Forskolin Enhances Urokinase Plasminogen Activator Secretion and Angiogenic Activity of Xeno‐Free Cultures of Human Adipose Tissue‐Derived Stem Cells

Abstract

Adipose tissue‐derived stem cells (ASCs) hold significant potential for treating various clinical conditions. To enhance their regenerative properties, ASCs can be chemically stimulated using various in vitro protocols. However, unsatisfactory results persist, partly due to the relatively costly long‐term methods. Furthermore, current culturing techniques often rely on the use of xenogenic fetal bovine serum that can be immunogenic, limiting clinical translations. To facilitate clinical translation of ASCs‐derived therapeutics, the effect of different stimulation protocols on human ASCs cultured in a xeno‐free medium (PRIME‐XV MSC Expansion XSFM) is investigated. The xeno‐free medium was supplemented with stimulants (forskolin (FSK), basic fibroblast growth factor, platelet‐derived growth factor‐AA, neuregulin‐1) in combinations or individually. Stimulation for 72 h in FSK alone, or together with the growth factors, enhanced the production of urokinase plasminogen activator (uPA), a serine protease involved in tissue remodeling processes. Conditioned medium derived from stimulated ASCs enhanced in vitro angiogenesis and endothelial cells migration. This study shows that pro‐angiogenic responses in human ASCs can be enhanced with a defined short stimulation protocol using a xeno‐free medium. The protocol, using readily available manufacturing cell therapy grade molecules, may boost the regenerative properties of ASCs secretome which could enhance their efficacy in clinical treatments.

The regenerative potential of adipose tissue‐derived stem cells can be enhanced through chemical stimulation in vitro. A short stimulation protocol using forskolin, either alone or in combination with other growth factors, under xeno‐free conditions enhanced the pro‐angiogenic responses in human ASCs. This in vitro technique could strengthen the regenerative properties of the ASCs secretome, potentially improving their efficacy in clinical treatments The graphical abstract was created with BioRender.com.

1. Introduction

Mesenchymal stem cells (MSCs) constitute a category of cells with remarkable therapeutic potential, found within fat, bone marrow, dental, and other tissues.^{[ 1 ]} Among these various stem cell types, adipose tissue‐derived stem cells (ASCs) stand out for their easy accessibility through liposuction, coupled with rapid in vitro expansion capacity.^{[ 2} , ³ , ^{4 ]} The primary characteristic of ASCs lies in their ability to differentiate into diverse cell lineages, including osteoblasts, adipocytes, and chondrocytes.^{[ 1} , ⁵ , ^{6 ]} However, it is important to note that the versatility of ASCs extends beyond their differentiation capacity. ASCs are also recognized for their rich secretome, which is composed of a range of paracrine bioactive molecules, growth factors, and extracellular vesicles.^{[ 7} , ⁸ , ⁹ , ^{10 ]}

The ASCs secretome has attracted a lot of attention in the field of regenerative medicine. Considered a cell‐free product, ASCs‐derived factors may be a safer therapeutic approach able to overcome potential risks associated with cell‐based therapies such as uncontrolled cell growth and immune rejection. ASCs secretome is rich in bioactive molecules that can mediate tissue regeneration and angiogenesis.^{[ 11} , ¹² , ^{13 ]} One of the factors released by ASCs and involved in these processes is urokinase plasminogen activator (uPA), a serine protease that converts plasminogen into plasmin.^{[ 14} , ^{15 ]} Plasmin activates a proteolytic cascade that ultimately leads to extracellular matrix degradation. The breakdown of the extracellular matrix is fundamental for tissue regeneration and repair because it allows cells to migrate toward the damaged tissue.^{[ 16} , ^{17 ]} Due to the importance of uPA in cancer development and vascular diseases, numerous studies have been conducted about its role in these conditions.^{[ 18} , ¹⁹ , ^{20 ]} However, little attention has been paid to the production of the urokinase and the regenerative signaling pathways in ASCs.

In prior research studies, we developed an in vitro stimulation protocol for MSCs.^{[ 21} , ²² , ^{23 ]} This method involved treating rat or human cells with a specific combination of chemical stimulants and growth factors for a period of up to 2 weeks. The outcome of the method is a significant amplification of the growth factor secretome of the ASCs. Furthermore, we and others have shown that this approach enhances the therapeutic potential of ASCs secretome for possible treatment of a wide range of clinical conditions.^{[ 24} , ²⁵ , ²⁶ , ^{27 ]} However, culturing ASCs in vitro typically involves the utilization of serum from a xenogeneic source such as fetal bovine serum (FBS) carrying inherent risks, including potential contamination with bacteria and viruses, as well as the recurring challenge of batch‐to‐batch variation.^{[ 28} , ^{29 ]} For this reason, it is imperative to develop well‐defined, safer, and shorter protocols for future clinical use. ASCs cultured in serum‐ or xeno‐free medium is a promising approach to address these concerns and ensure the safety and consistency of ASC‐based therapies.^{[ 30} , ^{31 ]} There are many commercially available xeno‐free expansion media formulations. In this study, we used PRIME‐XV MSC Expansion XSFM (FujiFilm Irvine Scientific) which has been shown to enhance ASCs expansion while maintaining their stem cell characteristics.^{[ 32 ]} We stimulated human ASCs under these xeno‐free conditions and investigated signaling pathways and the biological activity of the secretome.

2. Results

2.1. Forskolin Stimulation Enhances IGF‐I and PLAU Gene Expression and Promotes uPA Protein Production and Secretion in ASCs

We first examined the effect of individual chemicals (FSK) and growth factors (GFs) and thereafter combinations of these on a range of regenerative genes (Table S1, Supporting Information), many of which we have previously studied in ASCs cultured in the presence of FBS.^{[ 33 ]} Of these, the most pronounced changes were in IGF‐I and PLAU gene expression. Both genes showed an increased gene expression in ASCs treated with FSK alone and in combination with GFs. However, these increases were not statistically significant (Figure 1A). We investigated if the gene changes were associated with increased secretion of the proteins. We were unable to detect significant quantities of IGF‐1 in the cell‐conditioned medium (data not shown). However, in both the group exposed to FSK alone and the group with GFs with FSK, we observed statistically significant increases in secreted uPA protein when compared with unstimulated ASCs and ASCs treated with GFs only (Figure 1B). FSK alone and in combination with GFs also enhanced the intracellular levels of uPA indicating that the stimulation process did not simply affect the secretion of the existing pool of uPA (Figure 1B). These results indicated that FSK addition alone was sufficient to stimulate the stem cells to release and produce increased amounts of uPA.

Figure 1.

FSK stimulation enhances IGF‐I and PLAU gene expression and promotes uPA protein production and secretion in ASCs in xeno‐free conditions. (A) RT‐qPCR showing the expression of IGF‐I and PLAU gene in unstimulated ASCs (ASCs^{FSK−/GFs−}) and stimulated in the presence of single GFs (ASCs^FGF+ or ASCs^PDGF+ or ASCs^NRG−1+) or FSK alone (ASCs^FSK+) or combined with GFs (ASCs^FSK+/FGF+ or ASCs^{FSK+/FGF+/ PDGF+} or all GFs, ASCs^FSK+/GFs+). RPL13a was used as a reference gene. Each symbol represents an individual donor. The data are shown as mean ± standard deviation of relative expression (n = 6). (B) Secreted and intracellular uPA protein in unstimulated ASCs (ASCs^{FSK–/GFs−}), treated with FSK alone (ASCs^FSK+/GFs−) or GFs alone (ASCs^FSK–/GFs+) or in combination (ASCs^FSK+/GFs+). Each symbol represents an individual donor. The data are shown as mean ± standard deviation (n = 5). Statistical significance indicated as ns (non‐significant) or * p < 0.05.

2.2. ERK1/2 Signaling Pathways in Stimulated ASCs

To investigate the pathways responsible for the stimulation of ASCs, we performed a protein analysis using a Proteome Profiler Human Phospho‐Kinase Array Kit. This assay enabled us to gain insights into the various pathways that were activated when ASCs were exposed to FSK alone or in combination with different GFs. The most prominent change was the upregulation of the ERK 1/2 signaling pathway in the ASCs stimulated with FSK (Figure S1, Supporting Information). This result was subsequently confirmed through western blot analysis for phosphorylated‐p44/42 MAPK (ERK 1/2), providing evidence that the ERK 1/2 pathway was activated in ASCs when stimulated with FSK. However, the western blot also showed that the ERK1/2 signaling pathway was triggered by stimulation with all the GFs in the absence FSK (Figure 2A; Figure S2, Supporting Information). Since we showed that FSK stimulation of ASCs consistently induced upregulation of the PLAU gene and secretion of uPA, and we also identified activation of the ERK 1/2 pathway, we wanted to explore potential correlations between these findings. The addition of PD98059, an ERK inhibitor, had no effect on FSK‐stimulated PLAU gene expression but markedly downregulated uPA protein levels. The differences in uPA protein levels were however not statistically significant in either ASCs^{FSK+/GFs–/PD+} (p = 0.2907) or ASCs^{FSK+/GFs+/PD+} (p = 0.5255) groups compared with their respective controls (Figure 2C).

Figure 2.

ERK1/2 signaling pathways in stimulated ASCs. (A) Western blot analysis of p‐p44/42 MAPK (ERK1/2) in 1h treated ASCs. Beta‐tubulin was used as loading control protein. Pooled patients’ lysate was used (n = 6). (B,C) PLAU gene expression and uPA protein secretion in unstimulated and stimulated pooled ASCs treated with PD98059 (PD; ERK1/2 inhibitor). The experiment was repeated three times with pooled ASCs. The data are shown as mean ± standard deviation of relative gene expression (n = 6) or protein levels (pg/mL/10⁶ cells). Statistical significance is indicated as ns (non‐significant) between the corresponding groups. Each symbol represents the same pooled sample across the different conditions. Full‐length blots are presented in Figure S2 (Supporting Information).

2.3. Conditioned Medium from Stimulated ASCs Enhances Angiogenesis

An in vitro angiogenesis assay was used to assess the biological activity of the conditioned medium from ASCs. Human umbilical vein endothelial cells (HUVEC's) exposed to angiogenic stimulating factors form tube capillary‐like structures when seeded on nutrient‐rich extracellular matrix gel. Conditioned medium from ASCs exposed to GFs with FSK induced the formation of significantly more closed mature networks in the HUVECs when compared with their respective control media XSFM (Figure 3A,B). A similar trend was observed in HUVECs exposed to FSK stimulated ASCs‐conditioned medium, although the difference between the conditioned medium and XSFM with addition of FSK was not statistically significantly different (Figure 3A,B).

Figure 3.

Conditioned medium from stimulated ASCs enhances angiogenesis. (A) Representative images of HUVECs on ECMatrix treated with XSFM and CM from one donors unstimulated ASCs (CM^{FSK–/GFs−}), treated with FSK alone (CM^FSK+/GFs−) or GFs alone (CM^FSK–/GFs+) or in combination (CM^FSK+/GFs+). (B) Number of closed networks counted in HUVECs on ECMatrix treated with conditioned medium from unstimulated and stimulated ASCs (n = 5) and corresponding XSFM (n = 3). For the CM groups each symbol represents an individual donor. Data shown are mean ± standard deviation (n = 5) and statistical significance is indicated as ns (non‐significant) or ** p < 0.01.

2.4. Conditioned Medium from FSK‐Stimulated ASCs Promotes HUVECs Migration and Supports Human Dermal Fibroblasts Migration

In vitro migration of HUVECs and human dermal fibroblasts was used to assess the ability of ASCs‐conditioned medium to induce cell migration. HUVECs treated with conditioned medium from unstimulated and stimulated ASCs showed better in vitro wound closure capacities when compared with their respective unstimulated and stimulated control medium XSFM (Figure 4A,B). When comparing treatments with ASCs‐conditioned media to those with unstimulated conditioned medium, only the HUVECs treated with conditioned medium from ASCs exposed to FSK and GFs showed a statistically significant difference. However, a similar trend was observed in FSK‐treated ASCs conditioned medium (Figure 4B). Migration of human dermal fibroblasts was also assessed. Both unstimulated and stimulated control medium XSFM and ASCs‐conditioned media induced ≈30% gap closure (Figure S3A,B, Supporting Information).

Figure 4.

Conditioned medium from ASCs exposed to FSK induces HUVEC migration (A) Representative images after 8 h of HUVECs with CM from one donor unstimulated ASCs (CM^{FSK–/GFs−}), treated with FSK alone (CM^FSK+/GFs−) or GFs alone (CM^FSK–/GFs+) or in combination (CM^FSK+/GFs+) in the absence or presence of the uPA inhibitor BC‐11 hydrobromide. (B) In vitro wound closure percentage of HUVECs treated with conditioned medium from unstimulated and stimulated ASCs (n = 5) and corresponding XSFM (n = 3) in the presence and absence of BC‐11 (BC), a uPA inhibitor. For CM groups each symbol represents an individual donor. Data shown are mean ± standard deviation (n = 3 and 5) and statistical significance indicated as ns (non‐significant), *p < 0.05, ***p < 0.001, and ****p < 0.0001.

Due to the pronounced effect of conditioned medium from unstimulated and stimulated ASCs on HUVEC cells migration, we investigated whether uPA directly influenced endothelial cell migration by using BC‐11, a uPA inhibitor. The addition of BC‐11 did not affect cell viability (Figure S4, Supporting Information). Migration was significantly reduced by 60–80% when BC‐11 was added to the ASCs‐conditioned medium (Figure 4B).

3. Discussion

In this study, we investigated the effects of a short‐term stimulation protocol on adipose tissue‐derived stem cells under xeno‐free conditions. Our results showed a significant increase in the production and secretion of uPA in response to FSK treatment, coupled with the activation of the ERK1/2 signaling pathway. Angiogenesis and migration were assessed, and it was observed that the conditioned medium from FSK/GFs‐treated ASCs had a stronger effect on promoting both closed network formation and in vitro wound closure in HUVEC cells compared with unstimulated ASCs conditioned medium. Moreover, we have not only demonstrated that uPA was produced and secreted by FSK‐treated ASCs, but we also showed that it was biologically active in ASCs conditioned medium and played an important role in inducing HUVEC cell migration.

The potential benefits of the ASC secretome have been well discussed by previous researchers.^{[ 34} , ³⁵ , ^{36 ]} Numerous studies have shown that conditioned medium from ASCs holds a great potential for treating various conditions.^{[ 37} , ^{38 ]} However, despite researchers' hopes to uncover the therapeutic properties of ASCs secretome and harness them for the development of cell‐free regenerative therapies, limitations in the potency of ASCs paracrine factors persist. For this reason, scientists have been focusing on developing stimulation protocols for ASCs to enhance their proliferation and differentiation potential, as well as promote their secretion of various growth factors and cytokines.^{[ 24} , ²⁵ , ³⁹ , ⁴⁰ , ^{41 ]} However, current stimulation methodologies mainly rely on long‐term approaches, which can be both costly and inefficient for future clinical applications. We aimed to explore alternative approaches to the more standard protocols for ASCs stimulation.^{[ 24} , ⁴² , ^{43 ]} Initially, we examined the effect of FSK or single growth factors and a combination of GFs with FSK on ASCs in XSFM for 72 h. A novel finding is that PLAU and IGF‐I gene expression were upregulated in ASCs treated with only FSK in XSFM. Other studies have demonstrated that uPA is produced by MSCs but the signaling pathways are not well elucidated.^{[ 44} , ^{45 ]} Furthermore, research has been predominantly conducted in serum‐containing media.^{[ 24} , ⁴² , ^{46 ]} When we analyzed the conditioned medium from ASCs, we found that FSK‐treated ASCs produce and secrete higher levels of uPA. However, despite the upregulation of IGF‐I gene expression, we did not detect IGF‐I protein in the conditioned medium, possibly due to interference from IGF‐binding proteins.^{[ 47 ]} Our findings suggest that 72 h FSK treatment in xeno‐free conditions can effectively stimulate adipose stem cells to produce and secrete uPA which has important applications in various physiological processes, such as tissue repair and angiogenesis.

To investigate the signaling pathways involved with our stimulation protocol, we performed a Phospho‐Kinase array analysis. We observed that the ERK1/2 pathway was activated in FSK‐treated ASCs, which is consistent with previous studies.^{[ 48} , ^{49 ]} Forskolin is a potent activator of adenylate cyclase, leading to increased intracellular levels of cAMP.^{[ 50} , ^{51 ]} Subsequently, cAMP activates protein kinase A, which in turn phosphorylates and activates ERK1/2.^{[ 52} , ^{53 ]} Our findings in western blot analysis of the phosphorylated forms of ERK1/2 demonstrate that both FSK treatment and GFs alone treatment can activate the ERK1/2 signaling pathway in ASCs, albeit with different intensity. Pharmacological inhibition of phosphorylated ERK1/2 using specific inhibitors such as PD98059 allowed us to further investigate the role of ERK1/2 in the secretion of urokinase plasminogen activator.^{[ 54 ]} The FSK/FSK+GFs‐induced elevation of PLAU gene expression was not affected by the presence of PD98059 whereas uPA protein levels were reduced by ≈50%. This indicates that the stimulatory effect of FSK on uPA production is not entirely dependent on ERK1/2.

For demonstrating the effectiveness of our short‐term stimulation protocol and its potential impact on biological activities, we conducted an in vitro angiogenesis assay using HUVECs.^{[ 55 ]} Conditioned medium from ASCs can be used as a cell‐free based therapy to stimulate angiogenesis.^{[ 33} , ⁵⁶ , ^{57 ]} Under our experimental conditions, we observed that conditioned medium from FSK and GFs‐treated ASCs promoted angiogenesis to a greater extent compared with the unstimulated ASCs conditioned medium. Capillary morphogenesis is influenced by uPA, as it degrades the extracellular matrix and promotes endothelial cell migration.^{[ 58} , ⁵⁹ , ^{60 ]} We hypothesized that the migration properties of our conditioned medium could be due to the presence of uPA. We tested HUVECs migration capacities with ASCs conditioned medium in the presence or absence of FSK and GFs. In line with previous studies, ASCs conditioned medium induced in vitro wound closure, but we showed that our stimulation protocol of FSK+GFs could enhance this further.^{[ 61} , ^{62 ]} Under our experimental conditions, this effect was specific to HUVECs migration as migration rates were similar for dermal fibroblasts in the presence of unstimulated or stimulated cells‐conditioned medium. This could be attributed to the human dermal fibroblasts having a higher baseline migration in XSFM, as the medium is designed for human mesenchymal stem cells, which share characteristics with fibroblasts. In contrast, HUVEC migration was significantly lower in XSFM, with less than 20% migration observed after 8 h. To further validate the role of uPA in this process, we chemically inhibited its activity using BC‐11. This drastically reduced the migration effects of ASCs conditioned medium on HUVECs, providing evidence that uPA produced and secreted by the stem cells is indeed biologically active. This is an important finding in the understanding of the ASCs secretome which highlights its remarkable potential in the field of regenerative medicine.

4. Conclusion

We have demonstrated that short‐term FSK stimulation of adipose stem cells in xeno‐free conditions promotes the production of uPA. Stimulation with FSK alone or in combination with GFs enhances angiogenic activity in the ASC secretome and promotes increased endothelial cell migration which is in part mediated by uPA. The rather modest enhancement of biological activity with the addition of GFs might be attributed to the medium already containing high levels of the GFs; however interestingly, of these, we could only detect bFGF in the medium (Table S2, Supporting Information) and this was at a much lower concentration than we used in the stimulation protocol. In conclusion, the protocol uses readily available manufacturing cell therapy grade molecules, which can be used to boost the regenerative properties of the ASCs secretome and thereby potentially enhance their efficacy in clinical treatments.

5. Experimental Section

Isolation of Adipose Tissue‐Derived Stem Cells (ASCs)

Human adipose tissue samples were obtained from the abdominal fat of six patients undergoing elective surgery (mean age: 46, mean body mass index: 27.6 kg m⁻²). In this study the procedures were performed according to the principles of the Declaration of Helsinki and were approved by the Regional Ethics Committee in Umeå (Section for Medical Research) under the application “Evaluation of mesenchymal stem cells isolated from different organs with the aim of increasing the efficiency and reducing the morbidity of cellular therapies” (2013‐276‐31M, decisioned 4th September 2013). The patients provided written informed consent for participation in the study and the use of samples. To isolate the ASCs, lipoaspirates were digested for 1 h at 37 °C with collagenase NB4 (Serva/Nordmark) dissolved in saline solution at a concentration of 0.3U per mL of aspirate. Samples were then centrifuged at 300 × g for 5 min, the undigested fat tissue and collagenase solution was aspirated, and the pellet was resuspended in phosphate‐buffered saline. Red blood cells were lysed with ACK (Ammonium‐Chloride‐Potassium) Lysing Buffer (ThermoFisher Scientific), samples re‐centrifuged and the pellet resuspended in PRIME‐XV MSC Expansion XSFM (FujiFilm Irvine Scientific) together with 1% (v/v) penicillin/streptomycin (Invitrogen Life Technologies). The cells were cultured on Corning CellBIND flasks in PRIME‐XV MSC Expansion XSFM. Cells were seeded at a minimum starting density of 10 000 cells cm⁻² and subsequently incubated at 37 °C with 5% CO₂. The non‐adherent cells were removed after 24 h by washing with saline solution, followed by replenishment with fresh medium. This process was repeated over the next 2 days. On reaching ≈90% confluence the adherent cells were passaged by using a solution of TrypLE Express (Invitrogen). Cells were cryopreserved in Synth‐a‐Freeze (Thermo Fisher Scientific) cryopreservation Medium (minimum 1 × 10⁶ cells mL⁻¹) and stored in liquid nitrogen.

Flow Cytometry

The expression of mesenchymal stem cell‐associated surface markers (CD73, CD90, CD105) and the putative pericyte marker CD146 was determined on passage 1‐2 ASCs grown in PRIME‐XV MSC Expansion XSFM as previously described.^{[ 63 ]} Data were acquired using a BD Accuri C6 flow cytometer (BD Bioscience) and analyses were performed on a minimum of 10 000 cells (Figure S5A, Supporting Information).

Differentiation

To confirm tri‐lineage differentiation (Figure S5B, Supporting Information), passage 1 cells were thawed and plated at a density of 55 000 cells per well into either 24‐well CellBIND plates (for adipogenesis) or Corning plates coated with type I collagen and vitronectin (for osteogenesis). Cultures were treated with respective differentiation medium and stained with Oil Red O (adipocytes) or Alizarin Red (osteogenesis) as previously described.^{[ 63 ]} For chondrogenesis, 3D aggregate cultures were used. ASCs (250 000) were resuspended in ChondroMAX Differentiation Media (Merck Millipore), centrifuged at 200 × g for 5 min and the pellet resuspended with 0.5 mL ChondroMAX Differentiation Media. The cells were then re‐centrifuged and thereafter with the medium remaining, the cap of the tube was loosened to allow gas exchange, and the sample was incubated upright in an incubator at 37 °C with 5% CO₂. Once the cell pellet had formed a round ball the medium was changed every other day. After 21 days the micromass cultures were fixed, sectioned (12 µm) and stained with 1% Toluidine Blue.

Stimulation of ASCs

Passage 2–5 cells were quickly thawed at 37 °C and then seeded at 4000 cells cm⁻² in Corning CellBIND flasks in PRIME‐XV MSC Expansion XSFM together with 1% (v/v) penicillin/streptomycin and left overnight at 37 °C. To stimulate the cells, 10 µm forskolin (FSK; Sigma–Aldrich), 10 ng mL⁻¹ basic fibroblast growth factor (bFGF; Invitrogen/Fisher Scientific), 200 ng mL⁻¹ neuregulin1‐beta1 (NRG‐1; R&D Systems) and 5 ng mL⁻¹ platelet‐derived growth factor‐AA (PDGF‐AA; Invitrogen/Fisher Scientific) were added into the media as a combination or as single factors. Unstimulated stem cells were cultured in parallel in PRIME‐XV MSC Expansion XSFM only. After 72 h, the cell‐conditioned media from both unstimulated and stimulated cells and control media XSFM was collected, centrifuged at 300 × g for 5 min and then filtered with a 0.8 µm Millex‐AA Filter Unit (Millipore). The conditioned medium and XSFM were aliquoted and stored at −80 °C until needed for all assays. The cells were treated with TrypLE Express Enzyme, centrifuged and the cell pellet resuspended in either RIPA Lysis Buffer (Thermo Scientific) for protein analysis or RLT buffer (Qiagen) for preparation of RNA. In another series of experiments, ASCs from the 6 donors were pooled and seeded at 4000 cells cm⁻² in Corning CellBIND flasks in PRIME‐XV MSC Expansion XSFM and left overnight at 37 °C. The ERK1/2 inhibitor PD98059 (R&D Systems) was added at a final concentration of 10 µm simultaneously with the cell‐stimulating factors or to unstimulated ASCs. After 3 days, the cell pellets and conditioned medium were collected for further investigation. The experiment was repeated 3 times.

Quantitative Reverse Transcription Polymerase Chain Reaction (RT‐qPCR)

Total RNA isolation from unstimulated and stimulated cells was performed with RNeasy Mini Kit (Qiagen) according to the manufacturer's instructions. An iScript cDNA synthesis kit (Bio‐Rad) was used to convert 10 ng µL⁻¹ reaction mix into complementary DNA (cDNA). RT‐qPCR was performed using Myra Liquid Handling System (Bio Molecular Systems) and Mic qPCR cycler (Bio Molecular Systems) with SsoFast EvaGreen supermix (Bio‐Rad) and 1 ng of cDNA. All samples were analysed in quadruplicates. The reference gene used to normalize the expression levels was ribosomal protein L13a (RPL13a) gene. Primers were manufactured by Sigma, sequences and variable annealing/extension temperature are displayed in Table 1 . ΔΔC(t) principle was applied to calculate the relative expression data.

Table 1.

Primer sequences for RT‐qPCR and annealing temperatures used (°C).

Factor	Forward primer 5´‐3´	Reverse primer 5´‐3´	Temperature
BDNF	AGAGGCTTGACATCATCATTGGCTG	CAAAGGCACTTGACTACTGAGCATC	65 °C
GDNF	CACCAGATAAACAAATGGCAGTGC	CGACAGGTCATCATTCAAAGGCG	65 °C
IGF‐I	TGTGGAGACAGGGGCTTTTA	ATCCACGATGCCTGTCTGA	61.5 °C
NGF	ATACAGGCGGAACCACACTCAG	GTCCACAGTAATGTTGCGGGTC	64 °C
PLAU	GCTTGTCCAAGAGTGCATGGT	CAGGGTCGGTTCTCGATGG	64 °C
RPL13a	AAGTACCAGGCAGTGACAG	CCTGTTTCCGTAGCCTCATG	58 °C
VEGF‐A	ATCTGCATGGTGATGTTGGA	GGGCAGAATCATCACGAAG	61.4 °C

Enzyme‐Linked Immunosorbent Assay (ELISA)

Secreted urokinase plasminogen activator (uPA) protein was quantified in thawed conditioned medium collected from both unstimulated and stimulated stem cells (n = 5 donors), using the Human PLAU / Urokinase‐type Plasminogen Activator ELISA Kit (Sigma–Aldrich). Samples were analyzed in duplicates. Intracellular uPA was measured from corresponding cell lysates using the Human uPA Quantikine ELISA Kit (R&D Systems), also in duplicate.

To assess background protein levels in the medium, XSFM was thawed and tested for the presence of bFGF using the Human FGF basic Quantikine ELISA kit (R&D Systems), PDGF‐AA using the Human PDGFAA ELISA Kit (RayBiotech), NRG‐1 using the Human NRG1‐beta 1 ELISA Kit (RayBiotech), and uPA using the same uPA kit as above. These control samples were also analyzed in duplicates.

All absorbance measurements were performed at 450 nm using a BioTek Synergy (Aglient Technologies) microplate reader.

Human Phospho‐Kinase Antibody Array

Total protein was isolated from unstimulated and stimulated ASCs after 1h treatment using a buffer from the human phospho‐kinase antibody array (R&D Systems) according to the manufacturer's protocol. Total protein was quantified using a BioRad Dc Protein Assay (Bio‐Rad). Pooled protein lysate from the six donors was used for the array. The detection of the relative levels of phosphorylation of 37 kinases phosphorylation sites on the membranes was performed according to the manufacturer's protocol and analyzed in an Odyssey Fc Dual‐Mode Imaging System (LI‐COR Biosciences).

Western Blotting

Stimulated and unstimulated ASCs were lysed in RIPA buffer supplemented with protease inhibitors (Sigma–Aldrich) and total protein was determined. Samples were electrophoresed on 12% (v/v) pre‐cast gels at 100 V for 1.5 h, then transferred to 0.2 µm nitrocellulose membranes (Amersham Protran) at 40 V for 1 h. The membranes were blocked with 5% (w/v) bovine serum albumin (BSA; Sigma–Aldrich) in Tris‐buffered saline with 0.1% (v/v) Tween 20 detergent (TBST) for 1 h at RT. Membranes were then stained with antibody against p‐ERK1/2 (1:2000) (Cell Signaling; #4370) overnight at 4 °C and washed with TBST 3 times for 10 min. An HRP‐conjugated secondary antibody (1:2000) (Cell Signaling; #7074) was added for 1 h at RT. Finally, Enhanced Chemiluminescence solution (GE Healthcare; #RPN2232) was added to the membranes for 1 min and chemiluminescence analyzed in an Odyssey Fc Dual‐Mode Imaging System.

In Vitro Angiogenesis Assay

Passage 2 human umbilical vein endothelial cells (HUVECs; Invitrogen) were thawed and expanded in human large vessel endothelial cell basal medium supplemented (Gibco) with low serum growth supplement (LSGS; Gibco) together with 1% (v/v) penicillin/streptomycin (Gibco) in Corning CellBIND flasks. At 85% confluency, HUVECs were passaged using Trypsin‐EDTA (0.05%; Gibco) and seeded at a density of 2.5 × 10⁴ cells in 200 µL suspensions of cell‐conditioned unstimulated and stimulated media or unstimulated and stimulated XSFM media onto 175 µL ECMatrix (Sigma–Aldrich) plated in a 48‐well plate (VWR International). Individual donor cells‐conditioned medium (n = 5) was applied in triplicates to individual wells. After 4 h at 37 °C, the capillary‐like tube formation was evaluated using light microscopy (Olympus IX71 microscope). Images were captured from four randomly selected fields from each well using a ColorViewII digital camera (Soft Imaging System). A manual count of closed networks was performed on the randomized photographs. The data are presented as the mean values derived from four images for each individual patient.

In Vitro Migration Assays—HUVECs

Passage 2 HUVECs were thawed and expanded in human large vessel endothelial cell basal medium (Gibco) supplemented with low serum growth supplement (LSGS; Gibco) together with 1% (v/v) penicillin/streptomycin in Corning CellBIND flasks. Culture‐inserts 2 well (Ibidi) were positioned on a CellBIND 24‐well plate. At 85% confluency, HUVECs were passed using Trypsin‐EDTA (0.05%) and seeded at a density of 2.7 × 10⁴ cells in 100 µL suspensions of human large vessel endothelial cell basal medium supplemented with LSGS and 1% (v/v) penicillin/streptomycin in each well of the culture‐inserts and left at 37 °C for 48 h. The inserts were removed, and the cells were then washed once with DPBS. Unstimulated or stimulated cell‐conditioned media or unstimulated or stimulated XSFM media were thawed and added to each well. The same procedure was performed in the presence of 50 µm BC‐11 hydrobromide (Tocris Bioscience), a specific uPA inhibitor. Individual donor cell‐conditioned medium (n = 5) was applied in individual wells. Photographs were taken from each well using a ColorViewII digital camera at 0h (T0) and 8h (T8). Surface area measurements at T0 and T8 were analyzed using ImageJ software, and in vitro wound closure percentage was calculated as a measure of migration. To test potential cytotoxicity of the BC‐11 in further experiments the inserts were removed, and the HUVECs were washed once with DPBS. XSFM media was thawed and added to each well (n = 3) in the presence of 50 µm BC‐11 hydrobromide. After 8h, the cells were washed twice with DPBS, and a LIVE/DEAD cell imaging assay was performed according to manufacturer's instruction (ThermoFisher Scientific). Bright field and fluorescence photographs were taken from each well using a ColorViewII digital camera at 0 h (T0) and 8 h (T8).

In Vitro Migration Assays—Human Dermal Fibroblasts

Passage 3 human dermal fibroblasts (adult skin, single donor; Gibco) were thawed and expanded in Dulbecco's Modified Eagle Medium (DMEM; Gibco) containing 10% (v/v) fetal bovine serum (FBS; Sigma Aldrich) together with 1% (v/v) penicillin/streptomycin in tissue culture flasks (Nunc). Ibidi two‐well culture‐inserts were positioned in a 24‐well plate (VWR International). At 80% confluency, fibroblasts were passed using Trypsin‐EDTA (0.05%) and seeded at a density of 1 × 10⁴ cells in 80 µL suspensions of DMEM containing 10% (v/v) FSB with 1% (v/v) penicillin/streptomycin in each well of the culture‐inserts and left at 37 °C overnight. After removing the inserts, cells were washed once with DPBS. Unstimulated or stimulated XSFM media or unstimulated or stimulated cell‐conditioned media were thawed and added to each well. Individual donor cell‐conditioned medium (n = 5, n = 3 for XSFM control media) was applied in individual wells. Three random regions of interest (ROIs) within the gap were imaged at 0 (T0) h and 15 (T15) h using a ColorViewII digital camera. ImageJ software was used to measure the surface area at T0 and T15, and in vitro wound closure percentage was calculated.

Statistical Analysis

No pre‐processing of data was made. Data for RT‐qPCR, ELISA, angiogenesis, and migration assays were calculated as mean ± SD with a sample size of n = 5 or 6 independent donor samples. For experiments investigating the effect of PD98059 pooled samples were made and the experiments repeated three times. T test, or for multiple comparisons, one‐way analysis of variance (ANOVA) with Bonferroni post hoc test was used. Statistical significance was determined as *p < 0.05, **p < 0.01, ***p < 0.001 and ****p < 0.0001. GraphPad Prism software (GraphPad Software Inc.) was used to perform the statistical analyses.

Ethics Approval and Consent to Participate

In this study the procedures were performed according to the principles of the Declaration of Helsinki and were approved by the Regional Ethics Committee in Umeå (Section for Medical Research) under the application “Evaluation of mesenchymal stem cells isolated from different organs with the aim of increasing the efficiency and reducing the morbidity of cellular therapies” (2013‐276‐31M, decisioned 4th September 2013). The patients provided written informed consent for participation in the study and the use of samples.

Conflict of Interest

The authors declare no conflict of interest.

Author Contributions

M.V.G. designed and performed experiments, analyzed the data, and wrote the manuscript. A.T.L. performed experiments. R.W. contributed funding for the project. M.B. performed experiments and contributed funding for the project. G.A. analyzed the data and contributed funding for the project. P.J.K. designed experiments, analyzed the data and wrote the manuscript, and contributed funding for the project. All authors reviewed and agreed on the final draft of the manuscript.

Supporting information

Supporting Information

Giraudo M. V., Lauvrud A. T., Wiberg R., Brohlin M., Andersson G., and Kingham P. J., “Forskolin Enhances Urokinase Plasminogen Activator Secretion and Angiogenic Activity of Xeno‐Free Cultures of Human Adipose Tissue‐Derived Stem Cells.” Adv. Biology 9, no. 10 (2025): e00466. 10.1002/adbi.202400466

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.