extfieldsofstudy: Biology Medicine

text:
Cell aggregation promotes pyoverdine-dependent iron uptake and virulence in Pseudomonas aeruginosa

In Pseudomonas aeruginosa the Gac signaling system and the second messenger cyclic diguanylate (c-di-GMP) participate in the control of the switch between planktonic and biofilm lifestyles, by regulating the production of the two exopolysaccharides Pel and Psl. The Gac and c-di-GMP regulatory networks also coordinately promote the production of the pyoverdine siderophore, and the extracellular polysaccharides Pel and Psl have recently been found to mediate c-di-GMP-dependent regulation of pyoverdine genes. Here we demonstrate that Pel and Psl are also essential for Gac–mediated activation of pyoverdine production. A pel psl double mutant produces very low levels of pyoverdine and shows a marked reduction in the expression of the pyoverdine-dependent virulence factors exotoxin A and PrpL protease. While the exopolysaccharide-proficient parent strain forms multicellular planktonic aggregates in liquid cultures, the Pel and Psl-deficient mutant mainly grows as dispersed cells. Notably, artificially induced cell aggregation is able to restore pyoverdine-dependent gene expression in the pel psl mutant, in a way that appears to be independent of iron diffusion or siderophore signaling, as well as of recently described contact-dependent mechanosensitive systems. This study demonstrates that cell aggregation represents an important cue triggering the expression of pyoverdine-related genes in P. aeruginosa, suggesting a novel link between virulence gene expression, cell–cell interaction and the multicellular community lifestyle.

Introduction
Pseudomonas aeruginosa is a metabolically versatile Gram-negative bacterium and an opportunistic pathogen in cystic fibrosis (CF) and otherwise critical patients, causing both chronic and acute infections (Driscoll et al., 2007). The ability of P. aeruginosa to rapidly adapt to diverse ecological niches and to switch from acute to chronic infections is related to the tightly regulated expression of specific sub-sets of genes in response to environmental cues (Stover et al., 2000).
Characteristic traits of P. aeruginosa chronic infection are the microcolony and biofilm mode of growth (Parsek and Singh, 2003). Microcolonies are small aggregates of cells that open the way to the communal organization of biofilms, which are surface-associated communities of bacteria encased in a selfgenerated polymeric matrix (Zhao et al., 2013). Extracellular polysaccharides represent a key component of the P. aeruginosa biofilm matrix, and are involved in surface attachment and cellcell interaction (Ma et al., 2009).
Pseudomonas aeruginosa strains can produce three main exopolysaccharides, namely alginate, Pel, and Psl (Colvin et al., 2012). Alginate confers the typical mucoid phenotype to producing strains (Sherbrock-Cox et al., 1984), and plays a crucial role in CF lung colonization. Psl and Pel are normally produced by non-mucoid strains, and failure to produce Pel and/or Psl exopolysaccharides impairs biofilm formation in vitro (Colvin et al., 2012). The exopolysaccharide Psl consists of repeating pentamers of D-mannose, L-rhamnose, and D-glucose (Ma et al., 2007). The helical distribution of Psl on the cell surface promotes cell-cell and cell-surface interactions in microcolonies (Zhao et al., 2013). Psl also plays an essential role in the maintenance of the mature biofilm structure (Jackson et al., 2004;Ma et al., 2009). The exopolysaccharide Pel was originally identified by a transposon mutagenesis screening for the loss of surface pellicle formation in P. aeruginosa PA14 (Friedman and Kolter, 2004). Pel structure has not yet been determined, although carbohydrate analysis suggested a glucose-rich composition (Friedman and Kolter, 2004).
The production of the Pel and Psl exopolysaccharides is controlled by many regulatory networks at the level of transcription, translation, and biosynthesis (Goodman et al., 2004;Ventre et al., 2006;Lee et al., 2007;Sakuragi and Kolter, 2007;Gilbert et al., 2009). The Gac system and the signaling molecule cyclic diguanylate (c-di-GMP) are the best characterized regulatory networks involved in the regulation of pel and psl gene expression and, consequently, in the switch from the planktonic to the biofilm lifestyle. The Gac system relies on the sensor kinase GacS that, in response to a still unknown signal, activates the transcriptional regulator GacA, which in turn promotes the transcription of two small non-coding RNAs (sRNAs), RsmZ and RsmY . These sRNAs bind to and sequester the mRNA binding protein RsmA, thereby inhibiting its activity as translational repressor (Heeb et al., 2006). While psl gene expression is directly repressed by RsmA at the translational level (Irie et al., 2010), there is no evidence of a direct effect of RsmA on the pel genes, although transcriptomic analysis showed that the active state of the Gac network (i.e., rsmA mutation) also promotes transcription of the pel operon (Brencic and Lory, 2009). The Gac system has recently been shown to positively affect the intracellular levels of the signaling molecule c-di-GMP, which also induces exopolysaccharides production (Moscoso et al., 2011(Moscoso et al., , 2014Irie et al., 2012;Frangipani et al., 2014). This second messenger regulates Pel production both at the transcriptional and post-transcriptional level, by inhibiting the activity of the transcriptional repressor FleQ (Hickman and Harwood, 2008), and activating the Pel biosynthesis protein PelD (Lee et al., 2007). Although the molecular mechanism has not been elucidated yet, high intracellular levels of c-di-GMP also increase the expression of psl genes (Hickman et al., 2005).
Besides exopolysaccharides production and biofilm formation, Gac and c-di-GMP also act in a concerted way to promote the expression of pyoverdine genes (Frangipani et al., 2014), and it has recently been reported that the exopolysaccharides Pel and Psl are important for c-di-GMP regulation of pyoverdine production (Chen et al., 2015). Pyoverdine is a green fluorescent siderophore which plays a prominent role in P. aeruginosa pathogenicity Cornelis and Dingemans, 2013). Pyoverdine acts not only as a high-affinity iron scavenger, but also serves as a signal molecule to promote the expression of important P. aeruginosa virulence factors, via a cell-surface signaling cascade which involves the other membrane ferri-pyoverdine receptor FpvA and the cytoplasmic membrane-spanning antisigma factor FpvR, ultimately leading to the activation of the extracytoplasmic function (ECF) sigma factor PvdS (Lamont et al., 2002;Llamas et al., 2014). PvdS directs the transcription of almost thirty P. aeruginosa genes, including those involved in pyoverdine biosynthesis and transport, the gene for the extracellular protease PrpL and, indirectly, the exotoxin A gene toxA Llamas et al., 2014). The dual function of pyoverdine in iron uptake and virulence renders this siderophore essential for P. aeruginosa infectivity, as demonstrated in different mouse models (Meyer et al., 1996;Takase et al., 2000;Imperi et al., 2013). As for any iron-uptake system, pyoverdine production needs to be promptly shut down when intracellular iron levels are sufficiently high. This iron-mediated control occurs through the ferric uptake regulator Fur, an iron-sensing transcriptional repressor which binds to its co-repressor Fe 2+ and inhibits transcription of the sigma factor gene pvdS . Besides Fur-Fe 2+ mediated repression, pyoverdine production was found to be influenced by a number of environmental signals and regulatory pathways, including oxygen and nutrient availability, cellular communication and oxidative stress (reviewed in Llamas et al., 2014).
In the present study we demonstrate that the extracellular polysaccharides Pel and Psl are also essential for Gac-mediated regulation of pyoverdine production, which appears strongly repressed in a pel psl double mutant. We also provide evidence that artificially induced cell aggregation is able to restore pyoverdine-dependent gene expression in the Pel and Psldeficient mutant. Our findings support the hypothesis that cell aggregation, rather than polysaccharide production per se, is an important cue triggering production of pyoverdine and pyoverdine-controlled virulence factors in P. aeruginosa.

Bacterial Strains, Growth Conditions, and Plasmids
Bacterial strains and plasmids used in this study are listed in Supplementary Table S1. The iron-depleted complex medium TSBD (Ohman et al., 1980) or the M9 minimal medium supplemented with 20 mM sodium succinate (Sambrook et al., 1989) were used as iron-poor media, to which FeCl 3 was added at the indicated concentrations when required. Virulence factor production and gene expression assays were performed on bacteria grown at 37 • C in 96-well microtiter plates (250 μl of medium in each well) under static conditions, unless otherwise stated. When specified, TSBD medium was supplemented with L-arabinose, agar, phytagel, or sucrose (Sigma-Aldrich) at the indicated final concentrations. Polystyrene beads (Polybead R Microspheres 3.00 μm, Polysciences) were washed several times with sterile water and then resuspended in TSBD at the desired concentrations.

Generation of Deletion and Conditional Mutants
All the primers and restriction sites used for PCR and cloning are listed in Supplementary Table S2. Deletion mutants in pelABCD, pslABCD, fpvR, pilY1, or pilA were generated using specific derivatives of the pDM4 suicide plasmid (Supplementary Table  S1). The fur conditional mutant was generated using a recently described strategy (Lo Sciuto et al., 2014), based on the mini-CTX1-mediated insertion of the fur coding sequence under the control of an arabinose-dependent promoter into a neutral site of the P. aeruginosa genome, followed by the in-frame deletion of the endogenous fur gene using the pDM4 fur suicide plasmid (Supplementary Table S1), under permissive conditions (i.e., arabinose-containing media). Gene replacements and insertions were verified by PCR and DNA sequencing.

Growth and Pyoverdine Measurements
Growth was measured as the OD 600 of appropriate dilutions of bacterial cultures in sterile growth medium. Pyoverdine production was measured as the OD 405 of culture supernatants appropriately diluted in 0.1 M Tris-HCl (pH 8), and normalized to the OD 600 of the corresponding cultures (Imperi et al., 2009). When indicated, pyoverdine production was normalized to the number of colony forming units/ml.

β-Galactosidase and c-di-GMP Assays
The β-galactosidase activity from P. aeruginosa cells carrying reporter plasmids (Supplementary Table S1) was determined spectrophotometrically using o-nitrophenyl-β-Dgalactopyranoside as the substrate, normalized to the OD 600 of the bacterial culture and expressed in Miller (1972) units. Intracellular c-di-GMP levels were determined by liquid chromatography coupled with tandem mass spectrometry as described (Spangler et al., 2010), and normalized to the corresponding cellular protein content determined using the DC protein assay kit (Bio-Rad) and bovine serum albumin as the standard.

Western Blot Analysis of ToxA and PrpL Enzymatic Assay
For detection of ToxA, 100-μl aliquots of culture supernatants were supplemented with 20 μl of 6× SDS-PAGE loading dye [375 mM Tris-HCl (pH 6.8), 9% SDS, 50% Glycerol, 0.03% Bromophenol blue]. To normalize the amount of secreted proteins to bacterial growth, the volume of supernatants loaded into SDS-PAGE gels was calculated according to the formula: loading volume (μl) = 10/OD 600 of the corresponding bacterial culture. Proteins resolved by SDS-PAGE were electrotransferred onto a nitrocellulose filter (Hybond-C extra, Amersham), and probed for ToxA using a rabbit polyclonal anti-ToxA antibody (Sigma-Aldrich). Filters were developed with 5-bromo-4-chloro-3-indoyl-phosphate and nitro blue tetrazolium chloride reagents for colorimetric alkaline phosphatase detection (Promega).
PrpL activity was determined as previously described (Imperi et al., 2010), using the chromogenic substrate Chromozym PL (tosyl-Gly-Pro-Lys-p-nitroanilide; Sigma-Aldrich) which is specific for PrpL (protease IV) and is not cleaved by other P. aeruginosa proteases (Caballero et al., 2001). Briefly, 10 μl of culture supernatants were mixed with 10 μl of 2 mg/ml Chromozym PL and 180 μl of phosphate buffer (pH 7.0) in 96well microtiter plates. The OD 410 was read at 2 min intervals for 30 min in a Victor 3 V plate reader (Perkin-Elmer), and the change in optical density ( OD 410 ) per minute was determined. PrpL activity units were determined as: ( OD 410 × total assay volume/ml)/(sample volume × E410 × light path); where total assay volume was 200 μl, sample volume was 10 μl, the extinction coefficient (E) of the product (p-nitroaniline) at 410 nm was 9.75, and the light path was 0.35 cm. PrpL activity was then normalized to the OD 600 of the corresponding bacterial culture.

Quantitative Real-Time Reverse-Transcription PCR (qRT-PCR)
Total RNA was purified by using RNeasy minicolumns, treated with DNase and re-purified with the RNeasy MinElute cleanup kit (Qiagen). cDNA was reverse transcribed from 0.5 μg of total RNA with PrimeScript RT reagent Kit (TaKaRa). cDNA was then used as the template for qRT-PCR in a 7300 Real-Time PCR System (Applied Biosystems) using SYBR green with the ROX detection system (Bio-Rad). The primers used for qRT-PCR are listed in Supplementary Table S2. At least three wells were run for each cDNA sample. Relative gene expression with respect to the housekeeping gene rpsL was calculated using the 2 − Ct method (Livak and Schmittgen, 2001).

Confocal Microscopy
Thirty microliter of P. aeruginosa cultures in TSBD or M9 were spotted on a glass slide that was freshly coated with 0.5% agarose in water, and covered with a cover slip. Images were acquired with a Leica TCS SP5 inverted confocal microscope equipped with a HCX PLAPO lamba blue 40X/1.25 OIL objective (Zeiss). Images were recorded using specific sets for GFP (excitation at 488 nm, emission window from 500 to 600 nm).

Laser Diffraction Analysis (LDA)
Laser diffraction analysis was performed in a Mastersizer 2000 (Malvern Instrument, UK) as previously described (Schleheck et al., 2009) with minor modifications. In particular, the following settings were used: stirring speed 1000 rpm, laser intensity 79%, 5,000 size scan, Frauenhofer count mode. The amount of bacterial suspension was adjusted to result in an LDA-obscurity of 2-6%. Size distribution scans are quantified by LDA and expressed as volume percentage, corresponding to the volumetric contribution of each particle size class to the total volume of all particle size classes (Schleheck et al., 2009).

Statistical Analysis
Statistical analysis was performed with the software GraphPad Instat, using One-Way Analysis of Variance (ANOVA), followed by Tukey-Kramer multiple comparison tests.

Exopolysaccharide Production is Essential for Gac-Mediated Regulation of Pyoverdine Production
It has recently been demonstrated that the presence of the Pel and/or Psl polysaccharides is required for the ability of the second messenger c-di-GMP to positively regulate the production of the P. aeruginosa siderophore pyoverdine (Chen et al., 2015).
In order to verify whether the Pel and Psl exopolysaccharides are also involved in the Gac-mediated control of pyoverdine gene expression, we generated single and double deletion mutants in pel and psl genes in P. aeruginosa PAO1 (wildtype) and in isogenic mutants in which the Gac signaling is inactive (rsmY rsmZ mutant) or constitutively active (rsmA mutant; Supplementary Table S1). In agreement with previous observations (Frangipani et al., 2014), pyoverdine production in the iron-poor TSBD medium was higher in the rsmA mutant and strongly reduced in the rsmY rsmZ mutant compared with the wild type (WT; Figure 1A). This pyoverdine production profile was maintained in strains lacking either Pel or Psl, while pyoverdine production was drastically reduced in the pel psl double mutant irrespective of the activation state of the Gac system ( Figure 1A). This response appeared to be independent of the previously reported effect of the Gac system on c-di-GMP production (Moscoso et al., 2011;Frangipani et al., 2014), since mass spectrometry analysis revealed that the Gac system is still able to control intracellular c-di-GMP levels in a Pel/Psl deficient background ( Figure 1B). Thus, both the Gac system ( Figure 1A) and the c-di-GMP second messenger (Chen et al., 2015) require at least one of two exopolysaccharides Pel and Psl to exert their control on pyoverdine biosynthesis, indicating that these exopolysaccharides, or an exopolysaccharide-dependent phenotype, are implicated in pyoverdine gene regulation. Indeed, we observed that the expression of the pyoverdine biosynthetic gene pvdD was greatly reduced in the pel psl mutant relative to the WT, as well as that of the pyoverdine-dependent Frontiers in Microbiology | www.frontiersin.org virulence genes toxA and prpL ( Figure 1C). Accordingly, the levels of the corresponding virulence factors in culture supernatants were significantly lower in the pel psl double mutant as compared to the WT strain (Figures 1D,E), as also observed for pyoverdine ( Figure 1A). Therefore, lack of the Pel and Psl exopolysaccharides in P. aeruginosa causes a significant reduction not only in the production of the siderophore pyoverdine but also of pyoverdine-dependent virulence factors.

Exopolysaccharides Affect Pyoverdine Production Independently of Fur and Pyoverdine Signaling
The presence/absence of extracellular polysaccharides on the bacterial cell surface could influence the transport of nutrients and/or signal molecules. Since pyoverdine gene regulation is directly or indirectly controlled by the intracellular levels of iron, through the iron sensor Fur, and by the availability of ironloaded pyoverdine on the cell surface, through the pyoverdine signaling cascade, we investigated the role of Fur and pyoverdine signaling in the Pel and Psl control of pyoverdine-dependent gene expression.
Since Fur is considered an essential protein in P. aeruginosa due to the inability to obtain fur null mutants in this species (Barton et al., 1996;Cornelis et al., 2009), to verify the involvement of Fur in exopolysaccharide-mediated control of pyoverdine production we generated a fur conditional mutants in WT and pel psl backgrounds, by replacing the native fur gene with an arabinose-inducible allele. The PAO1 fur conditional mutant grew poorly in the iron-poor TSBD medium, unless arabinose was added to the medium (Supplementary Figure S1). Compared with the WT strain, pyoverdine levels were significantly higher in the fur conditional mutant grown without arabinose, and were not shut down by the addition of iron to the growth medium (Supplementary Figure S1). This evidence confirmed the suitability of our conditional mutagenesis strategy to obtain Fur-depleted P. aeruginosa cells. Notably, the differences in pyoverdine production between fur and pel psl fur conditional mutants in the absence of arabinose were comparable to those observed for the WT and the pel psl mutant (Figure 2A), clearly indicating that Pel and Psl control pyoverdine gene expression in a way that is independent of Fur. Although not strictly instrumental in the present study, the fur conditional mutant generated in this work will represent a valuable tool for better understanding iron uptake regulation and metabolism in P. aeruginosa.
To test the effect of exopolysaccharides on pyoverdine signaling, we deleted the fpvR gene in both the WT and Pel-and Psl-negative backgrounds, in order to obtain strains in which PvdS activity cannot be influenced by pyoverdine signaling through the anti-sigma factor FpvR (Lamont et al., 2002;Tiburzi et al., 2008). Again, deletion of fpvR did not restore pyoverdine production in the pel psl mutant (Figure 2B), ruling out the involvement of FpvR and, thus, of the pyoverdine signaling cascade on the exopolysaccharides-mediated control of pyoverdine production.

Artificially Induced Cell Aggregation Restores Production of Virulence Factors in Exopolysaccharide-Defective Cells
Our findings argue for a prominent role of extracellular polysaccharides in triggering the expression of pyoverdinedependent virulence genes. As shown in Figure 1A, each single exopolysaccharide has the ability to promote pyoverdine production, suggesting that a phenotype that depends on the presence of exopolysaccharide(s), rather than a specific exopolysaccharide molecule, could be important for pyoverdinerelated gene expression. It is well known that extracellular polysaccharides have a role in surface attachment and biofilm formation (Colvin et al., 2012). However, exopolysaccharides have also been proposed to mediate planktonic aggregation (Klebensberger et al., 2007;Alhede et al., 2011), as confirmed by our observation that Pel and Psl-deficient cells prevalently grow in TSBD medium as dispersed cells, while Pel and Pslproficient WT cells grow as large aggregates including hundreds of cells ( Figure 3A). These bacterial aggregates appear quite loose, and can be partially dispersed by vigorous pipetting (data not shown). This qualitative evidence was confirmed by determining the size and distribution of planktonic cells and cell aggregates by LDA. This analysis revealed that cell aggregates with a size ranging from 30 to 600 μm represent more than 85% of the bacterial population in planktonic cultures of the WT strain. In contrast, cell aggregates were not detectable in planktonic cultures of the pel psl mutant, which only contained single cells (Figure 3B). Such planktonic aggregates were also observed in cultures of the WT PAO1 grown in M9 minimal medium, while they were not detected in pel psl cultures (Supplementary Figure S2). Notably, also in this medium pyoverdine production was strongly reduced in the pel psl mutant with respect to the WT strain (Supplementary Figure S2). We thus hypothesized that cell-to-cell contacts and/or cell aggregation, instead of exopolysaccharides by themselves, could represent the signal triggering pyoverdine-dependent gene expression. This hypothesis was first tested by growing WT and pel psl mutant cells as colonies on the surface of TSBD medium solidified with different gelling substances, such as polyacrylamide and the polysaccharides agar and phytagel, and qualitatively assessing pyoverdine production by comparing fluorescence emission under UV light . While the fluorescence of the pel psl mutant growing as single cells in liquid medium was much lower than that of the aggregated WT and comparable to that of the pyoverdine-deficient mutant PAO1 pvdA , the Pel and Psl-deficient mutant showed fluorescence levels that were indistinguishable from those of the WT, and much higher than those of the pvdA mutant, during aggregated (colony) growth on solid surfaces ( Figure 4A). Although this assay clearly showed that growth on solid surfaces, where cells can interact with each other irrespective of aggregative polysaccharides, enhanced pyoverdine production in the pel psl mutant, the qualitative nature of the assay did not allow a quantitative comparison of pyoverdine levels between strains.
To overcome this limitation, strains were cultured in TSBD medium supplemented with different sub-gelling concentrations of agar (0.0125-0.2%, Figure 4B) or phytagel (0.08-0.125%, Supplementary Figure S3), thus allowing to quantitatively assess pyoverdine levels in culture supernatants. Pyoverdine levels in the supernatants of pel psl mutant cultures were increased by sub-gelling concentrations of agar in a concentration-dependent manner, while agar had negligible effects on pyoverdine production by the WT strain ( Figure 4B). Substantially similar results were obtained with phytagel (Supplementary Figure S3). LDA and microscopic analyses confirmed that the presence of sub-gelling concentrations of agar in the growth medium forced the pel psl mutant to grow as large clusters of cells comparable to those observed for the WT (Figure 4C), consistent with the conclusion that cell aggregation stimulates pyoverdine production.
Notably, pyoverdine production by the pel psl mutant in the presence of agar was abrogated by agar degradation with β-agarase I, which however, had no effect on pyoverdine production by WT PAO1 (Figure 4B). This result indicates that the effect of agar on the pel psl mutant is not related to any specific constituent of the polysaccharide matrix or to a general increase in the osmolarity of the growth medium. In fact, differently from agar and phytagel, high concentrations of sucrose (up to 10%) did not stimulate pyoverdine production by the exopolysaccharidesnull mutant (Supplementary Figure S3). These evidences are in line with the finding that gelling agents only promoted pyoverdine production in the pel psl mutant but not in the WT (Figure 4 and Supplementary Figure S3), indicating that this effect is specific to Pel and Psl-deficient cells. Coherent with the pyoverdine production profile, addition of 0.2% agar stimulated pvdD promoter activity (Figure 4D), as well as PrpL and ToxA production by the pel psl mutant (Figures 4E,F), while it had no effect on the expression of the housekeeping gene proC (Figure 4D). Notably, pvdD promoter activity in the pel psl mutant was also restored to WT levels by growing cells on the surface of TSBD solidified with polyacrylamide ( Figure 4G), indicating that bacterial aggregation and/or cell contacts can trigger pyoverdine gene expression in the absence of any endogenous or exogenously added exopolysaccharide.
It appears therefore that artificially induced cell aggregation, obtained by growing cells either as colonies on solid surface or in liquid cultures in the presence of aggregating agents, is able to restore pyoverdine-dependent phenotypes in the Pel and Psldeficient mutant, supporting the idea that cell-to-cell contacts and/or growth as aggregates are by themselves cues that stimulate pyoverdine production and virulence in P. aeruginosa.

Contacts with Abiotic Surface and the Mechanosensors PilY1 and Type IV Pili are not involved in Aggregation-Mediated Activation of Pyoverdine Production
In order to verify whether the stimulation of pyoverdine production by cell aggregation was due to specific cell-to-cell interactions or to an increase in surface contacts during growth as cellular aggregates, the pel psl mutant was cultured in the presence of 5 × 10 7 or 5 × 10 8 polystyrene beads per ml FIGURE 4 | Cell aggregation is involved in Pel and Psl-mediated control of pyoverdine-dependent virulence factors. (A) Fluorescent phenotype upon UV light exposure of P. aeruginosa PAO1, the pel psl mutant and the pvdA mutant (used as pyoverdine-deficient negative control) grown in liquid TSBD medium (Control) or on TSBD solidified with 1.5% agar, 1% phytagel, or 10% polyacrylamide. (B) Pyoverdine production by P. aeruginosa PAO1 and the pel psl mutant grown in TSBD supplemented with increasing concentrations of agar (0-0.2%) and/or β-agarase I (3.3 units/ml). (C) Representative LDA particle-size scans of P. aeruginosa PAO1 and pel psl cultures in TSBD supplemented with 0.2% agar after 14 h of growth at 37 • C. (D) Activity of the PpvdD::lacZ and PproC'-'lacZ reporter fusions, (E) PrpL enzymatic activity and (F) ToxA levels in culture supernatants from P. aeruginosa PAO1 and pel psl cultures in TSBD supplemented or not with 0.2% agar. Bacteria were cultured in TSBD at 37 • C under static conditions for 14 h. (G) Activity of the PpvdD::lacZ transcriptional fusion in P. aeruginosa PAO1 and pel psl cells cultured on TSBD solidified with 10% polyacrylamide. Values in (B,D,E,G) are the mean (±SD) of at least three independent assays. Asterisks indicate statistically significant differences with respect to the wild type strain (PAO1) grown under the same culture conditions ( * * p < 0.01, * * * p < 0.001). Images in (A,F) are representative of two independent experiments giving similar results.
(3-μm size, Polysciences), under vigorous shaking in order to maintain beads in suspension. We calculated that the addition of 5 × 10 8 beads/ml with a 3-μm diameter in the well of a 96-wells microtiter plate (containing 250 μl of medium) leads to >20-fold increase in the plastic surface available for cell contacts, relative to cultures without beads. While the WT strain formed large aggregates of bacterial cells around the beads, the pel psl mutant only attached to the beads without developing cell aggregates (Figure 5A), indicating that the presence of the beads actually increases the number of contacts between bacterial cells and the inert plastic material. However, polystyrene beads had no effect on pyoverdine production by the pel psl mutant or the WT used as control (Figure 5B), suggesting that cell aggregation, rather than non-specific surface contacts, represents the cue which triggers pyoverdine gene expression in P. aeruginosa. The inset is a 2× magnification of the highlighted area. (B) Pyoverdine production by P. aeruginosa PAO1 and pel psl in the presence or in the absence of 3-μm size polystyrene beads (5 × 10 7 or 5 × 10 8 beads/ml), normalized to the number of colony forming units/ml and expressed as percentage with respect to the untreated wild type. Bacteria were grown in TSBD at 37 • C for 14 h under vigorous shaking (220 rpm). (C) Pyoverdine production by P. aeruginosa PAO1, pel psl and their corresponding pilY1 or pilA deletion mutants after 14 h of growth in TSBD at 37 • C under static conditions. Values are the mean (±SD) of at least three independent assays. Asterisks indicate statistically significant differences with respect to the corresponding parental strain grown under the same culture conditions ( * p < 0.05, * * p < 0.01, * * * p < 0.001).
To further verify this hypothesis, we investigated the possible involvement in aggregation-mediated induction of pyoverdine production of two P. aeruginosa mechanosensors, namely PilY1 and type IV pili, which have recently been implicated in surface contact-dependent activation of virulence gene expression (Siryaporn et al., 2014;Persat et al., 2015). Should PilY1 or type IV pili also be involved in pyoverdine activation in response to planktonic aggregation, the deletion of pilY1 or pilA (encoding the major pilin subunit of type IV pili) should either reduce pyoverdine production in the WT or increase pyoverdine production in the pel psl mutant, in case these mechanosensors act as positive or negative aggregation-dependent regulators, respectively. However, deletion of pilY1 or pilA caused only minor reduction (about 10%) of pyoverdine levels in the WT background, while it was not able to restore pyoverdine production in the pel psl mutant background ( Figure 5C). This result indicates that formerly characterized mechanosensors, such as PilY1 and type IV pili, are not the effectors of aggregationdependent activation of pyoverdine genes.

Discussion
In the last decades, the old vision of bacteria as strictly unicellular organisms living in a planktonic single-cell status was swept away by the finding that bacterial cells in natural, industrial and many clinical settings predominantly exist as biofilms, i.e., structured microbial communities attached to a surface and encased in an extracellular matrix (Mann and Wozniak, 2012). Also during planktonic growth in liquid cultures bacteria can assemble into aggregates of densely packed cells, and it is believed that planktonic aggregation can play a role in resistance to stresses and antibiotics (Schleheck et al., 2009;Blom et al., 2010;Haaber et al., 2012), as well as in microbe-host cell interaction (Lepanto et al., 2011). Although hundreds of studies have investigated the physiology of biofilm-living bacterial cells, very little is known about the effects of cell aggregation during planktonic growth.
Here we provide evidence that growth as planktonic aggregates promotes production of three major virulence factors in the opportunistic pathogen P. aeruginosa, namely pyoverdine, extracellular protease PrpL and exotoxin A. Indeed, we observed that a Pel and Psl-null mutant unable to aggregate in liquid cultures is also defective in the expression of virulence factor genes, and that this effect can be rescued by artificially induced cell aggregation (Figure 4 and Supplementary Figure S3).
Although in this study, we did not elucidate the mechanism by which cell aggregation is perceived by bacterial cells and translated into a transcriptional response which affects pyoverdine-related genes, two hypotheses can reasonably be made to explain the observed effect of planktonic aggregation on gene expression. First, a sensory machinery could transduce a contact signal deriving from the cell envelope into a cytoplasmic response. Our experiments lead to exclude the involvement of physical changes in the cell envelope due to contacts with abiotic surfaces, as well as any role of the two mechanosensors PilY and type IV pili (Figure 5), which have recently been found to induce P. aeruginosa virulence in response to surface contacts (Siryaporn et al., 2014;Persat et al., 2015). Also the P. aeruginosa Wsp system, a chemotaxis-like signal transduction complex which promotes c-di-GMP production by activating the diguanylate cyclase WspR, is stimulated by growth on surfaces (Güvener and Harwood, 2007). However, it has recently been shown that exopolysaccharides depletion in a c-di-GMP overproducing P. aeruginosa strain abrogated the ability of c-di-GMP to promote pyoverdine production (Chen et al., 2015). Accordingly, we found that the overexpression of a constitutively active WspR variant, which results in almost 100-fold increase in intracellular c-di-GMP levels, is unable to induce pyoverdine production in Pel and Psl-deficient cells (Supplementary Figure S4), reasonably excluding also the involvement of the Wsp system. However, other mechanosensitive factors or contact-dependent systems, which likely respond to specific cell-to-cell interactions, could exist in P. aeruginosa. For instance, the P. aeruginosa PAO1 genome is predicted to encode 26 methyl accepting chemotaxis proteins (Whitchurch et al., 2004), 13 cell-surface signaling systems (Llamas et al., 2014), up to 50 canonical two-component systems and more than 10 orphan sensor kinases (Gooderham and Hancock, 2009), many of them being implicated in virulence gene regulation.
The alternative hypothesis is that growth as aggregates determines changes in the cell surrounding environment which could influence virulence gene expression. Studies on the physiology of planktonic aggregates are still at an early stage, but it has recently been reported that localized oxygen depletion occurs in aggregates of P. aeruginosa cells grown in a gelatinbased microtrap (Wessel et al., 2014). Notably, oxygen is a well known inducer of pyoverdine gene expression (Ochsner et al., 1996;Llamas et al., 2014); thus oxygen limitation cannot explain the observed induction of pyoverdine production in bacterial aggregates. On the other hand, since exotoxin A expression is induced under microaerobic conditions through a PvdS-independent mechanism (Gaines et al., 2007), we cannot exclude that the increased exotoxin A expression levels in planktonic aggregates (Figures 1 and 4) could be related, at least in part, to oxygen depletion. It has also been reported that pyoverdine concentration is heterogeneous in P. aeruginosa microcolonies, with a maximum at the colony center (Julou et al., 2013). Although this heterogeneity could influence the efficiency of pyoverdine signaling and/or iron uptake in cell aggregates, we have demonstrated that the aggregation-mediated effect on pyoverdine production is independent of pyoverdine signaling and the iron sensor Fur (Figure 2), ruling out that increased virulence gene expression in aggregated cells may be due to altered pyoverdine signal transduction or dysregulated iron homeostasis. Thus, it is plausible that still uncharacterized chemical and/or physiological changes occurring in densely packed bacterial cells are responsible for the observed activation of pyoverdine-related virulence genes in P. aeruginosa planktonic aggregates.
In summary, our work provides the first evidence that formation of planktonic aggregates stimulates the production of pyoverdine-dependent virulence factors in P. aeruginosa. Further studies are necessary to clarify the mechanistic link between cell aggregation and activation of virulence gene expression. Such a kind of cell contact-or aggregation-dependent induction of virulence could represent a further strategy to modulate bacterial pathogenicity in response to population density, additional or complementary to chemical signaling via quorum sensing. Cellular aggregation also represents the first committed step of biofilm formation. Although some recent transcriptomics and proteomics studies highlighted an overall attenuation of virulence gene expression in mature P. aeruginosa biofilms (Li et al., 2014;Park et al., 2014), which include many slowly growing or quiescent cells, our finding could imply that the virulence potential of P. aeruginosa is increased during the first stages of biofilm formation, when siderophores, extracellular enzymes, and toxins would provide cells with essential nutrients for the energy-demanding process of biofilm development. This hypothesis is in line with the evidence that the gene expression profile of developing P. aeruginosa biofilms is more similar to that of exponential phase cultures rather than mature biofilms (Waite et al., 2006). Finally, since planktonic aggregation seems to be widespread among bacteria (Fazli et al., 2009;Schleheck et al., 2009;Blom et al., 2010;Haaber et al., 2012), it would be interesting to verify whether the correlation between cell aggregation and virulence is conserved in other bacterial pathogens.

Author Contributions
Conceived and designed experiments: DV and FI. Performed the experiments: DV, MP, CB, and FI. Analyzed the data: DV, CB, VK, PV, and FI. Contributed reagents/materials/analysis tools: VK, PV, and FI. Wrote the paper: DV, PV, and FI. All authors read and approved the final manuscript.


β-Lapachone suppresses neuroinflammation by modulating the expression of cytokines and matrix metalloproteinases in activated microglia

Background β-Lapachone (β-LAP) is a natural naphthoquinone compound isolated from the lapacho tree (Tabebuia sp.), and it has been used for treatment of rheumatoid arthritis, infection, and cancer. In the present study, we investigated whether β-LAP has anti-inflammatory effects under in vitro and in vivo neuroinflammatory conditions. Methods The effects of β-LAP on the expression of inducible nitric oxide synthase (iNOS), cytokines, and matrix metalloproteinases (MMPs) were examined in lipopolysaccharide (LPS)-stimulated BV2 microglial cells and rat primary microglia by ELISA, reverse transcription polymerase chain reaction (RT-PCR), and Western blot analysis. Microglial activation and the expression levels of proinflammatory molecules were measured in the LPS-injected mouse brain by immunohistochemistry and RT-PCR analysis. The detailed molecular mechanism underlying the anti-inflammatory effects of β-LAP was analyzed by electrophoretic mobility shift assay, reporter gene assay, Western blot, and RT-PCR analysis. Results β-LAP inhibited the expression of iNOS, proinflammatory cytokines, and MMPs (MMP-3, MMP-8, MMP-9) at mRNA and protein levels in LPS-stimulated microglia. On the other hand, β-LAP upregulated the expressions of anti-inflammatory molecules such as IL-10, heme oxygenase-1 (HO-1), and the tissue inhibitor of metalloproteinase-2 (TIMP-2). The anti-inflammatory effect of β-LAP was confirmed in an LPS-induced systemic inflammation mouse model. Thus, β-LAP inhibited microglial activation and the expressions of iNOS, proinflammatory cytokines, and MMPs in the LPS-injected mouse brain. Further mechanistic studies revealed that β-LAP exerts anti-inflammatory effects by inhibiting MAPKs, PI3K/AKT, and NF-κB/AP-1 signaling pathways in LPS-stimulated microglia. β-LAP also inhibited reactive oxygen species (ROS) production by suppressing the expression and/or phosphorylation of NADPH oxidase subunit proteins, such as p47phox and gp91phox. The anti-oxidant effects of β-LAP appeared to be related with the increase of HO-1 and NQO1 via the Nrf2/anti-oxidant response element (ARE) pathway and/or the PKA pathway. Conclusions The strong anti-inflammatory/anti-oxidant effects of β-LAP may provide preventive therapeutic potential for various neuroinflammatory disorders. Electronic supplementary material The online version of this article (doi:10.1186/s12974-015-0355-z) contains supplementary material, which is available to authorized users.

Background
Microglia are innate immune cells of the central nervous system that constantly move through the brain parenchyma and constitute an immune surveillance system [1,2]. Microglia become activated in response to various stimuli or injury and produce inflammatory mediators such as nitric oxide (NO), cytokines, and matrix metalloproteinases (MMPs). Alternatively, activated microglia produce anti-inflammatory cytokines and lead to matrix deposition and wound healing [3,4]. Thus, the balance between the inflammatory M1 and the anti-inflammatory M2 phase of microglial activation is important to maintain homeostasis in the brain. However, prolonged and unresolved inflammatory response leads to destructive, chronic inflammation (neuroinflammation) that results in neuronal cell death and ultimately in the onset of neurodegenerative diseases [5,6]. Therefore, inhibition of exaggerated inflammatory responses by microglia has been suggested as an important strategy to develop therapeutic agents for various neuroinflammatory disorders.
β-Lapachone (3,4-dihydro-2,2-dimethyl-2H-naphtho[1,2b]pyran-5,6-dione; β-LAP) is a natural compound which was originally isolated from the bark of the South American lapacho tree (Tabebuia avellanedae) [7]. β-LAP has been reported to have a wide variety of pharmacological effects including anti-inflammatory, anti-cancer, antibacterial, anti-fungal, anti-platelet, and anti-angiogenic action [8][9][10]. In particular, β-LAP exerts antineoplastic effects against various human cancer cell lines, and it is now being used in clinical trials for the treatment of various forms of cancer [11][12][13]. β-LAP has topoisomerase inhibitory activity and elevates NQO1 levels, leading to a futile redox cycle and apoptosis of cancer cells [11,14]. A recent study reports that β-LAP attenuates cisplatin-mediated nephrotoxicity by increasing NAD + levels with elevated tumoricidal effects of cisplatin [15]. Several studies have reported anti-inflammatory effects of β-LAP. β-LAP suppresses inflammatory responses in activated macrophages and protects from lung edema and high mortality in septic mice [16]. β-LAP alleviates carrageenan-induced rat paw edema by suppressing neutrophil migration and cytokine production [17]. In addition, β-LAP induces anti-inflammatory heme oxygenase-1 (HO-1) via AMPK activation in RAW264.7 macrophages and endothelial cells [18,19]. A previous study has shown anti-inflammatory effects of β-LAP in activated microglia [20]. It was demonstrated that β-LAP inhibits inducible nitric oxide synthase (iNOS) and cytokine expressions in lipopolysaccharide (LPS)-stimulated BV2 cells. However, the in vivo effects of β-LAP and the detailed molecular mechanism underlying the antiinflammatory effects of β-LAP have not been fully elucidated.
Therefore, in the present study, we examined the anti-inflammatory effects of β-LAP under both in vitro and in vivo neuroinflammatory conditions and analyzed, in detail, the molecular mechanism. In particular, we investigated the effects of β-LAP on the gene expression and activity of MMPs, because our group recently demonstrated the proinflammatory role of MMPs in activated microglia [21][22][23]. Through this study, we report for the first time that β-LAP inhibits microglial activation and expression of iNOS, cytokines, and several MMPs in the LPS-injected mouse brain. Furthermore, we demonstrated that multiple signaling pathways are involved in the anti-inflammatory mechanism of β-LAP in activated microglia.

Microglial cell cultures
The immortalized mouse BV2 microglial cell line [24] was grown and maintained in Dulbecco's modified Eagle's medium (DMEM), supplemented with 10 % heatinactivated fetal bovine serum, streptomycin (10 μg/ml), and penicillin (10 U/ml) at 37°C under 5 % CO 2 . Primary microglial cells were cultured from the cerebral cortices of 1-to 2-day-old Sprague Dawley rat pups as described previously [21]. The purity of microglial cultures was >95 %, as confirmed by Western blot and immunocytochemistry analyses using an antibody specific to ionized calcium-binding adapter protein-1 (IBA-1) staining (data not shown).

Measurement of cytokines, nitrite, and intracellular ROS levels
Cells (1 × 10 5 cells per well in a 48-well plate) were pretreated with β-LAP for 1 h and further stimulated with LPS (100 ng/ml) for 16 h. Concentrations of TNF-α, IL-1β, IL-6, and IL-10 in conditioned medium (CM) were measured by ELISA using monoclonal antibodies and procedures recommended by the supplier (PharMingen, San Diego, CA). Accumulated nitrite in CM and intracellular accumulation of reactive oxygen species (ROS) were measured using Griess reagent (Promega) and H 2 DCF-DA (Invitrogen, La Jolla, USA), respectively, as previously described [25].
Assays for MMP-3, MMP-8, and MMP-9 activity BV2 cells were stimulated with LPS in the presence or absence of β-LAP for 24 h, and the supernatants were collected to measure MMP activity using the SensoLyte® 520 MMP assay system (AnaSpec, San Jose, CA, USA). MMP activity measurements were performed by continuous detection of peptide cleavage using a fluorescence plate reader (Molecular Devices, Sunnyvale, CA, USA). MMP activity units were expressed as a change in the fluorescence intensity at an excitation wavelength of 490 nm and an emission wavelength of 520 nm.

LPS-induced inflammation and administration of β-LAP
C57BL/6 mice (10-11 weeks old) were purchased from the Orient Co., Ltd. (Seoul, Korea). All animal experiments were approved by the Institutional Animal Care and Use Committee at the School of Medicine, Ewha Womans University. All efforts were made to minimize animal suffering, to reduce the number of animals used, and to utilize alternatives to in vivo techniques, if available. Systemic inflammation was induced by LPS administration (5 mg/kg, i.p.) to male C57BL/6 mice as previously described [26]. β-LAP (10 mg/kg, i.p.), dissolved in vehicle solution (1 % DMSO and normal saline), was given daily for 4 days before the LPS treatment. Samples were obtained 3 or 6 h after LPS treatment.

Immunohistochemistry
Three hours after LPS treatment, the animals were anesthetized with sodium pentobarbital (120 mg/kg i.p.) and perfused transcardially with normal saline containing heparin (5 U/ml), followed by 4 % paraformaldehyde (PFA) in 0.1 M sodium phosphate buffer (PBS), pH 7.2. The brains were removed and incubated overnight in fixatives and stored in a 30 % sucrose solution. Serial coronal brain sections of regions containing the hippocampus (20 μm thick, at 600-μm intervals) were collected using a cryostat. Brain sections were incubated in PBS containing 0.1 % Triton X-100, 5 % normal serum, and 1 % bovine serum albumin for 1 h, and then subsequently incubated with primary antibody. On the next day, sections were incubated in a 1:200 dilution of Alexa Fluor 488-labeled donkey anti-rabbit secondary antibody or Alexa Fluor 594-labeled chicken anti-goat antibody (Molecular Probes Inc., Eugene, OR, USA) for 60 min at room temperature, and then washed with 0.05 % Tween 20 in PBS three times, 5 min each. Sections were then stained with a 0.5-μg/ml DAPI staining solution for 20 min at room temperature and washed. The sections were mounted with Vectashield mounting medium (Vector Laboratories, Burlingame, CA, USA), and fluorescence microcopy images were obtained using confocal microscopy (TSC-SP, Leica, Heidelberg, Germany). Iba1-, MMP-3-, MMP-8-, and MMP-9positive cells were quantified using the Metamorph program (Carl Zeiss, Jena, Germany). Two serial brain sections from each animal were used for further analysis, and quantification of Iba1-, MMP-3-, MMP-8-, and MMP-9-positive cells was performed in three different areas (500 μm 2 in size) in the lateral cortex and dentate gyrus of the right hemisphere per brain section. The mean cell number from six 500-μm 2 areas per animal was calculated.

RT-PCR
Total RNA (1 μg) isolated from BV2 or primary microglial cells (4.5 × 10 5 cells on a 6-cm dish), or from the brain tissue of LPS-injected mice, was reverse transcribed, and synthesized cDNA was used as a template for PCR. RT-PCR was performed on a T100 Thermal Cycler (Bio-Rad) with GoTaq polymerase (Promega). The primer sets shown in Table 1 were used to detect specific PCR products, and their values were calculated as fold change relative to control after normalization to the GAPDH gene.

Transient transfection and luciferase assay
BV2 cells plated at 50-60 % confluence (2 × 10 5 cells per well) in 12-well plates were transfected with 1 μg of plasmid DNA (ARE-luc, CRE-luc) using the Con-voy™ Platinum transfection reagent (CellTAGen, Seoul, Korea). After 36 h of transfection, cells were treated with β-LAP and LPS (100 ng/ml), or β-LAP only, for 6 h. The luciferase assay was used to determine the effect of β-LAP on ARE or CRE promoter activity. The ARE-luciferase reporter gene was kindly provided by Dr. Young-Joon Surh (Seoul National University, Seoul, Korea). The sequence of the antioxidant response element (ARE) construct is as follows: 5′-CTCAGCCTTCCAAATCG CAGTCACAGT GACTCAGCAGAATC-3′ [27,28]. The CRE-luc vector, which contains four copies of the cyclic AMP response element (CRE, TGACGTCA), was obtained from Stratagene (La Jolla, CA).

Statistical analysis
Unless otherwise stated, all experiments were performed with triplicate samples and repeated at least three times. Data are presented as mean ± SEM, and statistical comparisons among groups were performed using one-way ANOVA followed by Newman-Keuls post hoc tests or t tests. Statistical significance was accepted for P values <0.05.

β-LAP showed anti-inflammatory effects in LPS-stimulated microglial cells
To investigate the anti-inflammatory effect of β-LAP, BV2 or primary microglial cells were treated with LPS in the presence or absence of β-LAP for 16 h. Consecutively, NO and cytokine levels in CM were measured. We observed that β-LAP significantly inhibited the LPSinduced production of NO and proinflammatory cytokines such as TNF-α, IL-1β, and IL-6, whereas it raised anti-inflammatory IL-10 in BV2 cells and primary microglia (Fig. 1a, b). β-LAP did not have any cytotoxicity in the concentrations used in this study in both the BV2 and primary microglial cells, at least for 48 h (MTT assay, data not shown). Subsequent RT-PCR analysis showed that β-LAP reduced the mRNA expression of iNOS, TNF-α, IL-1β, and IL-6, and increased IL-10 ( Fig. 1c, d). The data suggest that β-LAP modulates the expressions of iNOS and cytokines at the transcriptional level.
β-LAP suppressed the expression and activity of MMP-3, MMP-8, and MMP-9, while it enhanced TIMP-2 expression in LPS-stimulated microglia Our group recently reported that MMP-3, MMP-8, and MMP-9 are important proinflammatory mediators in activated microglia, while TIMP-2 plays an antiinflammatory role [21][22][23]29]. To investigate whether β-LAP affects the expression of MMPs and TIMP-2, we performed RT-PCR analysis and Western blotting using cell lysates and CM from LPS + β-LAP-treated microglial cells. We found that β-LAP inhibited LPS-induced expressions of MMP-3, MMP-8, and MMP-9 at the mRNA and protein levels in both BV2 cells and primary microglia (Fig. 2a, b). Moreover, β-LAP reduced the enzymatic activity of MMP-3, MMP-8, and MMP-9 in the CM (Fig. 2d). On the other hand, β-LAP restored the expression of TIMP-2 ( Fig. 2a-c). The results suggest that the inhibition of MMPs and the upregulation of TIMP-2 are at least partly involved in the antiinflammatory effects of β-LAP.
β-LAP inhibited microglial activation and the expression of iNOS, cytokines, and MMPs in the brains of LPSinduced systemic inflammation mice To verify the anti-inflammatory effects of β-LAP in vivo, β-LAP (10 mg/kg) was injected before LPS administration. After 3 h, microglial activation and the expression levels of proinflammatory cytokines and MMPs were measured in the LPS-injected mouse brain. Systemic LPS led to an increase in densely stained Iba1-positive cells in the cortex and dentate gyrus (Fig. 3a). This indicates the presence of activated microglia. Pretreatment of β-LAP, however, reduced their number. The smaller amount of densely stained activated microglia was significant as seen in the subsequent quantification of Iba1-positive cells. However, we did not observe any infiltration of neutrophils or monocytes into the brain at this time point (i.e., 3 h) (data not shown). In addition, we found that β-LAP inhibited the mRNA expression of proinflammatory cytokines (IL-6, IL-lβ, TNF-α), iNOS, and MMPs (MMP-3, MMP-8, MMP-9) that were elevated in the cortex region of the LPS-injected mouse brain (Fig. 3b, c). To determine whether MMPs, upregulated in the cortex of LPS-injected mouse, were co-localized with microglia, immunohistochemistry was performed using antibodies against Iba1 (red) and MMP-3, MMP-8, or MMP-9 (green), 24 h after the LPS challenge. The number of double-immunopositive cells was significantly increased by LPS injection, but it was reduced in the β-LAP-injected groups (Fig. 4). The results indicate that β-LAP inhibits the expression of MMP-3, MMP-8, and MMP-9 in the microglia of the LPS-injected mouse brain.
β-LAP inhibited the phosphorylation of MAPKs and AKT and the DNA binding activity of NF-κB and AP-1 in LPSstimulated BV2 cells To further investigate the anti-inflammatory mechanism of β-LAP, we examined the effects of β-LAP on MAPKs and PI3K/AKT signaling, which play a critical role in the process of microglial activation and the production of NO and cytokines [30,31]. We found that β-LAP significantly inhibited the phosphorylation of three types of MAPKs and AKT in LPS-stimulated BV2 cells (Fig. 5a, b). Furthermore, we found that β-LAP suppressed the DNA binding activity of NF-κB and AP-1, which are key transcription factors modulating cytokine and iNOS gene expression in microglia (Fig. 5c, d).
β-LAP suppressed ROS production and expression of NADPH oxidase subunits, whereas it enhanced antioxidant enzyme HO-1/NQO1 expression Next, we examined the effects of β-LAP on LPS-induced ROS production, which acts as a second messenger in inflammatory reactions and subsequent neuronal cell death [32,33]. β-LAP significantly inhibited LPSinduced ROS production in the BV2 cells and primary microglia (Fig. 6a-c). Since NADPH oxidase is a major enzyme for microglial ROS release, we examined the effect of β-LAP on membrane (gp91 phox , p22 phox ) and cytosolic (p47 phox , p67 phox ) components of NADPH oxidase. β-LAP suppressed the expression and phosphorylation of p47 phox (Fig. 6d, e). Moreover, β-LAP inhibited the mRNA expression of gp91 phox , without affecting p67 phox or p22 phox (Fig. 6d). We next examined the effects of β-LAP on HO-1 and NQO1, which mediate anti-inflammatory and anti-oxidant effects in the activated microglia [33,34]. We observed that β-LAP increased LPS-induced HO-1 and NQO1 expression at mRNA and protein levels (Fig. 7a, b). Interestingly, we found that β-LAP itself also increased HO-1 and NQO1 expression.

β-LAP activated Nrf2/ARE and PKA/CREB pathways
Because Nrf2 binds to ARE on the promoters of phase II anti-oxidant enzyme genes such as HO-1 and NQO1, and controls their expression [27,34], we examined the A B C D Fig. 1 Effects of β-LAP on iNOS and pro-/anti-inflammatory cytokines in LPS-stimulated BV2 cells and primary cultured microglia. BV2 cells (a) or primary cultured microglia (b) were pretreated with β-LAP (0.5, 1, and 2 μM) for 1 h and incubated with LPS (100 ng/ml for BV2, 10 ng/ml for primary microglia). After incubation for 16 h, the conditioned media were collected, and the amounts of NO, TNF-α, IL-1β, IL-6, and IL-10 were measured using Griess reagent or ELISA. The data are the mean ± SEM of three independent experiments. *P < 0.05, significantly different from LPS-treated samples. BV2 cells (c) or primary microglia (d) were pretreated with β-LAP (0.5, 1, and 2 μM) for 1 h, followed by LPS (100 ng/ml) for 6 h, and total RNA was isolated. The mRNA expressions of iNOS and cytokines were analyzed by RT-PCR. Representative gels are shown on the left panel, and quantifications of three independent experiments are shown in the right panel. Values correspond to the mean ± SEM of three independent experiments. *P < 0.05, significantly different from LPS-treated samples effects of β-LAP on the binding of Nrf2 to ARE on HO-1/NQO1 promoters. We observed that LPS induced Nrf2 binding to ARE, which was enhanced by β-LAP (Fig. 7c). Moreover, β-LAP increased ARE-driven luciferase activity in the absence and presence of LPS (Fig. 7d). Previous studies of our own and by other groups reported that the PKA/CREB pathway contributes to the resolution of inflammation and ROS detoxification, and that PKA is an upstream modulator of HO-1 expression in microglia [35,36]. In the present study, we observed that β-LAP itself increased the phosphorylation of CREB, which is a downstream target of PKA. In addition, β-LAP potentiated the CREB phosphorylation induced by LPS (Fig. 7e). β-LAP also increased the DNA binding and transcriptional activity of CREB in the absence or presence of LPS (Fig. 7f, g). Thus, the data suggest that the Nrf2/ ARE and PKA pathways are largely involved in the antiinflammatory/anti-oxidant mechanism of β-LAP, in

Discussion
Our present study demonstrates the anti-inflammatory properties of β-LAP in brain microglia and their underlying molecular mechanisms. β-LAP inhibited the expressions of iNOS and proinflammatory cytokines in LPS-stimulated microglia. In addition, β-LAP reduced the expression and activity of MMP-3, MMP-8, and MMP-9, which are inflammatory mediators in activated microglia [21][22][23]. By using a systemic inflammation mouse model, we confirmed the anti-inflammatory role of β-LAP. Thus, β-LAP inhibited microglial activation and the expression of proinflammatory molecules in the LPS-injected mouse brain. By mechanistic analysis, we showed that β-LAP inhibited the phosphorylation of MAPKs and AKT and the DNA binding activity of NF-κB/AP-1 induced by LPS. Furthermore, we found that β-LAP exerts anti-oxidant effects by reducing ROS production via suppression of NADPH oxidase subunit activity and/or expression, and upregulation of antioxidant enzymes such as HO-1 and NQO1. We showed that β-LAP activated Nrf2/ARE and PKA/CREB pathways, which are involved in the upregulation of HO-1/ NQO1 expression. Therefore, β-LAP appears to act as an anti-inflammatory/anti-oxidant agent by modulating multiple signaling pathways (i.e., inhibition of MAPKs and PI3K/AKT, upregulation of Nrf2/ARE and PKA). A previous study reported that β-LAP inhibits the mRNA expression of TLR4 signaling molecules in experimental autoimmune encephalomyelitis mice [37]. In the present study, however, β-LAP did not affect the mRNA expression of TLR4 or MyD88 in LPS-stimulated microglia (Additional file 1: Figure S1), suggesting that β-LAP exerts anti-inflammatory effects by modulating signaling downstream of TLR4/MyD88 and/or via a MyD88-independent pathway. In addition, we found that β-LAP also inhibited the inflammatory reactions induced by TLR2 or TLR3 agonists. Treatment with β-LAP inhibited NO and TNF-α production in BV2 cells stimulated with lipoteichoic acid (LTA; TLR2 agonist) or polyinosinic-polycytidylic acid (Poly I:C; TLR3 agonist) (Additional file 1: Figure S2). Thus, the data imply that the anti-inflammatory effect of β-LAP is not confined to TLR4 activation. Further studies are necessary to investigate the detailed mechanism underlying the effect of β-LAP on TLR signaling.
MMPs are zinc-dependent endopeptidases which are involved not only in normal brain development but also in various neuropathological conditions such as Alzheimer's disease, Parkinson's disease, stroke, and multiple sclerosis [38]. MMPs are aberrantly expressed in neuropathological conditions and cause breakdown of the blood-brain barrier (BBB), infiltration of peripheral immune cells, demyelination, and neuronal cell death [39,40]. Our group recently reported that MMPs play an important role in various neuroinflammatory conditions [21][22][23]. We showed that MMP-3, MMP-8, and MMP-9 are upregulated in LPS or α-synuclein-stimulated microglia and mediate neuroinflammatory reactions. Thus, the specific inhibition of MMP-3, MMP-8, or MMP-9 suppresses iNOS and cytokine expressions in LPS or α-synucleinstimulated microglia. We demonstrated that MMPs cleave the N-terminal extracellular domain of protease-activated receptor-1 and activate intracellular inflammatory signaling pathways in α-synuclein-activated microglia [21]. More recently, we showed that MMP-8 plays a pivotal role in neuroinflammation by activating TNF-α processing in microglia [22]. We also reported that TIMP-2, as an endogenous inhibitor of MMPs, has an anti-inflammatory effect by modulating MMP-3, MMP-8, and MMP-9 in activated microglia [29]. Based on these findings, our present study examined the effects of β-LAP on MMPs and TIMP-2 in LPS-stimulated microglia. We observed that β-LAP significantly suppressed the expression and activity of MMP-3, MMP-8, and MMP-9 with enhancement of TIMP-2 under in vitro and/or in vivo neuroinflammatory conditions, which may contribute to the antiinflammatory properties of β-LAP.
In this study, we found that β-LAP induced phase II anti-oxidant enzymes such as HO-1 and NQO1, which are regulated through the Nrf2/ARE signaling pathway [34]. Under normal conditions, Nrf2 is sequestered by cytosolic Keap1, which serves as an adaptor to link Nrf2 to the ubiquitin ligase Cul3-Rbx1 complex that ubiquitinates and degrades Nrf2. However, upon stimulation by electrophilic agents or ROS, Nrf2 dissociates from its Results are representative RT-PCR data in the cortex 3 h after LPS treatment. Values are expressed as the means ± SEM for three independent experiments. *P < 0.05 vs. saline or # P < 0.005 vs. LPS-treated mice cytosolic docking protein Keap1, translocates into the nucleus, and binds to the ARE site [41]. It has been suggested that Nrf2 phosphorylation is involved in this release process. In the present study, we observed that β-LAP increased Nrf2 binding to ARE, as well as AREmediated transcriptional activity. However, we did not further examine the effect of β-LAP on the Nrf2 release mechanism related to Keap1; this would be an interesting study in the future.
It is well-known that β-LAP is a substrate and activator of NQO1, which catalyzes the oxidation of NADH to NAD + . A recent study reported that β-LAP induces HO-1 expression by increasing NQO1 activity and AMPK phosphorylation in RAW264.7 macrophages [18,19]. As a mechanism to resolve the neurotoxic responses brought about by microglial activation, microglia usually express anti-inflammatory cytokines (i.e., IL-10, TGF-β1), suppressor of cytokine signaling (SOCS)-family proteins (i.e., SOCS1 and SOCS3), and anti-oxidant enzymes (i.e., HO-1, NQO1, SOD) [42,43]. In the present study, we found that LPS itself increased HO-1 expression, which is associated with AREdependent transcription. We previously reported that the activation of PKA/CREB signaling is upstream of HO-1 expression and that the upregulation of the HO-1 and PKA pathway plays a key role in mediating the antiinflammatory mechanism in LPS-stimulated microglia [35]. Therefore, the potentiation of HO-1/ARE and PKA/ CREB by β-LAP may at least partly contribute to the anti-inflammatory and anti-oxidant effects of β-LAP in LPS-stimulated microglia. Interestingly, a recent study demonstrated that HO-1 knockout markedly increases MMP-9 expression in arteriovenous fistulas in mice and that MMP-9 induction reflects the prooxidant and  Cell extracts were prepared from BV2 cells pretreated with β-LAP (0.5, 1, and 2 μM, for 1 h), followed by LPS (100 ng/ml, for 1 h), and then subjected to immunoblot analysis using antibodies against the phospho-or total forms of JNK, ERK, p38 MAPK, and Akt. The autoradiograms are representative of three independent experiments. b Quantification of Western blot data. Levels of the phosphorylated forms of MAPKs and AKT were normalized with respect to the level of each total form and expressed as relative fold changes vs. the control group. Data are the means ± SEM for three independent experiments. *P < 0.05, significantly different from the LPS-treated samples. c, d EMSA for NF-κB and AP-1 DNA binding activity. BV2 cells were pretreated with β-LAP (0.5, 1, and 2 μM, for 1 h), followed by LPS (100 ng/ml, for 3 h), and nuclear extracts prepared from BV2 cells were incubated with the NF-κB (c) or AP-1 (d) probes. The arrow indicates a DNA-protein complex of NF-κB or AP-1. F indicates a free probe proinflammatory effects recognized in a state of HO-1 deficiency [44]. Therefore, the upregulation of HO-1/NQO1 may at least partly contribute to the anti-inflammatory effects of β-LAP by suppressing proinflammatory molecules such as cytokines and MMPs in LPS-stimulated microglia.
A number of studies have reported therapeutic and health benefits of β-LAP supplementation in experimental animal models and clinical trials. The most extensively studied property of β-LAP is its anti-cancer potential, and β-LAP is currently being evaluated in clinical trials for treatment of cancer [9,11]. In addition, β-LAP has beneficial effects on metabolic syndromes such as obesity, diabetes, hypertension, arterial restenosis, and salt-induced renal injury [45][46][47][48]. The anti-inflammatory/ cytoprotective effects of β-LAP have also been reported in several disease models. β-LAP attenuates cisplatinmediated acute kidney injury in mice by suppressing critical mediators for inflammation and ROS [15]. β-LAP has shown therapeutic effects against rheumatoid arthritis by inhibiting synoviocyte proliferation and suppressing MMP expression in chondrocytes [49]. β-LAP also protects against renal ischemia/reperfusion injury in mice by inducing NQO1 activation and subsequent inhibition of ROS [50]. In experimental autoimmune encephalomyelitis, an animal model of multiple sclerosis, the administration of β-LAP ameliorates the development of EAE by inhibiting the production of IL-12 family cytokines [37]. Interestingly, a recent study demonstrated that potentiation of NQO1 activity by feeding β-LAP prevents the agedependent decline of motor and cognitive function in Fig. 6 β-LAP inhibited ROS production via suppression of NADPH oxidase subunits. a BV2 cells or b primary microglia were pretreated with β-LAP (0.5, 1, and 2 μM, for 1 h), followed by LPS (100 or 10 ng/ml, for 16 h), and stained with 50 μM H 2 DCF-DA. DCF fluorescence intensities were measured using a microplate fluorometer. The data are expressed as the means ± SEM of three independent experiments. *P < 0.05, significantly different from the LPS-treated group. c A representative confocal image of DCF-derived fluorescence (green) in BV2 cells (n = 3), with an original magnification of ×200. d RT-PCR analysis for NADPH oxidase subunits (p47phox, p67phox, gp91phox, p22phox). BV2 cells were pretreated with β-LAP (0.5, 1, and 2 μM, for 1 h) followed by LPS (100 ng/ml), and total RNA was isolated at 2 h after LPS treatment. Representative gels are shown in the left panel, and quantification data are shown in the right panel (n = 3). e Western blot analysis for phosphorylation of the p47phox subunit (n = 3). BV2 cells were pretreated with β-LAP (0.5, 1, and 2 μM, for 1 h), followed by LPS (100 ng/ml, for 30 min), and then subjected to immunoblot analysis using antibodies against phospho-p47phox. Quantification data are shown in the graph. *P < 0.05, significantly different from the LPS-treated group aged mice [51]. β-LAP also increases memory performance and prevents the loss of synapses in aged mice, suggesting the therapeutic potential of β-LAP for neurodegenerative diseases.
As to the BBB permeability of β-LAP, Huntingdon Life Sciences (UK) has reported that a minimal concentration of β-LAP penetrates into the rat brain, compared with other organs, under normal conditions (unpublished Fig. 7 β-LAP increased HO-1 and NQO1 expression via upregulation of Nrf2/ARE and PKA/CREB pathways in LPS-stimulated microglia. a Western blot analysis shows the effects of β-LAP on HO-1 and NQO1 protein expression. Cell lysates were obtained from BV2 cells treated with β-LAP (0.5, 1, and 2 μM) with or without LPS (100 ng/ml) for HO-1 or NQO1 (6-16 h). b RT-PCR was performed to determine the HO-1 and NQO1 mRNA expression. Cells were treated with β-LAP (0.5, 1, and 2 μM) for 1 h prior to treatment with LPS (100 ng/ml, for 6 h) and analyzed. Quantification data are shown in the graph (n = 3). c EMSA for Nrf2. Nuclear extracts were prepared from BV2 cells treated with LPS (100 ng/ml, for 3 h) or LPS + β-LAP (1 and 2 μM, pretreatment for 1 h) and incubated with the ARE probe. The arrow indicates a DNA-protein complex of Nrf2. d Effect of β-LAP on ARE-luc reporter gene activity. Cells transfected with the reporter plasmid (ARE-luc) were treated with β-LAP (0.5, 1, and 2 μM) with or without LPS (100 ng/ml) for 6 h, and the reporter gene assay was performed. e Effect of β-LAP on the phosphorylation of CREB. Cell lysates were obtained from BV2 cells treated with β-LAP (0.5, 1, and 2 μM) with or without LPS (100 ng/ml) for 1 h. Quantification data are shown in the graph (n = 3). f EMSA for CREB. Nuclear extracts were prepared from BV2 cells treated with LPS (100 ng/ml, for 3 h) or LPS + β-LAP (1 and 2 μM, pretreatment for 1 h) and incubated with the CRE probe. The bracket indicates a DNA-protein complex of CREB. g Effect of β-LAP on CRE-luc activity. Cells transfected with the reporter plasmid (CRE-luc) were treated with β-LAP (0.5, 1, and 2 μM), with or without LPS (100 ng/ml) for 6 h, and the reporter gene assay was performed. Data are the means ± SEM of three independent experiments. *P < 0.05 vs. control or # P < 0.05 vs. the LPS-treated group


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Inhibitory Mechanism of FAT4 Gene Expression in Response to Actin Dynamics during Src-Induced Carcinogenesis

Oncogenic transformation is characterized by morphological changes resulting from alterations in actin dynamics and adhesive activities. Emerging evidence suggests that the protocadherin FAT4 acts as a tumor suppressor in humans, and reduced FAT4 gene expression has been reported in breast and lung cancers and melanoma. However, the mechanism controlling FAT4 gene expression is poorly understood. In this study, we show that transient activation of the Src oncoprotein represses FAT4 mRNA expression through actin depolymerization in the immortalized normal human mammary epithelial cell line MCF-10A. Src activation causes actin depolymerization via the MEK/Erk/Cofilin cascade. The MEK inhibitor U0126 blocks the inhibitory effect of Src on FAT4 mRNA expression and Src-induced actin depolymerization. To determine whether actin dynamics act on the regulation of FAT4 mRNA expression, we treated MCF-10A cells with the ROCK inhibitor Y-27632. Y-27632 treatment decreased FAT4 mRNA expression. This suppressive effect was blocked by siRNA-mediated knockdown of Cofilin1. Furthermore, simultaneous administration of Latrunculin A (an actin depolymerizing agent), Y-27632, and Cofilin1 siRNA to the cells resulted in a marked reduction of FAT4 mRNA expression. Intriguingly, we also found that FAT4 mRNA expression was reduced under both low cell density and low stiffness conditions, which suggests that mechanotransduction affects FAT4 mRNA expression. Additionally, we show that siRNA-mediated FAT4 knockdown induced the activity of the Hippo effector YAP/TAZ in MCF-10A cells. Taken together, our results reveal a novel inhibitory mechanism of FAT4 gene expression through actin depolymerization during Src-induced carcinogenesis in human breast cells.

Introduction
Oncogenic cell transformation results from the summation of changes in cell growth, cell viability, cell motility and cell morphology. The v-Src oncogene, a product of the Rous sarcoma virus, is the constitutively active form of c-Src. Src has the ability to regulate various signal transduction pathways, including the Ras/MEK/Erk, PI3K/Akt, STAT3, and Rho/ROCK pathways [1][2][3][4]. More specifically, Src has been reported to induce alterations in cell morphology through actin dynamics and to depolymerize the actin cytoskeleton via the MEK/Erk/Cofilin cascade [5]. Additionally, members of the cadherin superfamily have been implicated in Srcinduced tumor transformation. Src downregulates E-cadherin expression and triggers morphological changes in multiple cancers [6][7][8]. These findings suggest the importance of both actin dynamics and the loss of cadherin-mediated cell-cell adhesion in Src-induced tumorigenesis.
FAT4, a protocadherin, is the human ortholog of Drosophila Fat [9,10]. Recent studies indicate that Drosophila Fat suppresses tumorigenesis through activation of the Hippo pathway. In support of this finding, PrognoScan, a new microarray database [11], and other recent studies [12][13][14] have demonstrated that human FAT4 gene expression is repressed in breast and lung cancers and in melanoma, which suggests that reduced FAT4 gene expression can trigger carcinogenesis. However, the molecular mechanisms underlying the down-regulation of FAT4 gene expression in human cancers remain unknown.
The Hippo pathway is involved in tumor suppressor signaling and regulates organ size, cell proliferation, apoptosis and stemness [9,10,15,16]. In mammals, the core components of the Hippo pathway are primarily composed of Mst1/2 and Lats1/2, which inactivate the Hippo effector Yap/Taz via phosphorylation [17][18][19][20][21][22]. Yap/Taz acts as a transcriptional co-activator by binding the transcription factors Tead1-4 to induce the expression of genes such as the connective tissue growth factor CTGF [18,23,24]. Fat4 has been shown to repress Yap activity and thereby negatively regulates neural progenitor cell proliferation during mammalian neurogenesis [25,26]. Thus, the FAT4-Hippo axis is believed to be functionally conserved in humans; however, the functional significance and the molecular mechanisms that confer this axis are poorly understood.
Here, we show that transient Src activation represses FAT4 mRNA expression and leads to tumor transformation in MCF-10A cells, which are immortalized normal human mammary epithelial cells. We further demonstrate that the inhibitory effect of Src on FAT4 mRNA expression depends on actin depolymerization that is induced by the MEK/Erk/Cofilin cascade. Finally, we demonstrate that FAT4 represses YAP/TAZ activity in MCF-10A cells.

FAT4 gene expression is repressed during human breast cell transformation by the Src oncogene
To analyze the mechanisms that repress FAT4 gene expression in human breast cancer, we generated a model of carcinogenesis using MCF-10A cells, an immortalized normal human mammary epithelial cell line [27][28][29][30][31]. In brief, we used retroviruses to create MCF-10A cells that stably express the Src kinase oncoprotein (v-Src) fused to the ligand binding domain of the estrogen receptor (ER). In MCF-10A v-Src:ER cells, the addition of 4-Hydroxytamoxifen (TAM) induces Src activation and their subsequent cell transformation. TAM treatment resulted in morphological transformation (Fig. 1A), enhanced cell proliferation (Fig. 1B), and promoted anchorage-independent growth (Fig. 1C). In this model, we found that FAT4 mRNA expression was repressed by Src activation in MCF-10A cells (Fig. 1D). On the other hand, TWIST1 mRNA expression was not altered under similar experimental conditions (S1 Fig.). This points to the specificity of the inhibitory effect of Src on FAT4 mRNA expression. Thus, these results indicate that Src induces tumorigenesis and suppresses FAT4 mRNA expression in MCF-10A cells.
Src activation induces both FAT4 gene repression and Cofilin-mediated actin depolymerization through the MEK/Erk pathway Src has been shown to elicit Cofilin-mediated disruption of the actin cytoskeleton through the MEK/Erk pathway [5]. The function of Cofilin is activated via its dephosphorylation [32]. Thus, we examined whether Src can disrupt the actin cytoskeleton through dephosphorylation of Cofilin in MCF-10A v-Src:ER cells. Indeed, phalloidin staining and Western blot analysis demonstrated that Src can depolymerize the actin cytoskeleton and dephosphorylate Cofilin (Fig. 2). We further examined the involvement of the MEK/Erk pathway in Src-mediated modulation of the actin cytoskeleton. Treatment of MCF-10A v-Src:ER cells with the MEK inhibitor U0126 completely inhibited Src-mediated phosphorylation of Erk1/2, a major target of MEK1/2 (Fig. 3A). Under this condition, U0126 significantly inhibited both Src-induced actin dynamics (Fig. 3B) and the dephosphorylation of Cofilin (Fig. 3C). Therefore, these results indicate that Src induces Cofilin-mediated actin depolymerization through the MEK/Erk pathway in MCF-10A cells.
We also investigated whether the MEK/Erk pathway is involved in Src-reduced FAT4 mRNA expression by RT-qPCR analysis; the results suggest that Src-mediated inhibition of FAT4 mRNA expression is repressed in MCF-10A cells by U0126 or PD0325901, a MEK inhibitor ( Fig. 3D and S2 Fig.). In addition, to emphasize the importance of the MEK/Erk pathway in FAT4 mRNA repression we evaluated the effect of U0126 on FAT4 mRNA expression in MDA-MB-231 cells, a malignant breast cancer cell line. Treatment of MDA-MB-231 cells with U0126 also up-regulated FAT4 mRNA expression (Fig. 3E). Thus, these results indicate that Src represses FAT4 mRNA expression through the MEK/Erk pathway. Intriguingly, siRNA-    (Fig. 3F), perhaps because the MEK/Erk pathway regulates various targets that include FAT4 gene. Collectively, these results suggest that Src activation induces not only Cofilinmediated actin depolymerization but also FAT4 mRNA repression through the MEK/ Erk pathway.

Src represses FAT4 gene expression through Cofilin1-mediated actin depolymerization
Next, we examined the causal relationship between FAT4 mRNA repression and Cofilinmediated actin depolymerization, both of which are downstream of the Src/MEK/Erk pathway. The ROCK pathway is a major pathway that regulates Cofilin-mediated actin depolymerization [33,34]. Thus, to determine whether Src downregulates FAT4 mRNA expression by altering actin structures, we examined the effect of the ROCK inhibitor Y-27632 on FAT4 mRNA expression. Treatment of MCF-10A cells with Y-27632 reduced FAT4 mRNA expression in a dose-dependent manner (Fig. 4A). The inhibitory effect of Y-27632 on FAT4 mRNA expression was abrogated by siRNA-mediated knockdown of Cofilin1 (Fig. 4B). Cofilin1 is a nonmuscle type of Cofilin [32]. Effective knockdown of Cofilin1 was supported by RT-qPCR ( Fig. 4B) and Western blot analysis (Fig. 4C). Moreover, simultaneous administration of the actin depolymerizing agent Latrunculin A, Y-27632, and siCofilin1 to the cells resulted in a striking decrease of FAT4 mRNA expression equal to that of the Y-27632 treatment alone ( Fig. 4B and S3 Fig.). Phalloidin staining showed that Latrunculin A sufficiently depolymerizes the actin cytoskeleton (Fig. 4D). Therefore, on one hand, these results suggest that ROCK1/2 regulates FAT4 mRNA expression via Cofilin1-mediated actin depolymerization. On the other hand, they suggest that inhibition of Cofilin1 significantly attenuates FAT4 mRNA repression by Src (Fig. 4E). Therefore, we propose that Src suppresses FAT4 mRNA expression through actin depolymerization by Cofilin1.

Mechanotransduction affects FAT4 gene expression
Recent studies have reported that mechanical stress on cells affects cellular behavior via the modulation of actin dynamics [35][36][37]. Based on our findings that FAT4 mRNA expression is regulated by actin depolmerization through Cofilin1 (Fig. 4), we examined whether mechanical stress also affects FAT4 mRNA expression. To test this hypothesis, we performed a Cell Density Assay and a Softwell Assay. In the Cell Density Assay, MCF-10A cells were seeded at high and low cell densities to evaluate the effect of cell-cell contact on FAT4 mRNA expression. The Softwell Assay used collagen type I-coated Softwell plates of varying degrees of stiffness (50 kPa and 0.5 kPa) to assess the effect of anchorage stiffness on FAT4 mRNA expression. FAT4 mRNA expression was reduced under both low cell density (Fig. 5A) and low anchorage stiffness conditions (Fig. 5B). These findings suggest that the mechanical stress that stems from the extracellular environment is intimately linked with FAT4 mRNA expression. Moreover, we found no significant difference in phoshorylated-Cofilin expression between high and low density cells (S4 Fig.). Thus, it may be assumed that the observed alteration in FAT4 mRNA expression under low density conditions is not mediated through the MEK/ERK/Cofilin cascade.

FAT4 suppresses the activity of the Hippo effector YAP/TAZ
Accumulating evidence suggests that the FAT4-Hippo axis is also conserved in humans [12][13][14]25,26]. We next examined the effects of FAT4 knockdown on the function of the Hippo effector YAP/TAZ. YAP/TAZ activity is suppressed by cytoplasmic retention and proteasomal degradation due to LATS1/2-mediated phosphorylation of YAP/TAZ [38][39][40][41][42]. FAT4 siRNA increased mRNA expression levels of CTGF, a YAP/TAZ target gene (Fig. 6A) [18][19][20][21]23,24], by promoting the nuclear accumulation of YAP/TAZ (Fig. 6B). Furthermore, we found that the oncoprotein Src promoted YAP/TAZ activity in MCF-10A cells (Figs. 6C and 6D). These results are consistent with those of a recent study that demonstrated Drosophila Src64B activates Yorkie, the mammalian Yap/Taz homologue, by repressing the core components of the Hippo pathway (Hippo/Warts) [43]. Thus, our results suggest that FAT4 represses CTGF mRNA expression through the inactivation of YAP/TAZ. We next examined the molecular basis of YAP/TAZ activation by FAT4 knockdown. Because YAP/TAZ activity is regulated by mechanotransduction via actin dynamics [35,36,38], we investigated the effects of FAT4 siRNA on actin dynamics. Phalloidin staining revealed that FAT4 knockdown induced abnormal spiny actin structures that radiated in all directions from the cells (Fig. 6E and S5 Fig.). However, this structural modulation of actin was independent of Cofilin because we observed no increase in phosphorylated Cofilin (Fig. 6F). Taken together, our results may suggest that FAT4 represses YAP/TAZ activity by regulating actin structures.

FAT4 knockdown does not induce cell transformation
Finally, we investigated whether FAT4 knockdown alone can transform MCF-10A cells because FAT4 knockdown activates YAP/TAZ, which elicits cell transformation (Figs. 6A and 6B) [23,24]. However, FAT4 knockdown alone did not cause increased cell proliferation nor anchorage-independent growth in MCF-10A cells (S6 Fig.). Therefore, combinatorial effects of reduced FAT4 gene expression and other factors could be necessary for tumor transformation in MCF-10A cells.

Discussion
Reduced FAT4 gene expression in human breast cancers has been reported in several studies [11,12]. Nevertheless, the molecular mechanism that governs this down-regulation in gene expression remains unknown. Here, we show that FAT4 mRNA expression is repressed in response to actin dynamics during Src-induced tumor transformation of MCF-10A cells (Fig. 7). Src-mediated actin depolymerization is regulated by the MEK/Erk/Cofilin1 cascade and results in the repression of FAT4 mRNA expression. Finally, we demonstrate that reduced expression of FAT4 induces the activity of the Hippo effector YAP/TAZ in MCF-10A cells. Altogether, these results propose a novel mechanism by which actin dynamics regulate cancer-related traits through the loss of FAT4-mediated cell-cell adhesion. In this study, we demonstrate that the Src/MEK/Erk/Cofilin1 cascade, which mediates actin depolymerization, represses FAT4 gene expression in MCF-10A cells (Fig. 7). The mechanism by which the MEK/Erk pathway represses FAT4 gene expression is conserved in even the malignant breast cancer cell line MDA-MB-231 cells as well as the normal breast cell line MCF-10A cells (Fig. 3D and 3E). Src activates Cofilin via dephosphorylation by components of the MEK/Erk pathway (Fig. 3C) and thereby induces actin depolymerization (Fig. 3B). Notably, the MEK/Erk pathway plays an important role in activating Cofilin in a variety of human cancer cell lines [5,[44][45][46][47]. Moreover, simultaneous administration of the ROCK inhibitor Y-27632 and siCofilin1 lead to restore FAT4 mRNA expression repressed by Y-27632 in the cells (Fig. 4B). Thus, these results suggest that Src-Cofilin1 axis plays an important role in the regulation of FAT4 mRNA expression in breast cancer cells. Unexpectedly, the siRNAmediated Cofilin knockdown did not completely block the reduction in FAT4 mRNA expression by Src, although it abolished the inhibitory effect of Y-27632 on FAT4 mRNA expression ( Fig. 4B and 4E). Generally, Src regulates a wider array of signaling pathways than ROCK. As such, this may result in the significant differences presented in these data. Taken together, these results clearly show the inhibitory effect of Src on FAT4 mRNA expression in response to actin dynamics during carcinogenesis.
The molecular mechanisms by which Src-induced actin depolymerization represses FAT4 mRNA expression have not yet been elucidated. The FAT4 gene is silenced by promoter hypermethylation in breast cancer cells [12]. Interestingly, Src elicits the overexpression of DNA methyltransferase (DNMT) 1 and induces DNA hypermethylation in rat 3Y1 fibroblasts [48]. Additionally, the MEK/Erk pathway induces DNA hypermethylation via the regulation of DNMT1 [49][50][51]. Therefore, Src-induced actin depolymerization has been speculated to repress FAT4 mRNA expression via DNMT1-induced promoter hypermethylation during carcinogenesis, but further studies are needed for clarification.
Our study demonstrates that siRNA-mediated knockdown of FAT4 induces YAP/TAZ activity in MCF-10A cells (Fig. 6A and 6B). In line with this view, recent studies utilizing in vivo electroporation have demonstrated that Fat4 downregulates Yap activity leading to the inhibition of neural progenitor cell proliferation during murine and avian neurogenesis [25,26]. In addition, it has been demonstrated that deletion of Fat4 in mice causes primarily the nuclear accumulation of Yap in the nephron progenitor cells [52]. As such, these in vivo findings suggest that the Fat4-Hippo axis is highly conserved in mammals. However, the molecular underpinnings of Fat4-mediated nuclear translocation of Yap/Taz in mammals still remain unknown. Because there is no mammalian counterpart of Drosophila Dachs, which is regulated by Fat to activate the Hippo pathway [9,10,18], Fat4 should modulate Yap/Taz activity differently from Fat. In this regard, our research demonstrates that YAP phosphorylation levels (at least at Ser127) in FAT4 siRNA-treated cells are consistent with those found in control siRNAtreated MCF-10A cells (S7 Fig.). Nevertheless, MST1 kinase (mammalian Hippo ortholog) expression is reduced in FAT4 siRNA-treated cells (S7 Fig.). Accordingly, it is possible that nuclear localization of YAP is regulated by FAT4 through a Ser127 phosphorylationindependent mechanism.
It is also known that FAT4 is involved in the maintenance of planar cell polarity (PCP) [9]. Indeed, Fat4 mutant mice and PCP gene mutant mice show similar defects of the inner ear such as significant disruptions in orientation of hair cells and elongation of the cochlea [53]. Moreover, PCP genes have also been shown to regulate actin dynamics [54]. Collectively, these findings suggest that Fat4 regulates actin dynamics as well as vertebrate PCP. Interestingly, YAP/TAZ activity can be modulated by cytoskeletal tension induced by cell spreading in a Hippo pathway-independent manner [10,21,35]. Therefore, we can surmise that FAT4 regulates the YAP/TAZ activity through modulating actin cytoskeleton. In fact, we show that FAT4 knockdown induces abnormal spiny actin protrusions, which may enhance cell spreading (Fig. 6E). Thus, FAT4 may negatively regulate YAP/TAZ function by modulating actin cytoskeletal tension. Altogether, our current study presents a novel model in which FAT4 represses YAP/TAZ activity by regulating actin structures, at least in breast cancer cells. Further examination is needed to elucidate the molecular basis of YAP/TAZ inactivation by FAT4.
Accumulating evidences suggest the pathophysiological relevance between FAT4 reduction and carcinogenesis. Qi et al. have clearly demonstrated that FAT4 mRNA is not expressed at all in many breast cancer cell lines (BT20, ZR75-1, and BT474 cells) [12]. We also found that MCF-7 breast cancer cells express far too low a level of FAT4 mRNA compared with the MCF-10A normal mammary cell line (data not shown). Moreover, FAT4 knockdown has been shown to induce the malignant phenotype of human gastric cancer cell lines [55]. However, we show that FAT4 knockdown alone is not sufficient to induce cell proliferation or anchorage-independent growth in MCF-10A cells despite promoting YAP/TAZ activity (S6 Fig.; Fig. 6A and 6B); nevertheless, overexpression of YAP/TAZ can transform the cells [23,24]. Additionally, FAT4 knockdown alone does not induce cell proliferation in malignant breast cancer MDA-MB-231 cells (Fig. 3F). These data strongly demonstrate that cooperative effects between FAT4 reduction and other factors induce tumor transformation in MCF-10A cells. We propose that the MEK/Erk pathway involves in the Src-induced tumorigenesis as well as FAT4 mRNA repression (Fig. 3A-D), because numerous studies have already established the importance of the MEK/Erk pathway for cell transformation [2,5,44,56,57]. Further comprehensive analyses are required to solve molecular mechanisms of Src-mediated carcinogenesis through repressing FAT4 gene expression.

siRNA knockdown experiments
Cells were cultured in Growth Medium without antibiotics at 37°C for 24 h and were then transfected with siRNA using both Lipofectamine RNAiMAX (Invitrogen) and Opti-MEM I Reduced Serum Medium (31985-070; Gibco) according to the manufacturer's protocol. The sequences of the siRNAs employed in this study are listed in S1 Table. The siRNA concentrations are described in the Figure Legends.

RNA extraction and real-time quantitative PCR (RT-qPCR)
Total RNA was extracted from cells with ISOGENII (311-07361; Wako) or the RNeasy Mini Kit (Qiagen) and was subjected to cDNA synthesis with random hexamer primers and Moloney murine leukemia virus (M-MLV) reverse transcriptase (Invitrogen) according to the manufacturer's protocols. RT-qPCR was performed with FastStart Universal SYBR (Roche) and a Thermal Cycler Dice Real Time System II (TP900; Takara Bio). The PCR primers that were used are listed in S2 Table. 18S rRNA was used for normalization.

Fluorescent staining
The cells were fixed with 4% formaldehyde solution in PBS on ice for 15 min, washed 3 times, and were permeabilized with 0.5% Triton X-100 in PBS for 10 min at room temperature (RT). After blocking cells with PBS containing 1% skim milk for 20 min at RT, the cells were incubated overnight at 4°C with anti-YAP antibody diluted in PBS containing 1% skim milk. After washing the cells 3 times with PBS, the cells were incubated with Alexa Fluor 546-conjugated secondary antibody (A-11030; Invitrogen) diluted in PBS containing 1% skim milk for 1 h at RT. To stain actin filaments (F-actin), the cells were blocked with PBS containing 5% BSA for 30 min at RT and were incubated with CF594 Phalloidin (00045; Biotium) in PBS containing 1% BSA for 20 min at RT, followed by PBS washes. The nuclei were stained with 4',6'-diamidino-2-phenylindole (DAPI) (100 ng/ml) for 10 min at RT. Finally, the cells were sealed with fluorescence mounting medium (S3023; Dako). Fluorescent images were captured with an Olympus LX71 fluorescence microscope.

Soft Agar Colony Formation Assay
The cells were seeded at 4.6×10 5 cells (6 cm dish), cultured in Assay Medium at 37°C for 24 h and were then treated with TAM at 37°C for 72 h before the colony formation assay. For siRNA-mediated FAT4 knockdown experiments, the cells were seeded at 1.8×10 5 cells (6 cm dish) and cultured in Growth Medium without antibiotics at 37°C for 24 h. The cells were then transfected twice with FAT4 or control siRNA (at 24 h and 48 h after plating) and cultured for a further 24 h after the last transfection. Subsequently, the cells were seeded at 7.0×10 2 cells per well (96-well plate) in Growth Medium containing 0.4% agarose (100 μl) onto a lower layer of Growth Medium containing 0.5% agar (100 μl). Cells were fed 2 times per a week. The number of colonies was counted after 14 days.

Cell Density Assay
The cells were seeded at 4.2×10 5 cells (High density) or 7.0×10 4 (Low density) cells per well (6-well plate), cultured in Assay Medium at 37°C for 24 h and subjected to RT-qPCR analysis.

Softwell Assay
Twenty-four well Softwell plates of two different qualities of stiffness (50 kPa: stiff or 0.5 kPa: soft) (Matrigen) were coated by Collagen I, rat tail (354236, BD Biosciences) (10°g/ml) for 1 h at RT. The cells were seeded at 8.1×10 4 cells per well in Softwell plates. The cells were cultured in Assay Medium for 48 h and subjected to RT-qPCR analysis.

Statistical analyses
Statistical analyses were performed using Student's t-test; a P < 0.05 was considered significant.