Calprotectin protects against experimental colonic inflammation in mice

Abstract

Background and Purpose

Calprotectin is a heterodimer composed of two myeloid‐related proteins, S100A8 and S100A9, that is abundant in neutrophils and monocytes/macrophages. Faecal levels of calprotectin are used routinely to monitor inflammatory bowel disease activity.

Experimental Approach

We aimed to assess the role of calprotectin in intestinal inflammation, using the dextran sulfate sodium model of colitis in mice. Calprotectin was administered (50 or 100 μg·day⁻¹) by the intrarectal or by i.p. injection (50 μg·day⁻¹ only). The condition of the mice was characterized by morphological and biochemical methods.

Key Results

Intrarectal calprotectin protected significantly against colitis, as shown by lower levels of macroscopic and microscopic damage, colonic myeloperoxidase activity and decreased expression of TNFα and toll‐like receptor 4. IL‐17 production by spleen and mesenteric lymph node cells was reduced. Calprotectin had no effect on body weight loss or colonic thickening. There were no effects of calprotectin after i.p. injection. Calprotectin had virtually no effects in control, non‐colitic mice. Calprotectin had almost no effect on the colonic microbiota but enhanced barrier function. Treatment of rat IEC18 intestinal epithelial cells in vitro with calprotectin induced output of the chemokines CXL1 and CCL2, involving the receptor for advanced glycation end products‐ and NFκB.

Conclusion and Implications

Calprotectin exerted protective effects in experimental colitis when given by the intrarectal route, by actions that appear to involve effects on the epithelium.

Abbreviations

AP

alkaline phosphatase

ConA

concanavalin A

CP

calprotectin

DSS

dextran sulfate sodium

IBD

inflammatory bowel disease

MLC2

myosin light chain 2

MLNC

mesenteric lymph node cells

MPO

myeloperoxidase

RAGE

receptor for advanced glycation end products

TLR4

toll‐like receptor 4

Introduction

Calprotectin (CP) is a heterodimer of two S100 EF‐hand calcium‐binding proteins, also known as myeloid‐related proteins, S100A8 (MRP8) or CP‐10, and S100A9 (MRP14), expressed in neutrophils, macrophages, epithelial cells (including intestinal epithelial cells; Lee et al. 2012), T cells (Nguyen et al., 2015) and other cell types. In neutrophils, CP comprises 20–45% of the cytosolic proteins and plays a role in the innate immune response to bacteria and other microbial pathogens via a mechanism termed nutritional immunity (Kehl‐Fie et al., 2011). Levels of CP are substantially raised in several inflammatory processes, and it has been widely studied as a biomarker of disease activity, particularly in inflammatory bowel disease (IBD), where CP is measured in stool for diagnosis (Tibble and Bjarnason, 2001). CP is generally considered to be a pro‐inflammatory protein, acting as an alarmin in inflammatory sites via activation of http://www.guidetopharmacology.org/GRAC/ObjectDisplayForward?objectId=1754 (TLR4) and the http://www.guidetopharmacology.org/GRAC/ObjectDisplayForward?objectId=2843 (RAGE) (Vogl et al., 2007; Loser et al., 2010).

The exact role of CP in intestinal inflammation is not clear. For instance, TLR4 ligation by CP may enhance inflammation in the gut. However, as experimental colitis has been shown to be exacerbated in TLR4 KO mice and TLR4 agonists tend to ameliorate colitis (Sánchez de Medina et al., 2014), the effect of CP on this receptor in the inflamed intestine may be protective rather than deleterious. Indeed, TLR4 has been involved in the inflammatory reaction in the gut and also in bacterial clearance and epithelial repair. One example of such opposing roles of CP are ventilator‐induced lung injury, which is aggravated by high levels of CP (Kuipers et al., 2013), and acute lung injury, where the S100A8 subunit can induce IL‐10 expression starting a feedback loop that attenuates inflammatory damage (Hiroshima et al., 2014). In this regard, it is noteworthy that CP is elevated in pre‐term infants during the neonatal period (Rouge et al., 2010), presumably acting as an innate host defence element by protecting the immunologically immature bowel.

Hence, we hypothesized that CP may exert protective effects in intestinal inflammation by enhancing barrier function. In the experiments described here, we aimed to test this hypothesis, using the dextran sulfate sodium (DSS) model of colitis in mice.

Methods

Production of calprotectin

Expression and purification of CP was performed using a modified version of the original protocol (Hunter and Chazin, 1998), as described in Kehl‐Fie et al. (2011). Briefly, the S100A8 and S100A9 proteins were overexpressed in Escherichia. coli BL21 rosetta cells into inclusion bodies. The protein was unfolded in guanidinium hydrochloride and refolded by dialysis at 4°C in buffer containing 20 mM phosphate at pH 7.5. Purification involves a series of chromatographic steps, involving hydroxylapatite anion exchange, Source Q and Superdex 75 size‐exclusion chromatography. The two proteins preferentially form heterodimers over homodimers and the heterodimer, once formed, remains in this state (Hunter and Chazin, 1998). To further ensure heterodimerization, each batch of CP produced was verified by electrospray mass spectrometry. Lipopolysaccharides were removed using the ToxinEraser™ Endotoxin Removal Kit (Genscript) following the manufacturer's instructions.

Induction of DSS colitis and experimental design

All animal care and experimental procedures were approved by the Animal Welfare Committee of the University of Granada (ref. 2011‐349). Animal studies are reported in compliance with the ARRIVE guidelines (Kilkenny et al., 2010; McGrath and Lilley, 2015). Female C57BL/6 mice (2 months old) were obtained from Jackson Laboratory (Bar Harbor, ME, USA) and maintained in SPF conditions at the University of Granada Animal Facility (Biomedical Research Center), with a 12 h light–dark cycle and free access to food and water, for 10 days before use. A total of 99 female C57BL/6 mice were used.

Colitis was induced by adding DSS (2% w·v⁻¹) to the drinking water for 8 days (Pérez‐Navarro et al., 2005). The status of the mice was monitored individually by general examination and by measuring body weight loss. In addition, the presence of diarrhoea and haematochezia was noted (Ito et al., 2009). Food intake and water consumption were controlled every day. Mice that lost more than 20% of the initial body weight or showed signs of excessive suffering were to be humanely killed, but no such cases were registered. Three experiments were carried out. In the first experiment, mice were randomly assigned to four different groups (n = 10). The control (C) group did not receive DSS. The remaining mice were exposed to DSS‐supplemented water and additionally received by the intrarectal route 50 μg·day⁻¹ of CP (CP50 group), 100 μg·day⁻¹ (CP100 group) or the vehicle (β‐mercaptoethanol 10 mM in distilled water, DSS group). CP administration started at the same time as DSS supplementation (day 0) and was continued until mice were killed at day 8 by cervical dislocation. In the second experiment, the 50 μg·day⁻¹ dose of CP was tested by the intrarectal route in DSS colitic mice (CP50 group) and in non‐colitic mice (C‐CP50 group) and compared with the vehicle‐treated colitic (DSS) and non‐colitic (C) groups (n = 8 for all groups).

In the third experiment, 27 mice were randomly allocated into a control and a DSS group as above (n = 9), plus a CP group that received DSS and 50 μg·day⁻¹ by i.p. injection. The DSS group was injected daily with vehicle. The animals were followed for 8 days and killed as described above.

Assessment of colonic damage

The entire colon was weighed, and its length was measured under a constant load (2 g). Macroscopic damage was scored as follows: adhesions (0–3); hyperaemia (0–3); fibrosis (rigidity, 0–3); thickening (0–2); and other features (0–3). Two small segments were dissected for RNA isolation and histology, and the remainder was cut longitudinally into several pieces and snap frozen for the determination of biochemical parameters. Formalin‐fixed colonic tissue was processed for haematoxylin and eosin staining and scored on a 0–8 scale [erosion (0–2); infiltration (0–2); loss of crypts structure (0–2); submucosal thickening (0–2)]. Immunohistochemistry was also carried out to detect S100A8 (MRP‐8 antibody ab92331, Abcam, Cambridge UK), http://www.guidetopharmacology.org/GRAC/FamilyDisplayForward?familyId=930 (ref. 329400, Life Technologies, Rockford, IL, USA), ZO‐1 (ref. 617300, Life Technologies) and phosphorylated myosin light chain 2 (pMLC2, ab2480, Abcam), using the VECTASTAIN Elite ABC kit (Vectashield, PK‐6100) or the DAKO Real kit (Glostrup, Denmark), with Harris haematoxylin counterstaining.

Colonic http://www.guidetopharmacology.org/GRAC/ObjectDisplayForward?objectId=2789 (MPO) and alkaline phosphatase (AP) activities were measured spectrophotometrically as described. In addition, the sensitivity of AP to the specific inhibitor http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=7210 in vitro was measured (Lopez‐Posadas et al., 2011; Ortega‐Gonzalez et al., 2014a).

Total RNA was isolated by the Trizol method (Invitrogen, Barcelona, Spain) and checked for integrity by electrophoresis in 1% agarose gel (w·v⁻¹). Orally administered DSS interferes with qPCR amplification of cDNA derived from several tissues (Kerr et al., 2012). Thus, the Dynabeads® mRNA Purification Kit was used for the purification of DSS colonic mRNA from total RNA; 1 μg mRNA was subjected to reverse transcription (iScriptTM cDNA Synthesis Kit), and iQ™ Sybr® Green Supermix (Bio‐Rad, CA, USA) was used for amplification, and specific DNA sequences were amplified with a Biorad CFX connect real time PCR device (Alcobendas, Madrid, Spain). Primers used are shown in Supporting Information Table S1. Results are expressed as 2^‐ΔΔCt using GAPDH,18S and peptidyl‐prolyl cis‐trans isomerase B as reference genes.

Western blot analysis

Tissue samples were homogenized in lysis buffer (0.1% w/v SDS, 0.1% w/v sodium deoxycholate, 1% v/v Triton X‐100 in PBS) with protease inhibitor cocktail 1:200 (v/v) (Sigma, P9599) and phosphatase inhibitor cocktail 1:100 (v/v) (SC‐45045, Santa Cruz, Heidelberg, Germany). Then homogenates were sonicated and centrifuged at 10 000× g for 10 min at 4°C. Protein concentration was determined by the bicinchoninic acid assay (Canny et al., 2006). Samples were boiled for 5 min in Laemmli buffer (Biorad), separated by SDS‐PAGE, electroblotted to nitrocellulose membranes (pore size 0.25 μm) (Millipore, Madrid, Spain) and probed with the corresponding antibody. The bands were detected by enhanced chemiluminescence (PerkinElmer, Waltham, MA, USA). The primary antibodies were generally used at a 1:1000 dilution except where indicated and were obtained from the following: Cell Signalling (Danvers, MA, USA) (cyclin D1, ref. 2922; p‐MLC2 ref. 3675; MLC2 ref. 3672); Thermo Fisher Scientific (Waltham, MA, USA) (claudin‐4, ref. 32–9400); and Abcam (Cambridge, UK) (β actin, ref. ab6276, 1:500). The bands were quantified with the National Institute of Health software (Image J).

Secretion of cytokines by mesenteric lymph node cells and splenocytes

Mesenteric lymph nodes and spleen were extracted from the mice using sterile technique and dissected mechanically. Cells were washed once with fresh medium and were filtered using a 70 μM filter to obtain a mononuclear cell suspension. The cells were incubated at 10⁶ cells·mL⁻¹ and stimulated with concanavalin A (ConA) at 5 μg·mL⁻¹ in RPMI 1640 medium containing fetal bovine serum (10%), 2 mM L‐glutamine, 100 U·mL⁻¹ penicillin, 0.1 mg·mL⁻¹ streptomycin, 2.5 mg·mL⁻¹ amphotericin B and 0.05 mM β‐mercaptoethanol. The cells were cultured. Cell culture medium was collected after 48 h and assayed for cytokine content by elisa. The cytokines assayed were http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=4998, http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=4975, http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=4982, http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=4968 and http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=5073.

Flow cytometry

Single cell suspensions of freshly isolated splenocytes were washed in staining buffer (5% FBS and 0.02% NaN₃ in PBS), blocked with anti‐CD16/32 antibody for 5 min at room temperature and then stained for 30 min at 4°C with the following surface antibodies: anti‐CD3 APC, anti‐CD4 PerCP, anti‐CD8 FITC and anti‐Ly6G PE‐Cy7 (BD Pharmingen). Cell suspensions were then washed and analysed using a FACsCalibur flow cytometer (BD Biosciences) and FlowJo software (Version 9.3.1, Treestar).

For the in vitro stimulation of isolated splenocytes, cells were cultured as above plus phorbol myristate acetate (0.5 μg·mL⁻¹), ionomycin (2.5 μg·mL⁻¹) and 2 μM monensin (GolgiStop®), which were added simultaneously at the beginning of cell culture. Samples were incubated for 4 h at 37°C in a 5% CO₂ atmosphere, and then cells were stained with anti‐CD3 FITC and anti‐CD4 PerCP, fixed and permeabilized and stained overnight at 4°C with the following antibodies: anti‐IFNγ PE and anti‐IL‐4 APC. Cell suspensions were then washed and analysed.

Intestinal epithelial cell culture

Non‐transformed rat small intestinal epithelial cells, IEC18 (ECACC 88011801) (passages 25–50) were obtained from the Cell Culture Service of the University of Granada and cultured in DMEM containing fetal bovine serum (10%), 2 mM glutamine, 100 U·mL⁻¹ penicillin, 0.1 mg·mL⁻¹ streptomycin and 2.5 mg·mL⁻¹ amphotericin B. All experiments were carried out with monolayers reaching confluence. TLR4 knockdown IEC18 cells were obtained using reagents supplied by Santa Cruz Biotechnologies (Heidelberg, Germany). In order to explore signalling pathways, IEC18 confluent monolayers were exposed to http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=5934 (10 μM), a selective inhibitor of IκBα phosphorylation that blocks the NFκB signalling pathway or FPS‐ZM1 (10 μM) (Merck Millipore, Billerica, MA), a RAGE antagonist. All the inhibitors were dissolved in DMSO and were added to the culture medium 2 h before challenge.

DNA extraction, 16S rRNA gene amplification and analysis

For analysis of the microbiota, faecal samples were collected and homogenized in PBS, and bacterial genomic DNA was extracted from 200 μL aliquots after proteinase K and RNAse digestion using G‐spinTM columns (INTRON Biotechnology, Gyeonggi‐do, Korea). DNA concentration was determined using Quant‐IT PicoGreen reagent (Thermo Fischer), and DNA samples (about 3 ng) were used to amplify the V3‐V4 region of 16 S rRNA gene (Caporaso et al., 2011). PCR products (~450 bp) included extension tails which allowed sample barcoding and the addition of specific Illumina sequences in a second low‐cycle number PCR. Individual amplicon libraries were analysed using a Bioanalyzer 2100 (Agilent, Las Rozas, Spain), and a pool of samples was made in equimolar amounts. The pool was further cleaned, quantified and the exact concentration estimated by real‐time PCR (Kapa Biosystems, Wilmington, MA, USA). Finally, DNA samples were sequenced on an Illumina MiSeq Instrument under a 2 × 300 protocol at the Unidad de Genómica (Parque Científico de Madrid, Spain). Following sequencing, reads were quality filtered according to Illumina standard values, and demultiplexed and fastq files were mapped against the GreenGenes database using current applications of Base Space (metagenomics 16S, Illumina).

Data and statistical analysis

The data and statistical analysis comply with the recommendations on experimental design and analysis in pharmacology (Curtis et al., 2018). In all the experiments, samples were run in triplicate, and results are expressed as mean ± SEM. Differences among the means were tested for statistical significance by one‐way ANOVA and a posteriori Fisher least significant difference tests on preselected pairs. All the analyses were carried out with the GraphPad Prism 6 program. Differences were considered significant at P < 0.05.

Materials

Except where indicated, all the materials and primers were obtained from Sigma (St. Louis, MO, USA). Mouse elisa kits were obtained from eBioscience (San Diego, CA, USA). DSS was obtained from MP Biomedicals Santa Ana, CA, USA).

Nomenclature of targets and ligands

Key protein targets and ligands in this article are hyperlinked to corresponding entries in http:/http://www.guidetopharmacology.org, the common portal for data from the IUPHAR/BPS Guide to PHARMACOLOGY (Harding et al., 2018), and are permanently archived in the Concise Guide to PHARMACOLOGY 2017/2018 (Alexander et al., 2017a, 2017b, 2017c).

Results

DSS colitis

DSS colitis presented with body weight loss from the day 5 onward, diarrhoea and rectal bleeding, plus a general ‘sick’ appearance (Figure 1A and data not shown). The large intestine was hyperaemic, thickened and rigid, with loss of mucosal vascular pattern, abnormal crypt structure and crypt loss, leukocyte (neutrophil) infiltration and submucosal enlargement (Figure 1B,C and Table 1). Mice with colitis exhibited colonic thickening and damage score (Figure 1E,F) and an increase in colonic AP activity, an indicator of epithelial stress and inflammation (Lopez‐Posadas et al., 2011), associated with augmented in vitro sensitivity to the specific inhibitor levamisole (Figure 1G,H). DSS colitis was also associated with an increase in the neutrophil markers MPO and S100A8 (Figure 2). Immunohistochemical analysis revealed that S100A8 bearing cells were infiltrating the mucosa and submucosa, with no expression at the epithelial level (Figure 2C). Colitis was associated also with splenomegaly (Figure 1D).

Figure 1.

Effect of intrarectal CP on body weight loss and colonic macroscopic parameters in DSS colitic mice. Colitis was induced with 2% DSS and treated with 50 or 100 μg·day⁻¹ of CP. (A) Body weight, expressed as percentage of initial weight. (B) Representative colon images. (C) Representative histology. (D) Spleen weight. (E) Colonic weight/length ratio. (F) Macroscopic score. (G) Colonic AP activity. (H) Sensitivity of AP activity to levamisole in vitro. Data shown are means ± SEM; n = 10 mice. ⁺ P < 0.05, significantly different from the control group; *P < 0.05, significantly different from the DSS group. All groups groups were significantly different from the control from day 5 onward in panel (A) (not shown).

Table 1.

Histological score of DSS colitic mice were treated with CP intrarectally

	Erosion	Infiltration	Loss of crypt structure	Submucosal thickening	Total score
Control	0 ± 0	0 ± 0	0 ± 0	0 ± 0	0 ± 0
DSS	1.2 ± 0.3	1.3 ± 0.3	1.3 ± 0.4	0.3 ± 0.2	4.0 ± 1.0*
CP 50	0.6 ± 0.3	1.1 ± 0.3	0.8 ± 0.2	0.1 ± 0.1	2.5 ± 0.8*
CP 100	0.5 ± 0.2**	0.9 ± 0.2	0.8 ± 0.2	0 ± 0	2.1 ± 0.4*,†

Data shown are means ± SEM; n = 10 mice. *P < 0.05, significantly different from the control group. ^† P < 0.05, significantly different from the DSS group.

Figure 2.

Effect of intrarectal CP on colonic microscopic parameters in DSS colitic mice. (A) Colonic MPO activity. (B) Colonic IL‐22, S100A8 and TLR4 mRNA levels. (C) MRP8 (S100A8) immunostaining. (D) Colonic mRNA levels of various cytokines and FOXP3. Data shown are means ± SEM; n = 10 mice. ⁺ P < 0.05, significantly different from the control group; *P < 0.05, significantly different from the DSS group.

Effect of intrarectal CP on DSS colitis in mice

CP‐treated mice showed a slight reduction in body weight loss, compared to the DSS control, which was significant only for the CP50 group on the last day (Figure 1A). CP had no effect on the colonic weight:length ratio (Figure 1E). The macroscopic damage score was halved with both doses of CP compared with the DSS group, but this failed to reach statistical significance (Figure 1F). A dose‐dependent reduction of the microscopic damage score was also observed (significant with the higher dose only, Table 1), corresponding to preservation of the mucosal structure, particularly at the epithelial level (Figure 1C). MPO activity was fully normalized at the 50 μg·day⁻¹ dose, and it was actually below the non‐colitic control at 100 μg·day⁻¹ (Figure 2A). Comparable results were observed with S100A8 (Figure 2B,C). Similarly, colonic AP activity and sensitivity to levamisole were generally lower in both treated groups, although only with the lower CP dose was significance achieved (Figure 1G,H). Spleen size was significantly reduced by the higher CP dose (Figure 1D).

Effect of intrarectal CP on colonic expression of inflammatory markers by RT‐PCR

The colonic expression of several cytokines was sharply up‐regulated, including IL‐10, TNFα, IL‐4 and IFN‐γ (Figure 2D). In addition, the colonic mRNA levels of the regulatory T cell marker Foxp3 and the antibacterial peptide Reg3γ were also markedly increased, but the high variability in the DSS group for the latter marker prevented significance difference from the control group. CP treatment resulted in a reduction of the expression of these inflammatory markers, which was significant for TNFα, IL‐10, FOXP3, REG3γ and IFN‐γ at one or both doses. mRNA levels of IL‐22 were also lower than in the DSS group but without reaching statistical significance (Figure 2B). Because CP has been described as a ligand for TLR4, we also measured the expression of this receptor (Figure 2B). There was no effect of DSS colitis on this parameter, but both doses of CP decreased its mRNA levels significantly.

Effect of intrarectal CP on cytokine secretion by splenocytes ex vivo

In order to evaluate the immunological state within the gut environment, we cultured mouse splenocytes and mesenteric lymph node cells (MLNC) ex vivo, either in the presence or absence of the T cell stimulator ConA. IL‐17A was significantly reduced in those groups that received CP when splenocytes (Figure 3A) and MLNC (Figure 3B) were exposed to ConA. Conversely, IL‐10 production was unimpaired in both splenocytes and MLNC under ConA stimulation (Figure 3C,D). Basal IL‐10 release was in turn reduced in splenocytes from CP‐treated mice, while that in MLNC was unchanged (Figure 3C,D). elisa measurement of cytokines in cell supernatants did not show a consistent trend in the case of IFN‐γ, TNFα and IL‐6 (Supporting Information Figure S1).

Figure 3.

Effect of intrarectal CP on spleen and mesenteric mononuclear cells and on spleen leukocyte populations. Cytokine secretion by spleen (A, C) or MLNC (B, D) ex vivo in mice with DSS colitis treated with intrarectal CP. (A, B) IL‐17 and (C, D) IL‐10 production. Panels (E, F): spleen leukocyte populations. Relative amount of (E) neutrophils, CD4 lymphocytes, CD8 lymphocytes and (F) expression of IFN‐γand IL‐4 by CD4 T lymphocytes (original scatter plots in Supporting Information Figure S2). (G) Histograms of peripheral blood Ly6G positive events in the control, DSS and CP50 groups. Data shown are means ± SEM; n = 10 mice. ⁺ P < 0.05, significantly different from the control group; *P < 0.05, significantly different from the DSS group.

Finally, splenocytes and whole peripheral blood cells were analysed by flow cytometry. DSS colitis was associated with a marked increase in blood neutrophil levels, which was reduced in the CP groups (Figure 3G). There were no significant changes in the numbers of CD4 and CD8 lymphocytes (Figure 3E). In the spleen, CP increased, dose‐dependently, the number of both IL‐4+ and IFN‐γ + CD4+ cells, even though this was unaffected by DSS colitis (significant for the 100 μg dose only, Figure 3F; see also Supporting Information Figure S2).

Effect of CP on non‐colitic, control mice

In order to test whether CP may exert pro‐inflammatory effects in vivo, we performed a second in vivo experiment in which CP (50 μg·kg⁻¹) or vehicle was administered intrarectally to normal, non‐colitic control mice. DSS colitic groups treated in parallel with CP or vehicle were also included. Administration of intrarectal CP had no effect on non‐colitic mice in terms of body weight, colonic damage score, histological analysis or weight:length ratio, colonic MPO or AP activity (Figures 4 and 5; Supporting Information Figure S3 and data not shown). The protective effect of CP on experimental colitis was confirmed in this second experiment, with improved histological scores and MPO activity (Figures 4 and 5). The same trend towards weight gain in the last day was observed, although in this case, it was not significant. AP activity was unchanged, unlike the first experiment (Supporting Information Figure S3).

Figure 4.

Effect of CP on non‐colitic mice and DSS colitic mice. Colitis was induced with 2% DSS in half of the mice and treated with 50 μg·day⁻¹ of CP or vehicle. The other half did not receive DSS and was treated with CP or vehicle. (A) Body weight, expressed as percentage of initial weight. (B) Colonic MPO activity. (C) Colonic claudin 4 expression and MLC2 phosphorylation by Western blot. (D) Colonic cyclin D₁ expression by Western blot. (E) Colonic expression of Ccl2/MCP‐1 and Cxcl1/GROαmeasured by RT‐PCR. Actin was used as loading control in claudin 4 and cyclin D₁ Western blots, while pan‐MLC2 was used in the case of pMLC2. Data shown are means ± SEM; n = 8 mice. ⁺ P < 0.05, significantly different from the control group; *P < 0.05, significantly different from the DSS group.

Figure 5.

Histological analysis of non‐colitic mice and DSS colitic mice treated with CP (50 μg·day⁻¹). (A) Haematoxylin and eosin staining. (B) Immunohistochemical staining for claudin 4, pMLC2 and ZO‐1. (C) Quantitation of immunostaining. Data shown are means ± SEM; n = 8 mice. ⁺ P < 0.05, significantly different from the control group; *P < 0.05, significantly different from the DSS group.

Colonic tissue samples were also used in this experiment to measure the expression (mRNA) of the chemokines http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=4423 and http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=4407 The mRNA for both chemokines were markedly up‐regulated in DSS colitic mice, but their levels were significantly reduced in CP‐treated mice (Figure 4E). Conversely, the colonic expression of IL‐1β, IL‐6 and IL‐12 was clearly up‐regulated by colitis but unaffected by CP treatment (Figure 6A). The expression of TLR4 was again lower in both CP groups, although this difference was significant only for those with colitis (Figure 6B). We additionally measured the expression of http://www.guidetopharmacology.org/GRAC/ObjectDisplayForward?objectId=1752, http://www.guidetopharmacology.org/GRAC/ObjectDisplayForward?objectId=1755 and http://www.guidetopharmacology.org/GRAC/ObjectDisplayForward?objectId=1759 (Figure 6B). TLR2 and TLR9 were significantly up‐regulated by DSS colitis, while TLR5 failed to reach significance due to the distorting effect of the disproportionate increase brought about by CP in colitic mice. In contrast, CP treatment resulted in a significant reduction of TLR2, while TLR9 levels were lower but without reaching statistical significance. There was no effect of CP on basal TLR expression.

Figure 6.

Colonic expression of various cytokines and TLRs in noncolitic mice and DSS colitic mice treated with CP (50 μg·day⁻¹). Colitis was induced with 2% DSS in half of the mice and treated with 50 μg·day⁻¹ of CP or vehicle. The other half did not receive DSS and was treated with CP or vehicle. (A) Cytokines. (B) TLRs. Data shown are means ± SEM; n = 8 mice. ⁺ P < 0.05, significantly different from the control group; *P < 0.05, significantly different from the DSS group.

We also looked at epithelial barrier integrity markers in both non‐colitic and colitic mice. The protein levels of claudin 4, a component of tight junctions, were lower in DSS colitic mice as assessed by Western blot, but without reaching statistical significance (Figure 4C). IHC analysis confirmed this finding and suggested a protective effect of CP (non‐significant, Figure 5). MLC‐2 phosphorylation as assessed by Western blot was augmented in CP‐treated colitic mice, even though DSS colitis had no effect at this level (Figure 4C). IHC images provide further detail, showing a shift of pMLC2 immunostaining from the bottom of the crypts to the surface in DSS colitis in vehicle‐ but not CP‐treated mice (Figure 5). On the other hand, ZO‐1 immunostaining was prominent at the crypt bottom in non‐colitic mice, and this signal was diminished with DSS colitis. This effect was of lower magnitude in the CP50 group. Thus, CP exerted some protective effects at the barrier function level.

In addition, DSS colitic mice exhibited a down‐regulated expression of cyclin D1 (Figure 4D). There was no effect of CP on cyclin D1, but protein levels were somewhat higher in both CP groups compared with the respective controls. We also measured the expression of IL‐22 and IL‐27, which have been involved in the epithelial adaptive response to colitis (Figure 6B). In both instances, there was a marked up‐regulation with colitis, but no effect of CP. Thus, CP appeared to have no effect on epithelial proliferation.

Effect of intrarectal CP on the colonic microbiota

Metagenomic analysis of faecal samples established the occurrence of a significant shift in the composition of the microbiota in mice with DSS colitis when compared with the corresponding controls. As shown in Figure 7A, DSS colitis was associated with a reduction of Actinobacteria and Verrucomicrobia (the latter just short of significance), and an increase in Chloroflexi and Deferribacteres, with no change in the majoritary phyla or Thermotogae (Figure 7A and Supporting Information Figure S4). The reduction of Actinobacteria and Verrucomicrobia is accounted for to a great extent by a lower count of Bifidobacterium and Akkermansia muciniphila respectively (the latter non‐significant, Figure 7B). Similarly, the increase in Deferribacteres is explained by the changes in the genus Mucispirillium. On the other hand, DSS colitis resulted in a dramatic reduction in Lactobacillus as well as Allobaculum (Erysipelotrichales), Anoxybacillus amylolyticus (Bacillales), Butyricimonas synergistica and Dysgonomonas wimpennyi (Bacteroidales, P = 0.08). Conversely, there was a higher presence of Bacteroides and Enterobacteriaceae (particularly Escherichia and Serratia), Odoribacter (Bacteroidales), Flavobacterium (Flavobacteriales), Coprobacillus (Erysipelotrichales), Blautia and Oscillospira, but not Clostridium (Clostridiales) (Figure 7B,C).

Figure 7.

Metagenomic analysis of faecal samples in noncolitic mice and DSS colitic mice treated with CP (50 μg·day⁻¹). (A) Phyla exhibiting significant modifications. (B) Selected bacterial families, genus and species significantly changed. Additional data pertaining taxonomical categories unaffected by the experimental interventions are shown in Supporting Information Figure S4. Data shown are means ± SEM; n = 8 mice. ⁺ P < 0.05, significantly different from the control group; *P < 0.05, significantly different from the DSS group.

CP increased the counts matching Akkermansia municiphila (and Verrucomicrobia) in non‐colitic mice, but it had little or no effect in colitic mice. In addition, CP treatment decreased Bacteroides in samples from colitic mice. There was no other effect of CP on the microbiota.

Effect of intraperitoneal CP on mouse DSS colitis

In order to identify the possible systemic effects of CP, we performed a third experiment, in which the protein was administered by i.p.injection to DSS colitic mice, following a protocol otherwise identical to that of the first experiment. As shown in Table 2, there was no effect of CP under these experimental conditions, except for a marginally higher spleen weight.

Table 2.

Effect of intraperitoneal CP in DSS colitic mice

	Weight/length ratio (mg·cm−1)	Colonic MPO activity (mU·mg·prot−1)	Colonic AP activity (mU·mg−1)	Percentage of initial weight	Spleen weight (mg)
Control	20.2 ± 1.2	227.6 ± 44.5	98.8 ± 8.8	102.2 ± 1.3	681.7 ± 2
DSS	33.1 ± 1.2*	746.3 ± 95.5*	318.5 ± 27.6*	78.8 ± 1.0*	697.7 ± 3
CP 50	32.0 ± 1.3*	713.4 ± 86.4*	265.4 ± 21.0*	80.2 ± 1.3*	806.7 ± 4*,†

Mean values were significantly different from those of the control group or DSS group. Data shown are means ± SEM from experiments conducted on nine mice. *P < 0.05, significantly different from the control group. ^† P < 0.05, significantly different from the DSS group.

Effect of CP on epithelial cells in vitro

We additionally examined the effect of CP on intestinal epithelial cells in vitro. We used IEC18 cells, a non‐tumoural cell line. CP elicited substantial cytokine secretion at 10 μM (equivalent to 360 μg·mL⁻¹), albeit of a lower magnitude than http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=5019 (Figure 8A,B). As CP has been described to act via TLR4 and RAGE ligation, we assessed the impact of receptor interference using gene knockdown and a specific antagonist respectively. Gene knockdown was confirmed at the mRNA level (Supporting Information Figure S5). As expected, TLR4 silencing resulted in significant inhibition of LPS‐elicited cytokine secretion, but it had no effect on CP‐evoked secretion of CXCL1 or CCL2 (Figure 8C,D). In turn, the RAGE antagonist FPS‐ZM1 almost abolished CXCL1 and CCL2 secretion elicited by CP, with a lower effect on the LPS response (also a RAGE ligand; Yamamoto et al., 2011). The NFκB inhibitor Bay11‐7082 produced an almost complete inhibition in both cases (Figure 8E,F).

Figure 8.

Effect of CP on IEC18 intestinal epithelial cells. (A, B) Effect on the secretion of CXCL1 and CCL2 (n = 7). (C, D) Effects of TLR4 knockdown on the effect of CP on CXCL1 and CCL2 secretion. (E, F) Effects of pharmacological inhibition of NFκB (Bay11‐7082, 10 μM) or RAGE (FPS‐ZM1, 10 μM) on the effect of CP on CXCL1 and CCL2 secretion. Data shown are means ± SEM; n = 9‐10 mice. ⁺ P < 0.05, significantly different from control; *P < 0.05 significantly different from the stimulated group.

Discussion

Our results demonstrate that exogenous CP exerts protective effects in DSS colitis mice when administered by the intrarectal route, consistent with a local rather than a systemic mechanism, although it remains possible that CP is not readily absorbed when given by i.p. injection. The main source of CP in IBD and experimental colitis are presumably infiltrating neutrophils, which are recruited avidly to active mucosal lesions in both cases, ultimately resulting in increased faecal CP levels (Krawisz et al., 1984; Striz and Trebichavsky, 2004; Guardiola et al., 2014) either by transepithelial migration of neutrophils or by direct protein leakage (Roseth et al., 1999). In addition, the intestinal epithelium may produce S100A8 and/or S100A9 (Lee et al., 2012; Zindl et al., 2013), although this is disputed (Holgersen et al., 2014) and was not detected by us.

Intrarectally administered CP may act by several mechanisms. First, CP may exert antibacterial actions, possibly in areas close to the epithelium or in sites of epithelial injury with bacterial translocation (Sánchez de Medina et al., 2014). A related possibility is modulation of the microbiota, that is, a prebiotic mechanism. Metagenomic analysis of faecal microbiota showed a profound alteration of bacterial composition, consistent with previous studies of the microbiota in DSS colitis and in colonic inflammation in general (Schwab et al., 2014; Zhang et al., 2016). CP treatment resulted in significant but rather limited effects at this level, that is, a reduction of only Bacteroides in the DSS CP group. While Akkermansia muciniphila, a bacteria implicated in the maintenance of mucosal barrier function (Reunanen et al., 2015), was increased by CP, this effect was restricted to non‐colitic mice. Thus, CP displays prebiotic actions, but these are far from prominent.

Second, CP may act as an antioxidant in injured mucosal areas (Lim et al., 2011). Third, CP may modulate epithelial cell function, presumably by activation of apical TLR4 or RAGE receptors (Ehrchen et al., 2009). The latter mechanism has been traditionally ascribed a pro‐inflammatory role. However, although mucosal CP may act as an alarmin and chemoattractant, recent studies suggest that the predominant role of CP and the subunit S100A8 and S100A9 homodimers may be anti‐inflammatory (Averill et al., 2011; Geczy et al., 2014; Hiroshima et al., 2014). The analysis is complicated by the particular characteristics of the gut, where immune receptor activation may actually result in protection from inflammation. Thus, although TLR4 ligation elicits cell activation in immune and epithelial cells, it also modulates epithelial dynamics to face the challenge imposed by mucosal inflammation, in which epithelial proliferation and restitution are important for healing and protection from ulceration. Thus, TLR4 KO mice are more susceptible to experimental colitis, and similar results have been obtained for a number of TLRs and related genes (Sánchez de Medina et al., 2014). In turn, RAGE has been linked to colitis or IBD, as well as colitis‐related cancer, and has been suggested to be involved in leukocyte transepithelial migration (Wang et al., 2015). However, RAGE KO mice show no change in sensitivity to DSS colitis (Däbritz et al., 2011).

Our in vitro data demonstrate that CP exerts direct actions on epithelial cells, suggesting that epithelial modulation may be involved in the anti‐inflammatory effect. Immunomodulation was attained at concentrations expected to be readily reached in vivo with the doses assayed (i.e. 100 μg in ~0.1 mL) and was dependent on RAGE ligation and activation of NFκB, and independent of TLR4. In connection with the latter, it is interesting to note that TLR4 expression was consistently down‐regulated by CP in vivo, even though it was not affected by DSS colitis. This was a specific effect, as other TLRs, namely, TLR2, TLR5 and TLR9, were augmented in colitic mice but less so in CP‐treated groups (with the notable exception of TLR5, which was further up‐regulated), and they were unchanged in the absence of inflammation. Although the consequences of enhanced secretion of cytokines, evoked by CP, from epithelial cells are debatable, it is important to note that CXCL1‐KO mice are more sensitive to DSS colitis (Shea‐Donohue et al., 2008). Of note, CP had no deleterious effects in non‐colitic mice, as shown in the second experiment, confirming that epithelial immunomodulation does not cause inflammation. Further, the colonic expression of CXCL1 and CCL1 was down‐regulated in DSS colitic mice receiving CP. It is possible that such chemokine up‐regulation leads to enhanced barrier function, ultimately resulting in a dampened inflammatory response (Sánchez de Medina et al., 2014). As observed with CP, non‐digestible oligosaccharides, which are non‐toxic and have prebiotic properties, increase these chemokines, acting as TLR4 ligands in the intestinal epithelium (Ortega‐Gonzalez et al., 2014b). The relevance of this action is currently unknown, but experiments are underway to further characterize the effects of CP at this level. Whether the mechanism of CP is based on epithelial modulation or not, it resulted in down‐regulated IL‐17A production by MLNC without reducing IL‐10, suggesting a shift of the immune balance, as described recently in the lung, where CP showed an IL‐10 inducing effect (Hiroshima et al., 2014).

It is relevant also to consider a possible effect on barrier function. CP had a significant effect on MLC2 phosphorylation, which plays a major role in the regulation of epithelial permeability by pulling actin filaments associated with the tight junctions (Cunningham and Turner, 2012). This action is normally associated with increased paracellular permeability. DSS colitis did not affect overall MLC2 phosphorylation, but IHC analysis revealed a shift in pMLC2 immunostaining, with augmented signal at the crypt surface compared with the crypt base. CP treatment resulted in increased MLC2 phosphorylation, but the crypt base/surface pattern appeared to be less disturbed. Thus, our data suggest that CP may enhance barrier function at the crypt surface. On the other hand, the results obtained point to a lower expression of both claudin 4 and ZO‐1 in DSS colitis, which was significant only for the latter. There was some evidence of protection by CP at this level, which was significant only for ZO‐1. Cyclin D1 was markedly down‐regulated in DSS colitis, consistent with reduced epithelial proliferation, and was not significantly modified by CP, indicating a lack of effect on epithelial proliferation. Conversely, IL‐22 and IL‐27, which are involved in the epithelial response to inflammation, were markedly up‐regulated in colitic animals, but again with no effect of CP. Hence, any effect of CP on barrier function is modest at best.

It is interesting to consider the biological significance of the amounts of CP measured. Assuming a CP faecal level of ~50 μg·g⁻¹ as the upper normal limit (Guardiola et al., 2014), the amount of CP in the colonic lumen in non‐colitic mice would be approximately 20 μg, since the normal faecal output in mice is about 0.4 g·day⁻¹. Thus, the doses assayed in our study correspond to higher than normal luminal levels and are probably in the range of those measured in colitis in humans. This raises the possibility that ‘physiological’ CP may have a limiting role in the colitic response. That is, CP would be protective despite its being up‐regulated as part of the inflammatory response, much in the same way that the anti‐inflammatory cytokine IL‐10 is increased in experimental colitis (Martinez‐Augustin et al., 2008). In this regard, it is interesting to consider the phenotype of S100a8/S100a9 KO mice. S100a8 KO mice are not viable and die before birth (Passey et al., 1999). The S100a9 KO mice are on the contrary viable and apparently normal, although they lack S100A8/S100A9 heterodimers (i.e. CP) (Hobbs et al., 2003; Manitz et al., 2003). The effects of the absence of CP on neutrophil biology is not settled, with almost no effect detected in one study (Hobbs et al., 2003) and reduced chemoattractant response and expression of CD11b in another (Manitz et al., 2003). In vivo effects are also controversial, with no differences observed in the models of peritonitis, skin inflammation and urinary tract infection (Hobbs et al., 2003; Manitz et al., 2003; Dessing et al., 2010), while a higher susceptibility of S100a9 KO mice to oedema formation after paw infection with Leishmania (Contreras et al., 2013) and impaired tissue repair after renal ischaemia–reperfusion injury (Dessing et al., 2014) have been observed. Unfortunately, there are no reports on the effects on experimental colitis.

In conclusion, our data indicated that CP exerted protective effects in experimental colitis when given by the intrarectal route, suggesting the possibility of a similar beneficial role in IBD patients. Our study opens up a possible clinical application of CP or CP related drugs, although further experimentation is required to explore the mechanistic and therapeutic aspects involved.

Author contributions

W.C., M.D.S., A.Z., F.S.M. and O.M.A. contributed to the conception and design of the study. C.J.A., B.O. and M.A.A. performed the experiments. W.C. provided CP. F.S.M. and O.M.A. wrote the first draft of the manuscript. All the authors contributed to the analysis and interpretation of data, critically reviewed and approved the final draft.

Conflict of interest

The authors declare no conflicts of interest. F.S.M. and O.M.A. have received lecture fees and/or research support from Hospira, Pfizer, Sanofi, Biosearch Life, Bioiberica, Amino Up Chemical, APC Europe and the Spanish Generics Association.

Declaration of transparency and scientific rigour

This http://onlinelibrary.wiley.com/doi/10.1111/bph.13405/abstract acknowledges that this paper adheres to the principles for transparent reporting and scientific rigour of preclinical research recommended by funding agencies, publishers and other organisations engaged with supporting research.

Supporting information

Figure S1 Cytokine secretion by spleen (A,B,C) or mesenteric lymph node cells (D,E,F) ex vivo in mice with DSS colitis treated with intrarectal CP. Colitis was induced with 2.5% DSS as described in Methods and treated with 50 or 100 μg·day⁻¹ of CP intrarectally. Cells were isolated and cultured, with or without concanavalin A, and the levels of cytokines in the supernatant were measured by elisa. (A,D) IFN‐γ, (B,E) TNFαand (C,F) IL6 production. Values are means ± SEM. ⁺ P < 0.05 versus Control group; *P < 0.05 versus. DSS group. Data points represent means ± SEM from experiments conducted on 10 mice.

Figure S2 Scatter plots corresponding to the IFN‐γand IL‐4 measurement in Figure 3F. Data correspond to experiments conducted on 10 mice.

Figure S3 Effect of CP on AP in noncolitic mice and DSS colitic Mice. Colitis was induced with 2% DSS in half of the mice and treated with 50 μg·day⁻¹ of CP or vehicle. The other half did not receive DSS and was treated with CP or vehicle. (A) Colonic AP activity. (B) Sensitivity of AP activity to levamisole in vitro. ⁺ P < 0.05 versus. Control group. Data points represent means ± SEM from experiments conducted on 8 mice.

Figure S4 Metagenomic analysis of faecal samples in noncolitic mice and DSS colitic mice treated with CP (50 μg·day⁻¹). Taxonomical categories unaffected by the experimental interventions are displayed. ⁺ P < 0.05 versus. Control group. Data points represent means ± SEM from experiments conducted on 8 mice.

Figure S5 Expression of TLR4 gene by TLR4 knockdown IEC18 cells versus. control. TLR4 mRNA levels measured by RT‐qPCR are shown. Values are means ± SEM. ⁺ P < 0.05 versus. Control group (n = 6).

Table S1 Primers (sequence 5′‐3′).

Aranda, C. J. , Ocón, B. , Arredondo‐Amador, M. , Suárez, M. D. , Zarzuelo, A. , Chazin, W. J. , Martínez‐Augustin, O. , and Sánchez de Medina, F. (2018) Calprotectin protects against experimental colonic inflammation in mice. British Journal of Pharmacology, 175: 3797–3812. 10.1111/bph.14449.