ThermoFisher), anti-mouse CD44 (553134, BD Biosciences), PE-conjugated anti-mouse CD31 (553373, BD Biosciences) and PE-conjugated anti-mouse CD34 (551387, BD Biosciences). Immunoglobulin isotype controls were used to distinguish specific from non-specific bindings during flow cytometry (402012, Biolegend; 553930, BD Biosciences). Alexa Fluor 488-conjugated secondary antibody was used for the unconjugated CD90.2 and CD105 primary antibodies (A11006, Molecular Probes)(please refer to table S2 for antibody details). The expression levels were determined by the median fluorescence intensity (MFI), using FlowJo v10 software (BD Biosciences). View this table: View inline View popup Download powerpoint Table S2. Antibodies used for flow cytometry analysis. RNA extraction and RT-qPCR Total RNA was isolated using tri reagent solution (AM9738, Life Technologies). Reverse transcription was performed using the High-Capacity cDNA reverse transcription kit (4368814, AppliedBiosystems) according to manufacturer’s guidelines. Quantitative real-time PCR was performed in a CFX384 Thermal Cycler (Bio-Rad) using TaqMan Gene Expression Master Mix (4369016, ThermoFisher) according to the manufacturer’s guidelines and 10 ng of cDNA. Gapdh and Tbp were used as housekeeping genes (please refer to table S3 for probe details). Statistical analysis was performed based on the difference between the Ct value of the target gene and the Ct value of the housekeeping gene (2 -ΔCt values). Gene expression was represented using 2 -ΔΔCt values. View this table: View inline View popup Download powerpoint Table S3. Taqman probes use for RT-qPCR analysis. Genotype stability over passages To assess the stability of the gene expression over the passages, cells were plated at 10.000 cells/cm 2 on 12-well plates and cultured at 37 °C until 80% confluence was reached. Then, cells were detached (using Trypsin-EDTA 0.25%), counted and a fraction of them were seeded (10.000 cells/cm 2 ) while the rest were lysed for RNA extraction at each passage until passage 6. Scratch-wound assay Cells (10.000 cells/cm 2 ) were plated on 24-well tissue culture plates and allowed to reach confluence (3-4 days). Once confluent, the media was aspirated and the cell layers wounded using a 20 µL micropipette tip. Then, the plates were rinsed with D-PBS to remove detached cells; fed with cell medium, and monitored until the wound had fully closed. Photographs were taken every 24 hours and images were analyzed using the Wound Healing tool plugin for ImageJ. Tube formation assay in vitro 10 µL of Matrigel (3 mg/ml, 354234, Corning) were added to each well (µ-slide 15 well, Ibidi) and left to harden for approximately 30 minutes. Once the coating had cross-linked, 10.000 cells in 50 µL were seeded directly onto the Matrigel coated wells, incubated at 37 °C for 5 hours, stained for LIVE/DEAD and photographed using Zeiss AxioObserver 7 (Zeiss). Three-dimensional (3D) culture in vitro 1×10 5 cells were embedded in 100µL of Matrigel (3 mg/ml, 354234, Corning), seeded in an 8-well culture slide (BD Falcon) and let to crosslink for approximately 30 minutes. Once the gel had cross-linked, culture media was added and the constructs were incubated at 37 °C for 10 days. After 10 days, bright field images were taken to assess the web formation (AxioObserver 7, Zeiss), cell viability by LIVE/DEAD staining (L3224, Invitrogen) and fixed for immunofluorescence staining using Hystofix as described before. RNA Sequencing Total RNA was extracted using TRI reagent solution. RNA libraries were sequenced on an Illumina NextSeq2000, generating 20-30 million paired-end reads (100 bp, 50 bp x 2). RNA-Seq data processing and differential expression analysis were performed using the standardized nf-core/rnaseq (v3.15.0) and nf-core/differentialabundance (v1.5.0) pipelines to ensure reproducibility and best practices in computational workflows. 39 Raw sequencing data were processed through nf-core/rnaseq. Low-quality bases filtering (Phred score < 15) using a sliding window approach and adapter trimming were performed using fastp v.0.23.4. 40 Ribosomal RNA (rRNA) was removed using SortMeRNA v.4.3.6. 41 Cleaned reads were aligned to the Mus musculus GRCm39 reference genome (ENSEMBL) using STAR aligner v.2.7.10a. 42 Transcript abundance was quantified at the gene level with Salmon v.1.10.1. 43 Count matrices generated by nf-core/rnaseq were analyzed for differential expression using nf-core/differentialabundance, which leverages DESeq2 (v1.44.0). 44 The design matrix explicitly accounted for technical variability from sample replication batches while estimating absolute expression levels across conditions. p -values were adjusted with the Benjamini-Hochberg (BH) method to reduce the number of false positives. Differentially expressed genes were called with thresholds of |log2(fold change)| ≥ 1 and adjusted p -value (FDR) < 0.05. Results MSC isolation is consistently more efficient for SVF than BM or dermis SVF and BM display better results when it comes to cell count after the extraction protocol (Sup. 1). Whole BM cell suspensions contained approx. 250×10 6 cells (>90% viability), while SVF approx. 2×10 6 cells (>90% viability), and the dermis around 5×10 5 with really poor viability (∼50%). Regarding these results dermis was discarded and our focus relied into BM and SVF. When cultured in complete Mesencult Medium, SVF-derived MSCs showed higher proliferation rate in comparison to BM-derived MSCs, showing twice the accumulated cell proliferation per mouse for almost half the time of culture (Sup. 1B). Moreover, cell culture showed less morphological heterogeneity in SVF-derived MSCs than BM-derived MSCs (Sup. 1A.II and C.II). Finally, MSC lineage was confirmed for both tissue sources by (a) analysis of MSC surface markers through flow cytometry and (b) their differentiation potential for adipogenesis and osteogenesis (Sup. 1A.III-V and C.III-V). Both SVF- and BM-derived cells showed typical MSC marker expression patterns: SVF-derived MSCs expressed high levels of CD90.2 (90.48% ± 4.54), CD105 (51.13% ± 4.87), and SCA1 (92.48% ± 2.10), along with moderate CD44 expression (28.5% ± 25.17); BM-MSCs exhibited lower CD90.2 (9.06% ± 0.23) and CD44 (12.42% ± 11.57) but maintained strong CD105 (61.05% ± 0.78) and SCA1 (82.45% ± 1.77) expression. Importantly, both populations showed minimal endothelial marker presence (SVF: CD31 0.08% ± 0.02, CD34 0.09% ± 0.05; BM: CD31 0.68% ± 0.39, CD34 0.8% ± 0.3). Both cell populations successfully underwent adipogenic and osteogenic differentiation, as evidenced by distinct histological staining patterns. Adipogenic differentiation was confirmed through the accumulation of lipid droplets that stained positively with Oil Red O, while osteogenic differentiation was verified by Alizarin Red S staining of calcium phosphate deposits and the morphological presence of mineralized nodules. Flow cytometry analysis shows that SVF-derived MSCs outperform BM-derived MSCs in lymphangiogenic differentiation potential Flow cytometry analysis of SVF- and BM-derived LEPCs showed a significant increase in the expression of PDPN for both cell types ( fig. 2A -B); nonetheless, SVF-derived LEPCs showed a higher increase than the BM (97.4% ± 2.2 of PDPN-positive LEPCs for SVF vs. 76.2% ± 19.5 for BM) ( fig. 2A .I and B.I). In addition, MFI analysis confirm a statistically significant change in SVF-derived LEPCs (5.5×10 5 MFI for LEPC vs. 1×10 5 MFI for MSC; p -value<0.01) ( fig. 2A .II). MFI results for BM-derived LEPCs also showed a statistically significant increase in PDPN expression (2.5×10 4 MFI for LEPC vs. 1×10 4 MFI for MSC; p -value<0.05) ( fig. 2B .II). Notably, SVF-derived LEPCs exhibited significantly greater upregulation of PDPN expression compared to their BM-derived counterparts, as demonstrated by both higher absolute expression levels and enhanced fluorescence intensity quantification. Given these pronounced differences, we prioritized comprehensive characterization of the SVF-derived LEPCs. Download figure Open in new tab Fig 1. Differentiation protocol illustration. VEGF-C : Vascular endothelial growth factor C. Angpt 2 : Angiopoietin 2. CCBE1 : Collagen and calcium binding EGF domains 1. FGF2 : Fibroblast growth factor 2. IGF2 : Insulin-like growth factor 2. Angpt 1 : Angiopoietin 1. Download figure Open in new tab Fig 2. Stromal vascular fraction (SVF)-derived lymphatic endothelial progenitor cells (LEPCs) show better differentiation by flow cytometry analysis than bone marrow (BM)-derived LEPCs. (A) Flow cytometry workflow of the SVF-derived LEPCs from the first population until the PDPN-positive population. (A.I) PDPN expression histograms showing the expression level for SVF-LEPCs and -MSCs. (A.II) Median fluorescence intensity (MFI) of PDPN in SVF-LEPCs and –MSCs (mean ± SD). (B) Flow cytometry workflow of the BM-derived LEPCs from the first population until the PDPN-positive population (B.I) PDPN expression histograms showing the expression level for BM-LEPCs and -MSCs. (B.II) MFI of PDPN in BM-LEPCs and –MSCs (mean ± SD). Statistical significance: * p -value<0.5, ** p -value<0.01. Mann-Whitney test. n=6 for SVF-LEPCs and n=5 for BM-LEPCs. Mean ± SD represented. LEPC lineage is confirmed by gene expression analysis and immunofluorescence staining of SVF-derived LEPC Our novel differentiation protocol mimics the natural process of lymphatic embryogenesis and incorporates key molecular factors involved in adult lymphangiogenesis. By using VEGF-C, CCBE1, Ang1, Ang2, IGF2 and FGF2, the protocol demonstrated strong potential due to the complementary and synergistic roles these factors play in lymphatic vessel development and function 45 . RT-qPCR analysis confirmed the lymphatic commitment of SVF-derived LEPCs ( Fig. 3A ), with significant fold-changes observed in all critical lymphatic markers ( Pdpn, Lyve1, Vegfr3, Igf2, Vegf-c ), establishing their successful differentiation from MSC precursors ( p -value<0.05). Immunofluorescence analysis of PDPN revealed a dramatic upregulation in SVF-derived LEPCs compared to their MSC progenitors, corroborating earlier findings ( Fig. 3B ). Quantitative assessment by ImageJ demonstrated a significant increase in MFI for SVF-derived LEPCs when normalized to nuclear counts within each sample ( p -value<0.01). Download figure Open in new tab Fig 3. Characterization of lymphatic endothelial progenitor cells (LEPCs) shows correct differentiation by overexpression of lymphatic lineage specific markers. (A) RT-qPCR shows statistically significant overexpression of key lymphatic markers such as Pdpn , Vegfr3 , Lyve1 , Vegf-c and Igf2 . n=4. MSC, mesenchymal stem cells. Mean ± SD. (B) Median fluorescence intensity analysis of immunofluorescence staining images shows statistically significant differential expression of PDPN between MSCs and LEPCs. Scale bar: 100 µm. n=6. Statistical significance: * p -value<0.05; ** p -value<0.01. Mann-Whitney test. SVF-derived LEPCs show a reliable lymphatic phenotype in vitro Our novel differentiation protocol was further characterized in vitro for genotype stability with 3 more passages after differentiation and analyzing the migratory phenotype via scratch-wound assay. The SVF-derived LEPCs showed little variation of gene expression of the key molecular markers of the lymphatic system for at least 1 passage after differentiation (P4) ( fig. 4A ). Moreover, SVF-derived LEPCs showed a very pro-migratory phenotype, closing the generated wound in less than 48 hours of culture ( fig. 4B ). Comparative analysis of cellular migration patterns revealed that our differentiated cells exhibited a migratory phenotype functionally equivalent to HDLECs. These control HDLECs served as our biological reference standard, with quantitative assessments demonstrating comparable migration rates between the two cell populations. Download figure Open in new tab Fig 4. Stromal vascular fraction (SVF)-derived lymphatic endothelial progenitor cells (LEPCs) show a reliable lymphatic phenotype in vitro. (A) Genotype stability results of the key overexpressed genes over passages (P), assessed by RT-qPCR. n=3. (B) Scratch-wound assay of the SVF-derived LEPCs in comparison to human dermal lymphatic endothelial cells (HDLECs) along two days in culture. n=3. Scale bar: 200 µm. Mean ± SD. Mann-Whitney test. SVF-derived LEPCs outperform HDLECs in a biomimetic environment in vitro In order to assess key functional characteristics of LEPCs, we assessed angiogenesis in vitro in 2D and 3D environments using Matrigel as the hydrogel matrix, and was compared with HDLECs as gold standards (control). We selected these cells as controls for several key reasons. First, they exhibit a well-defined lymphatic expression profile, with strong expression of established lymphatic markers. 46 Second, they are readily accessible, being commercially available and widely referenced in existing studies (over 200 studies). Third, they are functionally validated, demonstrating characteristics consistent with LECs. 47 Finally, we opted for human-derived cells over murine models to enhance clinical relevance, ensuring closer alignment with human physiology for translational applications. 48 In a 2D manner, our SVF-derived LEPCs showed a similar capacity for capillary-like structure formation as the control, showing no statistically significant differences in the number of nodes, number of segments, number of loops and number of meshes ( fig. 5A -B). Conversely, when embedded in a 3D Matrigel hydrogel, the SVF-derived LEPCs outperformed the control, showing significantly better tube-like formation ( fig. 5C -E). Specifically, when cultured up to 10 days, while HDLECs showed a little tube-like formation or reorganization, remaining static, SVF-derived LEPCs showed a readily assembled web-like structure that matures into thick structures ( fig. 5C -E). The average number of nodes, junctions, total number of meshes and master junctions for SVF-derived LEPCs greatly outperformed the dermal controls ( fig. 5E ). Finally, immunofluorescence staining of 3D cultured SVF-derived LEPCs expressed canonical lymphatic markers (PDPN, LYVE1, VEGFR3 and F-actin) at 10 days of culture ( fig. 6 ). Download figure Open in new tab Fig. 5. Stromal vascular fraction (SVF)-derived lymphatic endothelial progenitor cells (LEPCs) show better in vitro angiogenesis phenotype than human dermal lymphatic endothelial cells (HDLEC). (A) LIVE/DEAD staining images of SVF-derived LEPCs and HDLECs culture on a Matrigel (3 mg/mL) coating for 5 hours (2D angiogenesis assay). (B) Angiogenesis analysis of SVF-derived LEPCs compared to HDLECs in a 2D environment. N=5. (C) Bright field images of SVF-derived LEPCs and HDLECs embedded in a 3D Matrigel hydrogel (3 mg/mL) and cultured for 10 days. (D) LIVE/DEAD staining images of SVF-derived LEPCs and HDLECs embedded in a 3D Matrigel hydrogel (3 mg/mL) and cultured for 10 days. (E) Angiogenesis analysis of SVF-derived LEPCs compared to HDLECs in a 3D environment. n=4. Scale bars: (A) 500 µm; (C-D) 200 µm. Statistical significance: * p -value<0.05. Mann-Whitney test. Download figure Open in new tab Fig. 6. Stromal vascular fraction (SVF)-derived lymphatic endothelial progenitor cells (LEPCs) express canonical lymphatic markers in a 3D environment. Representative immunofluorescence staining images of LEPCs expressing lymphatic markers (PDPN, LYVE1, VEGFR3 and F-actin) embedded in Matrigel for 10 days. To the right, magnification of the white square in the left. Scale bar: 100 µm. RNA Sequencing confirms LEC lineage of SVF-derived LEPCs For RNA sequencing, we utilized matched RNA samples isolated from SVF-derived LEPCs and their progenitor cells (passage 3). Three independent biological replicates were processed, with two technical replicates per sample. All cells underwent lymphatic differentiation prior to RNA extraction using TRI reagent solution, ensuring consistent sample origin across both genomic and expression analyses. Principal Component Analysis (PCA) of variance-stabilized transformed counts was performed to assess the overall variability and clustering of samples based on gene expression profiles. The first two principal components, PC1 and PC2, explained 69.2% and 9.3% of the total variance, respectively ( fig. 7A ). The PCA showed distinct clustering of the samples based on their experimental condition, also showing significant clustering in each experimental condition by their duplicates ( fig. 7A ). Consistent with the PCA, the sample clustering dendogram of the top 500 genes confirmed clear separation between biological groups, while duplicates exhibited high concordance ( fig. 7B ). Differential expression analysis showed a total of 2.037 upregulated and 1.936 downregulated genes for the differentiated cells in comparison to the MSCs (|log2 fold change| ≥1, adjusted p -value ≤0.05) ( fig. 7C ). Among the upregulated genes, several canonical markers of LECs were identified, including Pdpn , Lyve1 , and Vegfr3 , among others ( fig. 7D ). Focusing on the top 50 LEC specific genes, the differentiated cells showed a significant upregulation on 7 genes ( Pdpn, Vegfr3, Lyve1, Bmp2, Notch1, Cd34, Flt1 ) as compared to their controls ( fig. 7E ). Additional details regarding the expression levels of these genes are shown in boxplots (comparing LEPCs and MSCs) in Sup. 2. Download figure Open in new tab Fig 7. RNA sequencing analysis of Stromal vascular fraction (SVF)-derived lymphatic endothelial progenitor cells (LEPCs) confirm lymphatic genotype in comparison to mesenchymal progenitor cells. (A) Variance-stabilized transformed (VST) counts were used for Principal Component Analysis (PCA) plot. Axes show the top two principal components (PC1: 69.2%, PC2: 9.3%). Samples are colored by condition. (B) Sample clustering of the top 500 most variable genes. Distances between genes were estimated based on spearman correlation, which were then used to produce a clustering via the ward.D2 method. (C) Volcano plot of the differentially expressed genes (DEGs) between LEPCs and MSCs. Gray: non-significant; red: significant DEGs (|log2FC|≥1, adjusted p -value<0.05). (D) Boxplot of the differential expression of canonic lymphatic markers. Significance: ***adjusted p -value<0.001 (DESeq2 Wald test). (E) Heatmap of top 50 lymphatic endothelial cell (LEC) markers. Z-score-scaled VST counts for the 50 most significant LEC markers (rows) across samples (columns). Bold genes: |log2 fold change|≥1, adjusted p -value≤0.05. Clustering uses Euclidean distance and Ward.D2 linkage. These findings demonstrate the potential to rapidly and precisely generate reliable lymphatic progenitors in vitro, which could outperform existing commercial cell lines for lymphatic-related applications. Discussion Lymphatic tissue engineering has been significantly underexplored, resulting in a substantial knowledge gap that hampers the development of engineered constructs for purposes such as disease modelling, implants, or cell therapies. Tissue engineering typically relies on three essential components: cells, biomaterials, and biomolecules. However, in the context of lymphatic vessels/system, research has predominantly focused on biomolecules, as in vivo studies have proven valuable in understanding the aetiologies of various lymphatic system diseases. 2, 49 Among these, VEGF-C has received the most attention due to its critical role in lymphatic embryogenesis, postnatal stability, and lymphangiogenesis. 12 , 50 , 51 Despite progress in understanding biochemical cues, much less attention has been given to the cellular aspects of lymphatic tissue engineering. Most studies involving cells have primarily examined their potential to regenerate dysfunctional or absent lymphatic vessels in vivo. 21 These studies often use cells from a wide range of tissues, with a focus on MSCs or differentiated cells. 21 For MSCs, the emphasis has typically been on their paracrine effects, which enhance lymphatic vessel formation, rather than on their direct differentiation and application. 25 , 52 While in vitro studies have predominantly used commercial cell lines 53 , reliable protocols for differentiating MSC into LEPCs remain scarce 21 . Recognizing this significant gap, we have successfully established a novel in vitro differentiation protocol mimicking lymphatic embryogenesis, focusing on three clinically relevant tissue sources, obtainable from patients for autologous use. Among these, dermal tissue yielded the lowest cell count, whereas SVF and BM performed better. Even though, BM provides highest initial cell quantities, its poor attachment and proliferation in culture limits its utility. Conversely, SVF-derived MSCs, despite starting with fewer cells, proliferated quickly, achieving higher cell counts in half of the time, making them ideal for time-limited treatments 54 . Both SVF- and BM-derived MSCs promote lymphangiogenesis through paracrine signalling 9, 32 , 55 . However, adipose tissue is very highly related to the lymphatic pathologies, with a quite clear correlation of fat adipose tissue sedimentation and lymphatic malfunction 2 . Clinically, BM harvesting is highly invasive, posing greater risks to patients, while SVF extraction avoids this entirely 56 – 58 . In addition, SVF is easier to handle, requiring only standard culture conditions, unlike BM, which requires hypoxic conditions. Hence, SVF constitutes a superior option due its rapid expansion, minimal invasiveness and simplified culturing process. Our novel differentiation protocol mimics the natural process of lymphatic embryogenesis and incorporates key molecular factors involved in adult lymphangiogenesis. VEGF-C is an early chemokine of the lymphatic embryogenesis (mice E9.5) that initiates the lymphatic commitment of venous endothelial cells, and driving the budding and sprouting of lymph sacs from the cardinal vein via VEGFR-3 signalling. 12 , 51 , 59 CCBE1 acts synergistically to enhance VEGF-C activity through proteolytic maturation by ADAMTS3. 60 Ang2 and 1 contribute to vascular remodelling by promoting vessel enlargement, branching and stabilizing the nascent vessels through TIE2 receptor. 50 , 61 , 62 IGF2 plays a supplementary role during by activating VEGFR3-independet pathways, such as Akt and ERK, ensuring lymphangiogenesis continues even in the absence of VEGF-C. 63 , 64 Finally, FGF2 promotes LEC tip cell formation, enabling vessel sprouting and elongation while stabilizing LECs via LYVE1 interaction. 65 – 67 Together, these factors orchestrate the progression from initial LEC specification to the stabilization and maturation of a functional lymphatic network. Our results following this differentiation protocol align perfectly with the above-mentioned in vivo embryogenesis stages, as our denominated differentiation stages (induction, differentiation and maturation) accurately resemble embryogenesis in vitro . Our RT-qPCR results show that in comparison with the MSCs, the SVF-derived LEPCs show significant upregulation of the key lymphatic factors by statistically significant changes for Pdpn , Lyve1 , Vegfr3 , Vegf-C and Igf2 . Based on the previous mentioned factors, the key LEC markers have always been PDPN, LYVE1 and VEGFR3. Overexpression of PDPN, LYVE1 and VEGFR3 is a hallmark of LEC identity and is pivotal for validating successful in vitro differentiation protocols. 68 , 69 Furthermore, the overexpression of other markers, such as VEGF-C and IGF2, indicates that the cells not only express the key molecular biomarkers associated with LECs but also demonstrate the expression of functional markers. The overexpression of VEGF-C sustains the LEC phenotype by VEGFR3 signalling. 12 , 70 , 71 Similarly, IGF2 can maintain lymphangiogenesis in a VEGF-C independent manner, suggesting that LEPCs have the capacity to self-sustain their lymphatic genotype even in the absence of VEGF-C. 64 Following the successful differentiation of SVF-derived LEPCs, we performed extensive characterization to assess their behaviour and confirm their LEC phenotype in vitro . Our scratch-wound assay demonstrated a highly migratory phenotype comparable to HDLECs, as the SVF-derived LEPCs effectively closed the wound within 48 hours, displaying not only molecular marker expression but also robust functional capabilities. Additionally, maintaining the desired phenotype and gene expression over passages post-differentiation is crucial for reducing costs and accelerating practical applications. 72 We observed that the expression of key markers, including Pdpn , Lyve1 , Vegfr-3 , and Igf-2 , remained stable across the first two passages post-differentiation, indicating the robustness of our protocol. This stability could be attributed to the secretion of VEGF-C and IGF-2 by the cells, potentially forming an autocrine positive feedback loop. 12 , 64 Furthermore, we assessed the ability of the cells to form angiogenic structures in both 2D and 3D environments. In 2D cultures, SVF-derived LEPCs readily formed web-like structures within 5 hours of Matrigel culture, mirroring angiogenesis assays and showing no significant difference compared to HDLECs. In 3D environments, our differentiated cells outperformed HDLECs, forming intricate tubular structures that simulated a lymphatic capillary network and expressing key lymphatic markers such as PDPN, LYVE1 and VEGFR3. These findings underscore the strength of our differentiation protocol, demonstrating the ability to derive functional LEPCs from SVF-derived MSCs in vitro , a breakthrough that holds promise for advancing disease modelling, tissue engineering, and cell therapy. For instance, disease modelling can drive therapeutic development for lymphedema and other traditional lymphatic diseases, as well as emerging conditions like obesity, where lymphatic dysfunction contributes to fat accumulation and inflammation, 73 – 75 and Alzheimer’s disease, where the glymphatic system may play a pivotal role in addressing amyloid and tau pathology. 2 Finally, the resulting transcriptomic analysis from the differentiation protocol provides very compelling evidence of successful differentiation, as demonstrated by the gene expression profiles, PCA clustering patterns, and differential expression analysis. Indeed, PCA results indicate that the first two principal components, PC1 (62.9%) and PC2 (9.3%), account for a large amount of the total variance. Such high levels of variance for PC1 suggests that the largest source of variability are the experimental conditions, i.e. the differentiation from MSCs into LEPCs. Furthermore, the significant clustering of the samples based on their experimental condition highlights the robustness and reproducibility of the differentiation protocol, which further supports the conclusion that the differentiation process induces significant changes in gene expression patterns that translate into phenotypical changes. The differential expression analysis revealed a very significant number of upregulated and downregulated genes in the differentiated cells compared to their mesenchymal progenitors, suggesting correct differentiation. When analysing the canonical genes for LECs, all genes confirmed our RT-qPCR results, indicating that there is a substantial upregulation of Pdpn , Lyve1 and Vegfr3 . The upregulation of these markers strongly suggests that the differentiated cells have acquired a LEPC phenotype. In addition, confirmation of the upregulation of most of the LEC specific markers, further confirms the correct differentiation of the mesenchymal stem cells into LEPCs. Collectively, our findings present a novel and previously unreported differentiation protocol for generating LEPCs in vitro, addressing a critical gap in lymphatic research. Additionally, we identify the optimal tissue source for LEPC isolation—a clinically accessible and highly relevant tissue with direct translational potential. Conclusion This study explored the potential to differentiate MSCs derived from three distinct tissues into LEPCs for applications in lymphatic tissue engineering and cellular therapy. The findings demonstrated that SVF and BM serve as more reliable sources for isolating MSCs than the dermis. Furthermore, SVF-derived MSCs exhibited superior proliferation potential compared to BM-derived MSCs. A novel differentiation protocol was developed, mimicking the molecular signals of lymphatic embryogenesis, successfully guiding MSCs into LEPCs. Our results indicated that SVF-derived MSCs achieved more efficient differentiation into LEPCs than BM-derived MSCs, as evidenced by flow cytometry analysis of PDPN expression. Additional characterization of SVF-derived LEPCs confirmed their LEC phenotype, displaying high migratory capacity, a strong propensity for angiogenic structure formation, and consistent expression of key lymphatic markers in both 2D and 3D environments. Future studies will focus on evaluating these cells in in vivo models to assess their lymphangiogenic capabilities as a preliminary step toward their application in cell therapy. Overall, this research highlights a novel differentiation protocol that replicates the embryonic development of the lymphatic system, enabling the transformation of MSCs into LEPCs. These cells hold significant promise for tissue engineering, disease modelling, and therapeutic applications. Download figure Open in new tab Supplementary 1. Mesenchymal stem cells (MSCs) isolation protocol and lineage confirmation. (A) Isolation and confirmation of stromal vascular fraction (SVF)-derived MSCs; (A.I) Illustration of the isolation protocol followed for obtaining SVF-derived MSCs; (A.II) Morphology of the SVF-derived MSCs in culture; (A.III) Expression of the canonical MSC markers by flow cytometry; (A.IV) Adipogenic differentiation of SVF-MSCs; (A.V) Osteogenic differentiation of SVF-MSCs. (B) Accumulated cell proliferation of SVF- and BM-derived MSCs over passages relative to mice. (C) Isolation and confirmation of BM-derived MSCs; (C.I) Illustration of the isolation protocol followed for obtaining BM-derived MSCs; (C.II) Morphology of the BM-derived MSCs in culture; (C.III) Expression of the canonical MSC markers by flow cytometry; (C.IV) Adipogenic differentiation of BM-MSCs; (C.V) Osteogenic differentiation of BM-MSCs. Download figure Open in new tab Supplementary 2. Boxplots of the top 50 lymphatic endothelial cell (LEC) markers expressed in lymphatic endothelial progenitor cells (LEPCs) in comparison to their mesenchymal progenitors. Funder Information Declared Instituto de Salud Carlos III, https://ror.org/00ca2c886 , PI22/01247 , CERT22/00032 , RD24/0014/0012 , DTS24/00167 , PT23/00142 Basque Government, https://ror.org/00pz2fp31 , KK-2024/00041 , 2020111004 Footnotes ↵ † POLYMAT, University of the Basque Country UPV/EHU, Avenida Tolosa 72, Donostia/San Sebastian 20018, Gipuzkoa, Spain. References 1. ↵ Petrova , T. V. & Koh , G. Y . Biological functions of lymphatic vessels . Science 369 , eaax4063 ( 2020 ). OpenUrl Abstract / FREE Full Text 2. ↵ Oliver , G. , Kipnis , J. , Randolph , G. J. & Harvey , N. L . The Lymphatic Vasculature in the 21st Century: Novel Functional Roles in Homeostasis and Disease . Cell 182 , 270 – 296 ( 2020 ). OpenUrl CrossRef PubMed 3. ↵ Jiang , X. , Nicolls , M. R. , Tian , W. & Rockson , S. G. Lymphatic Dysfunction, Leukotrienes, and Lymphedema . Annu. Rev. Physiol . 80 , 49 – 70 ( 2018 ). OpenUrl CrossRef PubMed 4. ↵ Hong , Y.-K. , Shin , J. W. & Detmar , M . Development of the lymphatic vascular system: A mystery unravels . Dev. Dyn . 231 , 462 – 473 ( 2004 ). OpenUrl CrossRef PubMed Web of Science 5. ↵ Rauniyar , K. , Jha , S. K. & Jeltsch , M . Biology of Vascular Endothelial Growth Factor C in the Morphogenesis of Lymphatic Vessels . Front. Bioeng. Biotechnol . 6 , ( 2018 ). 6. ↵ Simeroth , S. & Yu , P . The role of lymphatic endothelial cell metabolism in lymphangiogenesis and disease . Front. Cardiovasc. Med . 11 , ( 2024 ). 7. Lucas , E. D. & Tamburini , B. A. J . Lymph Node Lymphatic Endothelial Cell Expansion and Contraction and the Programming of the Immune Response . Front. Immunol . 10 , ( 2019 ). 8. ↵ Rouhani , S. J. , Eccles , J. D. , Tewalt , E. F. & Engelhard , V. H . Regulation of T-cell Tolerance by Lymphatic Endothelial Cells . J. Clin. Cell. Immunol . 5 , 1000242 ( 2014 ). 9. ↵ Dai , T. et al. The Roles of Podoplanin-Positive/Podoplanin-Negative Cells from Adipose-Derived Stem Cells in Lymphatic Regeneration . Plast. Reconstr. Surg . 145 , 420 – 431 ( 2020 ). OpenUrl PubMed 10. ↵ Bieniasz-Krzywiec , P. et al. Podoplanin-Expressing Macrophages Promote Lymphangiogenesis and Lymphoinvasion in Breast Cancer . Cell Metab . 30 , 917 – 936 .e10 ( 2019 ). OpenUrl CrossRef PubMed 11. ↵ Wu , M. et al. Low Molecular Weight Hyaluronan Induces Lymphangiogenesis through LYVE-1-Mediated Signaling Pathways . PLoS ONE 9 , e92857 ( 2014 ). OpenUrl CrossRef PubMed 12. ↵ Lin , Q. , Zhang , Y. , Bai , J. , Liu , J. & Li , H . VEGF-C/VEGFR-3 axis protects against pressure-overload induced cardiac dysfunction through regulation of lymphangiogenesis . Clin. Transl. Med . 11 , e374 ( 2021 ). OpenUrl 13. ↵ Urner , S. et al. Identification of ILK as a critical regulator of VEGFR 3 signalling and lymphatic vascular growth . EMBO J . 38 , e99322 ( 2019 ). OpenUrl Abstract / FREE Full Text 14. ↵ Rezzola , S. , Sigmund , E. C. , Halin , C. & Ronca , R . The lymphatic vasculature: An active and dynamic player in cancer progression . Med. Res. Rev . 42 , 576 – 614 ( 2022 ). OpenUrl CrossRef PubMed 15. Yamashita , M. et al. Development of Lymphatic Capillary Network Along the Alveolar Walls of Autopsied Human Lungs with Pneumonia . Lymphat. Res. Biol . 14 , 210 – 219 ( 2016 ). OpenUrl PubMed 16. ↵ Pei , G. et al. Lymphangiogenesis in kidney and lymph node mediates renal inflammation and fibrosis . Sci. Adv . 5 , eaaw5075 ( 2019 ). OpenUrl FREE Full Text 17. ↵ Goodell , M. A. & Rando , T. A . Stem cells and healthy aging . Science 350 , 1199 – 1204 ( 2015 ). OpenUrl Abstract / FREE Full Text 18. ↵ Brown , C. et al. Mesenchymal stem cells: Cell therapy and regeneration potential . J. Tissue Eng. Regen. Med . 13 , 1738 – 1755 ( 2019 ). OpenUrl CrossRef PubMed 19. Fan , X.-L. , Zhang , Y. , Li , X. & Fu , Q.-L . Mechanisms underlying the protective effects of mesenchymal stem cell-based therapy . Cell. Mol. Life Sci . 77 , 2771 – 2794 ( 2020 ). OpenUrl CrossRef PubMed 20. ↵ Chang , C. et al. Effects of Mesenchymal Stem Cell-Derived Paracrine Signals and Their Delivery Strategies . Adv. Healthc. Mater . 10 , 2001689 ( 2021 ). 21. ↵ Arriola-Alvarez , I. , Jaunarena , I. , Izeta , A. & Lafuente , H . Progenitor Cell Sources for 3D Bioprinting of Lymphatic Vessels and Potential Clinical Application . Tissue Eng. Part A 30 , 353 – 366 ( 2024 ). OpenUrl PubMed 22. ↵ Religa , P. et al. Presence of bone marrow–derived circulating progenitor endothelial cells in the newly formed lymphatic vessels . Blood 106 , 4184 – 4190 ( 2005 ). OpenUrl Abstract / FREE Full Text 23. Wang , C. et al. M2b macrophages stimulate lymphangiogenesis to reduce myocardial fibrosis after myocardial ischaemia/reperfusion injury . Pharm. Biol . 60 , 384 – 393 ( 2022 ). OpenUrl PubMed 24. Conrad , C. et al. Multipotent Mesenchymal Stem Cells Acquire a Lymphendothelial Phenotype and Enhance Lymphatic Regeneration In Vivo . Circulation 119 , 281 – 289 ( 2009 ). OpenUrl Abstract / FREE Full Text 25. ↵ Maertens , L. et al. Bone Marrow-Derived Mesenchymal Stem Cells Drive Lymphangiogenesis . PLoS ONE 9 , e106976 ( 2014 ). OpenUrl PubMed 26. ↵ Berebichez-Fridman , R. et al. The Holy Grail of Orthopedic Surgery: Mesenchymal Stem Cells—Their Current Uses and Potential Applications . Stem Cells Int . 2017 , 1 – 14 ( 2017 ). OpenUrl CrossRef 27. ↵ Knezevic , L. et al. Engineering Blood and Lymphatic Microvascular Networks in Fibrin Matrices . Front. Bioeng. Biotechnol . 5 , ( 2017 ). 28. ↵ Bourland , J. , Fradette , J. & Auger , F. A . Tissue-engineered 3D melanoma model with blood and lymphatic capillaries for drug development . Sci. Rep . 8 , 13191 ( 2018 ). 29. ↵ Kawai , Y. et al. Cell transplantation therapy for a rat model of secondary lymphedema . J. Surg. Res . 189 , 184 – 191 ( 2014 ). OpenUrl PubMed 30. ↵ Fraser , J. K. , Wulur , I. , Alfonso , Z. & Hedrick , M. H . Fat tissue: an underappreciated source of stem cells for biotechnology . Trends Biotechnol . 24 , 150 – 154 ( 2006 ). OpenUrl CrossRef PubMed Web of Science 31. Strassburg , S. , Torio-Padron , N. , Finkenzeller , G. , Frankenschmidt , A. & Stark , G. B. Adipose-Derived Stem Cells Support Lymphangiogenic Parameters In Vitro . J. Cell. Biochem . 117 , 2620 – 2629 ( 2016 ). OpenUrl PubMed 32. ↵ Marcozzi , C. et al. Paracrine Effect of Human Adipose-Derived Stem Cells on Lymphatic Endothelial Cells . Regen. Med . 15 , 2085 – 2098 ( 2020 ). OpenUrl PubMed 33. ↵ Takeda , K. , Sowa , Y. , Nishino , K. , Itoh , K. & Fushiki , S . Adipose-Derived Stem Cells Promote Proliferation, Migration, and Tube Formation of Lymphatic Endothelial Cells In Vitro by Secreting Lymphangiogenic Factors . Ann. Plast. Surg . 74 , 728 – 736 ( 2015 ). OpenUrl PubMed 34. ↵ Ahmadzadeh , N. et al. Human adipose-derived stem cells support lymphangiogenesis in vitro by secretion of lymphangiogenic factors . Exp. Cell Res . 388 , 111816 ( 2020 ). 35. ↵ Fong , C.-Y. et al. Human Wharton’s Jelly Stem Cells Have Unique Transcriptome Profiles Compared to Human Embryonic Stem Cells and Other Mesenchymal Stem Cells . Stem Cell Rev. Rep . 7 , 1 – 16 ( 2011 ). OpenUrl PubMed 36. ↵ Hsieh , J.-Y. , Fu , Y.-S. , Chang , S.-J. , Tsuang , Y.-H. & Wang , H.-W . Functional Module Analysis Reveals Differential Osteogenic and Stemness Potentials in Human Mesenchymal Stem Cells from Bone Marrow and Wharton’s Jelly of Umbilical Cord . Stem Cells Dev . 19 , 1895 – 1910 ( 2010 ). OpenUrl CrossRef PubMed Web of Science 37. ↵ Nguyen , V. A. et al. Endothelial cells from cord blood CD133 + CD34 + progenitors share phenotypic, functional and gene expression profile similarities with lymphatics . J. Cell. Mol. Med . 13 , 522 – 534 ( 2009 ). OpenUrl CrossRef PubMed Web of Science 38. ↵ Tan , Y. et al. CD34+VEGFR-3+ progenitor cells have a potential to differentiate towards lymphatic endothelial cells . J. Cell. Mol. Med . 18 , 422 – 433 ( 2014 ). OpenUrl CrossRef PubMed 39. ↵ Ewels , P. A. et al. The nf-core framework for community-curated bioinformatics pipelines . Nat. Biotechnol . 38 , 276 – 278 ( 2020 ). OpenUrl CrossRef PubMed 40. ↵ Chen , S. , Zhou , Y. , Chen , Y. & Gu , J. fastp: an ultra-fast all-in-one FASTQ preprocessor . Bioinformatics 34 , i884 – i890 ( 2018 ). OpenUrl CrossRef PubMed 41. ↵ Kopylova , E. , Noé , L. & Touzet , H . SortMeRNA: fast and accurate filtering of ribosomal RNAs in metatranscriptomic data . Bioinformatics 28 , 3211 – 3217 ( 2012 ). OpenUrl CrossRef PubMed Web of Science 42. ↵ Dobin , A. et al. STAR: ultrafast universal RNA-seq aligner . Bioinformatics 29 , 15 – 21 ( 2013 ). OpenUrl CrossRef PubMed Web of Science 43. ↵ Patro , R. , Duggal , G. , Love , M. I. , Irizarry , R. A. & Kingsford , C . Salmon provides fast and bias-aware quantification of transcript expression . Nat. Methods 14 , 417 – 419 ( 2017 ). OpenUrl CrossRef PubMed 44. ↵ Love , M. I. , Huber , W. & Anders , S . Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2 . Genome Biol . 15 , 550 ( 2014 ). 45. ↵ Hu , Z. et al. Lymphatic vessel: Origin, heterogeneity, biological functions and therapeutic targets . Signal Transduct. Target. Ther . 9 , 9 ( 2024 ). 46. ↵ Koukorava , C. , et al. Mesothelial Cells Exhibit Characteristics of Perivascular Cells in an In Vitro Angiogenesis Assay . Cells 12 , 2436 ( 2023 ). OpenUrl 47. ↵ Iesato , A. et al. Axillary adipose tissue–derived lymphatic endothelial cells exhibit distinct transcriptomic signatures reflecting lymphatic invasion status in breast cancer . Breast Cancer Res. BCR 27 , 109 ( 2025 ). 48. ↵ Medetgul-Ernar , K. & Davis , M. M . Standing on the shoulders of Mice . Immunity 55 , 1343 – 1353 ( 2022 ). OpenUrl CrossRef PubMed 49. ↵ Breslin , J. W. et al. Lymphatic Vessel Network Structure and Physiology . in Comprehensive Physiology (ed. Prakash, Y. S.) 207 – 299 (Wiley, 2018 ). doi: 10.1002/cphy.c180015 . OpenUrl CrossRef PubMed 50. ↵ Tammela , T. et al. Angiopoietin-1 promotes lymphatic sprouting and hyperplasia . Blood 105 , 4642 – 4648 ( 2005 ). OpenUrl Abstract / FREE Full Text 51. ↵ Li , W.-N. et al. Extracellular vesicle-associated VEGF-C promotes lymphangiogenesis and immune cells infiltration in endometriosis . Proc. Natl. Acad. Sci . 117 , 25859 – 25868 ( 2020 ). OpenUrl Abstract / FREE Full Text 52. ↵ Zhan , J. et al. Culture medium of bone marrow-derived human mesenchymal stem cells effects lymphatic endothelial cells and tumor lymph vessel formation . Oncol. Lett . 9 , 1221 – 1226 ( 2015 ). OpenUrl PubMed 53. ↵ Alderfer , L. , Wei , A. & Hanjaya-Putra , D. Lymphatic Tissue Engineering and Regeneration . J. Biol. Eng . 12 , 32 ( 2018 ). OpenUrl CrossRef PubMed 54. ↵ Mousaei Ghasroldasht , M. , Seok , J. , Park , H.-S. , Liakath Ali , F. B. & Al-Hendy , A. Stem Cell Therapy: From Idea to Clinical Practice . Int. J. Mol. Sci . 23 , 2850 ( 2022 ). OpenUrl PubMed 55. ↵ Saeedi , P. , Halabian , R. & Imani Fooladi , A. A . A revealing review of mesenchymal stem cells therapy, clinical perspectives and Modification strategies . Stem Cell Investig . 6 , 34 – 34 ( 2019 ). OpenUrl PubMed 56. ↵ Mastrolia , I. et al. Challenges in Clinical Development of Mesenchymal Stromal/Stem Cells: Concise Review . Stem Cells Transl. Med . 8 , 1135 – 1148 ( 2019 ). OpenUrl CrossRef PubMed 57. Bourin , P. et al. Stromal cells from the adipose tissue-derived stromal vascular fraction and culture expanded adipose tissue-derived stromal/stem cells: a joint statement of the International Federation for Adipose Therapeutics and Science (IFATS) and the International Society for Cellular Therapy (ISCT) . Cytotherapy 15 , 641 – 648 ( 2013 ). OpenUrl CrossRef PubMed Web of Science 58. ↵ Aithal , A. P. , Bairy , L. K. & Seetharam , R. N . Safety and therapeutic potential of human bone marrow-derived mesenchymal stromal cells in regenerative medicine . Stem Cell Investig . 8 , 10 ( 2021 ). 59. ↵ Zhang , Y. et al. Lymphangiogenesis in renal fibrosis arises from macrophages via VEGF-C/VEGFR3-dependent autophagy and polarization . Cell Death Dis . 12 , 109 ( 2021 ). 60. ↵ Jeltsch , M. , et al. CCBE1 Enhances Lymphangiogenesis via A Disintegrin and Metalloprotease With Thrombospondin Motifs-3–Mediated Vascular Endothelial Growth Factor-C Activation . Circulation 129 , 1962 – 1971 ( 2014 ). OpenUrl Abstract / FREE Full Text 61. ↵ Akwii , R. G. , Sajib , M. S. , Zahra , F. T. & Mikelis , C. M . Role of Angiopoietin-2 in Vascular Physiology and Pathophysiology . Cells 8 , 471 ( 2019 ). 62. ↵ Morisada , T. et al. Angiopoietin-1 promotes LYVE-1-positive lymphatic vessel formation . Blood 105 , 4649 – 4656 ( 2005 ). OpenUrl Abstract / FREE Full Text 63. ↵ Lee , J.-S. , Tocheny , C. E. & Shaw , L. M . The Insulin-like Growth Factor Signaling Pathway in Breast Cancer: An Elusive Therapeutic Target . Life 12 , 1992 ( 2022 ). OpenUrl PubMed 64. ↵ Björndahl , M. et al. Insulin-like growth factors 1 and 2 induce lymphangiogenesis in vivo . Proc. Natl. Acad. Sci . 102 , 15593 – 15598 ( 2005 ). OpenUrl Abstract / FREE Full Text 65. ↵ Yu , P. et al. FGF-dependent metabolic control of vascular development . Nature 545 , 224 – 228 ( 2017 ). OpenUrl CrossRef PubMed 66. Cao , R. et al. Collaborative interplay between FGF-2 and VEGF-C promotes lymphangiogenesis and metastasis . Proc. Natl. Acad. Sci . 109 , 15894 – 15899 ( 2012 ). OpenUrl Abstract / FREE Full Text 67. ↵ Platonova , N. et al. Evidence for the interaction of fibroblast growth factor-2 with the lymphatic endothelial cell marker LYVE-1 . Blood 121 , 1229 – 1237 ( 2013 ). OpenUrl Abstract / FREE Full Text 68. ↵ Lee , J. Y. et al. Podoplanin-Expressing Cells Derived From Bone Marrow Play a Crucial Role in Postnatal Lymphatic Neovascularization . Circulation 122 , 1413 – 1425 ( 2010 ). OpenUrl Abstract / FREE Full Text 69. ↵ Kerrigan , A. M. et al. Podoplanin-expressing inflammatory macrophages activate murine platelets via CLEC-2 . J. Thromb. Haemost . 10 , 484 – 486 ( 2012 ). OpenUrl CrossRef PubMed 70. ↵ Kuonqui , K. et al. Dysregulation of Lymphatic Endothelial VEGFR3 Signaling in Disease . Cells 13 , 68 ( 2023 ). 71. ↵ Leppänen , V.-M. et al. Structural and mechanistic insights into VEGF receptor 3 ligand binding and activation . Proc. Natl. Acad. Sci . 110 , 12960 – 12965 ( 2013 ). OpenUrl Abstract / FREE Full Text 72. ↵ Dashtban , M. , Panchalingam , K. M. , Shafa , M. & Ahmadian Baghbaderani , B. Addressing Manufacturing Challenges for Commercialization of iPSC-Based Therapies . in Stem Cells and Good Manufacturing Practices (ed. Turksen, K.) vol. 2286 179 – 198 (Springer US, New York, NY, 2020 ). OpenUrl 73. ↵ Chakraborty , A. , Scogin , C. K. , Rizwan , K. , Morley , T. S. & Rutkowski , J. M . Characterizing Lymphangiogenesis and Concurrent Inflammation in Adipose Tissue in Response to VEGF-D . Front. Physiol . 11 , ( 2020 ). 74. Westcott , G. P. & Rosen , E. D. Crosstalk Between Adipose and Lymphatics in Health and Disease . Endocrinology 163 , bqab224 ( 2021 ). OpenUrl 75. ↵ Savetsky , I. L. et al. Obesity increases inflammation and impairs lymphatic function in a mouse model of lymphedema . Am. J. Physiol.-Heart Circ. Physiol . 307 , H165 – H172 ( 2014 ). OpenUrl CrossRef PubMed View the discussion thread. Back to top Previous Next Posted September 25, 2025. Download PDF Email Thank you for your interest in spreading the word about bioRxiv. NOTE: Your email address is requested solely to identify you as the sender of this article. Your Email * Your Name * Send To * Enter multiple addresses on separate lines or separate them with commas. You are going to email the following A novel approach for reliable differentiation of lymphatic endothelial progenitor cells in vitro Message Subject (Your Name) has forwarded a page to you from bioRxiv Message Body (Your Name) thought you would like to see this page from the bioRxiv website. Your Personal Message CAPTCHA This question is for testing whether or not you are a human visitor and to prevent automated spam submissions. Share A novel approach for reliable differentiation of lymphatic endothelial progenitor cells in vitro Inazio Arriola-Alvarez , Maria Rodriguez-Hidalgo , Hector Lafuente , Sonia Alonso-Martin , Ander Izeta bioRxiv 2025.09.23.677051; doi: https://doi.org/10.1101/2025.09.23.677051 Share This Article: Copy Citation Tools A novel approach for reliable differentiation of lymphatic endothelial progenitor cells in vitro Inazio Arriola-Alvarez , Maria Rodriguez-Hidalgo , Hector Lafuente , Sonia Alonso-Martin , Ander Izeta bioRxiv 2025.09.23.677051; doi: https://doi.org/10.1101/2025.09.23.677051 Citation Manager Formats BibTeX Bookends EasyBib EndNote (tagged) EndNote 8 (xml) Medlars Mendeley Papers RefWorks Tagged Ref Manager RIS Zotero Tweet Widget Facebook Like Google Plus One Subject Area Bioengineering Subject Areas All Articles Animal Behavior and Cognition (7640) Biochemistry (17706) Bioengineering (13902) Bioinformatics (41978) Biophysics (21465) Cancer Biology (18611) Cell Biology (25528) Clinical Trials (138) Developmental Biology (13387) Ecology (19920) Epidemiology (2067) Evolutionary Biology (24332) Genetics (15615) Genomics (22519) Immunology (17747) Microbiology (40424) Molecular Biology (17194) Neuroscience (88662) Paleontology (667) Pathology (2839) Pharmacology and Toxicology (4827) Physiology (7650) Plant Biology (15160) Scientific Communication and Education (2046) Synthetic Biology (4302) Systems Biology (9826) Zoology (2271)