Abstract The development of intestinal permeability and the penetration of microbial products are key factors associated with the onset of metabolic disease. However, the mechanisms underlying this remain unclear. Here we show that, unlike liver or adipose tissue, high fat diet (HFD)/obesity in mice does not cause monocyte/macrophage infiltration into the intestine or pro-inflammatory changes in gene expression. Rather HFD causes depletion of intestinal eosinophils associated with the onset of intestinal permeability. Intestinal eosinophil numbers were restored by returning HFD fed mice to normal chow and were unchanged in leptin-deficient (Ob/Ob) mice, indicating that eosinophil depletion is caused specifically by a high fat diet and not obesity per se. Analysis of different aspects of intestinal permeability in HFD fed and Ob/Ob mice shows an association between eosinophil depletion and ileal paracelullar permeability, as well as leakage of albumin into the feces, but not overall permeability to FITC dextran. These findings provide the first evidence that a high fat diet causes intestinal eosinophil depletion, rather than inflammation, which may contribute to defective barrier integrity and the onset of metabolic disease.

Citation: Johnson AMF, Costanzo A, Gareau MG, Armando AM, Quehenberger O, Jameson JM, et al. (2015) High Fat Diet Causes Depletion of Intestinal Eosinophils Associated with Intestinal Permeability. PLoS ONE 10(4): e0122195. https://doi.org/10.1371/journal.pone.0122195 Academic Editor: Pratibha V. Nerurkar, College of Tropical Agriculture and Human Resources, University of Hawaii, UNITED STATES Received: October 21, 2014; Accepted: February 13, 2015; Published: April 2, 2015 Copyright: © 2015 Johnson et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited Data Availability: All relevant data are within the paper. Funding: This work was funded by the following grants D.K.0033651, D.K.074868, D.K.063491 and D.K.09062 (J.O.). A.M.F.J. was supported by an A.D.A mentor-based fellowship (7-09-MN-37). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Competing interests: The authors have declared that no competing interests exist.

Introduction The penetration of bacteria or bacterial products, such as LPS, across the intestinal barrier is associated with the development of chronic inflammation and metabolic disease in both obese humans and mice [1,2,3,4,5,6]. Evidence suggests that reduction of tight junction protein expression and degeneration of epithelial barrier integrity is a feature of this permeability [4,7,8,9]. However, the underlying and associated mechanisms remain ill-defined. The intestinal immune system is an effective immunological barrier to infection based on its juxtaposition to the extensive community of micro-organisms referred to as the intestinal microbiota [10]. Indeed, the intestinal immune system is often referred to as a “firewall” preventing the systemic penetration of bacteria. This barrier function has metabolic implications, since microbial signals in locations distinct from the gastrointestinal (GI) tract can trigger inflammatory processes, such as those which can cause insulin resistance. Indeed, intravenous infusion of low levels of endotoxin to mimic bacterial penetration is sufficient to render mice glucose intolerant [2]. It has previously been suggested that a high fat diet (HFD) or obesity results in pro-inflammatory changes in the ileum and proximal colon of rodents, contributing to intestinal permeability [11,12,13,14,15,16]. These changes include foci of NF-κB activation and 1.5–6 fold increases in the expression of TNF-α and IL-1β [13,15,16]. However, such increases in cytokine expression have not been observed in all published studies [16], and neither histological nor cellular analysis of the colon revealed significant pro-inflammatory responses [14]. Cellular analysis of the small intestine in HFD fed mice remains unreported. Residing within the lamina propria, macrophages, dendritic cells (DCs) and eosinophils are predominate cell populations. Intestinal macrophage/DC populations regulate both tolerance to the microbiota and host defense against infection. Specifically, macrophage/DC IL-10 production and induction of regulatory T (Treg) cells can promote tolerance, whereas depletion of intestinal macrophages renders mice susceptible to bacterial penetration [17,18]. Although quite abundant, intestinal eosinophils have been less extensively studied. Eosinophilia can be a feature of specific inflammatory conditions in mice and humans, such as food allergies, eosinophilic gastroenteritis, allergic colitis and inflammatory bowel disease (IBD) [19,20,21]. In these conditions, recruitment and expansion of eosinophils is promoted by cytokines (e.g. IL-5) and chemokines (e.g. eotaxins) and they play an active role in disease progression [19,20,21]. However, tissue-resident eosinophils can also promote epithelial repair and barrier function during homeostasis [22,23,24]. We sought to determine whether the intestinal permeability observed on HFD feeding and obesity is associated with alterations in small intestinal macrophage/DC or eosinophil populations.

Materials and Methods Mice C57Bl6/J and Ob/Ob were purchased from Jackson labs and CX3CR1GFP/+ mice [25] were bred in-house and experiments approved by UCSD Institute Animal Care and Use Committee (IACUC). Mice were maintained on normal chow diet (Lab Diet 5001) on a 12h/12h light/dark cycle up to 8–12 weeks of age prior to transfer to a high fat diet containing 60%kcal fat (Research diets, New Brunswick, NJ, D12492) and analyzed at the indicated time points. In diet switch studies, mice were provided with HFD for four weeks and the returned to normal chow for a further 12 days. Upon analysis, mice were euthanized with 75mg/kg Pentobarbital (Schering-Plough, Millsoboro, DE) by i.p. injection and the diaphragm cut. Fecal albumin ELISA Fecal pellets were collected prior to the application of a high fat diet (day 0) and on 3 proceeding days, snap frozen in liquid nitrogen and stored at -80°C. Pellets were resuspended at 10mg/ml in sterile phosphate buffered saline (PBS) and the concentration of albumin determined by ELISA in accordance with manufacturer’s instructions (Bethyl labs, Montgomery, TX, E90-134). In vivo FITC Dextran assay Permeability was determined using a protocol previously described [4]. In brief mice were fasted for 6 hours (7am-1pm) and orally gavaged with 600mg/kg 4kDa FITC dextran (Sigma-Aldrich, St. Louis, MO) from a 125mg/ml solution. 1 hour later 120μl blood was collected, stored on ice in the dark and centrifuged at 12000xg for 3mins. Plasma was then diluted 1:2 in PBS and the concentration of FITC dextran determined by fluorescence spectroscopy (excitation at 485nm and emission at 535nm) relative to a linear standard curve made using diluted FITC dextran solutions in plasma from untreated mice. Flow cytometry of small intestinal lamina propria leukocytes The small intestine was dissected, washed in cold PBS and mesenteric fat removed. Peyer’s patches were excised, the intestine cut longitudinally and contents removed by washing three times in fresh cold PBS. The intestine was blotted on paper towel, washed a final time in cold PBS, and cut into pieces approximately 1 cm in length. Pieces were incubated for 30 minutes in HBSS media (without Ca2+ or Mg2+) (Life Technologies, NY, 14175) containing 5% FCS, Penicillin-streptomycin, 5mMEDTA at 37°C with agitation at 200rpm. The supernatant containing removed epithelial layer was aspirated, fresh media applied and the process repeated. The intestine was shaken in three changes of HBSS media (with Calcium chloride, Magnesium chloride and Magnesium sulphate) (Life Technologies, NY, 14025) containing 5% FCS and the pieces incubated for 35 minutes in the same HBSS media containing 0.5mg/ml Collagenase type II (Sigma-Aldrich, St. Louis, MO) and 40μg/ml DNaseI (Sigma-Aldrich, St. Louis, MO). The cell suspension was filtered through 100μm and 70μm filters each washed with 5ml cold HBSS media containing 5mM EDTA and cells pelleted by centrifugation at 400xg for 5 minutes. The approximate yield was between 5–10 million viable cells per intestine. 2 x 106 cells were then incubated with 1:1000 dilution of Live dead aqua (Invitrogen, NY) and 1:100 dilution of anti CD16/32 antibody (eBioscience, San Diego, CA, 93) for 30 minutes on ice in PBS, washed twice and incubated with the following antibody panel in PBS containing 2%FCS, 2mM EDTA: CD45 NC605 (30-F11), F4/80PECy7 (BM8) (from eBioscience, San Diego, CA), CD11b-FITC (M170), Siglec F-PE (E50-2440), CD11c-APC or CD11c-v450 (HL3) (from BD Biosciences, San Jose, CA) and MHCII APCCy7 (Biolegend, San Diego, CA, M5/114.15.2). Cells were fixed overnight in BD stabilizing fixative (BD Biosciences, San Jose, CA), washed twice and acquired on a BD Canto RUO flow cytometer. During analysis all gates were set relative to “fluorescence minus one” controls containing all fluorochromes minus the one of interest. Fluorescence microscopy The small intestine was isolated, the distal third dissected and rinsed with PBS. Tissue was then opened longitudinally, rolled mucosa side outwards and embedded in O.C.T. compound (Tissue-Tek, Sakura Finetek USA, Torrance, CA). 10μm sections were cut using a Leica Cryostat (Buffalo Grove, IL) and fixed with 4% methanol-free formaldehyde (Sigma-Aldrich, St Louis, MO). Sections were immunostained with antibodies directed against CD11b (Biolegend, San Diego, CA), MHCII FITC (Biolegend, San Diego, CA) and Siglec F (BD Biosciences, San Jose, CA) and mounted with SlowFade Gold Antifade medium containing 4′-6-Diamidino-2-phenylindole (Invitrogen, Carlsbad, CA). Tunnel staining was conducted using TACS2 TgT-Fluor In Situ Apoptosis detection kit (Trevigen, Gaithersburg) according to manufacturer’s instructions with cytonin used for permeabilisation and cobalt cation in the labelling reaction. Images were digitally acquired (Zeiss AxioCam HRC, Thornwood, NY) and analyzed using Image J software (rsb.info.nih.gov/ij/). For cell quantification, villi length was determined using Image J software measure tool and the number of cells quantified per villi. Three individual of mice were used per group, with a minimum of 20 images acquired. Monocyte tracking studies Peripheral blood mononuclear cells were isolated from the blood of wildtype donor mice and monocytes enriched using EasySep mouse monocyte enrichment kit (StemCell technologies, Vancouver, Canada). Monocytes were labelled with PKH26 (Sigma-Aldrich, St. Louis, MO, PKH26GL), washed twice in PBS and resuspended at 2.5 x 107 cells per ml in PBS prior to injection. 200μl (5 x 106 cells) were injected i.v. retro-orbitally. The proportions trafficking to the intestine were determined 3–5 days later by cell isolation and flow cytomtery as described above. RNA extraction and quantitative PCR The small intestine was opened longitudinally, contents removed and mucosa scraped away from muscularis using a clean razor blade and snap frozen in liquid nitrogen. RNA was extracted using Trizol (Life Technologies, NY) according to manufacturer’s instructions. 2μg RNA was converted to cDNA using a High-capacity cDNA conversion kit (Life Technoliges, Applied Biosystems, NY). Gene expression was measured by Quatnitative PCR using iTAQ university SYBR green super mix (Bio-Rad, Hercules, CA). The following primer sets were used for SYBR green Quantitative RT-PCR (5’-3’): β-actin FWD GGTTCTTTGCAGCTCCTTCGT, β-actin REV ATATCGTCATCCATCGCGAAC, CD11b FWD TGTGAGCAGCACTGAGATCC, CD11b REV ATGAGAGCCAAGAGCACCAG, CD11c FWD ACACAGTGTGCTCCAGTATGA, CD11c REV GCCCAGGGATATGTTCACAGC, TNFα FWD CCAGACCCTCACACTGAGATC, TNFα REV, CACTTGGTGGTTTGCTACGAC, IL-1β FWD CTTGGGATCCACACTCTCCAG, IL-1β REV AAATACCTGTGGCCTTGGGC, MCP-1 FWD AGGTCCCTGTCATGCTTCTG, MCP-1 REV GCTGCTGGTGATCCTCTTGT. Eicosanoid analysis The distal third of the small intestine (ileum) was weighed, supplemented with a cocktail consisting of 26 deuterated internal standards, homogenized with 1 ml 10% methanol per 5mg tissue on ice and briefly sonicated. Samples were then purified by solid phase extraction on Strata-X columns (Phenomenex, Torrance, CA), following the activation procedure provided by the distributor. Samples were eluted with 1 ml of 100% methanol, the eluent was dried under vacuum and dissolved in 50 µl of buffer A consisting of 60/40/0.02 water/acetonitril/acetic acid = 60/40/0.02 (v/v/v) and immediately used for analysis. Eicosanoids used for primary standards in standard curves as well as their deuterated analogs were from Cayman Chemicals (Ann Arbor, MI) and Biomol (Enzo Life Science, Framingdale, NY). Eicosanoids were analyzed by reverse-phase liquid chromatography, using a 1.7uM 2.1x100 mm BEH Shield Column (Waters, Milford, MA) and an Acquity UPLC system (Waters, Milford, MA) and mass spectrometry, using an AB SCIEX 6500 QTrap mass spectrometer equipped with an IonDrive Turbo V source (AB SCIEX, Framingham, MA), as previously described. Eicosanoids were quantitated by the stable isotope dilution method using standard curves from the deuterated internal standards. To calculate the amount of eicosanoids in a sample, ratios of peak areas between endogenous eicosanoids and matching deuterated internal eicosanoids were calculated. Ratios were converted to absolute amounts by linear regression analysis of standard curves generated under identical conditions. Measurement of intestinal permeability in Ussing chambers Segments of mid-jejunum and distal ileum were cut along the mesenteric border and mounted in Ussing chambers (Physiological instruments, San Diego, CA) exposing 0.09 cm2 of tissue area to 4 mL of circulating oxygenated Ringer’s buffer maintained at 37°C. The buffer consisted (in mM) of: 140 Na+, 5.2 K+, 1.2 Ca2+, 0.8 Mg2+, 120 Cl−, 25 HCO 3 −, 2.4 H 2 PO 4 −, 0.4 HPO 4 2−. Additionally, glucose (10 mM) was added to the serosal buffer as a source of energy, osmotically balanced by mannitol (10 mM) in the mucosal buffer. Agar–salt bridges were used to monitor the potential difference across the tissue and to inject the required short-circuit current (Isc) to maintain the potential difference at zero. This was registered by an automated voltage clamp and continuously recorded by computer. Conductance (G) was determined at baseline as an indicator of paracellular ion flux and expressed as mS/cm2. FITC labeled dextran (4 kDa, Sigma-Aldrich) was used as a probe to assess macromolecular permeability, and was added (2.2mg/mL final concentration) to the luminal buffer once equilibrium was reached. Serosal samples (200 uL) were taken at 30 min intervals for 2h and replaced with fresh buffer to maintain constant volume. The flux of FITC-dextran from the mucosa to the serosa was calculated as the average value of two consecutive stable flux periods (60–90 and 90–120 min) and expressed as pmol/cm2/h [26].

Discussion Obesity-associated chronic tissue inflammation is an important component of insulin resistance in obese subjects [35]. Recent studies have indicated that events arising in the gastrointestinal tract may play an etiologic role in driving systemic inflammation [1,2,4]. Thus, it has been well described that the gastrointestinal microbiota differ between the lean and obese state, and many studies have documented dysbiosis in obese rodents and humans [36,37]. This dysbiosis is associated with increased gut “leakiness” such that bacterial products, like LPS, and perhaps bacteria themselves, gain access to the systemic environment more readily in obesity [1,2]. Thus, it is hypothesised that signals derived from a leaky gastrointestinal tract promote systemic inflammation and insulin resistance in obesity. Interestingly, unlike the liver or adipose tissue, we do not find a greater degree of inflammation in the small intestine or colon of HFD fed/obese mice compared to normal chow fed mice. Thus, we found no evidence for lamina propria macrophage accumulation in the small intestine, expression of standard inflammatory mRNA markers were either unchanged or reduced in the HFD small intestine and colon, and histologic analysis showed no evidence of tissue damage. Prior studies of the colonic response to a HFD showed similar data, with no immune cell accumulation or tissue damage with respect to cellular composition or histological analysis [14]. However, other studies have reported 1.5–6 fold increases in TNF-α expression in the proximal colon and ileum [12,13,15,16], and, using an NF-κBeGFP reporter mouse, Ding et al showed foci of NF-κB activation along the length of the gastrointestinal tract following HFD feeding, which they concluded to be indicative of an inflammatory response [12]. One possible reason for this varying data could be the presence of varying microbiota offering different inflammatory potential. Indeed the inflammatory responses reported by Ding et al, were dependent on the presence of bacteria [12]. Alternatively, the whole tissue analyses presented here may not be sensitive enough to pick up highly localized regions of inflammation. Rather they give a broader of view of the immune status of the gastrointestinal tract. The majority of studies on obesity-associated intestinal permeability have been conducted in rodent models, where increased permeability caused by HFD or obesity is reproducibly observed [1,2,5,8]. In man, one pilot study found no correlation between obesity and gut permeability [38], whereas, another found an association between small intestinal permeability and metabolic syndrome parameters, such as waist and abdominal circumference [6]. Interestingly, detection of 16s rDNA sequences in the circulation might be predictive of type II diabetes progression [3]. Here we show that intestinal permeability occurs rapidly after HFD feeding, preceding the onset of obesity. In addition, we show that, although overall permeability to FITC dextran can be observed in obese Ob/Ob mice in the absence of a HFD, specific measurements associated with paracellular permeability, such as albumin leakage into the feces and transepithelial conductance, are specific to the HFD fed condition. One of the reasons for the difference in paracellular permeability between HFD fed and Ob/Ob mice could be the depletion of intestinal eosinophils, which was caused by HFD feeding, but not obesity alone. However, at present this remains an association as we have been unable to delineate a causal mechanism. Interestingly, a similar depletion of eosinophils also occurs in the adipose tissue following HFD feeding, and this coincides with polarization towards an M1-like macrophage phenotype and insulin resistance [39]. Eosinophils are highly granular leukocytes, generally found in small numbers (<5% of leukocytes) in the circulation and peripheral tissues [21]. The gastrointestinal tract is a notable exception to this, however, where tissue-resident eosinophils are reported to represent between 20–40% of leukocytes, a figure consistent with our data [40]. During inflammatory responses, such as allergy, eosinophils accumulate and produce pro-inflammatory and cytotoxic mediators with pathogenic consequences [19,20,21]. In the gastrointestinal tract this is true for a variety of conditions, such as allergic colitis, eosinophilic esophagitis and inflammatory bowel disease [19,20]. However, in the normophysiologic setting, eosinophils play a role in maintaining gastrointestinal homeostasis, including the maintenance of IgA-producing plasma cells, CD103+ dendritic cell and T cells, production and activation of TGF-β, and maintenance of the mucus layer [22,23]. Thus, we hypothesize that the depletion of intestinal eosinophils caused by a HFD could result in a state of nutritionally-induced relative immune deficiency and defective barrier function. Such an hypothesis is analogous to models of intestinal immune-deficiency caused by genetic deletion (e.g. TLR5-deficiency, ASC-deficiency) which result in penetration of bacterial products and the development of metabolic disease [41,42]. Further characterization of the intestinal immune system in HFD fed mice, the exposure of HFD fed mice to infectious challenge, and gain/loss of function experiments with eosinophil-deficient mice will be required to investigate this hypothesis fully.

Acknowledgments This work was performed with the support of the Flow Cytometry Core at the UC San Diego Center for AIDS Research (P30 AI036214), the VA San Diego Health Care System, and the San Diego Veterans Medical Research Foundation. The authors also wish to thank Bernd Schnabl and Peng Chen for advice on the initial study design.

Author Contributions Conceived and designed the experiments: AMFJ JMJ OQ JO. Performed the experiments: AMFJ AC MGG AMA. Analyzed the data: AMFJ AC MGG OQ. Wrote the paper: AMFJ JO.