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. 2020 May 26;17(3):335–350. doi: 10.1007/s13770-020-00265-5

Hypoxia Pretreatment Promotes Chondrocyte Differentiation of Human Adipose-Derived Stem Cells via Vascular Endothelial Growth Factor

Ok Kyung Hwang 1,2,#, Young Woock Noh 1,#, Jin Tae Hong 2,✉, Je-Wook Lee 1,✉
PMCID: PMC7260353  PMID: 32451775

Abstract

BACKGROUND:

Human adipose tissue-derived stem cells (ADSCs) are attractive multipotent stem cell sources with therapeutic potential in various fields requiring repair and regeneration, such as acute and chronically damaged tissues. ADSC is suitable for cell-based therapy, but its use has been hampered due to poor survival after administration. Potential therapeutic use of ADSC requires mass production of cells through in vitro expansion. Many studies have consistently observed the tendency of senescence by mesenchymal stem cell (MSC) proliferation upon expansion. Hypoxia has been reported to improve stem cell proliferation and survival.

METHODS:

We investigated the effects of hypoxia pretreatment on ADCS proliferation, migration capacity, differentiation potential and cytokine production. We also analyzed the effects of vascular endothelial growth factor (VEGF) on osteogenic and chondrogenic differentiation of ADSCs by hypoxia pretreatment.

RESULTS:

Hypoxia pretreatment increased the proliferation of ADSCs by increasing VEGF levels. Interestingly, hypoxia pretreatment significantly increased chondrogenic differentiation but decreased osteogenic differentiation compared to normoxia. The osteogenic differentiation of ADSC was decreased by the addition of VEGF but increased by the depletion of VEGF. We have shown that hypoxia pretreatment increases the chondrogenic differentiation of ADSCs while reducing osteogenic differentiation in a VEGF-dependent manner.

CONCLUSION:

These results show that hypoxia pretreatment can provide useful information for studies that require selective inhibition of osteogenic differentiation, such as cartilage regeneration.

Electronic supplementary material

The online version of this article (10.1007/s13770-020-00265-5) contains supplementary material, which is available to authorized users.

Keywords: Adipose-derived stem cells (ADSCs), Hypoxia, VEGF, Differentiation, Cartilage regeneration

Introduction

Stem cells are distinguished from other cell types by two important characteristics: (1) they are unspecialized cells that are capable of self-renewal through proliferation (2) and can be induced to become tissue-specific cells under certain physiologic or experimental conditions [1]. Stem cells with these unique properties have great potential for tissue regenerative medicine [1].

Mesenchymal stem cells (MSCs) are pluripotent adult stem cells that can be isolated from various mesenchymal tissues in the adult body. MSC was first described by Friedenstein et al. as bone marrow-derived clonogenic: plastic-adherent cells capable of differentiating into osteoblasts, adipocytes and chondrocytes [2]. However, ideal stem cells for use in functional tissue engineering should be abundantly available, harvested with minimal morbidity, reliably differentiated in a variety of pathways, and safe and effective for transplantation. Recently, some reports have described the isolation of MSCs from human adipose tissue that appears to meet these criteria [3, 4]. These human adipose tissue-derived stem cells (ADSCs) are capable of forming mesodermal tissues such as bone, cartilage, tendons, skeletal muscle and fat when cultured under lineage-specific conditions, and can also form non-mesodermal tissues such as neurons and endocrine pancreatic cells [5, 6].

Recent studies revealed that low oxygen tension or hypoxia affects various types of stem cells, such as embryonic stem cells [7], induced pluripotent stem cells [8], and bone marrow-derived stem cells (BMSCs) [9–11]. A low oxygen environment is physiologically normal for most mammalian embryos as well as adult somatic stem cells [11]. In mammalian cells, the transcriptional response to oxygen deficiency is primarily mediated by hypoxia-inducible factor 1 (HIF-1), which gradually increases with decreasing oxygen concentration [12]. The expression of genes such as vascular endothelial growth factor (VEGF) and erythropoietin is induced to stimulate angiogenesis and hematopoiesis [12, 13]. In hypoxic conditions, ADSC proliferation is increased compared to normal oxygen conditions [10, 14]. Under hypoxic conditions, the secretion of VEGF and fibroblast growth factor (FGF)-2 proteins by ADSCs is increased [15].

Previous studies have shown that hypoxia promotes osteogenesis in human mesenchymal stromal cells [11, 16, 17]. However, another study reported that hypoxia inhibits bone formation in human mesenchymal stem cells [18]. Therefore, the effect of hypoxia on MSC differentiation remains ambiguous. We hypothesized that oxygen participates in the intricate balance between cellular proliferation and commitment toward differentiation. In this study, we investigated the effects of hypoxia pretreatment on ADCS proliferation, migration capacity, differentiation potential and cytokine production. We also analyzed the effects of VEGF on osteogenic differentiation of ADSC by hypoxia pretreatment and chondrogenic differentiation.

Materials and methods

Cell culture and in vitro differentiation

STEMPRO® human adipose-derived stem cells (hADSCs) purchased from Thermo Fisher Scientific (Carlsbad, CA, USA) were cultured in 75 cm2 flasks using a MesenPRO RS™ Medium Kit (Thermo Fisher Scientific) and l-glutamine (Thermo Fisher Scientific) according to the manufacturer’s instructions. Hypoxic culture experiments were performed in a multi-gas incubator (InvivO2; Baker Ruskinn, Sanford, ME, USA) at 37 °C in an atmosphere containing 5% CO2 balanced with nitrogen to reach an oxygen concentration of 1%. Normal oxygen culture experiments were performed in a standard incubator under conditions containing 5% CO2 and 20% O2. The ADSCs were differentiated by guidelines provided for inducing STEMPRO® ADSCs to differentiate towards osteoblasts, adipocytes, and chondrocytes using the StemPro Osteogenesis Differentiation Kit (Thermo Fisher Scientific), StemPro Adipogenesis Differentiation Kit (Thermo Fisher Scientific) and StemPro Chondrogenesis Differentiation Kit (Thermo Fisher Scientific), respectively, according to the manufacturer’s protocol. Alizarin Red (EMD Millipore, Billerica, MA, USA) staining was used to assess osteogenic differentiation, Oil Red O (Biovision, Mountain View, CA) staining to verify adipogenic differentiation, and Safranin-O (Sigma-Aldrich, St. Louis, MO, USA) staining to evaluate the ability of hADSCs to differentiate toward chondrocytes. To examine the direct effect of VEGF on osteogenic differentiation, we induced osteogenic differentiation by adding 50 ng/mL of VEGF (PeproTech, Rocky Hill, NJ, USA) and 50 µg/ml of anti-VEGF antibody (Avastin; Roche, Basel, Switzerland).

Proliferation assay

Cell proliferation under normoxic or hypoxic conditions was assessed using a Cell Counting Kit-8 (CCK-8, Dojindo Molecular Technologies, Kumamoto, Japan). Briefly, ADSCs were seeded in 96-well plates at a density of 2.5 × 103 to 1 × 104 per well and incubated under normoxic (20% O2) or hypoxic (1% O2) conditions for 24 or 48 h. The cellular proliferation was measured by absorbance at 450 nm using VersaMax (Molecular Devices, San Jose, CA, USA) apparatus (n = 3) after adding 1/10 volume of CCK-8 reagent and incubating for 2 h.

Western blotting

Nuclear protein extracts were obtained using the NE-PER Nuclear and Cytoplasmic Extraction Kit (Thermo Fisher Scientific) for the detection of HIF-1α and histone H3. The protein concentrations were measured using a Bradford Protein Assay Kit (Bio-Rad Laboratories, Richmond, CA, USA). Ten micrograms of protein extracts were separated by SDS-PAGE using a NuPAGE electrophoresis system (Thermo Fisher Scientific). The proteins were transferred to polyvinylidene difluoride membranes using the iBlot Dry Blotting System (Thermo Fisher Scientific) in accordance with the manufacturer’s protocol. The membranes were blocked with 5% skim milk in Tris-buffered saline with 0.05% Tween 20 for 1 h and incubated with primary antibody overnight: anti-HIF-1α (Cell Signaling Technology, Beverly, MA, USA). Next, the membranes were incubated with the corresponding horseradish peroxidase-conjugated IgG secondary antibody (Cell Signaling Technology) for 1 h at room temperature. The proteins were detected using ECL reagent (GE Life Sciences, Issaquah, WA, USA) and the images were obtained using a ChemiDoc MP (Bio-Rad Laboratories) controlled by Image Lab software (Bio-Rad Laboratories).

RNA isolation and quantitative polymerase chain reaction

Total RNA was isolated from the cells using an RNeasy Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer’s protocol. The RNA concentrations were assessed using a Nanodrop spectrophotometer (ND-1000, Thermo Fisher Scientific). cDNA was synthesized from 1 μg of RNA using a Reverse Transcription Master Premix 1st strand cDNA synthesis kit (ELPIS, Daejeon, Korea). Quantitative polymerase chain reaction (qPCR) was performed using Power SYBR-Green Master Mix (Applied-Biosystems, Foster City, CA, USA) with triplicate reactions containing 1 × SYBR-Green Master Mix, 500 nM of each forward/reverse primer, and 15 ng of template DNA. The reactions were performed on an Applied Biosystems 7500 Fast Real-Time PCR System for 40 cycles. The fold-change in target gene expression was calculated using the 2−ΔΔCt method and normalized to the control housekeeping gene GAPDH. The primer sequences are shown in Table 1.

Table 1.

List of primers and sequences used for quantitative polymerase chain reactions

Target Forward (5′–3′) Reverse (5′–3′)
Oct4 ACATCAAAGCTCTGCAGAAAGAACT CTGAATACCTTCCCAAATAGAACCC
Nanog CCAACATCCTGAACCTCAGC GCTATTCTTCGGCCAGTTG
Sox2 TGCGAGCGCTGCACAT GCAGCGTGTACTTATCCTTCTTCA
C-Myc GGCCCCCAAGGTAGTTATCCTT CGTTTCCGCAACAAGTCCTCT
P16INK4A GAAGGTCCCTCAGACATCCCC CCCTGTAGGACCTTCGGTGAC
Runx2 TGAGAGCCGCTTCTCCAACC GCGGAAGCATTCTGGAAGGA
ALP ACCATTCCCACGTCTTCACA CAGGCCCATTGCCATACA
Collagen1 CACCAATCACCTGCGTACAG GCAGTTCTTGGTCTCGTCAC
OPG GGCAACACAGCTCACAAGAA CTGGGTTTGCATGCCTTTAT
Osteocalcin AGCAAAGGTGCAGCCTTTGT CTGAAAGCCGATGTGGTCAG
Sialoprotein ACCAGTACCAACAGCACAGA TGGCTCCAGTGACACTTTCT
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Migration assay

ADSCs (5 × 105) were plated in 10 cm cell culture dishes (Corning, Corning NY, USA) and incubated for 48 h in 1% or 20% O2. Then, the ADSCs (1 × 103) were plated in 96-well plates and single cells were tracked using Operetta High‐Content imaging system (PerkinElmer, Waltham, MA, USA). We analyzed the movement of single cells over 12 h using Harmony software.

ELISA for secreted VEGF

ADSCs (4 × 105) were plated in 10 cm cell culture dishes and incubated for 48 h in 1% or 20% O2. The culture supernatants were collected and VEGF levels were analyzed using a Human VEGF Quantikine ELISA Kit (R&D Systems, Minneapolis, MN, USA) according to the manufacturer’s instructions. The VEGF concentrations were quantified with a VersaMax apparatus at OD450 (n = 3).

Flow cytometry

Single-cell suspensions were obtained after pretreatment in hypoxic and normoxic conditions. The cells were washed twice with FACS buffer containing 1% fetal bovine serum in phosphate-buffered saline. The cells were counted and suspended at a concentration of 5 × 105 cells in 100 µl FACS buffer and incubated for 30 min at 4 °C in the dark with the following fluorescent antibodies: CD73-PE, CD90-FITC, CD105-APC, and CD45-PEcy7 (BD Biosciences, San Diego, CA, USA). The cells were washed twice with FACS buffer and then analyzed using flow cytometry (FACS CantoII; BD). The data were acquired, exported as FCS 3.0 files, and analyzed using FlowJo software (Tree Star, Inc., Ashland, OR, USA).

Gene expression profiling

Genomic expression analysis was performed using the Human Mesenchymal Stem Cell RT2 Profiler PCR Array (PAHS-082Z, Qiagen). cDNA was synthesized from 800 ng of total RNA using the RT2 First Strand Kit (Qiagen). PCR arrays were performed in a CFX96 Touch™ Real-Time PCR Detection System (Qiagen) with RT2 SYBR® Green qPCR Master Mix (Qiagen). Each array consisted of a panel of 96 primer sets of 84 mesenchymal stem cell or cellular senescence genes, five housekeeping genes, and seven quality controls. The data were analyzed using web-based RT2 Profiler PCR Array Data Analysis v3.5 software (Qiagen). The fold-change in target gene expression was calculated using the 2−ΔΔCt method and normalized to the geometric mean of five housekeeping genes (ACTB, B2M, GAPDH, HPRT1, and RPLP0) according to manufacturer’s guide.

Cytokine profiling

Culture supernatants were obtained after the cells were incubated for 48 h in 1% or 20% O2. The cytokines secreted by ADSCs were measured using a Proteome Profiler Human XL Cytokine Array Kit (R&D systems) with 102 capture antibodies that were spotted in duplicate on a nitrocellulose membrane according to the manufacturer’s protocol. Then the immunospots were captured and processed using a ChemiDoc MP (Bio-Rad Laboratories) controlled by Image Lab software (Bio-Rad Laboratories). The intensities of the identified spots were quantified using NIH Image J software.

Transfection

VEGF siRNA (Santa Cruz) was transfected using HiperFect transfection reagent (Qiagen). Shortly before transfection, 2 × 105 cells were seeded per 6-well plate. siRNA (150 ng) was diluted in 100 μl of culture medium without serum. HiperFect Transfection Reagent (12 μl) was added to the diluted siRNA and mixed by vortexing. The samples were incubated for 5–10 min at room temperature to allow the formation of transfection complexes. The complexes were added dropwise to the cells and incubated for 24 h at normoxic conditions. Then, the medium was changed to osteogenic differentiation medium.

Statistical analysis

Student’s t test and one-way ANOVA were used for statistical analyses and significance was indicated by p < 0.05. All values are expressed as the means ± standard deviations. GraphPad Prism (GraphPad software) and Excel 2016 (Microsoft Corporation) were used for all the statistical analyses.

Results

Proliferation of ADSCs after hypoxia treatment

We first examined the effect of hypoxia on ADSC proliferation. As shown in Fig. 1A, the number of ADSCs at 1% O2 was higher than that at 20% O2 measured using the Cell Counting Kit-8. Compared to normoxia, the ADSCs cultured at 1% O2 showed significantly high proliferation at 24 and 48 h (Fig. 1A).

Fig. 1.

Fig. 1

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Effects of hypoxia pretreatment on the proliferation of ADSCs. A ADSC proliferation was significantly increased at 24 and 48 h by hypoxia compared to normoxia and measured using the CCK-8 kit. B Nuclear proteins were prepared from ADSCs exposed to hypoxia for 6 h and Western blotting was performed. The expression of HIF-1α was increased by hypoxia. The results are shown as the mean ± SD values. The statistical significance of hypoxia versus normoxia was determined by Student’s t-test (*p < 0.05 and **p < 0.01). N: normoxia (20% O2), H: hypoxia (1% O2)

HIF-1α expression in ADSCs after hypoxia treatment

We measured the amount of HIF-1α protein to demonstrate that the cells were actually exposed to hypoxia. HIF-1α protein was increased in cells incubated in hypoxic conditions for 6 h (Fig. 1B).

Expression of stem cell and senescence markers after hypoxia treatment

We analyzed the effects of hypoxia on the expression of stem cell markers such as Oct4, Nanog, Sox2 and c-Myc, and senescence marker p16INK4A in ADSCs. ADSCs have been reported to express several markers of pluripotent embryonic stem (ES) cells [19, 20]. We examined the expression of pluripotent marker Oct4, an important transcription factor in maintaining the undifferentiated status of ES cells. The expression of Oct4 was significantly decreased by hypoxia compared to normoxia. Similar patterns were observed in the expression of Nanog and Sox2 (Fig. 2A). In hypoxic conditions, the expression of c-Myc, one of stem cell markers, was significantly higher than in normoxic conditions (Fig. 2A). p16INK4A is known to contribute to cell and stem cell senescence and accumulates in many aged cells and tissues. Thus, p16INK4A is considered a biomarker of aging [21, 22]. The expression of senescence marker p16INK4a in hypoxia was significantly lower than in normoxia. The expression of mesenchymal surface markers in ADSCs under normoxic and hypoxic conditions was examined by FACS analysis (Fig. 2B). The expression of CD73, CD90, or CD105 in hypoxic conditions was similar to that in normoxic conditions (Fig. 2B).

Fig. 2.

Fig. 2

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Effect of hypoxia pretreatment on the properties of ADSCs. A The mRNA expression of the indicated gene was analyzed using real-time RT-PCR. The expression of stem cell markers Oct4, Nanog, and Sox2 in hypoxia was significantly lower than in normoxia. The expression of stem cell marker c-Myc in hypoxia was significantly higher than in normoxia. The expression of senescence marker p16INK4a in hypoxia was significantly lower than in normoxia. B In immunophenotype analysis, the expression of mesenchymal surface markers CD73, CD90, and CD105 was not significantly changed by hypoxia compared to normoxia. C The migration ability of ADSCs was increased in hypoxia compared to normoxia. D The current step and accumulated distance of ADSCs cultured in hypoxia were increased compared to normoxia at 12 h. The results are shown as the mean ± SD values. The statistical significance of hypoxia versus normoxia was determined by Student’s t-test (*p < 0.05 and **p < 0.01)

Increase in ADSC migration ability after hypoxia treatment

To evaluate the migration ability of ADSCs in hypoxic conditions, we incubated ADSC for 48 h in normal oxygen and hypoxic conditions and then plated single cells into 96-well plates to track the migration pathway for 12 h. The results showed that hypoxia increased the mobility of ADSCs by increasing the speed and cumulative distance (Fig. 2C, D).

Reduction in osteogenic differentiation capability of ADSCs after hypoxia treatment

ADSCs are multipotent stem cells that can differentiate into mesenchymal and non-mesenchymal lineages [23]. To investigate the effect of hypoxic conditions on the differentiation of ADSCs, we evaluated the differentiation into adipocytes, osteoblasts and chondrocytes using ADSCs cultured under normal oxygen and hypoxic conditions. As determined by oil red O staining, the differentiation of ADSCs into adipocytes was not significantly different in hypoxia compared to normal oxygen conditions. As measured by Safranin O staining, the differentiation of ADSCs into chondrocytes was significantly increased in hypoxia compared to normoxia. As determined by Alizarin Red staining, the differentiation of ADSCs into osteoblasts was reduced in hypoxic conditions compared to normal oxygen conditions (Fig. 3).

Fig. 3.

Fig. 3

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Effects of hypoxia pretreatment on the differentiation potential of ADSCs. ADSCs were cultured at 4.5 × 105 cells/100 mm dish in hypoxia or normoxia for 48 h and induced to differentiate along adipogenic, osteogenic, and chondrogenic lines for 21 days. A Hypoxia inhibited the osteogenic differentiation efficiency compared to normoxia as shown by Alizarin Red staining. B No significant difference was observed in adipogenesis between normoxia and hypoxia by Oil Red O staining. C Hypoxia increased the chondrogenic differentiation efficiency compared to normoxia as shown by Safranin O staining

To determine the role of hypoxic culture conditions on the osteoblastic differentiation of ADSCs, Alizarin Red staining was performed at 7, 14, and 21 days after differentiation. We reconfirmed the reduction in the differentiation of ADSCs into osteoblasts after hypoxia treatment (Fig. 4A). The positively stained Alizarin Red area was decreased in hypoxia compared to cells cultured under normoxic conditions. In particular, the osteogenic differentiation capacity of ADSCs cultured in hypoxic conditions was observed to be significantly decreased on day 14 after differentiation (Fig. 4A).

Fig. 4.

Fig. 4

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Hypoxia reduced the mineralization capacity and the expression of osteogenesis-related genes in ADSCs. A Bone mineralization was reduced by hypoxia compared to normoxia as shown by Alizarin Red staining at 7, 14, and 21 days after the induction of osteogenesis. B The mRNA expression of the indicated genes was analyzed using real-time RT-PCR. The results are shown as the mean ± SD values. The statistical significance of hypoxia versus normoxia was determined by Student’s t-test (*p < 0.05 and **p < 0.01)

To investigate the changes in osteogenesis-related gene expression in osteoblast differentiation [24–29], we examined the expression of Runx2, ALP, collagen1, OPG, osteocalcin, and sialoprotein in osteoblasts differentiated from ADSCs cultured under normal oxygen and hypoxic conditions. The expression levels of Runx2 and sialoprotein were significantly decreased by hypoxia on days 7, 14, and 21 compared to normoxia (Fig. 4B). The expression of ALP was significantly decreased by hypoxia on days 7 and 14 compared to normoxia (Fig. 4B). The expression levels of collagen 1, OPG, and osteocalcin mRNA were decreased by hypoxia on day 7 compared to normoxia (Fig. 4B). These results indicated that hypoxic culture conditions could inhibit the osteogenic differentiation potential of ADSC.

Comparative gene expression profiles of ADSCs in normoxic and hypoxic conditions

To understand the molecular basis of regulation of differentiation by hypoxia and to identify the stem cell-related genes that are selectively activated or repressed in response to hypoxia, ADSCs were analyzed using a Human Mesenchymal Stem Cell RT2 Profiler PCR Array. The expression profiles of 84 genes in ADSCs obtained after normoxia and hypoxia exposure are shown in Fig. 5. We identified 12 genes that were differentially expressed between normoxia and hypoxia with a minimal fold-change of 1.8 (Fig. 5, Table 2). Particularly, the expression of VEGFA gene was found to be upregulated in response to hypoxia. In contrast, EGF gene expression was mostly downregulated under hypoxic conditions (4.59-fold) (Fig. 5, Table 2).

Fig. 5.

Fig. 5

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Effect of hypoxia pretreatment on the differential expression of mesenchymal stem cell-related genes. The heat map shows the changes in mesenchymal stem cell-related genes in ADSCs cultured at 4.5 × 105 cells/100 mm dish for 48 h in hypoxia compared to normoxia. Gradient indicates the magnitude of the change in gene expression

Table 2.

Differential gene expression in ADSCs cultured in hypoxic conditions

No. Gene symbol Fold change Ref. seq. number
1 ZFP42 2.5 NM_174900
2 VEGFA 1.87 NM_003376
3 EGF − 4.59 NM_001963
4 GDF5 − 4.1 NM_000557
5 FGF10 − 2.63 NM_004465
6 GDF15 − 2.08 NM_004864
7 KITLG − 1.94 NM_003994
8 TGFB3 − 1.88 NM_003239
9 ERBB2 − 1.85 NM_004448
10 PPARG − 1.84 NM_015869
11 WNT3A − 1.82 NM_033131
12 FUT1 − 1.8 NM_000148
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Profile of cytokines secreted by ADSCs cultured under normoxic and hypoxic conditions

We further analyzed the cytokines secreted by ADSCs cultured under hypoxic culture conditions and compared them to those cultured under normoxic conditions using an antibody array. As shown in Fig. 6, the expression levels of several cytokines and growth factors were altered by hypoxia compared to normoxia (Fig. 6A, B). In particular, the production of VEGF and IGFBP-3 by ADSC was significantly higher in hypoxia compared to normal oxygen conditions (4.08- and 2.74-fold, respectively). Although not a significant increase, hypoxia increased the production of IL-6 by more than 1.2 fold. The amount of VEGF in conditioned medium was also measured by ELISA and the results were consistent with those of the cytokine array analysis (Fig. 6C). In all of the cytokine assays for VEGF, similar expression patterns were found at the mRNA and protein levels (Figs. 5, 6).

Fig. 6.

Fig. 6

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Effect of hypoxia on the secretion of cytokines and VEGF by ADSCs. ADSCs were cultured at 4.5 × 105 cells/100 mm dish for 48 h in hypoxia or normoxia. A Cytokines were measured in cell culture supernatants using the Human XL Cytokine Array Kit according to the manufacturer’s protocol. B The expression of angiogenin, Dickkopf-1, IGFBP-3, IL-6, thrombospondin-1, uPAR, and VEGF by ADSCs in hypoxia was higher than in normoxia. C The secretion of VEGF into the medium by ADSCs in hypoxia was significantly increased compared to normoxia. The results are shown as the mean ± SD values. The statistical significance of hypoxia versus normoxia was determined by Student’s t-test (*p < 0.05 and **p < 0.01)

Effects of VEGF on the osteoblastic differentiation of ADSCs

We examined whether increased VEGF inhibited the osteoblast differentiation by hypoxia. We assessed the direct effect of VEGF by adding VEGF to osteoblast differentiation medium of ADSC under normal oxygen conditions. The positively stained Alizarin Red area indicating osteogenic differentiation was smaller in the VEGF-treated group at days 14 and 21 compared to the control group (Fig. 7). In hypoxia, a decrease in the differentiation of ADSCs into osteoblasts was restored by Avastin (VEGF antibody) treatment (Fig. 7).

Fig. 7.

Fig. 7

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Effect of VEGF and anti-VEGF antibody on the differentiation of ADSCs into osteoblasts. A The differentiation of ADSCs into osteoblasts was induced by the addition of VEGF to the medium. The positively stained Alizarin Red area was smaller in cells treated with VEGF compared to the control at 14 and 21 days. B, C The differentiation of ADSCs into osteoblasts in hypoxia was restored by VEGF antibody. The results are shown as the mean ± SD values. The statistical significance of hypoxia versus normoxia was determined by Student’s t-test (*p < 0.05 and **p < 0.01)

We examined whether the inhibition of VEGF expression by transfected VEGF siRNA increases osteoblast differentiation. The differentiation of ADSC into osteoblasts was induced after transfection with VEGF siRNA. The positively stained Alizarin Red area was increased after transfection with VEGF siRNA at day 14 compared to the control (Fig. 8).

Fig. 8.

Fig. 8

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Effect of VEGF siRNA on the differentiation of ADSCs into osteoblasts. The differentiation of ADSCs to osteoblasts was induced after transfection with VEGF siRNA. A, B The positively stained Alizarin Red area was increased in cells transfected with VEGF siRNA (si-VEGF) compared to control at 14 days. C The mRNA expression of VEGF gene was analyzed using real-time RT-PCR after transfection with VEGF siRNA. The results are shown as the mean ± SD values. The statistical significance of hypoxia versus normoxia was determined by Student’s t-test (*p < 0.05 and **p < 0.01)

Increase in chondrogenic differentiation potential of ADSCs by hypoxia pretreatment

We investigated whether the differentiation of ADSC to chondrocytes was increased by hypoxia pretreatment. In H&E staining, the mass of chondrocytes differentiated from ADSC was increased by hypoxia compared to normoxia (Fig. 9A). In Safranin O staining (Fig. 9B) and immunohistochemical analysis for type II collagen (Fig. 9C), hypoxia pretreatment increased the efficiency of ADSC differentiation into chondrocytes compared to normoxia.

Fig. 9.

Fig. 9

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Effect of hypoxia pretreatment on the chondrogenic differentiation potential of ADSC. A The mass of cells differentiated into chondrocytes was increased by hypoxia compared to normoxia. B, C Hypoxia increased the chondrogenic differentiation efficiency compared to normoxia as shown by Safranin O staining and immunohistochemical analysis for type II collagen

Discussion

MSCs are of interest for therapeutic purposes for tissue repair, but an expansion of cells through in vitro culture is required for use in therapy. In vitro culture conditions, however, change at both cellular and molecular levels, reducing the proliferation and differentiation potential of MSCs [30, 31]. ADSC has recently been isolated from human subcutaneous adipose tissue [5, 32, 33] and is considered an alternative to BMSC in regenerative medicine in that cells are readily available (Fig. 10).

Fig. 10.

Fig. 10

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Schematic diagram of regulation by hypoxia pretreatment in the differentiation of ADSC into osteoblasts and chondrocytes. Hypoxia (1% O2) pretreatment increases VEGF through increased expression of HIF1a, and increased VEGF inhibits osteogenic differentiation but promotes chondrogenic differentiation

Changes in oxygen concentration in culture conditions affect stem cell proliferation and differentiation. The proliferation of human ADSCs has been reported to be increased by hypoxia (2% O2) [11, 15]. Rasmussen et al. reported that 5% of oxygen culture conditions increased the proliferation of ADSC [34]. In this study, it was confirmed that hypoxia (1% O2) increased the proliferation of ADSC compared to normoxia. From these results, it was confirmed that hypoxic culture conditions could be usefully used for regenerative medicine which requires the production of a large number of ADSCs from a small number of donor cells.

HIF-1α improves the oxygen environment by promoting angiogenesis through induction of expression of VEGF, platelet-derived growth factor and basic fibroblast growth factor [35]. Hypoxic conditions have been reported to increase the expression of HIF-1α in ADSC by increasing the expression of VEGF [11, 36]. In this study, we also confirmed that hypoxic conditions increased the expression of HIF-1α and VEGF in ADSC.

In this study, changes in stem cell characteristics of ADSC caused by hypoxia were investigated by expression of common phenotypic markers (CD73, CD90 and CD105) of MSC. Hypoxic conditions did not affect the expression level of stem cell common phenotypic markers. Next, we investigated whether hypoxia can affect the expression of stem cell-related genes in ADSC. Indeed, several previous reports have described that hypoxia (1–2% O2) increases the expression of stem cell-related genes including Oct4, Nanog and Klf4 in various types of stem cells [10, 11]. In this study, as in previous studies by Kwon SY et al. [16], the expression of Nanog, Sox2, and Oct4 in ADSC was significantly decreased in hypoxia compared to normoxia, while c-Myc expression was increased. The reason why the expression pattern of MSC stem cell markers in our results differs from the previous study results is presumed to be due to the difference between the stem cell origin used in the experiment, the concentration of oxygen in culture, and the timing and duration of hypoxia exposure. The difference between this study and the previous studies is thought to be that hypoxia was used only as a pretreatment condition in ADSC culture. Furthermore, it is assumed that the increased expression of c-Myc, a key regulator of cell survival, is associated with an increase in ADSC proliferation in hypoxia [37, 38].

In contrast, the expression of the senescence marker, p16INK4a, was significantly reduced. Therefore, hypoxic conditions could improve the viability of transplanted cells when ADSC is used for clinical treatment.

Hypoxia has been reported to significantly increase the migration of ADSC [10, 39]. In this study, we also found that hypoxia increased the migration of ADSC. Based on previous studies and our findings, it was confirmed that hypoxia influences the movement of ADSC as well as the secretion of various growth factors and cytokines.

Hypoxia is known to affect the expression of genes and proteins in cells. As previously reported [40–42], we found that the expression of VEGF by ADSC in hypoxia was increased at both gene (mRNA) and protein levels. Since VEGF is known to play a pivotal role in angiogenesis and cardiac repair, increased VEGF expression in hypoxia can be clinically important [43, 44]. We also confirmed that hypoxic culture conditions induced down-regulation of the EGF gene and up-regulation of IGFBP-3 protein in ADSC through PCR and cytokine array experiments, respectively. EGF has been reported to significantly increase BMP9-induced osteogenic differentiation of MSCs [45] and IGFBP-3 has been reported to reduce osteogenic differentiation [46]. In this study, we confirmed that hypoxia can inhibit the differentiation of ADSC into osteoblasts by regulating the expression of VEGF, EGF and IGFBP-3.

Hypoxia has been reported to inhibit [18, 47] or promote [10, 16, 17] osteogenesis in mesenchymal stem cells. In this study, we found that hypoxia pretreatment inhibited osteogenic differentiation of ADSC but increased chondrogenic differentiation (Fig. 10). We confirmed for the first time that VEGF increased by hypoxia pretreatment inhibited osteogenic differentiation and that decreased osteoblast differentiation was increased by treatment with Avastin, a VEGF inhibitor. In addition, we confirmed that the differentiation of ADSC into osteoblast was increased by inhibition of VEGF expression using VEGF siRNA. These results indicate that VEGF is a key regulator of osteoblast differentiation of ADSC in hypoxia pretreatment. Malladi et al. reported that severe hypoxic conditions strongly inhibited the osteogenesis and chondrogenesis of ADSC [48], but hypoxia pretreatment increased the chondrogenic differentiation of ADSC. In addition, the use of VEGF has been reported to directly improve the differentiation efficiency from adipose-derived stem cells to chondrocytes [49]. These results showed that the effects of hypoxia on MSC may vary depending on the origin of the cells and the culture conditions, and it is necessary to confirm the characteristics of the cells by the culture conditions in order to use stem cells for therapeutic purposes.

ADSC can be considered as a promising candidate for various therapeutic purposes, such as tissue regeneration because it is relatively easy to acquire and isolate and manipulate in vitro compared to other stem cells. A better understanding of the biological properties and differentiation potential of ADSC will make it possible to culture them in a form suitable for transplantation. In this study, we showed that hypoxia pretreatment of ADSC can promote chondrogenic differentiation while inhibiting osteogenic differentiation, which can be useful for cartilage regeneration (Fig. 10). In order to use ADSC for therapeutic purposes, further studies are needed to establish the culture strategies that allow for the expansion of ADSC while retaining their differentiation potential.

Electronic supplementary material

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Acknowledgements

This work was supported by the National Research Foundation of Korea (NRF) Grant funded by the Korean government (Grant Numbers 2016R1A6A3A04011000 and 2015M3A9C7030190). This research was supported by Chungbuk National University (2018).

Compliance with ethical standards

Conflict of interest

The authors have no conflicts to disclose.

Ethical statement

There are no animal experiments carried out for this article.

Footnotes

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Ok Kyung Hwang and Young Woock Noh have contributed equally to this work.

Contributor Information

Jin Tae Hong, Email: jinthong@chungbuk.ac.kr.

Je-Wook Lee, Email: jwl@kbiohealth.kr.

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