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The Journal of Immunology, 2007, 178: 3602-3611.
Copyright © 2007 by The American Association of Immunologists, Inc.

IFN Regulatory Factor-2 Regulates Macrophage Apoptosis through a STAT1/3- and Caspase-1-Dependent Mechanism1

Natalia Cuesta*, Quan M. Nhu*, Enrique Zudaire{dagger}, Swamy Polumuri*, Frank Cuttitta{dagger} and Stefanie N. Vogel2,*

* Department of Microbiology and Immunology, University of Maryland, School of Medicine, Baltimore, MD 21201; and {dagger} Cell and Cancer Biology Branch, National Cancer Institute, National Institutes of Health, Bethesda, MD 20892


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
IFN regulatory factor (IRF)-2–/– mice are significantly more resistant to LPS challenge than wild-type littermates, and this was correlated with increased numbers of apoptotic Kupffer cells. To assess the generality of this observation, and to understand the role of IRF-2 in apoptosis, responses of peritoneal macrophages from IRF-2+/+ and IRF-2–/– mice to apoptotic stimuli, including the fungal metabolite, gliotoxin, were compared. IRF-2–/– macrophages exhibited a consistently higher incidence of apoptosis that failed to correlate with caspase-3/7 activity. Using microarray gene expression profiling of liver RNA samples derived from IRF-2+/+ and IRF-2–/– mice treated with saline or LPS, we identified >40 genes that were significantly down-regulated in IRF-2–/– mice, including Stat3, which has been reported to regulate apoptosis. Compared with IRF-2+/+ macrophages, STAT3{alpha} mRNA was up-regulated constitutively or after gliotoxin treatment of IRF-2–/– macrophages, whereas STAT3beta mRNA was down-regulated. Phospho-Y705-STAT3, phospho-S727-STAT1, and phospho-p38 protein levels were also significantly higher in IRF-2–/– than control macrophages. Activation of the STAT signaling pathway has been shown to elicit expression of CASP1 and apoptosis. IRF-2–/– macrophages exhibited increased basal and gliotoxin-induced caspase-1 mRNA expression and enhanced caspase-1 activity. Pharmacologic inhibition of STAT3 and caspase-1 abolished gliotoxin-induced apoptosis in IRF-2–/– macrophages. A novel IFN-stimulated response element, identified within the murine promoter of Casp1, was determined to be functional by EMSA and supershift analysis. Collectively, these data support the hypothesis that IRF-2 acts as a transcriptional repressor of Casp1, and that the absence of IRF-2 renders macrophages more sensitive to apoptotic stimuli in a caspase-1-dependent process.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Innate immune responses are kept in check by specialized counterregulatory mechanisms. One mechanism involves apoptosis, which is induced in macrophages by bacterial toxins such as LPS (1, 2, 3). Apoptosis has been suggested as a mechanism for limiting the t1/2 of activated macrophages and restricting the expression of their inflammatory products at the site of tissue invasion (4, 5, 6). Two families of transcription factors contribute significantly to the regulation of host defense-related apoptosis, as follows: STATs and IFN regulatory factors (IRFs)3 (7, 8, 9, 10).

STATs are a family of latent cytoplasmic proteins that are involved in transmitting extracellular signals to the nucleus. Among the STATs, STAT3 has been shown to be involved in apoptosis (11, 12, 13, 14, 15). Three distinct isoforms of STAT3 have been identified, as follows: STAT3{alpha} (p92), the full-length isoform expressed in most cells; STAT3beta (p83), an alternatively spliced RNA form of STAT3{alpha} (16); and STAT3{gamma} (p72), a C-terminal truncated form of STAT3{alpha} derived posttranslationally through limited proteolysis (17). The ratio of the STAT3 isoforms appears to be related to the stage of cellular differentiation. STAT3{alpha} is more prevalent during early stages of granulocytic differentiation, whereas STAT3beta is present later during maturation (17, 18). Little is known about the function of STAT3{gamma}. STAT3 is activated through phosphorylation of Y705, leading to dimerization and nuclear translocation (19). STAT3 is often constitutively activated in cancers, where it functions as a critical mediator of oncogenic signaling through transcriptional activation of genes that encode inhibitors of apoptosis (e.g., Bcl-xL and survivin), cell cycle regulators (e.g., cyclin D1 and c-Myc), and inducers of angiogenesis (20, 21).

Another member of this family, STAT1, has also been implicated in modulating pro- and antiapoptotic genes (22). Kim and Lee (23) demonstrated that activation of STAT1 through serine phosphorylation by p38 MAPK modulates cell death in macrophages. Townsend et al. (24) linked STAT1-induced inhibition of cell growth and apoptosis to its ability to interact with p53.

The IRFs are also a large family of transcription factors that modulate the cellular response to IFNs and viral infection and have also been shown to modulate cell growth and apoptosis (25). IRF-1 and IRF-2 were first identified as activator and repressor, respectively, of the transcription of type I IFNs (26). Both IRF-1 and IRF-2 mRNAs are expressed at low constitutive levels in the cell, but the IRF-2 protein is more stable and accumulates in the nucleus and, thus, represses a number of promoters that are under the control of other IRF family members (27). When cells are stimulated by virus, IRF-1 is up-regulated and can compete with IRF-2 and stimulate transcription of many IFN-inducible genes (28). IRF-1 regulates DNA damage-induced cell cycle arrest (29, 30) and is involved in the regulation of apoptosis in several cell types (31, 32, 33). We suggested a role for IRF-2 in apoptosis after detecting greater numbers of apoptotic Kupffer cells in the livers of IRF-2–/– mice when compared with wild-type littermates (34). Recently, the absence of IRF-2 has been shown to cause premature apoptosis of NK cells (35); however, the mechanism(s) by which IRF-2 regulates apoptosis is unknown.

One of the apoptosis pathways triggered by activation of STATs and IRFs is caspase-1-dependent apoptosis (13, 30). Caspase-1 is the best-characterized inflammatory caspase, which is activated within an adapter complex termed the inflammasome (36, 37). Activated macrophages use caspase-1 (also known as IL-1-converting enzyme) to cleave pro-IL-1 and pro-IL-18 to their mature forms, although the role of caspase-1 in the induction of apoptosis seems to be independent of IL-1 or IL-18 secretion (38, 39). At least one binding site for IRFs (IFN-stimulated response element (ISRE)) has been identified in the 5' flanking region of CASP1 (40), and it has been proposed that Casp1 is regulated transcriptionally by up-regulation of IRF-1 and down-regulation of IRF-2 (32, 41). Another mechanism that has been reported to cause expression of caspase-1 and apoptosis depends on the activation of STAT1 (13).

In this study, we sought to dissect the molecular mechanisms that regulate the augmented response of macrophages derived from IRF-2–/– mice to apoptotic stimuli. We found that the lack of IRF-2 accelerates and enhances macrophage apoptosis in a caspase-1-dependent manner. The constitutive activation of STAT3 that is observed in IRF-2–/– cells appears to contribute to this process. Up-regulation of caspase-1 expression in the absence of IRF-2 suggests the role of IRF-2 as a transcriptional repressor. In support of this, we identified a novel ISRE in the Casp1 promoter by EMSA and supershift studies, and found it to bind strongly to IRF-2. Collectively, our data support a model in which IRF-2–/– macrophages are more sensitive to apoptotic stimuli due to an up-regulation of CASP1.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Mice

IRF-2–/– mice were originally obtained from T. Mak (Amgen Institute, Toronto, Canada) (42) and backcrossed to C57BL/6J for 5–7 generations. A second IRF-2–/– colony of mice, backcrossed to C57BL/6J for >20 generations, was used as well. The breeding pairs for this colony were a gift from G. Splitter (University of Wisconsin, Madison, WI). Mice were bred, as described previously (34), and each animal was genotyped by PCR. Six- to 8-wk-old mice were used in all experiments. All experiments were conducted with institutional approval.

Cells and reagents

Peritoneal exudate macrophages were collected 4 days after i.p. injection of 3% thioglycolate, and cultured in RPMI 1640 supplemented with 2% FBS, 100 U/ml penicillin, 100 µg/ml streptomycin, and 2 mM glutamine.

Gliotoxin from Gliocladium fimbriatum was purchased from Sigma-Aldrich. Escherichia coli K235 LPS was prepared using the method of McIntire et al. (43). Murine rIFN-{gamma} was provided by Genentech. Annexin V FITC was purchased from BD Biosciences. All phospho-specific Abs, anti-pY705 STAT3, anti-pS727 STAT3, anti-pY701 STAT1, anti-pS727 STAT1, anti-pT180/Y182-p38 MAPK, anti-pT202/Y204 ERK1/2, anti-pT183/Y185 JNK, as well as anti-Bcl-xL Abs were from Cell Signaling Technology. Anti-beta-actin, anti-total STAT3, anti-IRF-1, and anti-IRF-2 Abs were from Santa Cruz Biotechnology.

Caspase-3/7 activities were measured using a caspase assay kit (Apo-One Caspase-3/7 assay) from Promega. Activation of caspase-1 was measured using a colorimetric caspase-1 assay (R&D Systems). Results are expressed as fold increase in caspase activity of stimulated vs nonstimulated cells.

STAT3 inhibitor peptide was purchased from Calbiochem. The p38 inhibitor, SB203580, was obtained from Sigma-Aldrich. The caspase-1 inhibitor, Z-WEHD-FMK, was from R&D Systems. Each inhibitor was tested to insure its functionality, as follows: inhibition of caspase-1 activity was proven by means of the caspase-1 activation colorimetric assay; inhibition of p38 was proven by the decrease in STAT1 phosphorylation on S727 after exposure to the inhibitor; inhibition of STAT3 was proven by the decrease in mRNA gene expression of STAT3-regulated genes.

Mouse high-density oligonucleotide microarrays

Total RNA was isolated from liver samples, and 20 µg of RNA was converted to cDNA with StrataScript reverse transcriptase (FairPlay Microarray Labeling Kit; Stratagene), using oligo(dT)12–18 as a primer. cDNA was purified and labeled with Cy3 and Cy5 (Amersham Biosciences). The mixture of labeled cDNA probes was hybridized to a mouse oligonucleotide array on a glass slide (National Cancer Institute). Each slide contained 8272 murine genes and unnamed expressed sequence tags. Following hybridization, slides were air dried and scanned using a Genepix laser scanner (Molecular Devices). The images acquired were analyzed using GenePix Pro 3.0 software. The absolute feature (microarray spot) and background intensity of Cy3 and Cy5 for each feature on the array were obtained. The fluorescence intensity of each spot was calculated using the histogram quantitation method. After scanning, image acquisition, and normalization, genes that showed >3-fold (test/control) change were selected for further analyses. Genes were filtered to exclude those that were undetected on all arrays.

Real-time PCR

The relative expression of mRNA for each gene was determined by real-time PCR to confirm results of the microarray data. Total RNA was isolated from peritoneal macrophages using RNA STAT60 (Tel-Test). A total of 1 µg of RNA was used for oligo(dT)-primed cDNA synthesis (Promega reverse transcription system A3500). Real-time PCR was performed on an ABI Prism 7900HT Sequence Detection System (Applied Biosystems) using SYBR Green master mix (Applied Biosystems) and different sets of primers at a final concentration of 0.3 µM. mRNA gene expression profiles of the genes studied were normalized according to the hypoxanthine-guanine phosphoribosyltransferase (HPRT) concentration of each sample, and the fold increase was calculated using the 2–{Delta}{Delta}Ct method (44).

The following primer sequences were designed based on the Primer Express Software (Applied Biosystems): total STAT3 sense (5'-CCGTCTGGAAAACTGGATAACTTC-3'); total STAT3 antisense (5'-CCTTGTAGGACACTTTCTGCTGC-3'); STAT3{alpha} sense (5'-CAGGTAGTGCTGCCCCGTA-3'); STAT3{alpha} antisense (5'-CAGGTCAATGGTATTGCTGCA-3'); STAT3beta sense (5'-CGAAGCCGACCCAGGTAGT-3'); STAT3beta antisense (5'-AACTGCATCAATGAATGGTGTCA-3'); caspase-1 sense (5'-ATCTGTATTCACGCCCTGTTGG-3'); caspase-1 antisense (5'-CCCTCAGGATCTTGTCAGCC-3'); HPRT sense (5'-GCTGACCTGCTGGATTACATTAA-3'); HPRT antisense (5'-TGATCATTACAGTAGCTCT TCAGTCTGA-3').

Identification of candidate regulatory elements

Previous reports have suggested that specific transcriptional networks may be identifiable based on coordinate changes in gene expression (45). To identify potential cis elements of genes with similar patterns of expression after saline or LPS challenge of IRF-2–/– mice, we used the new version of the TraFac database, which can be found at http://trafac.cchmc.org/trafac/index.jsp.

Immunocytochemistry

Peritoneal macrophages were fixed in 10% buffered formalin (Sigma-Aldrich) for 10 min and permeabilized with 0.1% Triton X-100 in 0.1% sodium citrate. Endogenous peroxidase was blocked with 3% H2O2. Cells were exposed to 5% normal goat serum for 30 min and incubated overnight at 4°C with anti-pY705-STAT3 at a 1/100 dilution. Avidin-biotin histochemical staining was used for detection of primary Ab (46). Developing was performed using 0.5 mg/ml 3-3'diaminobenzidine tetrachloride (Vector Laboratories) and hydrogen peroxide as a substrate.

Western analysis

Macrophages were lysed in lysis buffer (100 mM Tris-HCl (pH 8.0), 50 mM NaF, 100 mM NaCl, 2 mM EDTA, 1% Triton X-100, 1 mM Na3VO4, 2 mM PMSF, and protease inhibitor mixture from Roche) for 10 min on ice. Twenty-five to 50 µg of protein was separated by 10% SDS-PAGE. Gels were transferred to a polyvinylidene difluoride membrane and placed in 5% nonfat milk in TBST for 1 h. Blots were incubated overnight with the primary Ab at 4°C and washed extensively with TBST. Following HRP-conjugated secondary Ab incubation, bound IgG was visualized using an ECL detection system (Amersham Biosciences). Gel bands were quantified using densitometry and image analysis (ImageJ 1.37 software, National Institutes of Health, which can be found at http://rsb.info.nih.gov/ij/) and normalized according to the beta-actin bands corresponding to each lane.

Apoptosis assay

For quantitative determination of apoptosis, flow cytometric DNA analysis was used (47). This method quantifies the percentage of apoptotic cells whose DNA content is lower than that of diploid cells. Cells were harvested by gentle scraping with a rubber policeman and then centrifuged at 1900 rpm for 10 min. The pellet was resuspended in 1 ml of hypotonic fluorochrome solution (50 µg/ml propidium iodide, 0.1% sodium citrate, 0.1% Triton-X-100). The red fluorescence (620 nm) of individual nuclei was measured by using a FACSort flow cytometer equipped with CellQuest acquisition software (BD Biosciences). All measurements were done under the same instrument settings, and at least 104 cells were measured in every sample. We previously showed that this method is directly comparable to results obtained by DNA fragmentation and annexin V labeling in murine macrophages (48). For annexin V FITC/propidium iodide staining, cells were washed twice in cold PBS and resuspended in binding buffer, according to manufacturer’s instructions. Annexin V FITC and propidium iodide were added to the solution at a final concentration of 1:20 and 5 µg/ml, respectively. Cells were incubated for 15 min at room temperature and analyzed by flow cytometry.

EMSA

Nuclear extracts from peritoneal macrophages were obtained using a nuclear extract kit from Active Motif. Oligonucleotides for Casp1 ISRE I (5'-ATGCTTTCAGTTTCAGTAGCTC-3' and complementary strand), Casp1 ISRE II (5'-TTAACTTTCTATTTTTTTAATT-3' and complementary strand), and negative control ISRE III (5'-CAGCTCTTTCTTTCTTGATGAC-3' and complementary strand) were annealed and 32P-end labeled with T4 polynucleotide kinase (Invitrogen Life Technologies). The negative control consisted of a sequence similar to the ISRE core (GAAANNGAA), but with mismatched bases (GAAAGAAAGAA). A total of 4 µg of nuclear extracts was incubated with the indicated DNA probe and 2 µg of poly(dI-dC) (Sigma-Aldrich) in a binding buffer containing 20 mM HEPES (pH 7.9), 4 mM MgCl2, 0.5 mM DTT, 0.1 mM EDTA, and 10% glycerol at room temperature for 10 min. DNA-protein complexes were resolved by electrophoresis in a 4% polyacrylamide gel. For supershift assays, 1 µg of anti-IRF-1 or anti-IRF-2 was also included in the reaction.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
IRF-2–/– macrophages exhibit increased apoptosis in response to several apoptotic stimuli

We have shown previously that IRF-2–/– Kupffer cells exhibit an aberrantly high apoptotic response both basally and after LPS or TNF-{alpha} challenge in vivo (34). Thus, we hypothesized that this dysregulation in apoptosis might be a general feature for macrophages deficient in the transcription factor IRF-2. IRF-2–/– vs IRF-2+/+ macrophage cultures were exposed to two different apoptosis inducers, as follows: purified gliotoxin from G. fimbriatum or a combination of E. coli LPS plus rIFN-{gamma}, which we demonstrated previously to synergize to induce macrophage apoptosis (48). We tested the effect of LPS alone as an inducer of apoptosis as well, but the effect was less obvious than when present concurrently with IFN-{gamma}. The percentage of subdiploid nuclei, characteristic of apoptotic cell populations, was measured by flow cytometry after staining with propidium iodide, as described by Nicoletti et al. (47) (Fig. 1A). Under basal conditions, IRF-2–/– macrophages displayed consistently higher levels of apoptosis (4.9 vs 2.0% in wild-type cells; p < 0.05). Treatment of cells with both apoptosis inducers led to more robust and accelerated apoptosis in IRF-2–/– macrophages. Fig. 1 confirms that gliotoxin (5 µg/ml) is a very potent inducer of apoptosis in both wild-type and IRF-2–/– macrophages (48). Macrophages from wild-type IRF-2+/+ mice, however, were significantly less sensitive to gliotoxin-induced apoptotic cell death than IRF-2–/– macrophages, as evidenced by a significantly greater number of diploid cells and a marked reduction of cells in the subdiploid fraction at both time points examined (~5 and ~51% apoptotic cells at 8 and 16 h, respectively, after gliotoxin exposure of wild-type macrophages vs ~25 and ~75% apoptosis in macrophages from IRF-2–/– mice at these same two time points). LPS plus IFN-{gamma} treatment of macrophages in vitro was also shown previously to induce macrophage apoptosis, but requires a longer time period than gliotoxin to elicit maximum apoptosis (48). Nonetheless, a similar trend was observed when macrophages were treated with 1 µg/ml LPS plus 50 U/ml rIFN-{gamma} for 48 h (24.7% apoptotic cells in IRF-2–/– macrophages vs 12% in wild-type cells; p < 0.005). The externalization of phosphatidylserine in the plasma membrane was also analyzed as a second approach to detect early events in apoptotic injury. Binding of annexin V to macrophages, concurrent with propidium iodide staining, confirmed the pattern of apoptosis that was observed with the method described by Nicoletti et al. (47) (Fig. 1B). Thus, compared with wild-type macrophages, IRF-2–/– macrophages exhibit increased basal and inducible apoptosis both in vivo (34) and in vitro.


Figure 1
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  		FIGURE 1. Quantitative determination of apoptosis by flow cytometric DNA analysis. Peritoneal macrophages from IRF-2+/+ and IRF-2–/– mice were treated with 5 µg/ml gliotoxin for 8 and 16 h, or 1 µg/ml LPS plus 50 U/ml IFN-{gamma} for 48 h. Bars represent the arithmetic means ± SE of the percentage of apoptotic cells with subdiploid nuclei from three independent experiments. *, Statistically significant difference compared with wild-type macrophages (n = 3; *, p < 0.05; **, p < 0.005; Student’s t test). A, Propidium iodide staining according to the method by Nicoletti et al. (47 ). B, Annexin V/propidium iodide staining.

 
Increased apoptosis in IRF-2–/– macrophages is caspase-3/7 independent

Because caspases fulfill critical roles in apoptosis in mammalian cells, we next tested the activity of two effector caspases, 3 and 7, in macrophages from IRF-2+/+ and IRF-2–/– mice in response to either gliotoxin (Fig. 2A) or LPS plus rIFN-{gamma} (Fig. 2B). A fluorometric assay was used to detect the cleavage and removal of a profluorescent DEVD peptide that is a specific substrate for caspase-3/7 activity (see Materials and Methods). Surprisingly, activation of these two effector caspases was significantly lower in macrophages from IRF-2–/– mice than in wild-type macrophages, suggesting that apoptosis in IRF-2–/– macrophages is unlikely to be attributable to an increase in caspase-3/7 activities.


Figure 2
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  		FIGURE 2. Measurement of caspase-3/7 activity in peritoneal macrophages from IRF-2+/+ and IRF-2–/– mice after gliotoxin treatment (5 µg/ml, 18 h) (A) or LPS plus IFN-{gamma} (1 µg/ml LPS, 50 U/ml IFN-{gamma}, 48 h) (B) (n = 2; *, p < 0.05; **, p < 0.005; Student’s t test).

 
Identification of genes in livers from IRF-2+/+ and IRF-2–/– mice that are differentially expressed and regulate apoptosis

The data presented to date indicate that the lack of IRF-2 has a profound impact on macrophage apoptosis both in vivo (34) and in vitro. To identify potential IRF-2 target genes, we used hybridization DNA microarrays to assess changes in RNA expression in IRF-2+/+ vs IRF-2–/– mice basally and after LPS treatment. Groups of three wild-type or IRF-2–/– mice at each of two time points, 3 or 6 h, were injected i.p. with saline or LPS (35 mg/kg; ~90% lethal dose in C57BL/6 mice). RNA was harvested from livers at 3 or 6 h after treatment. These time points were selected based on our previous observation that the number of apoptotic Kupffer cells was significantly higher in IRF-2–/– mice 6 h after LPS challenge (34). Two-color comparative hybridization experiments were used wherein wild-type and IRF-2–/– targets were cohybridized to oligonucleotide microarrays. Of the ~8000 known genes and expressed sequence tags monitored, 73 genes were down-regulated in IRF-2–/– mice that received saline (Supplemental4 Table Ia; GEO Series ID: GSE5907). Surprisingly, the lack of IRF-2 does not seem to have a large effect on the induction of the genes monitored, because no genes were identified as significantly up-regulated in IRF-2–/– mice. LPS administration significantly altered the pattern of gene expression in the liver, which was different in IRF-2–/– vs wild-type mice, as follows: 49 genes were down-regulated ≥3-fold in IRF-2–/– samples, and again, no genes were identified as up-regulated (Supplemental Table Ib; GEO Series ID: GSE5907).

Shared regulatory elements in genes whose expression is affected by LPS treatment in IRF-2–/– mice: STAT3{alpha} is up-regulated in IRF-2–/– macrophages

Using the TraFac database (http://trafac.cchmc.org/trafac/index. jsp), we identified consensus shared transcription factor binding sites within our list of genes (45). When comparing the genes that were down-regulated in IRF-2–/– mice basally or after LPS treatment, with respect to IRF-2+/+ mice, we identified several gene clusters that shared four to five cis elements, including ETS, STAT, and NKXH transcription factor binding sites. Interestingly, these regulatory elements are involved in the differentiation, maturation, and maintenance of the immune system (49, 50, 51). Most of these differentially expressed genes were found by TraFac to contain putative STAT binding sites, and therefore, could theoretically be affected by fluctuations in STAT protein levels as a result of IRF-2 deficiency. Among the genes identified as being down-regulated in the absence of IRF-2, we selected Stat3 to examine for its potential role in IRF-2-dependent apoptosis, given its already well-characterized role in apoptosis regulation (15, 52, 53).

Analysis of mRNA expression levels by quantitative real-time PCR revealed that total STAT3 mRNA expression profiles were significantly lower in IRF-2–/– macrophages than in wild-type cells both basally and in response to gliotoxin at all time points examined (Fig. 3A). These findings confirm the results observed in the microarray analysis after LPS challenge in vivo. However, when analyzing mRNA levels of the two splice variants of STAT3, we observed differential mRNA expression of STAT3{alpha} and STAT3beta in IRF-2+/+ and IRF-2–/– macrophages. STAT3{alpha} mRNA abundance in IRF-2–/– macrophages started out lower than the basal levels found in IRF-2+/+ macrophages; however, upon stimulation with gliotoxin, STAT3{alpha} was up-regulated in IRF-2–/– macrophages to approximately the same level induced by gliotoxin in wild-type macrophages (Fig. 3B). In contrast, STAT3beta mRNA remained down-regulated in macrophages from IRF-2–/– mice, both basally and after exposure to gliotoxin (Fig. 3C). Consistent with this observation, STAT3{alpha} and STAT3beta protein levels showed the same pattern when analyzed by Western analysis (Fig. 3D), with STAT3{alpha} protein levels being higher and STAT3beta protein levels being lower in IRF-2–/– macrophages, when compared with wild-type cells (densitometric analysis of blots revealed a 2-fold increase in STAT3{alpha} protein levels at 1 and 3 h after gliotoxin treatment in IRF-2–/– macrophages, and a 6-fold decrease in STAT3beta protein levels in basal conditions, plus a 1.5-fold decrease at 1 and 3 h after exposure to gliotoxin).


Figure 3
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  		FIGURE 3. Real-time PCR analysis of mRNA expression of total STAT3 (A), STAT3{alpha} (B), and STAT3beta (C) in IRF-2–/– macrophages compared with background-matched controls. Macrophages were treated with 5 µg/ml gliotoxin for 1–7 h. D, Western analysis of STAT3 expression in IRF-2–/– macrophages after stimulation with gliotoxin. beta-actin was used as a loading control. Results are representative of three independent experiments.

 
STAT3 is constitutively activated in IRF-2–/– macrophages

STAT3 activation is dependent upon phosphorylation of Y705, followed by homodimerization and nuclear translocation (19). An Ab specific for the tyrosine-phosphorylated form of STAT3 was used to investigate whether STAT3 was activated in peritoneal macrophages derived from IRF-2–/– mice. Wild-type macrophages showed no staining with anti-pY705-STAT3 (Fig. 4A). In contrast, despite the finding that total STAT3 mRNA was lower in IRF-2–/– macrophages, tyrosine-phosphorylated STAT3 was found in the nuclei of untreated IRF-2–/– macrophages (Fig. 4B), indicating that STAT3 is constitutively activated in IRF-2–/– macrophages that lack IRF-2. This was confirmed by Western analysis of whole cell lysates from peritoneal macrophages. Higher levels of pY705-STAT3 could be detected in IRF-2–/– macrophages under basal conditions (~4-fold increase in IRF-2–/– macrophages, according to densitometric analysis; Fig. 5, A and B), and the level of tyrosine phosphorylation decreased within 1 h after gliotoxin treatment, disappearing by 3 h (Fig. 5B). In contrast, wild-type macrophages showed fainter bands for activated STAT3 at all time points examined (Fig. 5, A and B).


Figure 4
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  		FIGURE 4. STAT3 is constitutively activated in macrophages from IRF-2–/– mice. Formalin-fixed macrophages from IRF-2+/+ (A) and IRF-2–/– (B) mice were stained with anti-pY705-STAT3 and counterstained with Harris hematoxylin. Data are representative of two independent experiments. Original magnification, x40 magnification.

 

Figure 5
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  		FIGURE 5. Western analysis of total protein extracts from IRF-2+/+ and IRF-2–/– macrophages. Macrophages were incubated for different periods of time with 5 µg/ml gliotoxin. Western analyses were performed using Abs against pY705-STAT3, pS727-STAT1, pT180/Y182-p38, pT202/Y204-ERK1/2, pT183/Y185-JNK, and beta-actin. A, Time course in minutes after gliotoxin exposure. B, Analysis of protein expression from 0 to 7 h of treatment with medium only (0 h) or gliotoxin. Results are representative of two independent experiments. C, Real-time PCR analysis of mRNA expression of the three following STAT-3-dependent genes: Bcl-xL, c-myc, and survivin.

 
Real-time PCR analysis of mRNA expression levels of STAT3-regulated genes (e.g., Bcl-xL, c-myc, and survivin) revealed that all of these genes were up-regulated in response to gliotoxin in IRF-2–/– vs IRF-2+/+ macrophages, further confirming the higher level of activated STAT3 in the IRF-2–/– macrophages (Fig. 5C).

Activation of the MAPK, ERK-1/2, was also more rapid in IRF-2–/– macrophages than in wild-type cells (Fig. 5B; especially compare 1-h time points, in which there is a difference of >5-fold in p-ERK protein levels, according to densitometric analysis), but no significant differences were found for JNK activation.

STAT1 and p38 MAPK activation are enhanced in IRF-2–/– macrophages

There is a close relationship between STAT3 and STAT1 in the regulation of apoptosis (54). STAT1 has been implicated in modulating expression of pro- and antiapoptotic genes following stress-induced responses (22). These effects are dependent on STAT1 phosphorylation at S727 by p38 MAPK (55), which is required for maximal transcriptional activity of STAT1 (56). Thus, we next sought to investigate the possible involvement of STAT1 in gliotoxin-induced apoptosis in IRF-2–/– macrophages. We examined the protein level of pY701- and pS727-STAT1 in IRF-2–/– macrophages and found that pS727-STAT1 levels were constitutively higher than in wild-type macrophages and significantly increased as early as 5 min after gliotoxin treatment of IRF-2–/– macrophages (densitometric analysis revealed a 2-fold increase in basal conditions, and a 5-fold increase in p-STAT1 levels at 5 min after gliotoxin treatment; Fig. 5A), and activation of STAT1 was sustained even after 7 h in IRF-2–/– macrophages (Fig. 5B). Wild-type cells responded to gliotoxin by activating STAT1 much more slowly (Fig. 5A) and failed to reach the activation levels detected in IRF-2–/– macrophages (Fig. 5, A and B). Concurrently, p38 MAPK activation was also higher in cells lacking IRF-2, as follows: significantly stronger bands were detected for phosphorylated p38 MAPK in IRF-2–/– macrophages at 5 min after exposure to gliotoxin (~10-fold increase over the p-p38 protein levels found in IRF-2+/+ macrophages; Fig. 5A). This stronger activation persisted until at least 7 h (Fig. 5B). Activation of phospho-p38 MAPK in wild-type macrophages only reached levels comparable to those seen in IRF-2–/– macrophages 3 h after gliotoxin treatment (Fig. 5B).

Casp1 gene expression is up-regulated, and caspase-1 activity is enhanced in IRF-2–/– macrophages

A key apoptosis-related gene that is regulated by STAT proteins is CASP1 (13). This gene has also been shown to be regulated transcriptionally by another member of the IRF family, IRF-1 (40). Because IRF-2 was originally demonstrated to counteract the activating effects of IRF-1 (26), we hypothesized that in the absence of IRF-2, IRF-1 might be responsible for heightened mRNA expression of Casp1. Real-time PCR analysis of macrophage RNA from IRF-2+/+ and IRF-2–/– mice showed significant up-regulation of Casp1 mRNA in IRF-2–/– macrophages, both basally and after gliotoxin treatment at all time points in IRF-2–/– macrophages (Fig. 6A). This finding was further confirmed by the finding that caspase-1 activity was significantly increased in IRF-2–/– macrophages 5 h after gliotoxin exposure, after which it dropped to baseline (Fig. 6B).


Figure 6
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  		FIGURE 6. Caspase-1 mRNA expression is up-regulated, and caspase-1 activity is enhanced in IRF-2–/– macrophages. A, Real-time PCR analysis of caspase-1 mRNA expression in IRF-2–/– macrophages vs background-matched controls. Cells were treated with 5 µg/ml gliotoxin for 1–7 h before RNA isolation. B, Caspase-1 activity in response to gliotoxin. Cells were treated with 5 µg/ml gliotoxin for 1–16 h and lysed, and cell lysates were tested for protease activity by the addition of a caspase-1-specific peptide, which is conjugated to a color reporter. Results are expressed as fold increase above the activity of IRF-2–/– macrophages. *, p < 0.05. C, Western analysis of Bcl-xL protein expression in control and IRF-2-deficient macrophages after stimulation with gliotoxin. beta-actin was used as a loading control. Results presented are representative of two independent experiments.

 
One way that caspase-1 contributes to apoptosis is by cleaving Bcl-xL, which converts it from a protective, antiapoptotic protein to a lethal protein (38). Western analysis of lysates from IRF-2+/+ and IRF-2–/– macrophages showed significant degradation of Bcl-xL in cells from IRF-2–/– mice 5 h after gliotoxin exposure (2-fold decrease in Bcl-xL protein levels in IRF-2–/– macrophages when compared with IRF-2+/+ cells; Fig. 6C), supporting our hypothesis that a higher level of caspase-1 activation in IRF-2–/– cells underlies their augmented levels of apoptosis.

Inhibition of caspase-1, STAT1, and STAT3 abolishes gliotoxin-induced apoptosis in IRF-2–/– macrophages

To inhibit STAT3 in macrophages, we pretreated macrophages with a cell-permeable dimerization-disrupting phosphopeptide that acts as a highly selective and potent blocker of STAT3 activation (57) before gliotoxin treatment of IRF-2+/+ and IRF-2–/– macrophages. To inhibit caspase-1 activity, we used a fluoromethyl ketone peptide that contains the amino acid sequence WEHD and binds preferentially to caspase-1. Fig. 7 shows that gliotoxin-induced apoptosis in IRF-2–/– macrophages was significantly decreased by pretreating cells with either the STAT3 inhibitory peptide or the caspase-1 inhibitor. It is interesting to note that whereas gliotoxin induced significant apoptosis in wild-type macrophages (as shown in Figs. 1 and 7), neither the caspase-1 inhibitor nor the STAT3 inhibitory peptide resulted in a significant decrease in the percentage of apoptotic cells in wild-type cells. Similarly, inhibition of p38 MAPK by pretreatment with SB203580 did not prevent gliotoxin-induced apoptosis in IRF-2+/+ macrophages, whereas it decreased the percentage of apoptotic cells in IRF-2–/– macrophages. Treatment of macrophages with the inhibitors alone did not alter the percentage of viable cells after 32 h in either cell population (data not shown).


Figure 7
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  		FIGURE 7. Effect of a STAT3 inhibitory peptide, a specific caspase-1 inhibitor (Z-WEHD-FMK), and a p38 MAPK inhibitor (SB203580) on gliotoxin-induced cell death in macrophages. IRF-2+/+ and IRF-2–/– macrophages were preincubated with 40 µM Z-WEHD-FMK, 50 µM STAT3 inhibitory peptide, 20 µM SB203580, or medium for 16 h. The cells were stimulated with gliotoxin for additional 16 h. The percentage of apoptotic cells was determined by propidium iodide staining and flow cytometric analysis. Bars represent the means ± SE of the percentage of apoptotic cells with subdiploid nuclei from three independent experiments. *, Statistically significant difference compared with medium-pretreated macrophages derived from the same strain of mice (n = 3; *, p < 0.05; Student’s t test). Differences are outside the limit of statistical significance (p = 0.059) for IRF-2–/– macrophages pretreated with SB203580.

 
A new ISRE site found in the Casp1 promoter

A computer search for potential ISREs in the murine Casp1 promoter region using Transplorer 1.4 (www.biobase.de) resulted in the identification of two nucleotide sequences that partially matched canonical ISREs (referred to as ISRE I for the more downstream site and ISRE II for the upstream site; Fig. 8A). These two sequences were also highly conserved in the human CASP1 promoter, especially in those nucleotides that define the putative ISREs. ISRE I was previously reported to bind IRFs (41), but the presence and functionality of ISRE II have not been described to date. EMSA was used to analyze the capacity of nuclear proteins derived from IRF-2+/+ and IRF-2–/– macrophages treated with medium only or with gliotoxin to bind to oligonucleotides that correspond to the sequences of ISRE I and ISRE II (Fig. 8B). Minimal binding was detected when ISRE I and the negative control oligonucleotide ISRE III were incubated with the nuclear extracts. In contrast, ISRE II formed complexes with nuclear proteins derived from murine macrophages, both constitutively and upon induction by gliotoxin. In addition, the absence of IRF-2 favors the formation of the nuclear complexes with ISRE II, because these bands are greatly enhanced in IRF-2–/– cells, without or with stimulation (Fig. 8B). Preincubation of the EMSA reactions for ISRE II with anti-IRF-1 Ab diminished the intensity of the bands in both IRF-2+/+ and IRF-2–/– macrophages, indicating the presence of IRF-1 in this complex (Fig. 8C). Abs against IRF-2 also produced a band with less intensity in IRF-2+/+ macrophages, making it almost disappear in medium-treated samples. This clear diminution of the basal band in IRF-2+/+ cells indicates that IRF-2 is a main component of this complex with ISRE II in basal conditions and forms part of the complex after gliotoxin treatment, in combination with other transcription factors, such as IRF-1. It also indicates that in the absence of IRF-2, binding of other transcription factors to ISRE II is enhanced, pointing to a possible role of IRF-2 as a transcriptional repressor of the CASP1 gene.


Figure 8
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  		FIGURE 8. Functional activity of two ISREs located in the promoter region of the gene that encodes caspase-1. A, Schematic representation of the sequence for the two ISREs found in the mouse Casp1 promoter and their conservation with those found in the human (ISREs I and II). Numbers represent their nucleotide position from the +1 transcription start site. ISRE III represents the negative control. B, EMSAs performed using probes containing the ISRE I and ISRE II from the murine Casp1 promoter and nuclear extracts from C57BL/6 (IRF-2+/+) and IRF-2–/– macrophages incubated with medium only (M) or gliotoxin (G) for 1 h. C, Supershift experiments on ISRE II in IRF-2+/+ and IRF-2–/– macrophages using IRF-1- and IRF-2-specific Abs, as indicated.

 

   Discussion
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 Abstract
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Disclosures
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Although IRF-2 has previously been related to apoptosis regulation (34, 35, 58), the mechanism still remains unclear. In this study, we show that the lack of IRF-2 renders macrophages significantly more sensitive to apoptotic stimuli, supporting our previous observations that Kupffer cells from IRF-2–/– mice are more sensitive to apoptosis basally and after LPS (34). Supporting the antiapoptotic function of IRF-2 presented in this work, Taki et al. (35) recently reported that bone marrow-derived NK cells from IRF-2–/– mice proliferated almost normally, but underwent accelerated apoptosis.

An important observation in our study is that the ratio of the STAT3{alpha} and STAT3beta isoforms is different in macrophages that lack IRF-2 compared with wild-type cells. Results from real-time PCR and Western analysis demonstrate a prevalence of STAT3{alpha} over the STAT3beta isoform in IRF-2–/– macrophages, both basally and in response to apoptotic stimuli. These findings suggest that IRF-2 participates in the generation of alternative splice variants of STAT3. In this regard, it has been reported that some transcriptional activators affect alternative splicing (59), although there are very few examples in the literature.

However, total STAT3 mRNA expression profiles were significantly lower in IRF-2–/– macrophages than in wild-type cells both basally and in response to gliotoxin (Fig. 3A). It has been reported that more forms of alternative splicing might exist for STAT3 (60), which might explain the significantly lower levels of total STAT3 mRNA in the absence of IRF-2.

Although STAT3beta was originally considered to be a negative regulator of transcription (16), it was later shown that the STAT3 isoforms {alpha} and beta have unique and specific functions (18). Both seem to be necessary for the correct balance of immune responses, because the specific ablation of STAT3beta impairs recovery from endotoxin shock and affects STAT3-dependent gene expression (61, 62). The STAT3beta isoform lacks the trans-activating domain and has been shown to inhibit apoptosis induced by ligation of MHC-I in Jurkat T cells (15). Therefore, the relative absence of STAT3beta in IRF-2–/– macrophages could possibly underlie the observed higher incidence of apoptosis.

STAT3 was found to be constitutively activated in IRF-2–/– macrophages. The relationship between STAT3 activation and apoptosis has been explored previously (53, 63, 64, 65, 66). For example, STAT3 has been shown to repress apoptosis by inhibiting caspase-3 and up-regulating Bcl-xL (14). Our results support and extend these findings: IRF-2–/– macrophages exhibit inhibition of caspase-3 activity and up-regulation of Bcl-xL mRNA in response to gliotoxin or LPS plus IFN-{gamma}. Our results further confirmed enhanced activation of STAT3 in IRF-2–/– macrophages as shown both by immunohistochemistry and by Western analysis and prompted us to investigate other caspase pathways that might be responsible for the higher incidence of cell death in these cells. Because blocking STAT3 activity protected IRF-2–/– macrophages from undergoing increased apoptosis after gliotoxin treatment, we conclude that STAT3 acts as a proapoptotic factor in the absence of IRF-2. Although STAT3 activation has been associated with proliferation, antiapoptosis, and cellular transformation (11, 12, 13, 15), there are a number of examples in which activated STAT3 appears to play a role in differentiation and promoting apoptosis (11, 67, 68, 69). Thus, STAT3 must work in concert with additional signaling pathways that dictate whether STAT3 will act as a pro- or antiapoptotic factor. This is consistent with our observation that the STAT3 inhibitory peptide did not significantly reduce gliotoxin-induced apoptosis in wild-type macrophages in contrast to IRF-2–/– cells. One of the mechanisms by which STAT3 may contribute to apoptosis is through the transcriptional regulation of CASP1. No STAT binding sites have been reported in the CASP1 promoter to date, but several studies point to their existence (13, 55, 70, 71).

The activation of STAT1 has been correlated with increased apoptosis in many cell types (22, 23, 72), and we report in this work that STAT1 activation on S727 was also augmented in IRF-2–/– macrophages in response to gliotoxin. STAT1 phosphorylation on S727 is mediated by p38 MAPK (73). In agreement with this result, significantly enhanced activation of p38 MAPK was detected in macrophages from IRF-2–/– mice in response to gliotoxin. Pretreatment of cells with a specific inhibitor of p38 MAPK did not have any effect on gliotoxin-induced cell death in wild-type cells, but it diminished the percentage of apoptotic cells in gliotoxin-treated IRF-2–/– macrophages, showing that the hyperactivation of p38 MAPK contributes to increased apoptosis detected in the absence of IRF-2. In vitro studies on the mechanisms by which STAT1 activation may trigger apoptosis have linked STAT1 to the induction of caspase-1 and Fas-Fas ligand (13, 55, 70, 71), but the precise mechanisms involved remain obscure.

Although up-regulation of caspase-1 in IRF-2–/– macrophages was demonstrated in our study by real-time PCR and a caspase-1 activation assay, the most compelling piece of evidence supporting the activation of caspase-1 is the observation that Bcl-xL is cleaved 5 h after gliotoxin treatment. Bcl-xL is a substrate for caspase-1, and its cleavage during the execution phase of cell death converts it into a potent prodeath molecule (38). Consistent with this model, pretreatment of macrophages with a specific caspase-1 inhibitor prevented the high incidence of gliotoxin-mediated apoptosis in macrophages that lack IRF-2, restoring them to untreated levels. Collectively, these data suggest that activation of caspase-1 is the main event that triggers the apoptosis response in these cells.

The studies reported in this work suggest that STAT and IRF transcription factors contribute to the transcriptional regulation of Casp1. Although the role of IRFs in the regulation of Casp1 gene expression has been explored previously (32, 41), this work sheds new light on the role of IRF-2 in the transcriptional regulation of this gene. Tamura et al. (30) first reported that CASP1 gene expression is up-regulated transcriptionally by IRF-1 in T lymphocytes. Horiuchi et al. (32) later demonstrated that the up-regulation of IRF-1 in vascular smooth muscle cells after serum deprivation induced an increase in caspase-1 mRNA, and they also observed that down-regulation of IRF-2 contributed to Casp1 expression and apoptosis. Iwase et al. (41) also published results suggesting that IRF-1 was able to activate the human CASP1 promoter through binding to an ISRE located in the initiator element of the gene, which we have designated ISRE I. By computer-assisted promoter analysis of the murine Casp1 and human CASP1 promoters, we identified another potential IRF binding site, designated ISRE II. EMSAs showed enhanced complex formation between ISRE II and nuclear proteins from IRF-2–/– macrophages, despite the fact that ISRE I was originally described as functionally relevant (41). These results could be interpreted as a higher binding of IRF-1 to the promoter sequence as would be expected in the absence of IRF-2. IRF-1 and IRF-2 have been shown to act as a transcriptional activator and repressor, respectively, of many genes (26, 28). The absence of IRF-2 would theoretically allow IRF-1 to bind to ISRE II and promote the transcription of CASP1, thus providing an explanation for the up-regulation of caspase-1 mRNA in IRF-2–/– macrophages. Therefore, we propose a model for the role of IRF-2 in apoptosis through inhibition of Casp1 transcriptional regulation (Fig. 9).


Figure 9
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  		FIGURE 9. Model of Casp1 transcriptional regulation, mediated by STAT and IRF members. STAT3 is activated by phosphorylation at Y705, and STAT1 by phosphorylation at Y701. This phosphorylation induces dimerization, nuclear translocation, and DNA binding. The ERK family of MAPKs phosphorylates STAT3 at S727. STAT1 is further phosphorylated through a p38 MAPK-dependent pathway at the same position. This second phosphorylation modulates transcriptional activation of STAT family members. In macrophages lacking IRF-2, STAT3 is constitutively activated and, therefore, may drive transcription of apoptosis-related genes, such as CASP1. At the same time, enhanced activation of p38 MAPK in IRF-2–/– macrophages in response to gliotoxin maintains higher levels of serine-phosphorylated STAT1, which may be responsible, in part, for the higher induction of caspase-1 expression. In contrast, binding of IRF-1 to ISRE II will be favored in the absence of the negative regulator IRF-2, thereby increasing caspase-1 expression and inducing apoptosis.

 

   Disclosures
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by National Institutes of Health Grant AI18797 (to S.N.V.). Back

2 Address correspondence and reprint requests to Dr. Stefanie N. Vogel, Department of Microbiology and Immunology, University of Maryland, 660 West Baltimore Street, Suite 324, Baltimore, MD 21201. E-mail address: svogel{at}som.umaryland.edu Back

3 Abbreviations used in this paper: IRF, IFN regulatory factor; HPRT, hypoxanthine-guanine phosphoribosyltransferase; ISRE, IFN-stimulated response element; Ct, cycle threshold. Back

4 The online version of this article contains supplemental material. Back

Received for publication September 5, 2006. Accepted for publication January 4, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

  1. Akira, S., K. Takeda, T. Kaisho. 2001. Toll-like receptors: critical proteins linking innate and acquired immunity. Nat. Immunol. 2: 675-680. [Medline]
  2. Takeda, K., S. Akira. 2005. Toll-like receptors in innate immunity. Int. Immunol. 17: 1-14. [Abstract/Free Full Text]
  3. Yokochi, T., A. Morikawa, Y. Kato, T. Sugiyama, N. Koide. 1998. Apoptotic cell death in response to LPS. Prog. Clin. Biol. Res. 397: 235-242. [Medline]
  4. Wesche, D. E., J. L. Lomas-Neira, M. Perl, C. S. Chung, A. Ayala. 2005. Leukocyte apoptosis and its significance in sepsis and shock. J. Leukocyte Biol. 78: 325-337. [Abstract/Free Full Text]
  5. Kim, S., E. Y. Chung, X. Ma. 2005. Immunological consequences of macrophage-mediated clearance of apoptotic cells. Cell Cycle 4: 231-234. [Medline]
  6. Xaus, J., M. Comalada, A. F. Valledor, M. Cardo, C. Herrero, C. Soler, J. Lloberas, A. Celada. 2001. Molecular mechanisms involved in macrophage survival, proliferation, activation or apoptosis. Immunobiology 204: 543-550. [Medline]
  7. Barber, G. N.. 2001. Host defense, viruses and apoptosis. Cell Death Differ. 8: 113-126. [Medline]
  8. Chawla-Sarkar, M., D. J. Lindner, Y. F. Liu, B. R. Williams, G. C. Sen, R. H. Silverman, E. C. Borden. 2003. Apoptosis and interferons: role of interferon-stimulated genes as mediators of apoptosis. Apoptosis 8: 237-249. [Medline]
  9. Bromberg, J., J. E. Darnell, Jr. 2000. The role of STATs in transcriptional control and their impact on cellular function. Oncogene 19: 2468-2473. [Medline]
  10. Calo, V., M. Migliavacca, V. Bazan, M. Macaluso, M. Buscemi, N. Gebbia, A. Russo. 2003. STAT proteins: from normal control of cellular events to tumorigenesis. J. Cell. Physiol. 197: 157-168. [Medline]
  11. Chapman, R. S., P. C. Lourenco, E. Tonner, D. J. Flint, S. Selbert, K. Takeda, S. Akira, A. R. Clarke, C. J. Watson. 1999. Suppression of epithelial apoptosis and delayed mammary gland involution in mice with a conditional knockout of Stat3. Genes Dev. 13: 2604-2616. [Abstract/Free Full Text]
  12. Ahmed-Choudhury, J., K. T. Williams, L. S. Young, D. H. Adams, S. C. Afford. 2006. CD40 mediated human cholangiocyte apoptosis requires JAK2 dependent activation of STAT3 in addition to activation of JNK1/2 and ERK1/2. Cell. Signal. 18: 456-468. [Medline]
  13. Chin, Y. E., M. Kitagawa, K. Kuida, R. A. Flavell, X. Y. Fu. 1997. Activation of the STAT signaling pathway can cause expression of caspase 1 and apoptosis. Mol. Cell. Biol. 17: 5328-5337. [Abstract]
  14. Haga, S., K. Terui, H. Q. Zhang, S. Enosawa, W. Ogawa, H. Inoue, T. Okuyama, K. Takeda, S. Akira, T. Ogino, et al 2003. Stat3 protects against Fas-induced liver injury by redox-dependent and -independent mechanisms. J. Clin. Invest. 112: 989-998. [Medline]
  15. Skov, S., M. Nielsen, S. Bregenholt, N. Odum, M. H. Claesson. 1998. Activation of Stat-3 is involved in the induction of apoptosis after ligation of major histocompatibility complex class I molecules on human Jurkat T cells. Blood 91: 3566-3573. [Abstract/Free Full Text]
  16. Caldenhoven, E., T. B. van Dijk, R. Solari, J. Armstrong, J. A. Raaijmakers, J. W. Lammers, L. Koenderman, R. P. de Groot. 1996. STAT3beta, a splice variant of transcription factor STAT3, is a dominant negative regulator of transcription. J. Biol. Chem. 271: 13221-13227. [Abstract/Free Full Text]
  17. Hevehan, D. L., W. M. Miller, E. T. Papoutsakis. 2002. Differential expression and phosphorylation of distinct STAT3 proteins during granulocytic differentiation. Blood 99: 1627-1637. [Abstract/Free Full Text]
  18. Maritano, D., M. L. Sugrue, S. Tininini, S. Dewilde, B. Strobl, X. Fu, V. Murray-Tait, R. Chiarle, V. Poli. 2004. The STAT3 isoforms {alpha} and beta have unique and specific functions. Nat. Immunol. 5: 401-409. [Medline]
  19. Darnell, J. E., Jr. 1997. STATs and gene regulation. Science 277: 1630-1635. [Abstract/Free Full Text]
  20. Garcia, R., T. L. Bowman, G. Niu, H. Yu, S. Minton, C. A. Muro-Cacho, C. E. Cox, R. Falcone, R. Fairclough, S. Parsons, et al 2001. Constitutive activation of Stat3 by the Src and JAK tyrosine kinases participates in growth regulation of human breast carcinoma cells. Oncogene 20: 2499-2513. [Medline]
  21. Wang, T., G. Niu, M. Kortylewski, L. Burdelya, K. Shain, S. Zhang, R. Bhattacharya, D. Gabrilovich, R. Heller, D. Coppola, et al 2004. Regulation of the innate and adaptive immune responses by Stat-3 signaling in tumor cells. Nat. Med. 10: 48-54. [Medline]
  22. Stephanou, A., D. S. Latchman. 2003. STAT-1: a novel regulator of apoptosis. Int. J. Exp. Pathol. 84: 239-244. [Medline]
  23. Kim, H. S., M. S. Lee. 2005. Essential role of STAT1 in caspase-independent cell death of activated macrophages through the p38 mitogen-activated protein kinase/STAT1/reactive oxygen species pathway. Mol. Cell. Biol. 25: 6821-6833. [Abstract/Free Full Text]
  24. Townsend, P. A., T. M. Scarabelli, S. M. Davidson, R. A. Knight, D. S. Latchman, A. Stephanou. 2004. STAT-1 interacts with p53 to enhance DNA damage-induced apoptosis. J. Biol. Chem. 279: 5811-5820. [Abstract/Free Full Text]
  25. Taniguchi, T., K. Ogasawara, A. Takaoka, N. Tanaka. 2001. IRF family of transcription factors as regulators of host defense. Annu. Rev. Immunol. 19: 623-655. [Medline]
  26. Harada, H., T. Fujita, M. Miyamoto, Y. Kimura, M. Maruyama, A. Furia, T. Miyata, T. Taniguchi. 1989. Structurally similar but functionally distinct factors, IRF-1 and IRF- 2, bind to the same regulatory elements of IFN and IFN-inducible genes. Cell 58: 729-739. [Medline]
  27. Watanabe, N., J. Sakakibara, A. G. Hovanessian, T. Taniguchi, T. Fujita. 1991. Activation of IFN-beta element by IRF-1 requires a posttranslational event in addition to IRF-1 synthesis. Nucleic Acids Res. 19: 4421-4428. [Abstract/Free Full Text]
  28. Harada, H., E. Takahashi, S. Itoh, K. Harada, T. A. Hori, T. Taniguchi. 1994. Structure and regulation of the human interferon regulatory factor 1 (IRF-1) and IRF-2 genes: implications for a gene network in the interferon system. Mol. Cell. Biol. 14: 1500-1509. [Abstract/Free Full Text]
  29. Tanaka, N., M. Ishihara, M. S. Lamphier, H. Nozawa, T. Matsuyama, T. W. Mak, S. Aizawa, T. Tokino, M. Oren, T. Taniguchi. 1996. Cooperation of the tumour suppressors IRF-1 and p53 in response to DNA damage. Nature 382: 816-818. [Medline]
  30. Tamura, T., M. Ishihara, M. S. Lamphier, N. Tanaka, I. Oishi, S. Aizawa, T. Matsuyama, T. W. Mak, S. Taki, T. Taniguchi. 1995. An IRF-1-dependent pathway of DNA damage-induced apoptosis in mitogen-activated T lymphocytes. Nature 376: 596-599. [Medline]
  31. Horiuchi, M., T. Yamada, W. Hayashida, V. J. Dzau. 1997. Interferon regulatory factor-1 up-regulates angiotensin II type 2 receptor and induces apoptosis. J. Biol. Chem. 272: 11952-11958. [Abstract/Free Full Text]
  32. Horiuchi, M., H. Yamada, M. Akishita, M. Ito, K. Tamura, V. J. Dzau. 1999. Interferon regulatory factors regulate interleukin-1beta-converting enzyme expression and apoptosis in vascular smooth muscle cells. Hypertension 33: 162-166. [Abstract/Free Full Text]
  33. Kano, A., T. Haruyama, T. Akaike, Y. Watanabe. 1999. IRF-1 is an essential mediator in IFN-{gamma}-induced cell cycle arrest and apoptosis of primary cultured hepatocytes. Biochem. Biophys. Res. Commun. 257: 672-677. [Medline]
  34. Cuesta, N., C. A. Salkowski, K. E. Thomas, S. N. Vogel. 2003. Regulation of lipopolysaccharide sensitivity by IFN regulatory factor-2. J. Immunol. 170: 5739-5747. [Abstract/Free Full Text]
  35. Taki, S., S. Nakajima, E. Ichikawa, T. Saito, S. Hida. 2005. IFN regulatory factor-2 deficiency revealed a novel checkpoint critical for the generation of peripheral NK cells. J. Immunol. 174: 6005-6012. [Abstract/Free Full Text]
  36. Petrilli, V., S. Papin, J. Tschopp. 2005. The inflammasome. Curr. Biol. 15: R581[Medline]
  37. Martinon, F., J. Tschopp. 2004. Inflammatory caspases: linking an intracellular innate immune system to autoinflammatory diseases. Cell 117: 561-574. [Medline]
  38. Clem, R. J., E. H. Cheng, C. L. Karp, D. G. Kirsch, K. Ueno, A. Takahashi, M. B. Kastan, D. E. Griffin, W. C. Earnshaw, M. A. Veliuona, J. M. Hardwick. 1998. Modulation of cell death by Bcl-xL through caspase interaction. Proc. Natl. Acad. Sci. USA 95: 554-559. [Abstract/Free Full Text]
  39. Tamura, T., S. Ueda, M. Yoshida, M. Matsuzaki, H. Mohri, T. Okubo. 1996. Interferon-{gamma} induces Ice gene expression and enhances cellular susceptibility to apoptosis in the U937 leukemia cell line. Biochem. Biophys. Res. Commun. 229: 21-26. [Medline]
  40. Tamura, T., M. Ishihara, M. S. Lamphier, N. Tanaka, I. Oishi, S. Aizawa, T. Matsuyama, T. W. Mak, S. Taki, T. Taniguchi. 1997. DNA damage-induced apoptosis and Ice gene induction in mitogenically activated T lymphocytes require IRF-1. Leukemia 11: 439-440.
  41. Iwase, S., Y. Furukawa, J. Kikuchi, S. Saito, M. Nakamura, R. Nakayama, J. Horiguchi-Yamada, H. Yamada. 1999. Defective binding of IRFs to the initiator element of interleukin-1beta-converting enzyme (ICE) promoter in an interferon-resistant Daudi subline. FEBS Lett. 450: 263-267. [Medline]
  42. Matsuyama, T., T. Kimura, M. Kitagawa, K. Pfeffer, T. Kawakami, N. Watanabe, T. M. Kundig, R. Amakawa, K. Kishihara, A. Wakeham, et al 1993. Targeted disruption of IRF-1 or IRF-2 results in abnormal type I IFN gene induction and aberrant lymphocyte development. Cell 75: 83-97. [Medline]
  43. McIntire, F. C., H. W. Sievert, G. H. Barlow, R. A. Finley, A. Y. Lee. 1967. Chemical, physical, biological properties of a lipopolysaccharide from Escherichia coli K-235. Biochemistry 6: 2363-2372. [Medline]
  44. Livak, K. J., T. D. Schmittgen. 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2–{Delta}{Delta} CT method. Methods 25: 402-408. [Medline]
  45. Jegga, A. G., A. Gupta, S. Gowrisankar, M. A. Deshmukh, S. Connolly, K. Finley, B. J. Aronow. 2005. CisMols analyzer: identification of compositionally similar cis-element clusters in ortholog conserved regions of coordinately expressed genes. Nucleic Acids Res. 33: W408-W411. [Abstract/Free Full Text]
  46. Hsu, S. M., L. Raine, H. Fanger. 1981. Use of avidin-biotin-peroxidase complex (ABC) in immunoperoxidase techniques: a comparison between ABC and unlabeled antibody (PAP) procedures. J. Histochem. Cytochem. 29: 577-580. [Abstract]
  47. Nicoletti, I., G. Migliorati, M. C. Pagliacci, F. Grignani, C. Riccardi. 1991. A rapid and simple method for measuring thymocyte apoptosis by propidium iodide staining and flow cytometry. J. Immunol. Methods 139: 271-279. [Medline]
  48. Lakics, V., S. N. Vogel. 1998. Lipopolysaccharide and ceramide use divergent signaling pathways to induce cell death in murine macrophages. J. Immunol. 161: 2490-2500. [Abstract/Free Full Text]
  49. Bassuk, A. G., J. M. Leiden. 1997. The role of Ets transcription factors in the development and function of the mammalian immune system. Adv. Immunol. 64: 65-104. [Medline]
  50. Akira, S.. 1999. Functional roles of STAT family proteins: lessons from knockout mice. Stem Cells 17: 138-146. [Abstract/Free Full Text]
  51. Hutton, J. J., A. G. Jegga, S. Kong, A. Gupta, C. Ebert, S. Williams, J. D. Katz, B. J. Aronow. 2004. Microarray and comparative genomics-based identification of genes and gene regulatory regions of the mouse immune system. BMC Genomics 5: 82[Medline]
  52. Bromberg, J.. 2000. Signal transducers and activators of transcription as regulators of growth, apoptosis and breast development. Breast Cancer Res. 2: 86-90. [Medline]
  53. Sommer, V. H., O. J. Clemmensen, O. Nielsen, M. Wasik, P. Lovato, C. Brender, K. W. Eriksen, A. Woetmann, C. G. Kaestel, M. H. Nissen, et al 2004. In vivo activation of STAT3 in cutaneous T-cell lymphoma: evidence for an antiapoptotic function of STAT3. Leukemia 18: 1288-1295. [Medline]
  54. Stephanou, A., B. K. Brar, R. A. Knight, D. S. Latchman. 2000. Opposing actions of STAT-1 and STAT-3 on the Bcl-2 and Bcl-x promoters. Cell Death Differ. 7: 329-330. [Medline]
  55. Stephanou, A., T. M. Scarabelli, B. K. Brar, Y. Nakanishi, M. Matsumura, R. A. Knight, D. S. Latchman. 2001. Induction of apoptosis and Fas receptor/Fas ligand expression by ischemia/reperfusion in cardiac myocytes requires serine 727 of the STAT-1 transcription factor but not tyrosine 701. J. Biol. Chem. 276: 28340-28347. [Abstract/Free Full Text]
  56. Decker, T., P. Kovarik. 2000. Serine phosphorylation of STATs. Oncogene 19: 2628-2637. [Medline]
  57. Turkson, J., J. S. Kim, S. Zhang, J. Yuan, M. Huang, M. Glenn, E. Haura, S. Sebti, A. D. Hamilton, R. Jove. 2004. Novel peptidomimetic inhibitors of signal transducer and activator of transcription 3 dimerization and biological activity. Mol. Canc