Background
Rotavirus (RV) can infect almost all mammalian and avian species. RV is very common gastrointestinal pathogen in infants and children under 5 [
1]. Despite the global introduction of vaccinations for human RV over a decade ago, RV infections still result in > 200,000 deaths annually [
2‐
4]. The efficacy of RV vaccines may result from low standards of hygiene [
5], malnutrition, and disorder of the intestinal microbiota [
6,
7].
The intestinal microbiota plays an important role in host health. It has been demonstrated that the ecology and function of the microbiota are related to enteric virus infections [
8] Moreover, virus infection can change the gut microbiota composition and activity. Some viruses, such as poliovirus [
9], reovirus [
9], norovirus [
10,
11] and murine RV [
12,
13], have been reported to influence the gut microbiota. Commensal bacteria have been shown to enhance the infectivity of enteric viruses through several mechanisms, such as bacterial stabilization of viral particles, help of viral adsorption target cells, and restraining of antiviral immune responses [
14].
Previous studies have revealed that human RV infection in infants reduces the fecal microbiota diversity as compared to healthy infants [
15,
16]. Microbiota ablation resulted in reduced RV-induced diarrhea in mice model and a more durable RV-targeted antibody response via germ-free or antibiotic approaches [
17]. Another study showed that segmented filamentous bacteria (SFB) could protect mice against RV infection and associated diarrhea sufficiently [
18].
Bifidobacterium and
Lactobacillus species have been showed to be associated with increased extent of specific anti-RV immune responses, and subsequently, a shorter duration and severity of RV infection [
19,
20]. The probiotic bacteria
Escherichia coli Nissle and
Lactobacillus rhamnosus strain GG have been shown to influence the binding, infectivity, and B cell immune response of human RV [
21]. Additionally, antibiotics have been shown to increase the fecal output of RV, but also changes the beta diversity of gut bacterial, which further demonstrates that modification of the intestinal microbiota alters the immune response [
17].
It have been shown that the gut microbiota modulates RV infection and the antibody response of the host against RV infection in animal models [
8‐
13]. However, little studies regard alteration in the gut microbial upon RV infection. Thus, it is urgent to set up the etiological link between RV infection and the gut microbiota.
In this study, we elucidated the relationship between the gut microbiota and RV infection by characterizing the intestinal microbiota via 16S rRNA sequencing in a RV-induced diarrhea model in suckling mice. Our findings provide information regarding the development of probiotic therapy to ameliorate the symptom caused by RV infection or the identification of a microbial target that can inhibit RV replication and infection in children.
Materials and method
Viruses, cells, and viral load quantification
RV SA11 strain (provided by Dr Kobayashi, Osaka University, Japan) and fetal African green monkey kidney cells (MA104 cells; Cell Resource Center, IBMS, and CAMS/PUMC, Beijing, China) were used in this study. MA104 cells were grown in Dulbecco’s modified Eagle’s medium (DMEM; Thermo Fisher Biochemical, Beijing, China) supplemented with 5% calf serum (FBS; GIBCO, Paisley, UK) and 1% penicillin–streptomycin (Sigma-Aldrich, St. Louis, MO, USA) in 5% CO2 at 37 °C in a humidified incubator. RV strain SA11 was propagated in MA104 cells as in previous studies
42. The virus titer was counted by fluorescent focus assay (FFA) in MA104 cells resulting of 10
6FFU/mL, and viral load was quantified by QPCR [
22].
Total RNA was extracted from the colorectal contents of mice by using RNAiso Plus (Takara Bio Inc, Dalian, China) following the manufacturer’s protocol. The RNA was quantified using a NanoDrop 1000 Spectrophotometer (Thermo Fisher Biochemical, Beijing, China). The cDNA was prepared from 2 μg of RNA using the following primers: forward: 5′-ATCAGCAAACTGACGAAGCG-3′; reverse: 5′-CCAACTTTTCAGCTGTCGCA-3′ (Takara Bio Inc, Dalian, China). In brief, the amplification was performed using a 10-μL volume reaction in a 96-well plate with the following conditions: 1 cycle at 94 °C for 30 s, followed by 36 cycles of 94 °C for 5 s, 60 °C for 30 s, and 72 °C for 30 s. The RV RNA copy levels were quantified by comparison with a standard curve generated using ten two-fold serial dilutions of a plasmid containing the RV VP7 gene.
Animal and experimental design
Twenty specific-pathogen-free female Kunming (KM) mice were provided by the Laboratory Animal Center at Jinzhou Medical University (Liaoning Province, China). All experimental groups were housed in the same specific-pathogen-free room. The living environment, feeding conditions and microbial conditions of animals in research were always consistent which was maintained on a 12-h light/dark cycle at 22 ± 2 °C with 40–70% humidity.
The female mice were paired with male mice upon delivery. The males were removed the next day, and the pregnant females were monitored daily and allowed to deliver at term. The day of birth was recorded as day 1 of life. Litter statistics and the ratio of males/females in each cage were not calculated. One lactating female mouse and her pups were maintained together in individually ventilated autoclaved cages (IVC). Suckling mice were divided into two groups: RV-infected and uninfected groups. Each group contained 10 litters of mice. There were 7–8 mice in each litter. RV-infected groups of suckling mice were orally administered 50 μL of 10
6 PFU/mL RV strain SA-11 [
12], and the uninfected group were orally administered 50 μL of phosphate-buffered saline (PBS) as a control.
All of the suckling mice were euthanized four days (the time point at which the most severe diarrhea symptoms presented) post-RV infection. Mixed intestinal contents of the colon and rectum were collected using autoclaved tweezers and stored in sterile tubes at − 80 °C. All animal experiments were performed in accordance with the Jinzhou Medical University guidelines. All of the animal experiments in this study were approved by the Animal Welfare and Ethical Review Board at Jinzhou Medical University (approval ID: 2019014). All animal infections and infectious work were performed in biosafety level 2 facilities.
Histopathology
Five sucking mice were randomly selected from each RV group and NC group for sample collection and pathological analysis. The duodenum was harvested from the abdominal cavity immediately after the suckling mice were euthanized. For the histopathological investigation, the duodenum was fixed in a solution of 4% paraformaldehyde (0.01 M PBS, pH 7.4). For paraffin section preparation, the duodenum was dehydrated with an increasing ethanol gradient, cleared with xylene, and then embedded in wax. Three consecutive paraffin sections (5 μm thick) were used for hematoxylin and eosin (H&E) staining. For each slice, fields were randomly selected.
DNA extraction and PCR amplification
The RV-infected and uninfected control mice groups were euthanize. Each group contained 10 litters of mice, and there were 7–8 mice in each litter. Genomic DNA was extracted from mixed intestinal contents using the QIAamp DNA Stool Mini kit (Qiagen Inc., Valencia, CA, USA) according to the manufacturer protocols. The V3–V4 region of the bacterial 16S ribosomal RNA genes (342F and 806R) was amplified by PCR using primers 341F 5′-CCTACGGGRSGCAGCAG)-3′ and 806R 5′-GGACTACVVGGGTATCTAATC-3′. The PCR protocol used in this study was as follows: 95 °C for 3 min, followed by 30 cycles at 98 °C for 20 s, 58 °C for 15 s, 72 °C for 20 s, and a final extension at 72 °C for 5 min. PCR reactions were performed in a 30-μL mixture containing 15 μL of 2× KAPA Library Amplification ReadyMix (Roche-KAPA, Shanghai, China), 1 μL of each primer (10 μM), 50 ng of template DNA, and ddH2O. Amplicons were extracted from 2% agarose gels and purified using the AxyPrep DNA Gel Extraction Kit (Axygen Biosciences, Union City, CA, USA) according to the manufacturer’s instructions. The resulting library was analyzed with a Thermo NanoDrop 2000 spectrophotometer (ThermoFisher, Shanghai, China) and 2% agarose gel electrophoresis. Once the library passed the quality inspection, it was quantified by Qubit and mixed according to the data requirements of the quantification and normalization of individual PCR products.
Sequencing of 16S rRNA gene amplicons
The PCR products were quantified using Qubit (Invitrogen, Carlsbad, CA, USA), multiplexed at an even concentration, and subjected to 400–450 bp pair-end sequencing. The adaptor was added to the products, and the samples were sequenced on an Illumina MiSeq platform (Illumina, Inc., San Diego, CA, USA). The reads were assembled and used for subsequent 16S analysis. The assembled reads were filtered to acquire clean reads. Reads with an average quality value below 20 and number N that was more than 3 was removed, then the sequences spanning the entire V3–V4 amplicon were matched using PANDAseq [
23]. Merged sequences with 97% nucleotide sequence identity (97% identity) were binned into operational taxonomic units (OTUs) using UPARSE [
24]. Based on the RDP classifier, a representative sequence of each OTU was assigned to a taxonomic level in the RDP database using 0.8 as the minimum confidence threshold [
25].
Statistical analysis of the data
Alpha diversity indices, including the number of OTUs, observed species diversity, and Shannon and Simpson indices, were calculated by normalizing the number of clean reads in all samples to 47,504 sequences using mother software [
26]. Rarefaction curves were analyzed with mothur and plotted using R. A representative sequence was chosen from each OTU by selecting the sequence that had the largest number of hits in the OTUs. The non-parametric Wilcoxon test (Wilcox. Test in R) was performed for each index of alpha diversity. Rank sum test was used to screen the alpha diversity indices with significant differences under different conditions.
Beta diversity was calculated for the normalized OTU table using UniFrac distance matrices [
27,
28] in order to determine the amount of bacterial diversity shared between the two groups. Principal coordinate analysis (PCoA) of bacterial communities was performed using weighted UniFrac distances based on the presence and absence of OTUs, and the plot was generated using PERMANOVA, which was also performed using weighted UniFrac distance to test for differences in bacterial community composition in the samples from the two groups.
Linear discriminant analysis effect size (LEfSe) analysis was used to determine the features most likely to explain differences between the RV-infected group and healthy control group. Different features with an LDA score were identified [
29]. The
p value was set at
p < 0.05, and the threshold on the logarithmic linear discriminant analysis (LDA) score was 2 [
30]. In addition, heatmap analysis was performed to compare the significant differences at the genus level in the RV-infected and uninfected groups. PICRUSt (version 1.0.0,
http://picrust.github.com/) [
31] was performed to predict the microbial community function of the two groups. Finally, a statistical analysis of the taxonomic and functional profiles (STAMP, version 2.1.3,
http://kiwi.cs.dal.ca/Software/STAMP) [
32] was used for further exploration in level 2 of the KEGG analysis using Reconstruction of Unobserved States (PICRUSt) analysis.
Discussion
Enteric viral infections remain a major public health challenge. Since enteric viruses first encounter host cells amidst the microbiota, the microbiota composition might influence RV infection [
18]. Whereas, only narrowed amount of information regarding the correlation between the gut microbiota composition and RV infection. Therefore, the effects of RV infection on the intestine and intestinal microbiome in suckling mice were investigated in this report.
A previous report showed that gut microbial diversity was cut down in infants infected with human RV, however the enrichment of diversity was observed in healthy children’s, such as phylum Firmicutes was abundant [
15]. Additionally, it has been shown that there is a reduction of microbial diversity in children with RV infection as compared to children without infection [
33,
34]. In this work, we also observed low bacterial diversity in the RV group as compared to healthy mice. Therefore, the gut microbiota appears to take an important role in diarrhea-associated processes. The intestinal symbiotic role in regulating viral infections has been realized. An RV-infected suckling rat model was formerly taken to assess the effects of prebiotic mixtures in fermented milk on the reforming the course of infectious and amendment the immune response [
13]. The gut microbiota has also been shown to affect intestinal virus replication in mice [
9]. These study demonstrated that as RV infection model, suckling mice was used to assess the change of intestinal flora is feasible.
Higher diversity and integrity of intestinal flora are beneficial to intestinal ecosystem [
35]. The integrity of the microbiota for more and more physiological process is indispensable. The breakdown of homeostasis of the microbiota is associated with a variety of pathological states [
35]. Consequently, interference of the microbial balance (dysbiosis) may have substantial consequences for the metabolism and adaptive immune responses of the host. Importantly, commensal microbiota protect against invading bacterial pathogens. Loss of intestinal flora diversity is the most common symptom of intestinal disorders. Restoration of the intestinal flora diversity may be an option for the treatment of people with risk [
36]. In order to discuss the influence of RV infection on the relative abundance of specific microbiome taxa, we identified the bacterial genera that were different between RV-infected suckling mice and healthy suckling mice. We found alterations in the gut microbiota of the RV group. The relative abundance of
Proteobacteria and
Candidatus saccharibacteria was increased, while the abundance of
Firmicutes,
Fusobacteria,
Bacteroidetes, and
Actinobacteria was decreased as compared to NC group, which is consistent with a previous report where the phylum
Bacteroidetes was reduced in RV-induced diarrheal disease children [
37]. The gut microbiota in healthy mice was dominated by
Firmicutes (59.29%), followed by
Proteobacteria (28.43%), and
Fusobacterial (9.29%) during the neonatal period.
Notably, there was significant increase in the genera
Escherichia/shigella in RV-infected mice. However,
Lactobacillus,
Fusobacteria,
Streptococcus, and
Helicobacter were decreased.
Lactobacillus was also found decreased in a BALB/c model with rhesus rotavirus (RRV) infection [
38]. The present LEfSe analysis showed that
Helicobacter,
Alloprevotrlla,
Brevundimonas,
Paenibacillus, and
Parabacteroides were completely undetectable in the RV-infected group. Former studies have shown that
Lactobacillus and
Helicobacter are regarded as probiotics with certain protective effects, which can eliminate infections, attenuating both GI diseases and produce lactate and butyrate [
39]. Probiotics have been increasingly used to enhance the oral vaccine reactions and treat some intestinal infection and inflammatory of GI in children [
40]. In probiotics, gram–positive (G+) probiotics, such as
Lactobacillus spp. or
Bifidobacteria spp., have been used in randomized clinical trials in humans [
41,
42] and experimental studies [
19,
20,
43,
44] to reduce the severity of the diarrhea caused by RV.
Lactobacillus rhamnosus GG (LGG) has been extensively investigated for its beneficial health effects, such as shortening the duration of HRV diarrhea and enhancing HRV-specific immune responses in children [
41,
45]. However, one study regarding
Lactobacillus probiotics showed that probiotic supplementation in Indian infants did not substantially boost RV vaccine response in a randomized controlled trial [
46]. Therefore, the effect of probiotics use for diarrhea is still unclear. A modest effect of probiotics supplementation deserves further investigation.
In this study, we observed a significant increase in the
Escherichia/shigella genera in the RV-infected group, suggesting that RV likely promotes the disruption of the epithelial integrity and further induces changes in the abundance of
Shigella. Although this study suggests that E. coli may have potentially detrimental effects on the intestine. However, some reports show that E. coli Nissile has been shown to ameliorate diarrhea in RV infection in neonatal gnotobiotic (Gn) piglets models. These studies highlight strain specificity in the Escherichia may play a role in rotavirus infection [
47‐
49].
The abnormal abundance of
Escherichia/
Shigella in the intestine may be closely associated with many diseases.
Some studies have shown that an increase in intestinal Shigella in patients with rheumatoid arthritis [
50]
and compensated heart failure [
51]
is accompanied by a significant decrease in Lactobacillus and Firmicutes. In the gut microbiota of mice infected with Shigella, Lactobacillus supplementation inhibited the proliferation of
Shigella within the intestinal flora of mice [
52]. In this study, these same changes in the bacterial flora were observed in the intestine of suckling mice infected with RV virus. These results showed that changes in the abundance of
Lactobacillus,
Firmicutes, and
Escherichia/
Shigella correlated in intestinal homeostasis.
In the present study, we predicted the unobserved character states in the bacterial community by using PICRUSt, which is generally applied to investigate the intestinal function of suckling mice. With respect to the results of the LEfSe and STAMP analyses, “carbohydrate metabolism” and “lipid metabolism” pathways were significantly enriched in the NC group. Multiple studies have suggested that the gut microbiota influences host metabolism and function [
53]. Recent research has identified many virus-specific metabolic pathways and that eukaryotic viruses induce mass changes of metabolism of the host [
54]. Lipid plays an important role in viral infection because lipids are structural elements of cells and viral membranes [
55], membrane fusion, envelopment, remodeling, and lipid compounds, such as cholesterol and sphingolipids, are vital for viral replication. Viruses also make lipids at sites of replication by promoting lipid biosynthesis [
56]. It has been reported that hepatitis C [
57] and poliovirus [
9] influence lipid metabolism and bind specific microbe-associated surface polysaccharides, enhancing viral attachment to host cells.
Virally-mediated manipulation of lipid metabolism is important for viral infection, and viruses have evolved multiple mechanisms to ensure that the host-cell lipid metabolism is successfully hijacked to support viral infection. A previous study reported that RV replication was susceptible to inhibitors targeting various lipid synthetic enzymes [
58]. According to our data, “carbohydrate metabolism” and “lipid metabolism” pathways are an important part of the host metabolism, which is potentially influenced by the response of the intestinal microbiota to RV infection.
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