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PPAR1 attenuates cytosol to membrane translocation of PKC to desensitize monocytes/macrophages

Institute of Biochemistry I, Faculty of Medicine, Johann Wolfgang Goethe University, 60590 Frankfurt, Theodor-Stern-Kai 7, Germany

Correspondence to Andreas von Knethen: v_knethen{at}zbc.kgu.de

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Recently, we provided evidence that PKC depletion in monocytes/macrophages contributes to cellular desensitization during sepsis. We demonstrate that peroxisome proliferator–activated receptor (PPAR) agonists dose dependently block PKC depletion in response to the diacylglycerol homologue PMA in RAW 264.7 and human monocyte–derived macrophages. In these cells, we observed PPAR-dependent inhibition of nuclear factor-B (NF-B) activation and TNF- expression in response to PMA. Elucidating the underlying mechanism, we found PPAR1 expression not only in the nucleus but also in the cytoplasm. Activation of PPAR1 wild type, but not an agonist-binding mutant of PPAR1, attenuated PMA-mediated PKC cytosol to membrane translocation. Coimmunoprecipitation assays pointed to a protein–protein interaction of PKC and PPAR1, which was further substantiated using a mammalian two-hybrid system. Applying PPAR1 mutation and deletion constructs, we identified the hinge helix 1 domain of PPAR1 that is responsible for PKC binding. Therefore, we conclude that PPAR1-dependent inhibition of PKC translocation implies a new model of macrophage desensitization.

Abbreviations used in this paper: 15d-PGJ₂, 15-deoxy-^12,14-prostaglandin J₂; AF, activating function; CHX, cycloheximide; DAG, diacylglycerol; DBD, DNA-binding domain; DGK, DAG kinase ; EMSA, electrophoretic mobility shift assay; HEK, human embryonic kidney; IL, interleukin; LBD, ligand-binding domain; MCS, multicloning site; NF-B, nuclear factor-B; PPAR, peroxisome proliferator–activated receptor ; ROS, reactive oxygen species.

	Introduction

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Monocyte/macrophage desensitization is characteristic for late-phase immune responses (Liew et al., 2005). Confined proinflammatory cytokine expression and mediator synthesis is important to avoid pathological settings, such as sepsis or atherosclerosis (Hotchkiss and Karl, 2003; Hansson, 2005). Down-regulating proinflammatory cytokine expression (TNF-, interleukin [IL]-1ß, and IFN) or proinflammatory mediator release (nitric oxide and reactive oxygen species [ROS]) concomitantly switches the proinflammatory phenotype toward an antiinflammatory one. The latter is characterized by the synthesis of antiinflammatory cytokines, such as TGF-ß or IL-10, and is often accompanied by cellular desensitization upon secondary proinflammatory stimulation (Docke et al., 1997; Kalechman et al., 2002). Therefore, the identification of molecular mechanisms contributing to cellular desensitization attracted growing interest (Docke et al., 1997; von Knethen and Brune, 2002).

One factor attenuating proinflammatory gene expression is peroxisome proliferator–activated receptor (PPAR). PPAR is a nuclear hormone receptor that, upon agonist binding, transactivates gene expression as a heterodimer bound to retinoic acid receptor- (Abdelrahman et al., 2005). Its role in blocking proinflammatory gene expression comprises several options, mainly antagonizing signaling cascades. Specifically, PPAR negatively regulates transcription factors by scavenging transcriptional coactivators, such as the cAMP-response element–binding protein or the steroid receptor coactivator-1 (Yang et al., 2000). However, a direct association with the transcription factors NF-B, NF of activated T cells, signal transducer, and activator of transcription or NF-E2–related factor 2 (Ikeda et al., 2000; Wang et al., 2001, 2004; Chung et al., 2003) blocks their recruitment to responsive elements in promoter structures of target genes. Recently, it has been shown that PPAR is targeted to nuclear receptor corepressor–histone deacetylase-3 complexes in response to ligand-dependent SUMOylation (Pascual et al., 2005), protecting these complexes from proteosomal degradation. Normally, histone deacetylase-3 removes a corepressor complex, provoking expression of proinflammatory genes. Additionally, PPAR represses activation of a mitogen-activated protein kinase, which keeps downstream transcription factors unphosphorylated and, consequently, inactive (Desreumaux et al., 2001). Moreover, PPAR influences the cell cycle by up- regulating p21 expression, which is an established cell cycle inhibitor (Han et al., 2004), or down-regulating phosphatase PPA2, which is known to adjust E2F/DP DNA-binding activity, which is necessary for the G₁ to S-phase transition (Altiok et al., 1997). In response to proinflammatory stimulation, PPAR-dependent gene transcription also contributes to cellular desensitization. PPAR agonists inhibit diacylglycerol (DAG)–PKC signaling by inducing DAG kinase- (DGK) expression (Verrier et al., 2004). This enzyme lowers the amount of DAG, which is an established PKC activator. Normally, DAG is released from membrane lipids and activates classical PKCs (Liu and Heckman, 1998). Based on gene induction of DGK as the underlying mechanism, this type of desensitization demands at least 6–15 h. Thus, it appears that PPAR transrepresses proinflammatory gene expression, often in a DNA-unbound state, by provoking direct protein–protein interactions.

We provide evidence for a new PPAR-dependent mechanism in blocking PKC signaling. Depletion of PKC is attenuated by PPAR1 activation in RAW 264.7 cells or human primary monocyte–derived macrophages. Cytosolic localization of PPAR1 interferes with PKC cytosol to membrane translocation, which is a prerequisite for its activation-dependent depletion. Translocation is restored in cells transfected with a dominant-negative PPAR1 mutant. Coimmunoprecipitation studies and a mammalian two-hybrid system revealed a direct PPAR1–PKC interaction as the underlying mechanism. PPAR1 deletion constructs support the idea that ligand-dependent PPAR activation is necessary for PKC binding, which is mediated by the helix 1 of the PPAR1 hinge domain. Our data suggest a new mechanism for how activation of PPAR1 blocks PKC translocation, thereby achieving cellular desensitization.

	Results

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PPAR agonists inhibit PKC depletion
Recent data demonstrate that monocyte/macrophage desensitization in response to phagocytosis of apoptotic cells is achieved by attenuating PKC signaling, which blocks NADPH oxidase–dependent formation of ROS (Johann et al., 2006). Therefore, we were interested in identifying molecular mechanisms interfering with PKC depletion. A potential candidate known to affect the pro- versus antiinflammatory phenotype in monocytes/macrophages is PPAR. Because controversial data exist concerning its expression in monocytic and macrophage cell lines, as well as in primary human monocytes and macrophages, we performed a first set of experiments determining PPAR expression in the monocytic cell lines and primary cells under investigation. As shown in Fig. 1 A, PPAR is constitutively expressed in murine RAW 264.7 macrophages. In contrast, in THP-1 cells, PPAR is only fractionally expressed, but differentiation toward macrophages with 100 nM PMA for 24 h provoked up-regulation of PPAR (Fig. 1 A, lane 2 vs. 3). A similar expression pattern is observed in primary monocytes and macrophages, respectively. PPAR is only marginally expressed in monocytes, but induced upon differentiation toward macrophages (Fig. 1 B). To identify the expressed PPAR isoform 1 or 2, we performed a Western blot using human PPAR1-transfected human embryonic kidney (HEK) cells as a positive control. Taking into consideration that murine and human PPAR1 are identical in size (475 aa), we conclude that PPAR1 is expressed in RAW 264.7 macrophages, differentiated THP-1 cells, and primary macrophages (unpublished data). Based on these results, we choose RAW 264.7 cells, differentiated human THP-1 cells, and primary monocyte–derived macrophages as experimental cell models.

View larger version (43K): [in this window] [in a new window] [Download PPT slide]   Figure 1. PPAR expression in monocytes/macrophages. (A) PPAR expression was determined in lysates of RAW 264.7 macrophages and in control versus differentiated THP-1 cells. For differentiation, cells were treated for 24 h with 100 nM PMA. Western blot was performed as described in Materials and methods. (B) PPAR expression was analyzed by Western analysis in primary human monocytes and macrophages, differentiated for 7 d with medium containing serum of AB-positive donors. Experiments were performed at least three times, and representative data are shown.

View larger version (24K): [in this window] [in a new window] [Download PPT slide]   Figure 2. PPAR agonist prestimulation inhibits PKC depletion. RAW 264.7 macrophages (A) and PPAR1 AF2–overexpressing RAW 264.7 cells (B) were prestimulated for 1 h with ciglitazone (1 or 10 µM), rosiglitazone (1 or 10 µM), or remained as controls, followed by the addition of 100 nM PMA for 1 h. (C) RAW 264.7 macrophages were stimulated for 1 h with 50 µg/ml CHX. Thereafter, 10 µM of rosiglitazone or ciglitazone were added for 1 h, followed by 100 nM PMA stimulation for 1 h. (D) Primary monocyte–derived macrophages were prestimulated for 1 h with 10 µM ciglitazone, 10 µM rosiglitazone, or remained as controls. Afterward, 100 nM PMA was added for 1 h. For all experiments, cells were harvested and lysed, and Western blot was performed as described in the Materials and methods. Experiments were performed at least three times, and representative data are shown.

The physiological significance of these results obtained in murine RAW 264.7 macrophages was verified in primary human monocyte–derived macrophages isolated from peripheral blood. Similar to RAW 264.7 cells, in primary macrophages, pretreatment with ciglitazone and rosiglitazone preserved PKC expression upon PMA addition (Fig. 2 D).

Antiinflammatory consequences of PPAR1–PKC interaction
To elucidate whether the PPAR1–PKC interaction shows an impact on PKC signaling in inflammatory gene expression in macrophages, we analyzed two proinflammatory markers of macrophage activation, i.e., NF-B DNA binding and TNF- expression in response to PMA in RAW 264.7 macrophages. To determine activation of the proinflammatory transcription factor NF-B, we performed a set of electrophoretic mobility shift assays (EMSAs), demonstrating the DNA-binding capability of the transcription factor. As shown in Fig. 3 A, 100 nM PMA supplied for 3 h significantly induced NF-B activation (Fig. 3 A, second lane) compared with the untreated control (Fig. 3 A, first lane). To elucidate the composition of the transcription factor complex, we used antibodies against the p50 (Fig. 3 B, left) and p65 subunits (Fig. 3 B, right) of NF-B. As shown in Fig. 3 B (left), the lower and the upper NF-B shifts contained the p50 subunit. Therefore, the two bands were significantly reduced when an -p50 antibody was included in the binding reaction and a new band, the p50 supershift, occurred. Only the upper NF-B shift included the p65-subunit, as indicated by the addition of the -p65 antibody, which provoked the reduction of the upper NF-B shift, but did not alter the lower NF-B shift (Fig. 3 B, right). As expected, a new band was detectable (the p65 supershift). Thus, we conclude that the lower NF-B shift is formed by a p50 homodimer, whereas the upper NF-B shift consists of a p50/p65 heterodimer.

View larger version (17K): [in this window] [in a new window] [Download PPT slide]   Figure 3. PPAR1-dependent attenuation of PKC translocation provokes a reduction of the proinflammatory response in macrophages. (A) Activation of NF-B in RAW 264.7 macrophages in response to PMA. RAW 264.7 macrophages were stimulated for 3 h with 100 nM PMA or left untreated as control. Afterward, cells were harvested, nuclear extracts were isolated, and NF-B EMSA was performed as described in the Materials and methods. (B) Supershift analysis of the active NF-B complex was described in the Materials and methods. Macrophages were stimulated with 100 nM PMA for 3 h. For supershift analysis, a p50 antibody (left, second lane) or a p65 antibody (right, second lane) was included. NF-B activation without antibody addition (left and right, first lane) is shown. (C) 15d-PGJ2 inhibited PMA-mediated NF-B activation. RAW 264.7 cells were pretreated with 10 µM of the endogenous PPAR ligand 15-dPGJ2 for 1 h, followed by the addition of 100 nM PMA for 3 h (middle lane). To ensure a PPAR-dependent effect, one sample was prestimulated before 15d-PGJ2 addition with 10 µM of the PPAR antagonist GW9662 for 1 h (right lane). One sample remained PMA treated only as a control (left lane). Cells were harvested, nuclear protein extracts were isolated, and NF-B EMSA was performed as described in Materials and methods. (D) PMA-mediated NF-B activation is inhibited in human primary macrophages in response to 15d-PGJ2. Primary monocyte–derived macrophages were treated as described in C. Cells were harvested and processed, and NF-B EMSA was performed as described in Materials and methods. (E) Inhibition of PMA-mediated PKC activation by PPAR reduced proinflammatory TNF- expression. RAW 264.7 cells were treated for 1 h with 10 µM rosiglitazone or remained as controls. Afterward, cells were incubated with 100 nM PMA for 6 h, and TNF- expression in the cell supernatant was analyzed using the CBA system. All experiments were performed at least three times. Data are the means ± the SD of the individual experiments (*, P < 0.05) or representative of three similar experiments.

PPAR1-dependent inhibition of PKC translocation
Considering that activation of PKC, followed by its translocation to the cell membrane, is a prerequisite for its depletion, we were interested to determine whether PPAR blocks PKC translocation. To follow PPAR and PKC distribution in RAW 264.7 cells, we stained for PPAR and PKC in paraformaldehyde-fixed cells (Fig. 4). As shown in Fig. 4 A (third panel), PPAR localizes in the cytosol and the nucleus in untreated cells, whereas PKC is localized in the cytosol (Fig. 4 A, second panel). The nucleus is counterstained, using DAPI (Fig. 4 A, first panel), and an overlay is provided in Fig. 4 A (fourth panel). To prove specificity of the secondary antibodies used, which were labeled with either Alexa Fluor 488 or 546, we used these antibodies alone without a first antibody. In both cases, no signal is observed (unpublished data). Activation of the cells with 100 nM PMA for 50 min provokes PKC translocation (Fig. 4 B, second panel), whereas localization of PPAR is not altered (Fig. 4 B, third panel). Pretreatment of RAW 264.7 macrophages with 10 µM of the synthetic PPAR agonist rosiglitazone for 1 h prevents PKC translocation in response to 100 nM PMA stimulation for 50 min (Fig. 4 C, second panel). Localization of PPAR remains unaltered (Fig. 4 C, third panel). To prove a PPAR-dependent effect, we used the PPAR-specific antagonist GW9662. Preincubation of the cells for 1 h with 10 µM GW9662, followed by rosiglitazone treatment (1 h, 10 µM), restores PKC translocation after 100 nM PMA addition for 50 min (Fig. 4 D, second panel). PPAR localization was not affected (Fig. 4 D, third panel). From these data, we conclude that activated cytosolic PPAR in RAW 264.7 macrophages inhibits PKC translocation in response to 100 nM PMA. Based on the aforementioned Western blot results, RAW 264.7 cells express isoform 1, which is partially located in the cytosol.

View larger version (54K): [in this window] [in a new window] [Download PPT slide]   Figure 4. PKC and PPAR localization in RAW 264.7 macrophages. To follow PKC and PPAR localization in RAW 264.7 macrophages, cells were seeded on slides and treated (B) for 50 min with 100 nM PMA, (C) preincubated with 10 µM rosiglitazone for 1 h followed by 100 nM PMA addition for 50 min, or (D) pretreated with 10 µM GW9662 before cells were stimulated as described in C. Afterward, cells were fixed and stained for PKC and PPAR as described in the Materials and methods. Cell nuclei were counterstained with DAPI. DAPI staining is shown in the first panel, PKC staining in the second, PPAR staining in the third, and an overlay to estimate cytosolic and nuclear region is provided in the fourth panel. All experiments were performed three times, and representative data are shown.

View larger version (52K): [in this window] [in a new window] [Download PPT slide]   Figure 5. PPAR1 inhibits PKC translocation. HEK293 cells were cotransfected with DsRed-PPAR1 wild type/PKC-EGFP (A and B [top two rows] and C and D) or DsRed-PPAR1 AF2/PKC-EGFP (A and B, bottom two rows). To follow PKC-EGFP translocation, 100 nM PMA was added to control cells (A, second and fourth row) or cells pretreated for 1 h with 10 µM rosiglitazone (B, second and fourth row). To verify the role of PPAR on PKC-EGFP translocation in DsRed-PPAR1 wild type/PKC-EGFP, cotransfected HEK293 cells were treated for 1 h with 10 µM of the PPAR antagonist GW9662 before stimulation for 1 h with 10 µM rosiglitazone (C, top row) followed by 50 min of 100 nM PMA addition (C, bottom row) or preincubated for 1 h with 10 µM of the PPAR agonist WY14643 (D, top row) before activation with 100 nM PMA for 50 min (D, bottom row). Cell nuclei were counterstained with DAPI. DAPI staining is shown in the first panel, PKC-EGFP in the second, DsRed-PPAR in the third, and an overlay to estimate cytosolic and nuclear region is provided in the fourth panel. All experiments were performed three times, and representative data are shown.

PPAR1 directly binds to PKC
To elucidate whether PPAR1 inhibits PKC translocation by a direct PPAR1–PKC interaction, we performed a set of coimmunoprecipitation experiments. Immunoprecipitation of PKC from lysates of differentiated THP-1 cells, which had been stimulated for 1 h with rosiglitazone or left untreated, was conducted. As shown in Fig. 6 A, immunoprecipitation of PKC resulted in coimmunoprecipitation of PPAR1 in THP-1 cells that had been challenged with a PPAR agonist (Fig. 6 A, lane 2). In the flowthrough, PPAR1 was only detected when agonist stimulation was omitted (Fig. 6 A, lane 1). After PPAR1 activation, PPAR1 was almost completely retarded in the immunoprecipitation column.

View larger version (33K): [in this window] [in a new window] [Download PPT slide]   Figure 6. PPAR1 directly interacts with PKC. (A) THP-1 cells were differentiated for 24 h with 50 nM PMA. To allow new synthesis of PKC, which is depleted in response to the differentation regime, cells were further cultured for 48 h in normal medium. Afterward, cells were treated for 1 h with 10 µM rosiglitazone or remained as controls. Cells were harvested and lysed, and PKC was immunoprecipitated as described in Materials and methods. Eluates and flowthroughs were separated by Western blotting and stained for PPAR and PKC as indicated. (B) COS-7 cells were transiently cotransfected with PPAR1 wild type/PKC-EGFP or PPAR1 AF2/PKC-EGFP. 24 h later, cells were treated for 1 h with 10 µM rosiglitazone or remained as controls. Cells were harvested and lysed, and PKC-EGFP was immunoprecipitated as described in Materials and methods. Input controls, eluates, and flowthroughs were separated by Western blotting and stained for PPAR and PKC as indicated. All experiments were performed at least three times, and representative data are shown.

To provide further evidence for a direct PPAR1–PKC interaction, we used the mammalian two-hybrid system. In COS-7 cells transiently transfected by electroporation with a combination of pCMV-AD-PPAR1, pCMV-BD-PKC, and the Gal4 reporter vector pFR-luc, addition of rosiglitazone or ciglitazone provoked induction of luciferase expression as determined by a luciferase assay. As shown in Fig. 7, addition of both PPAR agonists induce luciferase expression roughly threefold compared with untreated controls. A PPAR-dependent effect was verified because addition of the PPAR agonist WY14643 left basal luciferase activity unaltered. With this two-hybrid model, direct binding of target (PPAR1) to bait protein (PKC) is required to induce luciferase expression. Therefore, our data suggest that PPAR1 directly binds PKC upon agonist activation. This interaction inhibits PKC translocation to the cell membrane, and thus, PKC activation.

View larger version (26K): [in this window] [in a new window] [Download PPT slide]   Figure 7. PPAR directly binds to PKC. COS-7 cells were transiently transfected with a combination of a target (PPAR1), a bait (PKC), and a reporter construct, as described in Materials and methods. Afterward, cells were treated with 10 µM ciglitazone, 10 µM rosiglitazone, 10 µM WY14643, or remained as controls. 6 h later, cells were harvested and lysed for a reporter analysis as described in Materials and methods. Experiments were performed at least three times in duplicate. *, P < 0.05. Data are the means ± the SD.

Identification of PPAR1 domains involved in PKC binding

View larger version (40K): [in this window] [in a new window] [Download PPT slide]   Figure 8. Helix 4 of the LBD/AF2 domain does not mediate PPAR binding to PKC. (A) Scheme of the PPAR1 constructs. (B and C) HEK293 cells were cotransfected with DsRed-PPAR1 wild type/PKC-EGFP (B, first panel), DsRed-PPAR1 L309A/PKC-EGFP (B, second panel), DsRed-PPAR1 N310A/PKC-EGFP (B, third panel), DsRed-PPAR1 G312A/PKC- EGFP (B, fourth panel), PPAR1 V313A/PKC-EGFP (C, first panel), PPAR1 L316A/PKC-EGFP (C, second panel), PPAR1 K317A/PKC-EGFP (C, third panel) or DsRed-PPAR1 aa309-319/PKC-EGFP (C, fourth panel). To follow PKC-EGFP localization, 24 h after transfection, cells were treated for 50 min with 100 nM PMA (second row), for 1 h with 10 µM rosiglitazone (third row), pretreated for 1 h with 10 µM rosiglitazone followed by the addition of 100 nM PMA for 50 min (fourth row) or remained as controls (first row). Experiments were performed three times, and representative data are shown.

Based on these results, we decided to generate three PPAR1 deletion constructs (DsRed-PPAR1 aa32-198, DsRed- PPAR1 aa32-250, and DsRed-PPAR1 aa51-406) with the belief that ligand binding is necessary for PPAR1–PKC interactions. As shown in Fig. 9 A, all deletions lack the DNA-binding domain (DBD) of PPAR1. Furthermore, to characterize the role of the hinge domain in PKC binding, it was eliminated to variable extents. In the DsRed-PPAR1 aa32-198 construct, the first 26 aa of the hinge domain were deleted, and in the DsRed-PPAR1 aa32-250 construct, 78 aa of the hinge domain were deleted. The hinge domain was completely removed in the DsRed-PPAR1 aa51-406 construct. In this construct, a part of the LBD/AF2 domain was deleted as well (aa288-406). All constructs lack a part of the AF1 domain.

View larger version (47K): [in this window] [in a new window] [Download PPT slide]   Figure 9. Hinge domain mediates PPAR1 binding to PKC. (A) Scheme of the PPAR1 constructs. (B) HEK293 cells were transiently transfected with one of the PPAR1 constructs, as indicated. 24 h after transfection cells were lysed. Western blotting was performed, and blots were stained for DsRed. All experiments were performed at least three times, and representative data are shown. (C) HEK293 cells were transfected with PKC-EGFP only (first panel) or cotransfected with DsRed-PPAR1 wild type/PKC-EGFP (second panel), DsRed-PPAR1 32-198/PKC-EGFP (third panel), DsRed-PPAR1 32-250/PKC-EGFP (fourth panel), or DsRed-PPAR1 51-406/PKC-EGFP (fifth panel). To follow PKC-EGFP localization, 24 h after transfection, cells were treated for 50 min with 100 nM PMA (second row), treated for 1 h with 10 µM rosiglitazone (third row), pretreated for 1 h with 10 µM rosiglitazone followed by the addition of 100 nM PMA for 50 min (fourth row), or remained controls (first row). Experiments were performed three times, and representative data are shown.

To elucidate the role of these deletions on PKC translocation, HEK293 cells were transiently cotransfected with the shortened DsRed-monomer–tagged PPAR1 constructs in combination with a PKC-EGFP–encoding vector. To follow PKC translocation, 100 nM PMA was added to (1 h, 10 µM) rosiglitazone-pretreated cells. PKC localization was documented in untreated cells (Fig. 9 C, first row), cells treated for 50 min with PMA (Fig. 9 C, second row), for 1 h with rosiglitazone (Fig. 9 C, third row), or preincubated for 1 h with rosiglitazone, followed by the addition of PMA for 50 min (Fig. 9 C, fourth row). In cells transfected with the DsRed-tagged PPAR1 aa32-198 construct, PKC-EGFP did not translocate to the cell membrane. However, in cells expressing the DsRed-tagged PPAR1 aa32-250 or aa51-406 construct, PKC translocated to the cell membrane in response to 100 nM PMA.

From these data, we conclude that for PKC, binding a part of the hinge domain of PPAR1 is indispensable. To further narrow the involved region of PPAR1, we finally created the construct DsRed-PPAR1 aa206-224 (Fig. 10 A), containing a deletion of helix 1 (aa206-224) of PPAR1, which is located in the hinge domain (aa173-288). Helix 1 has already been identified to mediate the protein–protein interaction of PPAR with ERK5 (Akaike et al., 2004). Expression of the construct results as expected in protein, demonstrating a slightly reduced protein mass (Fig. 10 B, lane 2) because of the aa206-224 deletion compared with the DsRed-PPAR1 wild type (Fig. 10 B, lane 1). We transiently cotransfected HEK cells with the PPAR1 aa206-224 construct tagged with DsRed-monomer in combination with a PKC-EGFP–encoding vector. In cells expressing the DsRed-tagged PPAR1 aa206-224 (Fig. 10 C), PKC translocated to the cell membrane in response to 100 nM PMA.

View larger version (25K): [in this window] [in a new window] [Download PPT slide]   Figure 10. Hinge helix 1 mediates PPAR1 binding to PKC. (A) Scheme of the PPAR1 construct. (B) HEK293 cells were transiently transfected with the DsRed-PPAR1 wild type as control or the DsRed-PPAR1 aa206-224 construct as indicated. 24 h after transfection, cells were lysed. Western blotting was performed, and blots were stained for DsRed. (C) HEK293 cells were cotransfected with DsRed-PPAR1 aa206-224/PKC-EGFP. 24 h after transfection, cells were treated for 1 h with 10 µM rosiglitazone. To follow PKC-EGFP translocation, 100 nM PMA was added to cells, and localization of PKC-EGFP was examined 50 min thereafter. Experiments were performed three times and representative data are shown.

	Discussion

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Recently, we demonstrated that monocyte/macrophage desensitization at least partially attenuates PKC signaling (von Knethen et al., 2005; Johann et al., 2006). We provide evidence that PPAR agonists block PKC translocation to the cell membrane and concomitant protein depletion, which normally occurs after cell activation. In monocytic cell lines, PPAR expression has been previously described (McIntyre et al., 2003; Musiek et al., 2005; von Knethen et al., 2005), and it was verified using primary human monocyte–derived macrophages. These data corroborate the work of Tontonoz et al. (1998) and Chinetti et al. (1998), showing PPAR expression in differentiated macrophages. However, even if PPAR is expressed, PPAR agonists are known to mediate PPAR-dependent and -independent effects (Nosjean and Boutin, 2002). To this end, 15d-PGJ₂ has been described to directly modify H-ras, provoking a constitutively active enzyme (Oliva et al., 2003) or inhibiting I-B kinase, and thus suppressing NF-B signaling (Straus et al., 2000). Our approach, using cells expressing PPAR1 wild type or the PPAR1 agonist-binding mutant AF2, substantiates the need of PPAR activation in our system. Only in cells expressing PPAR1 wild type was translocation of PKC blocked by PPAR activation. The PPAR1 AF2 mutant did not prevent PMA-mediated PKC translocation. These data support the notion of a PPAR-dependent mechanism.

PPAR-mediated inhibition of classical PKCs has been previously described (Verrier et al., 2004). In their case, PKCß translocation was blocked by PPAR agonists via DGK up-regulation. DGK metabolizes DAG, which is an established activator of classical and novel PKC isoforms. Therefore, its induction/activation will remove the potential PKC activator, causing desensitization as seen in our experiments. However, in our experiments, a role of DGK up-regulation must be excluded because the protein-synthesis inhibitor CHX did not restore PKC translocation. In line with this, our PPAR1 aa32-198 construct, where the PPAR1 DBD was removed, still inhibits PKC translocation. Further support for our hypothesis, suggesting a direct PPAR1–PKC interaction in preventing PKC translocation, came from previous studies (Johann et al., 2006). In this case, PPAR was activated in response to apoptotic cells, attenuating PKC translocation and concomitant ROS production. In this study, the role of PPAR was verified using a PPAR d/n cell line. In these cells, pretreatment with apoptotic cells left PMA-mediated PKC translocation and subsequent ROS production unaltered. A premise for this assumption is that PPAR is expressed at least partially in the cytosol. Generally, the nuclear hormone receptor PPAR is described to be exclusively localized in the nucleus (Akiyama et al., 2002; Feige et al., 2005). In support of our hypothesis, suggesting cytoplasmatic localization as well, we noticed a minor amount of PPAR1 to remain in the cytosol. This is based on results using DsRed-PPAR1–transfected cells, as well as immunohistochemical detection of endogenous PPAR1 located in the cytosol of RAW 264.7 macrophages besides its major nuclear localization. It should be noted that cytoplasmatic distribution of PPAR is in line with the work of Abella et al. (2005). In their study, an approach similar to our experiments was used, with EGFP-tagged PPAR used to characterize intracellular distribution of PPAR. Results indicated that PPAR is not exclusively located in the nucleus. Furthermore, localization of PPAR in the cytoplasma in the promonocytic cell lines HL-60 and K-562 has been observed, especially in response to the PPAR agonist troglitazone (Liu et al., 2005). This work was done using immunohistochemical detection of endogenous PPAR. Therefore, side effects, such as unphysiological high expression or a modified protein behavior as a result of a tag or label (Feige et al., 2005), can be excluded. In addition, Burgermeister et al. (2006) recently provided evidence that PPAR is actively exported from the nucleus into the cytosol in a MEK1-dependent manner, further supporting our observed PPAR localization pattern. Furthermore, Patel et al. (2005) described cytoplasmatic localization of a different PPAR isoform, PPAR, when coexpressed with CAP350, which is a putative centrosome-associated protein of unknown function. Therefore, we propose that members of the PPAR family may localize in the cytoplasm, possibly after activation, when bound to cytoplasmic proteins such as PKC. Immunoprecipitation of PKC from lysates of differentiated THP-1 cells coimmunoprecipitated PPAR. Remarkably, PPAR1 coimmunoprecipitation was only seen once PPAR1 became activated. The requirement of PPAR1 activation was verified using an agonist-binding mutant of PPAR1, which did not block PKC translocation in response to PMA stimulation. A direct PPAR1–PKC interaction was further supported by a mammalian two-hybrid system with PPAR1 as the target and PKC as the bait construct, provoking luciferase reporter gene expression when target and bait proteins interact. To avoid autocrine activation of the reporter system, PPAR has to be cloned as a target protein linked to the NF-B transactivation domain, not allowing this hybrid protein to bind to the promoter of the reporter. However, DNA binding of PPAR1 to PPREs, and concomitant scavenging the NF-B-AD-PPAR1 hybrid protein from the two-hybrid assay, cannot be excluded.

Based on the well-established role of helix 4 of the PPAR LBD in mediating protein–protein interaction of PPAR with coactivators, such as CBP and SRC-2, or repressors, such as the nuclear receptor corepressor and the silencing mediator for retinoic acid receptor and thyroid-hormone receptor (Nolte et al., 1998; Westin et al., 1998; Perissi et al., 1999; Perissi and Rosenfeld, 2005), we first generated 6 PPAR1 constructs in which only 1 aa was exchanged and 1 construct in which helix 4 was completely removed. Unexpectedly, these constructs did not alter rosiglitazone-dependent inhibition of PKC translocation.

Taking into account that PPAR binding to other factors, such as adipocyte-type fatty acid–binding protein or extracellular signal-related kinase 5, which do not belong to the family of transcriptional coactivators, can be mediated by other PPAR domains, such as A/B/C and D/E/F (Adida and Spener, 2006) or the hinge domain (domain D; Akaike et al., 2004), we created three PPAR1 deletion constructs. All of them lack the entire DBD (domain C). In addition, different parts of the A/B and D domains have been removed, and one construct contained the C-terminal third of the E/F domains only. Based on our collective results, we provide evidence that a part of the hinge domain probably confers the PPAR1–PKC interaction, which is present in the PPAR1 aa32-198 construct but absent in the aa32-250 construct, when PPAR1 is activated by an agonist, thus requiring the LBD/AF2 domains. One known region of PPAR1 located in aa198-250 is the hinge helix 1 (aa 206–224). Therefore, we cloned a PPAR1 construct with helix 1deleted (DsRed- PPAR1 aa206-224). In cells transfected with this construct, PKC translocated even after rosiglitazone pretreatment in response to PMA. From these results, we conclude that PPAR1 binds to PKC via the hinge helix 1 domain, after PPAR1 has been activated by a ligand.

The proposed mechanism of PPAR1–PKC binding proceeds fast. 1 h of prestimulation with PPAR agonists is sufficient to inhibit PKC translocation in response to 100 nM PMA. However, PKC translocation by 1 µM PMA was not blocked. These results support the assumption that the capacity of cytoplasmatic PPAR to bind PKC correlates with the strength of PKC activation. Likely, very strong activation signals, such as 1 µM PMA, exceed the inhibitory impact of PPAR. Thus, the role of PPAR in blocking PKC signaling might be only transient, allowing PKC activation by a more stringent activator. This makes the mechanism more interesting for the development of new therapy strategies. Prolonged periods of PPAR activation, which provoke transcriptional control to target members of the NADPH oxidase system, have already been described (p22^phox, p47^phox, and gp91^phox; Inoue et al., 2001; von Knethen and Brune, 2002; Hwang et al., 2005). Consequently, in these cells PPAR contributes to an antiinflammatory phenotype by blocking NADPH oxidase-dependent ROS production.

An involvement of PPAR in attenuating inflammatory reactions to improve the clinical picture of sepsis has previously been shown (for review see Zingarelli and Cook, 2005). In line with this, our results add to this data. In our system, PMA-mediated NF-B activation was inhibited in response to PPAR agonist pretreatment to 50% in RAW 264.7 cells, as well as primary human macrophages. In accordance, PMA-induced TNF- expression was PPAR dependently reduced to 70%. It has been observed that PPAR activation inhibits multiple organ failure in an animal model (Abdelrahman et al., 2005), although the underlying mechanism remains unclear. The option to adjust a pro- versus antiinflammatory monocyte/macrophage phenotype will provide new possibilities for the development of therapies to control systemic inflammation. Our data add a new antiinflammatory role for PPAR based on the ability to scavenge PKC in the cytosol, thus, blocking membrane translocation and downstream signaling.

	Materials and methods

Top Abstract Introduction Results Discussion Materials and methods References

 
Monocyte isolation
We analyzed human cells from peripheral blood of healthy donors. For monocyte enrichment, we isolated PBMCs from donors using Ficoll-Hypaque gradients (PAA Laboratories). Cells were left to adhere on culture dishes (Primaria 3072; Becton Dickinson) for 60 min at 37°C. Nonadherent cells were removed. Afterward, cells were differentiated to macrophages by culturing them in complete RPMI containing 10% AB-positive human serum. Flow cytometry confirmed that the monocyte-like population was 90–95% pure (CD14⁺ vs. CD14^–).

Cell culture
We cultivated RAW 264.7 and THP-1 in RPMI 1640 (PAA Laboratories). HEK293 and COS-7 cells were cultured in DME high glucose (PAA Laboratories). Both media were supplemented with 100 U/ml penicillin (PAA Laboratories), 100 µg/ml streptomycin (PAA Laboratories), and 10% heat-inactivated fetal calf serum (PAA Laboratories). Ciglitazone (Biomol), rosiglitazone (Biomol), WY14643 (Biomol), and CHX (Sigma-Aldrich) were dissolved in DMSO. Appropriate vehicle controls were performed.

Immunofluorescence staining
To determine intracellular PPAR localization, we seeded RAW 264.7 macrophages directly on a slide. After 24 h, cells were treated as indicated and fixed on the slides by 1-h incubation in 4% paraformaldehyde at 4°C. Thereafter, cells were permeabilized in PBS containing 0.2% Triton X-100 for 15 min. After a washing step in PBS, cells were incubated for 2 h with a 1:250 dilution of a rabbit -PPAR antibody (Calbiochem) at 4°C. After three 5-min washing steps with PBS, cells were incubated with a secondary goat -rabbit antibody (1:250) labeled with Alexa Fluor 546 (Invitrogen) for 2 h at 4°C. Cells were incubated for 2 h with a 1:250 dilution of a mouse -PKC antibody (BD Biosciences) at 4°C. After three 5-min washing steps with PBS, cells were incubated with a secondary goat -mouse antibody (1:250) labeled with Alexa Fluor 488 (Invitrogen) for 2 h at 4°C. Again, cells were washed three times with PBS and counterstained with DAPI (1 µg/ml in PBS for 15 min). After a final 5-min washing step in PBS, cells were covered with Vectashield mounting medium (Linaris) and a coverslip. PPAR and PKC localization were determined using an AxioScope fluorescence microscope with the ApoTome upgrade (Carl Zeiss MicroImaging, Inc.; lens 63x/0.6 NA; ocular 10x) at room temperature, documented by a charge-coupled device camera (Carl Zeiss MicroImaging, Inc.) and AxioVision Software (Carl Zeiss MicroImaging, Inc.).

Vector construction, transient transfection, fluorescence microscopy, and reporter analysis
To examine cellular PPAR localization, we subcloned human PPAR1 into the DsRed-monomer–encoding vector pDsRed-Monomer-C1 (CLONTECH Laboratories, Inc.) using the infusion ligation kit (CLONTECH Laboratories, Inc.). To allow integration of the PPAR1 fragment, the vector was cut within the multicloning site (MCS) by BamHI and XhoI. To insert PPAR1 (provided by V.K.K. Chatterjee, University of Cambridge, Cambridge, UK), we used the pcDNA3-PPAR1 wild-type and AF2 vectors for PPAR1 amplification by PCR, using the following sequences based on the infusion ligation requirements (changed nucleotides are underlined): wild type, 5'-GGACTCAGATCTCGAATGGTTGACACAGAGATC GCATTCTG-3' and 3'-AGGACGTCCTCTAGATGTTCCTGAACATGCTAGGTGGCCT AGA T-5'; AF2 mutant, 5'-GGACTCAGATCTCGAATGGTTGACACAGAGATCGCAT- TCTG-3' and 3'-GAGACGTCCGCTAGATGTTCCTGAACATGCTAGGTGGCCT AGAT-5'. Annealing temperatures were 62°C for the first cycle and 72°C for the later ones and calculated using the Oligo software (MBI). Infusion reaction of the cleaved vector with the amplified PPAR1 wild-type or AF2 fragment was performed according to the distributor's instructions.

Site-directed mutagenesis to generate single aa exchanges (L309A, N310A, G312A, V313A, L316A, K317A) and deletion of helix 1 (aa206-224) or 4 (aa309-319) of PPAR1 were performed using the QuikChange XLII kit (Stratagene). The following primers were used (changed nucleotides are underlined): L309A, 5'-CCTGGTTTTGTAAATCTTGACGCGAACGACCAAGTAACTCTCCTC-3' and 5'-GAGGAGAGTTACTTGGTCGTTCGCGTCAAGATTTACTTTTCCAGG-3'; N310A, 5'-CC TGGTTTTGTAAATCTTGACTTGGCGGACCAAGTAACTCTCCTC-3' and 5'-GAGGAGAG TTACTTGGTCCGCCAAGTCAAGATTTACTTTTCCAGG-3'; G312A, 5'-GTAAATCTTG ACTTGAACGACGCGGTAACTCTCCTCAAA- TATGG-3' and 5'-CCATATTTGAGGAGAGT TACCGCGTCGTTCAAGTC- AAGATTTAC-3'; V313A, 5'-GTAAATCTTGACTTGAACGA CCAAGCGACTCTCCTCAAATATGG-3' and 5'-CCATATTTGAGGAGAGTCGCTTGGTCG- TTCAAGTCAAGATTTAC-3'; L316A, 5'-CTTGAACGACCAAGTAACTCTC- GCGAAAT ATGGAGTCCACGAG-3' and 5'-CTCGTGGACTCCATATTT- CGCGAGAGTTACTTGGTCG TTCAAG-3'; K317A, 5'-CTTGAACGACCAAGTAACTCTCCTCGCGTATGGAGTCCAC GAG-3' and 5'-CTCGTGGACTCCATACGCGAGGAGAGTTACTTGGTCGTTCAAG-3'; aa309-319, 5'-CCTGGTTTTGTAAATCTTGACCCGCTGACCAAAGCAAAG-3' and 5'-CTTT GCTTTGGTCAGCGGGTCAAGATTTACAAAACCAGG-3'. The pcDNA3-PPAR1 wild-type vector was used as a template. An initial denaturation step was performed at 95°C for 1 min, followed by 18 cycles at 95°C for 50 s, annealing at 60°C for 50 s, and extension at 68°C for 7 min. A final extension phase was performed at 68°C for 7 min.

DsRed-PPAR1 aa32-198 was constructed by deleting the EcoRV fragment in the DsRed-PPAR1 wild-type vector. DsRed-PPAR1 aa32-250 was constructed by deleting the EcoRV–EcoRI fragment in the DsRed-PPAR1 wild-type vector, blunting the sticky EcoRI end before religating the remaining plasmid. Finally, DsRed-PPAR1 aa51-406 was constructed by deleting the XmnI fragment in the DsRed-PPAR1 wild-type vector. Restriction enzymes were obtained from New England Biolabs. The Klenow fragment and T4 ligase were provided by Fermentas. All manipulations did not alter the open reading frame of PPAR1.

Correct orientation and sequence of the generated vectors was verified by restriction analyses and/or sequencing. The PKC-EGFP signaling sample (pPKC-EGFP) used was obtained from CLONTECH Laboratories, Inc.

To follow PKC translocation and PPAR distribution, HEK293 cells were seeded directly onto a slide, and then transiently transfected by CaPO₄-precipitation with combinations of pDsRed-Monomer-C1 PPAR1 wild type/pPKC-EGFP, pDsRed-Monomer-C1 PPAR1 AF2/pPKC-EGFP, or the generated deletion and mutation constructs together with pPKC-EGFP. 24 h after transfection, cells were used for experiments. Cells were treated as indicated. Afterward, cells were fixed on the slides by 1-h incubation in 4% paraformaldehyde at 4°C. Cells were washed three times with PBS and counterstained with DAPI (1 µg/ml in PBS for 15 min). After a final 5-min washing step in PBS, cells were covered with Vectashield mounting medium and a coverslip. Translocation of PKC-EGFP and DsRed-PPAR1 wild type/AF2 distribution was analyzed using an AxioScope fluorescence microscope with the ApoTome upgrade (lens 63x/0.6 NA; ocular 10x) at room temperature, documented by a charge-coupled device camera and the AxioVision Software.

For reporter analysis, HEK293 cells were transiently transfected by CaPO₄-precipitation with pDsRed-Monomer-C1 PPAR1 wild-type, -AF2, aa32-198, 32-250, 51-406 constructs, or the empty DsRed vector in combination with the PPRE-containing p(AOX)₃-TK-luc reporter plasmid. Transfection efficiency was normalized by cotransfecting a pRL-TK control vector encoding for Renilla reniformis luciferase. Transfections were performed in duplicate, and each experiment was repeated at least three times.

Coimmunoprecipitation
After THP-1, cells were differentiated for 24 h with 50 nM PMA, PMA was removed, and cells were incubated for an additional 48 h in complete medium. Afterward, cells were stimulated for 1 h with 10 µM rosiglitazone or remained as controls. Eventually, cells were harvested and lysed in lysis buffer (50 mM Tris, 5 mM EDTA, 150 mM NaCl, 0.5% Nonidet-40, and 1 mM PMSF, pH 8.0). To assure cell lysis, cells were sheared 10 times with a 16-gauge needle, followed by a brief 10-s sonication (Sonifier; Branson; duty cycle 100%, output control 60%). Cell debris was removed by centrifugation (10,000 g for 5 min), and 1 mg of protein was used for immunoprecipitation. Sample volume was adjusted with lysis buffer to 1 ml. 2 µg anti-PKC antibody (BD Biosciences) was added and incubated at 4°C overnight. Thereafter, 50 µl µMACS protein A microbeads (Miltenyi Biotech) were added and incubated for 6 h. Lysate was applied onto an equilibrated µ column, which was already placed in the magnetic field of a µMACS separator. The flowthrough was collected and saved for further analysis. The column was rinsed 4 times with 200 µl wash buffer (150 mM NaCl, 1% Igepal CA-630, 0.5% sodium deoxycholate, 0.1% SDS, and 50 mM Tris HCl, pH 8.0), followed by 2 washes with low ionic buffer (20 mM TrisHCl, pH 7.5). Afterward, the column was removed from the magnetic field and the remaining proteins were eluted using 50 µl of lysis buffer.

COS-7 cells were transiently transfected by electroporation (450 V/300 µF; Equibio Easyjet T Prima; Peqlab) with a combination of pcDNA3 PPAR1 wild-type or pcDNA3-PPAR1 AF2 and pPKC-EGFP. Immunoprecipitation was performed as described in the previous paragraph using µMACS anti–GFP-microbeads (Miltenyi Biotec)

Mammalian two-hybrid assay
To use PPAR1 and PKC in the mammalian two-hybrid system (Stratagene), PPAR1 was cloned into the BamHI–HindIII site of the pCMV-AD MCS, and PKC was cloned into the BamHI–HindIII site of the pCMV-BD MCS. PPAR was amplified from the pcDNA3-PPAR1 wild-type vector and PKC from the vector pPKC-EGFP. The following primers were used: pCMV-BD-PPAR1, 5'-GCCGGAA TTGGGATCCATGGTTGACACAGAGATGCCATTCTG-3' and 5'-ACGCGGCCGCAAGC TCTAGTACAAGTCCTTGTAGATCTCCTGCAGG-3'; pCMV-AD-PKC, 5'-CAGCGGCC AAGGAT- CCATGGCTGACGTTTTCCCGGG-3' and 5'-ACGCGGCCGCAAGC- TTCATA CTGCACTCTGTAAGATGGGGTGC-3'. Annealing temperatures were 62°C for the first cycle and 72°C for the later ones, and were calculated using the Oligo software (MBI). Infusion reaction of the BamHI–HindIII–cleaved vectors with the amplified PPAR1 wild-type- or PKC- fragment was performed according to the distributor's instructions. Correct orientation and sequence of the generated vectors was verified by restriction analyses and sequencing. COS-7 cells were transiently transfected by electroporation using a combination of the two constructed vectors, as well as the pFR-luciferase reporter vector (Stratagene). Afterward, cells were incubated for 24 h, and then stimulated for 6 h with 10 µM ciglitazone, 10 µM rosiglitazone, or 10 µM WY14643, or they remained as controls. Thereafter, cells were lysed and assayed for firefly luciferase activity by a luciferase assay (Promega).

Western blot analysis
Cell lysis was achieved with lysis buffer (50 mM Tris, 5 mM EDTA, 150 mM NaCl, 0.5% Nonidet-40, and 1 mM PMSF, pH 8.0) and 20-s sonication (Sonifier; duty cycle 100%, output control 60%). Whole-cell lysates were cleared by centrifugation (10,000 g for 5 min), and protein concentration was determined with the Lowry method. 80 µg of protein was resolved on 10% polyacrylamide gels and blotted onto nitrocellulose sheets, basically following standard methodology. Equal loading and correct protein transfer to nitrocellulose was routinely quantitated by Ponceau S staining. Filters were incubated with the anti-PKC antibody (1:500; BD Biosciences), anti-PPAR antibody (1:500; Santa Cruz Biotechnology, Inc.), anti-RFP antibody (1:1,000; MBL), or anti-actin antibody (1:2,000; GE Healthcare) overnight at 4°C. Horseradish peroxidase–conjugated polyclonal antibodies (1:5,000; GE Healthcare) were used for enhanced chemiluminescence detection.

Quantification of TNF- expression
Supernatants from RAW 264.7 macrophages treated as indicated were harvested after the indicated times. Content of TNF- was quantified using the BD Cytometric Bead Array TNF- Flex Set (BD Biosciences) according to the supplier's instructions using a FACSCanto flowcytometer. Interpretation of the results was performed with the FCAP Array software (Soft Flow, Inc./BD Biosciences).

EMSA
Nuclear extracts were prepared as previously described (von Knethen and Brune, 2001). An established EMSA method, with slight modifications, was used (Camandola et al., 1996). Nuclear protein (20 µg) was incubated for 30 min at room temperature with 2 µg poly(dI-dC) from GE Healthcare, 2 µl buffer D (20 mM Hepes/KOH, 20% glycerol, 100 mM KCl, 0.5 mM EDTA, 0.25% Nonidet P-40, 2 mM DTT, and 0.5 mM PMSF, pH 7.9), 4 µl buffer F (20% Ficoll-400, 100 mM Hepes/KOH, 300 mM KCl, 10 mM DTT, and 0.5 mM PMSF, pH 7.9), and 250 fmol 5'-IRD700–labeled oligonucleotide (Metabion) in a final volume of 20 µl. Specific p65 and p50 supershift antibodies (2 µg; Santa Cruz Biotechnology, Heidelberg, Germany) were added as indicated. DNA–protein complexes were resolved at 80 V for 1 h in a native 6% polyacrylamide gel, and visualized with the Odyssey infrared imaging system (LI-COR). Oligonucleotides with the consensus NF-B site (bold letters) were used (Peng et al., 1995): 5'-GCCAGTTGA GGGGACTTTCCCAGGC-3'; 3'-CGGTCAACTCCCCTGAAAG GGTCCG-5'.

Statistical analysis
Each experiment was performed at least three times. Statistical analysis was performed using the paired t test. We considered P values 0.05 as significant. Otherwise, representative data are shown.

	Acknowledgments

 
We thank Nadja Wallner for expert technical assistance.

This work was supported by grants from the Deutsche Forschungsgemeinschaft (Br999).

Submitted: 5 May 2006

Accepted: 24 January 2007

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