In Vitro Induction of Human Adipose-Derived Stem Cells into Lymphatic Endothelial-Like Cells

Abstract

Human adipose-derived stem cells (hADSCs) may provide a suitable number of progenitors for the treatment of lymphatic edema; however, to date the protocols for inducing hADSCs into this tissue type have not been standardized. We wished to investigate the induction of hADSCs into lymphatic endothelial-like cells using vascular endothelial growth factor-C156S (VEGF-C156S) and other growth factors in vitro. hADSCs from healthy adult adipose tissue were purified using enzyme digestion. Differentiation was induced using medium containing VEGF-C156S and bovine fibroblast growth factor (bFGF). Differentiation was confirmed using immunostaining for lymphatic vessel endothelial hyaluronan receptor (LYVE-1) and fms-related tyrosine kinase 4 (FLT-4), two lymphatic endothelial cell markers. The expression levels of LYVE-1, prospero homeobox 1 (PROX-1), and FLT-4 throughout induction were assessed using reverse transcriptase quantitative polymerase chain reaction. hADSCs were successfully obtained by trypsin digest and purification. Flow cytometry showed these cells were similar to mesenchymal stem cells, with a high positive rate of CD13, CD29, CD44, and CD105, and a low positive rate of CD31, CD34, CD45, and HLA-DR. Induction to lymphatic endothelial-like cells was successful, with cells expressing high levels of LYVE-1, PROX-1, and FLT-4. Adipose-derived stem cells can be induced to differentiate into lymphatic endothelial-like cells using a medium containing VEGF-C156S, bFGF, and other growth factors. This population of lymphatic endothelial-like cells may be useful for lymphatic reconstruction in the future.

Introduction

Limb lymphedema is a pathophysiological process of lymph stasis due to lymph reflux disorder, which is often secondary to parasitic infections, malignancy, surgery, and radiotherapy. Despite numerous efforts, no effective treatment has emerged (Campisi and Boccardo, 2002; Felmerer, 2012; Vignes and Trévidic, 2002). Many studies support the idea that reconstruction of the lymphatic circulation is an ideal treatment of lymphedema (Boccardo et al., 2013; Campisi et al., 2003). However, the challenge of promoting lymphatic endothelial proliferation or transformation has not yet been addressed.

Lymphatic reconstruction has been demonstrated in animal studies using mesenchymal stem cells (MSCs) isolated from the bone marrow (Lee et al., 2010). However, these bone marrow–derived mesenchymal stem cells (BM-MSCs) are technically challenging to isolate, requiring a painful and dangerous surgical procedure. Additionally, the BM-MSCs are too few in number to be useful as a source of seed cells for tissue reconstruction (Gimble and Guilak, 2003a).

One proposed alternative to BM-MSCs are adipose tissue-derived stem cells (ADSCs), adult stem cells that can proliferate, self-renew, and undergo many differentiation programs, such as adipogenesis, osteogenesis, chrondrogenesis, myogenesis, and neurogenesis, supporting their use in tissue and cell repair (Gimble et al., 2003a; Gimble et al., 2003b; Zuk et al., 2001; Zuk et al., 2002). ADSCs are similar to BM-MSCs in cell morphology, proliferation, senescence, and multipotential differentiation (De Ugarte et al., 2003). Unlike BM-MSCs, ADSCs can be acquired relatively simply from the abundant stores of adipose tissue in the body.

To date, the methodology for inducing differentiation of MSCs to lymphatic endothelium-like cells has not been standardized. Stimulation of BM-MSCs with vascular endothelial growth factor-C (VEGF-C), a ligand of VEGFR2 and VEGFR3, induces expression of the lymphangiogenic markers podoplanin and lymphatic vessel endothelial hyaluronan receptor (LYVE-1) (Lee et al., 2010); however, this expression was only temporary, suggesting that a different stimulus may be necessary. ADSCs pretreated with native VEGF-C and transplanted into a mouse model showed increased proliferation relative to untreated cells (Yan et al., 2011). However, the VEGF-C–treated ADSCs increased the frequency of angiogenic cells (5×) and lymphangiogenic cells (3×), demonstrating that native VEGF-C does not specifically induce lymphangiogenesis (Yan et al., 2011).

VEGF-C156s is a mutant form of VEGF-C that binds VEGFR3 but not VEGFR2. Therefore, it may be feasible to induce the selective differentiation of adipose stem cells into lymphatic endothelial cells (Achen et al., 2006; Joukov et al., 2006). This is because VEGFR-3 controls signaling that mediates lymphangiogenesis. This receptor is mainly expressed in the tissues of normal adults and lymphatic endothelial cells of embryos (Bhansali, 2012).

This study had two purposes. First, we sought to confirm the characteristics and multipotency of human (h) ADSCs in vitro. Second, we developed a protocol to differentiate hADSCs into lymphatic endothelial-like cells using a mutant VEGF-C (VEGF-C156s) that binds VEGFR3 but not VEGFR2. This binding specificity may allow for selective differentiation of hADSCs into lymphatic endothelial tissue, instead of producing both angiogenic and lymphangiogenic lineages (Yan et al., 2011). We analyzed in vitro differentiation potential using standard protocols for BM-MSC adipogenesis and osteogenesis. The results of this study demonstrated that hADSCs could be easily obtained from adipose tissue. These cells possessed multipotency and could be differentiated into lymphatic endothelial-like cells of mesenchymal origin, suggesting that this stem cell source might be suitable for tissue engineering of lymphatic vessels in vitro and reconstruction of the lymphatic duct after lymphedema in vivo. Thus, hADSCs hold the promise for the cure of limb lymphedema and lymphatic tube-related diseases.

Reagents and Methods

Collection of human adipose tissue

Adipose tissue was collected from patients using a trocar needle during standard liposuction procedures. All samples were obtained with written consent from the donors according to the instructions from the local institution review board at the first affiliated hospital, Sun Yat-sen University of Guangzhou.

Isolation and purification of hADSCs

Cells were isolated from adipose tissue by trypsin digestion. First, the tissue samples were washed extensively with phosphate-buffered saline (PBS) containing 5% penicillin/streptomycin (pen/strep). Upon removal of debris, the samples were digested in a sterile tissue culture plate with collagenase prepared in PBS with 1% pen/strep for tissue digestion. The samples were then minced with scalpels and pipetted up and down to mix. The samples were then incubated for 30 min at an incubator at 37°C and 5% CO₂. To neutralize the collagenase activity, 5 mL of Dulbecco's Modified Eagle Medium (DMEM) containing 20% heat-inactivated fetal bovine serum (FBS; Thermo Fisher Scientific, Waltham, MA, USA) were added to the tissue sample. The samples were then filtered through a 40-μm filter into a 50-mL centrifuge tube spun at 1500 rpm for 5 min. The cells from both the wash fraction and the digested fraction were resuspended in complete medium. A total of 1×10⁵ cells were cultured in a 25-cm² culture flask. After 3–4 days, the unattached cells were depleted by replacing the medium. The medium was subsequently changed twice a week.

Flow cytometry analysis

Rapidly growing cells were selected for flow cytometry using the following antibodies: CD13, CD29, CD44, CD105, CD31, CD34, CD45, and HLA-DR conjugated with fluorescein isothiocyanate (FITC) or phycoerythrin (PE) (R&D Company). Isotypic control analysis was performed in parallel. Flow cytometry was performed using a FACSCalibur Cytometer (BD Biosciences, San Jose, California) and was analyzed using Cellquest software.

Immunofluorescence

After 10 days of induction, hADSCs were fixed with 4% (vol/vol) paraformaldehyde for 10 min and permeabilized for 30 min in PBS containing 0.1% (vol/vol) Triton X-100, goat serum, and 1% (wt/vol) bovine serum albumin (BSA) (Sigma, St. Louis, MO, USA) at room temperature before exposure to goat anti-human LYVE-1 (1:100; R&D, Minneapolis, MN, USA) or goat anti-human VEGFR-3 (1:100; R&D, Minneapolis, MN, USA) overnight at 4°C. After washing with PBS three times, AlexaFluor 488–conjugated rabbit anti-goat antibody (Invitrogen, Grand Island, NY, USA) was added to the samples and then incubated at 37°C for 40 min. After being washed with PBS three times, cell nuclei were counterstained with 4′,6-diamidino-2-phenylindole (DAPI, 1:1000; Sigma, St. Louis, MO, USA), and the positively-stained cells were captured with an IPP 4.5 system (Pixera, Santa Clara, CA, USA) under a fluorescence microscope (OLYMPUS-600). hADSCs incubated in low-glucose minimal essential medium (LG-DMEM) were used as a negative control.

Adipogenic differentiation

Near-confluent (80–90%) cells were washed with PBS containing 1% streptomycin. Adipogenic differentiation medium (HG-DMEM, supplemented with 0.5 mM isobutylmethylxanthine, 0.2 mM indomethacin, 1 μM dexamethasone, and 0.01 mg/mL insulin). This medium was changed every 3 days until the adipocytes reached maturation. After 12–14 days of differentiation, the cells were fixed using a 10% formalin solution. The accumulation of neutral lipids was detected by staining the cells with a solution of 0.5% Oil Red O at room temperature for at least 60 min.

Osteogenic differentiation

Osteogenesis was induced using culture medium supplemented with 100 nM dexamethasone, 10 mM glycerol phosphate, and 0.02 mM ascorbate-2-phosphate that was replaced every 2 or 3 days for 14 days. Mineralization was assessed by staining the cells with 40 mM Alizarin Red (pH 4.1) after fixation in 10% formalin.

Lymphatic-like endothelial cells differentiation

After the conventional digestion for the third-generation cells, the cells were inoculated onto a 12-well plate coated with 0.5% gelatin. When they reached 60% confluence, LG-DMEM containing 1% green – streptomycin was added to the control group, while induction medium [LG-DMEM medium with 5%FBS, 50 ng/mL VEGF-C156s, 10 ng/mL basic fibroblast growth factor (bFGF), and 1% green – streptomycin] was added in the experimental group. The fluid was changed every 3 days for 10 days, and then the cells were fixed with 4% paraformaldehyde, permeabilized with 0.2% Triton X-100, and blocked with PBS containing 5% rabbit serum. Rabbit anti-sheep LYVE-1 and VEGF receptor-3 (VEGFR-3) antibodies (1:50) were incubated overnight. After that, the cells were incubated with fluorescein isothiocyanate (FITC)-conjugated anti-sheep secondary antibodies (1:300). DAPI was used for nuclear staining, and photos were taken using fluorescence microscopy.

Real-time PCR

Total RNA was isolated using TRIzol reagent (Life Technologies, Gaithersburg, MD, USA) according to the manufacturer's instructions. After digestion with DNase I (Fermentas, Hanover, MD, USA), 1 μg of total RNA was reverse transcribed using a RevertAid First Strand Complementary DNA Synthesis Kit (Fermentas, Hanover, MD, USA). The mRNA levels of the indicated genes were analyzed in triplicate using 2×SYBR Green master mixture (TOYOBO, Osaka, Japan), and reactions were carried out in an ABI 7500 Real-Time PCR instrument (Applied Biosystems) under the manufacturer's instructions. The mRNA levels were normalized to GAPDH (internal control), and gene expression was presented as -fold changes (ΔΔCt method). The primer sequences used in the PCR are listed as follows:

LYVE-1 (forward primer), 5′-AATTTCACAGAAGCTAAGGAGGC-3′,

LYVE-1 (reverse primer), 5′-TCAAGGCTGTTTCAACTTGGTC-3′,

PROX-1 (forward primer), 5′-CTGGGAAATTATGGTTGCTCCT-3′,

PROX-1 (reverse primer), 5′-AAAGTCAAATGTACTCCGCAAGC-3′,

FLT-4 (forward primer), 5′-TGCACGAGGTACATGCCAAC-3′,

FLT-4 (reverse primer), 5′-GCTGCTCAAAGTCTCTCACGAA-3′,

GAPDH (forward primer), 5′-GGATTTGGTCGTATTGGG-3′,

GAPDH (reverse primer), 5′-GGAAGATGGTGATGGGATT-3′.

Immunoblotting

Cells were washed with pre-chilled PBS and directly lysed in 1×Laemmli buffer. Lysate was sonicated and then centrifuged at 16,000×g for 10 min at 4°C. Supernatant was recovered as total cell lysate. Equal amount of proteins (20 μg) were separated by 8–10% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and then electrotransferred to 0.45-μm polyvinylidene difluoride membranes (Millipore, Bedford, MA, USA). After that, membranes were blocked with a solution of Tris-buffered saline and 0.1% Tween 20 (TBST) containing 5% nonfat milk for 1 h at room temperature and then incubated with appropriate primary antibodies overnight at 4°C. Specifically bound primary antibodies were detected with peroxidase-coupled secondary antibody and enhanced chemiluminescence (Cell Signaling Technologies, Beverly, MA).

Statistical analysis

Data are presented as mean±standard error of the mean (SEM), and the Student's t-test was used for testing statistical significance.

Results

Isolation of hADSCs

An important source of adult stem cells has long been the bone marrow (Gimble and Guilak, 2003). Although cells isolated from this compartment are useful for further experimentation, this compartment presents two problems. First, it is technically challenging as well as traumatic to the patient to remove bone marrow. Second, the amount of bone marrow in the body is very limited (Gimble and Guilak, 2003). Therefore, we wished to determine the ability to isolate stem cells from the adipose tissue compartment, which is a readily accessible and abundant tissue store in the body. hADSCs, obtained by separation and purification of primary adipose tissue, were resuspended in a 25-cm² culture flask at the density of 1×10⁵ cells/mL. After 8–10 h, a small number of cells began to adhere the surface of the flask. Eventually, the adherent cell number increased, reaching confluence with a fibroblast-like morphology (Fig. 1A). These data suggest that significant numbers of purified hADSCs can be obtained from adipose tissue.

FIG. 1.

Characterization of hADSCs. (A) Morphology of purified, confluent hADSCs at 10×magnification (left panel) and 20×magnification (right panel). hADSCs showed a fibroblast-like morphology in culture. (B) Representative graphs showing fluorescence-activated cell sorting analysis of cell-surface antigen expression. (C) Oil Red O staining after 15 days of stimulation with adipogenic differentiation medium. (D) Alizarin S Red staining of hADSCs cultured after 14 days of stimulation with osteogenic differentiation medium.

Characterization and differentiation of hADSCs

To ensure that our hADSCs were comparable with stem cells isolated from other tissue compartments, we evaluated the expression of cell-surface antigens on the isolated hADSCs by flow cytometry of at least three different samples. Flow cytometry results showed cells were strongly positive for the MSC markers CD13 (99±1.0%), CD29 (98.2±1.7%), CD44 (97.9±2.0%), and CD105 (98.5±1.5%). The cells were negative for the hematopoietic markers CD31 (0.811±0.03%), CD34 (0.28±0.01%), CD45 (0.1±0.011%), and HLA-DR (0.05±0.004%) (Fig. 1B). This pattern of expression was similar to previously characterized BM-MSCs, indicating that these two populations play similar roles in the body (Mundra et al., 2013; Tsai et al., 2014).

Next, we wished to assess the multipotency of the isolated hADSCs by promoting differentiation pathways like adipogenesis and osteogenesis. To induce adipogenesis, nearly confluent hADSCs were incubated with induction medium (HD-MEM supplemented with 0.5 mM isobutylmethylxanthine, 0.2 mM indomethacin, 1 μM dexamethasone, and 0.01 mg/mL insulin) for 12–14 days before staining with Oil Red O to visualize lipid droplets. Many cells showed large, rounded lipoid bodies in the cytoplasm, indicating successful induction of adipogenesis (Fig. 1C).

Osteogenesis was induced by incubating hADSCs in osteogenic induction medium (HD-MEM supplemented with 100 nM dexamethasone, 10 mM glycerol phosphate, and 0.02 mM ascorbate-2-phosphate) for 14 days before staining with Alizarin Red S to visualize calcium deposits. Many cells demonstrated obvious calcium deposition, indicating successful osteogenic induction (Fig. 1D). Taken together, these data indicate that the isolated and cultured hADSCs maintain multipotency, and standard induction protocols for adult stem cells are useful for inducing hADSCs.

Identification of lymphatic endothelial-like cells

Because the hADSCs showed multipotency, we wished to assess whether they could be induced to selectively differentiate into lymphatic endothelial-like cells. After 10 days of induction in a medium containing 5% FBS, 50 ng/mL VEGF-C156s, 10 ng/mL bFGF, and 1% green – streptomycin, the morphology of hADSCs gradually converted from a long and spindly shape to an oval shape, similar to an endothelial cell (Fig. 2). Immunofluorescence detection of the lymphatic-specific markers LYVE-1 and VEGFR3 identified these cells as lymphatic endothelial-like cells (Fig. 3), compared to the negative control group. Additionally, reverse transcriptase quantitative polymerase chain reaction (RT-qPCR) and immunoblot results showed these cells expressed the specific molecular signature of lymphatic endothelial cells (Figs. 4 and 5).

FIG. 2.

Morphological change of hADSCs after induction with VEGF-C156s. The morphology of hADSCs after induction with VEGF-C156s was photographed from day 0 to day 10.

FIG. 3.

Immunofluorescent staining of hADSCs induced by VEGF-C156S for 10 days. LYVE-1 (A) and VEGFR3 (B) are shown in green. Cell nuclei were stained with DAPI.

FIG. 4.

Expression of lymphatic endothelial cell-specific proteins LYVE-1, PROX-1, and FLT-4 in induced cells assessed by RT-qPCR. Values are means±SEM (n=3). (*) p<0.05; (**) p<0.01 for Student's t-test.

FIG. 5.

Expression of lymphatic endothelial cell-specific proteins LYVE-1, PROX-1, and FLT-4 in induced cells. The expression of LYVE-1, PROX-1, FLT-4, and glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was determined by western blotting. The densities of LYVE-1, PROX-1, and FLT-4 were quantified and normalized to the expression levels of GAPDH.

Discussion

In recent years, tissue engineering and materials science have enabled the manufacture of blood vessels (L'Heureux et al., 2006). Isolation, identification, and induction of seed cells have always been important for tissue engineering. In this study, we investigated the ability of hADSCs to differentiate into lymphatic vascular endothelium. ADSCs are a class of pluripotent MSCs derived from the mesoderm and ectoderm during early embryogenesis (Gimble et al., 2007).

Compared to BM-MSCs, which have previously been used as tissue engineering seed cells, ADSCs can be obtained without a bone marrow biopsy or other invasive surgery and with higher yield. Studies have shown that the ADSCs are pluripotent and can differentiate into mesodermal tissue-derived cell types, such as adipocytes, cartilage cells, bone cells, muscle cells, and endothelial cells (Gaiba et al., 2012; Huang et al., 2004; Katz et al., 1999; Sen et al., 2001; Urbich and Dimmeler, 2004). They have also been successfully differentiated into nerve and myocardial tissues (Planat-Bénard et al., 2004; Schäffler and Büchler, 2007), revealing very important potential in future clinical applications.

Many studies have demonstrated that MSCs can be induced into vascular endothelial-like cell types. However, few studies have been published showing the differentiation of MSCs into lymphatic endothelial-like cells, and selective differentiation has remained a technical challenge (Yan et al., 2011). To address this question, we modified the vascular endothelial-like cell induction protocol. VEGF-C156S, a growth factor mutant that promotes lymphangiogenesis, was used instead of VEGF-C, which promotes both angiogenesis and lymphangiogenesis (Mǎkinen et al., 2001; Veikkola et al., 2001).

Studies have shown that adipocytes and endothelial cells share a common progenitor (Planat-Benard et al., 2004), and VEGF-C plays a key role in the transformation of MSCs into lymphatic endothelium. VEGF-C156S maintains binding capacity to VEGFR3 while losing recognition of VEGFR2 (Joukov et al., 1998), conferring selective differentiation potential into lymphatic endothelial cells (Mǎkinen et al., 2001; Veikkola et al., 2001). Using endothelial cell medium and VEGF-C156S, MSCs can be induced to express endothelial markers like podoplanin and VEGFR3, with a corresponding gene expression signature (Conrad et al., 2009; Wei et al., 2012). In our experiment, VEGF-C156S was used as the main induction factor to induce hADSCs into lymphatic endothelial-like cells in vitro, which was verified by testing for the expression of LYVE-1, a specific lymphatic marker (Banerji et al., 1999). Thus, human adipose tissue represents a promising source of seed cells for lymphatic tissue reconstruction in vivo.

Prospero homeobox 1 (PROX-1) is a homeobox transcription factor related to lymphatic endothelium growth and extension. Homozygous knockout PROX-1 in mice leads to a complete loss of the lymphatic system; however, the PROX-1 heterozygote knockout developed chylous ascites and died within 2–3 days after birth. PROX-1 is closely related to the growth, extension of embryonic lymphatic bud, and the phenotypic changes of lymphatic endothelial cells. Although PROX-1 can be expressed in a variety of nonendothelial cells (e.g., lens, heart, liver, pancreas, and the nervous system), in endothelial cells it is specifically expressed only in the embryonic lymphangion and the normal tissue of adults, as well as tumor lymphangion. PROX-1 may regulate the differentiation or migration of lymphatic endothelial cells, and it plays an important role in the development of embryonic lymphangion. Infants lacking PROX-1 often die from defects in lymphatic sprouting and differentiation. The differential expression of PROX-1 in the original vein endothelial cells is related to differentiation direction; the original vein that expresses PROX-1 differentiates to the lymphangion, whereas the one does not express PROX-1 differentiates to blood vessel. Wigle et al. used gene knockouts to observe the role of PROX-1 in lymphatic development. When the the PROX-1 gene was excluded, the vein endothelial cells were unable to differentiate into the lymphatic endothelial cells as directed. During embryogenesis, the process of budding from vein endothelial cells to lymphangion was obstructed. The endothelial cells did not express VEGFR-3, LYVE-1, or other markers of lymphatic endothelial cells, but it seems that blood vessel formation was not affected. These data suggest that PROX-1 is the basic regulatory factors of lymphangiogenesis.

Reconstruction of the lymphatic vessels is the ideal therapy for lymphedema; however, no protocol for in vitro reconstruction of lymphatic tissue has yet been reported. One study has reported the use of hADSC within a hydrogel doped with VEGF-C in a mouse model of injury-induced lymphedema (Hwang et al., 2011). This hydrogel reduced the depth of dermal edema significantly compared to controls, and the hADSCs were found to express LYVE-1, suggesting that these cells induced lymphangiogenesis in vivo (Hwang et al., 2011).

Our results were promising in that we showed hADSCs could differentiate into lymphatic endothelial-like cells; however, this study was limited in the following ways. First, we only used three lymphatic markers to characterize the lymphatic endothelial cells. Other lymphatic-specific markers, such as secondary lymphoid chemokine (SLC) and podoplanin, could have been included to strengthen the evidence that hADSCs had been successfully induced to become lymphatic endothelial-like cells. Second, we did not address the role of pro- and anti-lymphangiogenic cytokines, such as interleukin-8 (IL-8) and transforming growth factor-β1 (TGF-β1, in the lymphatic endothelial cells derived from hADSCs. Additional studies are required to confirm and extend these results.

In conclusion, we have demonstrated for the first time selective differentiation of hADSCs into lymphatic endothelial-like cells in vitro. These findings may help in the development of tissue reconstruction techniques by providing a useful source of seed cells, which may help lead to an effective treatment for lymphedema.

Footnotes

Author Disclosure Statement

The authors declare that no conflicting financial interests exist.

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