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Effects of adropin on proliferation and differentiation of 3T3-L1 cells and rat primary preadipocytes
Molecular and Cellular Endocrinology 496 (2019) 110532
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Effects of adropin on proliferation and differentiation of 3T3-L1 cells and rat primary preadipocytes
T
Mariami Jasaszwilia, Tatiana Wojciechowicza, Maria Billerta, Mathias Z. Strowskib,c, Krzysztof W. Nowaka, Marek Skrzypskia,∗ a
Department of Animal Physiology and Biochemistry, Poznań University of Life Sciences, 60-637, Poznań, Poland Department of Hepatology and Gastroenterology, Charité-University Medicine Berlin, Berlin, 13353, Germany c Department of Internal Medicine-Gastroenterology, Park-Klinik Weissensee, 13086, Berlin, Germany b
A R T I C LE I N FO
A B S T R A C T
Keywords: 3T3-L1 Adipogenesis Adropin Differentiation Enho Preadipocytes Proliferation
Adropin is a protein encoded by Energy Homeostasis Associated (Enho) gene which is expressed mainly in the liver and brain. There is evidence that biological effects of adropin are mediated via GPR19 activation. Animal studies showed that adropin modulates adiposity as well as lipid and glucose homeostasis. Adropin deficient animals have a phenotype closely resembling that of human metabolic syndrome with are obesity dyslipidemia and impaired glucose production. Animals treated with exogenous adropin lose weight, in addition to having reduced expression of lipogenic genes in the liver and fat tissue. While it was shown that adropin may contribute to energy homeostasis and body weight regulation, the role of this protein in controlling fat tissue formation is largely unknown. Thus, in the present study we investigated the effects of adropin on adipogenesis using 3T3-L1 cells and rat primary preadipocytes. We found a low Enho mRNA expression in 3T3-L1 cells and rat primary preadipocytes. Adropin stimulated proliferation of 3T3-L1 cells and rat primary preadipocytes. Stimulation of 3T3-L1 cell proliferation was mediated via ERK1/2 and AKT. Adropin reduced lipid accumulation as well as expression of proadipogenic genes in 3T3-L1 cells and rat preadipocytes, suggesting that this protein attenuates differentiation of preadipocytes into mature fat cells. In summary, these results show that adropin modulates proliferation and differentiation of preadipocytes.
1. Introduction Adropin is a secreted protein encoded by Energy Homeostasis Associated (Enho) gene (Kumar et al., 2008). Enho gene is mainly expressed in the liver and brain (Kumar et al., 2008). The expression of Enho depends on nutritional and metabolic state. Fat-enriched diet upregulates, while fasting reduces hepatic expression of Enho (Kumar et al., 2008). In contrast, in diet- or genetically-induced obese animal models, Enho expression in the liver is low (Kumar et al., 2008). There is emerging evidence indicating that adropin may play a role in regulating metabolism and controlling energy homeostasis. Adropin knockout mice (AdrKO) display adiposity despite normal food intake and energy expenditure (Ganesh Kumar, Zhang et al., 2012). Furthermore, AdrKO mice on the C57BL/6 J background have dyslipidemia and impaired glucose tolerance when fed high-fat diet (Ganesh Kumar, Zhang et al., 2012). These abnormalities are more pronounced in male animals (Ganesh Kumar, Zhang et al., 2012). In addition, a recent human study suggested that adropin may contribute to cholesterol
∗
homeostasis. It was demonstrated that adropin levels are inversely correlated with low-density lipoprotein cholesterol (LDL-C) in males (Ghoshal et al., 2018). In humans, low plasma levels of adropin are associated with insulin resistance (Butler et al., 2012). Consistently, in gestational and type 2 diabetes adropin levels are low (Beigi et al., 2015; Celik et al., 2013; Wu et al., 2014). Overall, these results collectively suggest that adropin may be involved in controlling body weight as well as lipid and glucose homeostasis. It is well known that lipid-carbohydrate metabolism is modulated by white adipose tissue activity. In addition to storage of triacylglycerol, adipocytes also produce and release a large number of hormones and metabolic factors which interact with peripheral tissues and the brain (Choe et al., 2016; Shimizu and Mori, 2005). Excessive content of adipose tissue is a hallmark of overweight and obesity and is linked to the development of metabolic syndrome, insulin resistance, type 2 diabetes, and cancer formation (Choe et al., 2016; Greenberg and Obin, 2006; Luo and Liu, 2016). Adipose tissue development and biology are
Corresponding author. Department of Animal Physiology and Biochemistry, Poznań University of Life Sciences, Wołyńska 35, 60-637, Poznań, Poland. E-mail address: [email protected] (M. Skrzypski).
https://doi.org/10.1016/j.mce.2019.110532 Received 14 February 2019; Received in revised form 1 July 2019; Accepted 3 August 2019 Available online 07 August 2019 0303-7207/ © 2019 Elsevier B.V. All rights reserved.
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Abbreviations AKT BrdU Ccnd1 c-myc C/ebpα C/ebpβ DMEM Enho ERK 1/2 Fabp4 Fasn
FBS Fsp27 Gapdh GPR19 IBMX ORO PBS Pcna Plin1 Pparγ T3 Tip47
serine/threonine-protein kinase 5-bromo-2′-deoxyuridine cyclin D1 myelocytomatosis oncogene CCAAT/enhancer-binding protein alpha CCAAT/enhancer-binding protein beta Dulbecco‘s Modified Eagle‘s Medium Energy Homeostasis Associated extracellular signal regulated kinases 1/2 fatty acid binding protein 4 fatty acid synthase
fetal bovine serum cell death-inducing DFFA-like effector c glyceraldehyde 3-phosphate dehydrogenase G Protein-Coupled Receptor 19 3-isobutyl-1-methylxanthin oil red O phosphate-buffered saline proliferating cell nuclear antigen perilipin 1 peroxisome proliferator-activated receptor gamma triiodothyronine perilipin 3
the care and use of animals were followed.
influenced by a variety of hormonal factors (Bjorntorp, 1997; Lee, 2017). While there is convincing evidence indicating that adropin peptide may modulate body mass, as well as glucose and lipid metabolism, the role of adropin in controlling adipogenesis in white fat is unknown. Differentiation of fat precursor cells (preadipocytes) into mature adipocytes is pivotal process in fat tissue formation, biology and remodelling (Ali et al., 2013). Abnormalities in the adipocytes differentiation contribute to the development of obesity and obesity-associated metabolic abnormalities (Camp et al., 2002; Spiegelman and Flier, 1996). Noteworthy, not only an excess of fat tissue but also the loss of fat tissue such as in patients with lipodystrophia causes numerous metabolic abnormalities and complications (Fiorenza et al., 2011). Thus, in the present study we studied the effects of adropin on proliferation and differentiation of white rat primary preadipocytes and 3T3-L1 cells.
2.4. Cell proliferation 3T3-L1 cells or rat primary preadipocytes were seeded in 96-well plates and cultured for 24 h in the growth medium at a density of 2 × 103 cells/well. Next, cells were incubated in a serum-free medium for additional 24 h. After the incubation cells were treated with or without adropin (1–100 nmol/l) for 24 or 48 h. Thereafter, BrdU solution was added (10 μmol/l) and cells were incubated for 3 h. Incorporation of BrdU into DNA was evaluated using Cell Proliferation BrdU Elisa Kit (Roche Diagnostics, Penzberg, Germany) according to the manufacturer's protocol. 2.5. 3T3-L1 cells and rat primary preadipocytes differentiation Two days post confluency, 3T3-L1 cells were cultured in the DMEM growth medium supplemented in 1 μmol/l dexamethasone, 500 μmol/l IBMX and 1 μmol/l insulin in the absence or presence of adropin (1, 10 or 100 nmol/l) for 48 h. Thereafter, this medium was replaced by a medium containing 1 μmol/l insulin with or without adropin, and cells were incubated for additional 48 h. Next, this medium was replaced by the growth medium and cells were incubated for additional two days in the presence or absence of adropin. Primary rat preadipocytes cells were cultured in a differentiation medium (DMEM/F12, 10% FCS, 850 mmol/l insulin, 2 nmol/l T3, 10 nmol/l dexamethasone) in the absence or presence of adropin (10 or 100 nmol/l), as previously described (Wojciechowicz et al., 2015). Medium was replaced every 48 h.
2. Materials and methods 2.1. Materials Adropin (34–76) (human, mouse, rat) trifluoroacetate salt was obtained from Bachem AG (Bubendorf, Switzerland). Media and supplements for cell culture were from Biowest (Nuaillé, France). 3T3-L1 fibroblast cell line (ATCC® CL-173TM) was from ATCC, LGC Standards (Manassas, VA, USA). Phospho-AKT (#9275L), AKT (#9272S), phospho-ERK1/2 (#9101S) and ERK1/2 (#9102S) antibodies were from Cell Signalling Technology (Danvers, MA, USA). PPARγ (#SAB4502262) antibody was from Sigma Aldrich (St. Louis, MO, USA). LY290042 and U0126 were from Tocris Bioscience (Bristol, UK). Unless otherwise stated, reagents were purchased from Sigma-Aldrich.
2.6. Real time PCR Real Time PCR was performed as we previously described (Billert et al., 2018). In brief, total RNA was isolated using Extrazol reagent (DNA-Gdańsk, Gdańsk, Poland), according to the manufacturer's protocol. Quality and quantity of isolated RNA was measured using NanoDrop 1000 (Thermo Scientific, Wilmington, DE, USA). 1 μg of total RNA was used for reverse transcription using FIREScript RT cDNA Synthesis MIX with Oligo (dT) and Random primers (Solis BioDyne, Tartu, Estonia). Obtained cDNA was diluted 1:10 with nuclease free water and used in real-time PCR reactions performed on QuantStudio 12K Flex (Life Technologies, CA, USA) by using EvaGreen qPCR Mix (Solis BioDyne). The primer sequences are listed in sup. Table 1. The relative quantification of gene expression levels was assessed using double delta CT method. All data were normalized to Gapdh mRNA expression level.
2.2. Cell culture 3T3-L1 cells were cultured in growth (pyruvate-free) DMEM medium supplemented with 10% FBS and penicillin-streptomycin (100 kU/l penicillin, 100 mg/l streptomycin) at 37 °C in a humidified atmosphere (5% CO2, 95% air). Rat primary preadipocytes were cultured in DMEM/F12 medium containing 10% FBS and penicillin-streptomycin (100 kU/l penicillin, 100 mg/l streptomycin) in a humidified atmosphere (5% CO2, 95% air). 2.3. Isolation of rat primary preadipocytes Stromal-vascular cell fraction containing preadipocytes was isolated from epididymal adipose fat pads obtained from male Wistar rats weighing 80–100 g (age, 5–6 weeks). Preadipocytes were purified as previously described by our laboratory (Wojciechowicz et al., 2015). All applicable international, national and/or institutional guidelines for
2.7. Western blot Phosphorylated, and total ERK1/2, and AKT were detected by 2
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inverted microscope and AxioVision Rel. software, version 4.6 (Carl Zeiss, Oberkochen, Germany). To assess lipid content, cells were air dried and ORO was eluted using 100% isopropanol. Absorbance values of eluates were read using microplate reader (Synergy 2 Multi-Mode Microplate Reader (BioTek, Winooski, VT, USA)) at 520 nm wavelength.
Western blot technique. Proteins were isolated using RIPA buffer (50 mmol/l Tris-HCL, pH 8.0 with 150 mmol NaCl, 1.0% NP-40, 0.5% sodium deoxycholate, 0.1% SDS) containing protease and phosphatase inhibitor cocktails (Roche Diagnostics). Proteins (30 μg per line) were separated on 5–12% Tris-HCl SDS-PAGE gel, transferred onto nitrocellulose membrane and detected, as we previously described (Skrzypski et al., 2014). Primary antibodies were diluted to 1:1000 and secondary antibodies to 1:5000.
2.9. Statistical analysis 2.8. Oil red O staining
Statistical analysis was performed using ANOVA followed by the Bonferroni post hoc test. T-student test was used to compare two groups. Statistical significance was considered when P < 0.05 (*). Data are shown as mean ± SEM. Each experiment was repeated independently at least two times.
3T3-L1 cells or rat primary preadipocytes were washed with PBS and fixed in 10% formaldehyde in PBS for 1 h. Next, cells were washed with 60% isopropanol, dried and stained with Oil red O working solution for 10 min, as described (Wojciechowicz et al., 2015). After that ORO staining, cells were washed four times with double distilled water. Representative images of stained cells were taken using an LSM 510
Fig. 1. Expression of Enho and Gpr19 mRNA in 3T3-L1 cells and rat primary preadipocytes. Images of undifferentiated rat primary preadipocytes (a) and 3T3-L1 cells (c). Differentiated rat primary preadipocytes (b) and 3T3-L1 cells (d) imaged 7 days after induction of differentiation. Expression of Enho and Gpr19 mRNAs in murine hypothalamus as well undifferentiated and differentiated (day 7th) 3T3-L1 cells (e, f) and in rat hypothalamus as well undifferentiated and differentiated (day 7th) rat primary preadipocytes (g, h). Results are the mean ± SEM, (n = 4–6). 3
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3. Results
the brain (positive control (Hoffmeister-Ullerich et al., 2004)) and at 93- or 60-fold lower levels in undifferentiated and differentiated 3T3-L1 cells, respectively (Fig. 1f). Enho mRNA is also present in the rat hypothalamus. Undifferentiated and differentiated rat preadipocytes express Enho mRNA, however at approximately 166- and 192-fold lower levels as compared to the rat hypothalamus (Fig. 1g). In addition Gpr19 mRNA expression was detected in rat hypothalamus as well as in undifferentiated and differentiated rat preadipocytes at approximately 1,6- or 2,3-fold lower levels, respectively (Fig. 1h). All these results show that Gpr19 and Enho mRNAs are detectable in 3T3-L1 cells and rat primary adipocytes. However, their expression was lower as compared with that measured in the hypothalamus.
3.1. Enho and Gpr19 mRNA is expressed in 3T3-L1 cells and rat preadipocytes First, we tested the expression of adropin precursor Enho and adropin putative receptor Gpr19 mRNAs in undifferentiated and differentiated 3T3-L1 cells, and rat primary preadipocytes. As shown in Fig. 1, undifferentiated rat primary adipocytes (Fig. 1a) and 3T3-L1 cells (Fig. 1c) show fibroblast-like morphology. By contrast, both, rat preadipocytes (b) and 3T3-L1 cells (d) differentiated for 7 days display spherical shape and are filled with lipids indicating that differentiation process into mature adipocytes was effective. Enho mRNA was detectable in the murine hypothalamus (positive control (Kumar et al., 2008)). As compared to hypothalamus, undifferentiated and differentiated 3T3-L1 cells express Enho at approximately 62.5- or 148-fold lower levels, respectively (Fig. 1e). Gpr19 mRNA is highly expressed in
Fig. 2. Effects of adropin on 3T3-L1 and rat primary preadipocytes proliferation. 3T3-L1 cell proliferation tested after 24 (a) and 48 h (b) treatment with adropin or vehicle. Rat primary preadipocyte proliferation evaluated in cells treated for 24 (c) or 48 h (d) with adropin. Expression of c-myc, Ccnd1 and Pcna in 3T3-L1 cells treated with adropin for 24 h (e). Western blot detection and quantification ERK1/2 (f–g) and AKT (h–i) phosphorylation in 3T3-L1 cell treated for indicated time points. Effect of U0126 (j) and LY294002 (k) on adropin-induced 3T3-L1 cell proliferation. Results are the mean ± SEM, (n = 6–8). 4
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cells after 15 min of incubation. In the presence of pharmacological MEK1/2-dependent blocker of ERK1/2 (U0126) (Favata et al., 1998) adropin-induced 3T3-L1 cell proliferation was completely attenuated (Fig. 2j). PI3/AKT blocker LY294002 (Vlahos et al., 1994) also partially suppressed basal cell proliferation, while addition of adropin stimulated cell proliferation in the presence of LY294002 (Fig. 2k). These data show that adropin promotes 3T3-L1 cell proliferation via ERK1/2 and AKT dependent mechanism.
3.2. Adropin stimulates proliferation of 3T3-L1 cells and rat preadipocytes role of ERK1/2 and AKT Next, we tested the ability of adropin to modulate cell proliferation. Adropin (100 nmol/l) stimulated 3T3-L1 preadipocytes proliferation after 24 and 48 h (Fig. 2a–b). In addition, adropin at the concentration of 100 nmol/l, increased rat preadipocytes proliferation after 48 (Fig. 2d) but not 24 h (Fig. 2c). To confirm the mitogenic activity of adropin, we studied mRNA expression c-myc, Ccnd1 and Pcna which are implicated in cell proliferation and DNA synthesis (Zhu et al., 2015). We found that adropin (100 nmol/l) stimulated expression of all three genes after 24 h (Fig. 2e). Furthermore, adropin (100 nmol/l) stimulated ERK1/2 (Fig. 2f–g) and AKT (Fig. 2h–i) phosphorylation in 3T3-L1
3.3. Adropin modulates expression of adipogenic markers in 3T3-L1 cells and rat preadipocytes Furthermore, we studied the effects of adropin on mRNA expression
Fig. 3. Expression of adipogenic genes in 3T3-L1 or rat primary preadipocytes treated with adropin or vehicle. Expression of Pparγ, Fabp4, C/ebpα and C/ebpβ in 3T3L1 cells differentiated for 7 days (a, b, c, d). Expression of Pparγ, Fabp4, C/ebpα and C/ebpβ in rat primary preadipocytes assessed 7 days after the onset of differentiation process (e, f, g, h). PPARγ protein production in 3T3-L1 cells (i, k) and rat primary preadipocytes (j, l) differentiated for 7 days in the presence or absence of adropin (100 nmol/l). Results are the mean ± SEM, (n = 4–6 or n = 3–4 (Western blots)). 5
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Fasn which are implicated in lipid synthesis, lipid droplet formation as well as antilipolytic activity (Greenberg et al., 1991; Menendez et al., 2009; Puri et al., 2007; Wolins et al., 2005). As shown in Fig. 4f, adropin suppressed expression of Fsp27 and Fasn in 3T3-L1 cells. Furthermore, adropin downregulated expression of Fsp27, Plin1 and Fasn in rat primary preadipocytes (Fig. 4l).
of important transcription factors and proteins such as Pparγ, C/ebpα, C/ebpβ and Fabp4 which are considered as adipogenic markers (Rosen and Spiegelman, 2000). Adropin (100 nmol/l) decreased the expression of Pparγ and Fabp4 in 3T3-L1 cells differentiated for 7 days (Fig. 3a–b). Furthermore, 10 nmol/l adropin suppressed expression of C/ebpβ in cells differentiated for 7 days (Fig. 3c). Moreover, adropin failed to modulate C/ebpα mRNA expression, evaluated 7 days after the onset of differentiation process (Fig. 3d). In rat primary preadipocytes differentiated for 7 days, adropin at 10 and 100 nmol/l suppressed Pparγ mRNA expression (Fig. 3e). Furthermore, adropin (10 and 100 nmol/l) decreased the expression of Fabp4 and C/ebpα mRNA in cells after 7 days of differentiation (Fig. 3f and g). C/ebpβ assessed 7 days after the onset of differentiation process was not affected by adropin (Fig. 3h). Adropin failed to affect the expression of mRNA of all tested genes in 3T3-L1 cells and rat primary preadipocytes, differentiated for 1 or 3 days (data not shown). We also found that adropin (100 nmol/l) downregulated PPARγ protein production in 3T3-L1 and rat primary adipocytes differentiated for 7 days (Fig. 3i-l).
4. Discussion Our study shows that adropin modulates proliferation and differentiation of preadipocytes. We found that Enho mRNA and putative adropin receptor Gpr19 mRNAs are expressed in 3T3-L1 as well as in rat primary preadipocytes. Nevertheless, Enho mRNA expression was rather very low as compared to the hypothalamus. Importantly, in a previous study, Enho mRNA in mouse fat tissue and 3T3-L1 cells was undetectable (Kumar et al., 2008). However the authors of this study used Northern blot analysis, only, which is much less sensitive as compared to the PCR in our current study (Streit et al., 2009). Even though that we found low level of Enho mRNA expression in rat preadipocytes and 3T3-L1 cells production of adropin protein in fat adipocytes cannot be excluded. Adipose tissue formation encompasses two major processes: fat precursor cell proliferation and differentiation into mature adipocytes (Fajas, 2003). Our data clearly show that adropin stimulates proliferation of 3T3-L1 cells as well as rat primary preadipocytes. The role of adropin in modifying cell proliferation is poorly documented so far. Of note, adropin was found to promote human umbilical vein ECs (HUVECs) cell proliferation (Lovren et al., 2010). Previous studies showed that adropin can activate AKT as well as ERK1/2 pathways. For example, it was demonstrated that adropin activates ERK1/2 signalling
3.4. Adropin reduces lipid accumulation in 3T3-L1 cells and rat preadipocytes Accumulation of lipids in preadipocytes is a hallmark of differentiation into mature adipocytes (Fu et al., 2005). Therefore, we assessed intracellular lipid level in 3T3-preadipocytes exposed to adropin. Adropin at 100 nmol/l decreased intracellular lipid levels in 3T3-L1 cells (Fig. 4a–e) and rat primary preadipocytes (Fig. 4g–k) assessed 7 days after the onset of differentiation process. Furthermore, we assessed the effects of adropin on mRNA expression of Fsp27, Plin1, Tip47 and
Fig. 4. Lipid accumulation and expression of genes involved in lipid accumulation in 3T3-L1 and rat primary preadipocytes differentiated in the presence or absence of adropin. Representative images of ORO-stained 3T3-L1 cells differentiated for 7 days without (a) or with 1 (b), 10 (c) or 100 nmol/ l adropin (d). Intracellular lipid content in 3T3-L1 cells assessed 7 days after the onset of differentiation process (e). Expression of Fsp27, Plin1, Tip47 and Fasn in 3T3-L1 cells differentiated for 7 days (f). Representative images of ORO-stained rat preadipocytes differentiated for 7 days without (g) or with 1 (h), 10 (i) or 100 nmol/l adropin (j). Intracellular lipid content in rat preadipocytes assessed 7 days after the onset of differentiation process (k). Expression of Fsp27, Plin1, Tip47 and Fasn in rat preadipocytes differentiated for 7 days (l). Results are the mean ± SEM, (n = 4–6).
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enhance insulin sensitivity in animals fed high-fat diet (Challa et al., 2012; Choe et al., 2016). Notably, adropin deficient mice are insulin resistant (Ganesh Kumar, Zhang et al., 2012). However, the role of adropin in controlling white adipose tissue cellular composition needs to be investigated in-depth and our current understanding about this issue has to be interpreted with caution. Our study has several limitations. Despite detecting Gpr19 mRNA expression in both rat primary preadipocytes as well as in 3T3-L1 cells we do not provide any evidence showing that promitogenic and antiadipogenic effects of adropin are mediated via GPR19. It must be considered that peptide hormones often interact with several types of receptors (Brothers and Wahlestedt, 2010; Sakurai et al., 1998). Furthermore, since we used rodent fat precursor cells, the question addressing the role of adropin in controlling human fat tissue remains to be investigated. Finally, we did not identify mechanism by which adropin can negatively affect adipogenesis. However, it is worth to notice that adropin can activate ERK1/2 pathway in 3T3-L1 cells. While it was shown that early activation of ERK1/2 is rather required to induce differentiation of preadipocytes (Prusty et al., 2002), others reported that rather late activation of ERK1/2 may suppress differentiation of fat precursor cells (Kim et al., 2007). Thus, potential role of ERK1/2 signalling and its time-dependent effects in adropin-mediated adipogenesis cannot be excluded. Another limitation of our study is that the effects of adropin on most genes expression were studied on mRNA level, only. Although suppression of proadipogenic genes mRNA expression was accompanied by reduced intracellular lipid content it needs to be investigated whether adropin affects protein production of tested genes. In summary, we found that adropin promotes preadipocytes proliferation via ERK1/2-dependent mechanism. Furthermore, adropin treatment reduces proadipogenic gene expression and intracellular lipid content during preadipocytes differentiation. These data indicated that adropin may be involved in controlling white adipogenesis.
in mesenchymal-like MDA-MB-231, thereby promoting breast cancer invasion (Rao and Herr, 2017). In addition, activation of AKT by adropin is essential in adropin-regulated endothelial cell functions (Lovren et al., 2010). Since both, ERK1/2 and AKT, are involved in controlling preadipocytes proliferation (Dong et al., 2016; Gan et al., 2015; Kim et al., 2004; Skrzypski et al., 2012), we assessed the effects of adropin of ERK1/2 and AKT phosphorylation in 3T3-L1 cells. Adropin promoted ERK1/2 and AKT phosphorylation in 3T3-L1 cells. Consistently, treatment of 3T3-L1 cells with adropin in the presence of ERK1/2 blocker attenuated the effects of adropin on cell proliferation. Furthermore, AKT blockade alone suppressed basal cell proliferation, however addition of adropin caused stimulation of cell proliferation. However, these effects were less pronounced as compared to cells treated with adropin without AKT blocker. Overall, these data indicate that stimulation of 3T3-L1 cell proliferation by adropin depends on ERK1/2 and AKT activation. Next, we studied whether adropin can affect adipogenesis in preadipocytes. To address this question, the effects of adropin on expression of transcription factors and proteins (Pparγ, Fabp4, C/ebpα and C/ ebpβ) which serve as well-defined stimuli and markers of preadipocytes differentiation, were evaluated (Rosen and Spiegelman, 2000). Adropin suppressed the expression of Pparγ, Fabp4 and C/ebpα. In addition, adropin decreased PPARγ protein production in 3T3-L1 and rat preadipocytes. However, these effects were observed in cells differentiated for 7 days, but not after three or one day. Therefore, adropin may be rather involved in modulating late but not early events of preadipocytes differentiation. Furthermore, we found that adropin suppressed lipid content in both 3T3-L1 as well as rat primary preadipocytes. Importantly, we also found that the suppression of lipid accumulation was accompanied by reduced expression of Fasn mRNA, which plays a role in stimulating lipid synthesis (Menendez et al., 2009) as well as lipid droplets formation (Fsp27, Plin1, Tip47) (Grahn et al., 2014; Greenberg et al., 1991). Overall, the suppression of adipogenic and lipogenic genes expression as well as intracellular lipid accumulation by adropin strongly suggests that this protein may attenuate adipogenesis, lipid synthesis and storage. Notably, a contribution of adropin to lipid metabolism regulation is supported by a previous animal study which showed that transgenic mice overexpressing adropin have low expression of lipogenic genes such as Pparγ in the liver and adipose tissue (Ganesh Kumar, Zhang et al., 2012). Furthermore, adropin-overexpressing mice fed high-fat diet have attenuated body weight gain (Kumar et al., 2008). Consistently, it was shown that adropin deficiency is linked to increased fat mass in mice fed normocaloric but not high-fat diet (Ganesh Kumar, Zhang et al., 2012). In addition, a human study reported negative correlation between body mass index and serum adropin levels (Butler et al., 2012). Overall, the reduction of adipogenic gene expression as well as lipid content in 3T3-L1 and rat primary preadipocytes exposed to adropin in addition to protection from obesity in in vivo studies suggests that adropin may be involved in regulating lipid accumulation and adipose tissue formation. Nevertheless, assuming that adropin is able to prevent obesity, stimulation of preadipocyte proliferation by adropin appears to contradict this assumption. Increased fat mass in obesity results from both, a hypertrophy of pre-existing mature adipocytes as well as from increased proliferation of preadipocytes (hyperplasia) (Jo et al., 2009). In humans, metabolic disturbances associated with obesity are mainly caused rather by hypertrophy whereas hyperplasia of adipocytes is less important, if at all (Choe et al., 2016). Undoubtedly, numerous pro-inflammatory cytokines and other factors responsible for “inflamed fat” are also responsible for the obesity and obesity-associated complications such as insulin-resistance (Nishimura et al., 2009). Furthermore, stimuli of preadipocytes proliferation and differentiation into new small-size adipocytes can reduce hyperlipidemia and
Declarations of interest None. Funding This research was supported by grant from the National Science Centre (Poland) (2016/23/D/NZ4/03557 to M. Skrzypski). Acknowledgment MS designed the study and wrote the manuscript. MJ performed the experiments and wrote the manuscript. TW and MB contributed to the experiments. MZS and KWN discussed data and reviewed the manuscript. Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.mce.2019.110532. References Ali, A.T., Hochfeld, W.E., Myburgh, R., Pepper, M.S., 2013. Adipocyte and adipogenesis. Eur. J. Cell Biol. 92, 229–236. Beigi, A., Shirzad, N., Nikpour, F., Nasli Esfahani, E., Emamgholipour, S., Bandarian, F., 2015. Association between serum adropin levels and gestational diabetes mellitus; a case-control study. Gynecol. Endocrinol. 31, 939–941. Billert, M., Wojciechowicz, T., Jasaszwili, M., Szczepankiewicz, D., Wasko, J., Kazmierczak, S., Strowski, M.Z., Nowak, K.W., Skrzypski, M., 2018. Phoenixin-14 stimulates differentiation of 3T3-L1 preadipocytes via cAMP/Epac-dependent mechanism. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 1863, 1449–1457. Bjorntorp, P., 1997. Hormonal control of regional fat distribution. Hum. Reprod. 12 (Suppl. 1), 21–25.
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Effects of adropin on proliferation and differentiation of 3T3-L1 cells and rat primary preadipocytes
T
Mariami Jasaszwilia, Tatiana Wojciechowicza, Maria Billerta, Mathias Z. Strowskib,c, Krzysztof W. Nowaka, Marek Skrzypskia,∗ a
Department of Animal Physiology and Biochemistry, Poznań University of Life Sciences, 60-637, Poznań, Poland Department of Hepatology and Gastroenterology, Charité-University Medicine Berlin, Berlin, 13353, Germany c Department of Internal Medicine-Gastroenterology, Park-Klinik Weissensee, 13086, Berlin, Germany b
A R T I C LE I N FO
A B S T R A C T
Keywords: 3T3-L1 Adipogenesis Adropin Differentiation Enho Preadipocytes Proliferation
Adropin is a protein encoded by Energy Homeostasis Associated (Enho) gene which is expressed mainly in the liver and brain. There is evidence that biological effects of adropin are mediated via GPR19 activation. Animal studies showed that adropin modulates adiposity as well as lipid and glucose homeostasis. Adropin deficient animals have a phenotype closely resembling that of human metabolic syndrome with are obesity dyslipidemia and impaired glucose production. Animals treated with exogenous adropin lose weight, in addition to having reduced expression of lipogenic genes in the liver and fat tissue. While it was shown that adropin may contribute to energy homeostasis and body weight regulation, the role of this protein in controlling fat tissue formation is largely unknown. Thus, in the present study we investigated the effects of adropin on adipogenesis using 3T3-L1 cells and rat primary preadipocytes. We found a low Enho mRNA expression in 3T3-L1 cells and rat primary preadipocytes. Adropin stimulated proliferation of 3T3-L1 cells and rat primary preadipocytes. Stimulation of 3T3-L1 cell proliferation was mediated via ERK1/2 and AKT. Adropin reduced lipid accumulation as well as expression of proadipogenic genes in 3T3-L1 cells and rat preadipocytes, suggesting that this protein attenuates differentiation of preadipocytes into mature fat cells. In summary, these results show that adropin modulates proliferation and differentiation of preadipocytes.
1. Introduction Adropin is a secreted protein encoded by Energy Homeostasis Associated (Enho) gene (Kumar et al., 2008). Enho gene is mainly expressed in the liver and brain (Kumar et al., 2008). The expression of Enho depends on nutritional and metabolic state. Fat-enriched diet upregulates, while fasting reduces hepatic expression of Enho (Kumar et al., 2008). In contrast, in diet- or genetically-induced obese animal models, Enho expression in the liver is low (Kumar et al., 2008). There is emerging evidence indicating that adropin may play a role in regulating metabolism and controlling energy homeostasis. Adropin knockout mice (AdrKO) display adiposity despite normal food intake and energy expenditure (Ganesh Kumar, Zhang et al., 2012). Furthermore, AdrKO mice on the C57BL/6 J background have dyslipidemia and impaired glucose tolerance when fed high-fat diet (Ganesh Kumar, Zhang et al., 2012). These abnormalities are more pronounced in male animals (Ganesh Kumar, Zhang et al., 2012). In addition, a recent human study suggested that adropin may contribute to cholesterol
∗
homeostasis. It was demonstrated that adropin levels are inversely correlated with low-density lipoprotein cholesterol (LDL-C) in males (Ghoshal et al., 2018). In humans, low plasma levels of adropin are associated with insulin resistance (Butler et al., 2012). Consistently, in gestational and type 2 diabetes adropin levels are low (Beigi et al., 2015; Celik et al., 2013; Wu et al., 2014). Overall, these results collectively suggest that adropin may be involved in controlling body weight as well as lipid and glucose homeostasis. It is well known that lipid-carbohydrate metabolism is modulated by white adipose tissue activity. In addition to storage of triacylglycerol, adipocytes also produce and release a large number of hormones and metabolic factors which interact with peripheral tissues and the brain (Choe et al., 2016; Shimizu and Mori, 2005). Excessive content of adipose tissue is a hallmark of overweight and obesity and is linked to the development of metabolic syndrome, insulin resistance, type 2 diabetes, and cancer formation (Choe et al., 2016; Greenberg and Obin, 2006; Luo and Liu, 2016). Adipose tissue development and biology are
Corresponding author. Department of Animal Physiology and Biochemistry, Poznań University of Life Sciences, Wołyńska 35, 60-637, Poznań, Poland. E-mail address: [email protected] (M. Skrzypski).
https://doi.org/10.1016/j.mce.2019.110532 Received 14 February 2019; Received in revised form 1 July 2019; Accepted 3 August 2019 Available online 07 August 2019 0303-7207/ © 2019 Elsevier B.V. All rights reserved.
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Abbreviations AKT BrdU Ccnd1 c-myc C/ebpα C/ebpβ DMEM Enho ERK 1/2 Fabp4 Fasn
FBS Fsp27 Gapdh GPR19 IBMX ORO PBS Pcna Plin1 Pparγ T3 Tip47
serine/threonine-protein kinase 5-bromo-2′-deoxyuridine cyclin D1 myelocytomatosis oncogene CCAAT/enhancer-binding protein alpha CCAAT/enhancer-binding protein beta Dulbecco‘s Modified Eagle‘s Medium Energy Homeostasis Associated extracellular signal regulated kinases 1/2 fatty acid binding protein 4 fatty acid synthase
fetal bovine serum cell death-inducing DFFA-like effector c glyceraldehyde 3-phosphate dehydrogenase G Protein-Coupled Receptor 19 3-isobutyl-1-methylxanthin oil red O phosphate-buffered saline proliferating cell nuclear antigen perilipin 1 peroxisome proliferator-activated receptor gamma triiodothyronine perilipin 3
the care and use of animals were followed.
influenced by a variety of hormonal factors (Bjorntorp, 1997; Lee, 2017). While there is convincing evidence indicating that adropin peptide may modulate body mass, as well as glucose and lipid metabolism, the role of adropin in controlling adipogenesis in white fat is unknown. Differentiation of fat precursor cells (preadipocytes) into mature adipocytes is pivotal process in fat tissue formation, biology and remodelling (Ali et al., 2013). Abnormalities in the adipocytes differentiation contribute to the development of obesity and obesity-associated metabolic abnormalities (Camp et al., 2002; Spiegelman and Flier, 1996). Noteworthy, not only an excess of fat tissue but also the loss of fat tissue such as in patients with lipodystrophia causes numerous metabolic abnormalities and complications (Fiorenza et al., 2011). Thus, in the present study we studied the effects of adropin on proliferation and differentiation of white rat primary preadipocytes and 3T3-L1 cells.
2.4. Cell proliferation 3T3-L1 cells or rat primary preadipocytes were seeded in 96-well plates and cultured for 24 h in the growth medium at a density of 2 × 103 cells/well. Next, cells were incubated in a serum-free medium for additional 24 h. After the incubation cells were treated with or without adropin (1–100 nmol/l) for 24 or 48 h. Thereafter, BrdU solution was added (10 μmol/l) and cells were incubated for 3 h. Incorporation of BrdU into DNA was evaluated using Cell Proliferation BrdU Elisa Kit (Roche Diagnostics, Penzberg, Germany) according to the manufacturer's protocol. 2.5. 3T3-L1 cells and rat primary preadipocytes differentiation Two days post confluency, 3T3-L1 cells were cultured in the DMEM growth medium supplemented in 1 μmol/l dexamethasone, 500 μmol/l IBMX and 1 μmol/l insulin in the absence or presence of adropin (1, 10 or 100 nmol/l) for 48 h. Thereafter, this medium was replaced by a medium containing 1 μmol/l insulin with or without adropin, and cells were incubated for additional 48 h. Next, this medium was replaced by the growth medium and cells were incubated for additional two days in the presence or absence of adropin. Primary rat preadipocytes cells were cultured in a differentiation medium (DMEM/F12, 10% FCS, 850 mmol/l insulin, 2 nmol/l T3, 10 nmol/l dexamethasone) in the absence or presence of adropin (10 or 100 nmol/l), as previously described (Wojciechowicz et al., 2015). Medium was replaced every 48 h.
2. Materials and methods 2.1. Materials Adropin (34–76) (human, mouse, rat) trifluoroacetate salt was obtained from Bachem AG (Bubendorf, Switzerland). Media and supplements for cell culture were from Biowest (Nuaillé, France). 3T3-L1 fibroblast cell line (ATCC® CL-173TM) was from ATCC, LGC Standards (Manassas, VA, USA). Phospho-AKT (#9275L), AKT (#9272S), phospho-ERK1/2 (#9101S) and ERK1/2 (#9102S) antibodies were from Cell Signalling Technology (Danvers, MA, USA). PPARγ (#SAB4502262) antibody was from Sigma Aldrich (St. Louis, MO, USA). LY290042 and U0126 were from Tocris Bioscience (Bristol, UK). Unless otherwise stated, reagents were purchased from Sigma-Aldrich.
2.6. Real time PCR Real Time PCR was performed as we previously described (Billert et al., 2018). In brief, total RNA was isolated using Extrazol reagent (DNA-Gdańsk, Gdańsk, Poland), according to the manufacturer's protocol. Quality and quantity of isolated RNA was measured using NanoDrop 1000 (Thermo Scientific, Wilmington, DE, USA). 1 μg of total RNA was used for reverse transcription using FIREScript RT cDNA Synthesis MIX with Oligo (dT) and Random primers (Solis BioDyne, Tartu, Estonia). Obtained cDNA was diluted 1:10 with nuclease free water and used in real-time PCR reactions performed on QuantStudio 12K Flex (Life Technologies, CA, USA) by using EvaGreen qPCR Mix (Solis BioDyne). The primer sequences are listed in sup. Table 1. The relative quantification of gene expression levels was assessed using double delta CT method. All data were normalized to Gapdh mRNA expression level.
2.2. Cell culture 3T3-L1 cells were cultured in growth (pyruvate-free) DMEM medium supplemented with 10% FBS and penicillin-streptomycin (100 kU/l penicillin, 100 mg/l streptomycin) at 37 °C in a humidified atmosphere (5% CO2, 95% air). Rat primary preadipocytes were cultured in DMEM/F12 medium containing 10% FBS and penicillin-streptomycin (100 kU/l penicillin, 100 mg/l streptomycin) in a humidified atmosphere (5% CO2, 95% air). 2.3. Isolation of rat primary preadipocytes Stromal-vascular cell fraction containing preadipocytes was isolated from epididymal adipose fat pads obtained from male Wistar rats weighing 80–100 g (age, 5–6 weeks). Preadipocytes were purified as previously described by our laboratory (Wojciechowicz et al., 2015). All applicable international, national and/or institutional guidelines for
2.7. Western blot Phosphorylated, and total ERK1/2, and AKT were detected by 2
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inverted microscope and AxioVision Rel. software, version 4.6 (Carl Zeiss, Oberkochen, Germany). To assess lipid content, cells were air dried and ORO was eluted using 100% isopropanol. Absorbance values of eluates were read using microplate reader (Synergy 2 Multi-Mode Microplate Reader (BioTek, Winooski, VT, USA)) at 520 nm wavelength.
Western blot technique. Proteins were isolated using RIPA buffer (50 mmol/l Tris-HCL, pH 8.0 with 150 mmol NaCl, 1.0% NP-40, 0.5% sodium deoxycholate, 0.1% SDS) containing protease and phosphatase inhibitor cocktails (Roche Diagnostics). Proteins (30 μg per line) were separated on 5–12% Tris-HCl SDS-PAGE gel, transferred onto nitrocellulose membrane and detected, as we previously described (Skrzypski et al., 2014). Primary antibodies were diluted to 1:1000 and secondary antibodies to 1:5000.
2.9. Statistical analysis 2.8. Oil red O staining
Statistical analysis was performed using ANOVA followed by the Bonferroni post hoc test. T-student test was used to compare two groups. Statistical significance was considered when P < 0.05 (*). Data are shown as mean ± SEM. Each experiment was repeated independently at least two times.
3T3-L1 cells or rat primary preadipocytes were washed with PBS and fixed in 10% formaldehyde in PBS for 1 h. Next, cells were washed with 60% isopropanol, dried and stained with Oil red O working solution for 10 min, as described (Wojciechowicz et al., 2015). After that ORO staining, cells were washed four times with double distilled water. Representative images of stained cells were taken using an LSM 510
Fig. 1. Expression of Enho and Gpr19 mRNA in 3T3-L1 cells and rat primary preadipocytes. Images of undifferentiated rat primary preadipocytes (a) and 3T3-L1 cells (c). Differentiated rat primary preadipocytes (b) and 3T3-L1 cells (d) imaged 7 days after induction of differentiation. Expression of Enho and Gpr19 mRNAs in murine hypothalamus as well undifferentiated and differentiated (day 7th) 3T3-L1 cells (e, f) and in rat hypothalamus as well undifferentiated and differentiated (day 7th) rat primary preadipocytes (g, h). Results are the mean ± SEM, (n = 4–6). 3
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3. Results
the brain (positive control (Hoffmeister-Ullerich et al., 2004)) and at 93- or 60-fold lower levels in undifferentiated and differentiated 3T3-L1 cells, respectively (Fig. 1f). Enho mRNA is also present in the rat hypothalamus. Undifferentiated and differentiated rat preadipocytes express Enho mRNA, however at approximately 166- and 192-fold lower levels as compared to the rat hypothalamus (Fig. 1g). In addition Gpr19 mRNA expression was detected in rat hypothalamus as well as in undifferentiated and differentiated rat preadipocytes at approximately 1,6- or 2,3-fold lower levels, respectively (Fig. 1h). All these results show that Gpr19 and Enho mRNAs are detectable in 3T3-L1 cells and rat primary adipocytes. However, their expression was lower as compared with that measured in the hypothalamus.
3.1. Enho and Gpr19 mRNA is expressed in 3T3-L1 cells and rat preadipocytes First, we tested the expression of adropin precursor Enho and adropin putative receptor Gpr19 mRNAs in undifferentiated and differentiated 3T3-L1 cells, and rat primary preadipocytes. As shown in Fig. 1, undifferentiated rat primary adipocytes (Fig. 1a) and 3T3-L1 cells (Fig. 1c) show fibroblast-like morphology. By contrast, both, rat preadipocytes (b) and 3T3-L1 cells (d) differentiated for 7 days display spherical shape and are filled with lipids indicating that differentiation process into mature adipocytes was effective. Enho mRNA was detectable in the murine hypothalamus (positive control (Kumar et al., 2008)). As compared to hypothalamus, undifferentiated and differentiated 3T3-L1 cells express Enho at approximately 62.5- or 148-fold lower levels, respectively (Fig. 1e). Gpr19 mRNA is highly expressed in
Fig. 2. Effects of adropin on 3T3-L1 and rat primary preadipocytes proliferation. 3T3-L1 cell proliferation tested after 24 (a) and 48 h (b) treatment with adropin or vehicle. Rat primary preadipocyte proliferation evaluated in cells treated for 24 (c) or 48 h (d) with adropin. Expression of c-myc, Ccnd1 and Pcna in 3T3-L1 cells treated with adropin for 24 h (e). Western blot detection and quantification ERK1/2 (f–g) and AKT (h–i) phosphorylation in 3T3-L1 cell treated for indicated time points. Effect of U0126 (j) and LY294002 (k) on adropin-induced 3T3-L1 cell proliferation. Results are the mean ± SEM, (n = 6–8). 4
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cells after 15 min of incubation. In the presence of pharmacological MEK1/2-dependent blocker of ERK1/2 (U0126) (Favata et al., 1998) adropin-induced 3T3-L1 cell proliferation was completely attenuated (Fig. 2j). PI3/AKT blocker LY294002 (Vlahos et al., 1994) also partially suppressed basal cell proliferation, while addition of adropin stimulated cell proliferation in the presence of LY294002 (Fig. 2k). These data show that adropin promotes 3T3-L1 cell proliferation via ERK1/2 and AKT dependent mechanism.
3.2. Adropin stimulates proliferation of 3T3-L1 cells and rat preadipocytes role of ERK1/2 and AKT Next, we tested the ability of adropin to modulate cell proliferation. Adropin (100 nmol/l) stimulated 3T3-L1 preadipocytes proliferation after 24 and 48 h (Fig. 2a–b). In addition, adropin at the concentration of 100 nmol/l, increased rat preadipocytes proliferation after 48 (Fig. 2d) but not 24 h (Fig. 2c). To confirm the mitogenic activity of adropin, we studied mRNA expression c-myc, Ccnd1 and Pcna which are implicated in cell proliferation and DNA synthesis (Zhu et al., 2015). We found that adropin (100 nmol/l) stimulated expression of all three genes after 24 h (Fig. 2e). Furthermore, adropin (100 nmol/l) stimulated ERK1/2 (Fig. 2f–g) and AKT (Fig. 2h–i) phosphorylation in 3T3-L1
3.3. Adropin modulates expression of adipogenic markers in 3T3-L1 cells and rat preadipocytes Furthermore, we studied the effects of adropin on mRNA expression
Fig. 3. Expression of adipogenic genes in 3T3-L1 or rat primary preadipocytes treated with adropin or vehicle. Expression of Pparγ, Fabp4, C/ebpα and C/ebpβ in 3T3L1 cells differentiated for 7 days (a, b, c, d). Expression of Pparγ, Fabp4, C/ebpα and C/ebpβ in rat primary preadipocytes assessed 7 days after the onset of differentiation process (e, f, g, h). PPARγ protein production in 3T3-L1 cells (i, k) and rat primary preadipocytes (j, l) differentiated for 7 days in the presence or absence of adropin (100 nmol/l). Results are the mean ± SEM, (n = 4–6 or n = 3–4 (Western blots)). 5
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Fasn which are implicated in lipid synthesis, lipid droplet formation as well as antilipolytic activity (Greenberg et al., 1991; Menendez et al., 2009; Puri et al., 2007; Wolins et al., 2005). As shown in Fig. 4f, adropin suppressed expression of Fsp27 and Fasn in 3T3-L1 cells. Furthermore, adropin downregulated expression of Fsp27, Plin1 and Fasn in rat primary preadipocytes (Fig. 4l).
of important transcription factors and proteins such as Pparγ, C/ebpα, C/ebpβ and Fabp4 which are considered as adipogenic markers (Rosen and Spiegelman, 2000). Adropin (100 nmol/l) decreased the expression of Pparγ and Fabp4 in 3T3-L1 cells differentiated for 7 days (Fig. 3a–b). Furthermore, 10 nmol/l adropin suppressed expression of C/ebpβ in cells differentiated for 7 days (Fig. 3c). Moreover, adropin failed to modulate C/ebpα mRNA expression, evaluated 7 days after the onset of differentiation process (Fig. 3d). In rat primary preadipocytes differentiated for 7 days, adropin at 10 and 100 nmol/l suppressed Pparγ mRNA expression (Fig. 3e). Furthermore, adropin (10 and 100 nmol/l) decreased the expression of Fabp4 and C/ebpα mRNA in cells after 7 days of differentiation (Fig. 3f and g). C/ebpβ assessed 7 days after the onset of differentiation process was not affected by adropin (Fig. 3h). Adropin failed to affect the expression of mRNA of all tested genes in 3T3-L1 cells and rat primary preadipocytes, differentiated for 1 or 3 days (data not shown). We also found that adropin (100 nmol/l) downregulated PPARγ protein production in 3T3-L1 and rat primary adipocytes differentiated for 7 days (Fig. 3i-l).
4. Discussion Our study shows that adropin modulates proliferation and differentiation of preadipocytes. We found that Enho mRNA and putative adropin receptor Gpr19 mRNAs are expressed in 3T3-L1 as well as in rat primary preadipocytes. Nevertheless, Enho mRNA expression was rather very low as compared to the hypothalamus. Importantly, in a previous study, Enho mRNA in mouse fat tissue and 3T3-L1 cells was undetectable (Kumar et al., 2008). However the authors of this study used Northern blot analysis, only, which is much less sensitive as compared to the PCR in our current study (Streit et al., 2009). Even though that we found low level of Enho mRNA expression in rat preadipocytes and 3T3-L1 cells production of adropin protein in fat adipocytes cannot be excluded. Adipose tissue formation encompasses two major processes: fat precursor cell proliferation and differentiation into mature adipocytes (Fajas, 2003). Our data clearly show that adropin stimulates proliferation of 3T3-L1 cells as well as rat primary preadipocytes. The role of adropin in modifying cell proliferation is poorly documented so far. Of note, adropin was found to promote human umbilical vein ECs (HUVECs) cell proliferation (Lovren et al., 2010). Previous studies showed that adropin can activate AKT as well as ERK1/2 pathways. For example, it was demonstrated that adropin activates ERK1/2 signalling
3.4. Adropin reduces lipid accumulation in 3T3-L1 cells and rat preadipocytes Accumulation of lipids in preadipocytes is a hallmark of differentiation into mature adipocytes (Fu et al., 2005). Therefore, we assessed intracellular lipid level in 3T3-preadipocytes exposed to adropin. Adropin at 100 nmol/l decreased intracellular lipid levels in 3T3-L1 cells (Fig. 4a–e) and rat primary preadipocytes (Fig. 4g–k) assessed 7 days after the onset of differentiation process. Furthermore, we assessed the effects of adropin on mRNA expression of Fsp27, Plin1, Tip47 and
Fig. 4. Lipid accumulation and expression of genes involved in lipid accumulation in 3T3-L1 and rat primary preadipocytes differentiated in the presence or absence of adropin. Representative images of ORO-stained 3T3-L1 cells differentiated for 7 days without (a) or with 1 (b), 10 (c) or 100 nmol/ l adropin (d). Intracellular lipid content in 3T3-L1 cells assessed 7 days after the onset of differentiation process (e). Expression of Fsp27, Plin1, Tip47 and Fasn in 3T3-L1 cells differentiated for 7 days (f). Representative images of ORO-stained rat preadipocytes differentiated for 7 days without (g) or with 1 (h), 10 (i) or 100 nmol/l adropin (j). Intracellular lipid content in rat preadipocytes assessed 7 days after the onset of differentiation process (k). Expression of Fsp27, Plin1, Tip47 and Fasn in rat preadipocytes differentiated for 7 days (l). Results are the mean ± SEM, (n = 4–6).
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enhance insulin sensitivity in animals fed high-fat diet (Challa et al., 2012; Choe et al., 2016). Notably, adropin deficient mice are insulin resistant (Ganesh Kumar, Zhang et al., 2012). However, the role of adropin in controlling white adipose tissue cellular composition needs to be investigated in-depth and our current understanding about this issue has to be interpreted with caution. Our study has several limitations. Despite detecting Gpr19 mRNA expression in both rat primary preadipocytes as well as in 3T3-L1 cells we do not provide any evidence showing that promitogenic and antiadipogenic effects of adropin are mediated via GPR19. It must be considered that peptide hormones often interact with several types of receptors (Brothers and Wahlestedt, 2010; Sakurai et al., 1998). Furthermore, since we used rodent fat precursor cells, the question addressing the role of adropin in controlling human fat tissue remains to be investigated. Finally, we did not identify mechanism by which adropin can negatively affect adipogenesis. However, it is worth to notice that adropin can activate ERK1/2 pathway in 3T3-L1 cells. While it was shown that early activation of ERK1/2 is rather required to induce differentiation of preadipocytes (Prusty et al., 2002), others reported that rather late activation of ERK1/2 may suppress differentiation of fat precursor cells (Kim et al., 2007). Thus, potential role of ERK1/2 signalling and its time-dependent effects in adropin-mediated adipogenesis cannot be excluded. Another limitation of our study is that the effects of adropin on most genes expression were studied on mRNA level, only. Although suppression of proadipogenic genes mRNA expression was accompanied by reduced intracellular lipid content it needs to be investigated whether adropin affects protein production of tested genes. In summary, we found that adropin promotes preadipocytes proliferation via ERK1/2-dependent mechanism. Furthermore, adropin treatment reduces proadipogenic gene expression and intracellular lipid content during preadipocytes differentiation. These data indicated that adropin may be involved in controlling white adipogenesis.
in mesenchymal-like MDA-MB-231, thereby promoting breast cancer invasion (Rao and Herr, 2017). In addition, activation of AKT by adropin is essential in adropin-regulated endothelial cell functions (Lovren et al., 2010). Since both, ERK1/2 and AKT, are involved in controlling preadipocytes proliferation (Dong et al., 2016; Gan et al., 2015; Kim et al., 2004; Skrzypski et al., 2012), we assessed the effects of adropin of ERK1/2 and AKT phosphorylation in 3T3-L1 cells. Adropin promoted ERK1/2 and AKT phosphorylation in 3T3-L1 cells. Consistently, treatment of 3T3-L1 cells with adropin in the presence of ERK1/2 blocker attenuated the effects of adropin on cell proliferation. Furthermore, AKT blockade alone suppressed basal cell proliferation, however addition of adropin caused stimulation of cell proliferation. However, these effects were less pronounced as compared to cells treated with adropin without AKT blocker. Overall, these data indicate that stimulation of 3T3-L1 cell proliferation by adropin depends on ERK1/2 and AKT activation. Next, we studied whether adropin can affect adipogenesis in preadipocytes. To address this question, the effects of adropin on expression of transcription factors and proteins (Pparγ, Fabp4, C/ebpα and C/ ebpβ) which serve as well-defined stimuli and markers of preadipocytes differentiation, were evaluated (Rosen and Spiegelman, 2000). Adropin suppressed the expression of Pparγ, Fabp4 and C/ebpα. In addition, adropin decreased PPARγ protein production in 3T3-L1 and rat preadipocytes. However, these effects were observed in cells differentiated for 7 days, but not after three or one day. Therefore, adropin may be rather involved in modulating late but not early events of preadipocytes differentiation. Furthermore, we found that adropin suppressed lipid content in both 3T3-L1 as well as rat primary preadipocytes. Importantly, we also found that the suppression of lipid accumulation was accompanied by reduced expression of Fasn mRNA, which plays a role in stimulating lipid synthesis (Menendez et al., 2009) as well as lipid droplets formation (Fsp27, Plin1, Tip47) (Grahn et al., 2014; Greenberg et al., 1991). Overall, the suppression of adipogenic and lipogenic genes expression as well as intracellular lipid accumulation by adropin strongly suggests that this protein may attenuate adipogenesis, lipid synthesis and storage. Notably, a contribution of adropin to lipid metabolism regulation is supported by a previous animal study which showed that transgenic mice overexpressing adropin have low expression of lipogenic genes such as Pparγ in the liver and adipose tissue (Ganesh Kumar, Zhang et al., 2012). Furthermore, adropin-overexpressing mice fed high-fat diet have attenuated body weight gain (Kumar et al., 2008). Consistently, it was shown that adropin deficiency is linked to increased fat mass in mice fed normocaloric but not high-fat diet (Ganesh Kumar, Zhang et al., 2012). In addition, a human study reported negative correlation between body mass index and serum adropin levels (Butler et al., 2012). Overall, the reduction of adipogenic gene expression as well as lipid content in 3T3-L1 and rat primary preadipocytes exposed to adropin in addition to protection from obesity in in vivo studies suggests that adropin may be involved in regulating lipid accumulation and adipose tissue formation. Nevertheless, assuming that adropin is able to prevent obesity, stimulation of preadipocyte proliferation by adropin appears to contradict this assumption. Increased fat mass in obesity results from both, a hypertrophy of pre-existing mature adipocytes as well as from increased proliferation of preadipocytes (hyperplasia) (Jo et al., 2009). In humans, metabolic disturbances associated with obesity are mainly caused rather by hypertrophy whereas hyperplasia of adipocytes is less important, if at all (Choe et al., 2016). Undoubtedly, numerous pro-inflammatory cytokines and other factors responsible for “inflamed fat” are also responsible for the obesity and obesity-associated complications such as insulin-resistance (Nishimura et al., 2009). Furthermore, stimuli of preadipocytes proliferation and differentiation into new small-size adipocytes can reduce hyperlipidemia and
Declarations of interest None. Funding This research was supported by grant from the National Science Centre (Poland) (2016/23/D/NZ4/03557 to M. Skrzypski). Acknowledgment MS designed the study and wrote the manuscript. MJ performed the experiments and wrote the manuscript. TW and MB contributed to the experiments. MZS and KWN discussed data and reviewed the manuscript. Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.mce.2019.110532. References Ali, A.T., Hochfeld, W.E., Myburgh, R., Pepper, M.S., 2013. Adipocyte and adipogenesis. Eur. J. Cell Biol. 92, 229–236. Beigi, A., Shirzad, N., Nikpour, F., Nasli Esfahani, E., Emamgholipour, S., Bandarian, F., 2015. Association between serum adropin levels and gestational diabetes mellitus; a case-control study. Gynecol. Endocrinol. 31, 939–941. Billert, M., Wojciechowicz, T., Jasaszwili, M., Szczepankiewicz, D., Wasko, J., Kazmierczak, S., Strowski, M.Z., Nowak, K.W., Skrzypski, M., 2018. Phoenixin-14 stimulates differentiation of 3T3-L1 preadipocytes via cAMP/Epac-dependent mechanism. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 1863, 1449–1457. Bjorntorp, P., 1997. Hormonal control of regional fat distribution. Hum. Reprod. 12 (Suppl. 1), 21–25.
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