extfieldsofstudy: Medicine Biology

text:
The Neisseria meningitidis ADP-Ribosyltransferase NarE Enters Human Epithelial Cells and Disrupts Epithelial Monolayer Integrity

Many pathogenic bacteria utilize ADP-ribosylating toxins to modify and impair essential functions of eukaryotic cells. It has been previously reported that Neisseria meningitidis possesses an ADP-ribosyltransferase enzyme, NarE, retaining the capacity to hydrolyse NAD and to transfer ADP-ribose moiety to arginine residues in target acceptor proteins. Here we show that upon internalization into human epithelial cells, NarE gains access to the cytoplasm and, through its ADP-ribosylating activity, targets host cell proteins. Notably, we observed that these events trigger the disruption of the epithelial monolayer integrity and the activation of the apoptotic pathway. Overall, our findings provide, for the first time, evidence for a biological activity of NarE on host cells, suggesting its possible involvement in Neisseria pathogenesis.

Introduction
Neisseria meningitidis is a Gram-negative, aerobic, non-motile, non-sporulating, usually encapsulated and piliated bacterium. It is restricted to humans and generally colonizes the nasopharynx of 8-20% of healthy individuals, however in a small proportion of infected patients, the bacterium crosses the mucosal barrier and reaches the bloodstream, giving rise to meningitis or fulminant septicaemia [1]. Masignani et al. have identified, through a profile-based computational approach, an ADP-ribosyltransferase protein, NarE, which shares structural homologies with E. coli heat-labile enterotoxin (LT) and cholera toxin (CT) [2]. NarE possesses both ADPribosylating and NAD-glycohydrolase activities, confirmed by the evidence that, in the presence of an ADP-ribose acceptor, NarE acts as a transferase whereas in the absence of the acceptor it acts as a NAD glycohydrolase [3]. Furthermore NarE undergoes auto-ADP-ribosylation [4].
Mono ADP-ribosylation is a post-translational modification of proteins, shared by eukaryotes and prokaryotes, which modulates protein function [5]. Mono-ADP-ribosyltransferases (ADPRTs) catalyze the transfer of a single ADP-ribose group of β-nicotinamide adenine dinucleotide (NAD + ) to protein/peptide target acceptors with the release of nicotinamide (Nam) at the same time [6]. In pathogenic bacteria, proteins known to belong to this class of enzymes are generally classified as toxins since they alter or impair essential functions of host eukaryotic cells [7,8]. On the basis of the ADPRTs targets, at least three groups of ADP-ribosylating toxins can be identified. One group causes ADP-ribosylation of G proteins. Members of this group are cholera toxin (CT) [9], E. coli heat-labile enterotoxin (LT) [10] and pertussis toxin (PT) [11], which, through modification of regulatory G proteins, impair signal transduction. The second group includes diphtheria toxin (DT) [12] and Pseudomonas exotoxin A (ExoA) [13] that target elongation factor 2 (EF-2), thus inhibiting protein synthesis. A large third group of bacterial toxins modulates actin cytoskeleton directly, by covalent modification of actin, as C2 toxin of C. botulinum [14], Iota toxin of Clostridium [15], VIP2 toxin of Bacillus cereus [16], and SpvB of Salmonella [17], or indirectly, by covalent modification of Rho GTPases, as C3 exoenzymes of Clostridium, Bacillus cereus and S.aureus [18,19] exoenzyme S (ExoS) of P.aeruginosa [20].
Each group of toxins provides the bacterial pathogen with a selective advantage in modulating cell host response and resistance to infection, therefore they have been extensively characterized.
The narE gene is present only in a subset of hypervirulent clusters, ET-5 and Lineage 3 complexes, suggesting its involvement in Neisseria meningitidis pathogenesis [3]. However, no evidence of NarE toxic activity has been provided so far and its function remains to be fully elucidated. In the present report, we show that NarE targets Chang human epithelial cells. We demonstrated that NarE is internalized and gains access to the cytoplasm. Furthermore, through its ADP-ribosylating activity, NarE targets host cell proteins, alters epithelial monolayer integrity and initiates the apoptotic pathway responsible for cell death. Collectively, our data provide for the first time evidence of the biological role of this enzyme and suggest its potential contribution during colonization of upper respiratory tract and spreading of infection.

Cells, antibodies, reagents and recombinant proteins
Chang human epithelial cell line (HeLa contaminant) was purchased from the American Type Culture Collection (ATCC, CCL-20.2). Chang cells were maintained in minimum essential medium Eagle (MEM, Invitrogen Ltd, Paisley, UK) supplemented with 10% heat-inactivated fetal bovine serum (Invitrogen Ltd, Paisley, UK), 15mM L-glutamine and antibiotics. Cells were grown at 37°C in a humidified 5% CO 2 atmosphere. In order to produce NarE polyclonal antiserum, CD1 mice were immunized with 10 μg of purified protein formulated with Al (OH) 3 , as an adjuvant. The recombinant protein was given intraperitoneally (day1), a second (day 21) and a third (day 35) booster doses were administered. Blood samples were taken on day 49. Antibody against cleaved caspase-3, anti-GAPDH and anti-Lamin were from Cell Signaling Technology (Beverly, MA). Antibody anti-ADAM10 was from Abcam, anti-cytokeratin was from Invitrogen and anti-actin was from Biosource. Mouse antibodies against MHCI were from Biolegend, anti-Lamp1 from Abcam, Rabbit antibodies anti-EEA1 were from Novus Biologicals. Rabbit anti-VAP-A antibody was kindly provided by Antonella De Matteis, (Telethon Institute of Genetics and Medicine, Pozzuoli, Naples) and rabbit anti-Giantin was from Covance. Alexa 488-and Alexa 568-conjugated secondary anti-rabbit or anti-mouse goat antibodies were from Molecular Probes. Recombinant NarE was conjugated with fixable Alexa-Fluor-546 according to manufacturers' instructions (kit A10237, Life Technologies). DNase I, Fluorescent Deoxyribonuclease Conjugates, Phalloidin-TRIC as well as FITC-labeled dUTP (Click-iT TUNEL Alexa Fluor 488 Imaging Assay) were obtained from Molecular Probes. CHX and NAD were purchased from Calbiochem (Merck Biosciences GmbH, Schwalboch, Germany). NarE (accession number NMB1343) and NarE-R7K recombinant proteins were cloned, expressed and purified as described [3]. In both preparations the endotoxin content, determined using a LAL assay (QCL-1000 kit from Lonza), was below 0.5 EU/mg protein.

Ethics statement
Mouse studies were performed in accordance with Novartis Animal Welfare Policies and Italian laws on care and protection of laboratory animals. Mice were euthanized in accordance with experimental protocols. Experimental protocols were reviewed and approved by the Italian National Institute of Health and by the local Novartis Vaccines and Diagnostics Animal Welfare Body (authorization AEC 201101). CD1 mice (Charles River Laboratories) were used for the experiments.

FACS analysis
To measure NarE binding to Chang cells, 2x10 5 cells were incubated with different concentrations of NarE or NarE-R7K (ranging from 0,01 to 100 μg/ml) in serum-free DMEM in 96-well culture plates for 1h on ice. Cells were then washed and incubated on ice with anti-NarE serum for 30 min. After washing, cells were stained with FITC conjugated secondary antibodies. Cells were analysed with a Canto II flow cytometer (Beckton-Dickinson) by using Flowjo software. The geometric mean fluorescence intensity (MFI) for each population was calculated.

In vitro ADP-ribosylation assay
For cell-free ADP-ribosylation assay, 50 μg of whole-cell lysate were used. ADP-ribosylation reaction was performed for 30 min at 37°C with 1 μg NarE or NarE-R7K in a buffer containing 20 mM Tris-HCl (pH 8), 1 mM EDTA, 1 mM DTT, 5 mM MgCl 2 , 10 μM biotinylated NAD + (Trevigen) and Complete protease inhibitor (Roche, Basel, Switzerland) (according to the manufacturer's manual). The reaction was stopped by adding 4x NuPAGE LDS Sample Buffer (Invitrogen) and 10x NuPAGE Reducing Agent (Invitrogen) and boiling the samples for 5 min at 100°C. The samples were subjected to SDS-PAGE and subsequently the biotin-ADP-ribosylated proteins were transferred to a nitrocellulose membrane and visualized with HRPconjugated streptavidin.

Immunofluorescence microscopy and pulse-chase experiments
Chang cells (5x10 4 /well) were plated on 8-wells Lab-Tek II Chamber Slide System (Thermo Scientific) and incubated overnight at 37°C 5% CO2. For NarE binding and to visualize morphological effects, cells were incubated at for different times with NarE/NarE-R7K. After washing with PBS, cells were fixed in paraformaldehyde 2% for 20 min at RT (room temperature) and then, after three washes with PBS, permeabilized with TritonX 0.1% plus saponin 1% for 20 min at RT. Cells were washed three times with PBST (PBS 1x, TritonX 0,1%) and incubated for 30 min at RT with Duolink II Blocking Solution 1X (Olink Bioscience). After washing with PBST twice, cells were incubated for 15 min at RT with mouse anti-NarE serum. Cells were washed again with PBST twice and incubated for 10 min at RT with secondary antibody Alexa Fluor 568 anti-mouse. Subsequently, cells were washed with PBS twice and, after drying at RT, one drop of ProLong Gold Antifade Reagent with DAPI (Invitrogen) was added. For pulse-chase experiments, in order to detach not internalized NarE bound to the plasma membrane of Chang cells, they were first incubated with NarE for 1 h, then they were washed with ice-cold phosphate-buffered saline (PBS) containing 500 mM NaCl and 0.5% acetic acid and three times with complete medium; before fixation the cells were incubated for 1 h. For the analysis of the subcellular localization of NarE, the slides were analyzed by Confocal microscopy, performed on a LSM700 (Carl Zeiss, Inc.) and LSM 510 (Carl Zeiss, Inc.) confocal microscopes using a Plan-Neofluar 63X objective. Detectors were set to detect an optimal signal below the saturation limits. Images were processed with Zen 2009 image software (Carl Zeiss, Inc.).
xCELLigence system: time-and dose-dependent cell response profiles of Chang cells The effect of NarE on epithelial barrier function was measured with an electrical cell-substrate impedance sensing system (Applied BioPhysics). In total, 1 × 10 5 Chang cells were seeded in 8W10E cultureware and incubated at 37°C in a carbon dioxide incubator. Resistance of the monolayer was recorded until a stable resistance of approximately 600-1000 O was documented prior to the addition of purified protein.

Cell treatments, lysis, and immunoblots
For immunoblot analysis, cells (2x10 6 cells/well) were plated in a 6-well dish, grown overnight at 37°C to reach a sub-confluent condition, and incubated with NarE/NarE-R7K at 37°C in a humidified atmosphere with 5% CO2 for different time. Cells were recovered by centrifugation at 16,000Xg for 30 sec at 4°C, washed 2X in PBS and lysed in 1% (v/v) Triton X-100 in 20 mM Tris-HCl pH 8, 150 mM NaCl (in the presence of0.2 mg/ml Na orthovanadate, 1 mg/ml pepstatin, leupeptin, and aprotinin, and 10 mM phenyl methyl sulfonyl fluoride). Alternatively, for subcellular protein fractionation, cells were collected, washed in ice-cold PBS and processed by Thermo scientific subcellular protein fractionation Kit. Protein content of isolated fractions, corresponding to cytoplasmic, membrane, nuclear and pellet fraction, was determined by BCA assay. Equal amounts of proteins from were resolved by NuPage Novex 4-12% Bis-Tris Gels (Invitrogen) and transferred to 0.45-mm nitrocellulose filters nitrocellulose membrane by means of iBlot Gel Transfer Device (Invitrogen). Pre-stained molecular mass markers (Invitrogen) were included in each gel. Immunoblots were carried out using primary antibodies and peroxidase-labeled secondary antibodies according to the manufacturers' instructions and a chemiluminescence detection kit (Pierce). Blots were scanned using a laser densitometer (DuoscanT2500 Agfa, Milan, Italy) and quantified using the ImageQuant5.0 software (Molecular Dynamics, Sunnyvale, CA). Data were normalized to loading controls.

Statistical analysis
Mean values, standard deviation values and Student's t test (unpaired) were calculated using the Microsoft Excel application. A level of P <0.05 was considered statistically significant.

NarE binding to human epithelial cells leads to protein internalization
The biological function of NarE was assessed on Chang human epithelial cells, a well-established in vitro model to study N. meningitidis interaction with the host [21,22]. Purified recombinant protein was added at increasing concentrations to epithelial cells and the binding was measured by flow cytometric analysis. As shown in Fig 1A, NarE, is able to bind Chang cells in a concentration dependent manner, with a rapid increase in binding at lower concentrations, a kinetic probably reflecting NarE interaction with a high affinity receptor. Similar results were obtained when the enzymatically inactive variant of the toxin, NarE-R7K, which is devoid of the ADP-ribosylation activity, was used [23] (data not shown). The evidence that NarE is able to bind to host epithelial cells was further demonstrated by confocal analysis (S1 Fig). Of interest, we observed that NarE not only is able to bind to target cells but it can also be internalized. Indeed, confocal microscopy analysis of cells extensively washed with high-salt acidic buffer (which allows the removal of surface-associated proteins) revealed that NarE is found intracellularly within numerous perinuclear structures ( Fig 1B).

NarE is internalized through an endocytic traffic route
To further characterize the intracellular trafficking of NarE, Chang cells were incubated with recombinant fluorescently labelled NarE (Alexa 546-NarE) for 1h at 37°C. Cells were then processed by immunofluorescence analysis to assess the co-localization of NarE with established markers of intracellular compartments. Confocal analysis showed that NarE is distributed in punctate structures at the peripheral and juxtanuclear region of the cell (Fig 2A). Notably, these structures were found to co-localize with the major histocompatibility complex Class I (MHCI) protein (Fig 2B), a marker of recycling endosomes, and EEA1 (Fig 2C), a marker of early endosomes. No co-localization was observed with Golgi and ER specific markers, or with LAMP-1, a lysosomal-associated membrane protein ( S2 Fig and Fig 2D, respectively). These data postulate that NarE is actively internalized into epithelial cells through an endocytic traffic route.

NarE gains access to the cytoplasm and targets host cell proteins
We then hypothesized that NarE internalization and consequent association to the endocytic route, may allow this ADP-ribosyl transferase to gain access to the cytosol. To this end we Chang cells were plated on coverslips and incubated with 1 g/ml of recombinant NarE for 1 h. Next, the cells were washed with a high-salt and acid buffer to rinse the non-internalized NarE. Finally, the cells were fixed and stained with an anti-NarE antibodies (Fig 1B, left panel) or further incubated for 1h at 37°C before being processed (Fig 1B, right panel). All the images were acquired using with the same imaging conditions. Three independent experiments were performed. Scale bar is 10 μm. isolated cytoplasmic, membrane, nuclear soluble and cytoskeletal fractions from Chang cells incubated with NarE. Briefly, Chang cells were exposed to NarE for 1 h, washed, to remove unbound NarE, and further incubated for 1, 3 and 6h before being processed. As shown in Fig 3A, Western blotting analysis revealed that already at 2h, NarE accumulated in the cytosolic fraction. To identify putative cellular substrate/s ADP-ribosylated by NarE, whole-cell lysates were incubated with NarE or NarE-R7K in presence of biotinylated NAD + (or with 32 P-NAD + , data not shown). Interestingly, as shown in Fig 3B, incubation of cell extracts with NarE, but not with the genetically inactivated mutant NarE-R7K, resulted in the increase of ADP-ribosylated proteins, indicating that NarE targets Chang cell proteins. Of note, we detected bands of approximately 42-45 kDa, compatible with the molecular weight of actin, or of α-subunit of heterotrimeric G proteins, both well-known targets for modification of many bacterial toxins [8,7].

NarE impairs human epithelial monolayer integrity and induces cell rounding
Since NarE ADP-ribosylates Chang substrates, we then examined whether it could alter essential functions of eukaryotic cells. To this end electrical cell-substrate impedance sensing (xCEL-Ligence) was used to measure trans-epithelial electrical resistance. Impedance reflects the status of the cell monolayer, including confluence, viability, and junction functionality [24,25]. Chang cells were grown to confluence on xCELLigence inserts and the experiments were started when trans-epithelial resistance (TEER) was found constant between two measures taken 24 hours apart. Cells were then incubated with NarE or NarE-R7K. Vehicle was used as negative control. Changes in monolayer integrity were monitored every 15 min. As shown in Fig 4A Chang monolayer treated with 1μg/ml NarE demonstrated a rapid and progressive loss of resistance. Similar results were obtained when Clostridium difficile toxin, TcdA, a wellknown cytotoxin affecting actin cytoskeleton [7,26], was used. Neither vehicle nor NarER-7K altered monolayer resistance.
In the attempt to correlate changes in monolayer integrity with cytopathic events, Nar-E-R7K and NarE treated cells were visualized by light microscopy. As shown in Fig 4B NarE, as well as TcdA, our positive control, induced morphological changes as the cells rounded up within 4h (Fig 4B). On the contrary, host cells exposed to the mutant form of the protein did not show any changes in their morphology, indicating that induction of the cell-rounding phenotype of Chang cells required a functional ADP-ribosylating domain.

NarE impacts on actin cytoskeleton
The observation that one of the ADP-ribosylated substrates could be actin (Fig 3, panel B), along with the phenotype of intoxication (cell rounding), lead us to further assess NarE effect on actin cytoskeleton of Chang cells. To this end, after 4 hours of incubation of cells with the wild-type or the mutant protein, morphological alterations were analyzed by confocal microscopy using fluorochrome-conjugated phalloidin. F-actin depolymerization and dissolution of cytoskeleton was observed in cells treated with NarE, suggesting that actin could be, directly or indirectly, a target of NarE-ADP-ribosylating activity. Representative micrographs are shown in Fig 5A. These results were further corroborated by monitoring global changes in actin dynamics using the DNase I inhibition assay, a selective analysis in which G-actin is detected with the fluorophore-derivatized Dnase I. Fig 5B displays a significant increase in the amount of unpolymerized actin after incubation with NarE. Similar results were obtained when TcdA was used. Notably, any effect on actin cytoskeleton was detected in cells treated with NarE-R7K.

NarE induces apoptosis in Chang cells
We next investigated whether NarE-induced cell damages correlate with the activation of cell death pathways. Among a wide range of factors controlling apoptotic cell death, caspase-3 activation plays a key role and its activation requires proteolytic processing of its inactive zymogene into activated p17/19 and p12 fragments [27]. Therefore in order to assess NarE effect on Chang monolayer was treated either with NarE or NarE-R7K (1μg/ml) and barrier resistance was continuously measured using an electric cell-substrate impedance sensing (xCELLigence) system. TcdA (100 ng/ml) was used as positive control, while saline was used as negative control. CI, Cell Index (arbitrary unit for electric impedance measurement). Top right, Variation of Chang cells CI after 6 h of treatment with NarE or NarE-R7K (1μg/ml). The results are expressed as the difference between normalized CI before agents addition (CI = 1) and CI value after 6 h of incubation averaged from three independent experiments, each performed on duplicate samples. Error bars represent the SD. **, P 0.01. (B) Effect of NarE treatment on the morphology of Chang cells. Chang cells were incubated at 37°C with NarE, NarE-R7K (1μg/ml), TcdA as positive control (100 ng/ml) or left untreated as control. After 4 hours, pictures were taken to assess the change in morphology. Bar, 20 μm.  the apoptotic pathway, Chang cells were incubated with NarE, NarE-R7K or cycloheximide, CHX, (as positive control) for various times (2h to 24h) and then caspase-3 cleavage, which is an indication of its activation state, was evaluated. As reported in Fig 6, 2h and 6 h of incubation with the toxin were sufficient to induce caspase 3 cleavage. However, there was a peak activity after 24 h of treatment. As expected, NarE-R7K, which is devoid of the ADP-ribosylation activity, did not induce caspase-3 activation.
The development of apoptosis was further assessed using Terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling (TUNEL). To this end Chang cells were treated with NarE for up to 24h. The appearance of fluorescence in the nuclei of NarE-treated cells showed evidence of apoptosis (Fig 6B, right panel), whereas no DNA fragmentation was observed in cells treated with NarE-R7K or in control cells (vehicle only). Following incubation with NarE for 24h, the amount of apoptotic cells significantly increased as shown by elevation of fragmented DNA (calculated as specific apoptosis as percentage of total cells) in Fig 6B (left panel).

Discussion
In this paper, we focus on the biological functions of NarE when getting in contact to epithelial cells. In particular, we reported that NarE binds to Chang cells in a dose dependent and saturable manner, and that these events trigger the internalization of this ADP-ribosyltransferase that once in the cytoplasm, exerts its enzymatic activity on host cell proteins. This results in the alteration of cell morphology and epithelial monolayer integrity leading to the activation of caspase-3, an apoptosis executer (cytotoxic effect). Notably, similar cytopathic and cytotoxic effects are reported for various bacterial toxins targeting mammalian cells (e.g. C. botulinum C2, C. perfringens iota toxin and S. enterica SpvB), which by inducing loss of adhesion and cell rounding lead to the activation of the apoptotic pathway [28,29,7].
To demonstrate that ADP-ribosylating activity of NarE is crucial in inducing a toxic phenotype, a catalytically inactivated mutant, NarE-R7K, was used. As reported by Koehler C. et al. [23], direct mutagenesis of R7 residue, to K, results in a drastic reduction of ADP-ribosylation activity. The NarE mutant loses both transferase and glycohydrolase activities, likely due to a reduced NAD binding.
ADP-ribosyltransferase activities have been observed in many prokaryotic and eukaryotic species and in viruses. The best characterized mono-ADP-ribosylation reactions are those catalyzed by bacterial toxins, such as diphtheria, cholera, pertussis and clostridial toxins, which act by modifying crucial host cell proteins such as the α-subunit of heterotrimeric GTP-binding (G) proteins, the small GTPase Rho, monomeric actin and elongation factor 2 (EF-2), resulting in permanent activation or inactivation of the cell functions modulated by these protein substrates. Consequently, these toxins have been characterized extensively for their activity and represent some of the best understood microbial mediators of disease and have been used as components of vaccines able to prevent associated diseases [30,31].
By in silico studies Masignani and colleagues identified an ADP-ribosyltransferase in MC58 strain of N. meningitidis serogroup B. They demonstrated that this protein is able to hydrolase NAD and to transfer the ADP-ribose group to small guanidine compounds like agmatine and arginine and so they called it NarE, N. meningitidis ADP-ribosylating enzyme. Interestingly, they found that, despite the absence of a predicted leader peptide, NarE efficiently accumulates in the periplasm of Meningococcus [3]. Further, they showed that the narE gene is present only in a subset of hypervirulent strains of bacteria.
An open question is how NarE, being a periplasmic protein, could achieve contact with host cells. NarE lacks a leader peptide as well as a gene coding for the translocation/receptor-binding subunit B but since it is a really small proteins it maybe can reach the periplasmic space of Meningococcus by translocating across the cytoplasmic membrane through natural membrane pores [3]. However, in order to exert a toxic activity, NarE has to contact host cells. For this reason, once in the periplasm, a second step is required to allow the release of the protein. A possible hypothesis would be that NarE follows a pathway similar to that described for LT of E. coli [32] and it could be released in association with vesicles. Internalization of vesicles could then allow intoxication of the host cells. Actually, we have experimental data suggesting that NarE is present in the outer membrane vesicles (OMVs) generated following treatment with denaturing agents of the MC58 strain (Rossi Paccani S. et al., unpublished observation). Furthermore, recent observations indicate that NarE protein is expressed in the OMVs of hypervirulent clones and that fusion of OMVs with host immune cell allows delivery of NarE, which impairs cellular effector function [33]. These data strongly support the hypothesis that NarE intoxication of host cells occurs upon internalization of OMVs. However, we cannot exclude that NarE export through the outer membrane could occur following cell contact or could just happen upon lysis of the bacterium [3].
Even if the mechanism by which NarE reaches host cells is still unknown, our data clearly highlight that the protein is able to bind to epithelial cells and alter their integrity suggesting a possible role of NarE in first stages of N. meningitidis pathogenesis, indeed disruption of epithelial barrier is a key step for a successful colonization of upper respiratory tract and spreading of infection. Further investigations are needed to elucidate the pathway followed by this newly identified ADP-ribosylating enzyme to get to epithelial cells and its potential role in the virulence and pathogenesis of meningococcal species.
Supporting Information S1 Fig. NarE binds to Chang cells. Immunofluorescence analysis of NarE binding to Chang cells. Cells were either untreated or incubated with 1ug/ml NarE at 37°C (upper panel) or 4°C (lower panel) for 1 hour and then fixed and stained. Representative confocal images are shown. Cells were co-stained with anti-NarE (red), phalloidin (green) and DAPI (blue). Bar, 20 μm (TIF) S2 Fig. Immunofluorescence analysis of NarE subcellular localization. Chang cells were plated on coverslips and incubated for the indicated times with NarE. Vehicle was used as a negative control. Cells were fixed and labelled with an anti-NarE antibody and with an anti-Giantin antibody (left panel) or an anti-VAP-A antibody (right panel). The red channel corresponds to NarE whereas the green one to Giantin or VAP-A. Magnified areas are shown in the inserts. All the images were acquired using the same confocal settings. Scale bar is 10 μm. (TIF)


extfieldsofstudy: Biology Medicine

text:
MicroRNA analysis suggests an additional level of feedback regulation in the NF-κB signaling cascade.

It is increasingly clear that the biological functions of a transcription factor cannot be fully understood solely on the basis of protein-coding genes that fall under its control. Many transcription factors regulate expression of miRNAs, which affect the cell by modulating translation and stability of mRNAs. The identities and the roles of NF-κB-regulated miRNAs have been attracting research interest for a long time. We revisited this issue in a system with controlled expression of one of the key regulators of NF-κB, RIPK1. Several regulated miRNAs were identified, including miR-146a, miR-215 and miR-497. The miRNAs were also inducible by IL-1β, but not when NF-κB activity was repressed by mutant IκBα. The presence of a miR-497 site was predicted in the 3′-UTR of IKBKB gene, which encodes IKKβ. Using appropriately engineered reporters, we confirmed that this site can be a target of suppressive action of miR-497. Our findings suggest that NF-κB controls expression of a miRNA, which may reduce production of IKKβ. Considering the role of IKKβ in the canonical pathway of NF-κB activation, our observations may indicate a new mechanism that modulates the magnitude of such activation, as well as the propensity of a cell to engage canonical vs. non-canonical pathways.

INTRODUCTION
MiRNAs are short non-coding RNAs predominantly known for attenuating translation of various proteincoding RNAs [1]. Changes in the repertoire of miRNAs accompany such diverse phenomena as aging [2,3], cognitive impairment [4], and oncogenesis [5,6], and it is widely accepted that miRNAs control major physiological and pathophysiological processes. MiRNAs are generated from longer transcripts via a series of nucleolytic processing steps [7]. Each mature miRNA may target multiple mRNAs [8]. Short stretches of homology between the miRNA and the mRNA are believed to mediate this interaction, but the precise criteria that determine the specificity of this interaction are still unknown [9]. The repertoire and the abundance of miRNAs in a given cell are tightly regulated. In part, this is effected through RNA-processing enzymes [10], but the best studied mode of such a regulation is at the level of transcription [1]. A large fraction of all miRNAs are generated from RNA polymerase II-driven transcripts [11], and the same transcription factors that control protein-coding genes affect miRNA expressions as well. Indeed, it could be argued that a biological role of a transcription factor cannot be fully understood without considering the spectrum of non-protein-coding RNAs that it controls.
The signaling pathway, which culminates in activation of NF-κB family of transcription factors, was among the first to be explored for the effects on miRNA expression [12]. Typically, NF-κB factors act as dimers, which could be formed by various members of the family. RelA, RelB and c-Rel subunits possess both transactivation and DNA-binding activity, while p50 and p52 subunits can bind DNA, but are incapable of transactivation on their own. The activation of NF-κB is a key feature of immune response and inflammation [13]. The respective regulatory pathway is stimulated by a number of cytokines and other secreted factors, by pathogens and xenobiotics, as well as by certain intracellular stressors. When activated, NF-κB promotes expression of genes that contribute to immune response, angiogenesis, cell survival and many other processes [13]. Deregulation of NF-κB has been reported in autoimmune and inflammatory disorders [14][15][16] and aging [17,18]. Elevated activity of NF-κB is also often observed in human malignancies, where it is believed to contribute to growth, survival and spread of the cancer cells [19].
NF-κB is under an intricate control, which still remains a subject of intense research. The centerpiece of the canonical pathway of NF-κB activation is a protein kinase called IKK [20]. This enzyme, which is composed of IKKα, IKKβ and IKKγ subunits, phosphorylates IκB ("inhibitor of kappa B") proteins. The latter normally act to sequester NF-κB in the cytoplasm. Upon phosphorylation, IκB is targeted for degradation, while released NF-κB translocates towards its target genes in the nucleus.
A so-called "non-canonical" pathway relies on IKKα, but not the full IKK, to enhance production of NF-κB subunit p52 from its inactive precursor [21]. This eventually results in accumulation of p52/RelB dimer in the nucleus. Inducers of the canonical pathway are plentiful, with tumor necrosis factor alpha (TNFα) and interleukin-1 beta (IL-1β) being among the most notable examples [22]. Activation of the non-canonical pathway has been attributed to such molecules as lymphotoxin-α and BAFF [21]. Since various NF-κB dimers have overlapping, but not identical sets of targets, the distinction between canonical and non-canonical pathways may explain some of the differences in the biological functions of NF-κB-activating stimuli. Importantly, among the transcriptional targets of NF-κB are both the negative and the positive regulators of this pathway [23], and evidence of complex cross-talk between the canonical and non-canonical pathways have been reported [23,24].
While the interest towards identifying NF-κB target genes and their roles has remained steady over many years, the tools available to explore these issues have been rapidly evolving. For example, an early survey of NF-κBinducible miRNAs was limited to an array of 200 human miRNAs [12]. In contrast, miRBase, an online database of known miRNAs [25], currently contains approximately 2000 unique entries corresponding to Homo sapiens. In many cases, the ranks of the previously known miRNAs were expanded due to the discovery of new molecules. In other cases, what used to be considered a single miRNA species, proved to be a family of closely-related sequences, which originate from distinct genes, but are hard to distinguish in conventional array hybridization assays. Modern high-throughput sequencing techniques, coupled with powerful instruments of sequence analysis, are primarily responsible for the increased breadth and sensitivity of detection of small RNAs. The availability of these tools facilitates continuous exploration of miRNA profiles in new experimental systems, but also warrants a fresh look at the effects of previously identified regulators of miRNA expression. RIPK1 protein is a central and essential element of multiple signaling pathways, including those initiated by TNFα, various xenobiotics and stress factors [26]. Depending on the context, RIPK1-dependent signaling leads to inflammation, necroptosis or apoptosis [26]. TNFα, acting through its cognate receptor and RIPK1, is among the best-studied activators of NF-κB [27]. In earlier attempts to identify events that may lead to NF-κB deregulation, we observed that overexpression of both fulllength and short forms of RIPK1 mimic many aspects of TNFα treatment and sustains constitutive activation of NF-κB [28,29]. In the current study, we identify miRNAs that are up-regulated following up-regulation of short RIPK1, and we document that these miRNAs are induced by other NF-κB-stimulating events in an NF-κB-dependent manner. Furthermore, we demonstrate that one of the induced miRNAs, miR-497, is a likely negative regulator of IKKβ, providing yet another layer of regulation for the NF-κB signaling cascade.

Identification of RIPK1-inducible miRNA
We used sequencing-based high-throughput small RNA profiling technique to identify RIPK1-dependent changes in miRNA abundance. We relied on a previously established cell line ("clone 3B4.1"), which contains a tetracycline-regulated promoter inserted into RIPK1 gene [28]. Clone 3B4.1 was originally selected for high NF-κB activity from among C6TA4 cells, which were mutagenized with a promoter-bearing transposon [28]. In turn, C6TA4 are derived from HEK293 cells, preengineered with a tetracycline transactivation protein and two NF-κB-responsive markers (Zeocin resistance and ganciclovir sensitivity) [30,31]. The inserted promoter in 3B4.1 clone drives expression of a shorter form of RIPK1 protein, which potently activates NF-κB and mimics other effects of TNFα treatment [28]. Importantly, exposure to tetracycline or doxycycline inhibits the promoter and, eventually, brings the activity of NF-κB down to a basal level for the parental cell line [28].
The RNA samples from 3B4.1 cells with and without doxycycline treatment were enriched for small RNAs using size-fractionation and profiled using highthroughput sequencing as described in Materials and Methods (Supplementary Table 1). We have attempted to confirm the observed differences in the expression of miR-14a, -215, -381 and -497 ( Figure 1A) using quantitative PCR. In accordance with the sequencing results, the expression levels of three of the miRNAs (miR-146a, miR-215 and miR-497) were sensitive to manipulation of RIPK1 expression ( Figure 1B, Figure 1C and Figure 1D), and the differences between doxycycline-treated and untreated samples were significant for each of these miRNAs (p < 0.05). Importantly, miR-146a is a well-known transcriptional target of NF-κB [12], and may serve as an internal control for activation of the corresponding signaling cascade in our experimental conditions. The fourth miRNA, miR-381, was undetectable in these conditions (data not shown). Considering the low number of corresponding reads in the sequencing experiment, we find it very likely that miR-381 is expressed at a very low level and is unlikely to play a significant biological role in our experimental system.

The role of NF-κB in the regulation of miR-146a, miR-215 and miR-497
While NF-κB is prominently induced by RIPK1 in 3B4.1 cells, it is not the only signaling cascade activated in these cells. We undertook two complementary approaches in order to verify that the induction of the identified miRNAs is, indeed, NF-κB-dependent.
First, we examined whether the same miRNAs are inducible by IL-1β. TNFα, whose action is mimicked by overexpression of RIPK1 in 3B4.1 cells, and IL-1β signal through distinct intermediates and elicit non-identical responses in the affected cells. However, both cytokines are powerful inducers of NF-κB activity. Accordingly, we observed that IL-1β treatment of C6TA4 cells resulted in elevated expression of miR-146a, miR-215 and miR-497 ( Figure 2A, Figure 2B and Figure 2C, respectively). The differences between samples at the start (0 hr.) and at the end (8 hr.) of the experiment were significant (p < 0.05) for all the three miRNAs.
Second, we examined how the induction of miR-146a and miR-497 is affected by the presence of a "super-repressor" mutant of IκBα, which lacks the phosphorylation site of IKK. The unphosphorylatable mutant cannot be targeted by upstream signals for degradation and, hence, continues to sequester NF-κB in the cytoplasm even upon stimulation with cytokines [32]. Accordingly, we have previously observed that C6TA4 cells harboring mutant IκBα fail to activate NF-κB and are hypersensitive to pro-apoptotic effects of TNFα [28]. As expected, the ability of IL-1β to induce the miRNAs was diminished when the mutant IκBα was expressed (compare Figure 2A to Figure 3A, Figure 2B to Figure 3B, and Figure 2C to Figure 3C), and this effect was significant (p < 0.05) for all the three miRNAs. In fact, when NF-κB activity was suppressed, miR-497 expression was reduced, rather than elevated, following IL-1β treatment ( Figure 3B). A small increase in miR-215 levels after IL-1β treatment in these cells ( Figure 3C) may reflect a contribution of an NF-κBindependent mechanism or a residual NF-κB activity, which remains despite expression of the IκBα mutant.
Overall, our observations suggest that miR-146a, miR-215 and miR-497 are inducible by various stimuli in an NF-κB-dependent manner.

MiR-497 targets a site in the 3′-UTR of IKKβ mRNA
We used TargetScan [33] to predict possible targets of miR-497. The analysis revealed the presence of a putative miR-497 binding site in the 3′-UTR of IKKβ mRNA (encoded by IKBKB gene). We have tested this prediction by fusing the 3′-UTR fragment of IKKβ mRNA to the coding region of firefly luciferase ( Figure 4A). The resulting construct demonstrated a profound sensitivity to the presence of miR-497 in transient transfection assays ( Figure 4B), a feature not seen with the original luciferase vector ( Figure 4D). The latter observation indicates that the phenomenon was not due to a generic effect on the function of the RSV promoter or the luciferase itself. Furthermore, the inhibitory effect of miR-497 was greatly reduced when the putative miR-497 site in the IKKβ UTR was mutated ( Figure 4C). The fraction of pRSV-luc expression remaining after miR-497 co-transfection was significantly (p < 0.05) different from that of pRSV-luc-IKBKB, but not of pRSV-luc-mIKBKB. We concluded that the miR-497 site in the 3′-UTR of IKKβ mRNA acts as a bona fide regulator of protein expression in response to elevated levels of this miRNA.
Accordingly, transfection of miR-497 into C6TA4 cells resulted in a noticeable reduction of the levels of IKKβ protein, as seen by Western blotting ( Figure 4E). were measured by quantitative PCR at the indicated times after addition of IL-1β to C6TA4 cells. The levels of the miRNAs from triplicate experiments were normalized to those of an internal control (RNU6B) in the same samples and are shown relative to those in untreated cells. www.impactjournals.com/oncotarget DISCUSSION HEK293-derived cells, akin to the ones used in the current study, have been extensively exploited to study the function and regulation of NF-κB (e.g. [28][29][30][31]34]). In particular, there is evidence that overexpression of the short RIPK1 variant in clone 3B4.1 mimics many NF-κB-dependent and -independent consequences of TNFα treatment [28], and a transcript, which closely resembles the one being overexpressed in these cells, may naturally originate from intron 6 of RIPK1 gene [28]. It is important to note that the exact pattern of NF-κB-responsive miRNA genes is likely to vary depending on the biochemical, genetic and epigenetic environment of a cell, such as the abundance of various regulatory proteins (including individual NF-κB subunits), mutations and polymorphisms within promoter elements, as well as processing of the latter by chromatin-modifying enzymes. Accordingly, not all miRNAs, previously reported as NF-κB-responsive [35], were affected in our experiment (Supplementary Table 1). Additional factors contributing to such an apparent discrepancy could be a superimposition of multiple regulatory mechanisms, an inability to distinguish some closely-related miRNAs, which are encoded by distinctly regulated genes, and sampling errors due to very low expression of some of these molecules.
In the current study we specifically confirmed NF-κB-dependent regulation of three human miRNAs: miR-146a, miR-215 and miR-497. MiR-146a is a well-known transcriptional target of NF-κB [12]. This miRNA has attracted considerable attention due to its possible involvement in immunity, hematopoiesis and cancer [36]. The presence of a known NF-κB target on our list may serve as an internal control for activation of the corresponding signaling cascade in our experimental conditions. To the best of our knowledge, our study is the first to describe miR-215 and miR-497 as NF-κB targets. Importantly, miR-194 and -195, which are clustered together with miR-215 and miR-497 respectively, are also up-regulated in our experiments (Figure 1). This coordinated enrichment of clustered miRNAs in our experiments further supports the notion of bona fide transcriptional regulation of the corresponding genes. However, the precise molecular and miR-215 C. were measured in C6TA4 cells stably transduced with a stabilized ("super-repressor") mutant of IκBα, with or without an eight-hour treatment with IL-1β. The experiments were performed and analyzed as in Figure 2. www.impactjournals.com/oncotarget mechanisms of this regulation remain to be elucidated. The promoters of the corresponding miRNA genes have not been exhaustively characterized, and there is a formal possibility that the effect of NF-κB on their expression is indirect. Very intriguing is the observation that in the absence of NF-κB, IL-1β reduces the abundance of miR-497 ( Figure 3B). We can only speculate about the mechanism of this phenomenon. For example, it is possible that in IL-1β-stimulated cells NF-κB competes with a concomitantly activated transcriptional inhibitor; or

A.
A schematic depiction of the basic luciferase reporter (pRSV-luc), the same reporter supplied with 1.3kb fragment from IKBKB 3′-UTR carrying the predicted miR-497 recognition site (pRSV-luc-IKBKB), and the construct, in which the site has been mutated (pRSV-luc-mIKBKB; mutated bases shown in bold). B-D. The effect of miR-497 co-transfection on the luciferase expression from pRSV-luc-IKBKB (B), pRSV-luc-mIKBKB (C) and pRSV-luc (D) The activity of luciferase in transiently transfected C6TA4 cells was normalized to that of a co-transfected constitutive β-galactosidase reporter and compared between the cells transfected with miR-497 or a non-targeting control as described in Materials and Methods. E. The effect of miR-497 transfection on IKKβ level in C6TA4 cells. The cells were transfected with miR-497 or a negative RNAi control. Lysates were prepared 48 h post-transfection and probed by immunoblotting for the indicated proteins. Images from one of three consistent blots are shown. www.impactjournals.com/oncotarget that NF-κB provides a transcriptional activation domain to a DNA-bound complex, which otherwise interferes with basal transcription. It is also important to note that the abundance of a miRNA could be controlled posttranscriptionally, at the level of processing or stability.
It is commonly accepted that control of miRNA expression is an indelible function of many, if not all, transcription factors. Studies of the relationship between transcription factors and miRNAs reveal a recurrent theme of mutual regulation. MiRNAs commonly target the signaling pathway, or even the very transcription factor, which is primarily responsible for their own expression [37]. Such a mechanism has been validated or, at least, proposed for numerous transcription factors, such as SRF [38], E2F [39], FOXO [40] and others. Indeed, miR-146a represents a classic example of such a negative feedback loop [24]: its targets, TRAF6 and IRAK1, which are key elements of several NF-κB-activating pathways.
MiR-215 is often characterized as a putative tumor suppressor, decreased in cancer and capable of inhibiting cell proliferation [41][42][43][44][45], although the nature of the most relevant targets in this case remains open for debate. However, this anti-cancer role does not appear to be universal, as there are reports of malignancies with miR-215 overexpression in cells, as well as of accelerated cell growth upon miR-215 overexpression [46,47]. MiR-215 (but not the co-clustered miR-194) is reportedly upregulated by differentiation-inducing protein CDX1 and targets BMI to inhibit stemness in enterocytes [48]. The possibility that members of the same cluster could be regulated either coordinately or independently may account for some of the controversy about the correlation between miR-215 expression and cancer-related phenotypes in various systems. Of note, a sequence complementary to miR-215 seed (UCCAGU) is also present in RIPK1 mRNA. The functional significance of this observation is yet to be validated, but it opens up an intriguing possibility that elevated miR-215 expression accounts for an apparent discordance between RIPK1 mRNA and protein levels in 3B4.1 clone upon induction of the short form of RIPK1 [28].
Targeting of IKKβ by miR-497 may be viewed as yet another example of a negative feedback loop involving a miRNA and a pathway that regulates its expression. Indeed, it might be one of the contributors to the oscillatory waves of NF-κB activity, which are observed in some systems upon cytokine stimulation [49]. However, the consequences of specific reduction in the abundance of IKKβ are likely to be more complex. While the protein is essential for canonical NF-κB signaling, it is dispensable for the non-canonical pathway. Consequently, we would like to speculate that elevated expression of miR-497 would result not in a mere reduction of NF-κB activity, but in an altered propensity of a cell to activate one or the other branch of the NF-κB pathway. If the level of IKKα remains stable, while IKKβ declines, one may expect the cell to possess a larger pool of free IKKα and, hence, mount a stronger response toward non-canonical NF-κB stimuli and a weaker one towards the canonical ones. Hence, the type and the magnitude of NF-κB response may depend on the prior exposure of a cell to miR-497-inducing factors. Furthermore, a switch towards predominantly non-canonical response might ensue from re-challenging or prolonger treatment with the same cytokine. It would be interesting to explore whether the systems where a positive feedback loop is formed via NF-κB-dependent production of NF-κB-activating cytokines [30] also change the balance between the canonical and noncanonical pathways over time. Intriguingly, a gradual switch from canonical to non-canonical pathway has been well documented upon prolonged treatment with lymphotoxin-β [24]. The ability to reduce IKKβ levels may give miR-497 a beneficial role during aging, since selective loss of IKKβ is associated with reduced inflammation and a better health status in aging animals [50] , while the decline in miR-497 expression is seen in a mouse model of accelerated aging [51]. Of course, ultimately the physiological role of this potential regulatory mechanism should be proven using a real clinical model (chronic or acute inflammatory process, etc.) with an analysis of the endogenous NF-κB targets and corresponding biochemical and physiological changes.
It is noteworthy that NF-κB is not the only signal that co-regulates miR-497 and miR-146a. Both miRNAs belong to the clusters that are negatively regulated by Myc [52]. Their proposed tumor-suppressive properties, inferred from ectopic overexpression experiments [52], would be consistent with the role of negative regulators of NF-κB pathway, which is known to contribute to survival of cancer cells. The complex and coordinated control of these miRNAs could be taken as yet additional evidence of their significance for the cell fate and function, a topic worthy of further exploration.

Cell culture
3B4.1 and C6TA4 (including the derivatives that express mutant IκBα) have been described earlier [28]. Cell culture conditions were as described [53]. The cells were free of contamination by mycoplasma (MycoAlert test from Lonza Group Ltd.) and replication-competent retroviruses [54,55].
For IL-1β treatment, cells were plated at a density of 2 × 10 6 cells per 6cm plate and allowed to grow for about 16 hours before cytokine treatment. IL-1β was added at a concentration of 2 ng/mL. After IL-1β treatment, cells were trypsinized, washed, pelleted, and stored at -80°C.

microRNA profiling and sequencing
3B4.1 cells were cultured and treated with +/doxycycline. MiRNAs were prepared, sequenced and enumerated as described before [40]. miRNA sequencing data was normalized by scaling. Total number of reads for all miRNA transcript variants was tallied in each of the RNA samples and then counts for all transcript variants were multiplied by a factor to match the total number of counts in the RNA sample with the highest number of total counts (7, 742, 361). microRNA target prediction and pathway mapping miRNA target genes were predicted using the software TargetScan (http://www.targetscan.org). A spreadsheet of predicted target genes for their respective miRNA's was compiled using a custom R script. The compilation of predicted target genes was then mapped to independent cellular pathways using the KEGG Pathway Analysis tool (http://www.genome.jp/kegg/ pathway.html).

Quantitative PCR
RNA was isolated using mirVana miRNA Isolation Kit (Life Technologies, Grand Island, NY) following manufacturer protocol for Total RNA isolation without enrichment for microRNAs.
Primers and kits for reverse-transcription and quantitative-PCR were ordered. TaqMan Small RNA Assay kits were used for miRNAs 146a, 215, 381, 497, and RNU6B (Life Technologies). Quantitative PCR cycle threshold data was converted to change-fold in expression by the "delta delta Ct" method [56] using RNU6B as an internal control.
In order to construct pRSV-luc-IKBKB, a portion of the IKBKB 3′UTR was amplified from genomic DNA of LNCaP cells using the following primers: CGCACTTCTAGACGCCTGTCTGCACACTG and CCT TTCCTACAACCCGATTCCGGCCGGTCGATC. The PCR product was digested with FseI and XbaI and cloned between FseI and XbaI sites of pRSV-luc to create the final plasmid.
In order to construct pRSV-luc-mIKBKB, a two-step PCR strategy was used. First, separate PCR products were produced from pRSVluc-IKBKB template using two distinct pairs of primers (CTCATAAAGGCCAAGAAGGG and AGGT AAAAACACAATTTTCCGCGCAGTGAAATA, and TCTTTTTATTTCACTGCGCGGAAAATTGTGTTTT and TGTGATCATCTGAACTCATT). Second, the two PCR products were purified, combined and subjected to PCR with a new set of primers (CTCATAAAGGCCAAGAAGGG and TGTGATCAT CTGAACTCATT). The resulting product, along with pRSV-luc-IKBKB, was digested with XbaI and SpeI and used to replace the XbaI/SpeI fragment in pRSVluc-IKBKB.

Luciferase reporter assays
The cells were plated in 20 wells of a 96-well plate at a seeding density of 10,000 cells / well in 100 μL of antibiotic-free DMEM. 16 hours after plating, cells were transfected with miR-497 (1 pmol / well) or nontargeting siRNA (1 pmol / well), pRSV-βgal (constitutive β-galactosidase expression vector; 70 ng / well), and an appropriate luciferase reporter plasmid (30 ng / well) using Lipofectamine 2000 (Life Technologies) per manufacturer's protocol. Luciferase activity was measured and normalized for that of β-galactosidase as described before [57].

IKKβ detection by immunoblotting
500,000 cells/well of 6-well plate was plated overnight. Next day the cells were transfected with 100 nM of siGlo (fluorescently labeled negative RNAi control; GE Dharmacon, Lafayette, CO) or miR-497-5P using Lipofectamine 2000 reagent (Life Technologies). Typically, transfection rates of 80-90% were achieved. 48 hrs later the cells were lysed in RIPA buffer and protein concentrations were measured using DC Protein Assay (Bio-Rad). 30 ug of total proteins were then separated on 4-20% Mini-PROTEAN TGX Gel (Bio-Rad) and transferred to PVDF membrane. The membrane was blocked with 5% milk and hybridized with a rabbit polyclonal anti-IKKβ antibody (sc-7607; Santa Cruz Biotechnology, Dallas, TX), or with a mouse monoclonal anti-β-actin antibody as a loading control (A3854; Sigma-Aldrich, St. Louis, MO). After subsequent probing of the membrane with goat anti-rabbit IgG-HRP (sc-2004; Santa Cruz Biotechnology) and goat anti-mouse IgG-HRP (sc-2005; Santa Cruz Biotechnology), respectively, the proteins were visualized using Western Lighting PLUS-ECL reagents (Perkin Elmer, Waltham, MA).

ACKNOWLEDGMENTS
We would like to thank Drs. George Stark (Cleveland Clinic Foundation) and Andrei Bakin (Roswell Park Cancer Institute) for sharing cells and reagents.

CONFLICTS OF INTEREST
The authors declare no conflicts of interest.

GRANT SUPPORT
The work was supported in part by the funds from the Roswell Park Alliance Foundation. Core facilities at the Roswell Park Cancer Institute, which were used in this study, are funded by the NCI grant P30CA16056.