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First published online January 10, 2008
Stem Cells Vol. 26 No. 3 March 2008, pp. 682 -691
doi:10.1634/stemcells.2007-0738; www.StemCells.com
© 2008 AlphaMed Press

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TISSUE-SPECIFIC STEM CELLS

Functional Structure of Adipocytes Differentiated from Human Umbilical Cord Stroma-Derived Stem Cells

Sercin Karahuseyinoglua, Cetin Kocaefeb, Deniz Balcia, Esra Erdemlia, Alp Cana

aDepartment of Histology and Embryology, Ankara University School of Medicine, Ankara, Turkey;
bDepartment of Medical Biology, Hacettepe University School of Medicine, Ankara, Turkey

Key Words. Adipocyte • Differentiation • Human • Lipid • Mesenchymal stem cells • Umbilical cord

Correspondence: Correspondence: Alp Can, M.D., Department of Histology and Embryology, Ankara University School of Medicine, Sihhiye, 06100 Ankara, Turkey. Telephone: 90-312-3103010, ext. 369; Fax: 90-312-3106370; e-mail: alpcan{at}medicine.ankara.edu.tr

Received on September 6, 2007; accepted for publication on December 21, 2007.

First published online in STEM CELLS EXPRESS  January 10, 2008.

   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
It has been previously demonstrated that human umbilical cord stroma-derived stem cells (HUCSCs) are competent to differentiate into adipocytes. However, controversies have arisen as to whether HUCSCs can become mature adipocytes or not, and to what extent these cells can be induced in adipogenic pathway. Here, we extensively analyzed their adipogenic potency with a structural and functional approach by determining lipid formation dynamics in concordance to adipocyte-specific markers. During a 35-day period, HUCSCs respond to adipogenic induction, at which point 88% of cells exhibited multilocular lipid granules (LGs) having a mean diameter of 3 µm in round-shaped, F-actin-poor cells. Although the 1st week of induction did not generally display typical lipidogenic phenotypes, the degree of adipogenesis was dissected and confirmed by mRNA expressions of peroxisome proliferator-activated receptor {gamma}, C/EBP-β, sterol regulatory element-binding transcription factor 1, adipophilin, stearoyl-CoA desaturase, glycerol 3-phosphate dehydrogenase 1, LIPE, adiponectin, and leptin. All markers tested were found elevated in various amounts (3–70-fold) around day 7 and reached a plateau after day 14 or 21 (5–335-fold). Perilipin as a surface protein around the LGs was confined exclusively to the enlarging LGs. Conclusively, we propose that after the termination of proliferation, HUCSCs possess the biochemical and cellular machinery to successfully differentiate into maturing adipocytes under adipogenic conditions, and this feature will ultimately allow these fetus-derived stem cells to be used for various therapeutic or esthetic purposes.

Disclosure of potential conflicts of interest is found at the end of this article.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
One of the recent promising sources of fetus-derived stem cells is human umbilical cord stroma. After isolation from the cord matrix, human umbilical cord stroma-derived stem cells (HUCSCs) have been successfully differentiated into neuronal [1–4] and mesenchymal lineages [4, 5], including adipocytes [4, 6, 7]. The adipogenic induction in HUCSCs was simply verified by oil red O, a histological stain used to detect neutral lipids, mainly triglycerides, although in addition, a few markers of adipogenesis were confirmed by mRNA expression of adipocyte-specific genes, such as the nuclear hormone receptor, peroxisome proliferator-activated receptor {gamma} (PPAR{gamma}) [5, 8], adipsin [8], and plasminogen activator inhibitor-1 [4], as well as the enzyme lipoprotein lipase [9].

Adipogenesis is the process by which undifferentiated precursor cells differentiate into fat cells. Adipogenic differentiation is initiated by several transcription factors, including PPAR{gamma} and CCAAT/enhancer binding proteins (C/EBP{alpha}, β, {delta}) [10]. These master regulators of adipogenesis can be induced in vitro by a cocktail of a glucocorticoid (such as dexamethasone), a cyclooxygenase inhibitor (indomethacin), a phosphodiesterase inhibitor (3-isobutyl-1-methylxanthine [IBMX]), and insulin to generate committed adipocytes from preadipocytes or other pluripotent cells [11, 12]. During the course of differentiation, the committed cells exhibit a number of phenotypic changes, including biochemical, structural, and transcriptional reorganization. Initially, an adipogenically induced cell displays growth arrest and expresses a number of adipocyte-specific transcriptional regulators, such as Pref-1 (preadipocyte factor-1), C/EBP{alpha} and β, and PPAR{gamma}. Likewise, the sterol regulatory element-binding transcription factor 1 (SREBP1) mediates insulin action, such as lipogenesis and fatty acid and cholesterol synthesis [13, 14]. The expression of these adipogenic transcriptional factors allows these factors to stimulate themselves and each other [15], a process that is generally known as autocrine and paracrine regulatory mechanisms. Cellular transformation is accomplished by the expression of both structural proteins involved in cytoskeletal remodeling [16] and lipid granule (LG)-specific surface proteins, such as adipophilin and perilipin, followed by the biochemical adaptation of an adipocyte via expression of specific enzymes, such as glycerol 3-phosphate dehydrogenase 1 (GPD1), malic enzyme, hormone-sensitive lipase (LIPE), and fatty acid synthase [17, 18]. The final step in adipogenic differentiation is the accomplishment of endocrine function characterized by production of adipocyte-specific hormones, such as leptin, which is elevated during terminal differentiation of adipocytes [19], and adiponectin, which promotes adipocyte differentiation, insulin sensitivity, and lipid accumulation [20].

It has previously been documented that under appropriate conditions, mesenchymal stem cells (MSCs) [21], muscle precursors [22, 23], and embryonic stem cells [24] can undergo adipogenic commitment to form mature adipocytes. A growing body of evidence suggests that human umbilical cord stromal cells share a substantial number of common properties with MSCs [4, 7, 25]. However, no study has shown the spatiotemporal changes in lipid formation, phenotype, and expression of adipocyte-specific transcripts and hormones during adipogenesis of HUCSCs. Therefore, we aimed at focusing on the transcriptional changes in a series of adipocyte-specific structural proteins, enzymes, and hormones during in vitro adipogenic differentiation beginning at day 0 and extending to day 35. In addition to these experiments, we also addressed the structural changes, specifically the LG formation and the appearance of granule proteins in developing adipocytes in vitro. The immediate rises in the expressions of many specific markers were found to be temporally associated with a dramatic increase in the amount of lipid synthesis and storage as detected by direct fluorescence measurements. Interestingly, the expression levels of those mRNAs were found to reach a plateau after the 1st or 2nd week.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
Isolation and Expansion of HUCSCs
Umbilical cords of full-term deliveries (n = 26) were transferred to the laboratory under sterile conditions. Ethical approval was obtained from the Ankara University School of Medicine Ethical Review Board (approval no. 69-1780-2005). HUCSCs were isolated and expanded as described elsewhere [4]. After a few passages, cells were used for adipogenic differentiation. Fluorescence-activated cell sorting was applied to cell suspensions for further isolation of HUCSCs using certain MSC markers such as CD105, CD44, and CD73, all conjugated with either fluorescein isothiocyanate (FITC) or phycoerythrin.

Adipogenic Differentiation
Subconfluent (80%–90%) HUCSCs from passages 3–6 grown on 35-mm culture dishes and glass coverslips were induced for adipogenic differentiation by an administration of 1 µM dexamethasone, 500 µM IBMX, 60 µM indomethacin, and 5 µg/ml insulin in Dulbecco's modified Eagle's medium (DMEM)-low glucose (low glucose, 1 g/l) supplemented with 10% fetal bovine serum (FBS) [26]. Noninduced cells were cultured in DMEM-Ham's F-12 (1:1 [vol/vol]) culture medium supplemented with 10% (vol/vol) FBS. Noninduced and induced cells at days 1, 3, 5, 7, 14, 21, 28, and 35 were used for either RNA extraction or in situ detection of LG formation (described below). All chemicals were purchased from Sigma-Aldrich (St. Louis, http://www.sigmaaldrich.com) unless stated otherwise.

Oil Red O Staining, Immunocytochemistry, and Morphometry
Ten percent buffered formalin-fixed cells (30 minutes at room temperature) were stained by 1% (wt/vol) oil red O in 60% isopropanol for 40 minutes. Guinea pig polyclonal anti-perilipin antibody (Progen, Heidelberg, Germany, http://www.progen.de) (1:50; 90 minutes at 37°C) was applied, followed by incubation with goat anti-guinea pig IgG conjugated with FITC (1:200; 60 minutes at 37°C), for perilipin labeling. Anti-human nuclei mouse monoclonal antibody (Chemicon, Temecula, CA, http://www.chemicon.com) (1:200; 60 minutes at 37°C) was applied, followed by incubation with Cy5 goat anti-mouse IgG (Zymed Laboratories Inc., San Francisco, http://www.bioresearchonline.com/content/homepage/), for nuclear labeling. FITC-phalloidin labeling (specific to F-actin; 20 µg/ml; 20 minutes at 37°C) was applied to observe F-actin filaments in induced and control cells. All immunofluorescent antibody labelings and dyes were examined using a Carl Zeiss LSM 510 Meta confocal laser scanning microscope (Carl Zeiss, Jena, Germany, http://www.zeiss.com) equipped with 488-nm argon ion and 543-nm and 633-nm helium neon lasers. Three-dimensional images obtained by x63 plan-apo objective (x1.5 zoom) were reconstructed by consecutive optical sections of various thicknesses (0.25–0.38 µm).

Cellular morphometry was assessed by direct measurement of oil red O-stained LGs, as they finely emit a red signal using 543-nm laser. Line measurements and gray scale-based thresholding (≥200 was considered positive staining) tools in the LSM-510 software (version 3.2) were used to semiautomatically calculate the cell size, granule size, and total area of LGs. The detection parameters, such as laser intensity, amplifier offset and gain, and pinhole diameter, were fixed and kept at the same values for all specimens.

Determination of Cell Viability
After certain time points during in vitro adipogenesis, a certain number of cells tended to detach from the substrate but remained within the culture medium until they were fixed and washed away. To identify whether these round cells were dead or alive, we performed a propidium iodide (PI) dye exclusion assay. PI (2 µg/ml) (Invitrogen, Carlsbad, CA, http://www.invitrogen.com) in culture medium was applied for 30 minutes under culture conditions and was scoped under an Olympus IX-71 inverted microscope (Olympus, Tokyo, http://www.olympus-global.com) equipped with relief contrast and fluorescence imaging systems. Dual images were captured by Olympus DP-30BW cooled charge-coupled device camera (Olympus) using a x40 plan-apo objective and DP Manager (Olympus) software.

Electron Microscopy
On successive days of culture, adipogenically induced and noninduced control cells grown on 75-cm2 tissue culture flasks were detached from the substrate by the application of 0.05% trypsin/0.02% EDTA in phosphate-buffered saline (Biochrom AG, Berlin, http://www.biochrom.de) for 5 minutes at 37°C. Suspended cells were centrifuged at 120g for 10 minutes. The cell pellets obtained were processed as described elsewhere [27]. Grids were dried and examined under a Leo 906 E, 100-kV transmission electron microscope (Carl Zeiss).

RNA Isolation and Reverse Transcription-Polymerase Chain Reaction
All samples were treated together to avoid any experimental bias resulting from handling and extraction. Total RNA from the cell cultures were harvested as described previously [28]. Quality control was ensured by denaturing agarose gel electrophoresis and optical density measurements at 260/320. One microgram of total RNA was reverse-transcribed into cDNA using oligo(dT) primers and Improm II reverse transcriptase (Promega, Madison, WI, http://www.promega.com). Equal amount of cDNA was used for the real-time amplification of the target genes using Jumpstart SYBR Green mix (Sigma-Aldrich) according to the manufacturer's recommendations on a Rotorgene 6000 (Corbett Life Science, Sydney, New South Wales, Australia, www.corbettlifescience.com) real-time quantitative polymerase chain reaction instrument. The primer pairs and the reaction conditions were designed to include at least one intron to avoid misamplification of any contaminated DNA. The sequences of the primer pairs and the reaction conditions are available upon request by e-mail. The geometric mean of the expressions of transcription factor II D (TFIID) and hypoxanthine-guanine phosphoribosyl transferase (HPRT) have been used to normalize the expression of the adipogenic differentiation-related transcripts, as suggested by Vandesompele et al. [29].

Statistical Analyses
Cell morphometry measurements (1,500 measurements from each sample) and triplicate gene expression results were directly transferred to Excel (Microsoft, Redmond, WA, http://www.microsoft.com). Mean, SE, and SD were calculated, and the significance between groups was analyzed by one-way analysis of variance and pairwise t test using the SPSS software package (SPSS, Chicago, http://www.spss.com).


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
Characteristics of Lipid Synthesis and Storage in Adipogenically Induced HUCSCs
HUCSCs at passages 3–6 were used in the adipogenic experiments, in which 80%–90% subconfluence was found to be optimum for the differentiation of induced cells in the following 5-week period. Although adipogenic differentiation potency of HUCSCs was examined for almost 5 weeks, the onset of phenotypic changes was noticed as early as the first days of induction. Starting from day 3 of induction, an increase was noted in LGs, both in number and size, in induced cells compared with controls. After 48–72 hours following treatment, specific intracytoplasmic LGs that significantly grew further in the following days appeared, particularly in the central cytoplasm (Fig. 1, day 3). At this stage, cell shape began to transform into ovoid/round morphology. In the following days, the average size of LGs gradually increased, and they showed a tendency to accumulate in the cell periphery (Fig. 1, days 5 and 7). At around day 7, homogeneously scattered LGs also appeared in noninduced control cells, albeit in low numbers and small sizes (Fig. 1, control day 7). Lipid content of the cells after the 1st week of induction was characterized by remarkable location of granules at the cell periphery and did seem to combine to form larger granules (Fig. 1, day 14). Along with the enlargement in cell volume detected, particularly in the 3rd and 4th weeks, larger granules accumulated and invaded the entire cytoplasm, reaching a state after which no more granules were synthesized (Fig. 1, days 28 and 35). Little or no change was noted in control cells at these stages (Fig. 1, day 28 control).


Figure 1
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  		Figure 1. Lipid granule (LG) formation during the 1st week of induction and the following 4 weeks. A significant increase in LG number and size in induced cells as compared with controls was noticed from day 1 until day 35 of induction, as indicated by the appearance of oil red O-positive LGs. Specific intracytoplasmic LGs appeared in the central cytoplasm during the very early days of induction (day 3), while cells attained ovoid-round morphology. The number and average size of LGs gradually increased, as they tended to accumulate in the cell periphery (after day 5). Peripheral location of granules was prominent by the end of the 2nd week (day 14), followed by a significant enlargement of the cell volume detected during the 3rd and 4th weeks (days 28 and 35). Noninduced control cells showed few small LGs during the 1st week (days 3 and 7 control). Little or no change was noted in control cells in the following weeks (days 14 and 28). F-actin filaments are shown in green and nuclei in blue. Scale bar = 20 µm.

 
The ratio of adipogenically responsive cells to nonresponsive cells was assessed simply by counting 240 cells from each sample, and the results are summarized in Figure 2A. A steady increase was noticed in the number of responsive cells in concordance with the culture days, where almost 88% (± 9.1%) of cells were found to contain a substantial amount of adipogenically induced LGs around day 35 of culture.


Figure 2
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  		Figure 2. Results of morphometric analyses of adipogenically-induced HUSCSs throughout the 35-day culturing period. (A): The percentage of cell population positively responding to in vitro adipogenic induction from day 1 to day 35 cultures gradually increased and reached 88%, meaning that 88% of induced cells were able to differentiate into adipocytes. Control data display zero values. (B, C): Morphometric measurements on lipid granules (LGs) in induced (bullet) and noninduced ({blacksquare}) cells during the days of the 1st week and the following 4 weeks of differentiation. (B): Increase in the granule diameter. (C): Increase in the total area of LGs per cell. Note the sudden increase in granule diameter around days 3–5 and 14–21. SD bars indicate that there was a wide variation in the sizes of granules, especially in later weeks, whereas control samples did not show any significant change in granule size and cytoplasmic occupation. *, p < .05.

 
To quantify the changes in granule size and the total number of granules per cell, a series of morphometric measurements were applied to the oil red O-stained, fluorescently illuminated cell cultures starting from 24 hours after induction. As illustrated in Figure 2B, LG size was measured as 1.01 µm (±0.26) at day 1 in induced cells and 0.59 µm (±0.12) in control cells. A slight increase (1.30 ± 0.56 µm) was noted 2 days later (day 3) in induced cells. The growth rate of granule size was amplified in subsequent days and reached 1.82 µm (±0.60) and 1.85 µm (±1.19) in days 5 and 7, respectively. The largest granules were measured as 3.66 µm and 6.27 µm around these days. Although the average granule size was slightly elevated during the 1st week of adipogenic induction, larger LGs occasionally began to form by the unification of pre-existing granules, especially when they translocated to the cell periphery for storage. Interestingly, an increase was also noted in control cells, which reached an average size of 1.01 µm (±0.32) at day 3 but then remained relatively constant during the 1st week (0.95–0.99 µm). More importantly, granules in control cells always appeared in the central cytoplasm and never seemed to combine to form larger granules (Fig. 1).

The increase in the LG size continued in the following days and was measured at 1.94 µm (±1.19) on day 14. As the area and size of granules increased, the preferential location of granules began to disappear, since enlarging granules invaded almost the entire cytoplasm (Fig. 1, day 28), when a sudden rise was noted for both the average (2.62 ± 1.55 µm) and the maximum (7.9 µm) size of the granules. The rate of increase in granule diameter continued to rise, and diameter reached 3.03 µm (±1.59) on day 28 and 3.04 µm (±1.30) on day 35. Constitutive LGs in noninduced control cells ranged between 0.95 and 1.02 µm and never exceeded 1.24 µm during the entire culture period.

As microscopic observations demonstrated that the increase in the size of LGs may be coupled to the increasing number of granules, we calculated the total area of LGs within a cell relative the total area of a cell from three-dimensional reconstructed confocal image stacks (Fig. 2C). At day 1, the total area of granules was calculated as 110 µm2 (±26.4) and 32 µm2 (±4.7) in induced and noninduced cells, respectively. After the ratiometric calculation, 3% and 1% of the total cell area were determined to be occupied with LGs at day 1 in induced and control cells, respectively (Fig. 2C). Total cytoplasmic occupancy of LGs was recorded as 4.5% at day 3, 5.3% at day 5, 7.8% at day 7, 14.3% at day 14, 18.4% at day 21, 19.1% at day 28, and 20.4% at day 35 (Fig. 2C). In controls, LGs occupied only 3.2%–4.1% of the cytoplasm after the 1st week of culture (Fig. 2C).

For viewing the course of LG formation, we performed transmission electron microscopy, which also allowed us to observe the cytoplasmic organelle arrangement associated with lipid synthesis. As also demonstrated in Figure 1, after the synthesis of LGs in the juxtanuclear cytoplasm (Fig. 3A), where the cell's synthetic organelles were located, LGs translocated to the cell periphery, combined (Fig. 3B, 3C), and were stored for longer periods. They were characterized as highly osmiophilic, multiple, round, and smooth-contoured granules. Although the LGs displayed various degrees of electron opacity, they were predominantly composed of poly-unsaturated lipids, particularly larger ones. Supporting evidence was the existence of numerous long mitochondria with complex cristae and an electron-dense matrix, which are characteristic features of an adipocyte (Fig. 3A, 3B). Neighboring rough and smooth endoplasmic reticulum membranes were also localized in close proximity to LGs, a sign of high metabolic and synthetic activity (Fig. 3A, 3B).


Figure 3
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  		Figure 3. Location and formation of lipid granules (LGs) during adipogenic differentiation as shown by transmission electron microscopy (A, B) and high-power three-dimensional confocal image stack (C). (A): Newly formed, relatively smaller LGs at day 3 first appeared in the juxtanuclear cytoplasm in close contact with granular (arrowheads) and smooth (arrows) endoplasmic reticulum and with mitochondria (asterisks) having abundant cristae. (B): Immediately after synthesis, they translocated to the peripheral cytoplasm, where they combined to form larger granules, reaching an average diameter of 2.62 µm at day 21. Note the onset of fusion of formed granules with neighboring ones. (C): The majority of the oil red O-stained large granules were stored at the peripheral cytoplasm after day 21. Scale bars = 1 µm (A, B), 10 µm (C). Abbreviation: n, nucleus.

 
Perilipin, a phosphoprotein that is selectively expressed in adipocytes and steroidogenic cells [30], was found exclusively at the periphery of enlarging granules as patchy rims beginning from the early stages of differentiation, whereas smaller granules did not contain any perilipin staining (Fig. 4A). In later stages, perilipin intensity increased around the periphery of large granules (Fig. 4B).


Figure 4
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  		Figure 4. Perilipin (green signal) in 14-day (A) and 35-day (B) adipogenically induced oil red O-stained (red signal) cells. Perilipin staining was restricted to the periphery of enlarging granules as patchy rims (arrowheads). The intensity and the distribution of perilipin staining increased in parallel to the increasing number of large lipid granules. Note that smaller granules were totally devoid of perilipin staining. Scale bar = 10 µm.

 
During the expansion of HUCSCs, cells typically displayed a fibroblastic phenotype bearing broader cytoplasms decorated with intense F-actin filaments, as clearly defined in earlier studies. When cell density was allowed to reach to confluence, cells began to inhibit proliferation, possibly by a mechanism known as contact inhibition of growth. Proliferation of cells, as indicated by mitotic indices, was almost aborted in induced cells, whereas a steady proliferation rate was noticed in controls (data not shown). Upon transfer of cells to adipogenic media, they slowly but drastically changed their shape from spindle to ovoid or round (Figs. 1, 5A, 5B). In parallel to this shape transition, adipogenically induced HUCSCs altered their F-actin filament decoration of stress fibers. As they extended their broad and flat cytoplasm along the substrate, they exhibited fine arrays of F-actin filaments, particularly in the earlier stages of adipogenesis (Fig. 5A). Interestingly, stress fibers began to diminish and only remained in little arrays around the cell periphery (Fig. 5B) when cells converted into round shape, which coincided with the increase in the number and size of LGs. In addition, beginning at days 18–20, some round cells were found to be loosely attached to the substrate (Fig. 5C). They were filled with round LGs, which eventually give the cell a perfect round shape, but not any attachment surface. Therefore, we wanted to know whether these cells were alive or were dead after they detached from the culture surface. PI dye exclusion assay showed no sign of death (Fig. 5D), although they possessed smaller nuclei than the attached cells.


Figure 5
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  		Figure 5. F-actin expression and detachment of cells as they become mature toward the end of induction period. F-actin decorated stress fibers were typically organized in flat and well-anchored human umbilical cord stroma-derived cells at day 7 (A), whereas around days 20–30 (B), as the cells dramatically change their shape from spindle to cuboid or round, only a few F-actin filaments were detected along the cell bodies. (C): Occasionally, some cells were found to be detached from the substrate (arrows) beginning from days 18–20 (Differential interference contrast image); interestingly, however, they were found to be alive, as they possess intact cell membrane that was impermeant to propidium iodide (D). Scale bars = 10 µm (A, B) and 20 µm (C, D).

 
Time-Course Change in Adipogenic Differentiation Markers
The mRNA levels of PPAR{gamma}, C/EBPβ, SREBP1, adipophilin, stearoyl-CoA desaturase 1 (SCD1), GPD1, LIPE, adiponectin, and leptin were assessed, whereas the expression of TFIID and HPRT was used to normalize the expression of the above genes. The relative expression of the transcripts at various time points was further normalized to day 0 control values to represent a relative fold change of expression. As illustrated in Figure 6, the expression of PPAR{gamma} started to show significant values at day 2 (Fig. 6A) and then reached 27.7-fold at day 7, upon induction. The expression of C/EBPβ increased more than 13-fold during the same period and reached a steady state around day 14 (Fig. 6B). Similarly, SREBP1 exhibited a dramatic increase (16-fold) at day 5 and was then steadily induced more than 250-fold after day 14 (Fig. 5C). Expression of adipophilin gradually increased during the 1st week and exceeded 50-fold after 21 days (Fig. 6D). During the observed period, the expressions of three enzymes, SCD1, GPD1, and LIPE, were found elevated up to 34.4-, 6.7-, and 140-fold, respectively (Fig. 6E–6G). All three of these tested parameters almost reached their peak levels between day 7 and day 21 and remained relatively constant during the rest of induction period. Adiponectin and leptin, as adipocyte-specific hormones, significantly increased a few days later than the above mRNAs. Their expression remained relatively low (3.3- and 8.8-fold, respectively) during the 1st week, whereas their expression reached significantly higher levels especially after the 2nd week (Fig. 6H, 6I). The expression of leptin displayed a steady pattern following the 2nd week, with an upregulation of more than 20-fold (Fig. 6I). The relative expressions of the above genes did not display any significant changes in the time-matched control samples in which adipogenic differentiation was not induced.


Figure 6
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  		Figure 6. The mRNA levels of PPAR{gamma} (A), C/EBPβ (B), SREBP1 (C), adipophilin (D), SCD1 (E), GPD1 (F), LIPE (G), adiponectin (H), and leptin (G) were assessed following a reverse-transcriptase reaction by semiquantitative polymerase chain reaction in induced (bullet) and noninduced ({blacksquare}) samples (n = 12 for each group). The relative expression of the transcripts at various time points (days) was further normalized to day 0 control values to represent a relative fold change of expression. *, p < .05. Abbreviations: GPD1, glycerol 3-phosphate dehydrogenase 1; LIPE, hormone-sensitive lipase; PPAR{gamma}, peroxisome proliferator-activated receptor {gamma}; SCD1, stearoyl-CoA desaturase; SREBP1, sterol regulatory element-binding transcription factor 1.

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosure of Potential...
 Acknowledgments
 References
 
MSCs obtained from human bone marrow [31–33], human placenta [34, 35], umbilical cord blood [36, 37], umbilical cord matrix [4, 5, 8, 9, 38], and adipose tissue itself [39] were all shown to be able to differentiate into adipocytes successfully. In addition, preadipocytes and human MSCs can also be successfully stimulated within some scaffolds to proliferate and differentiate into mature adipocytes [40, 41]. However, the dynamics of lipid accumulation with regard to the changes in structural and biochemical parameters have not previously been assessed in MSC-like cells. One of the major findings of the present study was observing the time-phenotype parity during in vitro adipogenesis. With the exception of adipogenic hormones such as adiponectin and leptin, all markers tested exhibited a sudden rise during the 1st week of induction, although phenotypic alterations such as LG size and responsiveness of cells to adipogenic induction did not roughly indicate a clear difference between induced and control samples. Precise microscopic observations and measurements, however, revealed that adipogenic cells possessed some peculiar features, such as the translocation and unification of synthesized granules, which were never observed in noninduced cells. Therefore, induced cells were not practically considered mature adipocytes by the end of the 1st week, even though expression profiles displayed significantly high values. So, further treatment of the cells up to 3–5 weeks was required to observe whether any further increase, especially in the lipid content of the cells, was noted. At the end of 5th week, both structural and transcriptional data implied that a steady level was reached after which no more alterations would be noteworthy. As a result, it is possible to assume that during adipogenic differentiation of these particular human stem cells, critical phenotypic features follow the essential transcriptional changes and require a culturing period where the majority of cells respond positively to induction, as evidenced by the appearance of many fundamental changes around days 28–35 associated with the significant rise in production of adipocyte-specific hormones.

The time point at which the adipogenic induction should be terminated is a matter of debate. Like the data in many other stem cell trials, the present data indicate that HUCSCs require at least 4–5 weeks of induction to fulfill the criteria to be considered mature adipocytes. Apparently, the duration of in vitro differentiation is essentially dependent on the nature of the cells studied and could not be estimated without any detailed analysis. In general, in vitro adipogenic differentiation conditions could not be easily attributed to in vivo environments. However, given that mature adipocytes derived from clonal preadipocytes in vitro exhibited most of the morphological and biochemical characteristics of normal adipocytes in vivo and shared sensitivity similar to that of the hormones responsible for influencing the growth of adipose tissue [42], HUCSCs may behave in the same way, as they require some well-defined conditions for a successful in vitro transformation following expansion. In brief, in vivo trials are needed to further evaluate their adipogenic potencies where many uncontrolled signals spray the cells for relatively longer periods.

Adipogenesis is accompanied by dramatic alterations in cell shape, as well as by molecular changes that lead to dramatic increases in the cell's ability to synthesize lipids [43, 44]. Once triggered, transcription factors act cooperatively and sequentially to promote the necessary stages that drive the adipogenic program [45]. Depending on the proliferation characteristics summarized in this study, we assumed that at the beginning of adipogenic induction, the addition of specific stimulants and supplements to the cell culture media allowed cells to reach a state where proliferation was terminated and cells were prepared to respond to adipogenic transformation. This terminal differentiation is known to be closely associated with an increase in the cell's ability to engage in de novo lipogenesis [44].

One of the most prominent phenotypic indicators of adipogenesis is the change in cell shape from fibroblastic to ovoid-round, which was typical in our cells, particularly after day 7. However, specific LGs were formed in elongated cytoplasms before the cells had become round. When these data are taken together, it is reasonable to consider that a change in cell shape may not be correlated with the onset of lipid accumulation. Supporting evidence for this was the fact that lipid-synthesizing cells, during days 3–14, were finely decorated with F-actin-positive stress fibers that give cells a relatively cylindrical morphology. In addition, many round cells seen in days 7–21 were devoid of specific LGs, which means that there is no correlation between the shape and the degree of responsiveness. On the other hand, the increase in cell size was found to be tightly associated with the degree of formation of LGs, as matured adipocytes were the largest in our experiments. In addition to an increase in cell size, adipogenic response of HUCSCs increased during the 35-day culturing period, starting from 30% at day 1 and reaching almost 90% of cells at day 35, a finding that correlates with human bone marrow [32–34, 37], umbilical cord blood [36], or matrix-derived MSCs [8]. Although Sekiya et al. [32] found that 30% and 60% of cell colonies were filled with LGs on days 14 and 21, respectively, in human bone marrow MSCs, Kim et al. [8] reported that only 35% of HUCSCs showed lipid inclusions. More striking results were reported by Janderová et al. [33], who found that only 20% of human bone marrow MSCs were positively stimulated, whereas the ratio of responsive cells increased up to 80% when the cells were incubated with rabbit serum or PPAR{gamma} agonists, such as thiazolidinediones (TZD). Similar variations were not only restricted to cell responsiveness but also with regard to the cell morphology, granule size, and number that were reported by different groups. For instance, DMEM with low (1 g/l) or high (4.5 g/l) glucose in the adipogenic induction media used by different laboratories seems to cause various results in the onset of LG formation. Whereas Wang et al. [5] found LGs to appear at day 7 with low glucose, Yen et al. [34] and Erices et al. [37] found them on days 10 and 15, respectively, when they used 3.5 g/l glucose. It has been well established that a cocktail of dexamethasone, insulin, and IBMX, each of which regulates the expression of adipocyte-specific genes or proteins, is essential for adipogenic differentiation [45]. Indometacin, a nonselective inhibitor of cyclooxygenase, is used especially when human MSCs were induced to adipocytes [21, 46]. In addition, supplementary hormones and reagents were added to the induction medium of fibroblasts from the stroma-vascular fraction of human adipose tissue, such as triiodothyronine (T3), thyroxine (T4) [47], cortisol, transferrin, and biotin [39, 48]. Therefore, depending on the culture system and the supplementary hormones, the process of differentiation, as well as minor differences in cell morphology, is noticed, and more controlled studies may be needed to evaluate the specific outcomes for a specific cell type.

The intracytoplasmic distribution of LGs in our cells showed a spatiotemporal change during adipogenesis. Newly synthesized LGs were closely associated with the endoplasmic reticulum and mitochondria in the juxtanuclear cytoplasm, as also reported for 3T3-L1 cells [49]. Although they maintained the central to peripheral route during synthesis, maturing cells tended to translocate the granules to the cell periphery where they combined to form larger granules. Nevertheless, studies examining the adipogenic potential of human MSCs revealed that LGs do not fuse to form a unilocular morphology [5, 32, 36], with an exception recorded by Mackay et al. [31], who showed formation of unilocular phenotypes both in bone marrow-derived MSCs and, more prominently, in medullary adipocytes, especially in cultures spanning more than 21 days. We did not note any sign of granule exocytosis or release in any time point studied. However, toward the end of five-week induction period, the presence of a few detached live cells with lipid-laden cytoplasm indicated the possibility that some of the fully mature cells may have been missed during the culturing period because of their extremely round shape, as also reported by Mackay et al. [31]. This assumption needs to be further analyzed, and if it is proven that some of the cells do become fully mature before the majority of the cells and detach from the substrate, then adipogenically induced cells or at least HUCSCs would definitely require three-dimensional culture systems rather than monolayer cultures. Given that changes in adipogenic differentiation in cell shape are paralleled by dramatic changes in the level and type of cytoskeletal components, such as actin and tubulin [16], it was not uncommon to note that these detached cells did not contain F-actin filaments. Since the loss of stress fibers parallels the conversion of the highly spread and flattened cells to spherical adipocytes [16] and correlates with an 80% reduction in cellular actin content [50], it is noteworthy to postulate that as the cells become mature, they downregulate their F-actin expression, an adaptation mechanism by which growing size and number of spherical LGs are stored in an unpolarized fat cell.

C/EBP{alpha}, C/EBPβ, and PPAR{gamma} are among the central regulators of adipogenesis by transactivation of numerous adipocyte-specific genes. Their expression is considered to initiate adipogenesis [10, 51], which was confirmed by the findings of others [5, 31–33] and our findings since the expressions of C/EBPβ and PPAR{gamma} increased up to 10-fold by the 5th and 3rd days of induction, respectively, indicating that they play crucial roles in the transcriptional cascade of adipogenesis. Likewise, lying downstream from the above regulators, SREBP1 is also responsible for modulating the insulin action in adipose tissue [13]; therefore, severalfold increase in its expression starting at day 5 was not an uncommon finding.

One of the important structural proteins coating the LGs is adipophilin, the adipose-differentiation-related protein and an early marker of adipogenic differentiation [52]. Expression of adipophilin was tightly correlated to the accumulation of LGs in our observations; it was upregulated 25-fold as early as day 3 of differentiation and reached its peak levels (40–50-fold) by days 5–7, suggesting that the majority of the LGs appear and increase in number in the early stages, after which larger LGs are formed by the unification of existing ones. Perilipin, on the other hand, is likely to play a role in lipolysis. It is known that adipophilin coats the small LGs in preadipocyte, and when the cells are differentiated into mature adipocytes, adipophilin disappears as LGs expand, being partially or completely displaced by perilipin [52]. Although we did not show in situ localization of adipophilin, de novo synthesis of preferentially located perilipin supports the previous findings. Further evaluation of the adipophilin-perilipin transition is needed to clarify the roles in stem cell-derived adipocytes in human.

Cytosolic GPD1 is an important enzyme in carbohydrate metabolism and is known as a biochemical marker for adipogenesis [53], whereas SCD1 is a key catalyzer of the rate-limiting step in the synthesis of the polyunsaturated fatty acids. The expression of GPD1 reached significant levels at day 3 and remained stable around a sixfold upregulation level throughout our observations. Likewise, the expression pattern of SCD1 exhibited a similar curve, but the expression levels were quite a bit higher. These observations clearly reflect the degree of success in the biochemical adoption of a maturing adipocyte.

The endocrine function of the adipocytes is of particular interest to obesity and diabetes research, as reviewed by Fasshauer and Paschke [54]. Here, we investigated the expressions of the adipocytic hormones leptin and adiponectin, both of which are known to be synthesized and released by adipocytes. Leptin is the product of the OB gene and is predominantly but not exclusively synthesized by the WAT of the subcutaneous fat. Its plasma levels correlate with body fat mass and are known to exhibit a central anorexic effect. Adiponectin, also known as the adipocyte complement-related protein of 30 kDa (Acrp30), has been shown to be positively induced by PPAR{gamma} agonists such as TZDs, which also induce adiponectin up to 50-fold in 48 hours in preadipocytes [55]. We showed the upregulation of both of these hormones in HUCSCs following adipogenic induction; leptin reached a significant level by day 7, and then its transcription reached a steady state of approximately 20-fold in the following weeks. Adiponectin exhibited a similar but lower level of induction, reaching up to threefold at day 5, followed by its peak levels after day 14. These results indicate that the HUCSCs adopt endocrine functions more slowly than the preadipocytes, but they successfully exhibit the endocrine properties that are specific to preadipocytes and mature adipocytes.

It is important to note that all of the above parameters showing a linear increase, especially during the first 2 weeks of induction, followed by relatively constant levels may also be due to the increasing number of responsive cells rather than the increased expressions of the given factors. Whether the number of responsive cells and the measured amount of these factors are related or not, it is evident that HUCSCs should be cultured at least 4–5 weeks under adipogenic conditions to be considered fully mature adipocytes.

Although the mechanism of adipocyte transformation in human mesenchymal tissues is still largely unknown, adipocyte precursors are undifferentiated fibroblasts that can be stimulated to form adipocytes. Given that HUCSCs were historically considered "unusual fibroblasts" [56], which were then demonstrated to possess many cellular features of myofibroblasts [57–59] and, more recently, some embryonic stem cell properties [60], they are likely to be a true source of stroma-resident stem cells, which could in turn be differentiated into many cell types upon activation. Our previous results comparing the in situ HUCSCs with in vitro isolated undifferentiated HUCSCs [4] suggested that these cells reside in the umbilical cord in a semidifferentiated state and maintain their differentiation potency until term. Therefore, after isolation and induction with specific stimulants, they immediately give rise to targeted cell types with a high proliferation rate.

Although recent articles emphasize the multidifferentiation potency and thus the potential clinical use of these cells, no report has been published so far dissecting the plasticity of HUCSCs during the course of differentiation. The present study, therefore, not only serves to demonstrate the detailed steps of adipogenic potency of HUCSCs but also reveals many structural and functional aspects of adipogenic transformation in human MSC-like cells, which might be attributed to the stem cells from other sources. We believe that our results could also facilitate an understanding of the in vitro plastic behavior of human stem cells while transforming into an adipocyte. On the basis of our published and unpublished results, we propose here a differentiation model for a HUCSC from its "niche" to a mature adipocyte (Fig. 7).


Figure 7
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  		Figure 7. A hypothetical model of in vitro adipogenic differentiation of the MSC-like stem cells that reside in human umbilical cord stroma. Two types of cells reside in the Wharton's jelly: PVCs and IVCs [4]. Within their stromal niche, these myofibroblastic cells normally show no sign of mitosis. However, both cell types display different proliferation and differentiation potencies [25] when they are placed in cell culture. They rapidly enter the cell cycle and replicate every 2–4 days (doubling time), displaying multiform cell populations that might be due to the mixture of PVCs and IVCs [4]. One subpopulation of cells later disappears because of possible premature senescence caused by cell fusion. The remaining cells display fibroblastic morphology and maintain their proliferation capacity, albeit at a gradually reduced rate, and extend their life span up to 50–60 division cycles until they reach replicative senescence without undergoing spontaneous differentiation. When these proliferating cells are induced during any stage of in vitro differentiation, they gradually exit the cell cycle and enter G0, a prerequisite state from which adipogenic transformations can begin. This process explains the increasing rate of adipocyte maturation seen in culture during the course of differentiation (gray arrow). Adipogenic differentiation is exemplified in this figure, which shows that MSC-like human umbilical cord stroma-derived cells (HUCSCs) require a promoting signal. This signal is achieved by a cocktail of specific hormones and reagents (Materials and Methods) that rapidly transforms HUCSCs into adipoblasts, in which specific LGs begin to appear. In a few days, adipoblasts turn to preadipocytes, characterized by enlarged and fused LGs. After a relatively long incubation time, preadipocytes gradually transform into maturing adipocytes, usually in the 4th and 5th weeks. However, no HUCSC-derived fully mature adipocytes have been observed thus far, demonstrating either that adipogenic differentiation of in vivo cells does not proceed beyond the multilocular stage (dotted line) as they prematurely age or that an alternative induction regime is required to reach the final maturation state. Abbreviations: IVC, intervascular cell; LG, lipid granule; MSC, mesenchymal stem cell; P, passage; PVC, perivascular cell.

 

   DISCLOSURE OF POTENTIAL CONFLICTS OF INTEREST
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
Acknowledgments
 References
 
The authors indicate no potential conflicts of interest.


   ACKNOWLEDGMENTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
References
 
We thank Dr. Fadil Kara for generously providing human umbilical cords obtained from Cesarean sections (n = 26) and Gen-TR (http://www.gen-tr.gen.tr) for generous help on primer design. This study was financially supported by Ankara University Biotechnology Institute Grants 2005-180, TUBITAK-106S036-95, 105S364, and 107S088. S.K. is currently affiliated with the Infertility Clinics, Suleymaniye Maternity and Women's Health Research Training Hospital, Istanbul, Turkey.


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Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 

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