Cellular prion protein regulates the differentiation and function of adipocytes through autophagy flux

Molecular and Cellular Endocrinology 481 (2019) 84–94

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Molecular and Cellular Endocrinology journal homepage: www.elsevier.com/locate/mce

Cellular prion protein regulates the differentiation and function of adipocytes through autophagy flux

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Jae-Kyo Jeong1, Ju-Hee Lee1, Sung-Wook Kim1, Jeong-Min Hong, Jae-Won Seol, Sang-Youel Park∗ Biosafety Research Institute, College of Veterinary Medicine, Chonbuk National University, Iksan, Jeonbuk, 54596, Republic of Korea

ARTICLE INFO

ABSTRACT

Keywords: Autophagy PrP < C > Adipogenesis Differentiation Prnp-knockout mice

The role of autophagy modulation in adipogenic differentiation and the possible autophagy modulators targeting adipogenesis remain unclear. In this study, we investigated whether normal cellular prion protein (PrP < C >) is involved in the modulation of autophagy and affects adipogenic differentiation in vivo and in vitro. Surprisingly, autophagy flux signals were activated in the adipose tissue of prion protein-deficient mice and PrP < C > deleted 3T3-L1 adipocytes. The activation of autophagy flux mediated by PrP < C > deletion was confirmed in the adipose tissue via transmission electron microscopy. Adipocyte differentiation factors were highly induced in prion protein-deficient adipose tissue and 3T3-L1 adipocytes. In addition, deletion of prion protein significantly increased visceral fat volume, body fat weight, adipocyte cell size, and body weight gain in Prnp-knockout mice and increased lipid accumulation in PrP < C > siRNA-transfected 3T3-L1 cells. However, the overexpression of prion protein using adenovirus inhibited the autophagic flux signals, lipid accumulation, and the PPAR-γ and C/ EBP-α mRNA and protein expression levels in comparison to those in the control cells. Our results demonstrated that deletion of normal prion protein accelerated adipogenic differentiation and lipid accumulation mediated via autophagy flux activation.

1. Introduction The prevalence of obesity is increasing rapidly, and obesity has become a globally widespread disorder (Lei et al., 2007; O'Neill and O'Driscoll, 2015; Yatsuya et al., 2014). Additionally, metabolic syndromes, including type II diabetes and fatty liver, develop from obesity (Alexandre et al., 2014; Kloting and Bluher, 2014; Lopez-Soldado et al., 2015; Boyle et al., 2009). Several studies have shown that obesity is caused by dysregulation of adipogenesis, including changes in the number of adipocytes (adipogenic differentiation) and adipocyte size (lipid accumulation) (Lafontan, 2014; Kim et al., 2010; Gregoire et al., 1998). Some reports have shown that a change in the number of adipocytes during aging may influence the accumulation of lipid observed in older individuals (Yanagiya et al., 2007; Kirkland et al., 2002). Thus, adipogenesis might be a key factor in the rise in obesity. Mesenchymal stem cells (MSCs) have the capacity to differentiate into chondrocytes, osteoblasts, or adipocytes (Kmiecik et al., 2015; Romagnoli and Brandi, 2014; Baghaban Eslaminejad and Malakooty Poor, 2014). Specific differentiation into various tissues is regulated by

a variety of differentiation factors according to MSC conditions (To et al., 2014; Augello and De Bari, 2010). In particular, the differentiation factors that influence adipogenesis include peroxisome proliferator-activated receptor γ (PPAR-γ), adiponectin, and the CCAAT/ enhancer binding proteins C/EBP-α, C/EBP-β, and C/EBP-γ, which are regulatory factors that induce adipogenic differentiation from MSCs (Yu et al., 2012; Theurich et al., 2007; Qian et al., 2010). Autophagy is the main mechanism for degradation of cell organelles and protein aggregates via the lysosomal pathway (Eskelinen and Saftig, 2009; Mizushima, 2007; Pan et al., 2008) and is initiated by the formation of double-membrane vesicles, known as autophagosomes, which engulf part of the cytoplasm. The autophagosome fuses to a lysosome for degradation of the load (Mizushima, 2007; Glick et al., 2010). Thus, autophagy plays a pivotal role in adaptation to starvation, immunity, anti-cancer effects, neuroprotection, and differentiation. Novikoff et al. showed that differentiated 3T3-L1 cells have an increased number of autophagosomes (Novikoff et al., 1980). Some reports have shown that adipose tissue-specific knockout of autophagyrelated genes (ATGs) suppresses adipogenesis and lipid accumulation

Corresponding author. College of Veterinary Medicine, Chonbuk National University, Iksan, Jeonbuk, 54596, Republic of Korea. E-mail address: [email protected] (S.-Y. Park). 1 These authors have equally contributed to this work. ∗

https://doi.org/10.1016/j.mce.2018.11.013 Received 24 August 2018; Received in revised form 6 November 2018; Accepted 23 November 2018 Available online 01 December 2018 0303-7207/ © 2018 Elsevier B.V. All rights reserved.

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(Singh et al., 2009; Zhang et al., 2009). Cellular prion protein (PrP < C >) is the prion protein isoform found in normal tissues (Mediano et al., 2014; Llorens et al., 2013). It is a copper-binding glycosylate-phosphatidylinositol-anchored membrane protein that has various physical functions, including cellular uptake of copper ions, transmembrane signaling, protection against apoptotic signaling and regulation of autophagy (Oh et al., 2008; Barbieri et al., 2011; Shin et al., 2013; Wang et al., 2012; Li et al., 2011). Further studies have shown that Stress-inducible protein-1(STI-1) is the ligand for the PrP < C > , which promote stroke recovery into the ischemic brain (Americo et al., 2007). And PrP < C > regulates special AT-rich sequence-binding protein-1 (SATB1) and glucose transporter 1 (Glut1) expression to support colorectal cancer cells via promotion of Fyn signaling. Also, Fyn-overexpression induces autopahgic inhibition in skeletal muscle (Wang et al., 2012; Li et al., 2011). Also, some studies have suggested that PrP < C > exerts a neuroprotective effect by regulating autophagy (Oh et al., 2008; Barbieri et al., 2011). Oh et al. showed that PrP < C > prevents the activation of autophagy following serum deprivation in PrnP−/− hippocampal neurons compared to that in PrnP+/ + hippocampal neurons (Oh et al., 2008). Additionally, inhibiting PrP < C > expression in glial and non-glial cancer cells induces autophagy-dependent cell death (Barbieri et al., 2011). These studies suggest that PrP < C > is a key factor negatively regulating the activation of autophagy via modulation of transmembrane signaling. PrP < C > is expressed not only in neuronal tissues, but also in some extra-neuronal tissues, such as hematopoietic, mesenchymal, and lymphatic tissues, in lesser amounts (Mediano et al., 2014; Peralta and Eyestone, 2009; Liu et al., 2001). Some reports have shown that PrP < C > influences differentiation in various tissues (Lima et al., 2007; Dodelet and Cashman, 1998). Flavia et al. suggested that astrocytes obtained from the brains of Prnp-knockout mice were induced at a lower level of neuritogenesis than astrocytes from the brains of wildtype mice (Lima et al., 2007). In addition, lymphocytes and monocytes do not change the expression of PrP < C > during differentiation; however, granulocytes decrease the expression of PrP < C > in lymphoid tissues (Dodelet and Cashman, 1998). Mesenchymal stem cells, the origin of adipocytes, also express PrP < C > proteins (Mohanty et al., 2012). These observations suggest that modulating PrP < C > expression may influence adipogenic differentiation. However, the relationship between PrP < C > expression and adipogenesis is unclear. Thus, the present study focused on the influence of the PrP < C > related autophagy pathway on adipogenesis. The results show that adipogenesis and autophagy increased in systemic Prnp−/− mice compared to those in wild-type mice. Additionally, the overexpression of the Prnp gene inhibited adipogenesis and autophagy in adipocytes. These results suggest that controlling the negative regulation of the autophagy pathway by PrP < C > may be a viable therapeutic strategy for metabolic disorders such as diabetes and obesity.

(Fig. 1g) and micro-CT imaging of mouse fat volume were significantly higher in Prnp-knockout mice than those in normal mice after 21 weeks (Fig. 1h). The adipose tissue volume analysis indicated that depleting Prnp increased the visceral adipose tissue volume (Fig. 1i). A histological analysis showed that gonadal white adipose tissue from Prnpknockout mice contained significantly larger adipocytes than those of control mice (Fig. 2a and b). Moreover, a quantitative analysis of the adipocyte distribution and size showed clear differences, in which deletion of PrP < C > induced large adipocytes mediated by lipid droplet accumulation and decreased adipocyte number in the Prnp-knockout mice (Fig. 2c–e). These results provide further evidence that the alleviation of PrP < C > expression may up-regulate adipogenesis and weight gain. 2.2. Activation of autophagic flux caused by inhibition of PrP < C > expression was sufficient to trigger adipogenic differentiation Next, we evaluated the influence of Prnp expression on the autophagic flux pathway in the adipose tissue and the relationship between the two (Fig. 3). We first examined the influence of Prnp expression on autophagic flux in the adipose tissue. Adipose and brain tissues of 10week-old Prnp-knockout mice did not express PrP < C > proteins (Fig. 3a and b). At 10 weeks of age, Prnp-knockout mice showed increased LC3-II protein levels and decreased p62 protein levels compared to the FVB control mice (Fig. 3c). The pattern of LC3 and p62 protein expression did not change in adipose tissues of 21-week-old Prnp-knockout mice (Fig. 3d). These results were confirmed by measuring autophagic vesicles (AVs) and lipid-containing autolysosomes using transmission electron microscopy (Fig. 3e and f). The 21-week-old Prnp-knockout mice showed more typical autophagic vesicles with double membranes and autolysosomes compared to the FVB control mice (Fig. 3e and f). Taken together, these data suggest that downregulation of PrP < C > may increase autophagic flux in adipose tissue. Some reports have shown that adipogenic differentiation is regulated by the modulation of autophagic flux (Novikoff et al., 1980; Singh et al., 2009; Zhang et al., 2009). Thus, we investigated the influence of Prnp depletion on adipogenic differentiation in adipose tissues. First, we assessed the expression of adipogenesis markers PPAR-γ and C/EBP-α in adipose tissues of Prnp-knockout mice compared to that in FVB control mice. PPAR-γ mRNA and protein levels increased in 10-weekold Prnp-knockout mice (Fig. 4a and c). However, the expression of C/ EBP-α, a late-stage marker of adipogenesis, decreased the protein and mRNA levels in 10-week-old Prnp-knockout mice (Fig. 4b and c). PPARγ protein, C/EBP-α protein, and mRNA expressions increased in 21week-old Prnp-knockout mice compared to that in control FVB mice (Fig. 4d–f). Consistent with these, PPAR-γ, C/EBP-α, and adiponectin mRNA expressions increased and PrP < C > mRNA decreased in PrP < C > siRNA-transfected MDI-treated adipocytes (Fig. 5a and d). A western blot analysis showed that PrP < C > protein levels decreased in adipocytes treated with si-PrP < C > and that PrP < C > knockdown up-regulated the expression of PPAR-γ, C/EBP-α, LC3-II and p62 proteins (Fig. 5e and f). Moreover, PrP < C > siRNA-transfected 3T3L1 cells showed increased lipid accumulation induced by treatment of differentiation-medium (MDI), which contains a mixture of 3-isobutyl1-methylxanthine (IBMX), dexamethasone and insulin, compared to NCe transfected cells (Fig. 5g). These results indicated that knockdown of PrP < C > increased the adipogenic differentiation and autophagic flux. However, the link between PrP < C > expression and autophagic flux in adipocytes is not clear. Thus, we next examined the relationship between PrP < C > expression and autophagic flux in adipogenesis. PrP < C > siRNA-transfected 3T3-L1 cells showed increased lipid content compared to NC-transfected cells (Fig. 6a and b). However, the differentiation effect caused by the knockdown of PrP < C > was blocked by treatment with autophagy inhibitors 3-MA and chloroquine (CQ) (Fig. 6a and b). Western blot analysis showed that the expression levels of LC3-II and p62 proteins decreased in adipocytes treated with

2. Results 2.1. Prnp depletion markedly increased weight gain and adipose tissue volume In a previous study, depleting Prnp influenced autophagic flux in a hippocampal neuron cell line, but the relationship between autophagic flux and PrP < C > has not been studied in adipose tissue (Oh et al., 2008). Thus, we evaluated body weight gain, food intake, and body composition to determine whether depleting Prnp influenced adipogenic differentiation and body fat accumulation. Weight gains by 10week-old Prnp-knockout old mice were similar to those of control mice. FVB mice at 10 weeks weighed 26 ± 1.4 g, whereas Prnp-knockout mice weighed 25.7 ± 1.5 g (Fig. 1a and c). Daily food intake remained unchanged even after depleting Prnp (Fig. 1d). Fat mass weight (Fig. 1e) and body composition (Fig. 1f) in Prnp-knockout mice were also similar to those in normal mice after 21 weeks. However, gonadal fat size 85

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Fig. 1. Body weight gain significantly increased in Prnp−/− mice compared to Prnp+/+ mice. Prnp+/+ and Prnp−/−mice following euthanization were captured showing the appreciable differences in body size following 21 wk of normal diet (a). Changes in body weight were measured during the normal diet in Prnp+/+ and Prnp−/−mice and C57BL/6J mice for 21wks (b). Cumulative weight gain was measured from 9 wk to 15 wk in normal diet (c). Daily food intake based on consumed food weight (d). Bar graph indicates the mean ± S. E. M. (n = 4–8). The data were analyzed using student's ttest. **p < 0.01, significant differences between Prnp+/+ and Prnp−/−group. After euthanization, body fat weight and composition were measured by Body composition analyzer in Prnp+/+ and Prnp−/ − mice at 21 wk (n = 4–10) (e and f). Representative gonadal fat pads were captured from the 2 types of mice (g). Micro-CT image of visceral and subcutaneous fat (bottom panel) of mice were measured after 21 wk in Prnp+/+ and Prnp−/−mice. Arrows indicate gonadal adipose tissue (h). Mouse adipose tissue volume was analyzed by data analysis program (CTAn) from Micro CT. Arrows indicates adipose tissue area (i). Bar graph indicates the mean ± S. E. M. (n = 3). The data were analyzed using student's ttest. **p < 0.01, significant differences between Prnp+/+ and Prnp−/−group.

Fig. 2. Effects of PrP < C > deficient on adipose tissue in mice fed for 21 weeks. After euthanization, gonadal WAT was gained from Prnp+/+ and Prnp−/−mice at 21 weeks. Sections of WAT were photographed with a light microscope (a and b) and number of adipocytes per area was measured using Image J program (c). Bar graph displayed Frequency distribution of adipocyte surface area from figure A and B (d and e). Bar graph indicates the mean ± S. E. M. (n = 4–8). The data were analyzed using student's t-test. **p < 0.01, significant differences between Prnp+/+ and Prnp−/−group.

autophagy inhibitors 3-MA and CQ, and that the expression of PPAR-γ and C/EBP-α proteins decreased in PrP < C > siRNA-transfected 3T3L1 cells (Fig. 6c). Also, the immunocytochemistry assay results showed

that treatment with CQ increased p62 protein levels, indicative of autophagic inhibition, as compared to that in PrP < C > si-RNA-transfected cells (Fig. 6d). These results indicate that suppressing PrP < 86

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Fig. 3. Suppression of PrP < C > activated autophagy flux in adipose tissue and adipocyte. Western blotting of proteins isolated from the indicated adipose tissues and brain from Prnp+/+and Prnp−/− mice. PrP < C > protein levels were evaluated in adipose and brain tissue at 10 wk (a and b). LC3 and p62 protein levels were measured in adipose tissue at 10 wk (c) and at 21 wk (d). Bar graph indicated the averages of LC3 and p62 ratio (c and d). Gonadal Adipose tissue from Prnp+/+ (e) and Prnp−/− (f) mice were processed and examined by TEM. Asterisk (*) indicates lipid. Arrow head (▲) indicates double membrane of autophagic vesicle. Arrow (↑) indicates lipid-containing autolysosome.

C > expression activates autophagic flux and this induces pro-adipogenic effects in adipocytes.

demonstrate that overexpressing PrP < C > using adenoviral vectors inhibits adipogenic differentiation, indicating that PrP < C > may play a pivotal role in regulating adipogenic differentiation.

2.3. Limitation of autophagic flux mediated by PrP < C > overexpression suppressed adipogenic differentiation

3. Discussion

We used adenoviral vectors to determine the effects of PrP < C > overexpression on autophagic flux in 3T3-L1 preadipocytes. The cells were infected with adenoviruses expressing PrP < C > or an empty vector at an MOI of 500 and treated with MDI. 3T3-L1 preadipocytes showed increased LC3-II levels and decreased p62 protein levels compared to those in MDI treatment groups. Moreover, decreased LC3-II (Fig. 7d) and increased p62 protein levels were observed after a 2-day MDI treatment in 3T3-L1 adipocytes over-expressing PrP < C > via adenoviral vectors compared to those in control cells (Fig. 7d). Furthermore, overexpression of PrP < C > using adenoviral vectors reduced triglyceride accumulation and lipid content induced by MDImediated adipocyte differentiation, whereas empty vector-overexpressing cells accumulated lipids at the level of the MDI control (Fig. 7a–c). The real-time PCR data showed that overexpression of PrP < C > reduced the mRNA levels of PPAR-γ, C/EBP-α and adiponectin to those of the MDI control (Fig. 8a and c). Consistent with these results, PrP < C > overexpression blocked the PPAR-γ and C/EBP protein expression induced by MDI treatment (Fig. 8d). These results

Our results demonstrate that depleting Prnp induced autophagy and that increasing autophagic flux by depleting PrP < C > increased adipogenic differentiation in the adipose tissue. Notably, PrP < C > mediated regulation of autophagic flux was related to adipose tissue differentiation, which, in turn, conferred regulation of adipogenesis. Some studies have suggested that PrP < C > expression is related to the proliferation and differentiation of various cells, including stem cells, leukocytes, and mesenchymal cells (Mediano et al., 2014; Dodelet and Cashman, 1998; Mohanty et al., 2012; Heikenwalder et al., 2008; Peralta et al., 2011; Mouillet-Richard et al., 1999; Zhang et al., 2011). Depleting PrP < C > decreases the differentiation of embryonic stem and neural stem cells into neuronal progenitors (Peralta et al., 2011; Mouillet-Richard et al., 1999). PrP < C > expression does not changes in human leukocytes during the differentiation of lymphocyte or monocyte lineage, however, the expression level of PrP < C > decreased in granulocyte differentiation (Dodelet and Cashman, 1998). In addition, undifferentiated dental mesenchymal cells from Prnpknockout mice molars increase more rapidly than molar dental 87

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Fig. 4. Expression of adipocyte differentiation factors increased in Prnp−/−mice. The mRNA expression of PPAR-γ and C/EBP-α was analyzed by real-time RT-PCR in Prnp+/+ and Prnp−/−mice at 10 wk (a and b). Western blotting of proteins isolated from the indicated adipose tissues from Prnp+/+ and Prnp−/−mice at 10 wk. PPAR-γ and C/EBP- α protein levels were evaluated. β-actin was used as loading control. Bar graph indicated the averages of PPAR- γ and C/ EBP-α ratio (c). The mRNA expression of PPAR-γ and C/EBPα was analyzed by real-time RT-PCR in Prnp+/+ and Prnp−/ − mice at 21 wk (d and e). Western blotting of proteins isolated from the indicated adipose tissues from Prnp+/+ and Prnp−/−mice at 10 wk. PPAR-γ and C/EBP- α protein levels were evaluated (f). Bar graph indicates the mean ± S. E. M. (n = 3); each experiment was performed in triplicate. The data were analyzed using student's t-test. *p < 0.05, **p < 0.01, significant differences between Prnp+/+ and Prnp−/−group.

mesenchymal cells in wild-type mice (Zhang et al., 2011). Thus, regulation of PrP < C > expression may be a key factor during the differentiation of various cell types. Our results show more rapid increases in body and lipid contents in Prnp-knockout mice than those in wildtype FVB mice at 21 weeks of age (Figs. 1 and 2), suggesting that depleting PrP < C > may increase adipogenic differentiation. Several lines of evidence suggest that adipogenic differentiation is regulated by autophagic flux (Zhang et al., 2009; Singh et al., 2009; Nunez et al., 2013). Knockdown of ATG5 and ATG7 in 3T3-L1 preadipocytes blocks lipid deposition and decreases the expression of adipocyte differentiation factors, including C/EBP-α (Singh et al., 2009). In addition, targeted deletion of ATG5 and ATG7 in mice inhibits adipogenesis and lipid accumulation, and adipocyte-specific depletion of ATG7 in mice lead to similar results (Zhang et al., 2009). We hypothesized that depleting PrP < C > increases adipose differentiation by regulating autophagic flux. Our results show that the levels of autophagy markers LC3-II increased and p62 protein levels decreased in Prnp-knockout mice compared to those in wild-type mice, regardless of age (Fig. 3c and d). Additionally, the number of autophagic vesicles with double membranes and autolysosomes increased in the adipose tissue of 21-week-old Prnp-knockout mice compared to those in the adipose tissue of control mice (Fig. 3e and f). Furthermore, PrP < C > knockdown 3T3-L1 preadipocytes cultured in adipocyte induction media increased lipid accumulation (Fig. 5g) and LC3-II protein levels, and decreased p62 protein expression levels (Fig. 5f). However, the upregulation of adipose differentiation caused by PrP < C > knockdown

was inhibited by treatment with the autophagy inhibitors 3-MA and CQ (Fig. 6). In addition, the protein levels of LC3-II increased and those of p62 decreased in PrP < C > -knockdown 3T3-L1 preadipocytes cultured in adipocyte induction media, whereas overexpressing PrP < C > in 3T3-L1 cells blocked adipogenic differentiation caused by the adipogenic induction media (Fig. 7a–c). These observations support the hypothesis that PrP < C > expression influences adipogenic differentiation by regulating autophagic flux. The accumulation of lipid in adipose tissue requires the differentiation of preadipocytes into adipocytes, which is tightly controlled by the adipogenic factors PPAR-γ and C/EBP-α (Yu et al., 2012; Theurich et al., 2007; Eeckhoute et al., 2012; Tontonoz et al., 1994; Rosen et al., 2002). In particular, PPAR-γ is the main regulator of adipogenic differentiation, and overexpressing PPAR-γ induces adipogenic differentiation in fibroblasts; however, no change in adipogenesis differentiation occurs when PPAR-γ is depleted (Rosen et al., 2002). Protein and mRNA levels of the adipogenic differentiation marker PPAR-γ increased in Prnp-knockout mice compared to those in control mice (Fig. 4). Furthermore, C/EBP-α mRNA and protein levels in adipose tissues from 21-week-old Prnp-knockout mice increased (Fig. 4e and f). Additionally, deletion of prion protein increased adipocyte size rather than cell number (Fig. 2) and PrP < C > knockdown of 3T3-L1 preadipocytes increased lipid accumulation (Fig. 5g), mRNA levels (PPAR-γ, C/EBP-α, and adiponectin) and protein levels (PPAR-γ and C/ EBP-α) after inducing adipocyte differentiation (Fig. 5). In addition, decreased lipid accumulation (Fig. 7a–c) and PPAR-γ, C/EBP-α, and 88

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Fig. 5. Knockdown of PrP < C > enhanced the adipocyte differentiation. 3T3-L1 cells were transfected with either PrP < C > siRNA or negative control (NC) siRNA for 24 h, and then cells were induced to differentiate for 2 days and harvested at day 2 during the differentiation period. The mRNA expression of C/EBP- α (a), PPAR-γ (b), Adiponectin (c) and PrP < C > (d) were analyzed by real-time RT-PCR. Bar graph indicates the mean ± S. E. M. (n = 3); each experiment was performed in triplicate. 3T3-L1 cells were transfected with either PrP < C > siRNA or negative control (NC) siRNA for 24 h, and then cells were induced to differentiate for 2 days. PrP < C > , PPAR-γ, C/EBP- α, LC3 and p62 protein levels were evaluated with Western blotting. β-actin was used as loading control (e and f). Bar graph indicated the averages of LC3 and p62 ratio (f). 3T3-L1 cells were transfected with either PrP < C > siRNA or negative control (NC) siRNA for 24 h, and then cells were induced to differentiate for 7 days. The AdipoRed assays were performed on day 7 and were photographed with a light microscope ( × 200) (g). Each experiment was performed in triplicate.

adiponectin levels (Fig. 8) were observed in PrP < C > -overexpressing 3T3-L1 cells cultured in adipogenesis induction media. These observations suggest that the regulation of PrP < C > expression may influence adipogenic differentiation. A previous study suggested that adipogenic differentiation involves the interaction between transcription factors C/EBPs and PPAR-γ (Yu et al., 2012; Theurich et al., 2007; Qian et al., 2010, Hishida et al., 2009). C/EBP-β and δ play a pivotal role in activating PPAR-γ expression during the early stages of adipogenesis (Hishida et al., 2009; Park et al., 2012). PPAR-γ induces the expression of C/EBP-α in the late stage of adipocyte differentiation (Hishida et al., 2009). Through a positive feedback loop, C/EBP-α up-regulates the expression of PPAR-γ. PPAR-γ and C/EBP-α cooperate to promote adipocyte differentiation, including adipocyte gene expression and lipid accumulation (Park et al., 2012). A recent study showed that activation of autophagy decreased C/EBP-α expression in liver tissue and hepatic stellate cells (HSCs), and increased PPAR-γ activations in adipose tissues (Ho et al., 2017). Our results showed that mRNA and protein levels of the terminal stage adipogenic differentiation marker C/EBP-α decreased in adipose

tissues of 10-week-old Prnp-knockout mice compared to those in control mice (Fig. 4b and c). Moreover, body contents in Prnp-knockout mice were comparable to those in wild-type FVB mice at 10 weeks of age. However, 21-week-old Prnp-knockout mice showed increased lipid accumulation (Fig. 2), mRNA levels (PPAR-γ, C/EBP-α, and adiponectin) and protein levels (PPAR-γ and C/EBP-α). These data suggest that depletion of PrP < C > might regulate adipogenic differentiation, especially adipocyte size rather than cell number. An understanding of its mechanisms will require further investigation. Recent studies suggest that Prnp expression regulates autophagic flux in various brain cells (Oh et al., 2008; Barbieri et al., 2011; Shin et al., 2014; Oh et al., 2012). The hippocampal CA3 region and cerebral cortical neuropils of Zürich I Prnp-deficient mice have impaired autophagic flux (Shin et al., 2014). Oh et al. have suggested that H2O2induced oxidative stress impairs autophagic flux in Prnp−/− hippocampal cells, whereas Prnp+/+ cells exposed to H2O2 show increased autophagic flux (Oh et al., 2012). However, other studies have suggested that depleting Prnp increases autophagic flux in neurons (Oh et al., 2008). Serum depriving Prnp−/− hippocampal neural cells 89

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Fig. 6. Knockdown of PrP < C > increased the adipogenesis via activation of autophagic flux in adipocytes. 3T3-L1 cells were transfected with either PrP < C > siRNA or negative control (NC) siRNA for 24 h, and then cells were induced to differentiate for 7 days. The AdipoRed assays were performed on day 7 and were photographed with a light microscope ( × 200) (a). Fluorescence was measured with excitation at 485 nm and emission at 572 nm (b). Bar graph indicates the mean ± S. E. M. (n = 3). *p < 0.05, significant differences between control and each treatment group. The data were analyzed using student's t-test. 3T3-L1 cells were transfected with either PrP < C > siRNA or negative control (NC) siRNA for 24 h, and then cells were induced to differentiate for 2 days. PrP < C > , PPAR-γ, C/EBP- α, LC3 and p62 protein levels were evaluated by Western blotting. β-actin was used as loading control (c). Each experiment was performed in triplicate. The adiopocytes were immunostained with p62 antibody (green) and DAPI (blue) was examined. The fluorescence intensity of p62 was measured using image J (d).

increases LC3-II expression and accumulation of autophagosomes compared to those in wild-type cells (Oh et al., 2008). This up-regulation of autophagic activity is delayed by overexpressing Prnp in Prnp−/ − hippocampal cells (Oh et al., 2008). Additionally, knockdown of PrP < C > expression by DNA-antisense oligonucleotides increases LC3-II and decreases p62 protein levels (Barbieri et al., 2011). Thus, the influence of PrP < C > on autophagic flux is thought to differ by cell type. However, the relationship between autophagy and PrP < C > has not been studied, except in neurons. We examined the relationship between PrP < C > expression and autophagy during adipogenic differentiation. Depleting PrP < C > induced adipogenic differentiation by activating autophagic flux. Yamada et al. phosphorylation of Fyn suppressed autophagic flux by inhibiting AMP-dependent protein kinase (AMPK) activation

(Yamada et al., 2016). Also, some paper suggested that PrP < C > regulates transmembrane signaling by modulation of STI-1 and Fyn kinase activity, respectively (Americo et al., 2007; Larson et al., 2012). Thus, Further detailed studies of the PrP < C > -related downstream signaling network, including Fyn and STI-1, will be important in understanding the mechanism of adipogenic differentiation via autophagic flux caused by depleting PrP < C > in adipose tissue. To the best of our knowledge, this is the first report demonstrating that activating autophagy caused by depleting PrP < C > may be a key factor during the adipogenic differentiation of adipose tissue. These results suggest that the regulation of PrP < C > is the main mechanism of adipogenic differentiation caused by autophagic flux and that regulating PrP < C > expression may be a clinical therapeutic strategy for obesity. 90

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Fig. 7. Overexpression of PrP < C > blocked the adipocyte differentiation. 3T3-L1 cells infected with adenoviruses expressing empty vector (Adcon) or Prnp for 48 h were induced to differentiate for 7 days. AdipoRed assays were performed as described Fig. 1a. (a) Fluorescence was measured with excitation at 485 nm and emission at 572 nm (b). Triglyceride (TG) assay was assessed on day 7, and TG contents relative to the control were measured (c). Bar graph indicates the mean ± S. E. M. (n = 3). 3T3-L1 cells infected with adenoviruses expressing empty vector (Adcon) or Prnp for 48 h were induced to differentiate for 2 days and harvested at day 2 during the differentiation period. The protein levels of LC3, p62 and PrP < C > were analyzed by Western blotting. β-actin was used as loading control (d). Bar graph indicates the mean ± S. E. M. (n = 3). The data were analyzed using student's t-test. *p < 0.05, **p < 0.01, significant differences between control and each treatment group.

4. Materials and methods

4.3. Micro-computed tomography

4.1. Animal models

Scanning was performed using a cone-beam type in vivo micro-CT scanner (Skyscan model 1076; Skyscan). All animals were scanned under anesthesia (injection of xylaxine, 10 mg/kg). The acquisition settings were as follows: X-ray source voltage 50 kVp, current 200 A; a 1-mm thick aluminum filter was used for beam hardening reduction. The pixel size was 35 μm, the exposure time was 4.7 s, and the rotation step was 0.6° with a complete rotation over 360°.

Control mice (FVB/NJ, 4 weeks of age, stock no: 001800) and FVB.129S7(B6)-Prnptm1Cwe/J (stock no: 018122) were purchased from the Jackson Laboratory (Bar Harbor, ME, USA) and housed in a temperature-controlled (21 °C) room with a 12:12-h light-dark cycle. The mice were fed normal chow (D12450B, 10% fat, Research Diets). All protocols were established based on the Guide for the Care and Use of Laboratory Animals (2011-0008) and approved by the Chonbuk National University Institutional Animal Care and Use Committee.

4.4. Histological analysis Morphometric analysis of adipocytes was performed on normal sections using NIH ImageJ software. Perimeters of the individual adipocytes, identified by their characteristic homogenous density, were delineated with the free-hand selection tool, and the mean perimeter value of at least 25 adipocytes per section was determined. The total

4.2. Body composition Whole body fat compositions of live animals were determined by using an NMR analyzer (Minispec, Bruker Optic, Billerica, MA, USA). 91

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Fig. 8. Overexpression of PrP < C > inhibited the expression of adipogenesis-related factors in adipocytes. Cells infected with adenoviruses expressing empty vector (Adcon) or Prnp for 48 h were induced to differentiate for 2 days and harvested at day 2 during the differentiation period. The mRNA expression of C/EBP-α (a), PPAR-γ (b) and Adiponectin (c) were analyzed by real-time RT-PCR. Bar graph indicates the mean ± S. E. M. (n = 3). 3T3-L1 cells infected with adenoviruses expressing empty vector (Adcon) or Prnp for 48 h were induced to differentiate for 2 days and harvested at day 2 during the differentiation period. The protein levels of PrP < C > , PPAR-γ and C/EBP-α were analyzed by Western blotting (d). β-actin was used as loading control. Bar graph indicates the mean ± S. E. M. (n = 3). The data were analyzed using student's t-test. *p < 0.05, **p < 0.01, significant differences between control and each treatment group.

number of adipocyte was quantified with the analyzed particle tool and then normalized to the number of adipocyte nuclei per section. Adipocyte was scored by visual counting of individual adipose tissue.

insulin and 10% FBS. 3-MA and chloroquine (CQ) were added at the time of the differentiation induction. The AdipoRed Assay was performed on day 7. And qPCR and western blot was performed on two days after treatment of MDI induction media.

4.5. Transmission electron microscope (TEM)

4.7. Quantification of lipid content

After fixation of samples in 2% glutaraldehyde (EMS, USA) and 2% paraformaldehyde (EMS, USA) in 0.05 sodium carcodylate buffer (pH7.2) (EMS, USA), specimens were post fixed in 1% osmium tetroxide (EMS, USA), dehydrated in graded ethanol and propylene oxide (EMS, USA). The cells were embedded in Epoxy resin (Embed 812, NMA; Nadic methyl anhydride, DDSA; Dodenyl Succinic Anhydride, DMP-30) (EMS, USA). Ultrathin sections were cut on an LKB-III ultratome (LEICA, Austria) and were stained with 0.5% uranyl acetate (EMS, USA) and lead citrate (EMS, USA). Images were recorded on a Hitachi H7650 electron microscope (Hitachi, Ltd., Tokyo, Japan; magnification, x10,000) installed at the Center for University-Wide research Facilities (CURF) at Chonbuk National University.

Lipid content was quantified using a commercially available AdipoRed Assay Reagent (Lonza, Portsmouth, NH, USA), according to the manufacturer's instructions. In brief, preadipocytes were grown in 24-well plates and incubated in MDI medium alone or supplemented with test compounds during the adipogenic phase. The culture supernatant was removed on day 7, and the cells were carefully washed with 500 μl phosphate-buffered saline (PBS). The wells were filled with 300 μl PBS, and 30 μl of AdipoRed reagent was added and incubated for 10 min at 37 °C. Fluorescence was measured at an excitation wavelength of 485 nm and an emission wavelength of 572 nm. 4.8. Measurement of triglyceride content in adipocytes

4.6. Cell culture and differentiation

3T3-L1 adipocytes were harvested 7 days after the initiation of differentiation. After treatment, the medium was removed and cell extracts were used for TG determination. Mature cells were washed extensively with phosphate-buffered saline (PBS) and scraped on ice in 500 μl of sonication buffer (25 mM, Tris buffer and 1 mM ethylenediaminetetraacetic acid (EDTA), pH 7.5), and sonicated to homogenize the cell suspension. For protein determinations, cells were lysed in 0.3N NaOH, 0.1% SDS. The total TG content in cells was determined using TG determination kit (Sigma, St. Louis, MO, USA). Protein measurements were performed using the BCA reagent. TG content results were

3T3-L1 cells were maintained in Dulbecco's modified Eagle's medium (DMEM) containing 10% calf serum, and antibiotics (100 μg/ mL gentamycin and 100 μg mL−1 penicillin-streptomycin; Sigma, St. Louis, MO, USA). Two-day post confluent 3T3-L1 cells were incubated in MDI induction medium (DMEM containing 10% fetal bovine serum and 0.5 mM 3-isobutyl-1-methylxanthine; Sigma, St. Louis, MO, USA), 1 μM dexamethasone (Sigma, St. Louis, MO, USA), and 1 μg/mL insulin; Sigma, St. Louis, MO, USA) for 2 days to induce differentiation. Two days after MDI, the medium was changed to DMEM containing 1 μg/mL 92

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obtained as mmol glycerol/mg protein and were expressed as the ratio (%) to the control value.

4.13. Construction of recombinant adenoviruses PrP < C > -expressing adenoviruses and empty vector adenoviruses were produced from Genenmed (Genenmed, Seoul, Korea). Recombinant adenoviruses were amplified in human embryonic kidney (HEK-293) cells and purified using the Vivapure AdenoPACK Kit (Sartorius AG, Göttingen, Germany) according to the manufacturer's instructions (Seo et al., 2012).

4.9. Quantitative real-time polymerase chain reaction (qRT-PCR) Total RNA was extracted from 3T3-L1 cells treated with 18β-glycyrrhetinic acid using the Easy-Spin™ Total RNA extraction kit (GeneAll, Seoul, Korea). cDNA synthesis was carried out following the instructions of the TaKaRa Prime Script TM 1st strand cDNA synthesis kit (TaKaRa Bio, Shiga, Japan). A 1 μl aliquot of SYBR Green gene primers (Bio-Rad Laboratories, Hercules, CA, USA) in a 20 μl reaction volume was used for qRT-PCR. The primer sequences used for the realtime PCR were as follows:

4.14. Statistical evaluation All data are expressed as mean ± standard error, and the data were compared using Student's t-test, analysis of variance, and Duncan's test using the SAS statistical package (SAS Institute, Cary, NC, USA). A P < 0.05 was considered significant.

PPAR-γ (forward 5′CGGAAGCCCTTTGGTGACTTTATG3′, reverse 5′ GCAGCAGGTTGTCTTGGATGTC3′), C/EBP-α (forward 5′CGGGAACGCAACAACATCGC3′, reverse 5′ TGTCCAGTTCACGGCTCAGC3′), adiponectin (forward 5′TGACGGCAGCACTGGCAAG3′, reverse 5′ TGATACTGGTCGTAGGTGAAGAGAAC3′), and β-actin (forward 5′TGAGAGGGAAATCGTGCGTGAC3′, reverse 5′GCTCGTTGCCAATAGTGATGACC3′)

Ethical approval Ethical approval for the project was granted by the institutional review board of the Chonbuk National University. Conflicts of interest The authors declare no conflict of interest.

All reactions with iTaq SYBR Green Supermix (Bio-Rad Laboratories, Hercules, CA, USA) were performed on the CFX96 Realtime PCR Detection System (Bio-Rad Laboratories).

Author contributions J.K.J., J.H.L., S.W.K. and S.Y.P. designed, executed the study, analyzed data and wrote the manuscript; J.K.J., J.M.H., J.W.S. and S.Y.P. revised the manuscript and figures; All authors reviewed the manuscript.

4.10. Western blot 3T3-L1 cells were lysed in lysis buffer (25 mM HEPES; pH 7.4, 100 mM NaCl, 1 mM EDTA, 5 mM MgCl2, 0.1 mM dithiothreitol, and protease inhibitor mixture). The proteins were electrophoretically resolved by 8–15% sodium dodecyl sulfate-polyacrylamide gel electrophoresis and immunoblotted as described previously. Images were captured using the Fusion FX7 acquisition system (VilbertLourmat, Eberhardzell, Germany). Densitometry of the signal bands was conducted using Bio-1D (VilberLourmat). The immunoblotting antibodies were LC3 (Cell Signaling Technology, Beverly, MA, USA), p62 (Millipore, Milford, MA, USA), C/EBP-α (Santa Cruz Biotechnology, Santa Cruz, CA, USA), PPAR-γ (Santa Cruz Biotechnology, Santa Cruz, CA, USA), PrP < C > (Abcam, Cambridge, MA, USA), and β-actin (Sigma, St. Louis, MO, USA).

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