Cystic fibrosis (CF) is caused by mutations in the cystic fibrosis transmembrane conductance regulator (CFTR) gene. CFTR is a cAMP-regulated chloride channel expressed in the apical plasma membrane of many epithelia. In the absence of CFTR, exocrine secretions are of low volume and are abnormally acidic. Hallmarks of CF are accumulation of excessive mucus, microbial infection, and inflammation. During microbial infection, inflammation in CF-affected tissues may be greater than would be expected in a similarly infected non-CF individual. There is also evidence for inflammation even in the absence of active infection and, at the present time, the reason for this is not well understood (1,2). A current hypothesis is that lack of CFTR affects membrane lipid metabolism such that lipid-mediated signals are altered, resulting in a proinflammatory state (3–5).
Before the CF gene was cloned in 1989, it already was known that there are imbalances in essential fatty acids in CF (6–8). More recent work has shown there are decreases in docosahexaenoic acid in CF serum lipids (9) and this can be accompanied by increased arachidonic acid (AA) levels in CF-affected tissues (10). There are also several reports of increased AA production by CF cells (3,8,11). There is evidence that phospholipase A2 (PLA2) activity is increased in CF cells (12–14), which may account for the increased AA production and the resultant imbalances in membrane essential fatty acids. The increased amount of AA in CF cells is expected to be converted to various eicosanoids, including proinflammatory prostaglandins (PGs).
In patients with CF, the small intestine already is affected at birth and a variety of gastrointestinal (GI) problems are common lifelong issues (15). PGs are important in the healthy GI tract as well as in disease states. In health, PGs are beneficial because they stimulate mucus and bicarbonate secretion, which protect the epithelium from microbes and noxious stimuli such as gastric acid. However, during inflammation in the CF intestine, elevated PG levels may contribute to the excessive mucus secretion (16) that is an obvious contributor to the pathological state. In the healthy gut, PGs also have important roles in modulation of enteric smooth muscle activity and GI transit. Conversely, during inflammation, increased PGs can interfere with GI motility programs (17) and foster conditions that promote bacterial overgrowth and impair digestive function. GI dysmotility (18–24) and small intestinal bacterial overgrowth (22,25) are reported to occur in about half of all patients with CF. Overgrown bacteria in the small intestine will compete for nutrients (26) and can contribute to GI problems in CF including steatorrhea, poor weight gain, nausea, and anorexia (27).
The Cftr knockout mouse (Cftr tm1UNC, CF mouse) exhibits many of the same GI changes that occur in patients with CF, including excessive mucus accumulation, impaired intestinal transit, and bacterial overgrowth (28–30). The small intestine of the CF mouse exhibits an innate immune response with an influx of mast cells and neutrophils (31). Inflammation of the human CF intestine also occurs (32,33), but this is less well characterized mainly because of the inaccessibility of this organ in patients. A previous GeneChip microarray analysis of the CF mouse small intestine showed that, among the more than 150 genes with altered expression, several eicosanoid metabolic genes were affected (31). Therefore, an aim of the present study was a more thorough analysis of gene expression in the 3 major AA metabolic pathways that produce hydroxyeicosatetraenoic acids (HETEs), leukotrienes, and PGs. We also measured specific eicosanoids that can be produced in the 3 major pathways. There are significant changes in eicosanoid metabolic gene expression as well as altered levels of specific eicosanoids in the CF intestine. The changes we discovered are discussed in the context of intestinal dysfunction in CF.
MATERIALS AND METHODS
Cftr(+/−) mice (cftr tm1UNC) on the C57Bl/6J background were bred to obtain littermate wild-type [WT, Cftr(+/+)] and CF [Cftr(−/−)] mice as described (31). Mice were kept in a specific pathogen-free facility in filter-topped cages. Mice of both sexes were used at 6 to 8 weeks of age. To prevent lethal intestinal obstruction, all of the mice were maintained on a complete elemental liquid diet (Peptamen; Nestlé, Deerfield, IL) (31). All of the animal work was approved by the Institutional Animal Care and Use Committee.
Measurement of Gene Expression
Total RNA was prepared from the entire small intestine using Trizol (Invitrogen, Carlsbad, CA) (31). Real-time quantitative reverse transcription-polymerase chain reaction (qRT-PCR) was performed with an iCycler instrument (Bio-Rad, Hercules, CA) using a 1-step qRT-PCR kit (Qiagen, Valencia, CA). The gene-specific primers used are listed in Table 1. For normalization, qRT-PCR of the mRNA for the ribosomal protein Rpl26 was used. Data were analyzed by ΔΔCt (threshold cycle) method with correction for differential PCR efficiencies (28). Data are expressed relative to the WT average. For comparison of the relative abundance of the different transcripts, the average Ct per microgram total RNA for WT samples is given in Table 1 for each primer pair.
Western Blot Analysis and Immunohistochemistry
For Western blots, the small intestine was flushed with phosphate-buffered saline (PBS) then homogenized in 10 mmol/L Tris, pH 7.4 plus protease inhibitors. The DNA content of the samples was determined using a fluorometric assay (34). Appropriate sample loads were used to ensure the Western blot signals were in the linear range. Equal amounts of DNA were separated on sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to polyvinylidene difluoride membranes. The membranes were probed with rabbit anti-Cox1 (No. 160109, Cayman Chemical, Ann Arbor, MI), anti-Cox2 (No. 160106, Cayman Chemical), or an affinity purified rabbit antipeptide antibody to the amino terminus of murine Ltb4dh (custom made by 21st Century Biochemicals, Marlboro, MA). The blots for Cox1 and Ltb4dh were subsequently incubated with goat antirabbit alkaline phosphatase (Sigma, St Louis, MO) and color was developed with 5-bromo-4-chloro-3-indolylphosphate and nitroblue tetrazolium. The anti-peptide antibody to Ltb4dh reacted with a band of the expected size of ∼36 kDa as well as a second band of about 22 kDa. Both bands could be competed by the peptide antigen (data not shown). The smaller protein may represent a splice variant; if it does, then it is expected to be nonfunctional due to absence of most of the dehydrogenase catalytic domain. For Cox2, for which higher sensitivity was required, an avidin–biotin alkaline phosphatase approach was used (Vectastain ABC-AmP kit, Vector Laboratories, Burlingame, CA). The blots were imaged on a flatbed scanner (Hewlett-Packard, Palo Alto, CA) and relative intensities were determined using OptiQuant software (Packard Instrument, Downers Grove, IL). Data were normalized to the averaged WT value set equal to unity.
The same antibodies were used to perform immunohistochemistry on 5-μm paraffin sections of tissue fixed in Carnoy solution (ethanol-acetic acid-chloroform at a ratio of 6:3:1 by volume). After the primary antibody, slides were incubated with mouse-preadsorbed anti-rabbit-biotin secondary antibody (Vector Laboratories). This was followed by use of the Vectastain Elite ABC kit and color development with the VIP peroxidase substrate (Vector Laboratories). All of the samples were processed in parallel under identical conditions. Microscopic images were obtained using identical illumination intensities and exposure times on a Nikon Diaphot microscope equipped with a SPOT RT-KE digital camera (Diagnostic Instruments, Sterling Heights, MI).
Eicosanoid Enzyme Immunoassays
Mice were fasted overnight and killed between noon and 1:00 PM the next day. To avoid variability due to regional differences, the entire small intestine was used. The small intestine was removed and placed in a tray of ice-cold PBS. The mesentery was carefully trimmed to extend the intestine without making any nicks through the intestinal wall. The small intestine lumen was lavaged with 5 mL of ice-cold PBS followed by 5 mL of air. The flushed material was centrifuged at 4°C at 14,000g × 30 minutes. The resulting supernatants were stored at −70°C.
Before use, samples were incubated in a boiling water bath for 10 minutes to inactivate endogenous enzymes (eg, alkaline phosphatase and digestive enzymes), which may interfere with subsequent assays. The boiled samples were centrifuged at 14,000g × 5 minutes to remove precipitates. The resulting supernatants were diluted with enzyme immunoassay (EIA) buffers supplied with the assay kits as needed to ensure signals were within the assay limits. The following EIA kits were purchased from Cayman Chemical: PGE2-metabolites, which uses an alkaline incubation step to convert all metabolites to a single compound; 13,14-dihydro-15-keto-PGF2α (PGF2α metabolite); and 11-β-PGF2α (PGD2 metabolite). The 12(S)-HETE, 15(S)-HETE, and 6-keto-PGF1α (PGI2/prostacyclin metabolite) EIA kits were obtained from Assay Designs (Ann Arbor, MI). The leukotriene B4 (LTB4), cysteinyl-leukotrienes (LT C4/D4/E4), and lipoxin A4 (LXA4) kits were obtained from Oxford Biomedical Research (Ann Arbor, MI). The 20-HETE EIA kit was purchased from Detroit Research & Development (Detroit, MI). Data are expressed as the mass of the specific eicosanoid per milliliter of lavage fluid. Recovery of the lavaged volume was 102% ± 2%, and there was no difference in recovered volumes comparing WT to CF samples.
Data are presented as mean ± standard error (SE). Statistical outliers were identified using Systat software (San Jose, CA) and were omitted from the final analysis. Significance was determined by t test and P < 0.05 was considered significant.
Our previous GeneChip analysis of the CF mouse small intestine showed that among the more than 150 genes with altered expression levels, there were significant changes in several eicosanoid metabolic genes (31). Here, we used qRT-PCR to verify the microarray results and extend the investigation to include other genes involved in metabolism of bioactive eicosanoids (Table 1). We also measured eicosanoid levels.
We first investigated eicosanoid synthetic genes. The AA synthetic gene phospholipase A2, group V (Pla2g5) was expressed at moderate levels in the WT intestine (as indicated by its qRT-PCR Ct value of 25/μg total RNA, Table 1). Pla2g5 expression was upregulated almost 10-fold in the CF small intestine (Fig. 1A, n = 10 WT and 10 CF samples, P < 0.005). The leukotriene synthetic gene Ltc4s was expressed at high levels in the WT tissue (Table 1) and was downregulated by about half in the CF intestine (Fig. 1B, n = 10 WT and 10 CF samples, P < 0.05). The AA ω-hydroxylase Cyp2c40 gene was expressed at moderate levels in the WT intestine (Table 1) and was downregulated by about half in the CF intestine (Fig. 1C, n = 10 WT and 10 CF samples, P < 0.005). Another AA ω-hydroxylase gene, Cyp4a10, also was expressed at moderate levels in the WT intestine (Table 1), but it was dramatically downregulated >90% in the CF tissue (Fig. 1D, n = 10 WT and 10 CF samples, P < 0.005). Cyclooxygenase 1 (Ptgs1/Cox1) mRNA was moderately expressed in WT tissue, whereas Ptgs2/Cox2 was expressed at much lower levels (Table 1). There were modest changes in Cox gene expression in the CF intestine: Ptgs1/Cox1 was slightly and statistically downregulated (Fig. 1E, n = 10 WT and 10 CF samples, P < 0.005) and Ptgs2/Cox2 was slightly upregulated, but the change was not statistically significant (Fig. 1F, n = 10 WT and 10 CF samples, P > 0.05). To achieve a more comprehensive analysis of eicosanoid synthetic genes in the 3 major pathways from AA, we also measured mRNA levels of 5-, 12-, and 15-lipoxygenases (Alox5, Alox12, Alox15), thromboxane A synthase 1 (Tbxas1), and prostaglandin PGE and PGI synthases (Ptges, also known as microsomal PGE synthase-1, mPges-1; Ptges2, also known as microsomal PGE synthase-2, mPges-2; Ptges3, also known as cytosolic PGE synthase, cPges; and Ptgis) all of which were unchanged in the CF intestine as compared with WT (data not shown). Also, prostaglandin D synthase (Ptgds) could not be detected in any intestinal samples, whereas it was readily measured in brain total RNA samples (data not shown).
We next measured expression of 2 major PG degradative genes that were reported as decreased in the CF intestine by the microarray analysis. Both hydroxyprostaglandin dehydrogenase 15 (Hpgd, also known as prostaglandin dehydrogenase or Pgdh) and leukotriene B4 dehydrogenase (Ltb4dh, also known as leukotriene B4 12-hydroxydehydrogenase/15-oxo-prostaglandin 13-reductase, LTB4 HD/PGR) were expressed at high levels in the WT small intestine (Table 1). In the CF intestine, Hpgd expression was downregulated >70% (Fig. 2A, n = 10 WT and 10 CF samples, P < 0.005). There was also a statistically significant downregulation of Ltb4dh in the CF intestine (Fig. 2B, n = 10 WT and 10 CF samples, P < 0.05).
We attempted to confirm at the protein level by Western blot the changes in gene expression presented above. However, the only available antibodies that worked reliably for mouse proteins were against Cox1, Cox2, and the custom-made anti-Ltb4dh. There was not a significant difference in either Cox1 or Cox2 protein levels in the CF intestine as compared with WT (Fig. 3A–D). Additionally, immunohistochemistry for Cox1 and Cox2 did not reveal any differences between WT and CF (data not shown).
By Western blot, Ltb4dh protein was decreased by about 70% in the CF intestine as compared with WT (Fig. 3E–F, n = 10 WT and 10 CF samples, P < 0.01). By immunohistochemistry, Ltb4dh was strongly expressed in the epithelium and muscularis externa layers of the WT small intestine (Fig. 4A, representative of 7 WT). In the CF intestine, the signal was uniformly decreased in all tissue layers (Fig. 4B, representative of 7 CF samples), consistent with the Western blot data.
As a direct test of whether the changes in gene expression result in altered eicosanoid levels, specific eicosanoids were measured in lavage fluid from the intestines of WT and CF mice. When possible, assays were used that measure stable metabolites of these eicosanoids (see Materials and Methods). First, we analyzed lipoxygenase products. LTB4 was measured at high levels and there was not a statistically significant difference comparing WT to CF samples (Fig. 5A, n = 15 WT and 8 CF samples). The cysteinyl-LTs were measured at moderate levels, but again they were not altered in the CF intestine as compared with WT (Fig. 5B, n = 15 WT and 8 CF samples). Lipoxin A4 was at low levels in both WT and CF samples, and there was not a statistically significant difference (Fig. 5C, n = 15 WT and 8 CF samples).
Several cytochrome P450 genes were strongly downregulated in the CF intestine, including some known to be ω-hydroxylases, which metabolize AA to HETEs. There is limited availability of immunoassays to HETEs, so we used the 3 that we could obtain commercially. In the WT small intestine, 12-HETE and 15-HETE were measured at high levels, and 20-HETE was at an extremely high amount (Fig. 6A–C, n = 15 WT and 8 CF samples). In contrast to WT, in the CF intestine all 3 HETEs were strongly decreased: 12-HETE (P < 0.005) and 15-HETE (P < 0.005) were decreased more than 5-fold, and 20-HETE (P < 0.005) was decreased by about 30-fold in the CF intestine, as compared with WT (Fig. 6A–C).
Besides the lipoxygenase and ω-hydroxylase pathways, the other major arm of AA metabolism is that which produces PGs, which were measured next. PGE2, by enzyme immunoassay for a derivative of its metabolic products, was measured at moderate levels in the WT intestine and there was an almost 5-fold increase in the CF intestine (Fig. 7A, P < 0.005). PGF2α, measured as its metabolic product, was also moderately present in the WT intestine, and was increased 6-fold in the CF intestine (Fig. 7B, P < 0.005). The metabolic product of PGD2 was measured at levels near the lower limit of the assay in both the WT and CF samples, and there was not a significant difference (Fig. 7C, P > 0.05). PGI2, measured as its metabolic product, was at moderately high levels in both WT and CF intestine, and there was not a statistical difference (Fig. 7D, P > 0.05).
This study demonstrates that there are several changes in the expression of eicosanoid metabolic genes in the CF mouse small intestine that include both eicosanoid synthetic and degradative genes. The importance of these changes in gene expression were validated in that there were also significant alterations in specific eicosanoid levels, namely increased PG levels and decreased HETE levels. These changes are summarized schematically in Fig. 8A.
The initial step in eicosanoid synthesis is generation of AA from membrane lipids by cytosolic PLA2 (Fig. 8A), and expression of cPla2 was not altered in the CF intestine by microarray analysis (31). However, the secreted phospholipase A2 gene Pla2g5 was strongly upregulated in the CF intestine. Pla2g5 activity potentiates that of cytosolic PLA2, resulting in enhanced AA generation (35).
There are 3 major pathways that convert AA to various active eicosanoids, as shown in Fig. 8A. Expression of Cyp genes that catalyze ω-hydroxylation of AA to HETEs was decreased, especially the expression of Cyp4a10. Enzyme immunoassays showed decreased HETE levels in the CF intestine. We found that 12-HETE and 15-HETE were strongly decreased in the CF mouse intestine. Although the most recognized pathway for 12-HETE synthesis is by 12-lipoxygenase, we did not find decreased Alox12 expression (data not shown). However, there is strong evidence that cytochrome P450 enzymes also can synthesize this eicosanoid. Cytochrome P450 inhibitors were shown to block 12-HETE formation in 3T6 fibroblast cells, but lipoxygenase inhibitors did not (36,37). The specific Cyp involved in 12-HETE production has not yet been identified. Similarly, 15-HETE also is produced by both lipoxygenase- and Cyp-mediated mechanisms (38).
The third HETE we measured is 20-HETE, and it was found to be dramatically decreased in the CF intestine. It had been thought that 20-HETE was produced by the activity of murine Cyp4a10, which is strongly downregulated in the CF mouse small intestine, but the generation of a Cyp4a10 knockout mouse has shown that renal 20-HETE levels are unaffected (39). Recent in vitro transfection studies suggested that the major Cyp responsible for 20-HETE synthesis in the mouse is Cyp4a12a, and that Cyp4a10 is poor at metabolizing AA (40). We were unable to detect an RT-PCR product for Cyp4a12a (NM_177406) using small intestine RNA as template, whereas we measured high levels of expression using kidney RNA (average Ct = 17/μg total RNA; data not shown). It remains unclear what enzyme is responsible for producing high levels of 20-HETE in the WT small intestine. Cyp2c40, for which expression also is downregulated in the CF intestine, generates 16-HETE (41), but there is no available immunoassay for this eicosanoid, so it was not measured.
The HETEs have been most intensively studied in the kidney, where they are important regulators of electrolyte transport and blood pressure (42). The functional significance of decreased HETE levels in the CF intestine is unknown. Because they are often vasoactive, their decrease in the CF mouse would suggest alterations in blood flow through the intestine. A known action of 12-HETE is that it inhibits electrogenic Cl− secretion in the rat colon (43), so its decrease may be an attempt to compensate for loss of the Cftr Cl− channel in the CF intestine.
The other pathway from AA to eicosanoids in which we found differences in the CF intestine is that to PGs. Although there were no major changes in synthetic enzymes that convert AA to PGs, there were decreases in expression of the important PG degradative genes Hpgd and Ltbd4. For Ltb4dh protein, this was confirmed by Western blot and immunohistochemistry of CF intestinal samples. These changes in the PG degradative enzymes were accompanied by significant increases in PGE2 and PGF2α levels in the CF small intestine. Until recently, it had been thought that eicosanoid levels are only controlled by their synthesis because they are rapidly degraded under normal circumstances. It has now been found that the expression levels of eicosanoid degradative enzymes also can be regulated, thus affecting the half-life of PGs. Hpgd and Ltb4dh act in series to degrade PGs, whereas Hpgd catalyzes a reversible reaction and the subsequent action of Ltb4dh is irreversible (44) (Fig. 8B). Both PGE2 and PGF2α are metabolized by Hpgd and Ltb4dh (45,46).
A potential common factor in altered expression of eicosanoid metabolic genes in the CF small intestine is the bacterial product lipopolysaccharide (LPS). The CF mouse small intestine has bacterial overgrowth of gram-negative organisms (29,47), which produce high levels of LPS. LPS can induce Pla2g5 expression in the mouse intestine (48), and LPS also stimulates Pla2g5-dependent PGE2 production by intestinal epithelial cells (49). In a Pla2g5 knockout mouse, activation of macrophages by the yeast component zymosan is attenuated, and PGE2 production is reduced by about 50% (50). A more recent study showed that mast cells from these knockout mice produced less eicosanoids when an innate immune response, mediated by TLR2 and MyD88, was activated by zymosan (35). LPS has been reported to cause downregulation of various Cyp genes (51–53), which may explain the decreased expression of these genes in the CF intestine. Also, systemic administration of LPS to rats strongly decreases expression of the Hpgd gene in liver and lungs (54). The known effects of LPS on expression of eicosanoid metabolic genes provide a link between bacterial overgrowth and the altered levels of eicosanoids in the CF intestine.
Increased PGE2 and PGF2α may have several actions in the CF intestine. PGE2 is known to stimulate mucus secretion (55) and excessive mucus production is characteristic of CF (6), including both the human and mouse CF intestines (47,56). PGE2 also is a relaxant for the circular layer of intestinal smooth muscle (57). Increased PGE2 levels could contribute to the impaired GI motility in many patients with CF (18–24) and to the dramatically decreased small intestinal transit in the CF mouse (28). It is known that endotoxin has a role in small intestinal dysmotility in postsurgical ileus (58,59). Preliminary studies in our laboratory suggest that elevated PGE2 impairs the function of circular enteric smooth muscle in the CF mouse small intestine (60).
Elevated PGE2 also may have important trophic effects in the CF intestine. The HPGD gene is downregulated in inflammatory bowel disease, while PGE2 is an important proinflammatory signal (61). HPGD also is downregulated in colon cancer, and this gene acts as a tumor suppressor by controlling PGE2 levels in the healthy colon (62–65). Less is known about regulation of Ltb4dh expression. A possible consequence of elevated PGE2 is hypertrophy of the intestinal mucosa that occurs in the CF mouse (47,66). Also, intestinal cancer rates are increased in CF, and this is becoming more important as patients with CF live longer (67,68). Our data on the CF mouse intestine is a novel example of a situation in which PG levels are increased in a manner associated with a decrease in PG-degradative enzymes.
Elevated PGF2α in the CF intestine may affect motility. PGF2α is known to cause contraction of both circular and longitudinal layers of intestinal smooth muscle (69). It could be that PGF2α stimulation of the enteric smooth muscle may result in uncoordinated contractions, which contribute to intestinal dysmotility in CF. It also has been shown that anion secretion in the guinea pig colon can be stimulated by PGF2α, so increased PGF2α may be an attempt to improve Cl− secretion in the absence of Cftr.
We showed that Ltb4dh is strongly expressed in the WT epithelium and muscularis layers, and Hpgd has been shown in guinea pig small intestine to be widely expressed, with stronger immunoreactivity in the muscularis as compared with the epithelium (70). Normally, PGs are rapidly metabolized, which limits their range of action. For this reason, PGs are thought of as autocrine or paracrine signals. In the intestine, PGs can be synthesized in all of the tissue layers, including the mucosa and the external smooth muscle layers, and by the normal cellular constituents as well as infiltrating immune cells in the inflamed CF intestine. Because the PG-degradative enzymes are expressed in all of the tissue layers of the small intestine, their downregulation in the CF intestine is expected to cause elevated PG levels in all of the layers. Also, it is possible in the CF intestine that PGs may be able to diffuse farther from their sources and be active at greater distances because of downregulation of the degradative enzymes. Resolution of these possibilities will require further investigation.
Investigations of eicosanoid metabolism similar to that which we performed in the CF mouse have not been done with the human CF intestine. However, because bacterial overgrowth may occur in up to half of all patients with CF, it is expected that these individuals also will have an innate immune response accompanied by changes in eicosanoids in the intestine.
The changes in eicosanoid metabolism can be integrated into the following model for the pathogenesis of CF in the intestine. Abnormal bacterial growth occurs in the mucus that accumulates in the dehydrated CF intestine, which produces LPS and stimulates an innate immune response. This includes upregulation of Pla2g5 and downregulation of PG degradative genes. Pla2g5 amplifies AA production by cytosolic PLA2. Reduced Hpgd and Ltb4dh levels increase the lifetime and perhaps the range of diffusion of PGs. PGE2 can stimulate mucus production and may impair intestinal motility, thereby contributing to pathogenesis of CF in the intestine.
We thank Racquel Sewell for maintaining the CF mouse colony.
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