Mice became symptomatic, showing symptoms such as weight loss, loose stool, and bloody diarrhea by day 2–3 of intrarectal TNBS administration. The symptoms were significantly improved by F. prausnitzii and its supernatant. Compared with the control group, the mice in the TNBS model group exhibited a markedly shorter colon length. The gross observation of the bowel revealed that the intestines of the mice in the EtOH group were pink, smooth and intact, while the gut lumen of the mice from the TNBS group showed swelling, congestion, a thinner colon wall, disappearance of mucosal folds, and a shortened colon. In some segments, large amounts of loose, unshaped or bloody feces were deposited in the gut lumen. After longitudinal incision, obvious ulcers were observed, and a few of the specimens exhibited intestinal adhesions and intestinal obstruction. Most of the changes observed in the gross observation of the gut were mitigated by F. prausnitzii and its supernatant to some extent. Histological examination also showed that F. prausnitzii and its supernatant could also ameliorate mucosal inflammation and histopathologic damage compared with the TNBS model group. Compared with the TNBS model group, the serum TNF-α concentrations in F. prausnitzii (32.51 ± 16.41 vs. 96.55 ± 16.06) and its supernatant (39.27 ± 16.70 vs. 96.55 ± 16.06) were also significantly decreased. (Fig. 1)
F. prausnitzii and its supernatant increase gut microbiota diversity
Mounting evidence shows that gut microbiota dysbiosis plays a significant role in the occurrence and development of IBD. To further study the effects of F. prausnitzii and its supernatant on the gut microbiota in IBD, we used 16S sequencing to further clarify the differences in the gut microbiome. Our findings indicated that the microbial diversity in the TNBS model group was significantly different from that in the EtOH control group, which was generally accompanied by a decrease in the diversity (Shannon index and species rank abundance curve) of gut microbiota. Treatment with both F. prausnitzii and its supernatant increased the diversity of the gut microbiota, especially in the supernatant group (P < 0.05). Based on the analysis of the number of OTUs, the 5 groups shared 597 OTUs, 23 of which were unique to the EtOH control group, while 27 were unique to the TNBS model group, and 22 were unique to the TNBS + ASA group. There were 15 and 21 unique OTUs in F. prausnitzii and its supernatant, respectively. (Fig. 2)
F. prausnitzii and its supernatant restore gut microbiota dysbiosis and increase the abundance of SCFA- producing bacteria
We further explored the changes in the gut microbiota composition, as shown by the relative abundance (Fig. 3), OTU level (Fig. 4) and LefSc biomarkers (Fig. 5). Compared with the TNBS model group, treatment with both F. prausnitzii and its supernatant increased the abundance of Firmicutes, Cyanobacteria and Clostridiales, and decreased the abundance of Proteinbacteria, Acidobacteria, and Bacteroidetes. At the genus level, F. prausnitzii increased the abundance of Butyricicoccus, Roseburia, Ruminiclostridium, Lachnospiraceae, Oscillibacter, Anaerotruncus, Ruminococcaceae, Alistipes, Rikenella, Allspirophore, Deisulfoprecotella, and Mucispirllum, which was more obvious in the F. prausnitzii supernatant group than in the TNBS model group. Notably, the Butyricicoccus, Roseburia, Lachnospiraceae, Ruminococcaceae, Rikenella, and Eubacterium_xylanophilum are short-chain fatty acid (SCFA) -producing bacteria.
Next, we investigated significant taxonomic shifts in the microbial community tnrough MetaStats analysis. Compared with the TNBS model group, F. prausnitzii treatment significantly increased the abundance of Ruminococcaceae (P < 0.01) and decreased the abundance of Peptostreptococcaceae (P < 0.01), Ruminococcaceae_UCG (P < 0.05), Terrisporobacter (P < 0.05), and Turicibacter (P < 0.05). These changes were the most obvious in the F. prausnitzii supernatant group (TNBS + SN) and were characterized by increased the abundance of Firmicutes (P < 0.01), Clostridiales (P < 0.05), Ruminococcaceae (P < 0.05), Clostridiales_vadinBB60 (P < 0.05), Lachnospiraceae_NK4A136 (P < 0.05), Eubacterium_xylanophilum (P < 0.05), Lachnospiraceae_bacterium_COE1 (P < 0.05), Blautia_coccoides (P < 0.05), and reducing the abundance of Rhizobiales (P < 0.05), Peptostreptococcaceae (P < 0.01), Phyllobacteriaceae (P < 0.05 ), Ruminococcaceae_UCG-002 (P < 0.01), Terrisporobacter (P < 0.01), Turicibacter (P < 0.05), Clostridium_sensu_stricto (P < 0.05), Phyllobacterium (P < 0.05), Proteus_mirabilis (P < 0.05), Streptococcus_gallolyticus (P < 0.01), and Porphyromonadaceae_bacterium (P < 0.01) (Fig. 6). Interestingly, Lachnospiraceae, Ruminococcaceae, Blautia and Eubacterium_xylanophilum are also SCFAs-producing bacteria.