- Open Access
1,25(OH)2D3 deficiency-induced gut microbial dysbiosis degrades the colonic mucus barrier in Cyp27b1 knockout mouse model
- Wenjing Zhu†1,
- Jiayao Yan†2,
- Chunchun Zhi4,
- Qianwen Zhou1 and
- Xiaoqin Yuan1, 3Email authorView ORCID ID profile
© The Author(s) 2019
- Received: 1 November 2018
- Accepted: 11 February 2019
- Published: 20 February 2019
The relationship between disturbances of the gut microbiota and 1,25(OH)2D3 deficiency has been established both in humans and animal models with a vitamin D poor diet or a lack of sun exposure. Our prior study has demonstrated that Cyp27b1−/− (Cyp27b1 knockout) mice that could not produce 1,25(OH)2D3 had significant colon inflammation phenotypes. However, whether and how 1,25(OH)2D3 deficiency due to the genetic deletion controls the gut homeostasis and modulates the composition of the gut microbiota remains to be explored.
1,25(OH)2D3 deficiency impair the composition of the gut microbiota and metabolite in Cyp27b1−/− mice, including Akkermansia muciniphila, Solitalea Canadensis, Bacteroides-acidifaciens, Bacteroides plebeius and SCFA production. 1,25(OH)2D3 deficiency cause the thinner colonic mucus layer and increase the translocation of the bacteria to the mesenteric lymph nodes. We also found 1,25(OH)2D3 supplement significantly decreased Akkermansia muciniphila abundance in fecal samples of Cyp27b1−/− mice.
Deficiency in 1,25(OH)2D3 impairs the composition of gut microbiota leading to disruption of intestinal epithelial barrier homeostasis and induction of colonic inflammation. This study highlights the association between 1,25(OH)2D3 status, the gut microbiota and the colonic mucus barrier that is regarded as a primary defense against enteric pathogens.
- 1,25(OH)2D3 deficiency
- Inflammatory bowel disease
- Gut microbiota
- Colonic mucus barrier
Vitamin D is a prohormone that can be converted to the active form of 1,25-dihydroxyvitamin D3 [1,25(OH)2D3] by 1α-hydroxylase encoded by the Cyp27b1 gene . In addition to its role in regulating Ca2+ and Pi transport and bone mineralization, 1,25(OH)2D3 also possesses various biological activities through binding vitamin D receptor (VDR), a high-affinity nuclear receptor that transcriptionally regulates its target genes . There is growing epidemiological evidence demonstrating that vitamin D-deficiency (commonly defined as serum 25(OH)D < 20 ng/ml) or vitamin D- insufficiency (serum 25(OH)D < 30 ng/ml) is related to an increased risk of inflammatory bowel disease (IBD) [3, 4]. Several studies have reported that vitamin D deficiency is often observed in patients with newly diagnosed IBD [5–7]. Conversely, high vitamin D intake can lower IBD risk . In mouse models, 1,25(OH)2D3 deficiency or VDR knockout increased the risk of colitis [9–11]. In either trinitrobenzene sulphonic acid (TNBS)- or dextran sodium sulphate (DSS)-induced colitis mice models, administration of 1,25(OH)2D3 effectively reduced the disease severity [9, 12]. Therefore, vitamin D might play a protective role for IBD. The role vitamin D plays in the pathogenesis of IBD is complex and not well defined. Some investigations have shown that 1,25(OH)2D3 has a pivotal role in the development of IBD via regulating innate and adaptive immune response , autophagy  or gut barrier integrity . Our prior study revealed that Cyp27b1 disruption induced colon inflammation in mice by increasing oxidative stress and DNA damage consequently leading to induction of cell senescence and mass generation of senescence-associated secretory factors .
Inflammatory bowel disease, i.e. Crohn’s disease and ulcerative colitis, is chronic relapsing and incurable inflammatory conditions of the bowel with an increasing trend of incidence and prevalence . IBD is one of major health problems in the Western world as about 0.5% of the general population are afflicted with this disease . Although the pathogenic factors have not been clarified yet, recent studies have demonstrated that intestinal microbiota might have an essential role in the development of IBD. Many studies demonstrated a lower diversity of the microbiome in IBD patients as compared with healthy controls, but a higher abundance of certain bacterial strains such as Enterobacteriaceae, and a stronger mucosal adherence of the bacteria [18, 19]. Although the gut microbiota has a role in IBD pathogenesis, the exact role of dysbiosis is far from clear. Gut microbial composition could be impacted by environmental factors including diet, age or genetic factors. Some studies have shown that vitamin D influenced the function and composition of bacterial communities in the gut in a protective way against dysbiosis and experimental IBD in mouse DSS model . Our prior observation indicates that Cyp27b1−/− mice with 1,25(OH)2D3 deficiency displayed severe colonic inflammation at the age of 8–10 month . In an attempt to examine the effect of 1,25(OH)2D3 deficiency on gut microbiota in the Cyp27b1−/− mice, we used 16S rRNA sequencing to dissect the composition of the gut microbiota and found gut dysbiosis in KO mice with a thinner mucus layer. Therefore, in this study we made an hypothesis that 1,25(OH)2D3 deficiency may influence gut homeostasis and induce the enrichment of some strains of bacteria such as A. muciniphila, Bacteroides-acidifaciens in the Cyp27b1−/− mice, thereby damaging the colonic mucus barrier that would allow a greater microbial access to the intestinal mucosa further promoting colonic inflammation.
Induction of colonic inflammation by 1,25(OH)2D3 deficiency
Effects of 1,25(OH)2D3 deficiency on gut microbiota
1,25(OH)2D3 deficiency causes degradation of the colonic mucus barrier
1,25(OH)2D3 affects A. muciniphila colonization in gut
A large body of evidences have established a strong link of low-level vitamin D to high risk of colon cancer and colonic inflammatory disease. Epidemiologic studies have shown that decreased vitamin D levels may influence the onset of IBD , increase clinical disease activity [7, 23, 24] and have a higher risk of malignant transformation [25, 26]. We and others also documented that in mice models, 1,25(OH)2D3 deficiency or VDR knockout was correlated with an increased risk of colitis and 1,25(OH)2D3 supplement ameliorated DSS-induced colitis [3, 9–11]. In the present study, 1,25(OH)2D3 supplement was able to rescue the inflammation occurred in Cyp27b1−/− mice (Fig. 4f). While the underlying mechanism is still unclear, accumulating evidences indicate that vitamin D play a preventive role in IBD development via regulating immune response, modulating the release of inflammatory cytokines [27, 28], improving intestinal epithelial barrier function by increasing the expression of some tight junction proteins such as Occludin, Zo-1, Zo-2, Vinculin and Claudins [29, 30], inducing colon cells senescence to secret senescence-associated inflammatory cytokines , and increasing antimicrobial peptide synthesis and secretion . Metagenomic studies have shown that vitamin D deficient diet or VDR knockout could impact the gut microbiome [20, 32].
Inflammatory bowel disease has been associated with dysbiotic microbiota due to a balance switch between commensal and pathogenic microorganisms [33–35]. For instance, the phylum Firmicutes is often less colonies in the feces of patients with Crohn’s disease [35, 36] whereas members of the Proteobacteria phylum such as Escherichia coli are commonly more abundant in patients with IBD as compared with healthy subjects [36, 37]. Bowdish and his colleagues found that alterations in age-related microbiota influenced intestinal permeability, caused age-associated inflammation, and decreased macrophage function . Microbiome genome-wide association studies have discovered that defects in many human genes involving IBD are associated with an aberrant composition of the gut microbiome . For example, knockout of Nod2 in mice predisposed them to colitis with lower levels of antimicrobial defensins and a higher bacterial load as compared with the control mice . In the present study, we compared the microbiome composition between 1,25(OH)2D3 deficient Cyp27b1−/− mice and WT mice via 16S rRNA sequencing. Our results demonstrated that the microbiomes established in WT and Cyp27b1−/− were distinct (Fig. 2a), suggesting that 1,25(OH)2D3 did modulate the composition of the gut microbiota. While these associations are well fit with the roles of the gut microbiota in IBD pathogenesis, the exact mechanism underlying dysbiosis remains to be fully elucidated.
A mucus layer in the gut tract is generally considered as a protective barrier against pathogenic micro-organisms and various chemical, enzymic or physical damage. Mucus produced by goblet cells is a viscous gel that mainly consists of high-molecular-mass glycoproteins, named as mucins . During evolution some mucolytic bacterial species may gain the capacity of utilizing this nutrient source . Therefore, the integrity of the mucus layer is leveraged between degradation by gut bacteria and replenishment by goblet cells. The Gram-negative A. muciniphila is a strictly anaerobic bacterium and abundant in the human gut with the capability of degrading mucin . Seregin and his colleagues found that NLRP6, which is a member of Nod-like receptor (NLR) family and are involved in the formation of inflammasomes , its deficiency can increase the susceptibility to DSS-induced colitis  and induced the enrichment of Akkermansia muciniphila that could function as a pathobiont by promoting colitis in a genetically-susceptible host . In contrast, Lemire et al.  and Mamantopoulos et al.  found that NLRP6 did not significantly influence the intestinal microbiota at homeostasis. These differences may be resulted from several factors including the mouse lineages (NLRP6 conditional knock-out versus NLRP6 conventional knock-out) and location of mouse facilities. 1,25(OH)2D3 has been reported to be involved in the inflammasome , whether 1,25(OH)2D3 has a function on NLRP6 is worthy of further investigation. It has also been reported that fiber-free dietary promoted enrichment of mucus-degrading bacteria including A. muciniphila and B. caccae . Consistently, our data showed that A. muciniphila was significantly enriched in Cyp27b1−/− mice as compared to WT mice (Fig. 2b, c), and supplement of 1,25(OH)2D3 could reduce its enrichment (Fig. 4d). This indicated that 1,25(OH)2D3 could limit the colonization of A. muciniphila. However, vitamin D deficient high fat diet has been shown to decrease the abundance of A. muciniphila in ileum . Such discrepancy might be due to the different mouse model and location site of A. muciniphila. In our study, Cyp27b1−/− mice showed the long-time status of 1,25(OH)2D3 deficiency while 1,25(OH)2D3 deficient diet indicated the short-time 1,25(OH)2D3 deficiency, which might result in the different effects on gut microbiota. The role of A. muciniphila in colitis is not very clear. Some studies showed that it could promote colitis. For example, one study found that occurrence of colitis was substantially increased in SPF IL10−/− mice administered with repeated oral gavage of A. muciniphila . In the presence of A. muciniphila, Salmonella-induced colitis was worsen and ulcerative colitis patients was accompanied by active pouchitis and the IBD patients presented with treatment failure [50–52]. The mechanism underlying A. muciniphila-promoted colitis might be due to the degradation of the mucus layer that allows a greater microbial access to the gut mucosa. However, some studies showed that colitis was associated with a reduction in Akkermansia muciniphila in IBD patients [53, 54]. Therefore, a large scale of studies is needed to confirm the clinical relation of colitis and A. muciniphila. In fact, we found a thinner mucus layer in Cyp27b1−/− mice with alterations in bacterial species such as higher amount of A. muciniphila (Fig. 3a) and an increase of total bacterial translocation (Fig. 1c) leading to the inflammation (Fig. 1d). Our results also showed no significant changes in the number of goblets and the compositions of mucins such as Muc1 and Muc3 between WT and Cyp27b1−/− mice (Fig. 3b, c). Since the proliferation of goblet cells and the expression of mucin genes were not significantly altered, it is reasonable to conceive that thinner mucus layer in Cyp27b1−/− mice may result from faster degradation of mucus layer due to the enrichment of mucin-degraded A. muciniphila in the gut rather than a reduction of mucin production itself. We further found that 1,25(OH)2D3 supplement reversed the amount of A. muciniphila, recovered the mucus layer and relieved the colonic inflammation (Fig. 4d–f). These findings indicate that 1,25(OH)2D3 could limit the colonization of A. muciniphila in gut. We and others have shown that 1,25(OH)2D3 is an important regulator of immune systems that could elicit Th2 immune responses and decrease pro-inflammatory cytokines such as IL-1, IL-6, IL-8, IFNγ and TNFα . 1,25(OH)2D3 could also increase Tregs, downregulate T cell-driven IgG production, inhibit DC differentiation, and enhance protective innate immune responses . Moreover, it has also been reported that 1,25(OH)2D3 promotes the production of anti-microbial peptides (AMPs), including β-defensins and cathelicidin [56, 57]. Although the mechanism was unclear, we speculated that 1,25(OH)2D3-reduced colonization of A. muciniphila in gut might result from activation of immune response by 1,25(OH)2D3 or antimicrobial peptide induced by 1,25(OH)2D3. In order to exclude the influence of age on A. muciniphila, we checked the colon phenotype and A. muciniphila abundance between the young Cyp27b1−/− and WT mice of 10–12 weeks. Our data demonstrated that even in young mice, 1,25(OH)2D3 deficiency led to a higher A. muciniphila abundance in fecal sample (Fig. 4a) and increased the translocation of bacterial to MLNs and thinner mucus layer in Cyp27b1−/− mice (Fig. 4b, c). It may be noteworthy that the inflammation was not significant in Cyp27b1−/− mice (data not shown), which was in concert with our prior study showing that 1,25(OH)2D3 deficiency could induce colon inflammation with aging . Our present study suggests that 1,25(OH)2D3 deficiency-induced higher A. muciniphila location in gut was gene associated but not age-related.
The present study demonstrated that 1,25(OH)2D3 deficiency impacted gut homeostasis including an increased enrichment of A. muciniphila in Cyp27b1−/− mice that might degrade the mucus layer thus allowing a greater microbial access to the intestinal mucosa and promoting colonic inflammation. The effect of 1,25(OH)2D3 on limiting the colonization of A. muciniphila was genetic-associated but not age-associated. Thus, the observations obtained from this study may disclose a potential new mechanism of how 1,25(OH)2D3 protects against colitis.
Generation of Cyp27b1−/− (KO) mice and the confirmation of their genotypes were described previously . Wild-type (WT) littermates served as the controls. Animals were maintained under pathogen-free conditions on a 12-h light/12-h dark cycle. 10–12 weeks or 8–10 months of male Cyp27b1−/− and WT littermates were used in this study. After weaning, they were fed with rescue diet (TD96348 Teklad, Madison, WI) formulated with 1.25% phosphorus, 2% calcium and 20% lactose or injected subcutaneously with 1,25(OH)2D3 at the dose of 1 μg/kg (KO + VD) until 10–12 weeks or 8–10 months old. It was confirmed that in the Cyp27b1−/− mice serum phosphorus and calcium levels were normalized and the littermates fed with the rescue diet .
Assessment of colon inflammation
After euthanasia, full length of colon was taken out and washed in PBS to remove fecal matter and then opened longitudinally, and jelly-rolled for formalin fixation and paraffin embedding. Histological assessment of H&E sections was performed in a blinded fashion by a pathologist using a scoring system as previously described . Briefly, each 100 × microscopic field along the length of the colon was scored separately for the presence and severity of inflammatory cell infiltration, hyperplasia, or epithelial damage. A weighted average percent for each lesion was calculated by the equation: [(1 × # of fields with score = 1) + (2 × # of fields with score = 2) + (3 × # of fields with score = 3)]/3 × total # of fields. Colon excluding the cecum was weighed after removal of feces normalized by its length (cm).
Extraction of bacterial DNA and 16S rRNA sequence analyses
DNA was extracted from fecal samples and 16S rRNA analysis was performed. Total genomic DNA from feces was isolated by CTAB/SDS method. Amplification of the V4 region of the 16S rRNA gene was performed by PCR with Phusion® High-Fidelity PCR Master Mix (New England Biolabs, Ipswich, MA) using custom barcoded primers (16S V4:515F-806R). Sequencing libraries were constructed according to the manufacturer’s recommendations on TruSeq® DNA PCR-Free Sample Preparation Kit (Illumina, San Diego, CA) and index codes were added. The library quality was evaluated on the Qubit@ 2.0 Fluorometer (Thermo Scientific, Waltham, MA) and Agilent Bioanalyzer 2100 system. Finally, an Illumina HiSeq 2500 platform was used to sequence the library with 250 bp paired-end reads generated. Analysis of sequences was done through Uparse software (Uparse v7.0.1001,http://drive5.com/uparse/) . Sequences with the similarity greater than 97% were assigned to the same OTUs and representative sequence for each OTU was screened for further annotation. The GreenGene Database (http://greengenes.lbl.gov/cgi-bin/nph-index.cgi)  was employed for analysis of each representative sequence based on RDP classifier (Version2.2, http://sourceforge.net/projects/rdp-classifier/) algorithm to annotate taxonomic information. A standard of sequence number corresponding to the sample with the least sequences was used to normalize OTUs abundance information. Based on this output normalized data, subsequent analyses of both alpha and beta diversity were performed.
Bacterial DNA was extracted from fecal samples and its concentration was measured by Nanodrop (Thermo Scientific, Waltham, MA). Total 20 ng DNA was input for qPCR using the SYBR Green reagents (Takara Bio, Shiga, Japan) on an ABI 7300 sequence detector (Applied Biosystems, Foster City, CA). Relative abundance of A. muciniphila in stool samples was normalized to the universal 16S rRNA gene EUB primers. Primer sequences are as follows: EUB-F: 5′-AGAGTTTGATCCTGGCTC-3′, EUB-R: 5′-TGCTGCCTCCCGTAGGAGT-3′; A. muciniphila-F: 5′-AGAGGTCTCAAGCGTTGTTCGGAA-3′ A. muciniphila-R: 5′-TTTCGCTCCCCTGG CCTTCGTGC-3′.
Total RNA was extracted from colon tissues with TRIzol reagent (Invitrogen, Grand Island, NY, USA) with the manufacturer’s protocol. 1 μg of RNA was reverse transcribed with the PrimeScript™ 1st Strand cDNA Synthesis Kit (Takara Bio, Shiga, Japan) according to the user’s manual. cDNA was used for real-time PCR analysis with gene-specific primers to determine the relative expression of genes of interest using SYBR green reagents (Takara Bio) in an ABI 7300 sequence detector (Applied Biosystems, Foster City, CA, USA). The forward and reverse primers used are listed as follows: 5′-TGGATTTGGACGCATTGGTC-3′ and 5′-TTTGCACTGGTACGTGTTGAT-3′ for GAPDH; 5′-CGGGAGGAGACGACTCTAAAT-3′ and 5′-CACGAACAGTTGTGAATCTGAGA-3′ for IL-1ɑ; 5′-GAAATGCCACCTTTTGACAGTG-3′ and 5′-CTGGATGCTCTCATCAGGACA-3′ for IL-1β; 5′-CTGCAAGAGACTTCCATCCAG-3′ and 5′-AGTGGTATAGACAGGTCTGTTGG-3′ for IL-6; 5′-ATGTGGGGGACCAAACTTCTG-3′ and 5′-GGATGGCGACATGAAGCAG-3′ for HGF1-F; 5′-TTAAAGACAGGCACTTTTGGCG-3′ and 5′-CCCTCGTATAGCCCAGAACT-3′ for MMP3. The respective forward and reverse primers were used to detect the relative expression levels of the target genes as fold changes by the 2−△△ct method. The relative amount of target mRNA was normalized to GAPDH.
Assessment of bacterial translocation
Total DNA was isolated from mesenteric lymph nodes (MLNs) and the bacterial load was measured using qPCR analysis of the universal 16S rRNA gene EUB primers in 20 ng DNA.
Alcian blue staining, goblet cell count and mucus thickness
The tissue of colon for Alcian blue staining was fixed in Carnoy’s fixative solution (dry methanol: chloroform: glacial acetic in the ratio of 60:30:10) and embedded in paraffin following standard procedure and the paraffin-embedded tissues were then cut 5 μm thick for staining. Alcian blue staining was performed with Kit from Nanjing Jiancheng Bioengineering Institute in China in compliance with the manufacturer’s instructions. On each slide, 10 high-power fields (200× and 400× magnification) were selected randomly. Mucus layer thickness was measured according to the method previously described . Goblet cells were counted and averaged over five high power fields at 400× magnification.
Determination of butyrate/SCFA concentrations
Gas chromatography was used to analyze the lyophilized fecal samples. One gram of lyophilisate was dissolved in 5–10 volume of ddH2O and 1 ml supernatant was added to 0.2 ml crotonic acid/metaphosphoric acid and then centrifuged for 10 min at 12,000 rpm. Butyrate concentration in the supernatant was measured by using a GC-14B gas chromatograph (Shimadzu Deutschland GmbH, Duisburg, Germany) equipped with a flame ionization detection with a NUKOLTM capillary column (Supelco) 30 m × 0.32 mm × 0.25 μm. A combined standard solution containing acetic acid, propionic acid, isobutyric acid, butyrate, isovaleric acid, valeric acid and crotonic acid to identify the presence of butyrate. Butyrate concentration was determined by the formula: Butyrate (mM) = (Sample PA × Standard crotonic acid PA × Concentration of standard butyrate)/(Sample crotonic acid PA × Standard PA). PA: peak area.
Data are expressed as mean ± SEM. Statistical analyses were performed by SPSS 20.0 (Abbott Laboratories, Chicago, IL). ANOVA was employed to compare the difference between WT, KO and KO + VD.
Conceived and designed the experiments: XY. Performed the experiments: WZ, QZ. Analyzed the data: JY, CZ. Contributed reagents/materials/analysis tools: WZ, JY. Wrote the paper: XY. All authors read and approved the final manuscript.
We would like to thank Dr. Zan Huang’s lab from Nanjing Agricultural University for providing technical support in gas chromatography experiment.
The authors declare that they have no competing interests.
Availability of data
The datasets used and/or analysed during the current study are available from the corresponding author on reasonable request.
Consent for publication
Ethics approval and consent to participate
Experimental procedures and animal welfare were approved by the Institutional Animal Care and Use Committee of Nanjing Medical University (Approval ID 1601080).
This work was supported by National Natural Science Foundation of China (Grant No. 81572386).
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