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Originally published In Press as doi:10.1074/jbc.M513101200 on December 27, 2005

J. Biol. Chem., Vol. 281, Issue 11, 7189-7196, March 17, 2006
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Cytochrome P-450 4F18 Is the Leukotriene B4 {omega}-1/{omega}-2 Hydroxylase in Mouse Polymorphonuclear Leukocytes

IDENTIFICATION AS THE FUNCTIONAL ORTHOLOGUE OF HUMAN POLYMORPHONUCLEAR LEUKOCYTE CYP4F3A IN THE DOWN-REGULATION OF RESPONSES TO LTB4*

Peter Christmas{ddagger}, Karine Tolentino{ddagger}, Valeria Primo{ddagger}, Karin Zemski Berry§, Robert C. Murphy§, Mei Chen, David M. Lee, and Roy J. Soberman{ddagger}1

From the {ddagger}Renal Unit and Department of Medicine, Massachusetts General Hospital (East), Charlestown, Massachusetts 02129, the §Department of Pharmacology, University of Colorado Health Sciences Center, Aurora, Colorado 80045, and the Division of Rheumatology and Immunology, and Allergy, Brigham and Women's Hospital, Boston, Massachusetts 02115

Received for publication, December 8, 2005


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Leukotriene B4 (LTB4) is a potent chemoattractant for polymorphonuclear leukocytes (PMN) and other cells. Human PMN inactivate LTB4 by {omega}-oxidation catalyzed by cytochrome P-450 (CYP) 4F3A. The contribution of the enzymatic inactivation of LTB4 by CYP4Fs to down-regulating functional responses of cells to LTB4 is unknown. To elucidate the role of CYP4F-mediated inactivation of LTB4 in terminating the responses of PMN to LTB4 and to identify a target for future genetic studies in mice, we have identified the enzyme that catalyzes the {omega}-1 and {omega}-2 oxidation of LTB4 in mouse myeloid cells as CYP4F18. As determined by mass spectrometry, this enzyme catalyzes the conversion of LTB4 to 19-OH LTB4 and to a lesser extent 18-OH LTB4. Inhibition of CYP4F18 resulted in a marked increase in calcium flux and a 220% increase in the chemotactic response of mouse PMN to LTB4. CYP4F18 expression was induced in bone marrow-derived dendritic cells by bacterial lipopolysaccharide, a ligand for TLR4, and by poly(I·C), a ligand for TLR3. However, when bone marrow-derived myeloid dendritic cells trafficked to popliteal lymph nodes from paw pads, the expression of CYP4F18 was down-regulated. The results identify CYP4F18 as a critical protein in the regulation of LTB4 metabolism and functional responses in mouse PMN and identify it as the functional orthologue of human PMN CYP4F3A.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
How polymorphonuclear leukocytes (PMN),2 macrophages, and dendritic cells (DC) control the initiation and amplification of innate and adaptive immune responses is a critical question, and the 5-lipoxygenase product of arachidonic acid metabolism leukotriene B4 (LTB4)is central to the amplification process. LTB4 is equal to the most potent chemoattractant known for myeloid cells (1–6) and is synthesized from arachidonic acid by the action and interactions of 5-lipoxygenase, the five-lipoxygenase-activating protein, and leukotriene A4 hydrolase (7). LTB4 mediates its activity in these cells via the high affinity G protein-coupled receptor BLT1 (8–10).

LTB4 has been implicated in the pathogenesis of multiple inflammatory diseases including inflammatory bowel disease (11–14), glomerulonephritis (15, 16), allograft rejection in kidney transplant models (17, 18), and cardiac allograft rejection (19). Studies with knock-out mice for the five-lipoxygenase-activating protein combined with LTB4 receptor antagonists have supported a role for LTB4 in murine collagen arthritis (20), in the EAE model of multiple sclerosis (21), and in mediating a primate model of asthma (22). In atherosclerosis, 5-lipoxygenase has been identified as a risk gene in a mouse model, and 5-lipoxygenase-rich cells have been identified in atheroscleotic plaques of mice and humans (23, 24). Furthermore, a protein closely related to CYP4F3A, presumably a mouse member of the CYP4F family, was strongly induced in foam cells in mice (23). Blockade of BLT1 has been associated with decreased progression of atherosclerosis in APOE1-/- mice (25, 26). Understanding the molecular basis of LTB4 signal termination is critical to elucidating how animals control the amplitude of inflammation in LTB4-dependent settings.

There are two general cellular mechanisms that have the potential to terminate the responsiveness to LTB4 and to all other chemoattractant molecules for G protein-coupled receptors. The first is the enzymatic metabolism of ligands. The second is receptor desensitization, which is based on G protein-coupled receptor kinases and beta-arrestin (27, 28). The desensitization process has been elucidated in depth (27, 28), whereas the relative importance of ligand inactivation remains unknown.

The major pathway for the metabolism and inactivation of LTB4 in human PMN is the uptake of extracellular LTB4, which is initially converted to 20-OH LTB4 by CYP4F3A (29–34). In rat PMN, the initial {omega}-1 and {omega}-2 oxidation products of LTB4 are 19-OH and 18-OH LTB4 (35, 36). These products (and LTB4 itself) can be converted by a 12-hydroxydehydrogenase to their 11,12-dihydro products (35, 36). In metabolism studies of LTB4 introduced into humans intravenously, 19-OH and 18-OH LTB4 were detected as urinary metabolites (37, 38); these products are not formed in human PMN.

The enzymes that catalyze the {omega}-1 and {omega}-2 oxidation and the 12-hydroxydehydrogenase reactions of LTB4 in mouse leukocytes have not been identified (39–41). Whether there is a unique role for CYP4F metabolism in terminating cellular responses to LTB4 is also unknown. Based on these considerations and with the ultimate goal of understanding the role of CYP4Fs in terminating responses to LTB4 in vivo, we sought to identify the enzyme responsible for the {omega}-1 and {omega}-2 oxidation of LTB4 in mouse PMN and other leukocytes and to identify a functional role for CYP4F family members in the regulation of PMN responses to LTB4.

We identified CYP4F18 as the mouse LTB4 {omega}-1 and {omega}-2 hydroxylase in mouse PMN and macrophages. Inhibition of CYP4F18 by the irreversible inhibitor of fatty acid and eicosanoid {omega}-hydroxylases 17-ODYA resulted in a 220% increase in the chemotactic response of PMN to LTB4 and a marked increase in calcium flux in response to LTB4. Inhibition of human PMN CYP4F3A also resulted in clear augmentation of calcium flux in response to LTB4. In addition, CYP4F18 expression was induced in bone marrow-derived DC by bacterial lipopolysaccharide, a ligand for TLR4 and poly(I·C), a ligand for TLR3. However, when DC trafficked to popliteal lymph nodes, the pattern of CYP4F18 expression was down-regulated relative to CYP4F15. The results identify {omega}-1 and {omega}-2 oxidation by CYP4F18 as a step in controlling the inactivation of functional responses to LTB4 in mouse PMN.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Isolation of Mouse Cells and Tissues—9–12-week-old male and female Balb/c mice were used for experimentation and were euthanized with CO2 in accordance with the guidelines of the Massachusetts General Hospital/Partners Committee on Research Animal Care. Selected tissues (liver, kidney, spleen, thymus, heart, lung, brain, ovary, and bladder) were dissected for RNA analysis immediately after euthanization. Axillary, inguinal, and popliteal lymph nodes were dissected and pooled for analysis. Peripheral blood samples were drawn from the tail veins of mice prior to euthanization.

Isolation of Peritoneal PMN and Macrophages—To generate peritoneal PMN, mice were injected intraperitoneally with 1 ml of sterile 0.9% sodium caseinate. After 18 h the mice were euthanized, and 9 ml of sterile PBS was injected into the peritoneal cavity to harvest PMN by peritoneal lavage. To isolate peritoneal macrophages, mice were injected intraperitoneally with 1 ml of sterile 3% sodium thioglycollate, and macrophages were harvested 72 h after injection by peritoneal lavage.

In Vitro Differentiation and in Vivo Injection of Mouse DC—Immature mouse DC were obtained by a modification of the method of Inaba et al. (42, 43). Bone marrow cells were isolated by flushing the femurs and tibiae of mice with RPMI 1640 containing 10% heat-inactivated fetal bovine serum. The cells were filtered through a spleen mesh, centrifuged at 1200 rpm for 5 min, and resuspended in ACK buffer (BioWhittaker) for 5 min to lyse red blood cells. The cells at this stage were used to analyze the expression of CYP4F18 in total bone marrow. The cells were then washed and cultured (5 x 105 cells/ml) in RPMI 1640 supplemented with 25 mM HEPES, pH 7.4, 10% fetal bovine serum, 1 mM sodium pyruvate, 2 mM beta-mercaptoethanol, granulocyte macrophage colony stimulating factor (100 units/ml) and interleukin-4 (100 units/ml). On day 2 and 4, the media and nonadherent cells were replaced with fresh media. On day 6, nonadherent cells (DC) were removed and resuspended in fresh medium at the same cell density. Differentiation to DC was confirmed by analysis of CD11c expression by flow cytometry. For studies of DC activation in vitro, the cells were stimulated with 10–100 ng/ml LPS (Escherichia coli, 055:B5) or 25–50 µg/ml poly(I·C) (Amersham Biosciences) and analyzed for CD80, CD86, and CYP4F isoform expression after 24 h. To target DC to popliteal lymph nodes, immature bone marrow-derived DC were labeled for 5 min with 5 µM CFDA-SE (carboxyfluorescein diacetate, succinimidyl ester) dye from (Invitrogen). The cells were washed twice, and then 5 x 105 cells were mixed with 1 µg/ml LPS in a 10-µl volume and injected into the right paw pads of mice. The mice were euthanized 18 h after injection, and popliteal lymph node tissue was dissected, dissociated with 1000 IU/ml collagenase D (Roche Applied Science) for 2 x 1 h at 37 °C, and filtered through a spleen mesh (44). The cells were resuspended in ACK buffer (BioWhittaker) for 5 min, washed in PBS, and centrifuged on 35% bovine serum albumin gradients for 15 min at 7000 rpm, 4 °C. The cells were collected from the interface of the bovine serum albumin gradient, and CFDA-SE dye-stained cells were isolated by flow cytometry for RNA analysis.

RNA Blot Analysis—A CYP4F18 cDNA probe was synthesized by labeling a 297-bp PCR product with [{alpha}-32P]dCTP by random priming; the PCR product was generated from mouse PMN cDNA with primers F18-2F and F18-6R (Table 1). PCR conditions were 94 °C for 30 s, 60 °C for 30 s, and 72 °C for 30 s for 30 cycles, and the product was purified using a Geneclean Spin kit (Qbiogene) prior to radiolabeling. A mouse multiple tissue expression array from BD Biosciences (Clontech) was hybridized with the 297-bp probe (2 ng/ml, ~2 x 106 cpm/ng) for 6 h at 65 °C in 10 ml of ExpressHyb buffer. The array was washed in 2x SSC, 1% SDS at 65 °C five times for 20 min and then 0.1x SSC, 0.5% SDS at 55 °C twice for 20 min and was exposed to Kodak XAR film overnight at -70 °C with an intensifying screen. Hybridization signals were quantified with a PhosphorImager.


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  		TABLE 1
Summary of primers cDNA positions are relative to the ATG initiation codon (A = +1)

 
Isoform-specific RT-PCR—RNA was extracted from freshly isolated mouse cells and tissues using Tri reagent (Sigma). In addition, total RNA from selected mouse tissues (liver, ovary, and smooth muscle) was purchased from BD Biosciences Clontech. Reverse transcription was performed with a cloned avian myeloblastosis virus first strand cDNA synthesis kit (Invitrogen). Primers summarized in Table 1 (listed under "isoform-specific primers") were used to detect CYP4F13 (182-bp product), CYP4F14 (168-bp product), CYP4F15 (164-bp product), CYP4F16 (244-bp product), and CYP4F18 (177-bp product) by isoform-specific PCR. The same annealing temperature can be used for all primer sets, but different polymerases/buffer systems require independent optimization to determine the annealing temperature critical to achieve specificity. An annealing temperature of 66 °C was optimal for PCRs containing 1 unit of Taq polymerase and 10–100 ng of DNA in 10 mM Tris-HCl, pH 9, 50 mM KCl, 1.5 mM MgCl2, 400 nM each primer, and 200 µM dNTPs. The PCR conditions were 94 °C for 30 s, 66 °C for 30 s, and 72 °C for 30 s; 30 cycles were used unless stated otherwise. Semiquantitative PCR for CYP4F18 was performed by including Quantum RNA Classic 18 S internal standard primers (Ambion); 25 cycles of PCR was representative of the linear range, and the 18 S primers were combined with Competimers (Ambion) in a ratio of 3:7. Alternatively, glyceraldehyde-3-phosphate dehydrogenase RT-PCR Control Amplimers (BD Biosciences Clontech) were used as an internal control. PCR products were resolved on a 1% agarose gel, and the bands were quantified using image analysis software.

Flow Cytometry—DC cultures were stained with allophycocyanin, percyphycoerythrin, or fluoroisothiocyanate-conjugated control IgGs or monoclonal antibodies directed to CD11c, CD80, and CD86 (BD Biosciences) to define immature and mature populations using a double or tri-color labeling. The samples were analyzed by flow cytometry using a FACScalibur instrument and the Cellquest program (BD Biosciences).

Expression of CYP4F18 Protein—The full-length cDNA coding region of CYP4F18 was amplified from PMN cDNA by PCR using Pfu polymerase (Stratagene) with primers F18-1F (which places the sequence CGACC immediately upstream of the ATG initiation codon) and F18-13R (which comprises the last 33 bp inclusive of the TGA stop codon). The PCR conditions were 94 °C for 30 s, 60 °C for 30 s, and 72 °C for 2 min for 30 cycles. A PCR product of 1580 bp was cloned into pCR2.1-TOPO vector (Invitrogen), and sequence analysis confirmed its identity with CYP4F18. The cDNA for CYP4F18 was subcloned into p51polORbp bicistronic baculovirus vector designed for coexpression with NADPH cytochrome P-450 reductase. Preparation of baculovirus, infection of Sf9 cells (at the Protein Expression Center at California Institute of Technology and at Gentest Corporation, Woburn, MA), and fractionation of cell extracts and microsomes was performed as previously described (32). Microsomes were adjusted to a concentration of 7.7 mg protein/ml (Lowry assay), and CYP4F18 expression was confirmed by Western blotting (31).

Immunofluorescent Microscopy—For indirect immunofluorescent staining of CYP4F18, DC were washed in PBS and cytospun onto slides. The cells were fixed in 4% paraformaldehyde in PBS for 30 min, incubated with 50 mM ammonium chloride for 10 min, and permeabilized with 0.1% Triton X-100 in PBS for 4 min. The cells were then blocked in 10% goat serum for 30 min and labeled with affinity-purified rabbit anti-CYP4F3 (10 µg/ml) for 1 h at room temperature. The affinity purified antibody to CYP4F3 has been described previously (32, 34), and it reacts with both human (CYP4F3) and mouse (CYP4F18) proteins. The secondary antibody was fluorescein isothiocyanate-conjugated goat anti-rabbit IgG (Jackson Laboratories), used at a 1:250 dilution in PBS for 1 h at room temperature. Immunofluorescent microscopy was performed using a Nikon FXA photomicroscope and IP Spectrum (Scanalytics, Vienna, VA) acquisition analysis software.

Analysis of LTB4 and AA Metabolites by RP-HPLC—A 100-µl reaction mixture containing 1 mM NADPH, 10 µl of Sf9 microsomes expressing CYP4F18, 30 µM LTB4, and 2 µCi of [3H8]LTB4 in 100 mM potassium phosphate buffer, pH 7.4, was incubated at 37 °C for 30 min. After the incubation, the reaction was stopped by adding 100 µl of ice-cold ethanol and centrifuged (10,000 x g) for 10 min. The supernatant was removed, and 1 ml of 100 mM potassium phosphate buffer, pH 7.4, and 1 drop of formic acid was added. The products were extracted (twice) using 1 ml 1:1 (v/v) hexane/ethyl acetate. After vortexing four times for 15 s, the samples were centrifuged at 135 x g for 5 min to separate the two layers. The organic layer was removed and taken to dryness under vacuum. Metabolites were purified by RP-HPLC using a Chromolith Performance RP-18e (4.6 x 100 mm) column. The reversed phase solvents used were 8.3 mM acetic acid adjusted to pH 5.7 with ammonium hydroxide (solvent A) and 65:35 acetonitrile/methanol (solvent B). The initial mobile phase was 25% solvent B, which was held for 3 min. This was followed by a linear gradient to 100% solvent B over 32 min. Initially, 5% of the hexane/ethyl acetate extract was introduced onto the RP-HPLC column, and the effluent was monitored using UV detection at 270 nm and on-line radioactivity. The remainder of the hexane/ethyl acetate extract was then introduced onto the RP-HPLC column, and 2 fractions/min were collected for GC/MS analysis.

Gas Chromatography/Mass Spectrometry—The reversed phase HPLC fractions of interest were taken to dryness under vacuum and derivatized for GC/MS analysis by the addition of 50 µl of 5% N,N-diisopropylethylamine in acetonitrile and 50 µl of 5% pentafluorobenzyl bromide in acetonitrile. The samples were kept at room temperature for 30 min and evaporated under a stream of nitrogen. The samples were further derivatized with the addition of 50 µl of acetonitrile and 50 µlof bis(trimethylsilyl)trifluoroacetamide by incubating at 60 °C for 30 min followed by evaporation under nitrogen. The samples were reconstituted in 20 µl of acetonitrile and subjected to GC/MS analysis (35, 45–47). A gas chromatograph/mass spectrometer (Trace 2000; Thermo Finnigan, San Jose, CA) was used for the electron ionization analysis. The electron ionization spectra were obtained at an electron energy of 70 eV and provided structural information regarding the hydroxyl group position from fragmentation events adjacent to the trimethylsilyl ether positions.

Calcium Flux and Chemotaxis—For cell migration assays, bone marrow cells were isolated from the femurs and tibias of mice by perfusion with sterile PBS (48). In some experiments they were further isolated by a discontinuous Percoll gradient at 500 x g for 30 min at room temperature (49, 50). The three-step gradient was 55, 65, and 75% (v/v) Percoll in PBS. The mature PMN are recovered at the interface of the 65 and 75% fractions. PMN purity was >97% as determined morphometrically by Diff-Quik staining. The cells were fluorescently labeled with 5 µM calcein AM (Invitrogen), suspended in 20 mM HEPES, pH 7.5, 125 mM NaCl, 5 mM KCl, 1 mM MgCl2, 1 mM CaCl2, 0.5 mM glucose, 0.2% bovine serum albumin, at 1 x 107/ml (48). The cells were also incubated with 30 µM 17-ODYA or buffer containing 0.05% Me2SO (controls) for 30 min and then tested for chemotactic activity in modified Boyden chambers. The cells (0.1 ml) were placed in the upper wells of 3-micron, 6.4-mm FluoroBlok filter inserts (Falcon) with 0.6 ml of buffer containing 1.0, 10, or 100 nM of LTB4 or 20-OH LTB4. The chambers were incubated at 37 °C for 45 min, and chemotaxis was determined by measuring the fluorescence intensity (excitation, 485 nm; emission, 535 nm) passing to the underside of the filter. The measurements were determined in quadruplicate for independent experiments.

For calcium flux determination (51), mouse PMN and peripheral blood human PMN were counted and incubated with the indicator dye Fura-2 AM (Invitrogen) for 1 h at 37°C in 20 mM HEPES, pH 7.4, 1 mM CaCl2, 1 mM MgCl2, 125 mM NaCl, 5 mM KCl, 0.5 mM glucose, and 0.2% bovine serum albumin (51). The cells were washed twice and resuspended in the same buffer at 1 x 106/ml and then incubated with either 17-ODYA (30 µM) or Me2SO (final concentration, 0.05%) for 30 min. Changes in intracellular calcium concentration in response to LTB4 were determined fluorometrically at 37 °C by monitoring the emission at 510 nm and the excitation at 340 and 380 nm as a function of a time (51). The responses were quantified as the peak of the fluorescence ratio of 340/380 nm. These data are representative of two or three determinations.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
CYP418 Is the Mouse PMN LTB4 {omega}-1/{omega}-2 LTB4 Hydroxylase—The major pathway for the inactivation of LTB4 in human PMN is {omega}-oxidation catalyzed by CYP4F3A to generate 20-OH LTB4 (29–34, 52–54). The mouse genome includes multiple Cyp4f genes that code for proteins with 70–80% amino acid identity. Therefore, we used isoform-specific primers to identify the CYP4F family member(s) that were expressed in mouse peritoneal PMN elicited with sodium caseinate that could function as the mouse orthologue of CYP4F3A. As shown in Fig. 1, a 177-bp PCR product was identified that was confirmed as bp 93–269 of CYP4F18. In contrast, multiple CYP4Fs were expressed in liver.


Figure 1
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  		FIGURE 1.
CYP4F18 is expressed in mouse PMN. Isoform-specific PCR for CYP4F13 (182-bp product), CYP4F14 (168-bp product), CYP4F15 (164-bp product), CYP4F16 (244-bp product), and CYP4F18 (177-bp product) in mouse liver and peritoneal PMN was performed for 30 cycles. The identity of the 177-bp mouse PMN PCR product was confirmed to be CYP4F18 by sequencing.

 
Because rodent PMN generate 19-OH and 18-OH LTB4 as {omega}-1, and {omega}-2 oxidation products, we next determined whether CYP4F18 formed these products from LTB4. The cDNA for CYP4F18 was cloned from mouse peritoneal PMN and coexpressed with NADPH cytochrome P-450 reductase in Sf9 cells using a baculovirus expression system (see "Materials and Methods"). Microsomes were prepared from the Sf9 cells, and the reaction products of 30 µM LTB4 and tracer quantities of [3H] LTB4 were determined. As shown in Fig. 2, two peaks were identified by RP-HPLC by both radioactivity (Fig. 2A) and by the conjugated triene chromophore that absorbed at 270 nm (Fig. 2B). The major peak was identified by GC/MS as 19-OH-LTB4 after derivatization to the pentafluorobenzyl ester, trimethylsilyl ether derivative, and electron ionization (Fig. 2C). The origins of the major ions are indicated in the structural inset in Fig. 2 and were consistent with the previous identification of this metabolite in human urine extracts (37) as well as detailed mass spectrometric studies of the electron ionization mass spectrometry of the methyl ester, trimethylsilyl ether derivatives of LTB4 (46). Of particular importance was the ion at m/z 117, which has been found to be a definitive {alpha}-cleavage ion that establishes the site of hydroxylation of various {omega}-hydroxylated eicosanoids (37, 47). The {alpha}-cleavage ion was shifted to m/z 131 in the mass spectrum of the metabolite identified as 18-OH-LTB4 (data not shown) (37). When 30 µM arachidonic acid was utilized as a substrate, no product was formed. These results demonstrate a clear preference for LTB4 versus arachidonic acid as a substrate and identify 19-OH LTB4 as the major product and 18-OH LTB4 as the minor product, consistent with results seen in rodent PMN (35, 36, 41).

Because of its expression in several cell types, we next determined whether the Cyp4f18 gene contains alternately spliced exons similar to the human CYP4F3 gene. In humans, exons 3 and 4 code for amino acids 67–114 of the mature protein and are alternately spliced on a tissue-specific basis. We isolated and sequenced mouse genomic clones that span exons 1–5 of the Cyp4f18 gene and include 7 kb of the 5'-flanking sequence (17.2 kb total). Sequence analysis confirmed that no alternative exons are predicted. Therefore, a single exon codes for amino acids 67–114 of CYP4F18. This exon has 70% amino acid identity to exon 4 of CYP4F3 (Fig. 3). There is no other mouse gene with an exon that contains greater than 40% identity to exon 4.


Figure 2
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  		FIGURE 2.
Identification of the {omega}-oxidation products of LTB4 catalyzed by CYP4F18. A, 30 µM LTB4 and 2 µCi of [3H8]LTB4 were incubated with CYP4F18 microsomes for 30 min at 37 °C, and metabolites were separated by RP-HPLC with on-line radioactivity monitoring. Two LTB4 metabolites with retention times of 12.1 min (major) and 13.1 min (minor) were observed. The remaining unreacted LTB4 eluted at 22 min. B, on-line UV detection at 270 nm revealed that the metabolites retained the conjugated triene chromophore. C, positive ion electron ionization mass spectrum (70 eV) of the pentafluorobenzyl/trimethylsilyl-derivatized metabolite with a reversed phase retention time of 12.1 min. The retention time of this metabolite during gas chromatography was 10.01 min. The ion at m/z 117 supported the location of the 19-hydroxy moiety for derivatized 19-OH LTB4 as the major metabolite.

 


Figure 3
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  		FIGURE 3.
Relationship of human CYP4F3 and mouse Cyp4f18 genes. A, alignment of the exons between the Cyp4f18 and CYP4F3 genes. B, the amino acid identities between the residues coded for by exon 3 and exon 4 of CYP4F3 with amino acids coded for by exon 3 of mouse Cyp4fs.

 
Tissue-specific Expression of CYP4F18—Initially, we used a mouse multiple tissue expression array to investigate the tissue distribution of CYP4F mRNA (data not shown). The array was hybridized with a 32P-labeled cDNA probe for CYP4F18 (bp 212–508); other CYP4F members have ~65–70% nucleotide identity within the 297-bp region of the probe and may cross-hybridize. Hybridization signals were detected in multiple tissues, and quantification confirmed that the strongest signals were in liver, smooth muscle, epididymus, and ovary. No hybridization was observed in the negative control grids with the exception of mouse genomic DNA, which permits detection of the CYP4F multigene family on chromosome 17.

The tissue distribution of CYP4F family members was then analyzed with isoform-specific primers in semi-quantitative RT-PCR experiments, using 18 S RNA as an internal standard (Fig. 4A). CYP4F18 is expressed at low levels in liver, kidney, and smooth muscle and is barely detectable in these tissues under linear PCR conditions (25 cycles). In contrast, it is highly expressed in ovary. CYP4F18 is detected in a variety of hematopoietic tissues including lymph node, spleen, bone marrow, and peripheral blood (Fig. 4A, lower panel) and is expressed at the highest levels in cell populations enriched in peritoneal PMN and mature bone marrow-derived DC (see below).

Recent reports have indicated that DC have the ability to both synthesize and respond to LTB4 (55–59), which has the potential to amplify the recruitment of other DC and possibly Th1 or Th2 CD4+ and effector CD8+ T cells. The mechanisms by which DC inactivate LTB4 are not known. We therefore determined whether CYP4F18 could be detected in DC. The generation of immature myeloid DC from bone marrow was confirmed by flow cytometry analysis of CD11c expression (Fig. 4B, upper left panel). DC were then incubated with 25 µg/ml poly(I·C) or 100 ng/ml LPS for 24 h, resulting in an increased expression of CD80 and CD86 consistent with cell maturation (Fig. 4B, upper right panel), and a concomitant increase in CYP4F18 mRNA and protein expression was observed (Fig. 4B, lower panels).

We next investigated whether DC could modulate the expression of CYP4F18 during in vivo trafficking from foot pads to popliteal lymph nodes (Fig. 4C). Murine bone marrow cells were differentiated to myeloid DC. DC were then labeled with 5 µM CFDA-SE dye, mixed with 1 µg/ml LPS in a 10-µl volume, and injected into the right paw pads of mice. Right popliteal lymph nodes were isolated for analysis by dissection after 18 h and then digested with collagenase (44). After separation on 35% bovine serum albumin gradients, the viable cells were isolated from the interface and then broadly separated by FACS (Fig. 4C, left panel) based on the properties of size (forward scatter) and granularity (side scatter) to generate R1 fraction. This fraction was then analyzed for CFDA-SE expression using the percyphycoerythrin channel as a negative control (Fig. 4C, right panel). High CFDA-SE expressors (fraction R2) were isolated by sorting to greater than 99% purity for further analysis; this represented 2–5% of the total cell population. As shown by RT-PCR analysis (Fig. 4C, lower panels) mature DC prior to foot pad injection expressed only CYP4F18. However, when dye-stained cells were recovered from lymph nodes (fraction R2) and analyzed for CYP4F expression, the expression of CYP4F18 was down-regulated relative to the expression of CYP4F15. This clearly contrasts with the relative expression of these two enzymes in DC induced with LPS ex vivo.

CYP4F18 Is the Functional Orthologue of Human PMN CYP4F3A—We next investigated whether a role for CYP4F18 in regulating the LTB4-dependent chemotaxis of mouse PMN could be identified. We therefore inhibited this enzyme with 30 µM 17-ODYA, an irreversible acetylenic inhibitor of P-450 mixed function oxidases that targets enzymes utilizing eicosanoids and fatty acids, including CYP4Fs (60–65). As shown in Fig. 5, when mouse leukocytes were treated with 30 µM 17-ODYA, chemotaxis in response to LTB4 was elevated by 220% at 10 nM and 150% at 100 nM LTB4. Because LTB4-dependent chemotaxis is mediated by Ca2+ signal transduction, we next investigated whether 17-ODYA treatment augments Ca2+ fluxes in response to LTB4 stimulation (Fig. 5B). In mouse bone marrow-derived PMNs, an increase in Ca2+ flux was observed at concentrations as low as 1 nM LTB4 (and was also observed at 10 nM LTB4), and the response was enhanced in cells treated with 17-ODYA (Fig. 5B, upper panel). Similar results were obtained with human peripheral blood PMNs (Fig. 5B, lower panel). The effect of 17-ODYA was greatest at 10 nM LTB4, in which case the magnitude of the Ca2+ response was comparable with that normally observed at 100 nM LTB4. The data provide evidence that CYP4F3 and CYP4F18 operate as functional orthologues to down-regulate LTB4-dependent responses (via 17-ODYA-sensitive LTB4 hydroxylation) in human and mouse PMN, respectively.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The major pathway for the metabolism and inactivation of LTB4 in human PMN is {omega}-hydroxylation by CYP4F3A, which generates 20-OH LTB4 (30–34). A parallel hydroxylation pathway has been identified in rodent PMN, in this case generating 19-OH LTB4 and 18-OH LTB4. In this report we identify CYP4F18 as the mouse CYP4F enzyme expressed in elicited peritoneal PMN and peritoneal macrophages (Figs. 1 and 4). We demonstrate that the recombinant enzyme converts LTB4 to 19-OH LTB4 and 18-OH LTB4 in the expected ratio (Fig. 2).

The augmentation of chemotaxis and calcium flux in response to the inhibition of CYP4F18 and the augmentation of calcium flux seen by the inhibition of CYP4F3A demonstrate a link between CYP4F-mediated LTB4 metabolism and the down-regulation of LTB4-induced biological functions and indicates that CYP4F18 is a biochemical and functional orthologue of human CYP4F3A (Fig. 5). These data are parallel to that seen for inhibition of dipeptidyl peptidase IV (DPPIV), which catalyzes the proteolysis of CXCL12 and other chemokines (66–71). Inhibition of DPPIV using a chemical inhibitor augments the calcium flux of hematopoetic stem cells in response to CXCL12 and augments both chemotaxis in vitro and the recruitment and engrafting of stem cells in bone marrow after intravenous injection in mice. The data support a role for CYP4F18 analagous to DPPIV.


Figure 4
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  		FIGURE 4.
Tissue distribution of CYP4F18. A, quantitative RT-PCR analysis of CYP4F18 in mouse tissues and cells. Isoform-specific primers for CYP4F18 (177-bp product) were combined with primers for 18 S RNA as an internal standard (488-bp product) under linear PCR conditions (25 cycles). B, CYP4F18 expression is induced in myeloid DC. In vitro differentiation of mouse bone marrow cells to myeloid DC was confirmed by staining with an allophycocyanin-conjugated monoclonal antibody to CD11c and analysis by flow cytometry (upper left, control represents unstained cells). The dendritic cells were then stimulated with 25 µg/ml poly(I·C) or 100 ng/ml LPS for 24 h, and maturation was indicated by increased expression of CD80 and CD86 using percyphycoerythrin-conjugated antibodies (upper right, control, represents mock stimulated cells). DC RNA was analyzed by RT-PCR (25 cycles) using primer pairs for CYP4F18 and glyceraldehyde-3-phosphate dehydrogenase, confirming increased expression of CYP4F18 in cells treated with poly(I·C) and LPS (lower left). CYP4F18 expression was increased as detected by immunofluoresence in cells stimulated with 25 µg of poly(I·C) and by Western blot analysis of cells stimulated with 100 ng/ml LPS (lower right). Equal amounts of cell extracts (20 µg) from induced and uninduced DC were loaded for Western blot analysis, and both direct and inverse black and white imaging are shown to emphasize the contrast. C, in vivo regulation of CYP4F18 expression. Bone marrow-derived DC were stained with CFDA-SE dye, stimulated with LPS, and injected in the right hind paw pads of mice. 18 h later the dye-stained cells were isolated from popliteal lymph nodes. Lymph node cells were broadly separated by flow cytometry based on the properties of size (forward scatter) and granularity (side scatter) to generate fraction R1 (left panel). This fraction was analyzed for CFDA-SE expression (right panel), and high CFDA-SE expressors (fraction R2) were isolated for analysis. RT-PCR analysis of DC RNA prior to injection into mice and after recovery from lymph nodes is shown.

 


Figure 5
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  		FIGURE 5.
CYP4F18 regulates PMN responses to LTB4. A, increase in chemotactic response in mouse PMN after inhibition of CYP4F18 with 30 µM 17-ODYA. The data are the means ± S.D. for two experiments with each point determined in quadruplicate. B, calcium fluxes in human and mouse PMN with or without inhibition of CYP4F18 with 30 µM 17-ODYA.

 
DC move between blood and tissue microenvironments to acquire antigen, migrate to lymph nodes, and present processed antigen to lymphocytes. Recent results have shown that DC both respond to LTB4 and synthesize LTB4 (55–59). Thus, mechanisms that regulate the responsiveness of DC to LTB4 have the potential to play an important role in DC recruitment and trafficking. Our results (Fig. 4) place CYP4F18 in the position to play an important role in the responses of these leukocytes to LTB4. It is not clear at what point during the trafficking from foot pads to lymph nodes that the relative down-regulation of CYP4F18 occurs. This will be a critical question to answer. The induction of CYP4F15 in response to LPS supports the previous observation of induction of CYP4F15 in mouse liver subsequent to the injection of LPS (72). The biological importance of the differential regulation of distinct CYP4Fs remains to be determined.

There are two cellular mechanisms that have the potential to terminate responsiveness to chemoattractant ligands (including LTB4) for G protein-coupled receptors. The first is enzymatic conversion of ligands to alternate structures by covalent modification. The second is receptor desensitization, which involves G protein-coupled receptor kinases and beta-arrestin (27, 28). The relative importance of these two mechanisms in terminating responsiveness to LTB4 and the extent to which they are integrated in vivo are not established. Our studies are the first to demonstrate an important role for ligand metabolism as opposed to receptor desensitization for LTB4 and identify a unique role for CYP4Fs in down-regulating LTB4 function. Finally, the work establishes the basis for future genetic studies elucidating the role of CYP4F18 and LTB4 metabolism in the control of LTB4-mediated functions of leukocytes.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grants GM-61823 (to R. J. S.), DK59991 (to P. C.), 5R01HL025785-25 (to R. C. M.), 5R01HL068256-03 (to N. C.), R01HL25785 (to R. C. M.), K08 AR 02214-04 and R01 AI 059746-02 (to D. L.), R03DK067940 and the Patricia Weilder Young Investigator Award from the National Kidney Foundation (to B. N.), and a gift from the Jewish Communal Fund (to R. J. S). 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. Back

1 To whom correspondence should be addressed: Renal Unit and Dept. of Medicine, MA General Hospital (East), Bldg 149, Navy Yard, 13th St., Charlestown, MA 02129. Tel.: 617-726-3747; Fax: 617-726-5669; E-mail: soberman{at}helix.mgh.harvard.edu.

2 The abbreviations used are: PMN, polymorphonuclear leukocyte(s); DC, dendritic cells; CYP, cytochrome P-450; LTB4, leukotriene B4; 17-ODYA, 17-octadecynoic acid; CFDA-SE, carboxyfluorescein diacetate, succinimidyl ester; PBS, phosphate-buffered saline; LPS, lipopolysaccharide; RT, reverse transcription; HPLC, high pressure liquid chromatography; RP, reversed phase; GC/MS, gas chromatography/mass spectrometry. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Norma P. Gerard (Children's Hospital, Harvard) for assistance with the chemotaxis and calcium measurements. We thank Dr. Henry Strobel (University of Texas, Houston) for advice and assistance.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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