Our previous study has shown that DCA accelerated the progression of the intestinal adenoma–adenocarcinoma sequence under the hereditary background of Apc mutation. 16 Here, we provide evidence that DCA‐mediated intestinal dysbiosis has a crucial role in promoting intestinal tumorigenesis and show that imbalanced gut microbiota impair the intestinal barrier function, modulate the cytokines and chemokines expressing in the tumor microenvironment and ultimately promote intestinal carcinogenesis via activation of Wnt signaling. Our study also provides new insight of potential treatment targeting gut microbiota for intestinal cancer.

The crosstalk between bile acids and gut microbiota has been established, both of which have been studied extensively in intestinal carcinogenesis. 15 DCA can be generated from cholic acid by 7α‐dehydroxylating bacteria in association with a high‐fat diet and is implicated in causation of CRC. 16 , 17 In turn, studies have shown cholic acid‐diet has significant impact upon the community structure of gut microbiota. 18 - 20 However, the mechanisms of how gut microbiota and DCA affect intestinal tumorigenesis remain to be clarified.

For many years, a high level of deoxycholic acid (DCA, a secondary bile acid) in the intestinal lumen, mostly induced by a high dietary fat intake, was proposed as a potential pro‐carcinogenic agent. 9 - 11 Persistent and repeated exposure of intestinal epithelium to abnormally high concentrations of DCA appears to induce DNA damage and genomic instability. 12 Several epidemiological studies have highlighted the correlation between high physiologic levels of DCA and incidence of CRC, 13 , 14 although the mechanisms remain elusive.

The human large intestine is a microbial ecosystem populated with dense microorganisms, and also a common site for developing adenocarcinomas. 6 , 7 Studies support the concept that disturbances in the composition and function of the intestinal microbiota contribute to intestinal tumor development, 8 and gut microbiome is not just a bystander along for the ride, it might be a prime suspect dictating colonic carcinogenesis. However, it is challenging to determine the causality between changes of gut microbiota and intestinal tumorigenesis.

Greater than 95% of colorectal cancers (CRCs) are sporadic and develop from adenomas over a number of years. 1 , 2 Apart from hereditable components, environmental factors strongly determine the progression of intestinal neoplastic transformation, known as the adenoma–adenocarcinoma sequence. 2 , 3 Adenomatous polyposis coli ( Apc ) gene mutation is often an early event in the adenoma–adenocarcinoma sequence (where 45–60% of sporadic adenomas and 60–80% of carcinomas harbor one inactivated allele). 4 The Apc gene is a tumor suppressor gene, which regulates the Wnt/β‐catenin signal pathway. The Apc min/+ mouse model carries a heterozygous mutation at codon 850 of the Apc gene, and can spontaneously develop multiple intestinal adenomas. 5

Fecal DNA was extracted from stool sample using the E.Z.N.A. Stool DNA Kit QIAamp DNA Stool Mini Kit (Omega Bio‐Tek, Norcross, GA) according to the manufacturer's guidelines. Barcoded universal bacterial primers targeting V3‐V4 hypervariable region of the 16S rRNA gene was PCR amplified in triplicate. The resulting amplicons were pooled and sequenced on the Illumina 2 × 300 bp MiSeq platform according to published protocols. 26 , 27 The optimizing sequences were mapped into operational taxonomic units (OTUs) and picked at 97% similarity in Mothur (version v.1.30.1.). 28 , 29 Community diversity was estimated by the Shannon diversity index and the Simpson concentration index. Taxonomic analysis was based on the OTUs sequences using QIME platform and estimated by RDP Classifier (version 2.2). 30 Similarities were shown by dendrogram among the samples. Principal component analysis (PCA) was carried out on the resulting matrix of distances between groups.

The tumor tissues lysates from the distal small intestine were solubilized using RIPA buffer supplemented with protease inhibitors (Solarbio, Beijing, China) and homogenized. Nuclear protein was extracted for β‐catenin analysis using the Nuclear Extraction Kit (Signosis, Inc., Sunnyvale, CA) according to the manufacturer's protocol. The membranes were probed with the primary antibodies β‐catenin and cyclin D1 followed by horseradish peroxidase‐conjugated secondary antibodies (Cell Signaling Technology, Inc.). Internal control protein β‐actin was used for total protein and Histone 2A for nuclear protein. Proteins were quantified by densitometry using an Imaging processor program (Image J).

Total RNA was extracted using the RNeasy mini kit (Qiagen, Carlsbad, CA), and cDNA reverse transcription was carried out using the TIANScript RT Kit (TIANGEN, Inc. Beijing, China) according to the manufacturer's instructions. The Oligonucleotide primers for target genes were shown in supplement materials (Table S1). The ΔΔCt method was used to calculate relative mRNA expression.

Immunofluorescent staining was performed on formalin‐fixed tissues to evaluate classic markers of M2 tumor‐associated macrophage (TAM) polarization. Specific antibodies against macrophage marker F4/80 (ab6640, Abcam, Cambridge, MA) and MR (ab64457, Abcam, Cambridge, MA) were used for immunofluorescent staining on sections overnight. The sections were washed three times with 1 × PBS for 5 min and incubated 60 min with fluorochrome‐conjugated secondary antibodies Alexa‐568 (F4/80, red), Alexa‐488 (MR, green) diluted to 2 μg/mL in PBS in the dark. Subsequently the specific secondary antibodies were applied. DAPI (4, 6‐diamidino‐2‐phenylindole) was lastly applied on the sections.

Formalin‐fixed tissues were dehydrated and embedded in paraffin according to standard H&E protocols. Intraepithelial neoplasia (dysplasia) is characterized by morphological changes that include altered architecture and abnormalities in cytology and differentiation. Low‐grade dysplasia (LGD) is confined to the lower half of the epithelium, and in high‐grade dysplasia (HGD), the abnormal cells occur in the upper half and exhibit a greater degree of atypia. And intramucosal carcinoma is diagnosed when the tumor invades into the lamina propria, but not through the muscularis mucosae. The histopathologic analysis was performed in a blinded manner by the same pathologist (YJZ). The tissue sections were incubated with primary antibodies, rabbit monoclonal anti‐Ki‐67 (ab16667, Abcam, Cambridge, MA), F4/80 (ab6640, Abcam, Cambridge, MA), MCP‐1 (ab9669, Abcam, Cambridge, MA) E‐cadherin (CST8437, Cell Signaling technology, Boston, MA), β‐catenin (Santa Cruz Biotechnology, Inc., Santa Cruz, CA), cyclin D1(ab134175, Abcam, Cambridge, MA). The biotinylated anti‐rabbit secondary antibody was applied followed by horseradish peroxidase (HRP)–streptavidin solution. Finally, the sections were counterstained with hematoxylin. Five random areas from a single section were checked for the percentage of positive cells. Data were quantified by calculating the average percentages of positive cells in each mouse as the positive rate of cells.

Mice were sacrificed for intestinal tumor burden assessment and tissue collection as previously described. 16 , 25 Tissue sections were prepared for hematoxylin and eosin (H&E) and immunohistochemical staining. Adenomas of distal small intestinal section were excised, immediately frozen in liquid nitrogen and then stored at −80°C until analysis for mRNA and protein expression.

During feces transfer experiments, fresh stool pellets from DCA‐treated and ‐untreated mice followed by 2‐weeks' washout of DCA were suspended in reduced PBS (0.1 g/1 mL), homogenized, centrifuged at 800 g for 5 min and the supernatant was collected. Three hundred μL of supernatant was transplanted to 4‐week‐old Apc min/+ mice, which were given a gavage of streptomycin (20 mg) daily for 3 days before FMT to deplete the native gut microbiota as previously described. 23 , 24 During transfer experiments, the mice were inoculated for a total of 16 times over the subsequent 8 weeks.

To exclude the potential role of residual fecal DCA in intestinal tumorigenesis, liquid chromatography‐mass spectrometry (LCMS) was used to detect the concentration of fecal DCA as described previously. 22 The DCA concentration in each group was calculated based on the peak areas.

FITC‐conjugated dextran dissolved in water (4,000 MW) was administered rectally to mice at 2 mg/10 g body weight. Whole blood was collected using heparinized microhematocrit capillary tubes via eye bleed 2 hr after FITC‐dextran administration. Fluorescence intensity in sera was analyzed using a plate reader. The concentration of FITC‐dextran in serum was determined by comparison to the FITC‐dextran standard curve.

To further determine the crucial role of the intestinal dysbiosis in tumorigenesis, another group of Apc min/+ mice were administrated with a cocktail of antibiotics for depletion of gut microbiota. A mixture of 500 mg of ampicillin, 250 mg of vancomycin, 500 mg neomycin and 250 mg of metronidazole (Sigma‐Aldrich, St. Louis, MO) were supplemented by gavage for 12 weeks, in accordance to published studies. 21

Four‐week Apc min/+ mice on a C57BL/6J background were purchased from Jackson laboratory. The mice were provide with either sterile water or 0.2% DCA (C97% titration, Sigma‐Aldrich, St. Louis, MO) in drinking water under specific pathogen free (SPF) conditions for 12 weeks, as previously described. 16 Signs of illness were monitored daily and body weight was recorded weekly. Animal protocols were approved by the Institutional Animal Care and Use Committee at Tianjin Medical University, Tianjin, P. R. China.

To further determine the crucial role of the distinct shift of intestinal microbiota in tumorigenesis, another group of Apc min/+ mice were administered with a cocktail of antibiotics that would deplete their gut microbiota 38 , 39 (Fig. S2 a ). Antibiotic supplementation significantly reduced the DCA‐induced intestinal tumors. Concordantly, the proportion of more advanced tumors was largely decreased (Fig. S2b and S2c). Moreover, DCA‐induced intestinal low grade inflammation was alleviated by antibiotics (Fig. S2d). Then we draw a conclusion that antibiotics could abolish DCA‐induced tumorigenesis, further demonstrating that alteration of the microbial community induced by DCA played an important role in the intestinal carcinogenesis.

Deoxycholic acid (DCA)‐induced dysbiosis activated tumor‐associated signaling pathway. ( a 1 and a 2) Immunohistochemistry results showed that signaling molecular related to Wnt/β‐catenin pathway (E‐cadherin, β‐catenin and cyclin D1) were enhanced in Apc min/+ mice that received fecal microbiota from DCA‐treated mice. ( b 1) Intestinal protein levels of β‐catenin and cyclin D1 in mice with fecal microbiota transplantation from DCA‐treated or ‐untreated donors were analyzed by Western blot, using internal control protein β‐actin for total protein and Histone 2A for nuclear protein. ( b 2) Proteins were quantified by densitometry using an Imaging processor program (Image J). WD‐FMT group, transfer of fecal microbiota from DCA‐treated donors to wild‐type C57BL/6 J mice. FMT‐C group, transfer of fecal microbiota from DCA‐untreated donors to Apc min/+ mice. FMT‐D group, transfer of fecal microbiota from DCA‐treated donors to Apc min/+ mice. Scale bar: 50 μm. *, P < 0.05, **, P < 0.01, ***, P < 0.001.

Growing evidence suggests that some specific bacteria can activate the Wnt/β‐catenin pathway during intestinal tumorigenesis. 37 Loss of epithelial cell markers (such as E‐cadherin) is associated with the more advanced grade of neoplasia. As shown in Figures 6 a and 6 b, the expressions of E‐cadherin was downregulated, and Wnt signaling molecules (including β‐catenin and cyclin‐D1) ware activated in the group of FMT‐D mice. These data indicated that transfer of feces from DCA‐treated donors activated the Wnt/β‐catenin pathway.

F4/80, a widely‐used marker of murine macrophage populations, was also detected in neoplastic tissues and non‐neoplastic tissues. Consistently, fecal transplantation from DCA‐fed mice significantly increased the total number of macrophages as compared to FMT‐C and WD‐FMT groups (Fig. 5 B ). Recently, studies have indicated that TAMs can switch their phenotype in response to the microenvironment, and M2 TAMs play a predominant role in tumor development. 35 , 36 Therefore, we sought to investigate the M2 TAMs in Apc min/+ mice after FMT. The mRNA levels of M2 genes arginase 1 (Arg‐1) and mannose receptor (MR) were markedly upregulated in neoplastic tissues of group FMT‐D (Fig. 5 b 3). Consistently, immunofluorescence showed that a number of F4/80‐ and MR‐positive macrophages were highly infiltrated within the neoplastic stroma (Fig. 5 c ).

Deoxycholic acid (DCA)‐induced dysbiosis promoted M2 phenotype tumor‐associated macrophages (TAMs) polarization during the progress of intestinal tumor development. ( a 1 and a 2) Immunohistochemistry analysis of intestinal tissues showed the increased chemoattractant cytokine (MCP‐1) secretion in Apc min/+ mice that received fecal microbiota from DCA‐treated mice (FMT‐D group). (a3) Real‐time‐PCR results showed that MCP‐1 gene was up‐regulated in FMT‐D group. ( b 1 and b 2) Macrophages recruitment (F4/80) in the small intestine was assessed using immunostaining. (b3) The mRNA level of F4/80 and M2 TAMs (Arg‐1 and MR) were increased in FMT‐D group. ( c ) Immunofluorescence assay indicated the transfer of fecal microbiota from DCA‐treated donors promoted polarization of M2 TAMs. Paraffin sections of small intestinal tissues were immunofluorescence stained with primary antibodies against F4/80 to mark total macrophages (red) and MR to label M2 TAMs (green), and counterstained with DAPI to mark nuclei (blue). In (a3) and (b3), the mRNA expression level of FMT‐D group was compared to that of FMT‐C group. WD‐FMT group, transfer of fecal microbiota from DCA‐treated donors to wild‐type C57BL/6 J mice. FMT‐C group, transfer of fecal microbiota from DCA‐untreated donors to Apc min/+ mice. FMT‐D group, transfer of fecal microbiota from DCA‐treated donors to Apc min/+ mice. Scale bar: 50 μm.*, P < 0.05, **, P < 0.01, ***, P < 0.001.

Monocyte chemoattractant protein‐1 (MCP‐1) is one of the critical chemoattractant cytokines that recruit monocytes/macrophages to local inflammatory sites. 34 We next sought to determine whether the monomacrophage system was activated in the progression of tumor development after microbiota transfer from cancerous donors. Immunohistochemistry showed that the expression of MCP‐1 was significantly higher both in neoplastic tissues and non‐neoplastic tissues from the FMT‐D group, compared to those in the FMT‐C group. Similarly, the highest level of MCP‐1 mRNA was showed in FMT‐D group (Fig. 5 a ).

Inflammation is a hallmark of cancer, which contributes to the development of approximately 15% of malignancies. 32 We found that transfer of feces from DCA‐treated donors induced low grade inflammation of the intestine. Gene expression of inflammatory mediators was examined in non‐neoplastic tissue of the three groups as shown above. All data were normalized to the fold‐change from the group of wild‐type C57BL/6J mice which received feces from the DCA‐treated donors (WD‐FMT). Gene expression of IL‐1β, IL‐6, TNF‐α and IL‐23p19 were significantly higher in Apc min/+ mice that received feces from DCA‐treated mice than in those that did not (Fig. 1 f ). Moreover, IFN‐γ has anti‐cancer functions including cell cycle restriction, immune surveillance and pro‐apoptotic functions, 33 and the FMT‐D group exhibited a significantly lower expression of IFN‐γ compared to the FMT‐C mice. There was no significant change in the expression of TGF‐β among the three groups (Fig. 1 f ).

Deoxycholic acid (DCA)‐induced dysbiosis could be transmitted to Apc min/+ mice. ( a ) Principal component analysis (PCA) of gut microbiota. The PCA analysis focused on fecal bacterial communities in distinct groups with respect to normal diet or DCA treatment after fecal microbiota transplantation using principal components. The spatial distance showed the similarity degree of bacterial taxon between samples [The closer, the more similar (PC1 and PC2)]. DCA ( n = 3): red dots; transfer of feces from DCA‐untreated donors to Apc min/+ mice (FMT‐C, n = 4): blue squares; transfer of feces from DCA‐treated donors to Apc min/+ mice (FMT‐D, n = 6): green rhombus. ( b ) The phylum level of the FMT‐C and FMT‐D group after the Experiment (8 weeks). ( c and d ) Heatmap results showed bacteria at important genus and species levels that were significantly different in abundance between FMT‐C and FMT‐D group after the Experiment (8 weeks) Different color showed the relative abundance of the community (from cold to warm color means from low to high abundance).

PCA results showed that the luminal microbial community after receiving the fecal microbiota from DCA‐treated donors was significantly different from those receiving microbiota from DCA‐untreated mice, but with no difference from DCA‐treated Apc min/+ mice. PCA disclosed that the major alteration in the microbial community was promoted by distinct treatments as shown in Figure. 4 a . Further analysis of microbial phylum indicated that the proportion of Firmicutes was significantly decreased, whereas Bacteroidetes were increased in FMT‐D group (Fig. 4 b ). Moreover, analysis at the genus level showed that the relative abundance of Alistipes , Helicobacter , Desulfovibrio , Turibacter , Parabacteroides and Escherichia‐Shigella were significantly higher, while Roseburia , Bifidobacterium , Lactobacillus and Ruminococcus were lower in the FMT‐D group compared to the FMT‐C group (Fig. 4 c ). Furthermore, the transfer of feces from DCA‐treated mice markedly altered the microbiota community at the species level by reducing butyrate‐producing microbiota ( Clostridium‐leptum , Eubacterium‐corprostanoligenes , Ruminococcus‐ flavefaciens and Lachnospiraceae‐bacterium‐COE1 ), as shown in Figure 4 d .

Importantly, tumorigenesis was transmitted to Apc min/+ mice after the FMT from DCA‐treated donors, but not to C57BL/6J mice. Feces from DCA‐treated donors significantly increased the total number of intestinal tumors of Apc min/+ mice by 103.4% compared to that from DCA‐untreated donors (33.00 ± 6.60 vs . 16.20 ± 9.47, P < 0.01, Fig. 3 c ), and the numbers of tumors with large size (≥ 2 cm) were also increased (9.3 ± 4.0 vs . 1.8 ± 2.2, P < 0.01). Fecal microbiota from DCA‐treated donors promoted intestinal tumorigenesis mainly in the proximal portion of the small intestine (7.83 ± 2.14 vs . 3.40 ± 1.34, P <0.01). Histological analysis showed that Apc min/+ mice with FMT from DCA‐treated donors led to further intestinal carcinogenesis (HGD including intramucosal cancer) in 75% of Apc min/+ mice, whereas none of the Apc min/+ mice with FMT from DCA‐untreated donors had further intestinal carcinogenesis. No invasive carcinoma invading into the lamina propria was found in any group in our study (Figs. 3 d and 3 e ).

Deoxycholic acid (DCA)‐induced dysbiosis promoted intestinal adenoma–adenocarcinoma sequence and induced low grade inflammation. ( a ) The experimental scheme showed the timing of fecal microbiota transplantation (FMT). Streptomycin (20 mg) was gavaged daily for 3 days before FMT. The mice were inoculated fecal microbiota for a total of 16 times over the subsequent 8 weeks. ( b ) To exclude the role of fecal DCA in intestinal tumorigenesis, liquid chromatography‐mass spectrometry (LCMS) was used to analyze the fecal DCA concentrations from DCA group (before and after wash out period) and control group. After 2 weeks' washout, there was no significant difference of fecal DCA concentration between the DCA‐treated and ‐untreated groups. ( c ) The total number of intestinal tumors from Apc min/+ mice with FMT from DCA‐treated (FMT‐D) or ‐untreated donors (FMT‐C). ( d , e ) The representative and histologic appearance of intestinal tumors from Apc min/+ mice with FMT from DCA‐treated or ‐untreated donors. ( f ) Real‐time‐PCR of the small intestine tumors showed the host immune response was activated in Apc min/+ recipients that received fecal transplantation from DCA‐treated mice. The mRNA expression levels of FMT‐C and FMT‐D mice were compared to that of WD‐FMT mice. WD‐FMT, transfer of fecal microbiota from DCA‐treated donors to wild‐type C57BL/6 J mice. FMT‐C, transfer of fecal microbiota from DCA‐untreated donors to Apc min/+ mice. FMT‐D, transfer of fecal microbiota from DCA‐treated donors to Apc min/+ mice. Scale bar: 50 μm. *, P < 0.05, **, P < 0.01, ***, P < 0.001.

To exclude the role of residual fecal DCA before FMT, experiments were performed shown in Figure 3 a , and the concentration of DCA in feces was detected. It showed that after 2 weeks' washout, there was no significant difference of fecal DCA concentration between the DCA‐treated and ‐untreated groups (Fig. 3 b ). Hence, the microbiota of fresh fecal samples from DCA‐treated or ‐untreated donor mice were transplanted to recipient Apc min/+ or wild‐type C57BL/6J mice: transfer of feces from DCA‐treated donors to Apc min/+ mice (FMT‐D group, n = 12); transfer of feces from DCA‐untreated donors to Apc min/+ mice (FMT‐C group, n = 8); transfer of fecal microbiota from DCA‐treated donors to wild‐type C57BL/6 J mice (WD‐FMT group, n = 6).

Recent evidence has demonstrated that intestinal dysbiosis could affect mucus barrier biology. 31 Disrupted barrier function, as evidenced by increased FITC‐dextran in the serum, was found in the DCA group (Fig. S1 a ). Besides, in accordance with the decreased colonic mRNA expression of mucin (MUC2, the prominent component in the gut secreted from goblet cells), and periodic acid schiff (PAS) staining showed that DCA attenuated the number of goblet cells, which play a vital role in maintaining the thick layer of mucus to defend against the pathobionts. Moreover, on exposure to imbalanced gut microbiota, intestinal low grade inflammation was induced. Inflammation cytokines including IL‐1β, IL‐6 and TNF‐α were upregulated in the intestine of DCA‐treated mice (Fig. S1 b ).

Deoxycholic acid (DCA) aggravated the intestinal dysbiosis during adenoma–adenocarcinoma development. ( a ) 16S rRNA sequence analysis showed the phylum level of the DCA‐treated (DCA) and ‐untreated (Control) before and after the Experiment (0 week and 12 weeks). Heatmap results of 16S rRNA sequence analysis showed that bacteria at important genus ( b ) and species ( c ) levels were significantly different in abundance between DCA‐treated and ‐untreated Apc min/+ mice at 0 week and 12 weeks. Control, n = 6; DCA, n = 6.

Diversity and concentration estimators (Shannon and Simpson) showed that DCA supplementation significantly increased the value of the Simpson concentration index and decreased the value of the Shannon diversity index, suggesting that DCA altered diversity in the microbiota. Richness estimators (Ace and Chao) showed no significant difference between the Control group (Con, n = 6) and the DCA group (DCA, n = 6). The microbiota community structure for each group at the phylum level is shown in Figure 2 a . Between the two groups, there were four dominant phyla: Firmicutes , Bacteroidetes , Proteobacteria and Verrucomicrobia . The proportions of the Firmicutes phylum were 25.1% in DCA group, and 33.7% in Control group. For Bacteroidetes phylum, the proportions were 71.9% in DCA group, and 62.2% in Control group. A heatmap of the microbial compositions at the genera level was generated with 8 significantly different genera between Week 0 and Week 12 for the two groups. The levels of opportunistic pathogens, including Ruminococcus , Escherichia‐Shigella , Desulfovibrio , Dorea significantly increased during intestinal tumorigenesis due to DCA. Lactobacillus , Lactococcus , Roseburia were relatively less abundant in mice in the DCA‐supplemented group compared to mice in Control group (Fig. 2 b ). Moreover, there were significantly higher levels of Clostridium and Escherichia‐Shigella in the microbiota of the DCA supplemented group of mice compared to those in the Control group. At species level, the abundance of Lactobacillus_gasseri and mostly butyrate‐producing bacteria, such as Clostridium leptum Lachnospiraceae bacterium and Eubaterium coprostanoligenes were significantly lower during the progression of intestinal tumor development (Fig. 2 c ).

Deoxycholic acid (DCA) accelerated intestinal adenoma–adenocarcinoma sequence. ( a ) The experimental flow showed when DCA was administered for 12 weeks until sacrifice. ( b ) The total number of intestinal tumors. ( c ) Histological scores of the tumors in distal small intestine. Each point referred to one animal, and lines indicated the mean level. ( d , e ) The representative and histologic appearance of intestinal tumors from DCA‐treated or ‐untreated Apc min/+ mice. Control, n = 9; DCA, n = 10. ( f 1 and f 2) Immunohistochemistry results showed that the cell proliferation (Ki‐67) was significantly increased in Apc min/+ mice by DCA. ND, normal diet. DCA, deoxycholate acid. Scale bar: 50 μm. *, P < 0.05, **, P < 0.01, ***, P < 0.001.

No differences were found in the dietary intake and body weight between the Control and DCA group. Supplementation with 0.2% DCA in drinking water for 12 weeks led to intestinal tumor progression, and the total tumor number in DCA‐supplemented mice was almost twofold greater than in the mice without DCA treatment (Figs. 1 a and 1 b ). HGD, including intramucosal carcinomas, were detected in 80% (8/10) of Apc min/+ mice supplemented with DCA, whereas only 33% (3/9) of age‐matched Apc min/+ mice, on a normal diet developed adenomas with LGD, and adenomas without dysplasia were found in the remained 67% mice (Figs 1 c–e ). It indicated that DCA‐treated mice developed tumors along the carcinogenic sequence starting from adenomas, and with LGD to HGD. Moreover, cell proliferation significantly increased in DCA‐treated mice (Fig. 1 f ).

Discussion

Substantial evidence suggests that there is a reciprocal interaction between bile acid metabolisms and intestinal microbiota.15, 17 Our study combined genetic predisposition, bile acid and gut microbiota to better understand the interactions between host and environmental factors in the etiology of CRC. Here, we showed that DCA‐induced dysbiosis during intestinal carcinogenesis in conjunction with Apc gene mutation, as evidenced by the increased abundance of pathogens and decreased in probiotics, and this structure shift of microbial community was sufficient to transmit disease independent of DCA treatment. Furthermore, DCA‐induced dysbiosis also impaired gut barrier function and induced the inflammation, and mononuclear phagocyte recruitment, M2 phenotype TAMs polarization in tumor microenvironment, and Wnt signaling activation.

Despite only a minority of tumors located in the colon in Apcmin/+ mice, which is the limitation of this mouse model, it offers genetic advantages mimicking human intestinal adenoma–adenocarcinoma sequence.40 More importantly, a previous study has reported that wild‐type mice with a 0.2% DCA supplemented diet developed colon cancers in 45% of mice after 10 months.41 And the level of DCA in feces from DCA diet mice was significantly higher than that from control diet mice (4.6 vs. 0.3 mg/g dry weight).9 Similarly, in our study, the level of fecal DCA in the DCA‐treated mice was much higher than the Control group. However, our previous study did not determine whether there was any shift of the gut microbiota. Important features of our study, on the other hand, are that DCA changed the microbial community to accelerate dysbiosis, as well as causing an impaired intestinal barrier function during intestinal adenoma–adenocarcinoma sequence. Interestingly, the composition shift of the gut microbiota observed in DCA‐treated Apcmin/+ mice was similar to that seen in individuals with a western diet.42, 43

The antimicrobial activity of bile acids was first described in the last century,44 although the action of bile acids on typical intestinal bacteria has often been examined in vitro.45, 46 Previous studies have shown the mechanisms underlying the bactericidal action of bile acids through membrane damage.44, 47 The bactericidal activity of bile acids corresponds to its hydrophobicity, which increases its affinity for the phospholipid bilayer of the bacterial cell membrane.48 Among bile acids, DCA is most toxic, for example, inclusion of ∼1 mmol/L DCA in growth medium severely can inhibit the growth of many intestinal bacteria, including Clostridium perfringens, Bacteroides fragilis, lactobacilli and bifidobacteria.46 Therefore, it is reasonable to propose that DCA may participate in alteration of the composition of the gut microbiota. However, more studies are needed to elucidate how DCA modifies the composition of the gut microbiota during intestinal tumorigenesis.

To determine whether the gut microbiota can be carcinogenic, we used FMT for our subsequent investigation. Previous work showed that gut dysbiosis is the major factor for triggering low grade inflammation,49 known to promote tumor progression.50, 51 We found that the transfer of feces from DCA‐treated mice also induced intestinal low grade inflammation. In fact, all the proinflammatory cytokines were positively correlated with the increased tumor burden, suggesting that DCA‐induced dysbiosis mediated gut inflammation and could be a substantial factor in carcinogenesis.

Innate immune cells are highly represented in the tumor microenvironment, with macrophages being the most abundant. MCP‐1 expression produced by tumor cells is significantly correlated with the recruitment and regulation of TAMs.34 In our study, we found that DCA‐induced dysbiosis promoted the expression of MCP‐1 and infiltration of macrophages in the gut. Moreover, studies indicated that M2 phenotypic TAMs predominantly occur during the advanced stage of tumor progression and produce growth‐stimulating molecules that promote tumor growth.36, 52 Hence, dysbiosis‐mediated recruitment of M2 phenotype TAMs might be another substantial element conducive to tumor development by promoting cancer cell proliferation.

Dysregulation of Wnt signaling has been reported to play a major role in CRC development and progression. And enhanced nuclear β‐catenin in the setting of Apc mutation results in intestinal epithelial cell proliferation.53 A tumor suppressor of E‐cadherin, which is present in 75% of the tumor tissue and 100% of the normal mucosa, can function through β‐catenin.38 Previous studies have observed a correlation between gut bacteria, such as Fusobacterium nucleatum (Fn), and E‐cadherin/β‐catenin signaling.37, 53 FadA adhesin, which is required for Fn to attach E‐cadherin on epithelial cells and activate β‐catenin signaling, ultimately leads to increased expression of Wnt pathway genes.37 Consistent with this, our study showed that DCA‐mediated‐dysbiosis induced reduction of E‐cadherin, and augmented nuclear β‐catenin expression with induction of the downstream Wnt signaling molecules.

Collectively, our results pointed to a major role of DCA‐induced dysbiosis in the intestinal adenoma–adenocarcinoma sequence. We demonstrated that, in mutationally predisposed hosts, DCA‐induced dysbiosis might disrupt intestinal barrier function, promote the recruitment of tumor‐associated macrophages and polarization of M2 macrophages, and further accelerate the intestinal adenoma–adenocarcinoma sequence via activation of the Wnt/β‐catenin signaling pathway. These results highlight a predominant role of intestinal dysbiosis mediated by DCA‐supplementation in intestinal tumor development. Hence, personalized dietary interventions might modulate the gut microbiota to promote health in individuals with a high genetic risk of CRC.