Adipose tissue-derived stromal/stem cells (ASCs) represent a valuable tool for cell-based therapies because of their widely acknowledged capacity to exert beneficial functions in tissue regeneration or in tissue repair [1
]. This has been demonstrated for example for cell-assisted lipotransfer, where autologous ASCs added to lipografts have been shown to enhance vascularity, to improve the survival rate of grafts, and to reduce postoperative atrophy [7
]. The benefits of ASCs in this context were mainly attributed to secreted paracrine factors rather than to direct differentiation into tissue-specific cell types [10
]. In recent years, numerous studies have shown that ASCs secrete a complex panel of trophic factors, including growth factors, cytokines, chemokines, extracellular microvesicles and exosomes, that contribute to angiogenesis, anti-apoptosis, immunomodulation, and the activation of resident and circulating stem cells [10
]. However, in regenerative approaches such as tissue engineering or cell-assisted lipotransfer, the implanted cell-laden construct or lipograft is at least initially impaired by a lack of blood supply [18
]. This leads to an ischemic environment that is characterized by the deprivation of nutrients, oxygen, and growth factors. It has been shown that depletion of oxygen and nutrients, in particular glucose, significantly affects cell survival and function, as they are both critically required for energy-related pathways [20
To date, the response of ASCs to ischemic stress and the different components of ischemia remains poorly understood. Only a few studies so far examined the viability and metabolic response of ASCs to combined oxygen and glucose deprivation, but they did not focus on their secretory function under this condition [23
]. The modulation of the paracrine activity of ASCs by low oxygen concentrations is well documented. However, little is known about how nutrient deprivation, a further major component of ischemic stress, can affect their secretory potential.
In this context, the present study aimed to investigate the effect of glucose and oxygen deprivation on the viability, metabolic activity, and secretory capacity of ASCs. Specifically, we focused on glucose starvation in concert with hypoxia as a potential modulator of the paracrine function of ASCs. In response to combined glucose and oxygen deprivation, ASCs demonstrated increased levels of secreted angiogenic and anti-apoptotic mediators including vascular endothelial growth factor (VEGF), interleukin-6 (IL-6), IL-8, angiogenin (ANG), and stanniocalcin-1 (STC-1). We further investigated the impact of conditioned medium of ischemia-challenged ASCs on the viability and tube formation of endothelial cells, and the proliferation and migration of fibroblasts. The results of this study suggest that ASCs can maintain their secretory function and thus exert regenerative effects even under ischemia-like stress conditions.
2. Materials and Methods
2.1. Cells and Cell Culture
Human adipose-derived stem cells (ASCs) were obtained from Lonza (Walkersville, MD, USA). ASCs were expanded and cultured in 175 cm2 cell culture-treated plastic flasks in growth medium consisting of Dulbecco’s Modified Eagle´s Medium/Ham´s F-12 (DMEM/F-12) (Thermo Scientific, Waltham, MA, USA), supplemented with 1% penicillin/streptomycin (Thermo Scientific), 10% fetal bovine serum (FBS; Thermo Scientific), and 3 ng/mL basic fibroblast growth factor (bFGF; BioLegend, London, UK) dissolved in phosphate-buffered saline (PBS) containing 1% BSA (bovine serum albumin). Cultures were maintained under a sterile humidified 37 °C, 5% CO2, and 95% air environment. The culture medium was replaced every other day. At 80–85% confluence, the ASCs were detached using a 0.25% trypsin-EDTA solution (Thermo Scientific) and passaged. ASCs were used at passage 4 for the subsequent experiments. Human umbilical vein endothelial cells (HUVECs) were obtained from PromoCell (Heidelberg, Germany). HUVECs were plated in 25 cm2 flasks and cultured in endothelial cell growth medium 2 (PromoCell). When the cells reached 70–80% confluence, they were detached with DetachKit (PromoCell) and expanded to passages 2–3. NIH/3T3 fibroblasts were obtained from ATCC (Manassas, VA, USA). Fibroblasts were cultured in fibroblast growth medium (Dulbecco´s Modified Eagle´s Medium (Thermo Scientific), supplemented with 1% penicillin/streptomycin (Thermo Scientific), and 10% bovine calf serum (BCS; Sigma-Aldrich, Munich, Germany)). Cultures were maintained under a sterile humidified 37 °C, 5% CO2, and 95% air environment. The culture medium was replaced every other day. At 70–80% confluence, the fibroblasts were detached using a 0.25% trypsin-EDTA solution (Thermo Scientific) and passaged.
2.2. Ischemic Culture Conditions
ASCs were exposed to glucose and oxygen deprivation separately and in combination. Cells were seeded (25 × 103 cells/cm2) in growth medium, and after incubation overnight with 21% O2 for attachment of cells, ASCs were washed twice with PBS and then cultured in basal medium (d-glucose-, L-glutamine-, phenol red-, and sodium pyruvate-free DMEM) containing no serum. The medium was supplemented with D-glucose (0.1 or 1 g/L) according to the respective condition. The following conditions were examined: (1) 1 g/L glucose and normoxia (21% O2) (control); (2) 1 g/L glucose and hypoxia (0.2% O2); (3) 0.1 g/L glucose and normoxia (21% O2); (4) 0.1 g/L glucose and hypoxia (0.2% O2). Hypoxic conditions were achieved using the well-established and finely controlled proOx-C chamber system (C-Chamber, C-274; BioSpherix, New York, NY, USA). The oxygen concentration was maintained at 0.2% with the residual gas mixture composed of 5% CO2 and balance nitrogen. In order to ensure sustained hypoxic conditions, cell cultures were left undisturbed without medium changes.
2.3. Live/Dead Staining
ASCs and HUVECs were seeded in their respective growth medium and after attachment of cells, they were cultured according to the respective condition. Cell viability was determined using live/dead cell staining (PromoKine, Heidelberg, Germany) according to the manufacturer´s instructions. Living cells were stained with calcein (green) and dead cells were stained with ethidium bromide (red). Images were taken using an Olympus IX51 fluorescence microscope and analyzed with the Olympus CellSens™ Software v1.16 (Olympus, Hamburg, Germany).
2.4. Quantification of DNA
For determination of total DNA content, the intercalating dye Hoechst 33,258 was used (Polysciences, Warrington, PA, USA). Cells were harvested in phosphate-buffered saline (PBS) and sonicated with an ultrasonic homogenizer. Quantification of DNA content was carried out by measuring fluorescence intensities at an excitation wavelength of 365 nm and an emission wavelength of 458 nm with a fluorescence spectrometer (Infinite M200; Tecan, Crailsheim, Germany).
2.5. MTT Assay
ASCs, HUVECs, and NIH/3T3 fibroblasts were seeded in their respective growth medium and after attachment of cells, they were cultured according to the respective condition. At the indicated time points, cells were treated with MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) at a final concentration of 0.5 mg/mL and incubated for 3 h at 37 °C. Cells were then washed with phosphate-buffered saline (PBS), incubated with dimethyl sulfoxide (DMSO) for 5 min with gentle shaking, and mixed to ensure complete solubilization of the dye formed. The respective light absorbance of this solution was recorded using a microplate reader (Infinite M200; Tecan) at a wavelength of 570 nm. The mean value of day 0 samples was taken as reference and set as 100%.
2.6. Adipogenic Differentiation of Ischemia-Treated ASCs
To assess the adipogenic differentiation capability of ischemia-treated ASCs, cells were cultured under combined glucose/oxygen deprivation for four days. The cells were then trypsinized, re-seeded, and cultured in Preadipocyte basal medium-2 (PBM-2; Lonza) containing 10% FCS and 1% penicillin/streptomycin. After two days, adipogenic differentiation was induced by changing to differentiation medium (PBM-2 with 1.7 μM insulin (PromoCell, Heidelberg, Germany), 1 μM dexamethasone, 200 μM indomethacin (both from Sigma-Aldrich), and 500 μM 3-isobutyl-1-methylxanthin (IBMX; Serva Electrophoresis, Heidelberg, Germany)) for 14 days. Non-induced cells were cultured in PBM-2 during the differentiation period. In parallel, control ASCs were cultured in growth medium for four days and then treated in the same way for adipogenic differentiation. After 14 days of culture in adipogenic medium, lipid accumulation was histologically assessed by staining with Oil Red O solution (3 mg/mL Oil Red O in 60% isopropanol; Sigma-Aldrich) and cell nuclei were counterstained with hematoxylin (Bio Optica, Milan, Italy). Samples were imaged using an Olympus IX51 inverted light microscope and analyzed with the Olympus CellSens™ Software v1.16 (Olympus, Hamburg, Germany).
The quantitative determination of intracellular lipid accumulation was performed using the Serum Triglyceride Determination Kit from Sigma-Aldrich. Cells were harvested and sonified in Thesit solution (0.5% Thesit in H2O; Gepepharm, Hennef, Germany) and the triglyceride quantification was performed according to the manufacturer’s instructions and measured with a microplate reader (Infinite M200; Tecan) at a wavelength of 570 nm. Triglyceride contents were normalized to the DNA content of the respective samples.
2.7. Glucose and Lactate Determination
Exogenous glucose and lactate levels in the cell culture supernatants were measured with GLUC3 and LACT2 COBAS INTEGRA substrate reagents using the related COBAS INTEGRA 800 (Roche, Basel, Switzerland) robot.
2.8. Assays of Cytokines
Antibody array: To identify factors secreted by glucose/oxygen-deprived ASCs, a profiling of human cytokines was performed using an antibody array covering 80 cytokines (Human Cytokine Antibody Array C5; RayBiotech, Norcross, GA, USA). Cell culture supernatants from glucose/oxygen-deprived ASCs (0.1 g/L glucose, 0.2% O2) and ASCs cultured under the control condition (1 g/L glucose, 21% O2) were centrifuged at 1000× g for 5 min to remove cell debris. Array analyses were performed according to the manufacturer’s instructions. Briefly, the array membranes were blocked with a blocking buffer and incubated with 1 mL of each supernatant overnight at 4 °C. Subsequently, the membranes were assayed for chemiluminescence signals.
Enzyme-linked immunosorbent assays (ELISAs): The concentrations of individual cytokines in the cell culture supernatants from cells cultured under the different deprivation conditions and the control condition were determined using ELISA kits for vascular endothelial growth factor (VEGF), interleukin (IL)-6, IL-8, angiogenin (ANG), TIMP metallopeptidase inhibitor (TIMP)-1, monocyte chemoattractant protein (MCP)-1, and stanniocalcin (STC)-1 from R&D Systems (DuoSet ELISA; Minneapolis, MN, USA). Concentration levels were normalized to the total DNA content of the respective samples (pg/µg DNA).
2.9. RNA Isolation and Quantitative Real-Time PCR (qRT-PCR) Analysis
Total RNA from cultured cells was isolated using TRIzol® reagent (Invitrogen, Karlsruhe, Germany). First-strand cDNA was synthesized from total RNA with ImProm-II Reverse Transcription System (Promega, Mannheim, Germany). Quantitative PCR analyses were performed using the MESA GREEN qPCR MasterMix Plus with MeteorTaq polymerase (Eurogentec, Seraing, Belgium). cDNA for genes of interest was amplified with the PrimePCR™ SYBR® Green Assay using the following cycle conditions: 95 °C for 15 min initial denaturation followed by 40 cycles at 95 °C for 15 s, 60 °C for 30 s, and 70 °C for 30 s using the following primers: IL-6 (qHsaCID0020314, IL6, human), VEGF (qHsaCED0043454, VEGFA, human), and STC-1 (qHsaCID0006115, STC1, human), all from BioRad (Hercules, CA, USA). mRNA expression levels were normalized to the eukaryotic translation elongation factor 1 alpha (EF1α) (forward, 5′-ccccgacacagtagcatttg-3′; reverse, 5′-tgactttccatcccttgaacc-3′) (Biomers, Ulm, Germany). The relative expression levels were determined using the 2−ΔΔCT method and were further normalized to the respective day 0 sample.
2.10. Preparation of Conditioned Medium
ASCs were seeded at 25,000 cells per cm2 in growth medium and allowed to adhere overnight at 21% O2. ASCs were washed twice with PBS, and the medium was replaced with basal medium (D-glucose-, L-glutamine-, phenol red-, and sodium pyruvate-free DMEM) containing no serum and supplemented with 0.1 g/L glucose. Cells were incubated under 0.2% O2, to generate a conditioned medium (CM) of ASCs exposed to glucose/oxygen deprivation. After four days, the medium was harvested as ASC-CMischemic.
2.11. Tube Formation Assay
Angiogenesis µ-Slides (Ibidi, Gräfelfing, Germany) were coated with 10 µL of growth factor- reduced matrigel (BD Biosciences, San Jose, CA, USA). HUVECs were suspended in basal medium, ASC-CMischemic or endothelial growth medium and plated with 1 × 104 cells per well on top of the matrigel. After 4, 6, and 10 h of incubation at 37 °C under hypoxic conditions (0.2% O2), the formation of tube-like structures was examined microscopically. The tube length and branch count were quantified using the automated image analyzer ACAS from ibidi (Tube formation ACAS image analysis module) at the indicated time points.
2.12. Proliferation and Metabolic Activity of Fibroblasts
The conditioned medium from glucose/oxygen-deprived ASCs (ASC-CMischemic) was prepared as described. Fibroblasts were treated with basal medium (DMEM, w/o FBS) or ASC-CMischemic, each supplemented with 1 g/L glucose, under normoxic conditions. Proliferation and metabolic activity of the cells were analyzed at the indicated time points using a DNA and MTT assay as described above.
2.13. Fibroblast Migration Assay
The migratory activity of NIH/3T3 fibroblasts was assessed using a migration assay. Ibidi Culture-Inserts 2 well (Ibidi, Gräfelfing, Germany) were transferred into 6-well plates and 70 µL cell suspension containing 3 × 105 cells/mL was applied to each well. After an appropriate duration for cell attachment (24 h) the Ibidi Culture-Inserts were removed to create a cell-free gap of 500 µm. Cells were then washed with phosphate-buffered saline (PBS), and incubated with basal medium (DMEM, w/o FBS) or ASC-CMischemic, each supplemented with 1 g/L glucose, under normoxic conditions for 24 h. The fibroblast growth medium was used as positive control. To monitor the progress of gap closure, micrographs were taken at different time points.
2.14. Statistical Analysis
Quantitative results are presented as means ± SD. Statistical analyses of variance comparisons between groups were performed using the ANOVA-test in conjunction with Bonferroni post-hoc adjustment. For statistical analyses of endothelial tube formation an unpaired Student’s t-test was applied. For all analyses, differences at p < 0.05 were considered as statistically significant. All statistical analyses were performed using the GraphPad Prism Software 8.3 (GraphPad Software, San Diego, CA, USA).
The positive effects of ASC-based approaches in regenerative therapies have been demonstrated in preclinical and clinical studies, for example in cell-assisted lipotransfer or treatment of ischemic diseases [9
]. However, a substantial loss of implanted cells has been documented during the early phase of engraftment [12
]. Since ASCs, after transplantation into damaged tissues, are exposed to an ischemic environment characterized by the deprivation of nutrients and oxygen, a better understanding of the mechanisms underlying the beneficial effects in the early phase following transplantation is required. This would contribute to a more rational application of ASCs in cell-based approaches. The regenerative potential of ASCs nowadays is mainly attributed to their trophic activity through the secretion of angiogenic, anti-apoptotic, and immuno-modulatory factors. Hypoxia as one of the hallmarks of ischemia is well reported to enhance the secretion of such factors [32
]. However, little is known about the ability of ASCs to maintain their secretory function under starvation conditions. Thus, we investigated the viability and secretory function of ASCs in response to glucose deprivation and severe hypoxia and the combination of both as major stress conditions in an ischemic environment. To our knowledge, no study has yet been conducted to investigate the effect of glucose deprivation as one of the components of ischemia on the secretion capacity of ASCs.
The hallmarks of ischemia were simulated in our in vitro set-up by culturing the ASCs under glucose deprivation (0.1 g/L glucose) and severe hypoxia (0.2% O2
) alone and in combination over a period of seven days in serum-free medium to mimic ischemic conditions during the early post-transplantation phase. Serum-free culture in combination with oxygen and also glucose deprivation is a widely used experimental model to mimic ischemic conditions in vitro. Furthermore, serum-free culture is also a common approach in studies analyzing the effects of secreted proteins by using a conditioned medium in order to avoid interference with serum proteins [35
]. Viability and cell morphology were shown to be virtually not affected by the applied stress conditions over the culture period of seven days. Sustained viability of ASCs in response to adverse nutrient and oxygen levels was also reported by Mischen et al. [23
] in a comparable set-up and time frame. In contrast to the limited cell death, the metabolic activity of the cells was significantly affected by glucose limitation from day 1, whereas severe hypoxia did not particularly influence the metabolic activity. Under the glucose-limiting condition (0.1 g/L), cells faced a complete exhaustion of glucose from day 4 under hypoxia, whereas the glucose level decreased more slowly in the normoxic condition. One g/L glucose was not limiting in this setup. In general, glucose levels in the medium demonstrated a steeper decline of the available glucose, when cells were exposed to hypoxia. The corresponding increase of lactate as an important by-product in glycolysis indicated that ASCs increasingly rely on anaerobic glycolysis for their metabolic demands when exposed to hypoxic conditions. Several studies exploring the metabolism of bone marrow-derived mesenchymal stem cells (MSCs) under glucose and oxygen deprivation demonstrated the metabolic flexibility of MSCs under such adverse conditions and their ability to rely on anaerobic glycolysis for energy supply in a hypoxic environment [21
]. In this context, the crucial role of glucose for MSCs function in a hypoxic environment was emphasized. In the present study, the cells were able to maintain their viability for several days despite the complete exhaustion of glucose. This could possibly be due to an enhanced autophagic activity of the cells, as autophagy has been shown to be a survival mechanism for oxygen/glucose-deprived MSCs [36
]. A further observation was that those ASCs that survived under the harsh ischemic condition (glucose/oxygen deprivation) were not affected in their adipogenic differentiation capability. This finding additionally underlined the remarkable resilience of hASCs to an adverse environment.
The next step was to examine the secretion of the ASCs exposed to oxygen and/or glucose deprivation to determine whether cells under ischemic stress were able to maintain their secretory function. As displayed by a cytokine antibody array, ASCs were able to express a broad range of growth factors, cytokines, and chemokines, which in part appeared to be stimulated by hypoxia. When glucose deprivation was combined with hypoxia in order to mimic ischemia, ASCs were still able to maintain secretory function. Under this condition, the cells expressed growth factors and cytokines with angiogenic (VEGF, IL-6, IL-8, ANG) and matrix-remodeling (TIMP-1, TIMP-2) functions and chemokines (MCP-1/CCL2, IP-10/CXCL10) among others, while cytokines associated for example with the regulation of proliferation, cell division, and differentiation appeared to be expressed to a lesser extent (e.g., FGF-4 and -6, IGFBP-2, -3, and -4, NAP-2).
To reveal the impact of the individual stress condition on the expression level of selected factors (VEGF, IL-6, IL-8, ANG, TIMP-1, and MCP-1), their expression was investigated under glucose and oxygen deprivation separately and in combination. STC-1 was included in the analysis as a factor associated with the reduction of apoptosis, angiogenesis, and enhanced resistance of cells to metabolic stress [25
]. Furthermore, STC-1 appears to be closely related to cellular metabolism, as a role of STC-1 in the activation of AMP-activated protein kinase (AMPK) has been postulated. AMPK in turn is a key regulator in the cellular adaptive response to ischemia [39
]. STC-1 was shown to be expressed by different cell types including bone marrow-derived MSCs [25
] but is a still unknown factor in ASCs. Thus, to the best of our knowledge, the secretion of STC-1 by ASCs was demonstrated for the first time in this study. We found different response patterns of the investigated factors to the individual stress conditions. It is generally accepted that hypoxia triggers the expression of a variety of growth factors and cytokines in ASCs [32
]. Accordingly, we also found an increase in the expression of most of the factors investigated, when the cells were exposed to 0.2% O2
(VEGF, IL-6, IL-8, ANG, TIMP-1, STC-1). MCP-1 showed no response to reduced oxygen levels. However, when hypoxia was combined with glucose deprivation in order to mimic ischemia, the secretion of VEGF, IL-6, IL-8, ANG, and STC-1, which are all factors with angiogenic and/or anti-apoptotic properties, increased markedly compared to the hypoxic condition alone. Thus, glucose deprivation (in conjunction with hypoxia) proved to be a factor that positively influenced the secretion of angiogenic and anti-apoptotic cytokines. The availability of glucose as a variable that influences the secretion performance of cells has hardly been investigated so far. Bakopoulou et al. [44
] examined the secretion of human apical papilla mesenchymal stem cells subjected to glucose and oxygen deprivation and they also reported a stimulating effect of glucose deprivation on the secretion of angiogenic growth factors in conjunction with hypoxia. In addition, VEGF has been described in early studies with glioma tumor cells as a “classical stress-induced gene”, whose secretion was enhanced by oxygen and glucose deficiency [45
]. In contrast, Deschepper et al. [47
] considered glucose essential for the response of hMSC to near-anoxic conditions. They reported a moderate increase in VEGF-C secretion with increasing glucose concentrations under severe hypoxia in MSCs. With regard to IL-6, the elevated expression under both glucose and oxygen deprivation determined in the present study is in accordance with reports that glucose deprivation triggers IL-6 expression by activation of ER stress signaling pathways [48
]. The consideration of the time course of secretion additionally underlined the effect of glucose deficiency on the secretion of ASCs with a distinct increase in the levels of IL-6 and VEGF from the time point of complete glucose exhaustion in the culture (day 4). STC-1 secretion was detectable from day 3 under glucose/oxygen deprivation and was maintained until day 7 with a significant increase as compared to hypoxia alone. Gene expression of STC-1 was immediately upregulated in response to glucose/oxygen deprivation. This may indicate that STC-1 possibly elicits a stimulatory effect on VEGF expression. Several studies have shown that the expression of VEGF is associated with STC-1 and a positive feedback-loop between STC-1 and VEGF stimulation has been postulated [50
Maintaining endothelial function and promoting angiogenesis in an ischemic environment to ensure adequate blood supply are considered key processes in regenerative approaches such as cell-assisted lipotransfer or treatment of ischemic diseases. The angiogenic function of secreted growth factors and cytokines of (co-)implanted ASCs could play an important role in this context [30
]. For this reason, we investigated the impact of the secretome of ischemia-challenged ASCs on the viability, metabolic activity, and tube formation of endothelial cells (HUVECs). The results indicated that ASCs exposed to ischemia-mimicking conditions were able to restore endothelial cell viability, metabolic activity, and tube formation via their secretory function. The angiogenic response to VEGF, IL-6, IL-8, and ANG, which were shown to be major factors secreted under this condition, has been well documented [53
]. Further studies are necessary to clarify to what extent the newly detected STC-1 in ASCs may contribute to this effect.
ASCs have further been shown to improve wound healing by promoting angiogenesis and fibroblast activation through their paracrine action [58
]. Here, we demonstrated that factors secreted by ASCs in an ischemia-like environment, besides their angiogenic activity, stimulated the proliferation, metabolic activity, and migration of fibroblasts. IL-6 and IL-8 are known as factors that play an important role in wound healing by triggering fibroblast and keratinocyte migration, leukocyte infiltration, and collagen synthesis [61
]. MCP-1 (CCL-2), which was also prominently expressed by oxygen/glucose-deprived ASCs, is a further important chemokine involved in wound healing processes, acting mainly through the recruitment of macrophages [64
]. Altogether, the findings support the regenerative potential of ischemia-challenged ASCs in wound healing, for example in chronic ischemic wounds. In contrast, the proliferation of breast cancer cell lines (MCF-7 and MDA-MB-231) was not increased in the presence of trophic factors from ASCs cultured under ischemia-like stress conditions. This may be relevant in the context of cell-assisted lipografting after mastectomy, where ischemic conditions at the transplantation site are likely to prevail. This observation may also be in line with reports from clinical studies that have suggested no increase in cancer recurrence rates in breast cancer patients treated with ASC-enriched lipografts, but further studies would be needed to more specifically assess the effects of the secretome of ischemia-challenged ASCs on breast cancer cells [65