Physioxia: a more effective approach for culturing human adipose-derived stem cells for cell transplantation

Background Although typically cultured at an atmospheric oxygen concentration (20–21%), adipose-derived stem cells (ASCs) reside under considerable low oxygen tension (physioxia) in vivo. In the present study, we explored whether and how physioxia could be a more effective strategy for culturing ASCs for transplantation. Methods After isolation, human ASCs were cultured under physioxia (2% O2) and hyperoxia (20% O2) until assayed. WST-8, Transwell, tube formation, β-galactosidase staining, and annexin V-FITC/PI assays were used to evaluate cell proliferation, migration, angiogenesis, senescence, and apoptosis, respectively. Survivability was determined by an ischemia model in vitro and nude mouse model in vivo, and the underlying metabolic alterations were investigated by fluorescence staining, flow cytometry, and real-time polymerase chain reaction. Results Compared with those in the hyperoxia group, cells in the physioxia group exhibited increased proliferation, migration, and angiogenesis, and decreased senescence and apoptosis. The increased survival rate of ASCs cultured in physioxia was found both in ischemia model in vitro and in vivo. The underlying metabolic reprogramming was also monitored and showed decreased mitochondrial mass, alkalized intracellular pH, and increased glucose uptake and glycogen synthesis. Conclusions These results suggest that physioxia is a more effective environment in which to culture ASCs for transplantation owing to the maintenance of native bioactivities without injury by hyperoxia.


Background
Since first isolated in 1964 [1], human adipose-derived stem cells (ASCs) have garnered increasing attention [2]. Especially in the recent two decades, after the discovery of their stemness in 2001 [3], a growing body of research has indicated that ASCs possess properties of repair and regeneration, which include angiogenesis [4], multilineage differentiation [5], immunosuppression [6], and homing to ischemic tissues [7]. Consequently, there is great interest in and demand for utilizing ASCs in several clinical applications, such as osteoarthritis, heart failure treatment and wound healing, according to the clinicaltrials.gov database.
However, there are still several problems to resolve, such as the donor choice [8], therapeutic safety [9], and standard protocol for expanding ASCs [10]; among these problems, the most suitable strategy for culturing and expanding ASCs in vitro has been continuously studied. Several factors should be considered, such as the culture medium, serum replacements, and seeding density [11]. However, there is an extremely appropriate standard to which can be referred, the stem cell niche, which is the surrounding microenvironment and intrinsic factors that control the self-renewal and differentiation of stem cells [12,13].
A distinct difference between "standard culture conditions" and the ASC niche is the oxygen level [14]. Cell culture is typically performed at an atmospheric O 2 concentration (20-21%), i.e., the normoxia recognized by most researchers. However, there is a "mental shortcut" [15] neglecting the fact that the normoxia of 20-21% O 2 reflects the pathology of humans or animals, while the practical oxygen concentration of the ASC niche is lower, at 2% [16], which is called physioxia [17]. In other words, atmospheric normoxia represents a hyperoxic state for ASCs.
By comparison, previous studies have commonly used physioxia at 2% O 2 to culture ASCs [22][23][24] as a transitory approach to increase the expansion and angiogenesis of ASCs rather than as a culture standard through the entire in vitro period, except for some studies [25][26][27][28][29]; yet, these studies did not examine the angiogenesis or survival of ASCs under an ischemic environment. Thus, the aim of the present study was to explore the superiority of physioxia (2% O 2 ) compared with hyperoxia (20% O 2 ) throughout the in vitro culture of ASCs by examining discrepancies in proliferation, migration, senescence, apoptosis, angiogenesis, and survivability, as well as the underlying mechanism.

Cell isolation and culture
Subcutaneous adipose tissue was collected from the abdomen of four healthy females (age, 25 ± 5 years, body mass index [BMI]: [19][20][21][22] after obtaining their consent. After washing with phosphate-buffered saline (PBS), the tissue was minced and digested with 0.2% collagenase (Sigma-Aldrich, St. Louis, MO, USA)/PBS for 40 min at 37°C. The mixture was washed with PBS and centrifuged at 1000 rpm for 5 min, and the remaining pellet was cultured in α-modified Eagle's medium (α-MEM; HyClone, GE Healthcare, Marlborough, MA, USA), 10% fetal bovine serum (FBS; Gibco, San Jose, CA, USA), 100 IU penicillin, and 100 mg/mL streptomycin (Solarbio, Beijing, China). Cells in the physioxia group were cultured with 2% O 2 (using a modular chamber, Sanyo, Osaka, Japan) and 5% CO 2 at 37°C (physioxia ASCs, P-ASCs) until further analysis in the following tests at passage 3, with 20% O 2 and 5% CO 2 at 37°C as a control (hyperoxia ASCs, H-ASCs). Cells from different donors were mixed at passage 2 to explore the general effect on ASCs.

Cell characterization Flow cytometric analysis
Flow cytometry was used to analyze the surface markers of the ASCs. After detaching, 1 × 10 5 cells were incubated with PE-or FITC-conjugated antibodies against CD31, CD34, CD73, CD90, CD105, and HLA-DR for 30 min at 4°C. All antibodies were obtained from Abcam Biotechnology (Abcam, Cambridge, MA, USA). The cells were then analyzed using a BD Accuri™ C6 flow cytometer (BD Biosciences, San Jose, CA, USA).

Western blotting
Western blotting was performed as previously described [30], with slight modifications. After being dissolved in radioimmunoprecipitation assay (RIPA) buffer (KeyGEN, Nanjing, Jiangsu, China), 30 μg of protein, as detected by bicinchoninic acid (BCA) assay, was separated on a 10% polyacrylamide gel and blotted onto a polyvinylidene fluoride (PVDF) membrane. The membrane was blocked with 5% skim milk and then treated with primary antibodies against HIF-1 (1:1000, 14,179, Cell Signaling Technology, Beverly, MA, USA) and β-actin (1:1000, ab3280, Abcam, Cambridge, MA, USA) overnight at 4°C, followed by 1 h of incubation with horseradish peroxidase (HRP)-conjugated secondary antibodies at room temperature. The signals were detected with Amersham ECL Select Western Blotting Detection Reagent (GE, Waukesha, WI, USA) according to the manufacturer's protocol. The signals were visualized using an ImageQuant LAS 4000 mini (GE, Waukesha, WI, USA).

Cell doubling curve
ASCs were seeded onto six-well plates at a concentration of 3× 10 4 per well. The cells were collected at the indicated time points (1, 2, 3, 4, 5, 6, and 7 days), and the cell numbers were measured using an Automated Cell Counter (Bio-Rad, Hercules, CA, USA).
Determination of reactive oxygen species (ROS), mitochondrial mass, and glucose uptake

Transwell assay
After incubation in serum-free medium for 24 h, 1 × 10 5 cells were transferred to the upper chamber of a Transwell (Corning, Corning, NY, USA). Medium containing 10% FBS was added to the lower chamber as a chemoattractant. After 24 h, nonmigratory cells in the upper chamber were removed. The migrated cells were fixed with 4% paraformaldehyde for 30 min, followed by 0.1% crystal violet (Sigma-Aldrich, St. Louis, MO, USA) staining for 15 min. After images were captured, the crystal violet in the cells was extracted by 10% acetic acid for 15 min, and the absorbance at 600 nm was measured using a spectrophotometer (Multiskan GO, Thermo Fisher Scientific, Waltham, MA, USA).

Cell senescence
We assayed the ASCs for senescence-associated β-galactosidase (SA-β-Gal) activity using a Senescence β-Galactosidase Staining Kit (Beyotime, Shanghai, China) according to the manufacturer's instructions. The SA-β-Gal + area was calculated by ImageJ (NIH, Bethesda, MD, USA) using the ratio of Periodic acid-Schiff (PAS) + area to the total area of the image.

Cell apoptosis
ASC apoptosis was measured using an Annexin V-FITC/ PI Apoptosis Detection Kit (KeyGEN, Nanjing, Jiangsu, China) according to the manufacturer's instructions. Flow cytometry was conducted using the BD Accuri™ C6 flow cytometer (BD Biosciences, San Jose, CA, USA).

Cell survival
The survival assay was conducted as previously described [31].

PAS staining
We used PAS staining to explore the different expression levels of glycogen in P-ASCs and H-ASCs. Cells on six-well plates were fixed with 4% paraformaldehyde and incubation for 5 min with 0.5% periodic acid (Solarbio, Beijing, China), followed by Schiff 's reagent for 15 min. After the cells were imaged, the PAS + area was quantified by ImageJ (NIH, Bethesda, MD, USA).

Extracellular lactate assay
The lactate content of the culture medium was measured using a Lactate Assay Kit (KeyGEN, Nanjing, Jiangsu, China) according to the manufacturer's protocol.

In vivo experiment
Fibrin gel and TdT-mediated dUTP-biotin nick end labeling (TUNEL) assays were conducted as previously described [31]. The fibrin gel was composed of 25 mg/mL fibrinogen, 20 mM CaCl 2 , and 2.5 U/mL thrombin.

Statistics
Data were analyzed with GraphPad Prism 5.02 (GraphPad Software, San Diego, CA, USA) and are expressed as the mean ± standard deviation. Unpaired Student's t tests were performed, and statistical significance was considered at P < 0.05. At least three replicates were analyzed in each experiment.
Compared with the H-ASCs, the P-ASCs exhibited upregulated HIF-1 protein expression, as determined by Western blotting (Fig. 1e).

Angiogenic activities of ASCs were promoted under physioxia
Tube formation induced by Matrigel was employed to examine the angiogenic activities of the cells. The P-ASCs generated more meshes than the H-ASCs (Fig. 4a), and statistical analysis revealed significantly increased total mesh (Fig. 4b), branching length (Fig. 4c) and junction (Fig. 4d) values for P-ASCs than for H-ASCs (2.20-, 1.29-, and 1.41-fold greater, respectively). RT-PCR showed increased expression of the angiogenic genes vascular endothelial growth factor (VEGF), vascular endothelial growth factor receptor 2 (VEGF-R2) and von Willebrand factor (vWF) (Fig. 4e) in P-ASCs.

Survival of P-ASCs was strengthened under ischemic condition
After incubation in an ischemic environment (Fig. 5a) for 24 h, P-ASCs showed increased survival (Fig. 5B) and decreased death rates (Fig. 5A). A minor but significant difference was also detected under the hypoxic (Fig. 5b), acidic (Fig. 5c), and nutrient-depleted conditions (Fig. 5d).

Discussion
Offering safe and effective cell therapy products for clinical applications is consistent with good manufacturing practice (GMP) guidelines, which should be followed during the entire process of isolating, expanding and transplanting ASCs [32]. The present study compared ASCs cultured under hyperoxia (20% O 2 ) and physioxia (2% O 2 , oxygen concentration in situ) and provides compelling evidence that the latter could be a more effective approach owing to the advantages of retaining cell proliferation, migration, survival in ischemia and angiogenesis, and suppressing senescence and apoptosis.
There are no differences between P-ASCs and H-ASCs in terms of immunophenotype, morphology or adipogenesis, and a previous study [25] revealed that culturing ASCs under physioxia does not increase the risk of tumourigenesis associated with ASCs, indicating that P-ASCs are safe for clinical therapy. Physioxia promoted cell proliferation and migration, and many studies have attributed this effect to the stabilization of HIF-1 in the lack of O 2 [33,34]. However, the ROS level was also increased in P-ASCs, suggesting that transient physioxia can restore proliferation and migration through the augmentation of ROS [35]. Furthermore, we showed that without the injury caused by hyperoxia, physioxia is an appropriate condition for maintaining ASC proliferation and migration.
The relationship between physioxia and ROS is complicated [36]. In principle, HIF-1 decreases the ROS level [37,38], which should be lower in P-ASCs, but the results show the opposite effect. The underlying mechanism remains unknown, especially in stem cells.
Many studies have shown the ability of HIF-1 to enhance angiogenesis under transient physioxia [39,40], but consistent with most studies on physioxia and ASCs, the cells were isolated from a physioxic niche and then cultured under atmospheric hyperoxia, which could injure the bioactivity of the cells. Thus, the discrepancy of such bioactivity between P-ASCs and H-ASCs is not due to the acceleration of physioxia but reflects the damage caused by hyperoxia. Although transient physioxia preconditioning would be applied prior to transplantation for recovery, in the present method, culturing cells under physioxia through the entire in vitro period may be a better approach; however, further research is required.
To acquire an excellent stem cell product, cell viability should also be considered. Physioxia evidently suppressed senescence and apoptosis under nonstressful condition. The required survival of cells implanted in an ischemic environment composed of low oxygen, glucose, and pH levels is a main barrier for cell therapy [41]. Thus, we established an ischemic model and observed increased adaptability in P-ASCs; the same effect was observed in hypoxic, acidic, and nutrient-depleted environments, explaining the superiority of P-ASCs under these conditions and resulting in preferable adaptability in an ischemic environment.
This study shows for the first time that culturing ASCs under physioxia for the entire in vitro term could induce metabolic alterations and improve ASC survival in ischemic environment. Our observations illustrate the molecular, cellular, and in vivo biological effects induced by physioxia in ASCs, presenting a significant mechanistic basis for culturing ASCs under physioxia for cell therapy. However, longer culture periods should be examined to guarantee the security of cell properties under this condition. Moreover, specific cell therapy models should be constructed to verify the ultimate efficacy of the cells, including when applied in adipose regeneration, heart failure treatment, and wound healing. Increased glucose uptake and reserve of P-ASCs. a and b Fluorescent images and flow cytometry results of glucose uptake in ASCs determined by staining with 2-NBDG; the relative MFI was determined by MFI of P-ASCs versus that of H-ASCs. Scale bar = 100 μm. c Intracellular glycogen detected by PAS staining; the PAS + area was calculated as the PAS + area versus the total image area. Three fields were quantified. Scale bar = 50 μm. d Expression of HIF-1 target genes evaluated by qRT-PCR. Data are presented as the mean ± SD, *P < 0.05 (P-ASCs/H-ASCs), **P < 0.01 (P-ASCs/H-ASCs), Student's t tests, n = 3. 2-NBDG 2-[N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl) amino]-2-deoxy-D-glucose, ASCs adipose-derived stem cells, GLUT1 glucose transporter 1, glucose uptake, GLUT3 glucose transporter 3, glucose uptake, GYS1 glycogen synthase 1, glycogen synthesis, H-ASCs hyperoxia ASCs. PAS periodic acid-Schiff, P-ASCs physioxia ASCs, PGM phosphoglucomutase, glycogen synthesis, PYGL liver isoform of glycogen phosphorylase, glycogen breakdown

Conclusions
In summary, the present results suggest that culturing ASCs under physioxia (2% O 2 ) for the entire in vitro period, not under conventional hyperoxia (20% O 2 ), could be a more effective approach for cell therapy applications owing to the improvements in proliferation, migration, survival and angiogenesis, and suppression of senescence and apoptosis. Fig. 8 Physioxia increased ASC survivability in vivo. After mixing with 80 μL of fibrin gel, 1 × 10 6 P-ASCs or H-ASCs were subcutaneously transplanted into the dorsum of nude mice. The implants were extracted after 24, 48, and 72 h. a TUNEL assay was used to stain the nucleus of dead cells. The black arrows indicate dead cells. b The TUNEL + cell rate was determined by the ratio of TUNEL + cells versus total cells. Three fields were quantified. Data are presented as the mean ± SD, *P < 0.05, **P < 0.01, Student's t tests, scale bar = 100 μm. ASCs adipose-derived stem cells, H-ASCs hyperoxia ASCs, P-ASCs physioxia ASCs, TUNEL TdTmediated dUTP-biotin nick end labeling