extfieldsofstudy: Biology Medicine

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High Yield Production of a Soluble Human Interleukin-3 Variant from E. coli with Wild-Type Bioactivity and Improved Radiolabeling Properties

Human interleukin-3 (hIL-3) is a polypeptide growth factor that regulates the proliferation, differentiation, survival and function of hematopoietic progenitors and many mature blood cell lineages. Although recombinant hIL-3 is a widely used laboratory reagent in hematology, standard methods for its preparation, including those employed by commercial suppliers, remain arduous owing to a reliance on refolding insoluble protein expressed in E. coli. In addition, wild-type hIL-3 is a poor substrate for radio-iodination, which has been a long-standing hindrance to its use in receptor binding assays. To overcome these problems, we developed a method for expression of hIL-3 in E. coli as a soluble protein, with typical yields of >3mg of purified hIL-3 per litre of shaking microbial culture. Additionally, we introduced a non-native tyrosine residue into our hIL-3 analog, which allowed radio-iodination to high specific activities for receptor binding studies whilst not compromising bioactivity. The method presented herein provides a cost-effective and convenient route to milligram quantities of a hIL-3 analog with wild-type bioactivity that, unlike wild-type hIL‑3, can be efficiently radio-iodinated for receptor binding studies.

Introduction
Human interleukin-3 (hIL-3) is a four-helix bundle, short chain cytokine that is widely expressed in vivo, principally by hematopoietic cells, such as activated T-lymphocytes, mast cells and basophils [1]. This cytokine serves an important role in the regulation of proliferation, differentiation, survival and activation of a range of hematopoietic cell types, including progenitor cells, dendritic cells, basophils and mast cells [2][3][4]. In particular, IL-3 is known to stimulate the production and function of basophils and mast cells in the context of allergic inflammation in vivo [5,6], and is commonly used as a stimulus to culture these cell types in vitro (for example, [7][8][9]. Owing to its importance in immune cell stimulation and implication in the pathogenesis of acute myeloid leukemia and chronic myeloid leukemia [10][11][12][13], IL-3 signaling has emerged as a potential therapeutic target for the treatment of allergic diseases, including asthma, and hematopoietic malignancies. The effects of IL-3 on target cells are initiated by IL-3 binding to a transmembrane receptor system composed of an IL-3specific α-subunit (IL-3Rα) and a common β-subunit (βc) shared with the related cytokines, IL-5 and GM-CSF [14][15][16]. Engagement of these cell surface receptors by IL-3 leads to transmission of a signal across the cell membrane, via a poorly understood process, activation of intracellular signaling networks and subsequent cellular responses. A bottleneck in our efforts to characterize hIL-3 engagement of its receptor in vitro has been a paucity of efficient methods for hIL-3 production from E. coli. Wild-type recombinant hIL-3 expressed in E. coli exhibits limited solubility [17] and has been typically produced by oxidative refolding of material expressed in inclusion bodies, even by commercial suppliers. Here, we describe an economical, rapid and simple method to express and purify milligram quantities of soluble hIL-3 from E. coli. In addition to affording high yields (typically >3mg/L of shaking microbial culture), the resultant recombinant hIL-3 bears a non-native tyrosine at the N-terminus to allow efficient 125 Iradiolabelling for receptor binding studies. Despite truncation of the native sequence to enhance solubility, as described in the pioneering work of Olins et al. [17], and the introduction of the non-native N-terminal tyrosine, the hIL-3 expressed and purified from the soluble fraction of E. coli lysates using our method and commercially-sourced recombinant hIL-3 preparations were equipotent growth stimuli in TF-1 cell proliferation studies. Therefore, the method described herein provides a robust and convenient strategy to produce milligram quantities of fully bioactive soluble hIL-3 for a broad range of laboratory applications, including hematopoietic cell culture and molecular characterization of hIL-3 receptor binding.

Expression construct
A cDNA encoding hIL-3 residues 13-125, including the W13Y mutation, was amplified from an IMAGE clone template (IMAGE: 6971773; NCBI accession: BC066272) with Pfu polymerase using the primers: 5' CGC GGa TCc tAT gtt aac tgc tct aac atg atc gat g; 3' ATAAGAATGCGGCCG cTA ctg ttg agc ctg cgc att ctc. The PCR product was digested with BamHI and NotI restriction endonucleases before ligation into the corresponding restriction sites in the vector, pETNusH HTb [18], a derivative of pETM30 bearing a kanamycin-resistance marker. The insert was verified by Big Dye Terminator sequencing.

Protein expression and purification from E. coli
E. coli BL21 CodonPlus (DE3)-RIPL cells transformed with the expression construct described above were cultured at 37°C in 2 L flasks containing 0.6 L of TYH medium (20 g tryptone, 10 g yeast extract, 5 g NaCl and 1 g MgSO 4 per litre with 46 mM HEPES pH 7.4) and supplemented with 50 µg/ml kanamycin. Once the optical density at 600 nm reached 0.6-0.8, cultures were adjusted to 1.5 mM isopropyl β-D-1thiogalactopyranoside (IPTG) and incubated at ~23°C with shaking for ~16 hours. We found that induction for ~16 hours at 18-25°C gave comparable yields, although higher expression temperatures were not tested. Cells were harvested by centrifugation and lysed by sonication in NL buffer (0.2 M NaCl, 50 mM Na phosphate pH 8.0, 10% v/v glycerol, 0.05% v/v Na azide) freshly supplemented with 20 mM imidazole, 0.05% v/v Tween 20 and 1 mM phenylmethylsulfonyl fluoride (PMSF). The lysate was clarified by centrifugation at 26,800xg for 30 minutes followed by 0.45 µm filtration of the supernatant. Crude lysate was bound at 4°C and 2 ml/min to a 10 ml column of Ni-NTA Agarose (Qiagen) equilibrated in NL buffer that had been packed in an XK-26 column holder (GE Healthcare). The Ni-NTA agarose was washed in 200 ml NL buffer containing 20 mM imidazole at 5 ml/min and the bound NusA-His 6 -hIL-3 (13-125; W13Y) fusion protein eluted in 100 ml NL buffer containing 250 mM imidazole at 5 ml/min and collected in 10 ml fractions. Fractions containing purified NusA-His 6 -hIL-3 (13-125; W13Y) fusion protein were pooled and concentrated to a final volume of < 20 ml using Vivaspin 20, 30,000 molecular weight cut-off devices (Sartorius). The concentrated fusion protein was dialysed against 50mM Tris-HCl pH 7.4, adjusted to 1 mM dithiothreitol (DTT), 0.5 mM EDTA and mixed with purified TEV protease, produced in-house, at 23°C for ~16 hours. The amount of TEV protease used was empirically determined for each batch of the NusA-His 6 -hIL-3 (13-125; W13Y) fusion protein to achieve complete cleavage. Typically, complete cleavage required up to 0.1 mg TEV protease per 10 mg NusA-His 6 -hIL-3 (13-125; W13Y).
The TEV protease-digested NusA-His 6 -hIL-3 (13-125; W13Y) fusion protein was fractionated by size exclusion chromatography (SEC) using a Superdex 200 column (26 mm x 600 mm, GE Healthcare) fitted with a 10 ml Super Loop and operated at 4°C with a flow rate of 2 ml/min. SEC was performed using a running buffer composed of 150 mM NaCl, 50 mM Na phosphate pH 7.0. SEC fractions containing hIL-3 were established by SDS-PAGE analysis with Coomassie Blue staining and then pooled, with a portion sterilized using Spin-X filters (Corning).
In some cases, SEC purified hIL-3 (13-125; W13Y) was further purified by reversed-phase chromatography. Glacial acetic acid was added to the SEC purified hIL-3 (13-125; W13Y) to a final concentration of 1% (v/v) while trifluoroacetic acid was added to a final concentration of 0.1% (v/v). The sample was applied to an Aquapore RP300 reversed-phase column (4.6 mm x 100 mm) in buffer A (0.1% v/v trifluoroacetic acid in water) via a 2 ml injection loop fitted to a Breeze 2 semipreparative HPLC system (Waters). Bound proteins were eluted over 60 minutes using a linear gradient from 100% buffer A to 100% buffer B (0.085% v/v trifluoroacetic acid in acetonitrile) at 1 ml/min with the collection of 1 ml fractions into tubes containing 20 µl 2M Tris. Fractions containing purified hIL-3 (13-125; W13Y) were pooled, lyophilized, resuspended in PBS and sterilized using Spin-X filters.
Purified hIL-3 (13-125; W13Y) was quantitated by analytical SEC using a Superdex 200PC column (3.2 mm x 300 mm, GE Healthcare) and a 50 µl loop operated at 25°C and 40 µl/min with a running buffer composed of 150 mM NaCl, 50 mM Na phosphate pH 7.0. The area under the hIL-3 peak was integrated by using the calculated extinction coefficient of 0.644 M -1 cm -1 . The concentrations of commercially-sourced hIL-3 used to compare the bioactivity of hIL-3 (13-125; W13Y) was specified by the manufacturers.

Mass spectrometry analysis
hIL-3 (13-125; W13Y) was digested with trypsin using the FASP protocol [19] and eluted peptides were injected and separated by nano flow reversed-phase liquid chromatography on a nano LC system (1200 series, Agilent) using a nanoAcquity C18 150 mm × 0.15 mm I.D. column (Waters) with a linear 45 minute gradient from 5 to 100% buffer B set at a flow rate of 1.2 µl/min (Buffer A: 0.1% Formic acid in Milli-Q water; Buffer B: 0.1% Formic acid, 80% acetonitrile (Mallinckrodt, Baker) 20% Milli-Q water). The nano HPLC was coupled on-line to a Q-Exactive mass spectrometer equipped with a nano-electrospray ion source (Thermo, Fisher Scientific) for automated MS/MS. The Q-Exactive was run in a datadependent acquisition mode with the full scan resolution set at 30,000 and the top-ten multiply charged species selected for fragmentation using the high energy collision disassociation (HCD) (single charged species were ignored). Fragment ions were analysed with the resolution set at 17,500 and the ion threshold set to 1e5 intensity. The activation time was set to 30 ms and the normalized collision energy set to 24. Raw files consisting of full-scan MS and high resolution MS/MS spectra were converted to the MGF data format with Proteome Discoverer 1.4 and searched against UniProt database (2013/02) including the hIL-3 analog sequence, limiting the search to human taxonomy using Mascot v 2.4. Mascot parameters for each search included Trypsin/no P enzyme with three missed cleavages, a fixed modification in the form of carbamidomethyl at Cys residues and variable modifications of Acetyl at protein N-Terminal and oxidation at Met residues. Spectra were searched with a mass tolerance of 20 ppm in MS mode and 40 mmu in MS/MS mode.

Expression construct design
In an attempt to produce hIL-3 in the soluble fraction of E. coli lysates rather than in inclusion bodies, we prepared a fusion protein with NusA-His 6 [18], (Figure 1). To this end, we PCR amplified a fragment encoding the four-helix bundle core of hIL-3, residues 13-125, and ligated into the vector, pETNusH, as an in-frame fusion C-terminal to the 55kDa solubility tag, NusA, and a His 6 affinity tag to allow fusion protein purification by Ni 2+ chromatography. Between the His 6 tag and the hIL-3 sequence is a vector-encoded TEV protease recognition site. TEV protease cleavage of the NusA-His 6 tag from hIL-3 introduces a non-native GAMGS at the N-terminus owing to a cloning artifact. We chose to incorporate a TEV protease cleavage site, since this enzyme can be readily prepared in the laboratory to high yields [25] and cuts with high efficiency and sequence specificity unlike other enzymes, such as thrombin and Factor Xa. Importantly, TEV protease offers the flexibility of cleaving substrates at 25°C, typically within 2 hours, or at 4°C overnight should the target protein be temperature sensitive.
Full length hIL-3 is expressed from mammalian cells as a pro-protein that is cleaved during secretion (signal peptide shown in grey in Figure 1) to yield a mature polypeptide, the first residue of which we have numbered as residue 1 ( Figure   Figure 1. Alignment of amino acid sequences of human and mouse IL-3. The amino acid sequences of full length wild-type hIL-3, hIL-3(13-125; W13Y), the hIL-3 analog SC-65369 [41], and wild-type full length mouse IL-3 were manually aligned owing to low homology between mouse and human IL-3 (29% identity). Numbers above the sequence refer to the mature form of full length hIL-3 with dots above every 10 th residue. The sequences shown in gray for full length hIL-3 and full length mouse IL-3 are signal peptides that are cleaved during secretion. The key substitution, W13Y in hIL-3(13-125; W13Y), is shown in bold text and highlighted.
doi: 10.1371/journal.pone.0074376.g001 1). Prior studies have established that truncation of hIL-3 at either terminus enhances its solubility without compromising bioactivity [17] and the enhanced solubility of a truncated and highly-mutated hIL-3 analog, SC-65369 (Figure 1), allowed its solution structure to be determined [26]. Although the sequence of our hIL-3 protein is wild-type, but truncated at both N-and C-termini, we introduced a tyrosine in place of Trp13 (highlighted in Figure 1) in our expression construct to facilitate radio-iodination for receptor binding assays.

Purification of hIL-3(13-125; W13Y)
We developed a method to prepare hIL-3 to high-purity and yield (Figure 2A). The expression of NusA-His 6 fused hIL-3(13-125; W13Y) was carried out in E. coli BL21 Codon Plus (DE3) RIPL, followed by cell lysis and Ni 2+chromatography. Following cleavage of the fusion protein with TEV protease, the mixture was concentrated and the hIL-3(13-125; W13Y) separated from uncut fusion protein, the NusA-His 6 tag and TEV protease using preparative SEC. The molecular mass difference between NusA and hIL-3 was sufficiently large (55kD vs. 13.4kD) to permit resolution by Superdex-200 26/60 SEC, with NusA elution at 186 mL and hIL-3 at 254 mL ( Figure 2B). While the hIL-3 fractions were of high purity as gauged by SDS-PAGE with Coomassie Blue staining with no detectable NusA or TEV protease present ( Figure 2C), we proceeded to purify hIL-3 to homogeneity using reversed-phase high-performance liquid chromatography (RP-HPLC; Figure 2D). RP-HPLC was performed with a gradient of 0 to 100% acetonitrile with hIL-3 elution occurring at ~50% acetonitrile. As evidenced by the RP-HPLC chromatogram shown in Figure 2D, negligible quantities of proteinaceous contaminants were present in hIL-3 following SEC, underscoring that RP-HPLC can be considered a polishing step but does not vastly enhance the purity of hIL-3. Routinely, the yields per litre of shaking bacterial culture exceed 3mg, although in some instances we have obtained up to 7mg of purified hIL-3(13-125; W13Y) per litre of culture.
We verified the correct sequence of purified hIL-3(13-125; W13Y) protein and confirmed the introduced modifications by performing mass spectrometry. Purified protein was subjected to tryptic digestion and tandem mass spectrometry analysis. As summarized in Figure 2E, >90% sequence coverage was observed, verifying the composition of the purified product. The identified peptides are shown in Table S1.

Characterization of bioactivity relative to commerciallysourced hIL-3
Having established a high-yielding strategy to express and purify soluble hIL-3(13-125; W13Y) from E. coli, we proceeded to compare its potency as a growth stimulus in TF-1 cell proliferation assays relative to recombinant hIL-3 sourced from two different commercial vendors. TF-1 is a factor-dependent cell line that endogenously expresses the hIL-3 receptor, and is a cell line commonly used to assay hIL-3 bioactivity. As shown in Figure 3A were equipotent stimuli in TF-1 proliferation assays ( Figure  3A). These preparations of hIL-3(13-125; W13Y) exhibited comparable, if not slightly enhanced, potency compared to hIL-3 sourced from vendor A (black circles). Surprisingly, however, a second commercially-sourced sample (black squares; vendor B) was an order of magnitude less potent than each of our hIL-3(13-125; W13Y) preparations and the hIL-3 supplied by vendor A. Consequently, we concluded that not only are the bioactivities of our SEC product and our RP-HPLC product virtually indistinguishable, but their bioactivities in TF1 proliferation assays were equivalent or better than commercially-supplied hIL-3 from two different vendors.

Discussion
Since the human IL-3 gene was cloned in 1986 [28], highyielding, cost-effective approaches have been sought for the expression and purification of hIL-3 for biochemical, structural and biological studies, including as a growth stimulus for cell culture applications. While several different preparation methods from a variety of expression hosts have been described [29][30][31][32], E. coli remains the most rapid, accessible and inexpensive source of recombinant cytokines. Commonly, however, the preparation of recombinant cytokines from E. coli necessitates time-consuming and cumbersome oxidative refolding steps (e.g. [33]). In the present work, we demonstrate the efficacy of producing hIL-3 as a soluble fusion protein bearing a NusA-His 6 solubility/purification tag, which allows for convenient purification prior to cleavage with TEV protease. Consistent with an emerging role for the NusA tag as an alternative to GST fusions to enhance the solubility of poorlysoluble proteins [34,35], we have previously found that the NusA tag was effective in allowing the preparation of mouse IL-3 as a soluble protein from E. coli [18] (Figure 1) with yields and solubility in aqueous solvents sufficient for NMR spectroscopy studies [36][37][38]. In addition to maintaining hIL-3 in soluble form, the present method allows the preparation of milligram quantities of hIL-3 per litre of shaking E. coli culture, is rapid, simple and completely avoids refolding. To minimize losses during chromatographic steps, rather than dialyzing the TEV protease-cleaved hIL-3 and performing further Ni 2+chromatography, the cleavage reaction was concentrated and applied directly to a preparative SEC column. The large molecular weight difference between the NusA-His 6 tag (55 kDa) and hIL-3 (13.4 kDa) enabled resolution of the two species by SEC ( Figure 2B). Whilst the SEC product was highly purified ( Figure 2C) with no detectable NusA or TEV protease present, an additional polishing step, reversed-phase B) Elution profile of the TEV protease digested, NusA and hIL-3 analog mixture, following size exclusion chromatography (SEC) using a Superdex 200 column (26 mm x 600 mm) operated at 2 ml/min at 4°C with 50 mM sodium phosphate pH 7.0, 150 mM NaCl as running buffer. Free NusA eluted at ~186 mL and the hIL-3 analog eluted at 254 mL. The first peak at 116 mL contains aggregates while we suspect the last peak at 328 mL contains DTT from the digest. Molecular weight (kDa) marker elution positions are marked as dots above the elution profile. C) Analysis of hIL-3 analog purification by 15% acrylamide reducing SDS-PAGE with Coomassie Blue staining. NusA: hIL-3(13-125; W13Y) fusion protein was isolated by Ni-chromatography (Lane 1) prior to cleavage by TEV protease (Lane 2) to yield free NusA (55kDa) and the hIL-3 analog (13.4kDa). Fractions containing the hIL-3 analog that eluted around 254ml during SEC are shown (Lanes 4-9), illustrating that the hIL-3 analog was purified free from NusA (Lane 3). D) The SEC purified hIL-3 analog was applied to an Aquapore RP300 reversed-phase column (4.6 mm x100 mm) and bound proteins eluted using a 0-100% gradient of acetonitrile in 0.1% trifluoroacetic acid. The hIL-3 analog eluted at 34 min (~50% acetonitrile). E) Purified hIL-3(13-125; W13Y) was subjected to tryptic digestion and tandem mass spectrometry. The sequences of peptides not identified in this analysis are shown as lowercase italics. Asterisked methionine residues were oxidized. Sequence arising from the NusA-His 6 fusion overhang after TEV protease cleavage is underlined, while the residues are numbered according to the mature, full-length IL-3 reference sequence.  HPLC, can be employed to purify the hIL-3(13-125; W13Y) to homogeneity ( Figure 2D). Both the SEC and the RP-HPLC products exhibited comparable bioactivities in stimulating proliferation of the hIL-3-dependent cell lines, TF-1 ( Figure 3A). We consider the RP-HPLC step non-essential for structural and biochemical studies of hIL-3, but note it is likely to be advantageous for biological studies since RP-HPLC is known to largely eliminate lipopolysaccharide (LPS) [39], an agonist of the pro-inflammatory TLR4 receptor, which is a common contaminant of proteins prepared from Gram-negative bacterial hosts.
In designing our hIL-3 expression construct, we introduced a tyrosine in place of Trp13 (highlighted in Figure 1) to serve as a substrate for 125 I-labelling. Wild-type hIL-3 is a poor substrate for commonly employed tyrosine-radioiodination methods, such as Iodogen or iodine monochloride labeling, owing to the sole tyrosine being buried in the helical bundle core ( [26]; PDB, 1JLI), leading to low specific activities when native hIL-3 is tyrosine-radioiodinated [23]. We reasoned that introduction of a tyrosine residue peripheral to the four-helix bundle would overcome a longstanding technical hurdle and allow efficient radiolabelling of the hIL-3 analog. As anticipated, introduction of a non-native Tyr residue N-terminal to the hIL-3 four helix bundle does not impact the bioactivity ( Figure 3A). We found that hIL-3(13-125; W13Y) is an excellent substrate for 125 Ilabeling using the Iodogen method and we successfully used the radiolabelled product in receptor binding studies, affording K d values comparable to those reported in prior hIL-3 receptor binding studies [40].
In summary, herein we describe a simple and inexpensive procedure for preparation of a hIL-3 analog as a soluble protein, which exhibits wild-type bioactivity and improved radiolabeling capacity, from E. coli. Not only does this procedure typically yield >3mg per litre of shaking culture, but the whole procedure can be completed within 3 days without the necessity of oxidative refolding. Table S1.

Supporting Information
Tryptic peptide mass spectra identifying hIL-3(13-125; W13Y). High quality spectra identified 89% of the full length construct sequence using Mascot 2.4 upon searching the UniProt database (with the vector-derived overhang, GAMGS, included). Complete details of the experiment are shown in Materials and Methods. (XLS)


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In Vivo Cell Reprogramming towards Pluripotency by Virus-Free Overexpression of Defined Factors

The ability to induce the reprogramming of somatic mammalian cells to a pluripotent state by the forced expression of specific transcription factors has helped redefine the rules of cell fate and plasticity, as well as open possibilities for disease modeling, drug screening and regenerative medicine. Here, we hypothesized that the non-viral forced expression of the four originally discovered defined factors (OKSM) in adult mice could result in in vivo reprogramming of cells in the transfected tissue in situ. We show that a single hydrodynamic tail-vein (HTV) injection of two plasmids encoding for Oct3/4, Sox2, Klf4 and c-Myc respectively, are highly expressed in the liver tissue of Balb/C adult mice. Hallmark pluripotency markers were upregulated within 24–48 h after injection, followed by down-regulation of all major hepatocellular markers. Generation of transcriptionally reprogrammed cells in vivo was further confirmed by positive staining of liver tissue sections for all major pluripotency markers in Balb/C mice and the Nanog-GFP reporter transgenic strain (TNG-A) with concomitant upregulation of GFP expression in situ. No signs of physiological or anatomical abnormalities or teratoma formation were observed in the liver examined up to 120 days. These findings indicate that virus-free expression of OKSM factors in vivo can transcriptionally reprogram cells in situ rapidly, efficiently and transiently, absent of host tissue damage or teratoma formation.

Introduction
Forced reprogramming of somatic cells into a pluripotent, stem cell-like state by the ectopic expression of specific transcription factors results in the generation of induced pluripotent stem (iPS) cells. Such transcription factor cell reprogramming has been achieved today by viral [1,2,3] and non-viral [4,5,6,7] gene transfer, protein cytoplasmic translocation [8,9], miRNA [10] and is changing the landscape in developmental biology, can potentially resolve all ethical concerns about the use of embryonic stem cells and open further opportunities for regenerative medicine. The original discovery by Yamanaka and colleagues that the in vitro expression of four transcription factors, Oct3/4, Klf4, Sox2, c-Myc (OKSM) was capable to revert fully differentiated mouse and human skin fibroblasts into iPS cells [1,11] constitutes the most widely used transcription-based reprogramming technology today.
The initial reports of transcription-mediated somatic cell reprogramming involved the use of retroviruses to stably transduce skin fibroblasts with defined transcription factors [1,2,11]. This methodology of gene transfer is still today the most popular way to reprogram animal and human somatic cells despite the risks from insertional mutagenesis, stable transduction and long-term gene expression of known proto-oncogenes [12,13]. Moreover, the vast majority of current methodologies to generate iPS cells involve use of long-term culture conditions and treatment of cells with multiple rounds of gene transfer vectors, growth factors, antibiotics and other cell media cocktails to promote reprogramming and select for pluripotency. All of these are considered major culprits for the potential risks associated with the ensuing cells as recent studies investigating the genomic integrity of iPS have alluded to [14,15,16]. In terms of iPS generation using non-viral gene transfer vectors, plasmid DNA [4,5,6] or RNA [7,10] delivery using liposomes or electroporation have been reported. Compared to viruses, episomal vectors are generally considered safer, however transduction and reprogramming efficiencies are much lower [13]. Alternatively, Warren et al. reported somatic cell reprogramming in vitro by direct delivery of synthetic mRNAs [7]. Although this methodology offers significantly higher reprogramming efficiency, high RNA dosages, multiple rounds of transfection and complex cell culturing protocols are still needed [13].
Due to the paradigm-shifting nature of transcription-induced reprogramming to pluripotency there is still limited understanding of the exact mechanisms and pathways implicated in induced cell reprogramming, and the exact features of reprogrammed cells [17,18]. Morever, the majority of experimental evidence today is based on the concept of extraction and in vitro manipulation of the somatic cells to be reprogrammed, leading to the array of caveats mentioned above that make clinical translation of iPS cells seem distant [19,20,21]. In the present work, we hypothesized that in vivo forced expression of the OKSM transcription factors by non-viral transient over-expression within living tissue can induce cell reprogramming towards pluripotency. In order to test this hypothesis we chose the most naive, non-viral gene transfer technology available today: large-volume, rapid hydrodynamic tail vein (HTV) injection of plasmid DNA [22,23] encoding the originally proposed OKSM factors. This gene transfer methodology circumvents most complications or risks associated with viral gene transfer vectors, as has been previously shown in numerous preclinical [24,25] and clinical [26,27] studies allowing unprecedented levels of exogenous gene expression in hepatocytes.

Plasmids
Reprogramming plasmids pCX-OKS-2A encoding OCT3/4, KLF4, SOX2; pCX-cMyc encoding CMYC and pCAG-GFP encoding eGFP under the control of CAG promoter (as previously described by Okita et al. [4]) were obtained from Addgene (USA) as bacterial stabs. Research grade plasmid production was performed from these stabs (Plasmid Factory, Germany). pGFP-Luc plasmid (Clontech, USA) encodes for the eGFP and Luc transgenes under the control of a CMV promoter. Vector maps of all plasmids used in this work are included in Figure S5.

Animals and Hydrodynamic Tail Vein (HTV) Injection of Plasmids
All experiments were performed with prior approval from the UK Home Office under a Home Office project license (PPL 80/ 2296). Female Balb/C mice, 6 weeks old were purchased from Harlan, UK. TNG-A mice which carry the eGFP reporter inserted into the Nanog locus [28], were a kind gift from The Wellcome Trust Centre for Stem Cell Research, University of Cambridge, UK. TNG-A mice and WT controls, were of 129 background and were bred and genotyped at the UCL. Mice were allowed one week to acclimatize prior to use. Mice (4 animals/ group)were warmed in a 37uC heating chamber, anesthetized with isofluorane and were injected via tail vein in 5-7 sec with 1.5 ml of 0.9% saline including 75 mg of pCX-OKS-2A with and without 75 mg of pCX-cMyc or 150 mg pCAG-GFP plasmids or no plasmid. Mice were culled at different time points, such as 2, 4, 8, 12, 24, 50, 120 days after HTV injections.

Isolation of Hepatocyte Population
Mice livers were perfused as previously described [36] with some modifications. In brief, livers were first perfused with Ca 2+ and Mg 2+ free Hank's buffered salt solution (Sigma-Aldrich, UK) and then with liver digest medium (Gibco, UK) at 37uC. After digestion, liver was washed with Hepatocyte Wash Medium (HWM, Gibco, UK) and cell suspension was passed through a 100 ml cell strainer (VWR, UK) at 4uC. Cells were centrifuged at 50 g for 5 min to separate parenchymal cells (PC including hepatocytes) which were collected in pellet and non-parenchymal cells (NPC including Kupffer cells and epithelial cells) which stayed in supernatant. The hepatocyte fraction was collected after washing twice with HWM.

RNA Isolation and Reverse Transcription-quantitative PCR (RT-qPCR) Analysis
Following liver perfusion and isolation of hepatocytes, RNeasy Blood and Tissue Kit (Qiagen, UK) were used to isolate total RNA. cDNA synthesis from 1 mg of RNA sample was performed by iScript cDNA synthesis kit (Bio-Rad, UK) according to manufacturer's instructions. Two microliters of each cDNA sample was used to perform RT-qPCR reactions with iO SYBR Green Supermix (Bio-Rad, UK). Primer sequences are shown in Table S1. Samples were run on CFX-96 Real Time System (Bio-Rad, UK) with the following protocol: 95uC for 3 min, 1 cycle; 95uC for 10 sec, 60uC for 30 sec, -repeated for 40 cycles. b-actin was used as a housekeeping gene and gene expression levels were normalized to saline groups.

Flow Cytometry
Cell density was adjusted to 1610 7 cells/ml and 100 ml suspensions were aliquoted in microfuge tubes. BD Mouse Pluripotent Stem Cell Transcription Factor Analysis Kit (BD Biosciences, UK) were used to analyze OCT3/4-positive or NANOG-positive cells. In brief, cells were firstly fixed with BD Cytofix fixation buffer and permeabilized with 1X BD Perm/ Wash buffer. Then cells were incubated with either anti-mouse OCT4-PerCP-Cy5.5 or anti-mouse NANOG-PE for 30 min according to manufacturer's instructions. Negative and isotype controls were included in each experiment. Stained cells were then analyzed by CyAn TM ADP High-Performance Research Flow

Quantification of Serum Cytokine Levels
Blood was collected from mice at different time points. Serum levels of ALT, AST, ALP, GLDH were determined by Diagnostic Laboratories in the Royal Veterinary College, London, UK.

Immuno-histochemistry (IHC) of Mouse Livers
Livers were perfused with 10 mL HBSS, pre-warmed at 37uC, then immediately immersed into isopentane, pre-cooled in liquid nitrogen or into stabilized 4% paraformaldehyde for fixation. Frozen livers were stored at 280uC until further processing. Fourteen micron thick sections were prepared on a Cryomicro-tome (Leica Microsystems, CM3050S), air-dried for 1hour at room temperature, before storage at 220uC. Before staining, liver sections were post-fixed with methanol, pre-cooled at 220uC, for 10 min at 220uC, then air-dried for 15 min and finally washed twice with PBS for 5 min. For Oct4, Sox2 and Nanog, we used a conventional immunostaining protocol that consisted of 1 h incubation in blocking buffer (5% goat serum-0.1% Triton in PBS pH 7.3) at room temperature, followed by two washing steps with PBS (1%BSA-0.1% Triton, pH 7.3) before overnight incubation at +4uC with the different primary antibodies [(rabbit pAb anti-OCT4 (ab19857,3 mg/ml, Abcam, UK)/rabbit pAb anti-SOX2 (ab97959, 1 mg/ml, Abcam, UK)/rabbit polyclonal anti-NANOG (ab80892, 1 mg/ml, Abcam, UK]). Next day sections were washed (2 min each) with PBS and incubated (1.5 hours at room temperature) with the secondary antibody (goat polyclonal anti- rabbit IgG labeled with Cy3, 1/250, Jackson ImmunoResearch Laboratories Inc.). For SSEA1 immunostain, a mouse monoclonal antibody (ab16285, 20 mg/ml, Abcam,UK) was used, which make it more suitable to use a specific immunodetection kit to localize mouse antibodies on mouse tissues (vector MOM immunodection kit, Vector Laboratories). The endogenous liver avidin/biotin were blocked with Avidin/blocking Kit (Vector Laboratories, UK). The staining procedure was performed according to the supplier recommendations, using either the provided secondary biotinylated anti-mouse IgG revealed by fluorescein-tagged Avidin (1/250, Vector Laboratories) or a goat polyclonal anti-mouse IgG labeled with Cy3 (1/250, Jackson ImmunoResearch Laboratories Inc.). Then liver sections were washed with PBS and mounted in DAPI and antifade containing medium (Vectashield mounting medium, Vector Laboratories, UK). Slides were visualised under epi-fluorescence microscope (Zeiss Axio Observer). Paraformaldehyde fixed samples were paraffin-embedded and liver sections were stained with Haematoxylin and Eosin (H&E) by Diagnostic Laboratories in the Royal Veterinary College, London, UK. Random images were captured by light microscopy for different treatment groups.

Periodic Acid Schiff (PAS) Staining of Liver Sections
Liver samples were fixed in paraformaldehyde, paraffinembedded and liver sections were stained with PAS stain (Sigma, UK). Random images were captured by light microscopy for saline or OKSM treatment groups.

Alkaline Phosphatase (ALP) Staining of Cell Cultures and Liver Sections
ALP activity staining was performed using the BCIP/NBT liquid substrate system (Sigma UK). Methanol-fixed cell cultures were first washed with HEPES buffer and then incubated at 37uC for 30 min with the BCIP/NBT liquid substrate system. Color development was stopped by rinsing with water. In order to determine ALP activity in the tissue, frozen liver sections were prepared as described above and incubated with BCIP/NBT liquid substrate system for 30 min. Sections were washed with water and mounted. Random images were captured by light microscopy (106) for saline, GFP or OKSM treatment groups.

Statistical Analysis
Experiments were performed with at least four animals per group. Statistical analysis was performed by analysis of variance and Tukey's pairwise comparison using SPSS software, version 16.0.

Results
Mice were injected by HTV with an equimolar mix of two plasmids, pCX-OKS-2A and pCX-cMyc, encoding for the OKS and M reprogramming factors respectively. HTV injection of plasmid DNA has been known to result in high levels of gene expression in hepatocytes [22,23] and primary hepatocytes from OKSM-injected animals were extracted at different time points here. RT-qPCR and flow cytometry were used to analyze the expression levels for various reprogramming (Oct3/4, Sox2, c-Myc), pluripotency (Nanog, Ecat1, Rex1, Cripto, Gdf3 and endogenous Oct3/4, Klf,4 or Sox2) and hepatocyte markers (Alb, Trf, Aat) directly after extraction of the hepatocytes at different time points after HTV injection.
A significant increase in the gene expression of all transduced transcription factors was observed on day 2 that decreased over time (Figure 1a). At the same time endogenous pluripotency markers were upregulated, reached peak values on day 4 and decreased to background levels from day 8 onward (Figure 1b). Protein expression in the hepatocyte extracts from the transduced liver tissue using flow cytometry indicated that on day 1 only OCT3/4 was expressed, whereas by day 4 both OCT3/4 and NANOG positive cells were detected (Figure 1c). Examination of the expression levels of hepatocyte markers by RT-qPCR indicated that on day 2 they were similar to those of animals HTV injected with saline, while on day 4 significant downregulation in hepatocyte marker expression was considered as another indication of cell reprogramming, manifested by the dedifferentiation of hepatocytes (Figure 1d). Later than day 8, hepatocyte markers returned back to control levels, while on day 24 increased levels of these markers were obtained.
Transgene expression by HTV injection was monitored separately by injection of control plasmids encoding for GFPluciferase (pCMV-GFP-luc) and eGFP (pCAG-eGFP). Figure S1 confirms that hepatocytes were the targeted cell population in the liver following HTV injection as previously reported. The kinetics of transgene expression in the liver following HTV injection with the OKSM plasmids indicated that from day 9 onward, pluripotency factors began to be down-regulated reaching naive levels by day 22 (Figure S2). The effect of plasmid dose was then studied by HTV injections of OKSM against control (pCAG-eGFP) plasmid DNA at an escalated dose regime (Figure 2a). A sharp increase in the levels of gene expression profile of the transduced reprogramming and other endogenous pluripotent factors on day 4 after administration was observed only in the case of OKSM plasmid DNA injections, and the levels of up-regulation reached a plateau at 75 mg/animal. c-Myc is a known oncogene and its use in cell reprogramming (in particular for in vivo strategies) is preferable to be avoided. However, a decrease in the efficiency of reprogramming has been previously described in the absence of this factor [30]. The effect of the presence or absence of c-Myc in the reprogramming cocktail was also investigated in the present study (Figure 2b). In vivo cell reprogramming could be achieved without c-Myc, however the levels of pluripotency markers Nanog and Ecat1 observed were significantly lower compared to use of the OKSM cocktail, in line with the previous reports omitting c-Myc from in vitro cell reprogramming.
To further interrogate the occurrence of in vivo induced cell reprogramming in the liver by the forced expression of the OKSM transcription factors, tissue sections from transfected Balb/C mice were directly stained immunohistochemically (IHC) at day 4 (post- HTV) for different pluripotency markers (Oct3/4, Sox2, Nanog and ALP). Figure 3 shows that distinctive IHC-positive cells were obtained for all four markers only in the case of animals injected with the OKSM plasmids compared to saline or GFP plasmid HTV-injected groups. Positive staining for Nanog was obtained reproducibly in all liver sections in all OKSM-transfected animals indicating the presence of transcriptionally reprogrammed cells throughout the liver tissue ( Figure S3). Considering the critical role of Nanog in the control of pluripotency, we performed a separate experiment using the transgenic strain TNG-A that carries the eGFP reporter inserted into the Nanog locus [28]. HTV injection of the OKSM plasmids in TNG-A mice showed enhanced gene expression in both reprogramming and endogenous pluripotency markers on days 2 and 4 post-injection (Figure 4a & b). Hepatocyte extracts from OKSM injected TNG-A mice on day 4 were analyzed by flow cytometry for eGFP protein expression, revealing that 6-15% of the total cell population were eGFP positive (Figure 4c). Lastly, further confirmation of the generation of reprogrammed cells in the liver was offered by the presence of eGFP-positive cells in frozen tissue sections imaged by fluorescence microscopy (Figure 4d).
One of the key concerns by induction of reprogramming towards pluripotency in vivo may be the spontaneous occurrence of teratomas within the tissues where reprogrammed cells are generated [12,19]. To address this, animals HTV-injected with the reprogramming factors were kept for a period of 120 days and at frequent intervals (days 2, 4, 8, 12, 50 and 120) different groups were analyzed heamatologically and histologically (Figure 5a). HTV injection of plasmid DNA can result in moderate tissue damage manifested by increased serum levels of liver enzymes at early time points [26], which was also observed here at day 2 for both saline and OKSM injected groups (Figure 5a & S4). H&E staining of liver tissues indicated that HTV injection of OKSM plasmids did not lead to any tissue damage beyond day 2, nor development of teratomas at later time points, with all liver sections and animals exhibiting healthy structural morphology and behavior with no signs of dysplasia (Figure 5a). Serum levels were also analyzed for liver enzymes over the same period, with no aberrant change in the levels of ALT, AST and GLDH between the saline-injected and OKSM-injected groups (Figure 5b). Albumin levels and glycogen staining of liver sections (Figure 5c & d) further confirmed no hepatic structural or functional abnormality throughout the course of the study for any of the animals. These findings in conjunction with the gene and protein expression analyses of the liver tissue ( Figure 1) suggested that transcriptional cell reprogramming was occurring in vivo rapidly after HTV-injection of the OKSM factors, similar to recent direct reprogramming (transdifferentiation) studies [29,30,31]. We speculate that the endogenous tissue milieu (growth factors, homeostatic signaling cues) leads to rapid loss of the transcriptionally reprogrammed cells into phenotypically normal, functioning hepatocytes. However, much more work using lineage-tracing techniques is needed to elucidate the mechanisms and extent of in vivo cell reprogramming to a pluripotent state.

Discussion
This study provides previously unreported evidence that direct in vivo transcriptional cell reprogramming towards the pluripotent state in adult mammalian somatic (hepatocyte) cells is possible using non-viral transfection of plasmids encoding the OKSM (or OKS) transcription factors. This occurs following rapid kinetics (within 24-48 h) that are, in principle, reminiscent of the kinetics described for somatic cell reprogramming using the egg and oocyte nuclear transfer methodologies [32]. Zhou et al. have previously reported rapid and highly efficient in vivo conversion of pancreatic exocrine cells directly into insulin secreting b-cells without reverting to a pluripotent state (i.e. by transdifferentiation) using adenovirus-mediated transcription factor (not OKSM) overexpression [33]. Very recently, three studies have also offered proof-of-concept evidence that transcriptionally-induced, direct transdifferentiation from fibroblasts to cardiomyocyte-like cells in infracted heart tissue in situ can offer therapeutic benefits [29,30,31]. In all such previous work direct conversion to different lineages is reported without reprogramming towards a plutipotent state, shown to take place rapidly and efficiently (compared to in vitro transdifferentiation methodologies). From our studies herein, in vivo cell reprogramming by overexpression of OKSM transcription factors is also shown to be rapid and efficient enough to be detected in situ, however transiently maintained. According to our semi-quantitative analysis based on FACS and IHC of liver sections from transfected Balb/C and TNG-A mice, it took place efficiently in the order of 5-15% of the total hepatocyte population. Even though upregulation of eGFP and pluripotency markers in the TNG-A mice strongly suggest that HTV injection of OKSM plasmids can result in adult somatic cell reprogramming toward pluripotency in vivo, further studies are needed to determine the level of pluripotency achieved, similar to previous Nanog-activity studies using ES cells [34], as well as more detailed characterization of the extracted reprogrammed cells in relation to the loss of hepatocyte phenotype. Moreover, the in vivo cell reprogramming towards pluripotency in fully developed mammals shown here is also in agreement with recently reported cell reprogramming for non-mammalian tissue [35] and did not lead to any structural or functional side effects in the liver, nor did it lead to any manifestation of carcinogenesis or teratoma formation.
Further progress from this work will be the isolation and characterization of the transcriptionally reprogrammed cells from the primary hepatocyte extracts. Determination of the quality and level of functional pluripotent capacity of those cells, their genomic stability and epigenetic character, along with all the comparative studies currently undertaken by numerous laboratories to define the similarities and differences between iPS and ES cells will be needed. Also, the mechanistic basis of direct in vivo induced reprogramming of somatic cells to a pluripotent state and the dissection of the reasons behind the rapid kinetics observed, the extent and possible implications of transient cell reprogramming to ground-state pluripotency that could possibly take place transiently in situ, and the mechanisms by which the tissue microenvironment drives rapid re-differentiation in the host cells will have to be determined.
Enhancement of the level of in vivo cell reprogramming to a pluripotent state can also be envisaged by improvements in gene transfer methodology, dosing regimen and vector construct design. Finally, it can be envisioned that improved (and less invasive) protocols and technologies for the possible generation of in vivo induced pluripotent cells may lead to a general methodology for the extraction of autologous cells from tissue (liver or other) rapidly and with minimization of risks (mutagenesis, media contamination, xenobiotic reagents, etc) from culturing conditions and protocols. Overall, this study offers the basis of a rapid, virus-free, efficient, and in the absence of culturing complications in vivo platform to study reprogramming mechanisms in situ using adult, fully developed, somatic tissue. Figure S1 Transfection after HTV injection. Balb/C mice were HTV injected with pCMV?GFP-Luc in 0.9% saline and hepatocytes were isolated after 24 h and analyzed for (a) luciferase activity by luciferase assay (b) GFP and Luc gene expression by real-time PCR. Balb/C mice were HTV injected with pCAG?GFP in 0.9% saline and hepatocytes were isolated after 24 h and analyzed for (c) eGFP gene expression by real-time PCR (d) transfection efficiency by FACS. (e) Liver samples were frozen and sectioned to image transfected hepatocytes under (i) differential interference contrast (DIC) illumination or (ii) blue light excitation (10x).


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CRABP2 Promotes Myoblast Differentiation and Is Modulated by the Transcription Factors MyoD and Sp1 in C2C12 Cells

Cellular retinoic acid binding protein 2 (CRABP2), a member of a family of specific carrier proteins for Vitamin A, belongs to a family of small cytosolic lipid binding proteins. Our previous study suggested that CRABP2 was involved in skeletal muscle development; however, the molecular function and regulatory mechanism of CRABP2 in myogenesis remained unclear. In this study, we found that the expression of the CRABP2 gene was upregulated during C2C12 differentiation. An over-expression assay revealed that CRABP2 promotes myogenic transformation by regulating the cell cycle during C2C12 differentiation. The region from −459 to −4 bp was identified as the core promoter and contains a TATA box, a GC box and binding sites for the transcription factors MyoD and Sp1. Over-expression, site-directed mutagenesis and EMSA assays indicated that the transcription factors MyoD and Sp1 regulate CRABP2 expression and promote myoblast differentiation in C2C12 cells.

Introduction
Myogenesis can be regarded as a two-step process consisting of the determination, in which the satellite cells commit to the mature muscle lineage, and subsequent differentiation of mononuclear myoblasts to multinuclear myotubes [1]. A previous study documented that several myogenesis factors (MyoD, Myf5, MRF4 and MyoG as well as members of the MyoD gene family) are critical for the determination and terminal differentiation of skeletal muscle cells [2]. In cultured non-muscle cells, the exogenous expression of the MyoD gene family can induce myogenic differentiation [3,4,5,6]. The MyoD gene family proteins bind to the CANNTG sequence (also known as the Ebox sequence), which is present in the promoters and enhancers of many muscle-specific genes [7,8]. Through the DNA-protein interaction, the MyoD gene family induces a conformational change and promotes muscle cell-specific gene transcription. Additionally, the MyoD gene family also promotes cell differentiation and inhibits the cell cycle [9]. Sp1 belongs to a family of zinc-finger transcription factors involved in the early development of an organism [10,11]. It contains 3 zinc finger motifs, which bind to GC-rich sequences. Specifically, the Sp1 protein recognizes a motif of GGGCGG or other related GC-rich sequences [12]. The Sp1 protein regulates the expression of several genes involved in cell differentiation and embryonic development, enhancing or repressing gene activity [13,14,15,16].
Vitamin A and its related molecules (retinol, retinaldehyde and retinoic acid) specifically bind many distinct cytoplasmic proteins and play essential roles in vision, growth, reproduction and cell differentiation [17]. The cellular retinoic acid binding proteins (CRABPs) are well-characterised members of a large family of small proteins that specifically bind retinoic acid [18] and mainly exert their biological function within cells. The CRABPs are abundantly expressed in numerous developing tissues [19], which suggests that CRABPs may perform specific functions during morphogenesis [20,21]. CRABP2 is a low molecular mass (15 kDa) protein that belongs to the multi-gene family of cellular retinoic acid binding proteins [22]. During mouse embryonic development, CRABP2 was expressed in many organs [21], we used the GNF SymAtlas expression datasets ( http://symatlas.gnf.org/ SymAtlas/), which demonstrated that CRABP2 mRNA was upregulated from day 6.5 to day 10.5. These data indicate that the CRABP2 gene may play a vital role during embryonic development.
The precise biological function and regulatory mechanism of the CRABP2 gene was unknown. Our recent LongSAGE analysis indicated that the CRABP2 gene contributes to prenatal skeletal muscle development in pigs [23]. Myoblast differentiation is a critical molecular event during foetal muscle development. We hypothesised that the CRABP2 gene plays an important role in myogenesis. In this study, we analysed the biological function and regulatory mechanism of the CRABP2 gene in C2C12 cells.

Results
The CRABP2 gene was upregulated during C2C12 differentiation We analysed the expression of the genes (CRABP1, CRABP2, CRBP1, CRBP2) expressing carrier proteins for Vitamin A and related molecules by RT-PCR during C2C12 differentiation. The results shown reveal expression levels of CRABP1, CRABP2, CRBP1, CRBP2 observed. We found that the CRABP1 and CRBP2 genes were not detected and the mRNA expression of the CRBP1 gene was unchanged, but the CRABP2 gene was markedly upregulated (Figure 1). To further test the expression change of the CRABP2 gene during C2C12 differentiation, quantitative realtime PCR (qRT-PCR) was performed. The results suggested that CRABP2 was upregulated from day 0 to day 4 during myogenic differentiation in C2C12 cells ( Figure 2).

CRABP2 promotes C2C12 differentiation
To study the effects of CRABP2 in C2C12 cells, we constructed CRABP2 lentivirus vector for transit gene expression. Analysis by western blotting verified that CRABP2 was expressed in 293T cells ( Figure 3) and the titer of the lentivirus particle was 2E +9 TU/ml.
After transfecting C2C12 cells with CRABP2 expression vector, we observed cell morphology by fluorescence microscopy and studied a mixture of cell populations with various CRABP2 expression levels by qRT-PCR. We found that the expression levels of CRABP2 significantly increased ( Figure 4) and that the cell morphology was characteristic of differentiation ( Figure 5) in overexpression (OE) group compared with Blank control (CON) and Negative control (NC) groups.
In contrast with the CON and NC groups, FACS cell sorting results of the OE group showed no significant changes in the percentage of cells in the G1 phase, but the percentage of cells in the G2/M phase decreased and the percentage of cells of S phase increased (p,0.05) ( Figure 6). It has been reported that the number of cells in G2/M phase decreases and more cells remain in S phase during the early differentiation of C2C12 cells [24]. Because the percentage of S phase cells was higher and the percentage of G2/M phase cells was lower in OE group than control group, indicating that S phase arrested and the number of cells entered into G2 phase decreased. These data demonstrated that DNA synthesis reduced, C2C12 cell no longer continue to proliferate, may enter the cell differentiation.

Identification of the core promoter region
To determine the functional fragment of the CRABP2 promoter, we inserted a 2.4 kb fragment into the promoter-less vector pGL 3basic. The resulting plasmid (pGL 3 -A) was transfected into C2C12 cells, and the luciferase activity measured. Compared with cells transfected with pGL 3 -basic, the cells transfected with the pGL 3 -A plasmid had significantly increased luciferase activity, revealing that the 2.4 kb fragment was a functional promoter fragment. To further identify the precise promoter region, the four promoter luciferase reporter plasmids (pGL 3 -B, pGL 3 -C, pGL 3 -D and pGL 3 -E) were constructed by further deletion using pMD18 T-A as a template. As shown by transient transfection results, the pGL 3 -E plasmid contained the basal promoter activity (Figure 7).
Four promoter luciferase reporter plasmids (pGL 3 -a, pGL 3 -b, pGL 3 -c and pGL 3 -d) were constructed by further deletion of pGL 3 -E. The results indicated that the pGL 3 -E plasmid with the 2459 bp to 24 bp fragment contained the minimal core promoter region ( Figure 8).
Identification of MyoD and Sp1 as transcription factors of the CRABP2 gene Bioinformatic analyses using TFSEARCH, TESS, Web Promoter Scan Service and MatInspector programs indicated that the CRABP2 promoter contains a TATA box, a GC box and binding sites for the transcription factors MyoD and Sp1. The core promoter region contains the TATA box and the GC box, as well as MyoD and Sp1 binding sites.
To determine experimentally that MyoD and Sp1 are regulators of the CRABP2 gene, an over-expression vector of MyoD and a site-directed mutation vector of Sp1 were constructed. Cotransfection with the minimal core promoter vector was performed to verify the transcription factors MyoD and Sp1, respectively. The results showed that the MyoD and Sp1 sites in the promoter region were necessary and functional to drive the basal reporter expression in transient transfection assays in C2C12 cells ( Figure 9).

Validation of MyoD and Sp1 transcription factors as regulators for the CRABP2 gene
To further confirm the function of MyoD and Sp1, electrophoretic mobility shift assays (EMSA) were carried out to analyse binding capabilities in cell nuclear extracts.
Oligonucleotides were synthesized and BIO-labelled and then incubated with nuclear extracts from C2C12 cells. Specific DNAprotein complexes were identified from C2C12 nuclear extracts ( Figure 10). The binding activities were different between C2C12 myoblasts and C2C12 myotubes. In C2C12 myotubes, the binding activity of MyoD was decreased; however, the binding activity of Sp1 was enhanced. These results indicate that MyoD and Sp1 specifically bind to promoter elements to regulate CRABP2 expression in C2C12 cells at different times during differentiation.

Discussion
Our previous LongSAGE analysis suggested that CRABPs were involved in skeletal muscle development in the pig throughout embryonic development. 23 Therefore, the aim of this study was to investigate the function of CRABPs during the process of differentiation of C2C12 cells into myotubes. The RT-PCR and real time PCR analyses suggested that CRABP2 was increasingly expressed during C2C12 cell differentiation. Interestingly, we found that CRABP2 over-expression accelerated the process of C2C12 differentiation into myotubes in vitro. These observations suggest that CRABP2 contributes to the process of C2C12 myoblast differentiation into myotubes.
We then carried out a promoter analysis through a step-by-step deletion process to identify the core promoter region. Bioinformatics analyses showed that the promoter region contained a TATA box, a GC box, and MyoD and Sp1 transcription factor binding sites. We constructed a MyoD over-expression vector and an Sp1 site-directed mutation vector. Co-transfection was performed to demonstrate the effects of MyoD and Sp1 transcription factors on the CRABP2 gene. The EMSA results indicated that the predicted MyoD and Sp1 binding sites bound to the MyoD and Sp1 proteins, respectively, in nuclear extracts from C2C12 cells. Additionally, excess amounts of unlabelled oligonucleotides inhibited the binding activities in C2C12 cells. This result further proved that the MyoD and Sp1 binding sites were functional regulators for the CRABP2 gene. These results demonstrated that the putative MyoD and Sp1 binding sites in the core promoter of CRABP2 may be responsible for transcriptional regulation in C2C12 cells.
Transcription factors are trans-regulatory factors that exert biological function by interaction with cis-regulatory elements of structural genes. The MyoD family recognizes and binds the consensus sequence CANNTG (N for any nucleotide), and the promoter region of CRABP2 contains a CANNTG sequence. The MyoD gene family forms a very complex regulatory network by interacting with a number of positive and negative factors to regulate CRABP2 gene transcription, controlling cell growth and cell differentiation. The Sp1 transcription factor was involved in regulating cell growth and controlling morphogenetic pathways both in mammals and in invertebrates [25,26]. In our study, Sp1 regulated gene transcription by binding the GGGCGG sequence in specific regulatory sites of the CRABP2 gene. Therefore, MyoD and Sp1 are important regulators for the CRABP2 gene and directly bind to the promoter region to influence CRABP2 expression. CRABP2 is in turn involved in regulating C2C12 cell differentiation.
In summary, CRABP2 expression was upregulated from day 0 to day 4 at the mRNA level in differentiating C2C12 cells. The overexpression of CRABP2 led to cell cycle changes in C2C12 cells in vitro. We have identified a biological function for CRABP2 in C2C12 differentiation and found that MyoD and Sp1 play a regulatory role in CRABP2 gene transcription by binding in the core promoter region between 2459 bp and 24 bp upstream of the CRABP2 gene. These results give new insight into the biological function and regulatory mechanism of the CRABP2 gene.

Materials and Methods
Cell culture and RNA extraction   maintained at 37uC in 5% CO 2 . The C2C12 cells were seeded in six-well plates. After approximately 12-16 h, when cell confluence reached approximately 60%-70%, the differentiation of C2C12 myoblasts into myotubes was induced by the addition of differentiation medium (DMEM containing 2% horse serum instead of 10% FBS).
Starting at the beginning of differentiation, the C2C12 cells were cultured in six-well plates and harvested for RNA extraction at 0, 1, 2, 3 and 4 days. Total RNA samples were extracted using TRIZOL (Invitrogen, USA) according to the manufacturer's instructions. The cDNA samples were obtained by reverse transcription from 1 mg RNA using oligo(dT) and M-MLV reverse transcriptase (Promega, USA).

RT-PCR analysis
The RT-PCR (Reverse Transcription-Polymerase Chain Reaction) was performed in a volume of 10 ml containing 1 ml of 106 PCR buffer (plus Mg 2+ ), 100 mmol/L of each dNTP, 0.5 mmol/L of each PCR primer, 1.0 U Taq DNA polymerase (Takara, Japan) and 0.5 ml of cDNA template. The primers were designed and synthesized as shown in Table 1. The PCR reaction was 5 min at 95uC followed by 27 cycles of 30 s at 95uC, 30 s at the specific melting temperatures of each primer pair, 30 s at 72uC and a final extension time of 5 min at 72uC.

Real-time PCR analysis
Real-time PCR amplifications were carried out using the Light Cycler Real Time PCR instrument (model 7500; Applied Biosystems, Inc., Foster City, CA). The PCR reaction contained 506 ROX Reference DyeII, 26 SYBR Green Realtime PCR Master Mix (Takara, Japan), 10 mM primers and 1.0 ml of cDNA template. Pure water was added for a total volume of 20.0 ml. Gene amplification was performed under the following conditions: 95uC for 30 s, followed by 40 cycles of 95uC for 5 s, 60uC for 20 s and 72uC for 34 s. The PCR was performed in biological triplicate for each condition. The target gene data was analysed and normalized to GAPDH mRNA levels.

Construction and transfection of the CRABP2 lentivirus vector
The vector overexpressing CRABP2 was constructed to study its function in C2C12 cells. The cDNA fragment was isolated and ligated into the pGC-FU vector linearized by cutting with Age I restriction enzyme, purchased from GeneChem Co., Ltd, Shanghai, China. The expression plasmid containing the fusion protein was extracted using ultrapure endotoxin-free extraction kits (Omega, USA). The transfection of 293T cells was carried out using Lipofectamine 2000 according to the manufacturer's instructions. After a 48 h incubation, the cell supernatant containing virus-like particles was collected. After concentration, the viral titres were determined in the 293T cells.
The C2C12 cells were plated on 6-well plates and cultured overnight at 37uC in 5% CO 2 . When cell confluence reached approximately 60-70%, lentivirus of a specific quantity was added to cells. After 12 h, the transfection medium was removed and replaced with normal growth medium.     After 3-4 days, the GFP expression was tested by fluorescence microscopy to determine transfection efficiency. The cells were then harvested and used for the cell cycle assay and RNA extraction. All transfections were performed in triplicate for each plasmid.

Flow cytometry analysis (FCM)
The C2C12 cells were collected and centrifuged at 1200 rpm for 5 min. The cells were washed twice in 4uC pre-chilled PBS (phosphate buffered saline, pH = 7.2-7.4), fixed with 4uC pre-chilled 70% ethanol, centrifuged at 1500 rpm for 5 min to remove stationary phase cells and resuspended in PBS. The cells were filtered once through 400 mesh and centrifuged at 1200 rpm for 5 min, and after the PBS supernatant was removed, the cells were then stained with 1 ml of PI at 4uC in darkness for 30 min. Finally, the cells were analysed by flow cytometry (FACSCalibur, BD, USA).

Construction of the luciferase reporter plasmids
The 59-upstream 2.4 kb promoter fragment of the CRABP2 gene was amplified from mouse genomic DNA and then inserted into the pMD18-T vector. It was named pMD18 T-A.
The construction of the luciferase reporter gene plasmid was carried out as follows: the pMD18 T-A vector was cut using MluI and NcoI restriction enzymes and then ligated into linearized basic vector (named pGL 3 -A). Fragments of different lengths from the 59-upstream sequence were amplified using pMD18T-A as a template and then ligated into linearized pGL 3 -basic vector (named pGL 3 -B, pGL 3 -C, pGL 3 -D and pGL 3 -E). The plasmids pGL 3 -A, pGL 3 -B, pGL 3 -C, pGL 3 -D and pGL 3 -E were constructed to contain 22292 bp to +115 bp, 21646 bp to +115 bp, 21111 bp to +115 bp, 2719 bp to +115 bp and 2459 bp to +115 bp of the CRABP2 gene, respectively.
To identify more precisely the core promoter region, we performed further deletions on the promoter fragment pGL 3 -E: four luciferase reporter gene plasmids, pGL 3 -a (2290 bp to +115 bp), pGL 3 -b (2178 bp to +115 bp), pGL 3 -c (24 bp to +115 bp) and pGL 3 -d (+44 bp to +115 bp), were constructed using the above methods. Each promoter luciferase plasmid was constructed using a sequence-specific forward primer and a common reverse primer (shown in Table 2). The sequences were analysed by Invitrogen.
All experiments were performed in triplicate, and the data were presented as mean 6 SD. Student's t-test was used to determine statistical significance.  Luciferase assay The C2C12 cells were plated on 24-well plates and cultured overnight at 37uC to ensure approximately 60%-70% confluence. Co-transfections were performed using 2.0 ml of Lipofectamine 2000 reagent (Invitrogen, USA), 0.8 mg of the firefly luciferase plasmid DNA and 0.008 mg of pRL-TK plasmid DNA (Promega, USA) as an internal control. The transfection medium was removed and replaced with growth medium after 5 h.
The Firefly and Renilla luciferase activities were measured at 24 h after transient transfection using the Dual-Glo Luciferase assay system (Promega, USA) and a TD20/20 luminometer (Turner Designs) according to the manufacturer's instructions. Each plasmid was tested in three independent experiments. The luciferase activity was normalized using the Renilla luciferase activity levels and expressed as relative luciferase unites (RLU) to reflect the gene promoter activity.

Construction of the MyoD expression vector
The cDNA sequence of the MyoD gene was amplified using mouse muscle mRNA with the primers listed in Table 1. The PCR product was cloned into the pMD18-T vector (Invitrogen, USA) and then inserted into the EcoRV/NotI sites in the linearized pcDNA3.1(+) vector (Invitrogen, USA). The recombinant plasmid was sequenced by Invitrogen.

Construction of the Sp1 site-directed mutation vector
To identify Sp1 as a regulatory factor for the CRABP2 gene in C2C12 cells, we constructed a site-directed mutation vector of Sp1. The site-directed mutagenesis of the Sp1 binding site was obtained by a two-step PCR protocol using pMD18 T-A as a template. The original GGC site was mutated to TTA. The mutagenic primers were 59 AGGTGGTGTGGAAGGCGGT-TAGGGGGCGGGGCCGCCTCATGCACCAGCT 39 (forward primer) and 59 CCTAACCGCCTTCCACACCACCT 39 (reverse primer). Thermocycler parameters were set at 95uC for 5 min followed by 27 cycles of 95uC for 30 s, 60uC for 30 s and 72uC for 30 s with a final extension time of 5 min at 72uC. The final PCR product was cloned into pGL 3 -basic and named pGL 3 -muE.

Electrophoretic mobility shift assay (EMSA)
Nuclear extracts from C2C12 myoblasts and myotubes were prepared according to the instruction manual of the NProtein Extraction kit (Exprogen, Beijing, China). The proteins were measured using a BCA Protein Assay Kit (Exprogen, Beijing, China). The probes for MyoD and Sp1 were synthesized by Integrated DNA Technologies (IDT, USA).
The binding reactions were performed in a 10 ml mixture containing 1.0 ml of 106 Binding Buffer, 1.0 ml of Poly (dI:dC), 2 mg of nuclear extract and 0.5 ml of Bio-labelled probes. For the competition experiments, unlabelled probes were added to the binding reaction mixture and co-incubated. Subsequently, DNAprotein complexes were separated by 6.5% non-denaturing polyacrylamide gel electrophoresis in Tris-Boric acid (TBE) buffer. Electroblotting and chemiluminescence detection were performed according to the instructions of the BiotinLight TM EMSA Kit (Exprogen, Beijing, China). Results were observed by chemiluminescence imager.


s2fieldsofstudy: Biology Chemistry

text:
Aromadendrin Inhibits Lipopolysaccharide-Induced Nuclear Translocation of NF-κB and Phosphorylation of JNK in RAW 264.7 Macrophage Cells

Aromadendrin, a flavonol, has been reported to possess a variety of pharmacological activities such as anti-inflammatory, antioxidant, and anti-diabetic properties. However, the underlying mechanism by which aromadendrin exerts its biological activity has not been extensively demonstrated. The objective of this study is to elucidate the anti-inflammatory mechanism of aromadedrin in lipopolysaccharide (LPS)-stimulated RAW 264.7 macrophage cells. Aromadendrin significantly suppressed LPS-induced excessive production of pro-inflammatory mediators such as nitric oxide (NO) and PGE2. In accordance, aromadendrin attenuated LPSinduced overexpression iNOS and COX-2. In addition, aromadendrin significantly suppressed LPS-induced degradation of IκB, which sequesters NF-κB in cytoplasm, consequently inhibiting the nuclear translocation of pro-inflammatory transcription factor NF- κB. To elucidate the underlying signaling mechanism of anti-inflammatory activity of aromadendrin, MAPK signaling pathway was examined. Aromadendrin significantly attenuated LPS-induced activation of JNK, but not ERK and p38, in a concentration-dependent manner. Taken together, the present study clearly demonstrates that aromadendrin exhibits anti-inflammatory activity through the suppression of nuclear translocation of NF-κB and phosphorylation of JNK in LPS-stimulated RAW 264.7 macrophage cells.

INTRODUCTION
Macrophages play essential roles in host defense against bacterial infection (Rehman et al., 2012). Macrophages orchestrate the innate immunity response to infection by expressing various infl ammatory cytokines such as TNF-α, IL-1, NO, and PGE 2 . The most common cause of macrophage activation is an exposure to LPS, the principal component of the outer membrane Gram-negative bacteria (Rietschel and Brade, 1992). However, aberrantly over-activated macrophages have also been attributed to detrimental responses in infl ammatory diseases including sepsis, septic shock, or systemic infl ammatory response syndrome (Rackow and Astiz, 1991;Shapiro et al., 2010). Therefore, suppression of aberrant activation of macrophage may have valuable therapeutic potential for the treatment of infl ammatory diseases such as sepsis.
Aromadendrin is a fl avonol, which is a type of fl avonoids.
Recently, it has been reported that aromadendrin possesses anti-diabetic activity by stimulating glucose uptake and ameliorating insulin resistance (Zhang et al., 2011). In addition, it has been also reported that aromadendrin derivatives exhibit various biological activities such as anti-proliferative (Duh et al., 2012) and osteogenic actions (Swarnkar et al., 2011). However, the underlying mechanism by which aromadendrin exerts its pharmacological activity, especially anti-infl ammatory, has not been clearly demonstrated. Aromadendrin has been reported to be present in various medicinal herbs including Chionanthus retusus (Kwak et al., 2009), Cudrania tricuspidata
In the present study, to provide a novel pharmacological agent that could suppress aberrantly activated macrophages, the anti-infl ammatory activity of aromadendrin and its underlying mechanism were elucidated in LPS-stimulated RAW 264.7 macrophage cells.

Reagents and cell culture
Bacterial lipopolysaccharide (LPS) from Escherichia coli serotype 055:B5 was purchased from Sigma-Aldrich (St. Louis, MO, USA). Aromadendrin was isolated and identifi ed from Hemiptelea davidii (Fig. 1) (Chang et al., 2004). Aromadendrin was dissolved in dimethyl sulfoxide (DMSO) and added to the cell culture at the desired concentrations. The macrophage RAW 264.7 cell line was maintained in Dulbecco's modifi ed Eagle's medium (DMEM; Gibco BRL, Grand Island, NY, USA) containing 5% heat-inactivated fetal bovine serum and penicillin-streptomycin (Gibco BRL) at 37 o C, 5% CO 2 . In all experiments, cells were incubated in the presence of the indicated concentration of Aromadendrin before the addition of LPS (200 ng/ml).

Cell viability
Cell viability was determined by 3-(4,5-dimethylthiazol-2yl)-2,5-diphenyltetrazolium bromide (MTT) assay. RAW 264.7 macrophage cells were seeded at 5×10 5 cells per well and incubated with aromadendrin at various concentrations for 24 hr at 37 o C. After incubation, MTT (0.5 mg/ml in PBS) was added to each well, and the cells were incubated for 3 hr at 37 o C and 5% CO 2 . The resulting formazan crystals were dissolved in dimethyl sulfoxide (DMSO). Absorbance was determined at 540 nm. The results were expressed as a percentage of surviving cells over control cells.

Nitrite quantifi cation assay
The production of NO was estimated by measuring the amount of nitrite, a stable metabolite of NO, using the Griess reagent as described . After aromadendrinpretreated RAW 264.7 macrophage cells were stimulated with LPS in 12-well plates for 24 hr, 100 μl of the cell supernatant was mixed with an equal volume of Griess reagent. Light absorbance was read at 540 nm. The results were expressed as a percentage of released NO from LPS-stimulated RAW 264.7 cells. To prepare a standard curve, sodium nitrite was used to prepare a standard curve.

Isolation of nuclear extract
The isolation of the nuclear fraction was performed according to the protocol of NF-κB activation assay kits (FIVE photon).

Statistical analysis
All values shown in the fi gures are expressed as the mean ± SD obtained from at least three independent experiments.
Statistical signifi cance was analyzed by two-tailed Student's t-test. Data with values of p<0.05 were considered as statistically signifi cant. Single (* and # ) and double (** and ## ) marks represent statistical signifi cance in p<0.05 and p<0.01, respectively.

Aromadendrin inhibits NO and PGE 2 production in LPSstimulated RAW 264.7 macrophage cells
Given the previous reports that pro-infl ammatory mediators such as NO and PGE 2 play key roles in the progression of infl ammation (Ock et al., 2009;Lee et al., 2012), the inhibitory ef- fects of aromadendrin on NO and PGE 2 release in LPS-stimulated RAW 264.7 macrophage cells. Cells were incubated with aromadendrin (10, 50, 100, or 200 μM) for 1 hr prior to LPS treatment (200 ng/ml) LPS. LPS markedly increased NO and PGE 2 production in RAW 264.7 cells. However, aromadendrin signifi cantly inhibited NO and PGE 2 release in LPS-stimulated RAW 264.7 cells in concentration dependent manners (Fig. 2  A and B). Furthermore, to examine the anti-infl ammatory magnitude of aromadedrin, inhibitory potency of aromadendrin on LPS-induced NO production was compared to prototype fl avo-noids, such as wogonin and quercetin ( Fig. 2A). Although aromadendrin signifi cantly attenuated LPS-induced NO production in a concentration-dependent manner, the inhibitory effect of aromadendrin was less potent than those of wogonin and quercetin. In addition, aromadendrin showed no signifi cant cytotoxicity in concentration ranges used in the study (Fig. 2C).

Aromadendrin inhibits LPS-induced mRNA and protein expressions of iNOS and COX-2
As aromadendrin inhibited LPS-induced NO and PGE 2 production in RAW 264.7 cells (Fig. 2), whether the suppression of NO and PGE 2 were attributable to downregulation of iNOS and COX-2 expression was examined in the presence or absence of aromadendrin. LPS stimulation of the RAW  www.biomolther.org 264.7 cells strongly upregulated iNOS and COX-2 protein levels. Pretreatment of aromadendrin resulted in a signifi cant decrease in LPS-induced iNOS and COX-2 overexpression levels in a concentration-dependent manner (Fig. 3A). Consistent with the decrease of protein levels, LPS-induced up-regulation of iNOS and COX-2 mRNA expression was signifi cantly attenuated with aromadendrin (Fig. 3B), indicating that attenuated release of NO and PGE 2 is due to decreased expression of their responsible genes by aromadendrin.

Aromadendrin inhibits IκB degradation and NF-κB translocation in in LPS-stimulated RAW 264.7 cells
Activation of the transcription factor NF-κB has been reported to result in the increased transcription of pro-infl ammatory mediators including iNOS and COX-2 (Baldwin, 1996;Moynagh, 2005). IκB has been demonstrated to inhibit nuclear translocation of NF-κB by retaining it in cytoplasm Rehman et al., 2012). It has been also reported that LPS causes activation of IκB kinases, which phosphorylate IκB and then lead to proteasomal degradation of IκB (Baeuerle and Baichwal, 1997). Therefore, in the present study, whether aromadendrin could suppress LPS-induced IκB degradation was examined. LPS treatment resulted in obvious degradation of IκB. However, aromadendrin signifi cantly attenuated LPS-induced degradation of IκB in a concentration-dependent manner (Fig. 4A). Furthermore, whether aromadendrin could prevent nuclear translocation of the p65 subunit of NF-κB was examined. LPS treatment caused a signifi cantly increased nuclear translocation of NF-κB. However, aromadendrin resulted in a signifi cant reduction of LPS-induced nuclear translocation of NF-κB in a concentration-dependent manner (Fig. 4B).

Aromadendrin inhibits the phosphorylation of JNK in LPS-stimulated RAW 264.7 cells
It has been previously reported that MAP kinase signaling pathway is implicated in the LPS-induced infl ammatory responses (Guha and Mackman, 2001;Rushworth et al., 2005). In the present study, MAP kinase pathway was examined to understand the underlying signaling mechanism by which aromadendrin exhibits its anti-infl ammatory activity. The effect of aromadendrin on LPS-stimulated phosphorylation of JNK, ERK, and p38 kinase in RAW 264.7 macrophage cells was determined. Cells were pretreated with aromadendrin at indicated concentrations (10, 50, 100, and 200 μM), and then were treated with LPS (200 ng/ml) for 30 min. Aromadendrin signifi cantly suppressed the LPS-induced phosphorylation of JNK in a concentration-dependent manner in RAW 264.7 cells  ing pathway in RAW 264.7 macrophage cells. RAW 264.7 cells were stimulated with 200 ng/ml LPS in the absence or presence of aromadendrin. Western blot analysis was then performed to evaluate the activation of MAP kinase signaling pathway. LPSinduced phosphorylation of JNK was signifi cantly attenuated with aromadendrin treatment, but phosphorylation of ERK and p38 was not decreased with aromadendrin treatment, suggesting that JNK signaling might play a key role in the LPS-induced overactivation of RAW 264.7 cells. β-Actin was used as an internal control. Images are representative of three independent experiments that shows reproducible results. ARD stands for aromadendrin. http://dx.doi.org/10.4062/biomolther.2013.023 (Fig. 5). However, no signifi cant attenuation of ERK and p38 phosphorylation was observed with aromadendrin. The data strongly suggest that JNK signaling plays a key role in LPSinduced infl ammatory response in RAW 264.7 cells.

DISCUSSION
The present study clearly demonstrated that aromadendrin suppresses infl ammatory responses through the suppression of nuclear translocation of NF-κB and phosphorylation of JNK in LPS-stimulated RAW 264.7 macrophage cells. Aromadendrin signifi cantly inhibited LPS-stimulated excessive production of NO and PGE 2 with the suppression of their responsible proteins, iNOS and COX-2, respectively. Aromadendrin also attenuated LPS-induced IκB degradation, nuclear NF-κB translocation, and JNK phosphorylation.
Although macrophages play essential roles in innate immunity for the host defense against infections (Rehman et al., 2012), macrophages also play a key role in sepsis by producing pro-infl ammatory mediators, expression of adhesion molecules and coagulation factors, phagocytosis, and cytoskeletal rearrangement (Sweet and Hume, 1996). It has been reported that aberrantly activated macrophages are implicated in the pathogenesis of many infection-associated disorders including sepsis and septic shock (Rackow and Astiz, 1991;Shapiro et al., 2010). Therefore, regulation of aberrant macrophage activation may have valuable therapeutic potential for the treatment of infl ammatory diseases such as sepsis. It has been reported that LPS activates gene transcription by binding to its membrane receptor in macrophages, TLR4, which induces signal transcription pathways leading to the phosphorylation of kinases such as IκB kinases, protein kinase C, PI3K, and MAPKs. Subsequently, these kinases activate various transcription factors such as NF-κB and AP-1 families (O'Connell et al., 1998;Guha and Mackman, 2001). In accordance, the present study showed that aromadendrin resulted in the signifi cant attenuation of LPS-induced degradation of IκB and nuclear NF-κB translocation. However, effects of aromadendrin on other pro-infl ammatory transcription factors have not been studied. Therefore, further studies are necessary to clearly understand the role of aromadendrin on various infl ammation-related transcription factors.
It has been reported that LPS is a potent activator of all three MAPK pathways in human monocytic cells (Guha and Mackman, 2001). Many studies have shown that LPS activates ERK1/2, JNK, and p38 in macrophages (Sweet and Hume, 1996) and that many of the downstream targets of MAPK pathways are transcription factors, which regulate various genes encoding infl ammatory mediators (Zhang et al., 2006;Lee et al., 2009;Zhang et al., 2011). In the present study, LPS treatment exhibited increased phosphorylation of all three MAPKs. However, aromadendrin selectively attenuated LPS-induced JNK activation in a concentration-dependent manner. No noticeable changes were observed in phosphorylation of ERK and p38 with aromadendrin. Selective suppression of JNK activation appeared to be suffi cient to exert anti-infl ammatory effects of aromadendrin in RAW 264.7 cells, suggesting that JNK pathway rather than ERK and p38 plays a key role in pro-infl ammatory responses in RAW 264.7 cells.
NF-κB is a major transcription factor for the expression of pro-infl ammatory mediators such as iNOS, IL-1β, and TNF-α (Siebenlist et al., 1994;Kuprash et al., 1995) and aberrant activation of NF-κB and its downstream genes have been implicated in various pathological conditions including sepsis and autoimmune diseases (Karin et al., 2001;Li and Verma, 2002). LPS has been reported to cause the nuclear translocation of NF-κB through IκB degradation (Moon et al., 2007;Zheng et al., 2008). Accordingly, the present study showed that aromadendrin signifi cantly attenuated LPS-induced IκB degradation and nuclear translocation of NF-κB in LPS-induced RAW 264.7 macrophages.
In conclusion, the present data demonstrate that aromadendrin possesses anti-infl ammatory activity such as suppression of NO and PGE 2 production, and inhibition of nuclear transclocation of NF-κB and phosphorylation of JNK in LPS-stimulated RAW 264.7 macrophage cells. The present study strongly suggests that aromadendrin might be a valuable therapeutic agent in the treatment of infl ammation-related pathologies including sepsis. However, further studies are necessary to clearly elucidate the exact mechanism by which aromadendrin inhibits LPS-induced activation of JNK signaling pathway and the possible crosstalk between NF-κB activation and JNK signaling pathway.