Cystic fibrosis (CF) is one of the most common life-shortening genetic diseases in whites (http://www.CFF.org). In the absence of the functional cystic fibrosis transmembrane conductance regulator (CFTR) anion channel, affected epithelial surfaces are poorly hydrated and more acidic than normal (1). The altered luminal environment of affected epithelial organs impairs turnover and clearance of the normally protective mucus layer (2), and this situation fosters abnormal bacterial colonization (3). The major cause of mortality in CF is chronic airway infection, which leads to progressive damage and eventual respiratory failure. In addition, CF affects other organs, with important consequences for health and longevity, particularly the gastrointestinal system.
The intestinal tract is involved early in life in CF, and long-term gut dysfunction in CF is apparent as malnutrition. Poor nutrition in CF is strongly associated with airway disease severity and progression (4–7). A major factor of malnutrition in CF is exocrine pancreatic insufficiency; however, even with optimal pancreatic enzyme therapy, nutrition is often not fully corrected (8,9). Also, despite the fact that gene-targeted mouse models of CF are pancreatic sufficient (10,11), their major phenotype is in the intestines and they exhibit poor body weight gain (12). These facts suggest that there are functional defects in the CF small intestine, where digestion and absorption occur.
The CF small intestine exhibits mild inflammation and structural changes to the mucosa (13–15), which may affect digestive and absorptive functions. Such changes may be the result of dysbiosis of the normal enteric microbiota (altered bacterial composition and/or small intestinal bacterial overgrowth [SIBO]). Microbial dysbiosis is a likely consequence of abnormal mucus clearance in the CF intestine. In CF mice, SIBO occurs with colonic-type bacteria that colonize the accumulated mucus (16,17), and CF mice are more susceptible to colonization with pathogenic bacteria (18). Although the evidence is less direct, microbial dysbiosis is likely also common in human CF (19–22). Additionally, the mucosal barrier function of the intestine is compromised in patients with CF as shown by enhanced urinary excretion of orally administered permeability markers and elevated levels of serum albumin in the intestinal lumen (15,23–26). Increased permeability is expected to allow passage of danger signals such as the bacterial component lipopolysaccharide (LPS) from the intestinal lumen into the mucosa, which can trigger inflammatory reactions (27).
Besides the physical barrier comprising the lining epithelium and its mucus covering, there are other mechanisms that contribute to the mucosal barrier function. One of these is the enzyme intestinal alkaline phosphatase (IAP), which protects against LPS from Gram-negative bacteria. IAP dephosphorylates and thereby detoxifies LPS (28). IAP activity in biopsies of human CF duodenum is decreased by 20% to 60% of normal levels (29,30). This is expected to reduce the ability of the CF intestine to detoxify LPS.
We used the Cftr-knockout mouse (Cftrtm1unc, CF mouse) to determine whether this mouse is a good model to investigate intestinal mucosal barrier function in CF. We also tested the hypothesis that interventions known to ameliorate the CF intestinal phenotype would improve the barrier function. The CF mouse has many similarities to human CF with respect to effects on the small intestine. These include excessive luminal mucus accumulation (31,32), SIBO (16,17,20,31), altered innate defenses (18,33), and poor weight gain (9,16).
The CF intestinal phenotype in mice is significantly improved by eradication of SIBO using oral administration of broad-spectrum antibiotics (16,31) or by improving the hydration of the gut lumen with oral osmotic laxative (34). In this work we tested whether the intestinal mucosal barrier function could be improved by antibiotic or laxative treatments, and we also investigated the effects of inhibiting endogenous IAP or supplementation with exogenous IAP on permeability and bacterial load in the small intestine.
MATERIALS AND METHODS
Unless otherwise specified, all of the reagents were from Sigma (St Louis, MO).
Cftrtm1UNC+/− mice were originally obtained from Jackson Laboratories (Bar Harbor, ME). These mice have been bred onto the C57BL/6J background until congenic. They are periodically backcrossed with wild-type (WT) C57BL/6J mice to prevent genetic drift of our colony, and mice used in this study were at generation 31 of backcrossing. Cftrtm1UNC+/− mice were bred to obtain Cftrtm1UNC−/− (CF) and Cftr tm1UNC+/+ (WT) mice. Mice ages 6 to 12 weeks and of both sexes were used; no sex differences in the measured parameters were observed in this study. Cftr tm1UNC+/− mice are phenotypically normal and were used occasionally when needed as WTs; none of the parameters measured in this study were different between Cftr homozygous WT and Cftr heterozygous mice. Unless otherwise indicated, WT and CF mice were fed a liquid diet (Peptamen, Nestle Nutrition, Florham Park, NJ) from weaning, which prevents lethal intestinal obstruction in CF mice. Some mice received broad-spectrum antibiotics added to the liquid diet (ciprofloxacin, 0.05 mg/mL; metronidazole, 0.5 mg/mL) as previously described (16). Some mice received purified calf IAP (Lee Biosolutions, St Louis, MO) (35–38) at 13.3 U/mL in the liquid diet. Another group of mice received the alkaline phosphate (AP) selective inhibitor L-phenylalanine (L-Phe) (39) at 10 mmol/L in the liquid diet. Another group of mice was maintained on standard mouse chow and provided an osmotic laxative (Colyte formulation) in their drinking water (40). Before sacrifice, all of the mice were fasted overnight (<16 hours) with free access to water (supplemented with L-Phe as appropriate) or laxative solution as appropriate. All animal use was submitted to and approved by the University of Kansas Medical Center's institutional animal care and use committee.
Intestinal tissue was fixed in 4% paraformaldehyde overnight followed by paraffin embedding, sectioning, deparaffinization, and rehydration in saline. For conventional histochemistry of IAP, slides were incubated in 0.1 mol/L Tris-HCl, pH 9.5, 5 mmol/L, MgCl2, 0.1 mol/L NaCl containing 0.19 mg/mL 5-bromo-4-chloro-3-indolyl-phosphate and 0.5 mg/mL nitroblue tetrazolium. WT and CF samples were processed in parallel using identical conditions and times of incubation. For histochemistry using LPS as substrate, slides were processed according to Poelstra et al (28). Briefly, slides were incubated with 50 μg/mL LPS and lead nitrate at pH 7.6, plus or minus the selective inhibitor of IAP L-Phe (10 mmol/L) (28). The lead precipitate was converted to a visible product with ammonium sulfide.
Quantitative reverse transcription-polymerase chain reaction (qRT-PCR)
The entire small intestine was flushed with ice-cold saline and the mesentery was trimmed off. The tissue was then processed with TRIzol (Invitrogen, Carlsbad, CA) to isolate total RNA as previously described (16). Real-time qRT-PCR was performed with an iCycler instrument (Bio-Rad, Hercules, CA) with a 1-step RT-PCR kit (Qiagen, Valencia, CA). The following primers were used for Akp3, the mouse IAP gene: forward 5′-CAT GGA CCG CTT CCC ATA-3′ and reverse 5′-CTT GCA CTG TCT GGA ACC TG-3′, product = 72 bp; and for Akp6: forward 5′-AGG ATC CAT CTG TCC TTT GGT-3′ and reverse 5′-CAG CTG CCT TCT TGT TCC A-3′, product = 73 bp. The mRNA for ribosomal protein L26 (Rpl26) was used as a housekeeping gene for normalization as previously described (34). Expression levels were calculated using the ΔΔCt method after correcting for differences in PCR efficiencies, and were expressed relative to WT control levels.
Enzyme Activity Measurement
Tissues were homogenized by sonication on ice in 10 mmol/L Tris, pH 7.0, plus protease inhibitors at 10 mL buffer per gram wet weight of tissue. Alkaline phosphatase activity was measured using di-Tris p-nitrophenyl phosphate (18 mmol/L final) as substrate in 0.1 mol/L Tris-HCl, pH 10, 0.1 mmol/L MgCl2, 0.1 mol/L NaCl buffer (41). The reaction was measured at 405 nm using zero-order kinetics on a synergy HT microplate reader (BioTek, Winooski, VT) at 30°C. Where indicated, the IAP selective inhibitor L-Phe was included in the assay. Alkaline phosphatase activity data are presented as micromoles of product (p-nitrophenol) per minute, normalized to DNA content of the homogenates. DNA was measured using a fluorometric assay (42).
Western Blot of Serum Albumin
Mice were fasted overnight with free access to water. The next morning mice were sacrificed; the entire small intestine was removed and lavaged with 5 mL ice-cold saline. The lavaged fluid was centrifuged to pellet debris and the supernatant was saved. Equal volumes of supernatant (12 μL) were separated on 7.5% sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to polyvinylidene fluoride membrane. The membranes were probed with an antibody to mouse serum albumin (#ab19194; Abcam.com).
Intestinal Permeability Measured In Vivo
Mice were fasted overnight with free access to water or laxative solution as appropriate. In the morning, they were gavaged with a 0.1-mL solution of 1.5% methylcellulose (to mimic the viscosity of digesta) in saline with 25 mg/mL rhodamine-dextran (70 kDa). Ninety minutes later, the mice were sacrificed and blood was collected in EDTA tubes. The samples were centrifuged and plasma was collected. The fluorescence in plasma samples was measured on the plate reader, and concentrations of rhodamine-dextran in plasma were determined using a standard curve of known concentrations of rhodamine-dextran.
Estimation of Bacterial Load
The bacterial 16S rRNA gene was used as an estimate of bacterial load in the small intestine as previously described (34). Briefly, mice were fasted overnight with free access to water. The small intestine was resected and flushed with phosphate-buffered saline containing the mucolytic agent dithiothreitol (10 mmol/L). The flushed material was centrifuged and the pellet was processed to extract bacterial DNA, using the QIAamp DNA Stool Mini Kit (Qiagen, Valencia, CA) with minor modifications as previously reported (16). The DNA was used to amplify the bacterial 16S rRNA gene with universal primers by real-time PCR and quantified by comparison with a standard curve of known amounts of a cloned PCR 16S product (16).
Data are expressed as mean ± SE, and the number of animals used for each group is given in the figure legends. Statistical analysis was by analysis of variance with post-hoc Tukey test using Systat software (SPSS Inc, Chicago, IL).
IAP Activity on the Brush Border Membrane Is Decreased in the CF Intestine
IAP is a brush border enzyme that is most heavily expressed in the proximal small intestine (43). We performed histochemical staining for IAP using conventional reaction conditions (5-bromo-4-chloro-3-indolyl-phosphate as substrate at pH 9.5, which when cleaved by alkaline phosphatase precipitates nitroblue tetrazolium). In the WT mouse, the reaction product was strong in the duodenum (Fig. 1A) on the brush border surface (Fig. 1A′), as expected. Under identical reaction conditions and time of development using CF tissue, the reaction product was still localized to the brush border surface but was noticeably weaker (Fig. 1B, B′) as compared with WT controls (Fig. 1A). To verify others’ work that IAP can use LPS as a substrate, we also performed the reaction at physiological pH (7.6) using LPS as substrate. Again, the product in WT intestine was on the brush border surface (Fig. 1C, C′). We also used the selective IAP inhibitor L-Phe (10 mmol/L) (44) to demonstrate specificity of the reaction using LPS as substrate. As shown in Fig. 1D, L-Phe totally inhibited formation of reaction product in WT intestinal tissue with LPS as substrate.
CF Mice Have Decreased IAP (Akp3) Gene Expression, and Interventions That Improve the CF Phenotype Increase IAP Expression
The gene encoding intestinal alkaline phosphatase is Akp3 and we measured its mRNA levels using qRT-PCR. As shown in Figure 2A, CF control mice express less than one-third as much Akp3 as do WT controls. To further test whether Akp3 expression is associated with the CF intestinal phenotype, we used interventions previously shown to improve intestinal function in CF mice. One of these is oral administration of broad-spectrum antibiotics, which eradicates SIBO and improves several aspects of the CF phenotype (16,31). When CF mice were treated with antibiotics, there was a 3.8-fold increase in Akp3 expression as compared with CF controls (Fig. 2A). When WT mice were treated with antibiotics, there was a 2.1-fold increase in Akp3 mRNA expression as compared with WT controls (Fig. 2A).
Another intervention is use of oral osmotic laxative (Colyte formulation), which better hydrates the gut lumen, preventing intestinal obstruction and allowing CF mice to be maintained on standard solid chow (40). Laxative treatment was shown to improve several aspects of the CF intestinal phenotype (34), so we tested its effects on Akp3 gene expression. Treatment of CF mice with laxative increased Akp3 expression more than 5-fold as compared with CF controls (Fig. 2A). When WT mice were maintained on laxative there was a 1.9-fold increase in Akp3 expression as compared with WT controls (Fig. 2A).
Recently, a second intestine-specific alkaline phosphatase gene was discovered in mice, Akp6, whose expression is increased in Akp3-knockout mice (43). To determine whether there are any compensatory changes in Akp6 expression in control CF mice or after experimental interventions, we measured its expression levels using qRT-PCR. In control WT mouse small intestine, Akp6 expression was 6.5-fold less than that of Akp3, based on comparison of Ct values; the PCR efficiencies for the 2 genes were identical (data not shown). There was no difference in Akp6 expression comparing control CF to control WT mice (Fig. 2B). After treatment with antibiotics, there was a small but significant increase in Akp6 expression in treated WT mice as compared with control (Fig. 2B). Antibiotic treatment did not change Akp6 expression in CF mice (Fig. 2B). In mice treated with laxative, there was no significant change in Akp6 expression in either WT or CF mice (Fig. 2B).
CF Mice Have Decreased IAP Enzyme Activity, and Interventions That Improve the CF Phenotype Increase IAP Activity
We next measured IAP enzyme activity to compare expression levels of Akp3 mRNA to actual enzyme activity. Specific IAP activity was considered to be that which was sensitive to inhibition by 10 mmol/L L-Phe (44). As expected, IAP activity was strongest in the proximal small intestine, with little activity in the WT mouse after the first tenth of the intestine (Fig. 3A). IAP activity in the CF intestine also was similarly localized to the first tenth of the intestine, but it was less than one-third of the WT control activity (Fig. 3A), essentially the same as the respective mRNA levels. In CF mice treated with antibiotics, IAP enzyme levels were now the same as in WT control mice (Fig. 3B), which is consistent with the increase in Akp3 mRNA in antibiotic-treated CF mice (Fig. 2A).
Antibiotic treatment of WT mice did not significantly affect IAP enzyme levels as compared with WT controls (Fig. 3B). Laxative treatment of CF mice increased IAP activity to a level that is comparable to that of control WT mice (Fig. 3C), similar to the effect of laxative on Akp3 mRNA in the CF intestine (Fig. 2A). When WT mice were treated with laxative, there was a small decrease in IAP activity as compared with WT controls (Fig. 3C), in contrast to the significant increase observed in Akp3 mRNA levels in laxative-treated WT mice (Fig. 2A).
To determine whether a non-IAP AP activity was increased in the control CF intestine or after the specific interventions, we also calculated the L-Phe insensitive activity. As shown in Fig. 4, there was little L-Phe insensitive activity in either the WT or CF small intestine under control or experimental conditions. Because the enzyme encoded by Akp6 has not been characterized biochemically, it is not known whether this enzyme is L-Phe sensitive, and we could not investigate its enzymatic activity in WT and CF mice.
CF Mice Have Increased Intestinal Permeability, and Interventions That Improve the CF Phenotype Decrease Permeability
As an indication of increased intestinal permeability, the presence of serum albumin in the luminal content of the small intestine was assessed by Western blot. As shown in Figure 5, there was more immunoreactive serum albumin in intestinal lavage fluid from CF mice as compared with WT. Note that there were several reactive bands smaller than the expected size of intact serum albumin, indicating proteolytic activity in the lumen of the intestine of both WT and CF mice, which are pancreatic sufficient. When this antibody was used on mouse plasma, a single immunoreactive band at the expected location was observed (data not shown). To determine intestinal permeability more quantitatively, we measured passage of a nondigestible fluorescent tracer (rhodamine-dextran) from the intestine into the blood circulation. Mice were fasted overnight to empty the gastrointestinal tract followed by gavage of the tracer into the stomach. After 90 min the mice were sacrificed and blood was collected to measure levels of fluorescence as an indicator of intestinal permeability. Fluorescence in the plasma of CF mice was about 5-fold greater than WT mice (Fig. 6).
When CF mice were treated with broad-spectrum antibiotics to eradicate SIBO, plasma fluorescence after gavage was reduced as compared with CF controls and this difference was borderline significant (P = 0.07 vs CF control). The plasma fluorescence of antibiotic-treated CF mice was not significantly different compared with WT mice (P = 0.22 vs antibiotic-treated WT) (Fig. 6). Antibiotic treatment of WT mice did not affect plasma fluorescence after gavage compared with WT control (Fig. 6).
We next tested the effect of laxative, which aids normal hydration of the CF gut lumen and improves several aspects of the CF intestinal phenotype, for potential effects on intestinal permeability. When CF mice were treated with laxative, the amount of plasma fluorescence after gavage was less than half that of CF controls (P = 0.045; Fig. 6). This value was not significant as compared with WT plasma fluorescence (P = 0.73 vs laxative-treated WT; Fig. 6). When WT mice were treated with oral laxative solution, there was no change in plasma fluorescence as compared with WT control (Fig. 6).
IAP Is Not Directly Related to Intestinal Permeability
To gain insight into whether IAP has a direct role in intestinal permeability, 2 experimental manipulations were used. First, exogenous-purified calf IAP was added to the liquid diet for 3 weeks followed by measurement of intestinal permeability. Exogenous IAP has been shown to be protective in experimental colitis (35,37) and necrotizing enterocolitis (38) and in human patients with inflammatory bowel disease (36). In WT mice exogenous IAP caused a nonsignificant decrease in permeability (Fig. 6). In CF mice, there was a significant decrease in permeability after IAP treatment, and the level of fluorescence in the plasma was not significantly different from the WT group (P = 0.79 vs WT IAP treated) (Fig. 6).
Second, to test whether loss of IAP activity results in increased permeability, endogenous IAP activity was inhibited by addition of the IAP-selective inhibitor L-Phe to the liquid diet for 3 weeks. Previous work showed that oral L-Phe inhibited IAP activity and increased the severity of experimental colitis (39). In mice treated with L-Phe, there was no significant change in permeability in either WT or CF mice (Fig. 6). Interestingly, 2 of the 6 CF mice administered L-Phe died within 1 week of starting the treatment, whereas none of the 9 L-Phe-treated WT mice died. The postweaning death rate of CF control mice on Peptamen is <1 of 5 that observed in the L-Phe-treated mice; only 2 of 31 CF control mice in recent litters died after weaning on Peptamen, and no WT mice died.
Exogenous IAP Affects Bacterial Load
Because it was recently reported that the fecal microbiota is altered in IAP-deficient Akp3-knockout mice (45) and CF mice have small intestinal bacterial overgrowth, it was of interest to measure bacterial load in the small intestine after experimentally manipulating IAP levels. In WT mice treated with oral exogenous IAP, there was no significant effect on the normal low bacterial load in the small intestine (Fig. 7). In contrast, in IAP-treated CF mice, there was >80% decrease in bacterial load (Fig. 7). Administration of the IAP inhibitor L-Phe had no significant effect on bacterial load in either WT or CF mice (Fig. 7).
In the present study, we set out to determine whether the CF mouse was a suitable model to investigate altered mucosal barrier function of the CF intestine and whether interventions known to improve the CF intestinal phenotype would also improve barrier function. We demonstrated that CF mice have impairments of the intestinal mucosal barrier that are similar to those reported in human patients with CF. We show that oral broad-spectrum antibiotics or osmotic laxative, interventions that improve the CF intestinal phenotype, also improve mucosal barrier function. Also, administration of exogenous IAP to CF mice improved intestinal permeability and reduced small intestinal bacterial overgrowth >80%.
Investigation of the LPS detoxifying enzyme IAP showed that IAP activity was correctly localized to the brush border of villus enterocytes in the CF mouse, but the strength of the reaction was reduced as compared with WT. Expression of the murine IAP gene (Akp3) in the CF mouse intestine was also reduced to less than one-third of WT. This was paralleled by an equal decrease in IAP enzyme activity. The regulation of IAP expression is complex and is affected by many conditions in the gut (46). Changes in IAP expression are not directly linked to Cftr. IAP levels can be brought back to normal in the CF mouse by antibiotics or laxative, so the decrease in IAP in CF is not a direct consequence of loss of Cftr, but rather is secondary to its loss. A common result in the CF intestine because of oral antibiotics and laxative treatments is that both eradicate bacterial overgrowth. Hence, it is proposed that as part of the intestinal response to bacterial overgrowth IAP expression is decreased in the CF intestine.
Because a second intestine-specific alkaline phosphatase gene, Akp6, was recently discovered whose expression is increased in Akp3-knockout mice (43), we also measured RNA levels for this gene in CF mice with and without the interventions used here. There was a modest increase in Akp6 mRNA levels in antibiotic-treated WT mice (175% of WT control) but no changes in CF mice under any conditions. Because the Akp6-encoded enzyme has not been characterized, it is not known whether it is L-Phe sensitive and it was not possible to measure its enzymatic activity. In any case, unlike the Akp3-knockout mouse, Akp6 gene expression is not significantly different in the CF mouse, which has a deficiency in Akp3 expression and activity.
Two situations that may be relevant to the CF phenotype that have been shown to reduce IAP expression are inflammation and severe malnutrition. The CF intestine has mild inflammation that could contribute to decreased IAP expression. It is known that inflammation of the gut decreases IAP expression, and this can be mediated by the inflammatory cytokines interleukin (IL)-1β and TNF-α (47). When CF mice are treated with antibiotics to eradicate SIBO, mast cell and neutrophil infiltration is reduced and expression of innate immune markers is more normal (16). Also, when treated with laxative, CF mice have normal numbers of small intestinal bacteria and, again, the innate immune changes are ameliorated (34). Although there are many changes in innate immune markers in the CF small intestine, our previous microarray study did not show changes in IL-1β or TNF-α (33). Therefore, it is uncertain whether the changes observed in IAP expression are related to the mild immune response in the CF mouse intestine.
An alternative possibility is that decreased IAP in the CF intestine is a result of malnutrition. It has been shown in rodents that prolonged fasting or starvation strongly decreases IAP expression (48,49). Also, early weaning in pigs, which involves nutritional stress, results in decreased IAP levels (50). CF mice receiving the liquid diet are about 70% the weight of age- and diet-matched WT mice (16); however, the mechanism linking nutrition to IAP expression is unknown, and although CF mice grow poorly, it is not clear whether this degree of malnutrition is sufficient to result in the strong decrease in IAP in CF.
In addition to its ability to detoxify LPS, IAP has been shown to have other important physiological functions (46). One is that IAP dephosphorylates luminal ATP, thereby contributing to a feedback loop controlling P2Y1 purinergic signaling and influencing bicarbonate secretion in the intestine (51). Thus, a consequence of decreased IAP in the intestine would be to increase bicarbonate secretion; however, bicarbonate secretion in the intestine is CFTR-dependent, so a decrease in IAP in CF is not productive in this respect. An additional consideration is that ATP in high enough concentrations is considered an endogenous danger signal that has proinflammatory effects (52). Thus, a decrease in IAP activity in the CF intestine may contribute to an inflammatory process.
Another role of IAP is in dietary lipid assimilation, and IAP-deficient (Akp3 null) mice show enhanced lipid uptake (53). In CF, malnourishment is a significant problem and fat maldigestion and malabsorption in particular are common (9). Poor fat assimilation also occurs in CF mice (10). A decrease in IAP in the CF intestine may be an adaptive change in an attempt to increase dietary lipid assimilation. How IAP participates in fat assimilation is not well understood, and whether a deficiency in dietary fat assimilation is important in regulating Akp3 gene expression is unknown.
The other aspect of the mucosal barrier we investigated in the CF mouse was permeability of the gut to macromolecules. We observed increased amounts of serum albumin in the lumen of the small intestine and greater passage of fluorescent dextran from the lumen to the blood in CF mice as compared with WT. After antibiotic treatment of CF mice, passage of the fluorescent dextran from the intestine into the blood was reduced by about half. A similar degree of reduced permeability occurred in laxative-treated CF mice. An even greater decrease in permeability was observed in CF mice provided exogenous IAP, and these mice also had a >80% reduction in small intestinal bacterial overgrowth. Impaired intestinal permeability appears to be a ubiquitous feature of intestinal dysfunction and it occurs even with mild inflammation (54). Additionally, increased permeability is a common feature of microbial dysbiosis. In patients with SIBO with colonic-type bacteria, there is increased intestinal permeability (55), and CF mice also have SIBO with predominantly colonic-type bacteria (16,17). Because the interventions we used dramatically decrease bacterial load in CF mice (16,34), it is suggested that signals from the microbial dysbiosis in CF affect the epithelium to make it more permeable.
Although IAP affects the gut microbiota (45), how it does so is unknown. A possible mechanism of the effects of exogenous IAP in the CF intestine could involve a decrease in bacterial LPS, which in turn reduces stimulation of immune responses. Previous work showed that eradication of bacterial overgrowth in the CF mouse reduced mucus accumulation (31,34), and bacteria colonize this mucus. Thus, it is possible that bacterial LPS increases mucus production, which in turn enhances the niche for bacterial growth. More work is needed to investigate the effects of exogenous IAP on the CF intestinal phenotype.
An interesting question is the extent to which IAP levels and epithelial permeability are causally related. Although IAP is considered part of the mucosal defenses of the intestine because of its LPS detoxifying activity, there are also data suggesting that IAP is more directly involved in the physical barrier of the epithelium. After ischemia reperfusion, IAP-deficient mice exhibit significantly increased bacterial translocation across the gut wall to mesenteric lymph nodes (49). It has not been reported whether epithelial permeability to tracer macromolecules is altered in the IAP-deficient mouse. To address this, we used the IAP inhibitor L-Phe, added to the liquid diet. Previous work showed that oral L-Phe increased passage of gavaged LPS into blood in rats (56) and increased the severity of experimental colitis induced with dextran sodium sulfate in mice (39). When we treated mice with oral L-Phe, intestinal permeability was not affected in either WT or CF mice. Therefore, it appears that IAP is not required for normal low permeability, and that its inhibition alone is not sufficient to increase permeability. There was more than a 5-fold increase in postweaning deaths of L-Phe-treated CF mice as compared with CF controls. This was observed in a small sample size, but it may indicate the importance of the residual IAP activity in the CF intestine in which increased permeability is also present. Further work is needed to address this issue.
In summary, CF mice, like human patients with CF, have significantly decreased IAP levels and increased intestinal permeability. Interventions that improve other aspects of the CF intestinal phenotype also increase IAP levels and decrease intestinal permeability. In the untreated CF mouse small intestine, there is SIBO with Gram-negative bacteria, decreased IAP activity, and increased permeability. These conditions are expected to allow greater biologically active LPS to become systemic in CF, which may have a significant effect on distant sites such as the airways. Another important novel finding was that exogenous IAP reduced permeability in the CF intestine to WT levels and reduced bacterial overgrowth >80%. Thus, exogenous IAP may be a new therapy for CF intestinal disease.
Because systemic exposure to gut bacterial products such as LPS can affect distant organs, our work suggests that therapies that improve the mucosal barrier function of the intestine could have broader beneficial effects such as lessening airway inflammation in CF.
1. Barraclough M, Taylor CJ. Twenty-four hour ambulatory gastric and duodenal pH profiles in cystic fibrosis: effect of duodenal hyperacidity on pancreatic enzyme function and fat absorption. J Pediatr Gastroenterol Nutr 1996; 23:45–50.
2. Quinton PM. Cystic fibrosis: impaired bicarbonate secretion and mucoviscidosis. Lancet 2008; 372:415–417.
3. Boucher RC. Airway surface dehydration in cystic fibrosis: pathogenesis and therapy. Annu Rev Med 2007; 58:157–170.
4. Houwen RH, van der Doef HP, Sermet I, et al. Defining DIOS and constipation in cystic fibrosis with a multicentre study on the incidence, characteristics, and treatment of DIOS. J Pediatr Gastroenterol Nutr 2010; 50:38–42.
5. Courtney JM, Bradley J, McCaughan J, et al. Predictors of mortality in adults with cystic fibrosis. Pediatr Pulmonol 2007; 42:525–532.
6. Milla CE. Association of nutritional status and pulmonary function in children with cystic fibrosis. Curr Opin Pulm Med 2004; 10:505–509.
7. Stallings VA, Stark LJ, Robinson KA, et al. Evidence-based practice recommendations for nutrition-related management of children and adults with cystic fibrosis and pancreatic insufficiency: results of a systematic review. J Am Diet Assoc 2008; 108:832–839.
8. Baker SS, Borowitz D, Duffy L, et al. Pancreatic enzyme therapy and clinical outcomes in patients with cystic fibrosis. J Pediatr 2005; 146:189–193.
9. Borowitz D, Durie PR, Clarke LL, et al. Gastrointestinal outcomes and confounders in cystic fibrosis. J Pediatr Gastroenterol Nutr 2005; 41:273–285.
10. Bijvelds MJ, Bronsveld I, Havinga R, et al. Fat absorption in cystic fibrosis mice is impeded by defective lipolysis and post-lipolytic events. Am J Physiol Gastrointest Liver Physiol 2005; 288:G646–G653.
11. De Lisle RC, Isom KS, Ziemer D, et al. Changes in the exocrine pancreas secondary to altered small intestinal function in the CF mouse. Am J Physiol Gastrointest Liver Physiol 2001; 281:G899–G906.
12. Grubb BR, Boucher RC. Pathophysiology of gene-targeted mouse models for cystic fibrosis. Physiol Rev 1999; 79:S193–S214.
13. Werlin SL, uri-Silbiger I, Kerem E, et al. Evidence of intestinal inflammation in patients with cystic fibrosis. J Pediatr Gastroenterol Nutr 2010; 51:304–308.
14. Raia V, Maiuri L, De Ritis G, et al. Evidence of chronic inflammation in morphologically normal small intestine of cystic fibrosis patients. Pediatr Res 2000; 47:344–350.
15. Hallberg K, Grzegorczyk A, Larson G, et al. Intestinal permeability in cystic fibrosis in relation to genotype. J Pediatr Gastroenterol Nutr 1997; 25:290–295.
16. Norkina O, Burnett TG, De Lisle RC. Bacterial overgrowth in the cystic fibrosis transmembrane conductance regulator null mouse small intestine. Infect Immun 2004; 72:6040–6049.
17. Canale-Zambrano JC, Auger ML, Haston CK. Toll-like receptor-4 genotype influences the survival of cystic fibrosis mice. Am J Physiol Gastrointest Liver Physiol 2010; 299:G381–G390.
18. Clarke LL, Gawenis LR, Bradford EM, et al. Abnormal paneth cell granule dissolution and compromised resistance to bacterial colonization in the intestine of CF mice. Am J Physiol Gastrointest Liver Physiol 2004; 286:G1050–G1058.
19. Lisowska A, Wojtowicz J, Walkowiak J. Small intestine bacterial overgrowth is frequent in cystic fibrosis: combined hydrogen and methane measurements are required for its detection. Acta Biochim Pol 2009; 56:631–634.
20. Fridge JL, Conrad C, Gerson L, et al. Risk factors for small bowel bacterial overgrowth in cystic fibrosis. J Pediatr Gastroenterol Nutr 2007; 44:212–218.
21. Lewindon PJ, Robb TA, Moore DJ, et al. Bowel dysfunction in cystic fibrosis: importance of breath testing. J Paediatr Child Health 1998; 34:79–82.
22. O’Brien S, Mulcahy H, Fenlon H, et al. Intestinal bile acid malabsorption in cystic fibrosis. Gut 1993; 34:1137–1141.
23. Reims A, Strandvik B, Sjovall H. Epithelial electrical resistance as a measure of permeability changes in pediatric duodenal biopsies. J Pediatr Gastroenterol Nutr 2006; 43:619–623.
24. Dalzell AM, Freestone NS, Billington D, et al. Small intestinal permeability and orocaecal transit time in cystic fibrosis. Arch Dis Child 1990; 65:585–588.
25. Escobar H, Perdomo M, Vasconez F, et al. Intestinal permeability to 51Cr-EDTA and orocecal transit time in cystic fibrosis. J Pediatr Gastroenterol Nutr 1992; 14:204–207.
26. Hendriks HJ, van Kreel B, Forget PP. Effects of therapy with lansoprazole on intestinal permeability and inflammation in young cystic fibrosis patients. J Pediatr Gastroenterol Nutr 2001; 33:260–265.
27. Purohit V, Bode JC, Bode C, et al. Alcohol, intestinal bacterial growth, intestinal permeability to endotoxin, and medical consequences: summary of a symposium. Alcohol 2008; 42:349–361.
28. Poelstra K, Bakker WW, Klok PA, et al. Dephosphorylation of endotoxin by alkaline phosphatase in vivo. Am J Pathol 1997; 151:1163–1169.
29. Van Biervliet S, Eggermont E, Carchon H, et al. Small intestinal brush border enzymes in cystic fibrosis. Acta Gastroenterol Belg 1999; 62:267–271.
30. Van Biervliet S, Eggermont E, Marien P, et al. Combined impact of mucosal damage and of cystic fibrosis on the small intestinal brush border enzyme activities. Acta Clin Belg 2003; 58:220–224.
31. De Lisle RC, Roach EA, Norkina O. Eradication of small intestinal bacterial overgrowth in the cystic fibrosis mouse reduces mucus accumulation. J Pediatr Gastroenterol Nutr 2006; 42:46–52.
32. Jeffrey I, Durrans D, Wells M, et al. The pathology of meconium ileus equivalent. J Clin Pathol 1983; 36:1292–1297.
33. Norkina O, Kaur S, Ziemer D, et al. Inflammation of the cystic fibrosis mouse small intestine. Am J Physiol Gastrointest Liver Physiol 2004; 286:G1032–G1041.
34. De Lisle RC, Roach E, Jansson K. Effects of laxative and N-acetylcysteine on mucus accumulation, bacterial load, transit, and inflammation in the cystic fibrosis mouse small intestine. Am J Physiol Gastrointest Liver Physiol 2007; 293:G577–G584.
35. Bol-Schoenmakers M, Fiechter D, Raaben W, et al. Intestinal alkaline phosphatase contributes to the reduction of severe intestinal epithelial damage. Eur J Pharmacol 2010; 633:71–77.
36. Lukas M, Drastich P, Konecny M, et al. Exogenous alkaline phosphatase for the treatment of patients with moderate to severe ulcerative colitis. Inflamm Bowel Dis 2010; 16:1180–1186.
37. Ramasamy S, Nguyen DD, Eston MA et al. Intestinal alkaline phosphatase has beneficial effects in mouse models of chronic colitis. Inflamm Bowel Dis 2010;17:532–42.
38. Whitehouse JS, Riggle KM, Purpi DP, et al. The protective role of intestinal alkaline phosphatase in necrotizing enterocolitis. J Surg Res 2010; 163:79–85.
39. Campbell EL, Macmanus CF, Kominsky DJ et al. Resolvin E1-induced intestinal alkaline phosphatase promotes resolution of inflammation through LPS detoxification. Proc Natl Acad Sci U S A 2010;107:14298–303.
40. Clarke LL, Gawenis LR, Franklin CL, et al. Increased survival of CFTR knockout mice with an oral osmotic laxative. Lab Anim Sci 1996; 46:612–618.
41. Bowers GN Jr, McComb RB. A continuous spectrophotometric method for measuring the activity of serum alkaline phosphatase. Clin Chem 1966; 12:70–89.
42. Cesarone CF, Bolognesi C, Santi L. Improved microfluorometric DNA determination in biological material using 33 258 Hoechst. Anal Biochem 1979; 100:188–197.
43. Narisawa S, Hoylaerts MF, Doctor KS, et al. A novel phosphatase upregulated in Akp3 knockout mice. Am J Physiol Gastrointest Liver Physiol 2007; 293:G1068–G1077.
44. Shephard MD, Peake MJ, Walmsley RN. Quantitative method for determining serum alkaline phosphatase isoenzyme activity. II. Development and clinical application of method for measuring four serum alkaline phosphatase isoenzymes. J Clin Pathol 1986; 39:1031–1038.
45. Malo MS, Alam SN, Mostafa G, et al. Intestinal alkaline phosphatase preserves the normal homeostasis of gut microbiota. Gut 2010; 59:1476–1484.
46. Lalles JP. Intestinal alkaline phosphatase: multiple biological roles in maintenance of intestinal homeostasis and modulation by diet. Nutr Rev 2010; 68:323–332.
47. Malo MS, Biswas S, Abedrapo MA, et al. The pro-inflammatory cytokines, IL-1beta and TNF-alpha, inhibit intestinal alkaline phosphatase gene expression. DNA Cell Biol 2006; 25:684–695.
48. Hodin RA, Chamberlain SM, Meng S. Pattern of rat intestinal brush-border enzyme gene expression changes with epithelial growth state. Am J Physiol 1995; 269:C385–C391.
49. Goldberg RF, Austen WG Jr, Zhang X, et al. Intestinal alkaline phosphatase is a gut mucosal defense factor maintained by enteral nutrition. Proc Natl Acad Sci U S A 2008; 105:3551–3556.
50. Lackeyram D, Yang C, Archbold T, et al. Early weaning reduces small intestinal alkaline phosphatase expression in pigs. J Nutr 2010; 140:461–468.
51. Akiba Y, Mizumori M, Guth PH, et al. Duodenal brush border intestinal alkaline phosphatase activity affects bicarbonate secretion in rats. Am J Physiol Gastrointest Liver Physiol 2007; 293:G1223–G1233.
52. Ishii KJ, Akira S. Potential link between the immune system and metabolism of nucleic acids. Curr Opin Immunol 2008; 20:524–529.
53. Nakano T, Inoue I, Koyama I, et al. Disruption of the murine intestinal alkaline phosphatase gene Akp3 impairs lipid transcytosis and induces visceral fat accumulation and hepatic steatosis. Am J Physiol Gastrointest Liver Physiol 2007; 292:G1439–G1449.
54. Bjarnason I, Takeuchi K, Bjarnason A, et al. The G.U.T. of gut. Scand J Gastroenterol 2004; 39:807–815.
55. Riordan SM, McIver CJ, Thomas DH, et al. Luminal bacteria and small-intestinal permeability. Scand J Gastroenterol 1997; 32:556–563.
56. Koyama I, Matsunaga T, Harada T, et al. Alkaline phosphatases reduce toxicity of lipopolysaccharides in vivo and in vitro through dephosphorylation. Clin Biochem 2002; 35:455–461.