Abstract Background & Aims Cirrhosis profoundly impacts extrahepatic vasculature, particularly altering the portal venous system, leading to increased portal pressure, portosystemic collaterals, and portal vein thrombosis, which heightens morbidity and reduces survival in liver disease. Although intrahepatic vascular changes in cirrhosis are well studied, molecular insights into extrahepatic alterations in the splanchnic region remain limited owing to the inaccessibility of the human portal vein and suboptimal preclinical models. Here, we aim to isolate, characterize, and immortalize primary human portal vein endothelial cells (PVECs) to enhance understanding of pathophysiological changes during liver disease and establish a platform for future drug testing. Methods PVECs (n = 12) and inferior cava vein (ICV, n = 9) endothelial cells (ECs) were isolated from human portal vein or ICV, obtained during hepatic transplantation, using trypsinization, mechanical scratching, and FACS. EC identity was confirmed through characterization of gene and protein marker expression as well as functional assays assessing angiogenic capacity (tube formation), migratory ability (wound closure), and acetylated low-density lipoprotein uptake. PVECs were immortalized (iPVECs) with lentiviral particles expressing the SV40 large T-antigen. Results Isolated PVECs confirmed classical endothelial morphology and functionality, expressing hallmark proteins and functions. PVECs exhibited a distinct transcriptomic profile from ICVEC and systemic ECs, enriched in pathways for vascular remodeling and stress response. iPVECs retained endothelial identity and preserved the PVEC-specific transcriptomic traits across more than 20 passages. Conclusions We successfully isolated, characterized, and immortalized PVECs, creating a novel tool to study splanchnic vascular diseases. These cells retain transcriptomic uniqueness distinct from systemic venous ECs, enabling investigation of vascular dysfunction mechanisms in liver disease and supporting translational research. Impact and implications Portal hypertension and vascular complications are major drivers of morbidity in cirrhosis, yet extrahepatic vascular mechanisms remain poorly understood due to limited access to human portal vein tissue and inadequate models. By isolating, characterizing, and immortalizing primary human portal vein endothelial cells, we establish the first renewable, disease-relevant platform for studying splanchnic vascular biology. These immortalized portal vein endothelial cells preserve endothelial identity and transcriptomic signatures distinct from systemic venous cells, providing unique insights into vascular remodeling and stress responses in liver disease. This resource enables mechanistic discovery and drug testing aimed at improving outcomes in portal hypertension and related complications. Keywords: Cirrhosis, Vascular dysfunction, Splanchnic system, Vascular endothelium Graphical abstract [50]Image 1 [51]Open in a new tab Highlights * • Isolation and characterization of primary human portal vein endothelial cells. * • Identification of differences in endothelial cells between the splanchnic and systemic territories. * • Immortalized PVECs provide a novel tool for studying splanchnic vein vasculature. Introduction Portal hypertension (PHT) induces profound changes in the portal venous system (PVS), impairing vascular function and leading to major complications. Although intrahepatic vascular changes in cirrhosis and PHT are well studied,[52]^1 molecular insights into extrahepatic vascular remodeling in the splanchnic territory have been largely neglected. This gap is critical, as the splanchnic circulation is the site of key PHT-related complications, including portosystemic collateral formation and portal vein thrombosis (PVT). The PVS differs markedly from the systemic venous system. Embryologically, it originates from the vitelline and umbilical veins, forming the main portal vein via selective involution and anastomoses. Anatomically, it lacks venous valves, drains the capillary network (hepatic sinusoids), and operates under low pressure, slow flow, and high compliance.[53]^2 Its position also exposes it to high concentrations of metabolites and toxins,[54]^3 likely shaping a distinctive endothelial phenotype. Understanding the PVS endothelium at the molecular level is key for developing better therapies, particularly for cirrhosis-related vascular complications, such as PHT and portosystemic collaterals formation, given the pivotal role of endothelial cells (ECs) in maintaining vascular homeostasis.[55]^4^,[56]^5 The role of the extrahepatic endothelium in liver disease-related vascular alterations and frequent vascular events such as PVT, remains poorly understood despite its major impact on outcomes in liver transplant candidates. Endothelial dysfunction may disrupt vascular homeostasis and accelerate liver disease progression, yet the underlying pathways are still unclear.[57][6], [58][7], [59][8] Existing animal models of PHT fail to replicate the slow, progressive nature of human cirrhosis and its vascular changes. They lack key factors such as metabolic influences, obesity, alcohol use, and infections, which drive proinflammatory activation and exacerbate PH. Moreover, rodent models present distinct anatomical and cellular characteristics in their vasculature,[60]^9^,[61]^10 limiting their relevance—they do not develop spontaneous PVT or reproduce the human rebalanced coagulation system—and underscoring the urgent need for more representative models.[62]^11^,[63]^12 Studies on the PVS endothelium have been limited by the inaccessibility of the portal vein. Most insights rely on the systemic venous system, ECs from the umbilical cord (e.g. human umbilical vein endothelial cells [HUVECs]) or intrahepatic ECs from the sinusoid (liver sinusoidal endothelial cells), which may not fully reflect the physiological and pathological characteristics unique to the extrahepatic PVS. This gap hinders our understanding of the complex vascular changes in cirrhosis and limits therapeutic discovery. To overcome this, we aim to isolate, characterize, and immortalize primary human portal vein endothelial cells (PVECs) to create a novel model for studying cellular mechanisms underlying splanchnic vascular dysfunction. Materials and methods Human portal vein tissue obtention We prospectively collected extrahepatic portal vein and inferior cava vein (ICV) samples from explanted livers of 12 patients with cirrhosis undergoing liver transplantation at Hospital Clínic de Barcelona (2020–2022). Paired PVECs and inferior cava vein endothelial cells (ICVECs) were obtained from nine patients, whereas only PVECs were isolated from the remaining patients. Patients with active HCV, HBV, HDV, or HIV infections at the time of surgery or PVT were excluded. The study was approved by the Hospital Clínic ethics committee (HCB/2018/1246) and conducted according to the Declaration of Helsinki, with written informed consent obtained from all participants. Within 10–60 min post resection, surgeons excised 1- to 2-cm tubular portal vein segments, which were placed in sterile Falcon tubes containing EGM™-2 medium (Cultek, Madrid, Spain, CC-3162) at 4 °C. Samples were processed within 1–6 h. EC isolation and culture from the human portal vein and ICV For EC isolation from vein tissue, excess surrounding fat tissue was meticulously removed from the vein using sterile scissors, and any residual blood was washed away with DPBS. Subsequently, the tubular vein was cut open along its long axis using small scissors and placed in a cell culture dish, exposing the inner part of the vein facing upwards. Trypsin EDTA 0.05% (Gibco, Paisley, UK, #25300054) was then added to the upper surface and incubated for 5 min at 37 °C. EGM-2 medium was then added abundantly over the tissue, and a cell scraper (TPP-Reactiva, Trasadingen, Switzerland, #99002) was used to carefully scrape the surface of the inner wall several times to detach the ECs from the vein wall. The resulting cell suspension was then centrifuged at 300 × g for 5 min. The pellet was resuspended in 2 ml of fresh EGM-2 and seeded onto a six-well plate precoated with human plasma fibronectin (1 μg/cm^2;Sigma, Darmstadt, Germany, #F0895). The media was refreshed every other day until confluence ([64]Fig. S1). To isolate ECs from other cell types scraped during tissue processing, cells were trypsinized, stained with anti-CD31-FITC and CD144-BV786 antibodies, and FACS-sorted. ECs were selected as CD31^+ CD144^+ double-positive cells ([65]Fig. S2A) and seeded back to either a six-well plate or T25 flask based on yield. A reanalysis of the sorted population was consistently performed to ensure the purity of our EC fraction after sorting ([66]Fig. S2B). Once confluent, one-third of the ECs were harvested in RLT buffer (Qiagen, Hilden, Germany) for RNA extraction (passage 1), one-third were used for flow cytometry purity analysis, and the remaining one-third were expanded further. PVECs were then passaged at a 1:5 ratio in fibronectin-coated flasks until senescence. At each passage, EC identity and function were validated by flow cytometry and tube formation assays. All the aforementioned steps were identically applied to isolate both PVECs and ICVECs. The fraction of non-ECs obtained from the sorting (double negative for CD31 and CD144) was cultured in EGM-2 media but without fibronectin coating and was passaged at a ratio of 1:10 owing to their faster growth, serving as a negative control. HUVECs (Lonza, Basel, Switzerland, #CC-2517) cultured in EGM-2 on fibronectin-coated flasks and passaged at a ratio of 1:5 were used as a positive EC control as a widely recognized EC model. Flow cytometry and FACS Expression of endothelial markers CD31 and CD144 was analyzed by flow cytometry at each passage. Briefly, cells were washed, trypsinized, and centrifuged at 300 × g for 5 min. Cells were then resuspended in 100 μl of PBS + 5% FBS (staining buffer), and unspecific binding was blocked with 10% goat serum (Dako, Glostrup, Denmark, #X0907) at 4 °C for 10 min. Cells were then stained with 1 μl of CD31-FITC (BD Bioscience, San Jose, CA, USA#555445) and 1 μl of CD144-BV786 (BD Bioscience, #565672) antibodies at 4 °C in the dark for 20 min. Antibodies were washed by adding 1 ml of staining buffer and centrifuged at 300 × g for 5 min. Cells were finally resuspended in 200 μl of staining buffer containing 1 μg/ml DAPI (4′,6-diamidino-2-phenylindole; Thermo Fisher, Waltham, MA, USA, #D1306) and either analyzed using a FACS Canto III flow cytometer (BD Bioscience) or sorted in a FACS Aria II cell sorter, as described above. The same protocol was applied when performing the expression analysis of CLEC2 (BD #755872). Data were analyzed using Flowjo 10.10 software. Immortalization of PVECs Passage 1 PVECs were seeded in 24-well plates at 1 × 10^4 cells/ml. At 70% confluency, cells were transduced with 1 μl/ml of lentiviral particles (Multiplicity of infection of 5) expressing the SV40 large T-antigen and containing the GFP–puromycin fusion dual marker (Amsbio, Abingdon, Oxfordshire, UK, #LVP016-GP-PBS). After 72 h, transduced cells were selected with EGM-2 medium containing 1.5 μg/ml puromycin, and GFP^+ expression was confirmed by flow cytometry. Immortalized PVECs (iPVECs) were expanded for 25–40 passages and continuously characterized as described for primary PVECs. Bioinformatics analysis Passage 1 PVECs (n = 12), ICVECs (n = 9), and passage 20 iPVECs (n = 6) were collected in RLT buffer combined with β-mercaptoethanol, and RNA was extracted using the RNeasy Micro Kit (Qiagen) with on-column DNAseI digestion according to the manufacturer’s instructions. RNA quality was assessed by a Bioanalyzer, and only samples with an RNA integrity number (RIN) >8 were processed further. Libraries were generated using the NEBNext Ultra II Directional RNA Library Prep Kit for Illumina (New England Biolabs, Ipswich, MA, USA) and sequenced in paired-end mode (2 × 50 bp) on an Illumina NovaSeq 6000 sequencer, achieving at least 30 million reads per sample. The quality of the sequences was assessed using FastQC (version 0.11.9, Babraham Institute, Cambridge, UK). Reads were pseudoaligned to the human genome (version GRCh38.p13) using the Kallisto pseudoaligner (version 0.46.1, Pachter Lab, Caltech, Pasadena, CA, USA). Transcript and gene-level quantifications, including transcripts per million (TPM) calculations and differential expression analysis, were performed using the sleuth R package (version 0.30.1; Integrated Development Environment for R. Posit Software, PBC, Boston, MA, USA), with an exclusive focus on protein-coding genes. Differentially expressed genes (DEGs) were identified using the Wald test on aggregated transcript p values, with significance thresholds set at p value <0.05 and fold discovery range (FDR) <0.2. RNA-sequencing (RNA-seq) data have been deposited in NCBI’s Gene Expression Omnibus (GEO accession number [67]GSE296600). GEO data sample collection We retrieved RNA-seq data for human EC lines from various vascular territories, including venous, arterial, and lymphatic origins, from the NCBI GEO database (see GEO accession numbers in [68]Table S1). Only untreated control samples were included. FASTQ files were processed using Kallisto pseudoalignment as described above. To enable data visualization, all samples were normalized together using the sleuth package, and the resulting normalized counts and TPM values were used for subsequent analyses. Statistical analysis Analyses were performed using R (version 4.3.1; R Foundation for Statisitcal Computing). Normality and homoscedasticity were assessed using Shapiro’s and Levene’s tests, respectively. For normally distributed, homoscedastic data, group comparisons were made using ANOVA followed by Tukey’s post hoc test. For non-normal or heteroscedastic data, the Kruskal–Wallis test with Dunn’s post hoc correction was applied. Continuous variables are presented as raw values and visualized in box plots showing median and IQR. Statistical significance was set at p ≤0.05. Results Isolation of ECs from human portal vein tissue ECs were isolated from fresh portal vein or inferior cava vein tissue collected immediately after hepatectomy during liver transplantation. Within 2–6 h after tissue processing, cells attached, spread, and became visible in culture. They began proliferating and typically formed a confluent monolayer within a median of 10 days. This monolayer contained both EC and non-EC populations. [69]Fig. 1A shows the different cell types observed after the isolation procedure. Although ECs would grow in colonies (arrows), mesenchymal cells would grow all along the plate surrounding endothelial colonies (arrow heads). ECs were purified from this mixed population by FACS, selecting CD31 and CD144 (VE-Cadherin) double-positive cells, and then seeded onto fibronectin-coated plates. Fig. 1. [70]Fig. 1 [71]Open in a new tab Isolated human PVECs display characteristic endothelial features. (A) Cells 7 days post isolation showing the typical coexistence of ECs (arrow) and non-ECs (arrowhead). (B, C) Post-sorting PVECs initially growing from small colonies of tightly clustered cells (B) until reaching confluence (passage 1) and (C) exhibiting a uniform monolayer of cobblestone-shaped ECs. (D) PVECs start to exhibit larger and irregular shapes (indicated by arrows) around passages 6–7. (E) By passages 8–9, the cultures typically become senescent, displaying aberrant morphological phenotypes characterized by flattened and enlarged cells. (F) Representative images of immunofluorescence staining of eNOS and vWF in passage 3 cultured PVECs. (G) PVECs from different passages form tube structures when cultured in a Matrigel matrix, maturing into vascular nets after 24 h. Images were acquired at either 5 × or 10 × magnification using an Olympus IX71 microscope. All these images are representative of experiments performed in all PVECs isolated (n = 12). (H–L) qRT-PCR gene expression quantification of hallmark (H) endothelial genes, (I) mesenchymal markers, (J) vasoactive and coagulation molecules, (K) adhesion molecules, and (L) angiogenic factors in HUVECs (n = 5), PVECs (n = 9), ICVECs (n = 5), and non-ECs (n = 5). All qRT-PCR experiments were conducted using cells at passage 2. Quantification of target genes is relative to Gapdh. Results are presented as box plots. Statistical analysis was performed using ANOVA and Tukey’s post hoc test (∗p <0.05). EC, endothelial cell; HUVEC, human umbilical vein endothelial cell; ICVEC, inferior cava vein endothelial cell; PVEC, human portal vein endothelial cell; qRT-PCR, quantitative reverse transcription PCR. After purification, PVEC cultures displayed a uniform cobblestone morphology, initially distributed as small, tightly clustered proliferating colonies ([72]Fig. 1B), which grew to form uniform monolayers ([73]Fig. 1B). The double-negative fractions for CD31 and CD144 ([74]Fig. S3) were used as controls for non-ECs. PVECs maintained growth and expansion in culture for approximately 60 days, with passages occurring every 5–6 days on average at a 1:5 split ratio. Between passages 6 and 9, PVECs began to exhibit a senescent phenotype characterized by enlarged, flattened, multinucleated cells with cytoplasmic granularities and bigger morphologies ([75]Fig. 1D). By passages 7–9, cells ceased proliferating ([76]Fig. 1E). Endothelial characterization of PVECs To confirm that PVECs maintained endothelial identity in vitro, we assessed both functional traits and surface marker expression over time. Flow cytometry analysis at each passage showed stable expression of CD31 and CD144, indicating sustained endothelial phenotype ([77]Fig. S4). Similarly, immunocytochemistry further confirmed membrane expression of eNOS and vWF, two hallmark endothelial proteins, in PVECs ([78]Fig. 1F). To assess angiogenic potential, PVECs were cultured on Matrigel and monitored for tube formation. As shown in [79]Fig. 1G, PVECs formed tubes within 4 h, which matured into vascular networks within 24 h. This ability was sustained at least until passage 6. We analyzed expression of key endothelial transcripts in PVECs by quantitative reverse transcription PCR (qRT-PCR) and compared them with two positive controls—primary ECs from other venous territories (ICVECs and HUVECs)—and with the non-EC fraction (putative mesenchymal cells) as a negative control. As expected, PVECs exhibited expression levels of PECAM1, CDH5, and vWF that were comparable to those in ICVECs and HUVECs, which were higher than those observed in the non-EC fraction ([80]Fig. 1H). Conversely, non-ECs showed high expression levels of the mesenchymal markers CD90 and Col1a1, which were both absent in the three EC lines ([81]Fig. 1I). Likewise, PVECs exhibited production levels of vasoactive and coagulation molecules comparable to those of ICVECs and HUVEC, with subtle non-significant differences in NOS3, EDN1, SERPINE1, PLAT, and TBXAS1 ([82]Fig. 1J). They also expressed the adhesion molecules ICAM1 and SELL ([83]Fig. 1K), as well as the angiogenic factors KDR and ANGPT2 ([84]Fig. 1L), indicating robust endothelial functionality. PVECs show transcriptomic differences with EC from other territories The heterogeneity of human organ-specific ECs is well established, with distinct organ- and vascular bed-specific subtypes. ECs differ not only between arterial and venous origins but also among venous territories. However, the molecular and functional characteristics of PVECs have not been previously characterized. To investigate whether PVECs differ from ECs in the systemic circulation (ICVECs), we conducted bulk RNA-d on passage 1 cells from 12 PVEC and nine ICVEC samples. Principal component analysis of RNA-seq data revealed that samples tended to cluster according to their vascular territory of origin ([85]Fig. 2A). We identified 233 differentially expressed genes between PVECs and ICVECs, of which 164 and 69 were significantly upregulated and downregulated, respectively, in PVECs (fold change ≥1.5; [86]Fig. 2B and [87]Table S2). Differentially expressed genes included genes involved in EC proliferation, growth, and survival (e.g. FIGN and SULF1); cytoskeletal dynamics (e.g. KALRN and TPM2), cellular migration and immune cell trafficking (e.g. SLIT2 and LYVE1); and cell adhesion and structural integrity (e.g. FN1 and ADAMTS23). Fig. 2. [88]Fig. 2 [89]Open in a new tab PVECs exhibit differential gene expression in endothelial markers and functions compared with systemic ECs. (A) Principal component analysis plot showing the distribution of the PVEC and ICVEC samples according to the first two principal components (PCs), which explain 50% of the total variance. (B) GSEA results represented in a dot plot. Significant enriched upregulated or downregulated pathways between the comparisons of PVECs and ICVECs. (C–E) Expression levels of endothelial cell markers for (C) NRP2 (venous), (D) EPHB2 (arterial) and (E) PROX1 (lymphatic) in the different sets of ECs confirm successful isolation of vein ECs for our PVECs and ICVECs. Gene expression levels are presented as TPM for each group. (F) Heatmap showing the DEGs between PVECs and ICVECs when including how these genes are expressed in HUVECs, LECs, and arterial ECs. DEG, differentially expressed gene; EC, endothelial cell; ICVEC, inferior cava vein endothelial cell; LEC, lymph node endothelial cell; PVEC, human portal vein endothelial cell; TPM, transcripts per million; GSEA, gene set enrichment analysis. To explore fundamental differences between PVECs and ICVECs, we performed pathway enrichment analysis. PVECs exhibited upregulation of pathways related to biosynthesis, developmental plasticity, structural remodeling, vascular repair, stress response, and immunity ([90]Fig. 2B and [91]Table S3). These PVEC adaptations may support resistance to mechanical stress, promote tissue repair, and preserve vascular hemostasis, functions that are particularly critical under chronic stress conditions of elevated portal pressure observed in cirrhosis. To further contextualize PVECs, we expanded our analysis beyond ICVECs by incorporating publicly available RNA-seq datasets of primary ECs from other vascular territories obtained from the GEO. These datasets included primary ECs of venous origin (HUVECs), arterial origin (human aortic endothelial cells, human coronary artery endothelial cells, and human pulmonary artery endothelial cells), and lymphatic origin (human dermal or lymph node endothelial cells [LECs]). The venous identity of PVECs was confirmed by evaluating the expression of the venous marker NRP2 across the different cell types. NRP2 was highly expressed in both PVECs and ICVECs but showed minimal expression in arterial cell lines ([92]Fig. 2C). Conversely, markers specific to arterial (EPHB2) or lymphatic (PROX1) ECs exhibited relatively low expression in both PVECs and ICVECs ([93]Fig. 2D and E). Next, we analyzed the gene expression patterns of the 233 DEGs previously identified between PVECs and ICVECs across EC samples from different vascular beds. As shown in [94]Fig. 2F, hierarchical clustering of these DEGs revealed three main clusters: on the right, arterial ECs displayed a highly similar expression profile among themselves, forming a cohesive group. Notably, they exhibited a consistent pattern of gene regulation compared with PVECs, with numerous upregulated genes showing significantly higher expression levels. LECs and ICVECs formed a mixed cluster, with a highly similar expression profile between them. Meanwhile, PVECs grouped closely with HUVECs, forming a distinct intermediate cluster between the arterial and lymphatic clusters. Functionally, ICVECs and LECs displayed upregulation of genes such as PIEZO2, TFPI2, and NOSTRIN, which are involved in mechanostransdution,[95]^13 coagulation inhibition,[96]^14 and nitric oxide regulation.[97]^15 The arterial cluster was characterized by upregulation of genes associated with cellular migration and vascular remodeling (SEMA7A and SLIT2),[98]^16^,[99]^17 consistent with the greater mechanical demands of arterial circulation. PVECs and HUVECs showed distinct upregulation of genes such as IGF2BP1, CCND2, and IL7R, indicative of heightened transcriptional activity and immune-related activity.[100][18], [101][19], [102][20] Moreover, genes commonly upregulated in both arterial and PVECS—such as SULF1 and TWIST—further suggest a more active and remodeling-prone phenotype[103]^21^,[104]^22 compared with systemic ECs. These results support the notion that PVECs represent a unique subset of ECs with distinct molecular characteristics. This highlights their specialized functional traits, likely shaped by the unique microenvironment of the portal vein and the hemodynamic forces of their respective vascular beds, underscoring their value for studying the splanchnic venous territory. Generation of iPVECs To address the limited lifespan of primary ECs, we aimed to develop a more robust model for extended in vitro studies of the splanchnic vasculature endothelium. We immortalized primary PVECs (n = 6) by transducing passage 1 PVECs with lentiviral particles carrying the SV40 large T-antigen and a GFP–puromycin resistance fusion dual marker. Transduced cells were initially selected using puromycin, followed by GFP-positive selection through FACS ([105]Fig. S5A). Successful transduction was also analyzed by measuring SV40 expression by qRT-PCR, which, as expected, was clearly expressed only in transduced PVECs ([106]Fig. S5B). iPVECs displayed consistent proliferation and extended their lifespan to at least 30–40 passages, in contrast to primary PVECs, which began showing signs of senescence by passages 6–7 and rarely exceeded nine passages. Although PVECs maintained normal growth for 11 doublings before slowing, iPVECs preserved a stable doubling rate beyond 40 doublings ([107]Fig. S5C). Overall, iPVECs preserved their phenotype and growth characteristics for at least 140 days in culture, consistently displaying cobblestone morphology similar to primary PVECs. Minor morphological changes, such as cell elongation, were observed beginning around passage 25 ([108]Fig. S5D). iPVEC maintains EC features iPVECs maintained stable protein expression of key endothelial markers CD31 and CD144 across passages, as confirmed by flow cytometry ([109]Fig. 3A). Although gene expression of some endothelial markers declined by passage 20 ([110]Fig. 3B), there was no induction of mesenchymal markers, indicating preserved endothelial identity ([111]Fig. 3C). Although gene expression related to vasoactivity, coagulation, adhesion, and angiogenesis displayed some variability over time, several important markers remained stably expressed ([112]Fig. 3D–F). At the protein level, iPVECs from both early and late passages retained an expression of eNOS and CD36 comparable to that of primary PVECs. Although a trend toward reduced expression of the endothelial marker vWF was observed ([113]Fig. 3G and [114]Fig. S6), this difference was not statistically significant at the protein level. However, as shown before, vWF expression was significantly decreased at the RNA level. The reduction in vWF expression was consistently observed during in vitro culture and after the immortalization process (data not shown). Functionally, iPVECs retained their ability to form tubes when seeded on Matrigel ([115]Fig. S7). Fig. 3. [116]Fig. 3 [117]Open in a new tab iPVECs retain the expression of endothelial markers and functional genes. (A) Representative histogram comparing the analysis of CD144 expression levels between non-ECs, passage 1 PVECs, and iPVECs at different passages. All PVEC (n = 12) and iPVEC (n = 6) samples were tested for the expression of CD144. (B–F) qRT-PCR gene expression quantification of hallmark (B) endothelial genes, (C) mesenchymal markers, (D) vasoactive and coagulation molecules, (E) adhesion molecules, and (F) angiogenic factors in iPVECs (n = 6), PVECs (n = 9), ICVECs (n = 4), and non-ECs (n = 5). Quantification of target genes is relative to Gapdh, and results are presented relative to the same controls shown in [118]Fig. 1. Control data are not included in this graph to avoid redundancy. Results are presented as box plots. Statistical analysis was performed using the Mann–Whitney U test (∗p <0.05). (G) Representative images of Western blots of eNOS, CD36, and vWf proteins from HUVECs (n = 3), ICVECs (n = 3), primary PVECs (n = 4), and iPVECs at different passages (n = 4). Results were normalized to β-actin and are shown as box plots for each group. Statistical analysis was performed using ANOVA and Tukey’s post hoc test (∗p <0.05). (H) Representative images for ac-LDL uptake assay in primary PVECs (n = 3) and iPVECs at early (passage 5, n = 4; passage 10, n = 4) and late passages (passage 15, n = 3; passage 20, n = 4). Data are presented as box plots for each group showing no significant differences in ac-LDL uptake between groups. Statistical analysis was performed using ANOVA and Tukey’s post hoc test. (I) Representative images for wound healing assay in primary PVECs and iPVECs at passage 20. Results are presented as a time graph for each group (PVEC, n = 3; iPVEC p5, n = 3; iPVEC p10, n = 3; iPVEC p15, n = 3; iPVEC p20, n = 3) showing a similar pattern and rates of scratch closure. EC, endothelial cell; HAEC, human aortic endothelial cell; HCAEC, human coronary artery endothelial cell; HPAEC, human pulmonary artery endothelial cell; HUVEC, human umbilical vein endothelial cell; ICVEC, inferior cava vein endothelial cell; iPVEC, immortalized portal vein endothelial cell; LEC, lymph node endothelial cell; PVEC, human portal vein endothelial cell. Moreover, the ac-LDL uptake assay[119]^23^,[120]^24 demonstrated that iPVECs maintained a comparable ability to internalize ac-LDL, as observed in primary PVECs, with no significant differences between the cell lines ([121]Fig. 3I). Similarly, the wound healing assay demonstrated that iPVECs displayed a comparable ability to migrate and repair damage, showing very similar closure rates and patterns ([122]Fig. 3J). These findings suggest that although iPVECs present some genotypic alterations, potentially associated with prolonged culture and immortalization, they preserve critical endothelial properties, validating their continued biological relevance after immortalization. Importantly, iPVECs did not exhibit tumorigenic potential, as they failed to form colonies in a soft agar anchorage-independence assay. Both PVECs and iPVECs remained as single cells as opposed to the human epithelial cells from pancreatic duct carcinoma, used as a positive control ([123]Fig. S8). Maintenance of portal vein-specific transcriptomic traits in iPVECs To assess whether iPVECs retained portal vein-specific traits, we performed RNA-seq on six iPVEC lines and compared them with primary PVECs. To obtain a global picture on immortalization-induced alterations in gene expression of iPVECs, we performed a gene set pathway enrichment analysis. Results revealed that most transcriptional changes in iPVECs were related to cell cycle regulation, DNA repair, and cellular maintenance ([124]Fig. S9 and [125]Table S4), hallmarks of the immortalization process. Analysis of previously identified DEGs between PVECs and ICVECs showed that iPVECs closely mirrored the PVEC expression profile, retaining key portal-specific signatures and remaining distinct from ICVECs ([126]Fig. 4A). Fig. 4. [127]Fig. 4 [128]Open in a new tab iPVECs maintain their distinctive gene expression profile. (A) Heatmap showing the DEGs between PVECs and ICVECs with iPVECs incorporated. (B) qRT-PCR gene expression quantification of the selected genes from the DEGs shown in (B), ICVECs (n = 3), PVEC (n = 9), and iPVECs (n = 6). iPVECs display consistent expression of these genes when compared with PVECs. Quantification of target genes is relative to Gapdh. Results are presented as box plots. Statistical analysis was performed using ANOVA and Tukey’s post hoc test (∗p <0.05). (C) Percentage of membrane protein CLEC2 in the three different groups (PVECs, n = 4; iPVECs, n = 5; ICVECs, n = 5) assessed by flow cytometry. Results are shown as box plots for the geometrical mean. (D) Western blot analysis of ABCB1, TPM2, and FN1 proteins normalized to β-actin in ICVECs (n = 3), primary PVECs (n = 4), and iPVECs at different passages (n = 6). Results are shown as box plots for each group. DEG, differentially expressed gene; EC, endothelial cell; ICVEC, inferior cava vein endothelial cell; iPVEC, immortalized portal vein endothelial cell; PVEC, human portal vein endothelial cell; qRT-PCR, quantitative reverse transcription PCR. Finally, RNA-seq results were validated at both gene and protein levels using qRT-PCR, flow cytometry, and Western blot. Consistent with the RNA-seq results, qRT-PCR confirmed that iPVECs retained the DEG expression patterns observed in primary PVECs compared with ICVECs, indicating that immortalization did not significantly alter their portal-specific expression profile ([129]Fig. 4B). Similarly, we confirmed at the protein level the differences between PVECs and ICVECs in the expression of genes related to platelet-endothelium interactions (CLEC2) ([130]Fig. 4C), the endothelial barrier (ABCB1), cytoskeletal dynamics (TPM2) and extracellular matrix assembly and cell adhesion (FN1) ([131]Fig. 4D). As shown in [132]Fig. 4C and D, we confirmed that these differences were maintained also in iPVECs. These findings indicate that iPVECs retain the key molecular and functional features of primary PVECs despite immortalization, preserving their endothelial identity. As such, iPVECs represent a robust and reliable preclinical model for studying splanchnic vascular diseases such as PVT and hold promise for drug screening and therapeutic testing. Discussion During chronic liver disease, the splanchnic endothelium undergoes significant changes that exacerbate and perpetuate the disease.[133]^25^,[134]^26 Increased pressure in the portal vein and the development of portosystemic collaterals and hyperdynamic circulation are hallmarks of the disease, profoundly impacting prognosis.[135]^27 Although drugs that modify vascular tone in the PVS are available,[136]^28 the molecular regulation of endothelial changes during disease progression and regression remains poorly understood. Most current knowledge derives from animal models or indirect surrogates of vascular dysfunction, as studying the PVS in living patients is ethically impractical because of its remote location and the invasiveness of required procedures. We present a novel biological tool to advance research on splanchnic venous alterations in cirrhosis. By isolating and immortalizing PVECs from liver transplant patients, we established and comprehensively characterized cell lines at the functional, phenotypic, and molecular levels. These iPVECs offer a valuable platform for studying the mechanism underlying extrahepatic vascular pathology and hold strong potential for preclinical drug testing. Historically, PVECs have primarily been obtained from non-human species such as dogs, pigs, or rats.[137][29], [138][30], [139][31] However, alterations in the PVS typically result from chronic processes that are challenging to replicate in model organisms. Although rodents have been extensively used to study liver cirrhosis and PHT, their disease progression does not fully mimic the natural course observed in humans, especially in the PVS. Human cirrhosis develops over an extended period with distinct underlying causes that significantly alter the portal circulatory environment and are very difficult to reproduce in animal models, which undergo a more acute onset of liver damage. These factors contribute to the vascular activation of proinflammatory immune modulators, driving the development or worsening of portal hypertensive syndromes, processes that animal models fail to capture and underscoring the critical need for isolating and investigating human PVECs to conduct pathophysiological studies that accurately reflect human conditions. Here, we demonstrated that primary PVECs maintain their endothelial identity both functionally and transcriptionally in vitro. Importantly, PVECs exhibit distinct transcriptomic profiles compared with venous ECs from other vascular territories. The unique environment and localization of PVECs, characterized by high compliance capacity in health and altered flow dynamics during liver disease, seem to shape their gene expression. Transcriptomic comparisons with non-venous ECs underscore the unique molecular profile of PVECs, likely influenced by their embryonic origin,[140]^32 splanchnic localization, and exposure to gut-derived factors. These findings emphasize the need to study PVECs specifically in the context of cirrhosis. Using non-PVEC endothelial models risks overlooking critical transcriptomic distinctions unique to the PVS. This underscores the unique biology of the PVS and the importance of dedicated, PVEC-specific research rather than generalizing splanchnic complications as typical venous disorders. We acknowledge several limitations in this study. Firstly, ECs may undergo phenotypic changes in vitro, potentially masking subtle in vivo differences between EC types. However, even when PVECs and ICVECs were isolated and cultured under identical conditions, they consistently maintained distinct gene expression profiles, supporting the robustness of their intrinsic molecular identity. This observation suggests that differences between these cell types are substantial enough to persist in vitro, at least for a certain period. Notably, the transcriptomic analyses were conducted in early passage 1 cells, which likely contributed to unveiling these differences. Secondly, the ECs used in this study were obtained from liver transplant candidates. Therefore, the findings may not fully represent less advanced stages of the disease or the context of disease regression. However, as accessing the portal vein endothelium is a highly invasive procedure, typically only feasible during liver transplantation, and the absence of suitable animal models for PVECs limits translational research, our study holds significance as the best available approach for obtaining human-derived data on this pathology. A major challenge in this study was the inability to isolate PVECs from healthy donor livers, likely because of prolonged and variable cold ischemia times, compromising cell viability. In contrast, portal vein remnants from cirrhotic explants—processed without such ischemic stress—enabled consistent and efficient PVEC isolation for downstream analyses. To address the challenge of limited proliferation capacity in primary cultures[141]^33^,[142]^34 and guarantee the use of PVECs for future studies, we immortalized six primary PVEC lines from patients with cirrhosis using SV40T. These iPVECs sustained at least 35 passages without signs of senescence or mesenchymal transformation, enabling the generation of substantial cell stocks for extended experimentation. Concerns regarding SV40T-induced alterations in cell identity[143]^35^,[144]^36 were mitigated as iPVECs maintained protein expression of most key endothelial markers and preserved most of their original characteristics over extended passages. Although some genotypic and transcriptional changes associated with immortalization were observed, these alterations did not compromise iPVEC functionality or endothelial identity. Notably, expression of most PVEC-specific genes remained preserved. In conclusion, iPVECs are a robust tool for both basic and translational research. They retain portal vein-specific features after immortalization and overcome key limitations of existing endothelial models by providing a physiologically relevant platform for studying splanchnic vascular complications such as PHT. Their applicability to drug screening and integration into advanced systems such as organoids or microfluidics may further enhance their value for developing targeted therapies for region-specific vascular diseases. Their derivation from cirrhotic livers enables the study of mechanisms underlying cirrhosis-associated vascular complications and advances the development of targeted, precision-based therapeutic strategies. Abbreviations DEG, differentially expressed gene; EC, endothelial cell; FDR, fold discovery range; HUVEC, human umbilical vein endothelial cell; ICV, inferior cava vein; ICVEC, inferior cava vein endothelial cell; iPVEC, immortalized portal vein endothelial cell; LEC, lymph node endothelial cell; PCA, principal component analysis; PHT, portal hypertension; PVEC, human portal vein endothelial cell; PVS, portal venous system; PVT, portal vein thrombosis; qRT-PCR, quantitative reverse transcription PCR; RNA-seq, RNA-sequencing; TPM, transcripts per million. Financial support This work was supported by FIS PI20/00569 and FIS PI23/00997, funded by the “Instituto de Salud Carlos III” and co-funded by the European Union; by Ministerio de Economía y Competitividad (SAF2019: PID2019-105148RB-I00); and by “CIBEREHD,” funded by the “Instituto de Salud Carlos III”. AA has an FPI grant from Ministerio de Economía y Competitividad with file code PRE2020-093316 (AEI_FPI20) related to Grant SAF2019: PID2019-105148RB-I00. RM has an PFIS grant from Ministerio de Economía y Competitividad with file code FI21/00064 (FIS_FI21). SS has a Rio Hortega Grant. The current contract is funded by the “Instituto de Salud Carlos III” with charges to the European funds of the Recovery, Transformation, and Resilience Plan (Plan de Recuperación, Transformación y Resiliencia), with file code CM23/00068, pursuant to the Resolution of the Instituto de Salud Carlos III of December 2023, granting the Rio Hortega Contracts, and “Co-Financed by the European Union.” OT-C was supported by funding from a Fondo de Investigación Sanitaria (FIS) Contratos Miguel Servet tipo II, CPII22/00006. VH-G was supported by a research intensification grant (INT21/00011) from the “Instituto de Salud Carlos III” (AES 2021). Authors’ contributions Contributed to study design: AA, GC, OT-C, JA. Contributed to sample collection: AB, YF, JCo. Conducted experiments: AA, GC, RM, HG-C, LS, JCa. Performed data analysis: AA, GC. Contributed to results discussion: SS, JCG-P. Wrote the manuscript: AA, GC, SS, VH-G. Provided economic support: JCG-P. Provided study guidance: JCG-P. Provided critical manuscript revision: JCG-P. Designed the research: VH-G. Conceived ideas: VH-G. Obtained funding: VH-G. Directed the study: VH-G. Edited and reviewed the final manuscript: all authors. Data availability All sequencing data generated by this study have been deposited in the Gene Expression Omnibus (GEO), accession number [145]GSE296600. All relevant data are within the manuscript and Supplementary data. Conflicts of interest The authors have nothing to report. Please refer to the accompanying ICMJE disclosure forms for further details. Acknowledgements