Decelerating Mature Adipocyte Dedifferentiation by Media Composition

Abstract

The establishment of adipose tissue test systems is still a major challenge in the investigation of cellular and molecular interactions responsible for the pathogenesis of inflammatory diseases involving adipose tissue. Mature adipocytes are mainly involved in these pathologies, but rarely used in vitro, due to the lack of an appropriate culture medium which inhibits dedifferentiation and maintains adipocyte functionality. In our study, we showed that Dulbecco's Modified Eagle's Medium/Ham's F-12 with 10% fetal calf serum (FCS) reported for the culture of mature adipocytes favors dedifferentiation, which was accompanied by a high glycerol release, a decreasing release of leptin, and a low expression of the adipocyte marker perilipin A, but high expression of CD73 after 21 days. Optimized media containing FCS, biotin, pantothenate, insulin, and dexamethasone decelerated the dedifferentiation process. These cells showed a lower lipolysis rate, a high level of leptin release, as well as a high expression of perilipin A. CD73-positive dedifferentiated fat cells were only found in low quantity. In this work, we showed that mature adipocytes when cultured under optimized conditions could be highly valuable for adipose tissue engineering in vitro.

Introduction

Besides the storage of energy in the form of triglycerides and its endocrine and metabolic functions, fatty tissue is important for the padding of inner organs, and therefore, their protection from mechanical stress. However, fatty tissue is often involved in diseases. Several studies have shown that besides obesity, adipose tissue plays a role in acute pancreatitis, Morbus Crohn, cancer diseases, or nonalcoholic liver steatosis.^1–4 Test systems are needed to investigate the cellular and molecular interactions responsible for the pathogenesis as well as to examine the effect of drugs, which might have a positive influence on fatty tissue and could relieve pain or promote recovery. To date, mainly preadipocytes differentiated to adipocytes have been used to investigate mechanisms occurring in fatty tissue in vitro, which is very time consuming as well as material and cost-intensive. Preadipocytes can be isolated in large amounts from fatty tissue, but need to be differentiated into the adipogenic lineage for at least 14 days. It is known that only around 42% of the cells are differentiating into the adipogenic lineage.⁵ In contrast, a high amount of adipocytes could be harvested from small tissue samples, since ∼50% of fatty tissue is composed of mature adipocytes. Since adipocytes are terminally differentiated, they can be used immediately for test purposes.⁶ Several studies have been performed to establish the isolation and characterization of mature adipocytes in between the 1960s and 1990s.^7–10 Since then, mature adipocytes were only rarely used, probably because of their mainly cited drawbacks such as vulnerability and dedifferentiation under in vitro conditions. To make these adipocytes available for several applications, an optimized culture medium is needed to keep them in their quiescent status in vitro over a long period of time. To date, no literature is available on the optimal long-term culture conditions for mature adipocytes. It is well known that these cells start to dedifferentiate after 1 or 2 weeks,^7,11,12 at which point, the cells reach a multivacuolar morphology while they are diminishing their cell volume.^9,13 Elongated, nonlipid-filled dedifferentiated fat cells (DFAT) are further able to proliferate and exhibit multilineage potential.^14,15 These cells are known to express stem cell markers, such as CD73.¹⁶ An improved media composition, which minimizes lipolysis and therefore the dedifferentiation process, could stabilize mature adipocytes and keep them in their quiescent status in vitro.

Several substances are known to reduce or inhibit lipolysis and favor lipogenesis.^17–23 Insulin, as an anabolic hormone, is known to play a multipotent role in glucose and lipid metabolism, where it is mainly responsible for the uptake and storage of glucose and fatty acids.²⁴ Dexamethasone activates transcription factors upregulating enzymes that promote lipid uptake and adipogenesis.²⁵ Biotin, also known as vitamin H, serves as a prosthetic group for carboxylases, which are involved in gluconeogenesis and fatty acid biosynthesis.^26,27 Pantothenic acid, also known as Vitamin B5, is important for the synthesis of coenzyme A, which is needed for the synthesis of fatty acids.^28,29 Fetal calf serum (FCS) has contrary effects on mature adipocytes, depending on the used concentration. On the one hand, serum contains some insulin-like factors that have comparable effects on glucose as insulin does.¹⁷ On the other hand, a high amount of serum favors the dedifferentiation of mature adipocytes because it also contains several factors inhibiting lipogenesis, like tumor necrosis factor-alpha, growth hormones, and epidermal growth factor.²³

In this study, we evaluated two commercially available media concerning their ability to maintain the morphology and functionality of mature adipocytes for up to 3 weeks.

Materials and Methods

Human tissue samples

All research was carried out in accordance with the rules for investigation of human subjects as defined in the Declaration of Helsinki. Patients gave a written agreement according to the permission of the Landesärztekammer Baden-Württemberg (F-2012-078; for normal skin from elective surgeries).

Adipocyte isolation and measurement of cell size

Mature adipocytes were isolated from human fatty tissue of plastic surgeries received from Dr. Ziegler (Klinik Charlottenhaus). The isolation was based on the procedure previously described by Zhang et al.³⁰ Briefly, the connective tissue was removed and adipocytes were isolated by collagenase digestion (125 U mL⁻¹ in Dulbecco's Modified Eagle's Medium [DMEM; Biochrom] containing 1% bovine serum albumin [BSA; Sigma-Aldrich]) for 1.5 h at 37°C under agitation. The cell suspension was filtered through a 500-μm meshed sieve, centrifuged (19 g, 1 min), and washed twice.

The mean size of mature adipocytes and DFAT was determined of at least 60 cells from three different donors, whereby the inner diameter of the adipocytes and the longitudinal axis of the DFAT were evaluated.

Encapsulation of mature adipocytes into collagen type I hydrogels

Mature adipocytes were seeded in a collagen type I hydrogel (9 mg mL⁻¹ from rat tail; Fraunhofer IGB) in 24-well plate inserts (Brand). A neutralization buffer consisting of 10 × DMEM/Ham's F-12 (Biochrom) and 50 mM NaOH in Aqua dest (1:1) with 0.2 M NaHCO₃ and 0.225 M HEPES (Roth) was prepared. Four portions of collagen gel were mixed with four portions of packed mature adipocytes and one portion of neutralization buffer. Three hundred microliters of the mixture was pipetted into each well and gelled for 10–20 min at 37°C.

Cells were then cultured for 3 weeks in three different media compositions: Medium 1 consisted of DMEM/Ham's F-12 and 10% FCS as a control medium. As medium 2, AM-1 (ZenBio) was used, consisting of DMEM/Ham's F-12 medium, HEPES pH 7.4, fetal bovine serum, biotin, pantothenate, human insulin, and dexamethasone. Medium 3 was the Adipocyte Nutrition Medium (PromoCell) supplemented with 0.03 mL mL⁻¹ FCS (Gibco), 8 μg mL⁻¹ d-biotin, 0.5 μg mL⁻¹ human insulin, and 400 ng mL⁻¹ dexamethasone (all purchased from Sigma-Aldrich).

Immunofluorescence staining

Hydrogels were fixed using Bouin's fixation (Sigma-Aldrich) for 1 h and watered with tap water for several hours. Gels were embedded into paraffin and slices of 5 μm were generated. Deparaffinization was done according to a standard protocol. Sections were heat demasked with a target retrieval solution, pH 9 (Dako), for 20 min in a preheated steamer (Braun). Cells were blocked with 3% BSA in PBS⁺ for 30 min. Primary antibody (perilipin A: 1:200; Sigma-Aldrich, CD73: 1:25; Santa Cruz) was diluted with an antibody diluent (Dako) and incubated overnight at 4°C. Tissues were then stained using the EnVision+ System-HRP (Dako) according to the manufacturer's protocol.

Live cell staining

Cytoplasm/lipid droplet/nuclei staining

Nonfixed hydrogels were stained with 1 μg mL⁻¹ Hoechst 33342 in PBS⁺ for 15 min at 37°C. Thirty μL mL⁻¹ Nile Red (Lonza) and 10 μg mL⁻¹ fluorescein diacetate (FDA; Sigma-Aldrich) were added and incubated for another 15 min at 37°C. Images were taken in PBS⁺ using a laser scanning microscope (Zeiss).

Cytoplasm/nuclei staining

Cells were stained with FDA and Hoechst 33342 as described above. The total cell number was determined as the number of nuclei from 5 × magnified 100 μm z-Stack images using the Cell Profiler Cell Images Analysis Software (Broad Institute, United Kingdom). The ratio of univacuolar and multivacuolar cells and DFAT was analyzed manually. For every analysis day, at least three representative images of three hydrogels were analyzed per medium composition.

Functionality tests

Leptin enzyme-linked immunosorbent assay

An enzyme-linked immunosorbent assay (Pepro Tech) was performed according to the manufacturer's protocol. For color development, 100 μL tetramethylbenzidine (TMB) substrate (Sigma-Aldrich) for each well was used. Incubation took place at room temperature (RT) for ∼15 min. The enzymatic reaction was stopped with 50 μL 1 M sulfuric acid, and the color development was measured at 450 nm with a wavelength correction set at 570 nm (Tecan).

Glycerol release

Lipolysis was detected with a glycerol kit (Randox). Briefly, 5 μL samples were mixed with 100 μL reagent and incubated for 10 min at RT. The color development was analyzed at 520 nm.

Leptin and glycerol measurements were performed 24 h after the last media exchange. Values were normalized to the number of univacuolar and multivacuolar adipocytes, named as fat cells.

Statistics

All experiments were repeatedly performed, successively using cells from at least three different donors. Data were compared by a one-way analysis of variance with a Tukey post hoc test using Origin Pro 8.5 G. Statistic significances were stated as p ≤ 0.05.

Results

Mature adipocytes dedifferentiated during in vitro culture

Mature adipocytes were encapsulated in a collagen type I hydrogel and cells cultured in medium 1 dedifferentiated over time (Fig. 1). After encapsulation, they showed a round morphology with one huge lipid droplet filling the whole cytoplasm. The nucleus was compressed to the cell membrane. Over the culture period, cells were elongating and stored their lipids in several lipid droplets. The nucleus migrated to the center of the cell. When reaching a fibroblast-like morphology, these DFAT did not store any lipid vacuoles in their cytoplasm and started to proliferate. Mature adipocytes had a diameter of 79 ± 19.0 μm, whereas DFAT had a nonsignificantly smaller cell length with 70 ± 24.4 μm (Fig. 1D).

FIG. 1.

Mature adipocytes lose their lipid droplets during dedifferentiation. Cells were cultured in medium 1. Lipid vacuole was stained with Nile Red and is shown in red, nuclei were stained with Hoechst 33342 in blue, and cytoplasm was marked with fluorescein diacetate (FDA) in green. (A) Mature adipocytes. (B) Multivacuolar dedifferentiated cells (image taken on day 14). (C) DFAT (image taken on day 14). (D) The mean size of mature adipocytes and DFAT was determined of at least 60 cells from three different donors. The inner diameter of the adipocytes and the longitudinal axis of the DFAT were evaluated. DFAT, dedifferentiated fat cells. Scale bar: 20 μm. Color images available online at www.liebertpub.com/tec

Optimized media reduced the dedifferentiation of mature adipocytes

The total number of cells encapsulated in a collagen type I hydrogel was evaluated and shown in Figure 2 as a percentage of the control medium (medium 1) on day 1. A similar cell amount (112% ± 43.6%) could be found in medium 2, whereas in medium 3, significantly fewer cells were available (24% ± 4.4%). Only a slight decrease in the cell number could be seen in medium 1 and 2 on day 7, whereas the number of cells in medium 3 stayed constant up to day 21. In medium 1 and 2, cells proliferated on day 14, to a level of 125% ± 43.1% and 152% ± 23.8%, respectively. The cell amount further increased on day 21 to 133% ± 65% in medium 1 and 173% ± 62.7% in medium 2.

FIG. 2.

Total cell number per hydrogel. To evaluate the total cell number, the amount of nuclei was counted from 5× magnified 100 μm z-Stack images and given as a percentage of the control medium 1 on day 1 (p ≤ 0.05).

Figure 3A shows the percentile number of univacuolar and multivacuolar adipocytes and DFAT when culturing mature adipocytes for 21 days in three different media. On day 1, only mature adipocytes could be found (Fig. 3B–D). The number of mature adipocytes on day 7 in medium 1 with 62.3% ± 25.37% was significantly lower when compared with medium 3 with 85.7% ± 10.29%. In medium 2, about 83.2% ± 13.36% mature adipocytes could be found. The number of multivacuolar adipocytes increased only slightly in all three media. The number of DFAT was significantly higher in medium 1 (37.2% ±24.94%) compared with medium 2 (12.1% ± 11.58%) and medium 3 (13.3% ± 10.18%).

FIG. 3.

Evaluation of mature adipocytes cultured in different media for up to 21 days. (A) The number of univacuolar and multivacuolar adipocytes and DFAT cultured for 21 days in three different media was determined microscopically from tissue equivalents which were stained with FDA (cytoplasm, green) and Hoechst 33342 (nuclei, blue). (B–D) Cells on day 1 (top) and day 21 (bottom). (B) Adipocytes cultured in medium 1, (C) medium 2, and (D) medium 3. Living cells are seen in green and nuclei are shown in blue. Scale bar: 50 μm; p ≤ 0.05. Color images available online at www.liebertpub.com/tec

On day 14, only 49.0% ± 28.89% of the cells were mature adipocytes in medium 1, whereas medium 2 and 3 contained 72.8% ± 19.63% and 72.8% ± 17.01% mature adipocytes, respectively. The number of multivacuolar adipocytes was significantly higher in medium 2 (23.1% ± 17.85%) compared with medium 1 and 3. DFAT increased significantly on day 14 in medium 1 compared with medium 2 and 3: 48.4% ± 26.58% DFAT could be found in medium 1, 4.1% ± 3.22% were available in medium 2, and 20.8% ± 15.89% have been found in medium 3.

On day 21, the highest amount of mature adipocytes could be found in medium 3 with 71.5% ± 21.14%, followed by medium 2 with 45.6% ± 28.27% and medium 1 with 39.9% ± 27.51%. The number of multivacuolar adipocytes was highest in medium 2 with 47% ± 33.47%, but very few could be found in medium 1 and 3. The significantly highest amount of DFAT was seen in medium 1 on day 21 with 58.9% ± 27.64%, whereas in medium 2, only 7.4% ± 9.88% DFAT could be found. Medium 3 had 20.8% ± 16.94% DFAT. This is visualized in Figure 3B–D.

Specific marker expression is shown in Figure 4. On day 1, perilipin A was expressed from adipocytes cultured in all media, where there was a lower expression in medium 3. Only slight CD73 staining was found on day 1. On day 21, cells expressed perilipin A to a lower extent than on day 1. Perilipin A could also be found in multivacuolar fat cells. A high amount of CD73-positive cells was detected in medium 1 on day 21, whereas the level was lower in medium 2 and 3.

FIG. 4.

Perilipin A and CD73 expression of cells in different media. (A–D) Medium 1, (B–E) medium 2, and (C, F) medium 3. Scale bar: 50 μm. Color images available online at www.liebertpub.com/tec

Mature adipocytes remained functional when cultured in an optimized media

The lipolytic activity of cells cultured in different media was analyzed by measuring the glycerol release and is shown in Figure 5. The values were normalized to 10,000 fat cells, whereas univacuolar as well as multivacuolar cells were considered since both cell types are able to undergo lipolysis.

FIG. 5.

Lipolytic activity of cells cultured in different media by measurement of the glycerol release. The values were normalized to 10,000 fat cells, where univacuolar and multivacuolar cells were considered to undergo lipolysis (p ≤ 0.05).

On day 1, only a slight release of glycerol occurred in all tested media, whereas the amount of glycerol in medium 3 was significantly higher than in medium 1 and 2. On day 7, a significantly higher amount of glycerol was measured from cells cultured in medium 1 with 57 ± 19.4 nmol. In medium 2, 13 ± 6.4 nmol glycerol could be measured in the supernatants. A higher amount of glycerol of 27 ± 14.6 nmol could also be found in medium 3 on day 7. On day 14, the glycerol release was reduced again. Glycerol release decreased further on day 21 to 14 ± 5.0, 8 ± 5.4, and 19 ± 11.7 nmol in all tested media, respectively.

Cell functionality was analyzed by measuring the leptin release, which is seen in Figure 6; again, the data were normalized to 10,000 fat cells. On day 7 to 21, a significant higher release of leptin was found in medium 2 and 3 compared to medium 1. A similar release of leptin was seen in medium 1, 2, and 3 on day 1 with 0.7 ± 0.30, 0.3 ± 0.31, and 0.2 ± 0.13 ng leptin, respectively. In medium 1, the leptin release stayed constantly on this low level. Cells cultured in medium 2 released 3.3 ± 2.08 ng leptin on day 7, which only decreased slightly up to day 21. In medium 3, 4.0 ± 2.0 ng leptin was released on day 7 and on day 14 and 21, a slightly lower amount with 3.2 ± 1.48 and 2.8 ± 1.47 ng leptin, respectively, were measured.

FIG. 6.

Leptin release from cells cultured in different media for up to 21 days. The values were normalized to 10,000 fat cells, where univacuolar and multivacuolar cells were considered to produce leptin (p ≤ 0.05).

Discussion

Mature adipocytes are involved in various diseases, such as obesity or cancer. To date, mainly differentiated preadipocytes have been used for investigations in adipocyte-related cross talks, since mature adipocytes dedifferentiate under in vitro conditions. In this study, two media compositions were evaluated in comparison to a standard medium for their suitability to culture mature adipocytes.

Mature adipocytes dedifferentiated during in vitro culture

It was previously demonstrated that mature adipocytes dedifferentiate in vitro.^10,14,31,32 DFAT are then able to proliferate.^33,34 Adipocytes are thought to be very plastic cells and depending on the nutrition status, they can store or release triglycerides, which has a drastic impact on their cell size.³⁵ It is thought that by means of lipid droplet fragmentation, and therefore, surface enlargement, lipases have a facilitated contact to triglycerides, which gives a direct link to lipolysis.^36,37

Optimized media reduced the dedifferentiation of mature adipocytes

Triggers for the dedifferentiation of mature adipocytes are still mainly unknown, and only a few hypotheses are available. Some authors suggested the susceptibility of adipocytes against hypoxia.^23,38 Others proposed that a high cell density is important to decelerate the dedifferentiation process.³⁹ Therefore, we used the maximal number of fat cells possible to incorporate into a stable collagen type I hydrogel. It is also believed that the culture of human fat cells in the absence of stimulatory factors results in spontaneous basal lipolysis.⁴⁰ We were now able to show that the morphological and functional stability of mature adipocytes could be largely maintained by using an optimized media in vitro.

The total cell number was increasing slightly in medium 1 and 2 over the culture period. Since mature adipocytes are in a terminal differentiation status, it is likely that in medium 1, DFAT started to proliferate and therefore increased the cell number. In medium 2, only a small amount of DFAT were available. We propose that multivacuolar adipocytes were able to divide, which was already shown by Sugihara et al.¹⁰ In medium 3, there was an initial drastic decrease in the cell number, which then stayed stable over the culture period. This led us to the assumption that for the initial cell survival, either a high FCS content (medium 1) or the addition of pantothenate, the supplement that distinguished medium 2 and 3, might be indispensable. Further research is needed to elucidate the roles of FCS and pantothenate in initial mature adipocyte cell survival.

We have seen that cells cultured in medium 3 had the highest percentage of mature adipocytes on day 21. In medium 1, mature adipocytes were mainly replaced by DFAT, which could be confirmed by immunofluorescence staining of CD73. In medium 2, however, only a small amount of DFAT could be detected, and most of the cells were multivacuolar. The decelerated dedifferentiation in medium 2 and 3 is thought to rely on the content of FCS, biotin, insulin, dexamethasone, and pantothenate (only in medium 2).

Justesen et al. showed that mature adipocytes were dedifferentiating in the presence of 10% FCS, which could be confirmed by us in medium 1.⁴¹ It is thought that FCS contains many insulin-like factors, and Gliemann proposed that 2% serum has the same effect as insulin on the intake and metabolism of 0.1 mg mL⁻¹ glucose in mature adipocytes.¹⁷ This supports our results that a high FCS content of 10% in medium 1 favored dedifferentiation, whereas a low content in medium 3 promoted the maintenance and viability of mature adipocytes. We also suggest that medium 2 has a low FCS content. Other groups have performed their studies with a low serum content between 1% and 3% FCS, but did not evaluate the long-term culture of adipocytes.^42–45 Sugihara et al. reported that 20% FCS showed the best adhesion and proliferation of DFAT.⁸

Several researchers could show that insulin had an antilipolytic effect on mature adipocytes in short-term culture in vitro, where they defined a wide range of concentrations as optimal.^{17–19,46,47} Generally, the glycerol release in medium 2 and 3 was lower compared with medium 1 on all analyzed days (except medium 3 on day 21), which led us to the assumption that insulin had a positive effect on the lipolytic activity of adipocytes.

An enhanced effect could be seen by the combination of supplements. It was shown that the combination of insulin and dexamethasone resulted in an increased incorporation of glucose into lipids compared with insulin alone.²⁰ Also, medium 2 and 3 in our study contained insulin and dexamethasone, which suggests a synergistic effect.

The effect of biotin on mature adipocyte behavior has not yet been examined intensively, but was used in several studies for the differentiation of preadipocytes into adipocytes. Kuri-Harcuch et al. have shown that 3T3 adipocytes did not accumulate triglycerides without biotin during differentiation.²¹ Others showed that lipid accumulation was increased using biotin-supplemented media.^22,48,49 Janke et al. cultured mature adipocytes for a short term while adding 4 μg mL⁻¹ biotin to the culture medium.⁵⁰ Medium 2 and 3 were also supplemented with biotin, of which we used 8 μg mL⁻¹ in medium 3, but the concentration for medium 2 was unknown. According to these data, a positive effect of biotin on the stability of mature adipocytes was seen.

The effect of pantothenate and dexamethasone on mature adipocytes is hardly known. Many investigations have been performed with a coapplication of the two supplements. Wang et al. reported a synergistic effect of dexamethasone with insulin on glucose incorporation and lipogenesis.²⁰ Furthermore, insulin and dexamethasone were responsible for an increased leptin release, but were not thought to be the main modulators.⁵¹ Also, biotin combined with panthothenate improved lipogenesis.²³

Mature adipocytes remained functional when cultured in an optimized media

To evaluate cell functionality, the release of glycerol and leptin was measured. We observed that cells cultured in medium 1 had a high release of glycerol and expressed leptin only in very small amounts. Cells cultured in medium 2 produced lower amounts of glycerol, but higher amounts of leptin at all analyzed days compared with medium 1. The glycerol release of cells in medium 3 stayed on an average level from day 7 to 21, whereas the leptin production was higher compared with medium 1 and 2.

Glycerol is thought to be a product of lipolysis.^39,52 Therefore, it is likely that the number of mature adipocytes in medium 1 decreased because of lipolytic activities, which was shown in a high release of glycerol from these cells and appearance of a fibroblast-like morphology. We could see that dedifferentiation also occurred in cells cultured in medium 2, but mainly to the level of multivacuolar adipocytes. The trigger to hold these cells in a multivacuolar state instead of completing the dedifferentiation process still needs to be determined. The glycerol release on day 1 in medium 3 was significantly higher compared to medium 1 and 2. This might be a result of elevated lipolysis during cell death. In this study, a still higher glycerol content was expected; we propose that initial cell death further resulted in disintegration of the cells and nonactive release of triacylglycerides from the lipid vacuoles, which cannot be measured with the method used here. In general, cells in medium 3 seemed to have a general turnover of fatty acids and glycerol because the total number of mature adipocytes decreased less compared with medium 1 and 2.

In general, we were able to detect a higher release of leptin from cells cultured in medium 2 and 3. It was previously described that the expression and secretion of leptin correlated proportionally with body fat mass and adipocyte size.^53–55 This supports our findings that cells cultured in medium 2 and 3 were able to keep a higher fat mass and adipocyte typical morphology compared with cells cultured in medium 1. Furthermore, insulin and dexamethasone are thought to be potent stimulators for the leptin expression.^{44,53,56–58} The effect of insulin is thought to rely on a higher glucose transport and metabolism and less on a direct insulin effect.⁵⁹ Another study proposed that insulin regulates the release of leptin from intracellular storages.⁵⁶ The higher leptin release from cells cultured in medium 2 or 3 could therefore be an additional result of insulin and dexamethasone in the culture medium.

On the contrary, a high leptin expression is thought to stimulate basal lipolysis.^60–62 Siegrist-Kaiser et al. have shown in sugar rats that leptin increased lipolysis in white adipose tissue dosis- and time-dependent. They concluded that leptin has an auto-/paracrine effect on adipocytes, which might be the weight- and fat-reducing activity of leptin.⁶¹ These data show that there might be a cascade of insulin–leptin–lipolysis signalling controlling fatty tissue storage and release. Because of the high production of leptin from cells cultured in medium 2 and 3, this might be another explanation for the elevated glycerol release in these media.

Conclusion

Mature adipocytes were cultured in two commercially available media compared with a control culture medium used in the literature and known to promote dedifferentiation. It was shown that the media supplemented with adipocyte-specific factors prevent cells from dedifferentiation and had a greater effect on cell viability, stability, and functionality compared with the nonsupplemented medium. For the composition and culture of an in vitro adipose tissue equivalent, cell survival and cell functionality are both of great importance. Since medium 2 favors high initial adipocyte survival, it is superior to medium 3. Mature adipocytes can now be used in a wide range of studies and also on a long-term basis to answer questions involving adipocyte metabolism and function. These cells might be highly valuable for the implementation of test systems to investigate cell–cell cross talk in monocultures and cocultures. In the future, mature adipocytes could be used for the composition of implants because of their promising properties concerning their high cell yield during isolation, easy handling, and stability under in vitro conditions.⁶³

Footnotes

Acknowledgments

The authors thank Dr. Ziegler (Klinik Charlottenhaus, Stuttgart, Germany) for the kind provision of human fatty tissue samples from plastic surgery. We gratefully acknowledge the European Commission for funding the ArtiVasc 3D project under the Seventh Framework Program (Grant Agreement No. 263416).

Disclosure Statement

No competing financial interests exist.

References

Cohen

J.C.

, Horton

J.D.

, and Hobbs

H.H.

Human fatty liver disease: old questions and new insights. Science, 332, 1519, 2011.

Park

, Euhus

D.M.

, and Scherer

P.E.

Paracrine and endocrine effects of adipose tissue on cancer development and progression. Endocr Rev, 32, 550, 2011.

Kredel

, Batra

, and Siegmund

Role of fat and adipokines in intestinal inflammation. Curr Opin Gastroenterol, 30, 559, 2014.

Yashima

, Isayama

, Tsujino

, Nagano

, Yamamoto

, Mizuno

, Yagioka

, Kawakubo

, Sasaki

, Kogure

, Nakai

, Hirano

, Sasahira

, Tada

, Kawabe

, Koike

, and Omata

A large volume of visceral adipose tissue leads to severe acute pancreatitis. J Gastroenterol, 46, 1213, 2011.

Zuk

P.A.

, Zhu

, Mizuno

, Huang

, Futrell

J.W.

, Katz

A.J.

, Benhaim

, Lorenz

H.P.

, and Hedrick

M.H.

Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Eng, 7, 211, 2001.

Rodbell

Localization of lipoprotein lipase in fat cells of rat adipose tissue. J Biol Chem, 239, 753, 1964.

Sugihara

, Yonemitsu

, Miyabara

, and Yun

Primary cultures of unilocular fat cells: characteristics of growth in vitro and changes in differentiation properties. Differentiation, 31, 42, 1986.

Sugihara

, Funatsumaru

, Yonemitsu

, Miyabara

, Toda

, and Hikichi

A simple culture method of fat cells from mature fat tissue fragments. J Lipid Res, 30, 1987, 1989.

Sugihara

, Yonemitsu

, Toda

, Miyabara

, Funatsumaru

, and Matsumoto

Unilocular fat cells in three-dimensional collagen gel matrix culture. J Lipid Res, 29, 691, 1988.

10.

Sugihara

, Yonemitsu

, Miyabara

, and Toda

Proliferation of unilocular fat cells in the primary culture. J Lipid Res, 28, 1038, 1987.

11.

Vierck

J.L.

, McNamara

J.P.

, and Dodson

M.V.

Proliferation and differentiation of progeny of ovine unilocular fat cells (adipofibroblasts). In Vitro Cell Dev Biol Anim, 32, 564, 1996.

12.

Matsumoto

, Kano

, Kondo

, Fukuda

, Iribe

, Tanaka

, Matsubara

, Sakuma

, Satomi

, Otaki

, Ryu

, and Mugishima

Mature adipocyte-derived dedifferentiated fat cells exhibit multilineage potential. J Cell Physiol, 215, 210, 2008.

13.

Sugihara

, Toda

, Yonemitsu

, and Watanabe

Effects of fat cells on keratinocytes and fibroblasts in a reconstructed rat skin model using collagen gel matrix culture. Br J Dermatol, 144, 244, 2001.

14.

Shigematsu

, Watanabe

, and Sugihara

Proliferation and differentiation of unilocular fat cells in the bone marrow. Cell Struct Funct, 24, 89, 1999.

15.

Nobusue

, and Kano

Establishment and characteristics of porcine preadipocyte cell lines derived from mature adipocytes. J Cell Biochem, 109, 542, 2010.

16.

Dominici

, Le Blanc

, Mueller

, Slaper-Cortenbach

, Marini

, Krause

, Deans

, Keating

, Prockop

D.J.

, and Horwitz

Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy, 8, 315, 2006.

17.

Gliemann

Insulin-like activity of dilute human serum assayed by an isolated adipose cell method. Diabetes, 14, 643, 1965.

18.

Rodbell

, and Jones

Metabolism of isolated fat cells. 3. The similar inhibitory action of phospholipase C (Clostridium perfringens alpha-toxin) and of insulin on lipolysis stimulated by lipolytic hormones and theophylline. J Biol Chem, 10, 140, 1966.

19.

Rodbell

The metabolism of isolated fat cells I. Effects of hormones on glucose metabolism and lipolysis. J Biol Chem, 239, 375, 1964.

20.

Wang

, Jones Voy

, Urs

, Kim

, Soltani-Bejnood

, Quigley

, Heo

Y.R.

, Standridge

, Andersen

, Dhar

, Joshi

, Wortman

, Taylor

J.W.

, Chun

, Leuze

, Claycombe

, Saxton

A.M.

, and Moustaid-Moussa

The human fatty acid synthase gene and de novo lipogenesis are coordinately regulated in human adipose tissue. J Nutr, 134, 1032, 2004.

21.

Kuri-Harcuch

, Wise

L.S.

, and Green

Interruption of the adipose conversion of 3T3 cells by biotin deficiency: differentiation without triglyceride accumulation. Cell, 14, 53, 1978.

22.

Ramírez-Zacarías

J.L.

, Castro-Muñozledo

, and Kuri-Harcuch

Quantitation of adipose conversion and triglycerides by staining intracytoplasmic lipids with Oil red O. Histochemistry, 97, 493, 1992.

23.

Flynn

, and Woodhouse

K.A.

Adipose tissue engineering with cells in engineered matrices. Organogenesis, 4, 228, 2008.

24.

Dimitriadis

, Mitrou

, Lambadiari

, Maratou

, and Raptis

S.A.

Insulin effects in muscle and adipose tissue. Diabetes Res Clin Pract, 93, S52, 2011.

25.

Gregoire

F.M.

, Smas

C.M.

, and Sul

H.S.

Understanding adipocyte differentiation. Physiol Rev, 78, 783, 1998.

26.

Zempleni

, and Mock

D.M.

Biotin biochemistry and human requirements. J Nutr Biochem, 10, 128, 1999.

27.

Wakil

S.J.

, and Abu-Elheiga

L.A.

Fatty acid metabolism: target for metabolic syndrome. J Lipid Res, 50, S138, 2009.

28.

Brown

G.M.

The metabolism of pantothenic acid. J Biol Chem, 234, 370, 1959.

29.

Kornberg

, and Pricer

W.E.

Enzymatic synthesis of the coenzyme A derivatives of long chain fatty acids. J Biol Chem, 204, 329, 1953.

30.

Zhang

H.H.

, Kumar

, Barnett

A.H.

, and Eggo

M.C.

Ceiling culture of mature human adipocytes: use in studies of adipocyte functions. J Endocrinol, 164, 119, 2000.

31.

Poloni

, Maurizi

, Leoni

, Serrani

, Mancini

, Frontini

, Zingaretti

M.C.

, Siquini

, Sarzani

, and Cinti

Human dedifferentiated adipocytes show similar properties to bone marrow-derived mesenchymal stem cells. Stem Cells, 30, 965, 2012.

32.

Tholpady

S.S.

, Aojanepong

, Llull

, Jeong

J.H.

, Mason

A.C.

, Futrell

J.W.

, Ogle

R.C.

, and Katz

A.J.

The cellular plasticity of human adipocytes. Ann Plast Surg, 54, 651, 2005.

33.

Watson

J.E.

, Patel

N.A.

, Carter

, Moor

, Patel

, Ghansah

, Mathur

, Murr

M.M.

, Bickford

, Gould

L.J.

, and Cooper

D.R.

Comparison of markers and functional attributes of human adipose-derived stem cells and dedifferentiated adipocyte cells from subcutaneous fat of an obese diabetic donor. Adv Wound Care, 3, 219, 2014.

34.

Ono

, Oki

, Bono

, and Kano

Gene expression profiling in multipotent DFAT cells derived from mature adipocytes. Biochem Biophys Res Commun, 407, 562, 2011.

35.

Jernås

, Palming

, Sjöholm

, Jennische

, Svensson

P.A.

, Gabrielsson

B.G.

, Levin

, Sjögren

, Rudemo

, Lystig

T.C.

, Carlsson

L.M.

, and Lönn

Separation of human adipocytes by size: hypertrophic fat cells display distinct gene expression. FASEB J, 20, 1540, 2006.

36.

Marcinkiewicz

, Gauthier

, Garcia

, and Brasaemle

D.L.

The phosphorylation of serine 492 of perilipin a directs lipid droplet fragmentation and dispersion. J Biol Chem, 281, 11901, 2006.

37.

Prins

J.B.

, Walker

N.I.

, Winterford

C.M.

, and Cameron

D.P.

Apoptosis of human adipocytes in vitro. Biochem Biophys Res Comm, 15, 500, 1994.

38.

Von Heimburg

, Hemmrich

, Zachariah

, Staiger

, and Pallua

Oxygen consumption in undifferentiated versus differentiated adipogenic mesenchymal precursor cells. Respir Physiol Neurobiol, 146, 107, 2005.

39.

Lafontan

Historical perspectives in fat cell biology: the fat cell as a model for the investigation of hormonal and metabolic pathways. Am J Physiol Cell Physiol, 302, C327, 2012.

40.

Large

, Peroni

, Letexier

, Ray

, and Beylot

Metabolism of lipids in human white adipocyte. Diabetes Metab, 30, 294, 2004.

41.

Justesen

, Pedersen

S.B.

, Stenderup

, and Kassem

Subcutaneous adipocytes can differentiate into bone-forming cells in vitro and in vivo. Tissue Eng, 10, 381, 2004.

42.

Engeli

, Gorzelniak

, Kreutz

, Runkel

, Distler

, and Sharma

A.M.

Co-expression of renin-angiotensin system genes in human adipose tissue. J Hypertens, 17, 555, 1999.

43.

Moustaïd

, Jones

B.H.

, and Taylor

J.W.

Insulin increases lipogenic enzyme activity in human adipocytes in primary culture. J Nutr, 126, 865, 1996.

44.

Bruun

J.M.

, Pedersen

S.B.

, Kristensen

, and Richelsen

Effects of pro-inflammatory cytokines and chemokines on leptin production in human adipose tissue in vitro. Mol Cell, 190, 91, 2002.

45.

Chazenbalk

, Bertolotto

, Heneidi

, Jumabay

, Trivax

, Aronowitz

, Yoshimura

, Simmons

C.F.

, Dumesic

D.A.

, and Azziz

Novel pathway of adipogenesis through cross-talk between adipose tissue macrophages, adipose stem cells and adipocytes: evidence of cell plasticity. PLoS One, 6, e17834, 2011.

46.

Claycombe

K.J.

, Jones

B.H.

, Standridge

M.K.

, Guo

, Chun

J.T.

, Taylor

J.W.

, and Moustaïd-Moussa

Insulin increases fatty acid synthase gene transcription in human adipocytes. Am J Physiol, 274, R1253, 1998.

47.

Perry

W.F.

, and Bowen

H.F.

Factors affecting the in vitro production of non-esterified fatty acid from adipose tissue. Can J Biochem Physiol, 40, 749, 1962.

48.

Xue

, and Zemel

M.B.

Relationship between human adipose tissue agouti and fatty acid synthase (FAS). J Nutr, 130, 2478, 2000.

49.

Sengenes

, Bouloumie

, Hauner

, Berlan

, Busse

, Lafontan

, et al. Involvement of a cGMP-dependent pathway in the natriuretic peptide-mediated hormone-sensitive lipase phosphorylation in human adipocytes. J Biol Chem, 278, 48617, 2003.

50.

Janke

, Engeli

, Gorzelniak

, Luft

F.C.

, and Sharma

A.M.

Mature adipocytes inhibit in vitro differentiation of human preadipocytes via angiotensin type 1 receptors. Diabetes, 51, 1699, 2002.

51.

Hardie

L.J.

, Guilhot

, and Trayhurn

Regulation of leptin production in cultured mature white adipocytes. Horm Metab Res, 28, 685, 1996.

52.

Burns

T.W.

, Langley

P.E.

, and Robison

G.A.

Site of free-fatty-acid inhibition of lipolysis by human adipocytes. Metabolism, 24, 265, 1975.

53.

Houseknecht

K.L.

, Baile

C.A.

, Matteri

R.L.

, and Spurlock

M.E.

The biology of leptin: a review. J Anim Sci, 76, 1405, 1998.

54.

Skurk

, Alberti-Huber

, Herder

, and Hauner

Relationship between adipocyte size and adipokine expression and secretion. J Clin Endocrinol Metab, 92, 1023, 2007.

55.

Klok

M.D.

, Jakobsdottir

, and Drent

M.L.

The role of leptin and ghrelin in the regulation of food intake and body weight in humans: a review. Obes Rev, 8, 21, 2007.

56.

Lee

M.J.

, Wang

, Ricci

M.R.

, Sullivan

, Russell

C.D.

, and Fried

S.K.

Acute and chronic regulation of leptin synthesis, storage, and secretion by insulin and dexamethasone in human adipose tissue. Am J Physiol Endocrinol Metab, 292, E858, 2007.

57.

Russell

C.D.

, Ricci

M.R.

, Brolin

R.E.

, Magill

, and Fried

S.K.

Regulation of the leptin content of obese human adipose tissue. Am J Physiol Endocrinol Metab, 280, E399, 2001.

58.

Kristensen

, Pedersen

S.B.

, and Richelsen

Regulation of leptin by steroid hormones in rat adipose tissue. Biochem Biophys Res Commun, 259, 624, 1999.

59.

Baile

C.A.

, Della-Fera

M.A.

, and Martin

R.J.

Regulation of metabolism and body fat mass by leptin. Annu Rev Nutr, 20, 105, 2000.

60.

Wang

M.Y.

, Lee

, and Unger

R.H.

Novel form of lipolysis induced by leptin. J Biol Chem, 274, 17541, 1999.

61.

Siegrist-Kaiser

C.A.

, Pauli

, Juge-Aubry

C.E.

, Boss

, Pernin

, Chin

W.W.

, Cusin

, Rohner-Jeanrenaud

, Burger

A.G.

, Zapf

, and Meier

C.A.

Direct effects of leptin on brown and white adipose tissue. J Clin Invest, 100, 2858, 1997.

62.

Frühbeck

, Aguado

, Gómez-Ambrosi

, and Martínez

J.A.

Lipolytic effect of in vivo leptin administration on adipocytes of lean and ob/ob mice, but not db/db mice. Biochem Biophys Res Commun, 250, 99, 1998.

63.

Huber

, Borchers

, Tovar

G.E.M.

, and Kluger

P.J.

Methacrylated gelatin and mature adipocytes are promising components for adipose tissue engineering. J Biomater Appl pii: 0885328215587450, 2015.