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ScienceDirect Journal of Nutritional Biochemistry 26 (2015) 36 – 43

Polyunsaturated fatty acid supplementation reverses cystic fibrosis-related fatty acid abnormalities in CFTR −/− mice by suppressing fatty acid desaturases☆ Sarah W. Njoroge, Michael Laposata, Kelli L. Boyd, Adam C. Seegmiller⁎ Department of Pathology, Microbiology, and Immunology, Vanderbilt University School of Medicine, Nashville, TN, USA

Received 3 April 2014; received in revised form 7 August 2014; accepted 2 September 2014

Abstract Cystic fibrosis patients and model systems exhibit consistent abnormalities in metabolism of polyunsaturated fatty acids that appear to play a role in disease pathophysiology. Recent in vitro studies have suggested that these changes are due to overexpression of fatty acid desaturases that can be reversed by supplementation with the long-chain polyunsaturated fatty acids docosahexaenoate and eicosapentaenoate. However, these findings have not been tested in vivo. The current study aimed to test these results in an in vivo model system, the CFTR−/− knockout mouse. When compared with wild-type mice, the knockout mice exhibited fatty acid abnormalities similar to those seen in cystic fibrosis patients and other model systems. The abnormalities were confined to lung, ileum and pancreas, tissues that are affected by the disease. Similar to in vitro models, these fatty acid changes correlated with increased expression of Δ5- and Δ6desaturases and elongase 5. Dietary supplementation with high-dose free docosahexaenoate or a combination of lower-dose docosahexaenoate and eicosapentaenoate in triglyceride form corrected the fatty acid abnormalities and reduced expression of the desaturase and elongase genes in the ileum and liver of knockout mice. Only the high-dose docosahexaenoate reduced histologic evidence of disease, reducing mucus accumulation in ileal sections. These results provide in vivo support for the hypothesis that fatty acid abnormalities in cystic fibrosis result from abnormal expression and activity of metabolic enzymes in affected cell types. They further demonstrate that these changes can be reversed by dietary n-3 fatty acid supplementation, highlighting the potential therapeutic benefit for cystic fibrosis patients. © 2015 Published by Elsevier Inc. Keywords: Cystic fibrosis; Fatty acid; Desaturase; Docosahexaenoate; Eicosapentaenoate; Arachidonate

1. Introduction Cystic fibrosis (CF) is a common inherited disorder, with a prevalence of approximately 1 in 3000 among Caucasian Americans [1]. CF is caused by mutations in the gene encoding the CF transmembrane regulator (CFTR) protein, an anion channel predominantly expressed by epithelial cells [2]. The clinical features of CF are protean, but the most common and severe effects include exocrine pancreatic insufficiency, malabsorption and malnutrition, intestinal obstruction, and chronic pulmonary obstruction, infection and bronchiectasis [1]. Patients have shortened average life spans, averaging approximately 40 years in the United States. Beginning in the 1960s [3], a series of studies demonstrated consistent alterations in levels of polyunsaturated fatty acids (PUFAs) in CF patients when compared with healthy controls [reviewed in



Funding sources: Edward and Nancy Fody Endowed Chair in Pathology (M.L.) and the Vanderbilt Physician Scientist Training Program (A.C.S.). ⁎ Corresponding author. Department of Pathology, Microbiology, and Immunology, Vanderbilt University School of Medicine, 4918B TVC, 1301 Medical Center Dr., Nashville, TN 37027, USA. E-mail address: [email protected] (A.C. Seegmiller). http://dx.doi.org/10.1016/j.jnutbio.2014.09.001 0955-2863/© 2015 Published by Elsevier Inc.

Refs. [4–6]]. The most common of these changes are decreased linoleate (18:2n-6; LA) and docosahexaenoate (22:6n-3; DHA). Some studies have also shown increased arachidonate (20:4n-6; AA). These changes are independent of absorption status [7], and the degree of PUFA alteration correlates with the severity of disease [8–13]. Animal models have been useful in understanding disease mechanisms in CF. At least 13 mouse models of CF have been generated [14–16]. PUFA alterations similar to those of CF patients have been noted in the tissues of at least two of these, the CFTRtm1/UNC knockout model [17,18] and the CFTRtm1/EUR model [19] that carries a ΔF508 mutation, although these changes have not been consistent in every study [20,21]. In both of these models, dietary supplementation with large doses of free DHA has been successful in reversing the characteristic PUFA alterations [17,19]. In mixed background CFTRtm1/UNC mice model, such supplementation also ameliorated CF-related histopathologic features, including pancreatic duct dilation, intestinal villus hypertrophy and increased pulmonary neutrophils in response to inflammatory stimulus [17], although these effects were not seen in mice with a congenic C57BL/6J background [22]. However, the clinical application of these studies is questionable, as the dose is extremely high, the duration is short and the formulation (free fatty acid) is not amenable to clinical use. The underlying mechanisms responsible for these PUFA alterations in CF, including the relationship between CFTR mutations and

S.W. Njoroge et al. / Journal of Nutritional Biochemistry 26 (2015) 36–43

A

37

B 18:2n-6 (LA)

3.0

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0.8

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0.0

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CF

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75

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60

6

P=.005

P=.041

45

4

30

2

15

n.s.

P=.001

0

CF

Δ5D

Δ6D

0

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EL5

Fig. 1. Expression and activity of fatty acid desaturases in n-6 PUFA metabolic pathway in CF mice. (A) Schematic diagram of the n-6 PUFA metabolic pathway, including pertinent enzymes, through which LA is converted to AA. (B) Activity of the n-6 PUFA pathway in selected mouse tissues. WT (CFTR+/+) and CF (CFTR−/−) mice were produced, maintained on a Peptamen liquid diet post-weaning for 14 days, and then sacrificed as described in Methods and materials. Lung, ileum and pancreas tissues were harvested, processed, and total tissue fatty acids measured as described in Methods and materials. Relative activity of the n-6 PUFA pathway was estimated using the ratio of product (20:4n-6; AA) to substrate (18:2n-6; LA). Ratios for individual mice (n=6 or 7) are shown with the mean indicated by a horizontal line. P values were determined by t test. (C) RNA was harvested from lung and ileum of WT and CF mice generated and processed as in (B) above. Relative mRNA level of Δ5-desaturase (Δ5D), Δ6-desaturase (Δ6D) and elongase 5 (EL5) genes was determined by quantitative RT-PCR as described in Methods and materials. Bars indicate mean±SEM of six or seven animals. P values were determined by Mann–Whitney rank-sum test. n.s., not significant.

PUFA metabolism, have been largely unknown. However, there have been recent advances utilizing cell culture models of CF that exhibit the characteristic PUFA abnormalities [23,24]. These studies demonstrated clear correlations between PUFA abnormalities and the expression and activity of fatty acid metabolic enzymes [25–28]. In particular, increased Δ5- and Δ6-desaturase stimulate metabolism of LA to AA in the n-6 pathway (Fig. 1), leading to decreased LA and increased AA in CF compared to wild-type cells [25]. Importantly, these changes were reversed by DHA supplementation, correcting the metabolic abnormalities [28]. The application of these findings was limited, however, because they came from an in vitro system and it was not clear whether they would translate to a more complex in vivo model. Accordingly, the current study investigates the mechanisms of PUFA metabolic abnormalities in the CFTRtm1/UNC mouse model of CF. To compare with previous studies, we correlated fatty acid levels and gene expression at baseline and after dietary supplementation with high-dose DHA given as a free fatty acid (as in Ref. [17]). However, to improve clinical relevance, we repeated the studies with an existing clinical formulation of lower-dose fish oil-derived DHA plus eicosapentaenoic acid (20:5n-3; EPA) in predominantly triglyceride form for a longer duration. We observed that the typical PUFA alterations are restricted to tissues that express high levels of CFTR and that these abnormalities are tightly correlated to increased expression of Δ5and Δ6-desaturases and elongase 5. PUFA abnormalities are reversed by DHA supplementation, which suppresses expression of these enzymes in intestine and liver, but not in lung. Lower-dose DHA plus EPA is also effective in correcting PUFA changes and gene expression. However, pathologic features of CF in the small intestine are reversed only by high-dose DHA. These findings shed light on the in vivo

mechanisms of PUFA abnormalities in CF and provide further evidence supporting a role for PUFA therapy of CF patients. 2. Methods and materials 2.1. Mouse breeding and genotyping All experiments were carried out under supervision of the Vanderbilt Division of Animal Care using protocols approved by the Institutional Animal Care and Use Committee. CFTRtm1/UNC heterozygous mice (CFTR+/−) on a C57BL/6J genetic background (B6.129P2-Cftrtm1Unc/J, stock number 002196) were purchased from The Jackson Laboratory (Bar Harbor, ME, USA). The mice were housed within a specific pathogen-free barrier facility with a 12-h light/dark cycle. Heterozygous mice were bred to obtain both homozygous wild-type (CFTR+/+) and knockout (CFTR−/−) mice. Ear-clip samples of 14-day-old mice were used for genotype analysis as previously described [29]. 2.2. Mouse feeding Wild-type and knockout mice were weaned at 23 days of age and placed on one of three liquid diets for 14 days. Each diet was based on Peptamen (Nestle Clinical Nutrition, Deerfield, IL, USA), a complete liquid enteral formulation composed mainly of medium-chain triglycerides, carbohydrates and hydrolyzed protein. Diet 1 (Peptamen) consisted of the Peptamen diet alone without additions. Diet 2 (Peptamen plus DHA) consisted of Peptamen alone given for 7 days, followed by Peptamen supplemented with 40 mg/day of DHA in free fatty acid form, prepared as a stable emulsion, given for an additional 7 days. Diet 3 (Peptamen AF) consisted of Peptamen Advanced Formula (AF) given for 14 days. The ingredients of Peptamen AF are essentially identical to those of Peptamen except that the AF formulation contains soluble fiber and refined fish oil (from anchovy and sardine), including 2.4 g/L of DHA and EPA primarily in triglyceride form. Complete ingredient lists and nutritional information for the Peptamen diets are available at http://www.nestle-nutrition.com/ products/. All three diets contain antioxidants in the form of vitamin A, vitamin C, vitamin E and selenium, to minimize fatty acid oxidation. Mice were fed fresh food

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daily and had access to water ad libitum. The liquid food volume consumed and body weight were measured daily. 2.3. Tissue harvest and processing The mice were sacrificed after receiving the above diets for 14 days post-weaning. Plasma and tissue from various organs (lung, pancreas, ileum, liver, kidney and heart) were collected, flash frozen in liquid nitrogen, and stored at −80°C prior to analysis. For lung tissue, cell suspensions enriched in respiratory epithelial cells were prepared as previously described [17]. Briefly, the lung was flushed with Krebs–Henseleit buffer (KHB) containing 0.5% bovine serum albumin (BSA) and then minced and transferred to a tube containing 10 ml KHB with 2000 units DNase (Sigma-Aldrich, St. Louis, MO, USA), 0.5 units thermolysin (Sigma-Aldrich) and 1000 units collagenase (SigmaAldrich). The tissue was incubated in a shaker at 37°C for 30 min. Following incubation, KHB containing 4% BSA was added to the cell suspension and centrifuged at 500×g for 10 min. The supernatant was removed and the cells were washed once in KHB and resuspended in phosphate-buffered saline (PBS). Pancreatic cell suspensions were prepared by mechanical dissociation and addition of collagenase as described by Bruzzone et al. [30]. Briefly, Krebs–Ringer Hepes (KRB-Hepes) buffer, adjusted to pH 7.4, containing 12.5 mM Hepes, 135 mM NaCl, 4.8 mM KCl, 1.0 mM CaCl2, 1.2 mM KH2PO4, 1.2 mM MgSO4, 5.0 mM NaHCO3, 5 mM glucose and 0.01 mg/ml aprotinin was used as the dissociation medium. The pancreas was chopped into small pieces and transferred to a glass tube. Two milliliters of KRB-Hepes containing 1 mg/ml collagenase was added to the tube and shaken vigorously until the tissue suspension appeared homogenous. Fresh KRB-Hepes with 0.1% human serum albumin (HSA) was added and the tissue centrifuged at 500×g for 5 min. The digested tissue was washed twice with repeated centrifugations, resuspended in fresh KRB-Hepes-HSA and filtered consecutively through a 300-μm and then a 70-μm filter. The pancreatic cells that passed through the filter were centrifuged and resuspended in PBS. To enrich for ileal epithelial cells, sections of ileum were rinsed with PBS, sliced open and cells obtained by scraping the inner mucosal surface with a razor blade. Tissue homogenates were prepared from the kidney, heart, liver and brain by mincing and homogenizing the tissue samples in PBS.

paraffin, sectioned at 4 microns, and stained with hematoxylin and eosin, periodic acid-Schiff (PAS) and mucicarmine. Slides were evaluated on an Olympus BX41 microscope by a veterinary pathologist (K.L.B.) who was blinded to both genotype and diet of the mice. To analyze impaction of mucus accumulation and impaction in the ileal crypts, at least two entire ileal sections were examined and each specimen was given a semiquantitative score based on the following scale: 0, no impaction of secretory products in the crypts; 1, impacted secretory material in the deep portion of the crypts in b10% of the crypts; 2, impaction of secretory material in the deep portion of the crypts in 10%–30% of crypts, with rare accumulation of material in the bottom third of the villus; 3, impaction of secretory material in 30%–70% of the crypts, with accumulation of material commonly observed up to the top third of the crypt; and 4, impaction of secretory material in N70% of the crypts, with frequent dilation of the crypts and presence of material in the top third of the villus. 2.7. Statistics Individual quantitative characteristics between wild-type and knockout mice were evaluated using the Mann–Whitney rank-sum test. This nonparametric test was chosen because the numbers of mice are too small to formally establish normality of distribution. Comparisons of quantitative characteristics between wild-type and knockout mice treated with different diets were performed using two-way analysis of variance (ANOVA). The variables for these analyses were genotype (wild-type vs. knockout) and diet (Peptamen vs. Peptamen plus DHA or Peptamen AF). For these analyses, results from mice fed diets 2 and 3 were compared to those from mice fed the control diet 1 individually. Results from mice fed diets 2 and 3 were not compared to each other. Individual pair-wise comparisons were made using a Bonferroni posttest. In most cases, Pb.05 was considered statistically significant. For individual fatty acid levels (see Supplemental Tables 1 and 2), a Bonferroni correction for multiple comparisons was applied, and Pb.002 was considered significant. Statistical analysis was performed using Prism software (v5.04; GraphPad Software, La Jolla, CA, USA).

3. Results 3.1. CFTR knockout mice

2.4. Fatty acid analysis Cellular fatty acids were extracted and methylated using the method of Folch et al. [31]. Briefly, heptadecanoic acid (17:0) was added as an internal standard to cell suspensions in PBS. Six volumes of chloroform–methanol (2:1) were added, and the mixture was vortexed and incubated on ice for 10 min. The mixture was then centrifuged at 1100×g for 10 min, and the lower organic phase transferred to a new glass tube and dried down completely under nitrogen gas. To methylate the fatty acids, 0.5 ml of 0.5 N methanolic NaOH (Acros Organics, Geel, Belgium) was added to the dried-down lipids, vortexed and heated at 100°C for 3 min. Then 0.5 ml boron trifluoride (BF3-methanol; Sigma) was added to the mixture and incubated at 100°C for 1 minute. The resulting fatty acid methyl esters (FAMEs) were extracted using 1 ml hexane, followed by 6.5 ml of saturated NaCl solution. The mixture was vortexed and centrifuged at 500×g for 4 min, and the upper hexane layer was transferred to a new glass tube. Total FAMEs contained in the hexane layer were analyzed by gas chromatography (GC) using an Agilent 7980A GC system (Agilent Technologies, Santa Clara, CA, USA) equipped with a Supelcowax SP-10 capillary column (Supelco, Bellefonte, PA, USA) coupled to a mass spectrometer (model 5975c; Agilent Technologies). Individual FAMEs were identified by comparison to FAME standards (NuChek Prep, Elysian, MN, USA). The mass of the FAMEs was determined by comparing areas of unknown FAMEs to that of the 17:0 internal standard. To facilitate comparison to prior studies [17–19], results were expressed as the molar percentage (mol%) of each FAME relative to the total FAME mass of the sample. 2.5. Gene expression analysis Total RNA was isolated from homogenized mouse tissues using TRIzol reagent (Invitrogen, Carlsbad, CA, USA), following the manufacturer's instructions. Complementary DNA was generated from 2 μg of total RNA with random hexamers using TaqMan reverse transcription reagents (Applied Biosystems, Foster City, CA, USA). Quantitative real-time PCR (qRT-PCR) was done on mouse cDNA using Taqman gene expression assays (Applied Biosystems), universal PCR master mix (Bio-Rad Laboratories, Hercules, CA, USA) and a CFX96 Real-Time PCR system (Bio-Rad) with Taqman commercial primers and probes (Applied Biosystems), according to manufacturer's instructions. Data were analyzed using CFX Manager software (Bio-Rad). The relative expression of each target gene was calculated using the comparative Ct method and normalized to a reference gene, GAPDH. The levels of GAPDH expression remained essentially constant across genotypes and diets with less than 1 average Ct unit variance among all conditions for each tissue examined. 2.6. Histologic analysis For sample preparation, ileum tissues were submerged in 10% neutral-buffered formalin and fixed overnight. The tissues were processed routinely, embedded in

CFTR knockout (CFTR−/−) mice were generated as described in Methods and materials. These mice were significantly smaller at weaning than their wild-type littermates (6.5±0.3 g vs. 10.5±0.3 g; Pb.01). Both groups of mice gained weight at approximately the same rate over the 14 days of the experiment with a nearly constant ~4-g average difference between the wild-type and knockout animals (data not shown). Despite being fed a liquid diet (Peptamen) to prevent intestinal obstruction, a significant fraction of the knockout mice (9/28; 32.1%) died in the first 5 days post-weaning, compared with only 1/19 (5.3%) of wild-type littermates (P=.03). These findings are consistent with failure to thrive that is characteristic of this mouse model [32]. 3.2. n-6 PUFA metabolism in CF mice Fatty acid measurements were performed on various tissues from 35-day-old wild-type and CFTR −/− mice. Among the n-6 PUFAs, LA was significantly decreased and AA significantly increased in the lung, ileum and pancreas of CF mice (Supplemental Table 1). There was no difference in n-6 PUFAs in the liver, heart or kidney (Supplemental Table 2). In in vitro experiments, these changes have been attributed to increased metabolism of LA to AA, due to increased expression and activity of Δ5- and Δ6-desaturase enzymes [25,27] (Fig. 1A). This metabolism can be estimated by calculating the ratio of products to substrates (e.g., AA/LA) [33]. Accordingly, this ratio is significantly increased in the lung, ileum and pancreas of CFTR−/− animals compared with wild-type controls (Fig. 1B). There is no difference in these ratios in the liver, kidney and heart (not shown). In the lung and ileum, these changes correlate with significant increases in the mRNA expression of Δ5- and Δ6-desaturases in CFTR−/− mice (Fig. 1C). Elongase 5 mRNA is also increased in the lung only. There was insufficient material to assess gene expression in the pancreas. There are no differences in the expression of these enzymes in the liver, kidney and heart (not shown).

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Supplemental Table 3. Neither EPA nor DHA was detected in Peptamen. Supplementation with free DHA increased its concentration to 32.5%. In contrast, Peptamen AF contained only 3.8% DHA, with more than twice as much EPA (8.2%). The n-6:n-3 PUFA ratio, which was 5.7 in the Peptamen diet, was reduced to 0.7 and 1.4 in the Peptamen plus DHA and Peptamen AF diets, respectively. There was no difference in plasma DHA (5.2±0.5 mol% vs. 6.2± 0.5 mol%) levels between wild-type and CFTR−/− mice. Plasma DHA levels increased in all mice from 5.7±0.4 in control mice to 16.3± 0.3 mol% in Peptamen plus DHA-treated mice and to 9.0±0.3 mol% in Peptamen AF-treated mice (Pb.05 for both comparisons). Baseline DHA levels were lower in the lung and ileum in CFTR−/− animals compared with wild-type littermates (Fig. 3A). There was no difference in liver DHA levels. Mice fed DHA-supplemented Peptamen exhibited increased DHA in all tissues, reversing the baseline deficiency seen in CFTR−/− mice. Peptamen AF also reversed the deficiency by increasing DHA levels in knockout mice without increasing baseline levels in the wild-type mice in any tissue except liver. There were no significant differences in EPA levels between CFTR−/− and wild-type animals at baseline in any tissue (Fig. 3B). Mice fed DHA-supplemented Peptamen showed increased EPA levels in all tissues except pancreas, presumably due to DHA→EPA retroconversion [34,35]. The increase was greater in mice fed Peptamen AF. However, there were no differences between wild-type and knockout mice. There were marked decreases in the predicted metabolism of LA to AA in mice fed either DHA-supplemented Peptamen or Peptamen AF (Fig. 4A). Mice fed these diets showed significant decreases in the AA/LA

3.3. n-7 and n-9 metabolism in CF mice Fatty acid analysis also revealed differences in fatty acids of the n-7 and n-9 pathways, although these were seen only in ileum and pancreas, and not in the lung, liver, kidney or heart (Supplemental Tables 1 and 2). The primary changes were increased 18:0 and 18:1n-7 and decreased 16:1n-7 and 18:1n-9. The metabolic pathways and enzymes involving these fatty acids are shown in Fig. 2A. From this, one can hypothesize that these patterns are caused by increased elongase 6 activity and decreased Δ9-desaturase activity. Accordingly, ratios of 18:0/16:0 and 18:1n-7/16:1n-7, representative of elongase 6 activity, are significantly increased in the ileum and pancreas (Fig. 2B). However, ratios of 16:1n-7/16:0 and 18:1n-9/18:0, representative of Δ9-desaturase activity, are decreased. 3.4. Effects of dietary PUFA supplementation on metabolism and gene expression Prior in vitro studies have shown that the metabolic abnormalities characteristic of CF can be reversed by supplementation with DHA or EPA [27]. To test this in vivo, we fed CFTR−/− knockout mice and wildtype littermates one of two PUFA-supplemented diets. The first was the previously described Peptamen liquid diet supplemented with DHA in free fatty acid form (as in Ref. [17]). The concentration was adjusted to give each mouse approximately 40 mg/day DHA. The second diet consisted of Peptamen AF, which contains DHA and EPA in triglyceride form. Measured fatty acid concentrations are shown in

A

Elongase 6 (EL6) 16:0

18:0

B 18:0 16:0 (EL6)

18:1n-9 18:1n-7 16:1n-7 (EL6) 16:1n-7

18:1n-7 16:1n-7 16:0 (Δ9D)

18:1n-9 18:0 (Δ9D)

39

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Ileum

Pancreas

0.8

2.0

0.8

0.6

1.5

0.6

0.4

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0.4

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0.2

P=.001

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P=.004

0.0

0.0

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0.4

0.3

0.3

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0.1

0.1

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0.0

WT

CF

P=.005

0.0

P=.005

0.0 0.2

0.1

P=.003

P=.016

0.0

0.4

P=.001

WT

CF

0.0

WT

CF

Fig. 2. Activity of enzymes in the n-7 and n-9 unsaturated fatty acid metabolic pathways in CF mice. (A) Schematic diagram of the n-7 and n-9 unsaturated fatty acid metabolic pathways, including the pertinent enzymes, elongase 6 (EL6) and Δ9-desaturase (Δ9D). (B) Estimated activity of EL6 and Δ9D in selected mouse tissues. WT (CFTR+/+) and CF (CFTR−/−) mice were produced, maintained on a Peptamen liquid diet post-weaning for 14 days, and then sacrificed as described in Methods and materials. Lung, ileum and pancreas tissues were harvested and processed, and total tissue fatty acids were measured as described in Methods and materials. Relative activity of EL6 was estimated using the ratio of product (18:0 or 18:1n-7) to substrate (16:0 or 16:1n-7). Relative activity of Δ9D was estimated using the ratio of product (16:1n-7 or 18:1n-9) to substrate (16:0 or 18:0). Ratios for individual mice (n=6 or 7) are shown with the mean indicated by a horizontal line. P values were determined by Mann–Whitney rank-sum test. Comparisons without P values were not significant.

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Lung *,†

A DHA (mol%)

16

WT CF

Ileum 8

Pancreas 8





30 25

12

6 ‡



8

20

4



4



2 5

0

0

B 3



15 10

2

*



6



* 4

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† †

0

EPA (mol%)

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6

WT CF



0

8

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5

‡ ‡



8



6

6

4

4

2

2

0

0

4 †

3 1 †



† †

2



1 0

0

Fig. 3. DHA and EPA levels in mouse tissues treated with PUFA-supplemented diets. WT (CFTR+/+) and CF (CFTR−/−) mice were produced and maintained on a Peptamen, Peptamen plus DHA or Peptamen AF liquid diet post-weaning for 14 days, and then sacrificed as described in Methods and materials. Lung, ileum, pancreas and liver tissues were harvested and processed, and total tissue fatty acids were measured as described in Methods and materials. Graphs show relative DHA (A) or EPA (B) levels in tissues from WT (white bars) or CF (gray bars) mice. Bars indicate mean±SEM of six or seven animals. P values were determined by two-way ANOVA with Bonferroni posttest for WT vs. CF pair-wise comparisons. Significant interaction effect (Pb.05) between treatment and genotype was seen for ileum DHA (both Peptamen plus DHA and Peptamen AF), pancreas DHA (Peptamen AF only) and liver DHA (Peptamen plus DHA only). Statistical significance (Pb.05) is indicated on the figure for the following pair-wise comparisons: *CF vs. WT mice within each diet; †mice with common genotype fed Peptamen vs. Peptamen plus DHA; ‡mice with common genotype fed Peptamen vs. Peptamen AF.

A

Lung

20:4n-6/18:2n-6

3

Ileum 1.0

*

0.8

2 †





1

1.2



0.4





0.0

Peptamen Peptamen Peptamen +DHA AF

9 6

* *‡

12

*,‡

* * * *

3 0

Δ6D





Peptamen Peptamen Peptamen +DHA AF

15

60

12

45

9

30 15

Δ5D



*,‡

+DHA AF

*

75

*

Peptamen



0.0

Peptamen Peptamen Peptamen +DHA AF

B WT CF WT CF WT CF

0.8 0.4

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15

WT CF

0.6

0

Relative mRNA Level

Liver 1.6

*

0

EL5

6

† ‡

*



Δ5D



†† †





‡ ‡ ‡



0

EL5





3 †

Δ6D

*

Δ5D

Δ6D

EL5

Fig. 4. Expression and activity of fatty acid desaturases in n-6 PUFA metabolic pathway in CF mice treated with PUFA-supplemented diets. (A) Activity of the n-6 PUFA pathway in selected mouse tissues. WT (CFTR+/+) and CF (CFTR−/−) mice were produced, maintained on a Peptamen, Peptamen plus DHA or Peptamen AF liquid diet post-weaning for 14 days, and then sacrificed as described in Methods and materials. Lung, ileum and liver tissues were harvested and processed, and total tissue fatty acids were measured as described in Methods and materials. Relative activity of the n-6 PUFA pathway was estimated using the ratio of product (20:4n-6; AA) to substrate (18:2n-6; LA). Ratios for individual mice (n=6 or 7) are shown (WT, closed circles; CF, open circles) with the mean indicated by a horizontal line. (B) RNA was harvested from the lung, ileum and liver of WT and CF mice generated and processed as in (A) above. Relative mRNA level of Δ5-desaturase (Δ5D), Δ6-desaturase (Δ6D) and elongase 5 (EL5) genes was determined by quantitative RT-PCR as described in Methods and materials. Bars indicate mean±SEM of six or seven animals. For both panels, P values were determined by two-way ANOVA with Bonferroni posttest for pair-wise comparisons. For AA/LA ratios, significant interaction effect (Pb.05) between treatment and genotype was seen in the lung and ileum for both treatments. For gene expression, significant interaction effect (Pb.05) between treatment and genotype was seen for ileum Δ5D and Δ6D with both treatments. Statistical significance (Pb.05) is indicated on the figure for the following pair-wise comparisons: *CF vs. WT mice within each diet; †mice with common genotype fed Peptamen vs. Peptamen plus DHA; ‡mice with common genotype fed Peptamen vs. Peptamen AF.

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ratio compared with mice fed Peptamen alone, eliminating the difference between wild-type and knockout animals in the lung, ileum and pancreas. A similar decrease was seen in the liver, although there was no difference between mice with different genotypes at baseline. Neither of the supplemented diets reversed the changes in n-7 and n-9 metabolism described above (data not shown). In the ileum and liver, reversal of LA→AA metabolism was accompanied by significant decreases in the expression of Δ5- and Δ6-desaturases and elongase 5 (Fig. 4B). In contrast, DHA supplementation appeared to have no effect on expression of these genes in the lung, while expression of these genes was actually increased with Peptamen AF administration compared with mice fed unsupplemented Peptamen.

3.5. Effects of n-3 fatty acid supplementation on ileal pathology Examination of histologic sections of the knockout mouse ileum revealed evidence of hypersecretion and dysmotility. Specifically, the deep crypts between ileal villi showed accumulation of impacted eosinophilic material that occasionally dilated the crypts (Fig. 5B). This material also stained positive for mucicarmine and PAS, consistent with mucus (not shown), and was not found in the ileum of homozygous wild-type animals (Fig. 5A). This histologic evidence of mucus accumulation and impaction was scored semiquantitatively (see Methods and materials) in Peptamen-fed wildtype and knockout mice and compared with those fed Peptamen plus DHA (Fig. 5C) or Peptamen AF (Fig. 5D). Mice fed Peptamen plus DHA showed significantly lower levels of mucus impaction than those fed with unsupplemented Peptamen (Fig. 5E). There was no difference between mice fed unsupplemented Peptamen and Peptamen AF. Histologic examination of pancreas and lung tissue did not reveal any additional significant differences between wild-type and CF mice.

A

B

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4. Discussion The data in this report provide in vivo evidence supporting the hypothesis that the n-6 PUFA alterations observed in CF are due to increased expression and activity of the enzymes in the metabolic pathway. The predicted activity of these enzymes, as measured by AA/LA ratio, was significantly increased in the lung, ileum and pancreas of CFTR−/− mice (Fig. 1). This correlated perfectly with increased expression of Δ5- and Δ6-desaturases in the lung and ileum. Elongase 5, another enzyme in this pathway, was also increased in the lung. Importantly, there was no difference in expression of the enzymes between knockout and wild-type animals in the liver, heart or kidney, tissues that do not express significant levels of CFTR or exhibit the characteristic PUFA abnormalities of CF. These data are similar to those described in in vitro cell culture models of CF [25], and they provide a clear connection between loss of CFTR expression and function and the fatty acid abnormalities. We also noted differences in metabolism of n-7 and n-9 fatty acids that suggested increased activity of elongase 6 and decreased activity of Δ9-desaturase in the ileum and pancreas only (Fig. 2). These findings are different from those noted in cell culture models, which exhibited increased expression and activity of both enzymes [26]. In these tissues, this resulted in decreased levels of 16:1n-7 and 18:1n-9, both of which have been shown to be increased in the plasma of CF patients in a subset of studies [11,36,37]. The significance of these differences is unclear. However, they may reflect differences in human versus mouse physiology or differences between tissues and plasma. In in vitro models, administration of DHA or EPA reduced LA→AA metabolism by suppressing expression of Δ5- and Δ6-desaturases, supporting the role of these enzymes in the PUFA abnormalities of CF [27]. In vivo, supplementation with high doses of DHA or lower doses of DHA plus EPA (Peptamen AF) reduced LA→AA metabolism in all tissues examined, leading to normalization of PUFA levels in CFTR−/−

E4 ***

C

D

Impaction Score

3

2

1

0 Peptaman Peptaman Peptaman +DHA AF

Fig. 5. Mucus accumulation and impaction in ileal crypts of mice treated with PUFA-supplemented diets. WT (CFTR+/+) and CF (CFTR−/−) mice were produced, maintained on a Peptamen, Peptamen plus DHA or Peptamen AF liquid diet post-weaning for 14 days, and then sacrificed as described in Methods and materials. Ileal tissue was collected and processed for histology as described in Methods and materials. (A–D) Sections of ileum from WT mice fed Peptamen alone (A) or CF mice fed Peptamen alone (B), Peptamen plus DHA (C) or Peptamen AF (D) were stained with hematoxylin and eosin and imaged at 400× magnification. Arrows indicated accumulated and impacted mucus. (E) A histologic impaction score was generated for WT (circles) or CF (squares) mice fed with each diet as described in Methods and materials. The median score for CF mice on each diet is indicated by the horizontal line. ***Pb.001 for CF mice treated with Peptamen vs. Peptamen plus DHA.

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mice (Fig. 4A). In the ileum, this correlated with normalization of desaturase and elongase gene expression in knockout mice. Expression of these genes was also significantly decreased in the liver, as has been previously described [38]. In the lung, however, Peptamen plus DHA had no effect on enzyme gene expression in wild-type or knockout animals. This result was unexpected, since the supplement caused complete normalization of AA/LA ratios in the lung. One possible explanation is that PUFA levels in the lung are controlled to a larger extent by exogenous fatty acid uptake rather than metabolic enzyme activity. The source of exogenous fatty acids would be primarily chylomicrons and very low density lipoproteins, originating in the intestinal epithelium and liver hepatocytes, respectively, both of which show significant downregulation of enzyme gene expression and AA metabolism in DHAsupplemented mice. Interestingly, metabolic gene expression in the lung was actually increased in mice treated with Peptamen AF. This suggests that this diet may contain some tissue-specific inducer of enzyme gene expression. Besides the differences in PUFA content, Peptamen AF contained higher levels of saturated and monounsaturated (particularly 16:1n-7) fatty acids (Supplemental Table 3). It is possible that these changes are responsible for the gene expression differences in the lung. Other differences between Peptamen plus DHA and Peptamen AF should be noted. In the former, DHA is given in its free fatty acid form, while the fat content of Peptamen AF is primary triglyceride. Given that Peptamen AF was also effective in reducing desaturase enzyme expression and activity, these data suggest that either form of delivery is effective. This matches data from previous reports showing reversal of PUFA abnormalities in CF animal models using either free fatty acid [17] or glycerophospholipid forms [19]. The PUFA content of these diets also differed. Peptamen plus DHA contained high concentrations (14.19 mmol/ml) of DHA only. The peptamen AF diet contained less than one-tenth as much DHA (1.38 mmol/ml) and much more EPA (2.98 mmol/ml). Since Peptamen AF is also effective at reducing the abnormally increased metabolism in CF mice, this suggests that lower doses of PUFAs are sufficient to observe the desired effect. In fact, it is possible that the effect of PUFAs can be maximized at a much lower levels. It is difficult to translate these diets into an equivalent human dosage as feeding behaviors differ between the species and the amount of dietary lipid absorbed is not known. However, human studies using a standard DHA dose of 1.0–1.2 g/day were sufficient to increase plasma DHA by 4–5 mol%, similar to the increment seen in mice fed Peptamen AF [39,40]. However, Peptamen plus DHA increased plasma DHA levels by more than 10 mol% in mice, a level unlikely to be achieved by standard oral dosage in humans. In addition, it is likely that the EPA component of Peptamen AF contributes to the effect on desaturase gene expression in vivo, as it does in vitro[27]. However, it is difficult to separate the individual effects of DHA and EPA as they are metabolically interconvertible. Further study is required to determine optimal dietary dosage for humans. There was one notable difference between the Peptamen plus DHA and the Peptamen AF treated mice. The mice treated with Peptamen plus DHA showed a significant reduction in impacted mucus in the ileal crypts, while there was no change in the mice treated with Peptamen AF (Fig. 5). This finding suggests that the histologic abnormality is corrected by high-dose DHA, but not lower-dose DHA plus EPA. This difference occurred despite the fact that the n-6 PUFA abnormalities were corrected by both diets and may indicate that the pathologic factors contributing to this histologic abnormality, such as hypersecretion or decreased motility, are more associated with deficiency of DHA rather than with excess AA production. These findings are similar to those of Freedman et al. [17], which noted improvement of pancreatic duct dilation with DHA supplementation, but not with EPA, despite the fact that both supplements reduced AA production.

The current study extends the analysis of these previous reports [17,19] in several important ways. First, it demonstrates that the fatty acid abnormalities are associated with increased expression and activity of fatty acid metabolic enzymes. Further, it shows that correction of the fatty acid abnormalities with PUFA supplementation is accompanied by suppression of these enzymes. These data provide in vivo evidence to support the hypothesis first advanced in in vitro cell culture studies [23–27] that the n-6 PUFA abnormalities consistently observed in CF blood and tissues are caused by increased expression and activity of fatty acid desaturases. Second, it demonstrates that lower doses of PUFAs and mixtures of DHA and EPA are as effective in correcting metabolism as the higher doses of DHA only given in prior studies [17]. Thus, these results suggest that more therapeutic doses may still be effective. Third, while other studies focused on AA and DHA only, the current data indicate that there are also changes in other pathways, such as the n-7 and n-9 pathways that have been described in previous in vitro experiments [26]. Taken together, these studies provide correlative evidence suggesting that changes in enzyme expression may underlie fatty acid abnormalities in CF and establish an experimental model for further studies aimed at connecting these metabolic alterations with mutation and dysfunction of the CFTR protein and with CF pathophysiology. They also reveal potential targets for pharmacologic and/or dietary therapy that may have a positive impact on the disease course of CF patients. Acknowledgments This work was funded by the Edward and Nancy Fody Endowed Chair in Pathology (M.L.) and the Vanderbilt Physician Scientist Training Program (A.C.S.). Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.jnutbio.2014.09.001. References [1] O'Sullivan BP, Freedman SD. Cystic fibrosis. Lancet 2009;373(9678):1891–904. [2] Riordan JR, Rommens JM, Kerem B, Alon N, Rozmahel R, Grzelczak Z, et al. Identification of the cystic fibrosis gene: cloning and characterization of complementary DNA. Science 1989;245(4922):1066–73. [3] Kuo PT, Huang NN, Bassett DR. The fatty acid composition of the serum chylomicrons and adipose tissue of children with cystic fibrosis of the pancreas. J Pediatr 1962;60:394–403. [4] Al-Turkmani MR, Freedman SD, Laposata M. Fatty acid alterations and n-3 fatty acid supplementation in cystic fibrosis. Prostaglandins Leukot Essent Fatty Acids 2007;77(5–6):309–18. [5] Innis SM, Davidson AG. Cystic fibrosis and nutrition: linking phospholipids and essential fatty acids with thiol metabolism. Annu Rev Nutr 2008;28:55–72. [6] Strandvik B. Fatty acid metabolism in cystic fibrosis. Prostaglandins Leukot Essent Fatty Acids 2010;83(3):121–9. [7] Rogiers V, Vercruysse A, Dab I, Baran D. Abnormal fatty acid pattern of the plasma cholesterol ester fraction in cystic fibrosis patients with and without pancreatic insufficiency. Eur J Pediatr 1983;141(1):39–42. [8] Guilbault C, Wojewodka G, Saeed Z, Hajduch M, Matouk E, De Sanctis JB, et al. Cystic fibrosis fatty acid imbalance is linked to ceramide deficiency and corrected by fenretinide. Am J Respir Cell Mol Biol 2009;41(1):100–6. [9] Strandvik B, Gronowitz E, Enlund F, Martinsson T, Wahlstrom J. Essential fatty acid deficiency in relation to genotype in patients with cystic fibrosis. J Pediatr 2001; 139(5):650–5. [10] Olveira G, Olveira C, Acosta E, Espildora F, Garrido-Sanchez L, Garcia-Escobar E, et al. Fatty acid supplements improve respiratory, inflammatory and nutritional parameters in adults with cystic fibrosis. Arch Bronconeumol 2010;46(2):70–7. [11] Van Biervliet S, Vanbillemont G, Van Biervliet JP, Declercq D, Robberecht E, Christophe A. Relation between fatty acid composition and clinical status or genotype in cystic fibrosis patients. Ann Nutr Metab 2007;51(6):541–9. [12] Maqbool A, Schall JI, Garcia-Espana JF, Zemel BS, Strandvik B, Stallings VA. Serum linoleic acid status as a clinical indicator of essential fatty acid status in children with cystic fibrosis. J Pediatr Gastroenterol Nutr 2008;47(5):635–44.

S.W. Njoroge et al. / Journal of Nutritional Biochemistry 26 (2015) 36–43

[13] Ollero M, Astarita G, Guerrera IC, Sermet-Gaudelus I, Trudel S, Piomelli D, et al. Plasma lipidomics reveals potential prognostic signatures within a cohort of cystic fibrosis patients. J Lipid Res 2011;52(5):1011–22 [PMCID: 3073467]. [14] Carvalho-Oliveira I, Scholte BJ, Penque D. What have we learned from mouse models for cystic fibrosis? Expert Rev Mol Diagn 2007;7(4):407–17. [15] Guilbault C, Saeed Z, Downey GP, Radzioch D. Cystic fibrosis mouse models. Am J Respir Cell Mol Biol 2007;36(1):1–7. [16] Wilke M, Buijs-Offerman RM, Aarbiou J, Colledge WH, Sheppard DN, Touqui L, et al. Mouse models of cystic fibrosis: phenotypic analysis and research applications. J Cyst Fibros 2011;10(Suppl 2):S152–71. [17] Freedman SD, Katz MH, Parker EM, Laposata M, Urman MY, Alvarez JG. A membrane lipid imbalance plays a role in the phenotypic expression of cystic fibrosis in cftr(−/−) mice. Proc Natl Acad Sci U S A 1999;96(24):13995–4000 [PMCID: 24179]. [18] Ollero M, Laposata M, Zaman MM, Blanco PG, Andersson C, Zeind J, et al. Evidence of increased flux to n-6 docosapentaenoic acid in phospholipids of pancreas from cftr−/− knockout mice. Metabolism 2006;55(9):1192–200. [19] Mimoun M, Coste TC, Lebacq J, Lebecque P, Wallemacq P, Leal T, et al. Increased tissue arachidonic acid and reduced linoleic acid in a mouse model of cystic fibrosis are reversed by supplemental glycerophospholipids enriched in docosahexaenoic acid. J Nutr 2009;139(12):2358–64. [20] Zaman MM, Martin CR, Andersson C, Bhutta AQ, Cluette-Brown JE, Laposata M, et al. Linoleic acid supplementation results in increased arachidonic acid and eicosanoid production in CF airway cells and in cftr−/− transgenic mice. Am J Physiol Lung Cell Mol Physiol 2010;299(5):L599–606 [PMCID: 2980390]. [21] Tiesset H, Bernard H, Bartke N, Beermann C, Flachaire E, Desseyn JL, et al. (n-3) long-chain PUFA differentially affect resistance to Pseudomonas aeruginosa infection of male and female cftr−/− mice. J Nutr 2011;141(6):1101–7. [22] Beharry S, Ackerley C, Corey M, Kent G, Heng YM, Christensen H, et al. Long-term docosahexaenoic acid therapy in a congenic murine model of cystic fibrosis. Am J Physiol Gastrointest Liver Physiol 2007;292(3):G839–48. [23] Andersson C, Al-Turkmani MR, Savaille JE, Alturkmani R, Katrangi W, Cluette-Brown JE, et al. Cell culture models demonstrate that CFTR dysfunction leads to defective fatty acid composition and metabolism. J Lipid Res 2008;49(8):1692–700 [PMCID: 2444007]. [24] Al-Turkmani MR, Andersson C, Alturkmani R, Katrangi W, Cluette-Brown JE, Freedman SD, et al. A mechanism accounting for the low cellular level of linoleic acid in cystic fibrosis and its reversal by DHA. J Lipid Res 2008;49(9):1946–54 [PMCID: 2515522]. [25] Njoroge SW, Seegmiller AC, Katrangi W, Laposata M. Increased Delta5- and Delta6-desaturase, cyclooxygenase-2, and lipoxygenase-5 expression and activity are associated with fatty acid and eicosanoid changes in cystic fibrosis. Biochim Biophys Acta 2011;1811(7–8):431–40.

43

[26] Thomsen KF, Laposata M, Njoroge SW, Umunakwe OC, Katrangi W, Seegmiller AC. Increased elongase 6 and Delta9-desaturase activity are associated with n-7 and n-9 fatty acid changes in cystic fibrosis. Lipids 2011;46(8):669–77. [27] Njoroge SW, Laposata M, Katrangi W, Seegmiller AC. DHA and EPA reverse cystic fibrosis-related FA abnormalities by suppressing FA desaturase expression and activity. J Lipid Res 2012;53(2):257–65 [PMCID: 3269161]. [28] Katrangi W, Lawrenz J, Seegmiller AC, Laposata M. Interactions of linoleic and alpha-linolenic acids in the development of fatty acid alterations in cystic fibrosis. Lipids 2013;48(4):333–42. [29] Zeng W, Lee MG, Yan M, Diaz J, Benjamin I, Marino CR, et al. Immuno and functional characterization of CFTR in submandibular and pancreatic acinar and duct cells. Am J Physiol 1997;273(2 Pt 1):C442–55. [30] Bruzzone R, Halban PA, Gjinovci A, Trimble ER. A new, rapid, method for preparation of dispersed pancreatic acini. Biochem J 1985;226(2):621–4 [PMCID: 1144753]. [31] Folch J, Lees M, Sloane Stanley GH. A simple method for the isolation and purification of total lipides from animal tissues. J Biol Chem 1957;226(1):497–509. [32] Snouwaert JN, Brigman KK, Latour AM, Malouf NN, Boucher RC, Smithies O, et al. An animal model for cystic fibrosis made by gene targeting. Science 1992;257 (5073):1083–8. [33] Vessby B, Gustafsson IB, Tengblad S, Boberg M, Andersson A. Desaturation and elongation of fatty acids and insulin action. Ann N Y Acad Sci 2002;967:183–95. [34] Gronn M, Christensen E, Hagve TA, Christophersen BO. Peroxisomal retroconversion of docosahexaenoic acid (22:6(n-3)) to eicosapentaenoic acid (20:5 (n-3)) studied in isolated rat liver cells. Biochim Biophys Acta 1991;1081(1): 85–91. [35] Brossard N, Croset M, Pachiaudi C, Riou JP, Tayot JL, Lagarde M. Retroconversion and metabolism of [13C]22:6n-3 in humans and rats after intake of a single dose of [13C]22:6n-3-triacylglycerols. Am J Clin Nutr 1996;64(4):577–86. [36] Roulet M, Frascarolo P, Rappaz I, Pilet M. Essential fatty acid deficiency in well nourished young cystic fibrosis patients. Eur J Pediatr 1997;156(12):952–6. [37] Aldamiz-Echevarria L, Prieto JA, Andrade F, Elorz J, Sojo A, Lage S, et al. Persistence of essential fatty acid deficiency in cystic fibrosis despite nutritional therapy. Pediatr Res 2009;66(5):585–9. [38] Nakamura MT, Cheon Y, Li Y, Nara TY. Mechanisms of regulation of gene expression by fatty acids. Lipids 2004;39(11):1077–83. [39] Van Biervliet S, Devos M, Delhaye T, Van Biervliet JP, Robberecht E, Christophe A. Oral DHA supplementation in DeltaF508 homozygous cystic fibrosis patients. Prostaglandins Leukot Essent Fatty Acids 2008;78(2):109–15. [40] Alicandro G, Faelli N, Gagliardini R, Santini B, Magazzu G, Biffi A, et al. A randomized placebo-controlled study on high-dose oral algal decosahexaenoic acid supplementation in children with cystic fibrosis. Prostaglandins Leukot Essent Fatty Acids 2013;88(2):163–9.

- mice by suppressing fatty acid desaturases.

Cystic fibrosis patients and model systems exhibit consistent abnormalities in metabolism of polyunsaturated fatty acids that appear to play a role in...
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