{'fraction_non_alphanumeric': 0.03751893424476638, 'fraction_numerical': 0.023575562201421523, 'mean_word_length': 4.956049040018506, 'pattern_counts': {'":': 0, '<': 3, '<?xml version=': 0, '>': 1, 'https://': 1, 'lorem ipsum': 0, 'www.': 6, 'xml': 0}, 'pii_count': 0, 'substrings_counts': 0, 'word_list_counts': {'cursed_substrings.json': 1, 'profanity_word_list.json': 0, 'sexual_word_list.json': 0, 'zh_pornsignals.json': 0}}
The methylation status of the chemerin promoter region located from − 252 to + 258 bp regulates constitutive but not acute-phase cytokine-inducible chemerin expression levels

Chemerin is a chemoattractant protein with adipokine properties encoded by the retinoic acid receptor responder 2 (RARRES2) gene. It has gained more attention in the past few years due to its multilevel impact on metabolism and immune responses. However, mechanisms controlling the constitutive and regulated expression of RARRES2 in a variety of cell types remain obscure. To our knowledge, this report is the first to show that DNA methylation plays an important role in the cell-specific expression of RARRES2 in adipocytes, hepatocytes, and B lymphocytes. Using luciferase reporter assays, we determined the proximal fragment of the RARRES2 gene promoter, located from − 252 to + 258 bp, to be a key regulator of transcription. Moreover, we showed that chemerin expression is regulated in murine adipocytes by acute-phase cytokines, interleukin 1β and oncostatin M. In contrast with adipocytes, these cytokines exerted a weak, if any, response in mouse hepatocytes, suggesting that the effects of IL-1β and OSM on chemerin expression is specific to fat tissue. Together, our findings highlight previously uncharacterized mediators and mechanisms that control chemerin expression.

IL-1β and OSM stimulation upregulates RARRES2 expression in murine adipocytes but not hepatocytes. Next, we questioned how chemerin expression can be regulated in cells with apparent constitutive expression of RARRES2. WAT and the liver have been reported by multiple studies to be key sites of chemerin production 1 . Both organs contribute to fatty acid metabolism, respond to numerous cytokines and are involved in the pathogenesis and pathophysiology of obesity 23 which is linked to elevated systemic levels of chemerin 22 . Among the potential regulators of systemic chemerin levels are the acute-phase mediators OSM and IL-1β, which, as we showed previously, regulate chemerin levels in human epidermis-like cultures 2 . Moreover, liver and fat tissue are known to be responsive to both OSM and IL-1β [24][25][26][27] , suggesting that chemerin expression will be also regulated by these mediators in both adipocytes and hepatocytes. Treatment of epididymal white adipose tissue (eWAT)-derived mouse adipocytes with OSM and/or IL-1β resulted in statistically significant and comparable upregulation of RARRES2 mRNA by each stimulus (Fig. 1C). In parallel, secreted chemerin protein levels tended to be higher after 48-h of stimulation with the cytokines as compared with the control (Fig. 1D). In contrast with the adipocytes, downregulation of RARRES2 mRNA was detected in hepatocytes in response to OSM or OSM + IL-1β but not IL-1β alone (Fig. 1C). Likewise, cytokines did not affect the chemerin protein levels in hepatocyte-conditioned media (Fig. 1D). These in vitro results were corroborated by in vivo findings. RARRES2 mRNA was upregulated only in eWAT ( Fig. 1F) but not in the liver (Fig. 1G) after in vivo IL-1β + OSM administration. We confirmed that primary hepatocytes and mouse liver tissue responded to stimulation with IL-1β and OSM, since SAA3 mRNA levels, encoding an acute-phase protein, was markedly elevated (Fig. 1E,H). Together, these results suggest that the effect of cytokines that results in the upregulation of chemerin expression is specific to fat tissue.
Adipocytes are key cells expressing chemerin after stimulation with acute-phase cytokines. The stromal vascular fraction (SVF) of adipose tissue consists of a heterogeneous population of cells that includes adipocyte precursors, hematopoietic stem cells, endothelial cells, fibroblasts, and immune cells 28 . Therefore, we next asked whether adipocytes are the main cells that express chemerin in response to acute-phase cytokines. Indeed, we found a similar chemerin expression pattern and levels in both eWAT-derived adipocytes and in adipocytes derived from sorted SCA-1 + adipogenic progenitors ( Fig. 2A-D). Chemerin transcript levels continued to rise throughout a five-day time course. Stimulation with IL-1β and IL-1β + OSM yielded the highest mRNA levels (characterized by an approximately 12-to 13-fold increase relative to the oneday control) in both types of adipocyte cell culture ( Fig. 2A,C). However, statistically significant upregulation of chemerin protein levels was observed only in the eWAT derived adipocytes treated by IL-1β + OSM or in the cell culture of sorted adipogenic precursors following IL-1β and IL-1β + OSM stimulation. Accordingly, we concluded that IL-1β and OSM regulate chemerin expression primarily in adipocytes. Chemerin protein production was highest in response to IL-1β + OSM, suggesting the existence of additive effects. DNA methylation affects constitutive RARRES2 expression. We next investigated DNA methylation of the RARRES2 promoter region as a possible mechanism underlying differential 29 constitutive and Chemerin is constitutively expressed in the liver and WAT. Acute-phase cytokines upregulate chemerin expression in the adipocytes of WAT but not in hepatocytes. The Human Protein Atlas was used to compare human chemerin mRNA levels across multiple tissues and cells (A). B-cells, WAT, and liver tissue were chosen for further studies. Afterward, lymph nodes, liver tissue, and eWAT depots were excised from C57Bl6 mice and subjected to RT-QPCR analysis or isolation of B-cells, primary hepatocytes, or the SVF of eWAT. Relative chemerin mRNA levels across murine B-cells, liver tissue, and WAT are shown (B). SVF cells were differentiated to obtain a mature adipocyte cell culture. Then, the cells were treated with IL-1β (10 ng/mL), OSM (50 ng/mL), or a combination for 48 h. The levels of chemerin (C) and SAA3 (E) mRNA were determined using RT-QPCR. The relative expression of stimulated cells over the control is shown. Levels of secreted chemerin were determined in parallel in conditioned media by ELISA (D). Data are presented as the mean ± SD of at least three independent experiments. Statistical significance between the control and treated cells is shown by an asterisk; *p < 0.05 by ANOVA followed by a Bonferroni post-hoc test. In vivo, IL-1β and OSM were injected intraperitoneally at doses of 10 μg/ kg BW and 160 μg/kg BW, respectively. After 48 h, liver tissue and eWAT were isolated and subjected to RT-QPCR analysis. The levels of chemerin mRNA in eWAT (F) or liver tissue (G) and SAA3 (H) were determined. Data are presented as the mean ± SD of at least three independent experiments. Statistical significance between the control (PBS) and the cytokinetreated animals is indicated by an asterisk; *p < 0.05 by the two-tailed Student's t-test. All experiments were repeated at least three times. www.nature.com/scientificreports/ regulated chemerin expression in various cell types. We studied DNA fragments ranged ranging in location from − 735 to + 258 bp of the RARRES2 gene because promoter sequences are typically defined as being 100 to 1,000 bp upstream of the transcription start site and 100 bp downstream of the transcription start site 30 . Moreover, this DNA region contains all the binding sites of previously characterized transcription factors (TFs) of RARRES2 [18][19][20] .

Scientific RepoRtS
Using bisulfite sequencing we determined the methylation status of the murine RARRES2 gene in B-cells that do not produce chemerin, as well as that in unstimulated and cytokine-treated adipocytes and hepatocytes. We identified 17 CpG sites located within − 735/+ 258 bp of RARRES2 (Fig. 3A). However, a computational analysis of this sequence did not identify any CpG island (data not shown). The chemerin promoter was found to be highly methylated in B lymphocytes. In contrast, our results suggested a much lower methylation status of the chemerin promoter in unstimulated adipocytes and hepatocytes relative to in B-cells (Fig. 3B). Interestingly, the upregulation of chemerin expression after stimulation of adipocytes with IL-1β and OSM is correlated with the statistically significant increase in the average methylation level of the RARRES2 gene promoter within − 735/− 253 bp ( Fig. 3C) but not − 252/+ 258 bp (Fig. 3D). However, the methylation pattern of the chemerin promoter was not altered in the cytokine-treated but chemerin-unresponsive primary cultures of mouse hepatocytes. We concluded that DNA methylation plays a role in controlling the constitutive expression of chemerin and affects cytokine-regulated expression in adipocytes. the proximal part of the RARRES2 promoter is a key regulator of transcription. To test how cytokines or DNA methylation can regulate the transcription of RARRES2, we established the following three reporter constructs containing different portions of chemerin promoter sequences: Chemerin_Full (position − 735/+ 258 bp relative to the transcription start site), Chemerin_Proximal (position − 252/+ 258 bp), and Chemerin_Distal (position − 735/− 253 bp) (Fig. 4A). The promoter was divided into two parts based on the methylation levels of RARRES2 (Fig. 3B). The average percentage methylation levels for the proximal region was less than 14% in both unstimulated adipocytes and hepatocytes whereas the distal part of the promoter showed greater average methylation, varying from 28% (hepatocytes) to 40% (adipocytes). We used 3T3-L1 adipocyte precursors for transient transfection since these cells produce chemerin and respond to cytokine stimulation in a manner similar to that of differentiated adipocytes (Fig. 4B).

Figure 2.
Adipocytes are key cells expressing chemerin following stimulation with IL-1β + OSM. eWAT or SCA-1 + APs were isolated. The cells were differentiated to obtain a mature adipocyte cell culture. Then, the cells were treated with IL-1β (10 ng/mL), OSM (50 ng/mL), or a combination for up to five days. The levels of chemerin mRNA (A, C) were determined using RT-QPCR. The relative expression of stimulated cells over the control is shown. Levels of secreted chemerin were determined in parallel in conditioned media by ELISA (B, D). Data are presented as the mean ± SEM of at least three independent experiments. Statistical significance between the control and the treated cells is indicated by an asterisk; *p < 0.05 by analysis of covariance followed by a Bonferroni post-hoc test. www.nature.com/scientificreports/ First, we considered whether the Chemerin_Full construct, which covers all previously characterized TF binding sites [18][19][20] , responds to IL-1β + OSM stimulation. However, cytokine treatment did not reach statistical significance relative to in the untreated controls (Fig. 4C). To investigate whether inflammation-responsive TFs may bind to − 735/+ 258 bp of the RARRES2 promoter sequence in murine adipocytes and hepatocytes, we queried ChIP-Atlas 31 , a resource that gathers publicly available results from chromatin immunoprecipitationsequencing experiments. The results showed that inflammation-responsive TFs, including NFκB, AP-1, SAF-1, c-Jun/c-Fos, and STATs, do not bind to the − 735/+ 258 bp of the RARRES2 promoter sequence either in adipocytes or in hepatocytes (Table 1).
To determine which part of the RARRES2 promoter region is linked with constitutive chemerin expression, 3T3-L1 cells were transfected with the Chemerin_Full, Chemerin_Proximal, and Chemerin_Distal constructs, respectively. Chemerin_Proximal and Chemerin_Full showed the highest promoter activity as determined by luciferase assay (Fig. 4D). The Chemerin_Distal construct had similar promoter activity in comparison with the empty vector.
To investigate the role of DNA methylation in the regulation of chemerin expression, we excised the promoter sequence from pNL1.1[Nluc] constructs with restriction enzymes, treated the eluted DNA with SssI methylase or left it untreated, and relegated the methylated and unmethylated DNA inserts into the parent vector. Promoter activity of the methylated Chemerin_Proximal construct was five times lower than that of the unmethylated construct (Fig. 4E). In line with the results of bisulfite sequencing (Fig. 3), promoter activity of the methylated Chemerin_Distal construct was higher than that of the unmethylated construct.
Together, these data suggest that the DNA sequence of the RARRES2 promoter region located from − 252 to + 258 bp is critical for constitutive chemerin expression and that its methylation suppresses the transcription. www.nature.com/scientificreports/ Interestingly, the distal part of the chemerin promoter (position − 735/− 253 bp) shows minimal activity which is increased when DNA is methylated.

Discussion
Control of gene expression plays a key role in a variety of physiological and pathophysiological processes, ranging from cell differentiation, cellular stress responses, and immunity, to tissue homeostatis. Many mechanisms contribute to the regulation of gene expression to ensure coordinated cellular behaviors and fate decisions. This includes modifications of DNA (e.g. DNA methylation), binding of TFs to a gene promoter, alternative splicing, miRNAs, and many others 32 . To date, the molecular mechanisms controlling constitutive and regulated chemerin expression are still poorly understood 1 . Further, the functional significance of DNA methylation in the regulation of RARRES2 transcription has not yet been elucidated. www.nature.com/scientificreports/ Here, we determined, for the first time, that CpG sites located between − 735 and + 258 bp of the RARRES2 promoter are highly methylated in B-cells but not in hepatocytes and adipocytes. This suggests that DNA methylation controls constitutive chemerin expression in cells of different origins. B-cells do not secrete chemerin; meanwhile, hepatocytes and adipocytes are known to express high levels of chemerin and to release the chemerin protein 1 . This constitutive expression controls tissue homeostasis by affecting physiological processes like immune cell migration, angiogenesis, adipogenesis, and energy metabolism 33 . Epigenetic regulation by CpG methylation in the promoter region is often associated with both tissue-specific and heterogeneous expression of genes 29 . DNA methylation can cause transcriptional silencing of genes by inhibiting the binding of TFs to regulatory sequences 34 . Even the methylation status of a single CpG locus can modulate gene expression 35,36 . However, we have also observed variations in the DNA methylation pattern between the proximal (− 252/+ 258 bp) and distal (− 735/− 253 bp) parts of the RARRES2 promoter region in adipocytes and hepatocytes. Therefore, we divided the chemerin promoter into two parts to study activity. Luciferase assays revealed that the proximal part of the chemerin promoter is a key regulator of constitutive chemerin expression. Methylation of the CpG sites located within this region diminished luciferase activity. Finally, the distal part of the chemerin promoter is poorly characterized and did not show any important activity in luciferase assays. Taken together, the DNA methylation of the RARRES2 promoter region located from − 252 to + 258 bp regulates the constitutive expression of chemerin in a cell-type dependent manner.
The expression of chemerin may be also regulated by various mediators including IL-1β and OSM acute-phase cytokines. Both IL-1β, and OSM, are essential mediators of adaptive and innate immune responses 37,38 , and both have potential roles in the pathology of psoriasis 39,40 , rheumatoid arthritis 41 , cancer 42 , and obesity 43 , which are diseases with postulated chemerin involvement 6,21, 22,44 . We previously showed that IL-1β and OSM upregulated Table 1. TFs that may bind the RARRES2 promoter region located from − 735 to + 258 bp in murine adipocytes and hepatocytes. ChIP-Atlas 31 raw data linked to the selected records were downloaded and submitted to CRUNCH online tool 59 to identify proteins bound to the RARRES2 genomic loci with the binding score value of − 10 × log10[q] > 700. www.nature.com/scientificreports/ chemerin expression in human skin cultures 2 . Here, our research revealed that a combination of both cytokines considerably increased RARRES2 mRNA and protein levels in cell cultures of murine eWAT-derived adipocytes and differentiated SCA1 + APs but not in primary hepatocytes. This result is in agreement with those reported by Kralisch et al. 45 who found that IL-1β increased chemerin expression in 3T3-L1 adipocytes. Also, these in vitro results were corroborated by in vivo findings: RARRES2 mRNA was elevated in eWAT but not in the liver and chemerin was up-regulated, on average, by 1.5-to 2.0-fold in eWAT. Although this increase in chemerin levels was not robust, eWAT-mediated local production and/or activation of chemerin may play an important role in maintaining tissue homeostasis and should be considered in addition to systemic levels of circulating chemerin 1,46,47 .
In contrast with adipocytes, IL-1β and OSM did not affect chemerin expression in hepatocytes. The existence of differential regulation of chemerin expression in the liver and adipose tissue in response to other mediators has also been reported by other research groups 17,48 . So far, only FFAs and GW4064, a synthetic FXR agonist, have been shown to influence chemerin expression in hepatocytes 19 .
We also asked how the RARRES2 promoter region affects the regulated expression of chemerin. Analysis of the Chemerin_Full construct did not reveal any statistically significant degree of activation over the unstimulated control when transfected cells were stimulated with IL-1β and OSM. The Chemerin_Full vector, which covers − 735 to + 258 bp region of the murine RARRES2 gene, contains all previously characterized functional TF binding sites: including the PPAR response element 20 , FXR response element 19 , and SREBP2 binding site 18 . Acute phase cytokines can impact the expression of inflammatory-responsive genes by a variety of TFs including NFκB, AP-1, c-Jun/c-Fos, IRF1 or STAT1 [49][50][51][52] . However, our search of the ChIP-Atlas database 31 did not reveal that any of the listed TFs were able to bond to the − 735/+ 258 bp of the RARRES2 promoter region either in adipocytes or hepatocytes. This may indicate that inflammation-responsive TFs binding sites are located outside of the investigated promoter region or that other mechanisms and/or cis-regulatory elements affect chemerin expression.
Interestingly, the upregulation of chemerin expression after stimulation of adipocytes with cytokines correlated with a statistically significant increase in the average methylation level of the distal region of the RARRES2 promoter. The methylation pattern was not altered in cytokine-treated but chemerin-unresponsive primary cultures of mouse hepatocytes. Methylation of the Chemerin_Distal construct increased luciferase activity when compared with the unmethylated vector, but such was still far below the activity of the unmethylated proximal region. DNA methylation is typically associated with gene silencing although CpG methylation of the DNA and, like methylation of the CRE sequence, may enhance the DNA binding of TFs 53 .
In summary, our studies reveal novel insights into the mechanisms and factors regulating chemerin expression and secretion. For the first time, we show that DNA methylation may control the constitutive expression of RARRES2, and that the proximal region of the gene promoter is the key regulator. Acute-phase cytokines affect chemerin expression in a cell-type dependent manner, both in vitro and in vivo. The investigated RARRES2 promoter region was unresponsive to acute-phase cytokine stimulation. These findings provide a basis for further investigations of the regulation of chemerin transcription. Animal studies. Male eight-to 12-week-old C57BL/6 mice were used for these investigations. The mice were maintained under specific pathogen-free conditions at the Faculty of Biochemistry, Biophysics, and Biotechnology of Jagiellonian University animal care facility. IL-1β and OSM were injected intraperitoneally at doses of 10 μg/kg BW and 160 μg/kg BW, respectively. After 48 h, the liver and eWAT were isolated and subjected to RT-QPCR analysis. All experimental procedures were approved by the First Local Ethical Committee on Animal Testing at the Jagiellonian University in Krakow, Poland (permit no. 41/2014), in accordance with the Guidelines for Animal Care and Treatment of the European Community. The mice were sacrificed by an overdose of anesthesia (a mixture of ketamine and xylasine), followed by cervical dislocation. primary hepatocytes isolation and culture. Primary hepatocytes were isolated from C57BL6 mice with a modified two-step perfusion method according to the protocol described by Seglen 54 . Briefly, the mice were anesthetized with ketamine (100 mg/kg) and xylazine (10 mg/kg) intraperitoneally and the abdomen was opened under sterile conditions. Following cannulation of the portal vein, the liver was perfused with solution I [100 µM of EGTA in Krebs-Ringer (K-R) buffer], followed by solution II (1 mg/mL of collagenase D in K-R buffer supplemented with 150 µM of CaCl 2 ). Then, the liver was dissected, passed through a 100-µm cell strainer, and centrifuged (60 g, five minutes, 10 °C). The isolated hepatocytes were suspended in DMEM:F12 medium supplemented with 10% FBS, 50 µg/mL of gentamycin, 6 ng/mL of insulin, and 400 ng/mL of dexamethasone. Viable cells were counted using trypan blue staining and seeded on a collagen-coated plate at a den-Scientific RepoRtS | (2020) 10:13702 | https://doi.org/10.1038/s41598-020-70625-7

Materials
www.nature.com/scientificreports/ sity of 1 × 10 5 cells/cm 2 . Cells were cultured at 37 °C in the presence of 5% CO 2 for four hours. Afterward, plated cells were washed with DMEM:F12 medium and stimulated with IL-1β (10 ng/mL) and OSM (50 ng/mL) for 48 h. At the time of harvest, cell culture media supernatants were collected and RNA lysis buffer (Fenozol Plus; A&A Biotechnology, Gdynia, Poland) or radioimmunoprecipitation assay buffer containing protease inhibitors (Roche Holding AG, Basel, Switzerland) were added to the specified wells. Collected samples were subjected to RT-QPCR or enzyme-linked immunosorbent assay (ELISA) analysis.

Isolation and culture of adipose tissue-derived stromal vascular fraction (SVF). eWAT depots
were isolated from 8-to 10-week-old male C57BL6 mice, minced, and digested with collagenase D (3.5 mg/mL) in K-R buffer supplemented with 2% BSA and 150 μM of CaCl 2 for one hour in 37 °C water bath with shaking every five minutes. Digested tissue was then centrifuged (280 g, 10 min, 15 °C), washed, filtered through 100 μm of cell strainer, and centrifuged. Then, the pellet was suspended in red blood cell lysis buffer (155 mM of NH 4 Cl, 12 mM of NaHCO 3 , 0.1 mM of EDTA) for three minutes at room temperature, centrifuged, and suspended in DMEM:F12 medium (20% FBS, gentamycin 50 μg/mL). Isolated SVF cells were then seeded at a density of 9 × 10 4 cells/cm 2 on a culture plate. Attached cells were washed and replenished with fresh medium after 24 h to discard unattached dead cells or immune cells and, by the third day, more than 90% of cells displayed typical fibroblastic morphology. Then, the SVF culture was subjected to adipocyte differentiation.
isolation and culture of adipogenic progenitors. Adipogenic progenitors (APs) were isolated from C57BL6 mice according to the protocol described by Lee et al. 55 . eWAT was digested, filtered, and washed as described above. APs were then isolated first using negative selection of CD45 and CD31 and then positive selection for SCA-1. Briefly, SVF cells were suspended in MACS buffer (0.5% BSA, 2 mM of EDTA, 50 µg/mL of gentamycin) and preincubated with Fc block [rat-anti-mouse CD16/32 monoclonal antibody (mAb), 45 µg/mL], which was followed by incubation with biotin-conjugated antibodies against CD45 (rat-anti-mouse CD45 mAb) and CD31 (rat-anti-mouse CD31 mAb), both 7.5 μg/mL. Then, SVF cells were incubated with streptavidinconjugated magnetic beads, washed, and passed over a lineage depletion column to exclude endothelial cells and leukocytes. The flow-through was collected, washed, and incubated with PE-conjugated anti-SCA-1 (rat-antimouse Ly-6A/E mAb, 4.8 μg/mL), followed by anti-PE-conjugated magnetic beads. Cells were then washed and passed over a lineage selection column. Labeled cells were collected, suspended, in DMEM:F12 medium (20% FBS, gentamycin 50 μg/mL), counted, seeded on a cell culture flask, and expanded. Then, the cells were replated at a density of 2.5 × 10 4 cells/cm 2 and subjected to adipocyte differentiation in adipocyte maintenance medium.   . The integrity and orientation of the inserts were confirmed by sequencing. Plasmids were amplified in E. coli and purified using Plasmid MIDI AX kit (A&A Biotechnology, Gdynia, Poland). For in vitro methylation studies, the constructs were digested with NcoI and BglII (New England BioLabs, Ipswitch, MA, USA) and inserts and vectors bands were separated and extracted from the agarose gel. Inserts were then incubated for one hour at 37 °C in the presence or absence of SssI methylase (New England BioLabs, Ipswitch, MA, USA). The efficiency of the methylation reaction was verified by resistance to cleavage using the methylation-sensitive restriction enzyme HpaII (New England Bio-Labs, Ipswitch, MA, USA). The methylated and mock-methylated chemerin promoter fragments were re-ligated into the parent vector, purified using Clean-Up Concentrator (A&A Biotechnology, Gdynia, Poland), and used for transfection.