Next Article in Journal
Prevalence of Antibiotic-Resistant Escherichia coli Isolated from Beef Cattle and Dairy Cows in a Livestock Farm in Yamagata, Japan
Previous Article in Journal
Universal Lineage-Independent Markers of Multidrug Resistance in Mycobacterium tuberculosis
Previous Article in Special Issue
Effects of Age in Fecal Microbiota and Correlations with Blood Parameters in Genetic Nucleus of Cattle
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Microorganisms
Volume 12
Issue 7
10.3390/microorganisms12071341
microorganisms-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Effect of Probiotic Bacteria on the Gut Microbiome of Mice with Lipopolysaccharide-Induced Inflammation

by
Mariya Gryaznova
1,
Inna Burakova
1,
Yuliya Smirnova
1,
Polina Morozova
1,2,
Egor Chirkin
2,
Artem Gureev
2,
Evgeny Mikhaylov
3,
Olga Korneeva
1 and
Mikhail Syromyatnikov
1,2,*
1
Laboratory of Metagenomics and Food Biotechnology, Voronezh State University of Engineering Technologies, 394036 Voronezh, Russia
2
Department of Genetics, Cytology and Bioengineering, Voronezh State University, 394018 Voronezh, Russia
3
FSBSI All-Russian Veterinary Research Institute of Pathology, Pharmacology and Therapy, 394061 Voronezh, Russia
*
Author to whom correspondence should be addressed.
Microorganisms 2024, 12(7), 1341; https://doi.org/10.3390/microorganisms12071341 (registering DOI)
Submission received: 21 May 2024 / Revised: 27 June 2024 / Accepted: 28 June 2024 / Published: 30 June 2024
(This article belongs to the Special Issue Probiotics, Prebiotics, and Gut Microbes)
Browse Figures
Versions Notes

Abstract

:
The role of lipopolysaccharide (LPS) in the development of diseases is clear, but the specific mechanisms remain poorly understood. This study aimed to investigate the microbiome aberrations in the guts of mice against the background of LPS, as well as the anti-inflammatory effect of probiotic supplementation with Lactobacillus plantarum from the gut, a mix of commercial probiotic lactic acid bacteria, and Weissella confusa isolated from milk using next-generation sequencing. LPS injections were found to induce inflammatory changes in the intestinal mucosa. These morphological changes were accompanied by a shift in the microbiota. We found no significant changes in the microbiome with probiotic supplementation compared to the LPS group. However, when Lactobacillus plantarum and a mix of commercial probiotic lactic acid bacteria were used, the intestinal mucosa was restored. Weissella confusa did not contribute to the morphological changes of the intestinal wall or the microbiome. Changes in the microbiome were observed with probiotic supplementation of Lactobacillus plantarum and a mix of commercial probiotic lactic acid bacteria compared to the control group. In addition, when Lactobacillus plantarum was used, we observed a decrease in the enrichment of the homocysteine and cysteine interconversion pathways with an increase in the L-histidine degradation pathway.

1. Introduction

Lipopolysaccharide (LPS or endotoxin) is a thermostable component of the outer part of the cell membrane of most Gram-negative microorganisms [1]. This macromolecule can lead to dysregulation of the immune system, which plays an important role in the development of a wide range of pathologies, including neurodegenerative, metabolic, and cardiovascular diseases [2].
LPS consists of three covalently linked components: lipid A, which after the destruction of the bacterial cell is released into blood and can have a severe toxic effect; a central oligosaccharide, which is also an endotoxin and, when released into blood, causes a toxic effect, but to a lesser extent than lipid A; and O-antigen is the most immunogenic part of LPS, which is easily recognized by the host immune system [3].
By its nature, LPS is not toxic, but its effect is mediated by the activation of the immune system and depends on many factors that alter the host’s susceptibility [4]. Small amounts of LPS are released from the surface of bacteria, while large amounts are released upon bacterial cell lysis or death [5].
LPS is involved in the development of various diseases; for example, its role has been shown in Parkinson’s disease, Alzheimer’s disease [6], diabetes, obesity [7], and atherosclerosis [8], among others.
The search for a therapeutic approach is mainly associated with the neutralization of LPS or the inhibition of LPS-mediated activation of immune cells. However, such experiments did not lead to the expected positive results [3]. The intestinal microbiota is capable of producing various metabolites, regulating the number of pathogenic bacteria, and modulating the immune response, for example, by reducing intestinal permeability to LPS [9]. All of this suggests that taking probiotic bacteria may be able to correct the toxic effect caused by LPS.
Probiotic supplements are known to have the ability to restore gut microbiome composition and impart beneficial functions to host gut microbial communities, which in turn reduces or prevents the development of intestinal inflammation and other intestinal or systemic disease phenotypes [10].
Lactobacillus plantarum is a widely used probiotic with specific beneficial immunoregulatory properties, including regulation of the immune response [11]. In general, bacteria of the genus Lactobacillus are capable of modulating the immune system and exerting anti-inflammatory effects [12]. Although some studies indicate that the bacterium Weissella confuse can cause sepsis and other serious infections in humans and animals [13], it is often considered as a probiotic supplement, as it has high survival rate in gastrointestinal diseases and antimicrobial activity against foodborne pathogens [14]. All this makes these microorganisms potential probiotic candidates to reverse the toxic effects caused by LPS.
Currently, the effect of probiotic bacteria on inflammatory processes in the gut, which can be induced by the LPS, has been poorly studied. The purpose of this study was to examine the effect of probiotic supplementation with Lactobacillus plantarum from the gut, a mix of commercial probiotic lactic acid bacteria with five cultures of Lactobacillus bacteria, and also Weissella confusa isolated from cow’s milk on the development of LPS-induced inflammation in mice.

2. Materials and Methods

2.1. Study Objectives

Two-month-old male C57BL/6 mice were obtained from the Stolbovaya nursery (Stolbovaya, Russia) and kept at the animal facility at 25 °C and 40% humidity on a 12 h light/dark cycle. The mice were divided into five groups. The control mice (n = 10) group (C) received a standard laboratory diet (maintenance diet for rats and mice, Altromin Spezialfutter GmbH) throughout the entire duration of the experiment. In the 4th week, the mice received intraperitoneal saline injections for 7 days. The second group (LPS) of mice (n = 10) received a standard laboratory diet. In the 4th week of the experiment, the mice received intraperitoneal injections of lipopolysaccharides (LPS) (trade name Pyrogenal, N.F. Gamaleya Federal Research Center for Epidemiology & Microbiology, Russia) at 375 μg/kg/day. LPS is known to cause inflammatory responses in the gut [15]. The 3rd–5th groups of mice also received LPS injections in the 4th week of the experiment for 7 days (according to previous studies [16,17]) at a concentration of 375 μg/kg/day. The 3rd group (LA) of mice (n = 10) received a laboratory diet mixed with probiotic lactic acid bacteria (Lactobacillus delbrueckii subsp. bulgaricus VSUET15; Lactiplantibacillus plantarum VSUET13; Lactobacillus acidophilus VSUET12; and Limosilactobacillus fermentum VSUET14) from a commercial probiotic product [18,19] in a ratio of 1 × 108 CFU/g of diet. The 4th (LP) and 5th (W) groups of mice received a laboratory diet mixed with Lactobacillus plantarum VSRVR2 isolated from the gut of a 5-day-old pig (n = 10) and Weissella confusa VSRVR4 isolated from cow’s milk (n = 10), respectively, in a ratio of 1 × 108 CFU/g of diet. It was previously shown that this amount of lactic acid bacteria in the feed is considered as medium [20,21]. The average daily feed consumption by mice was 5.1 ± 0.08 g, which corresponds to each mouse consuming on average 5.1 × 108 bacteria daily.
On the 29th day of the experiment, fecal samples were collected for analysis of the bacterial composition of the gut microbiome. A total of one fecal sample was collected from each mouse, which was then used for DNA extraction and sample preparation for subsequent sequencing. DNA sequencing of fecal samples using next-generation sequencing tools to assess the bacterial composition of the gut microbiome is a widely accepted practice [22,23,24]. After the course of injections, the mice were sacrificed. The mice were anesthetized with isoflurane before the sacrificing. For morphological studies of the small intestine, tissue samples were fixed in a 10–12% solution of neutral formalin HistoSafe (Biovitrum, Moscow, Russia).
The animal study protocol was approved by the Ethics Committee of FSBSI “All-Russian Veterinary Research Institute of Pathology, Pharmacology and Therapy” (protocol number 1-02/23, 10 February 2023). All procedures were in accordance with the “Animal Research: Reporting of In Vivo Experiments” (ARRIVE) guidelines and took into account the 3R principles.

2.2. Histological Examination

For morphological studies of the mouse small intestine, tissue samples were fixed in a 10–12% HistoSafe neutral formaldehyde solution (Biovitrum, Moscow, Russia). The intestinal tissue was dehydrated with increasing concentrations of ethyl alcohol, followed by embedding in histological paraffin “Histomix” (Biovitrum, Moscow, Russia). Sections of 3–5 µm thickness were cut from the paraffin blocks using a Hospitex Diagnostics rotary microtome. Sections were stained using classical histological methods (haematoxylin and eosin). Histological examination was performed using a Hospitex Diagnostics microscope with a built-in camera with a resolution of 5 megapixels.
Morphometric analysis was performed in the intestinal mucosa in 10 fields of view of the microscope (eyepiece 10, objective 40).

2.3. Whole-Genome Sequencing

Total DNA was isolated using a commercial HiPure Microbiome DNA Kit (Magen, Guangzhou, China) according to the manufacturer’s protocol.
Sequencing was performed on the DNBSEQ-G50 platform (MGI, Shenzhen, China) according to the MGIEasy Fast FS DNA Library Prep Set protocol. The process includes fragmentation of the total DNA set MGIEasy Fast FS Library Prep Module (MGI, China), adapter ligation, and ligation of barcodes with PCR. The PCR protocol involves 95 °C for 3 min, followed by 7 cycles of 98 °C for 20 s, 60 °C for 15 s, and 72 °C for 30 s. After finishing the cycles, the samples were incubated for an additional 10 min at 72 °C. The next steps include a magnetic cleaner, monitoring the yield of the PCR product using electrophoresis in a 2% agarose gel, measuring concentration with a fluorometer, calculating the mass of the PCR product, calculating the amount of sample to be added to the pool, holding the pools, circularization of single-stranded DNA using the MGIEasy Dual Barcode Circularization Module kit (MGI, Shenzhen, China), cleaning the resulting product with magnetic particles, measuring DNA concentration with a fluorometer and QuDye ssDNA Assay Kit (Lumiprobe, Moscow, Russia), calculating the volume in μL of each pool that must be added when compiling a super pool, measurement of super pool concentration, calculating the volume in μL of the super pool covering 60 fmol, creation of DNB, measuring the concentrations of the resulting DNB using a fluorometer, and loading the cartridge.

2.4. Bioinformatics and Statistical Analysis

The quality of raw reads was assessed using FastQC [25]. Technical sequences and low-quality bases (Q < 30) were trimmed with fastp [26]. To remove human and mouse genome sequences, reads were mapped to reference genomes using Bowtie2 [27]. Taxonomic profiling of bacteria, viruses, and eukaryotes that form the microbiome of the studied samples was performed using MetaPhlAn4 [28]. HUMAnN3.0 was used for the functional profiling of metabolic pathways [29].
Statistical manipulations were performed in the R environment. Phyloseq was used to calculate microbiome diversity measures. The Shannon index was used to assess alpha diversity, and the Bray–Curtis dissimilarity metric was used for beta diversity. Differences in alpha diversity were evaluated using the Wilcoxon rank sum test [30]. The ADONIS function was used to estimate between-group diversity differences. Species differential abundance analysis was conducted using the MaAsLin2 package, which employed a multivariate regression model [31]. The Tukey test was used to statistically analyze the differences between groups, based on the normal distribution of the values of the morphometric parameters. A statistically significant result was considered to be an adjusted p-value of ≤0.05.

3. Results

3.1. Histological

In the control group of mice, the epithelial lining of the villi was preserved, with no signs of infiltration. The submucosa consisted of connective tissue with blood vessels. Crypts were visualized (Figure 1A).
In the LPS group, focal infiltration of lymphocytes and plasma cells with moderate neutrophil content was observed, mostly scattered in the villi. The epithelial lining of the villi was disrupted with signs of infiltration. The submucosa was represented by connective tissue with vessels in which signs of infiltration were observed. These changes were indicative of inflammation (Figure 1B).
In the LP group, the epithelial lining of the villi was preserved without signs of infiltration. The submucosa consisted of connective tissue. The crypts in the small intestine were flat and visualized. Secretion from goblet cells was observed (Figure 1C).
In group W, the intestinal architecture was the same, the mucosa had uneven oedema, the integumentary epithelium was largely preserved, in some places, the apical part of the villi was partially desquamated, and the rest of the area consisted of tall prismatic enterocytes with a well-defined brush border. The epithelial lining of the intestine was intact, and the cells were tightly packed (Figure 1D).
In the LA group, the epithelial lining of the villi was desquamated with signs of infiltration. The submucosa consisted of connective tissue. The crypts were shallow and slightly tortuous, and mitotic activity was low. Desquamation of the epithelial layer was noted in the apical part (Figure 1D).
Significant differences between the study groups were confirmed by morphometric analysis of the mice’s small intestines (Table 1).
Morphometric examination of intestinal tissue samples from the groups of mice revealed a statistically significant decrease in villus height in the LP group compared to the control group (102.27 ± 22.28 vs. 144.23 ± 23.21, respectively, p = 0.008). We also observed that the mean villus height in the LPS group was 121.24 ± 13.27 microns, which was lower compared to the control and some probiotic groups. At the same time, the LPS group showed a thickening of the intestinal villi of 28.72 ± 2.25 microns compared to the other study groups, and the values of the creep depth index in the LPS group were the smallest at 35.66 ± 7.65 microns. Thus, the difference between the intestinal morphometric parameters in the W, LA, and LP groups relative to the LPS group was 12.56%, and the overall difference between the LPS and control groups was 23.3%.

3.2. Microbiome Composition

Whole-genome shotgun sequencing on the DNBSEQ-G50 platform identified 446 bacterial and 2 eukaryotic species in the samples studied. In total, 290 bacterial species were identified as previously uncharacterized, reflecting the fact that the mouse fecal microbiome is a poorly studied consortium (with many uncultured and therefore few correctly named species). The abundance of 58 species was greater than 1%; they were the most abundant species in the study groups; all other species were grouped as “Others” (Figure 2).
The most abundant species in the control, LPS, and W groups was the uncharacterized GGB50207_SGB70279 belonging to the phylum Bacteroidetes. Its abundance was 10.66% ± 0.31, 9.90% ± 0.54, and 17.34% ± 0.30 for the control, LPS, and W groups, respectively. Lactobacillus taiwanensis was the most commonly described species in the control group (10.30% ± 0.19), Limosilactobacillus reuteri in the LPS group (5.80% ± 0.01), and Bacteroidales bacterium in the W group (4.33% ± 0.02). The most abundant species in the LP group was also Bacteroidales bacterium (10.93% ± 0.16). Ligilactobacillus murinus was the most abundant species in the LA group (7.68% ± 0.37).
We also conducted an alpha diversity analysis using a measure of observed species as well as the Shannon index (Figure 3).
The alpha diversity of the mouse fecal microbiome was characterized as high, as the Shannon index value was greater than three for all study groups (Table 2).
The state of the fecal microbiota was not statistically different between the groups of mice studied and can be characterized as balanced and stable due to high bacterial diversity.
For the study groups, we also examined the beta diversity of the mouse fecal microbiome (Figure 4).
A statistically significant bacterial clustering between the control and LP groups has been observed (p = 0.03).
Differential abundance analysis has revealed statistically significant differences in the abundance of 16 bacterial species between the groups studied (Figure 5).
It has been found that the abundance of Adlercreutzia equolifaciens (0.04% ± 0.02 vs. 0.58% ± 0.20, p = 0.01), Lactococcus lactis (0.04% ± 0.02 vs. 1.02% ± 0.40, p = 0.02), and GGB75046_SGB102341 belonging to Eggerthellaceae (0% vs. 0.54% ± 0.20, p = 0.01) was lower in the LP group compared to the control group. At the same time, the abundance of Duncaniella muris (1.95% ± 0.40 vs. 0.50% ± 0.12, p = 0.02), Muribaculaceae bacterium (0.54% ± 0.17 vs. 0.004% ± 0.003, p = 0.02), Staphylococcus nepalensis (0.35% ± 0.27 vs. 0.00005% ± 0.00005, p = 0.03), Staphylococcus saprophyticus (1.11% ± 0.43 vs. 0%, p = 0.03), GGB18827_SGB27766, which is closest to Bacteroides acidifaciens (1.49% ± 0.97 vs. 0.09% ± 0.08, p = 0.01), GGB24134_SGB35937 belonging to Lactobacillaceae (1.27% ± 0.28 vs. 0.09% ± 0.04, p = 0.004), GGB27849_SGB40283 belonging to Bacteroidales (3.23% ± 0.69 vs. 0.55% ± 0.22, p = 0.03), GGB27872_SGB40306 belonging to Bacteroidetes (0.42% ± 0.07 vs. 0.06% ± 0.02, p = 0.002), GGB27892_SGB40326 belonging to Selenomonadaceae (1.04% ± 0.26 vs. 0.04% ± 0.02, p = 0.002), GGB27925_SGB40363 belonging to Muribaculaceae (0.30% ± 0.14 vs. 0%, p = 0.02), GGB27929_SGB40368 belonging to Staphylococcaceae (0.48% ± 0.20 vs. 0.0008% ± 0.0008, p = 0.002), and GGB3583_SGB21441 belonging to Muribaculaceae (2.96% ± 0.80 vs. 0.09% ± 0.03, p = 0.00009) was higher in the LP group compared to the control group.
The abundance of GGB27929_SGB40368 belonging to Staphylococcaceae (1.35% ± 0.57 vs. 0.0008% ± 0.0008, p = 0.004) and GGB3583_SGB21441 belonging to Muribaculaceae (3.91% ± 1.31 vs. 0.09% ± 0.03, p = 0.01) was higher in the LPS group compared to the control group.
The abundance of GGB27929_SGB40368 belonging to Staphylococcaceae (0.26% ± 0.07 vs. 0.0008% ± 0.0008, p = 0.02) was higher in the LA group compared to the control group, but the abundance of GGB31189_SGB59906 belonging to Erysipelotrichaceae (0.18% ± 0.14 vs. 6.83% ± 2.22, p = 0.04) was lower in the LA group compared to the control.
Bacterial abundance in group W was not significantly different from the control.
Profiling of metabolic pathways in the studied microbiome community was also carried out. In total, 335 functional profiles were identified. However, the frequency of unintegrated and unmapped pathways was around 97%. The statistical analysis showed that there was a significant difference in the abundance of three metabolic pathways between the control group and the LP group (Figure 6).
It can be observed that the homocysteine and cysteine interconversion pathway (0.0003% ± 0.0001 vs. 0.003% ± 0.001) was less abundant in the LP group compared to the control. On the other hand, the L-histidine degradation pathway (0.17% ± 0.002 vs. 0.003 ± 0.001) is most abundant in the LP group compared to the control (in all cases, p = 0.03).

4. Discussion

A morphological study of mouse intestinal sections showed no changes in the morphological structure of the intestine in the control group. The presence of inflammatory sites, as well as the desquamation of intestinal villi, the release of secretions from goblet cells, and hemorrhages were found in intestinal sections obtained from mice in the LPS group. A reduction in the inflammatory process in the small intestine was also seen when people took probiotic supplements containing Lactobacillus plantarum and a mixture of lactobacilli.
To study the mouse fecal microbiome, we used the DNBSEQ-G50 whole-genome sequencing platform (MGI, China). This approach is more sensitive to small changes, has higher resolution, and provides a more complete picture of the structure and function of host microbial communities than 16S rRNA gene sequencing, which is used in most gut microbiota studies [32]. The microbiome composition of the study groups was characterized as rich and diverse. In addition to high numbers of bacteria, two closely related eukaryotic species were also identified in the mouse feces: Cryptosporidium hominis and Cryptosporidium parvum, although at low concentrations. Cryptosporidium, a ubiquitous protozoan parasite of vertebrates, causes cryptosporidiosis, a diarrheal disease that can become chronic and life-threatening in the absence of a competent immune response. C. hominis and C. parvum are known to be responsible for most human cases of cryptosporidiosis. Domestic and wild rodents have been reported to be susceptible to infection with Cryptosporidium spp., although they are usually asymptomatic [33,34,35]. We observed the presence of both parasite species in group W (one infected mouse). C. hominis was also detected in the LA group (one mouse only). No signs of infection were observed in these rodents.
Analysis of the differential abundance of bacterial species present in the fecal microbiome of mice from the studied groups showed the presence of statistically significant differences. We observed the greatest number of differences between the control and LP groups. We also observed changes in bacterial abundance between the control and LPS and LA groups. However, the bacterial composition of group W did not differ from that of the control group. We also observed no differences in the microbiome composition between the LPS group and the groups in which the mice were fed different probiotics. Thus, although there were morphological signs of intestinal wall regeneration in the LA and LP groups, we cannot isolate specific microorganisms that contributed to the pro-inflammatory and regenerative process.
The bacterium Adlercreutzia equolifaciens, belonging to the family Mycobacteriaceae, is known to contribute to the metabolism of polyphenols and is capable of forming bioactive molecules that reverse metabolic disorders in obesity and diabetes. Along with this, A. equolifaciens is an equol-producing bacterium and is also responsible for the production of dihydroresveratrol [36,37]. Previously, through analysis of the intestinal microbiota, it was found that, compared with a healthy control group, the number of A. equolifaciens bacteria decreased in a group of patients with liver diseases. This species has also been shown to decline as the disease progresses and eventually disappears in the final stages, indicating a strong association with disease severity. In addition, A. equolifaciens bacteria have been shown to have anti-inflammatory properties [38]. In a study of changes in the gut microbiota in men with hyperuricemia, it was shown that the bacterium A. equolifaciens may alleviate the damage from the disease by somehow protecting against estrogen [39]. The results of our study are consistent with previously obtained data. We showed that the bacterial species A. equolifaciens decreased in the group taking a bacterial supplement of Lactobacillus plantarum relative to the control group.
The bacterial species Duncaniella muris is widespread in the intestinal tract of mice [40]. It is known that this bacterium is capable of breaking down complex plant polysaccharides, such as hemicelluloses and pectins [41]. It was also previously shown that exposure to the antibiotic sulfamethazine significantly increased the relative abundance of D. muris bacteria in the offspring of female mice [42]. It has been previously revealed that one of the taxa that predicted the effectiveness of the combination of early inulin and pantoprazole was D. muris, which is known to produce short-chain fatty acids [43]. It was also noted that colonization with D. muris increased susceptibility to sodium dextran sulfate-induced intestinal inflammation in pathogen-free wild-type mice, a finding that may indicate that the reduction in acetate production is an indirect result of the dominance of specific Bacteroidales groups typically associated with increased inflammation after intestinal injury [44]. According to the results of our study, it was found that in the group of mice taking the Lactobacillus plantarum supplement, the number of the bacterial species D. muris was higher than in the control group. Possibly, the injections of LPS into this group of mice might be the cause.
Analysis of the results also showed an increase in the abundance of previously uncharacterized species found in the feces of mice that took a supplement consisting of Lactobacillus plantarum. These species were GGB18827 SGB27766 and GGB27872 SGB40306, belonging to the phylum Bacteroidetes. In turn, the uncharacterized species GGB18827 SGB27766 is closest to Bacteroides acidifaciens. One of the most common bacterial phyla present in the gut microbiota is Bacteroidetes [45]. Representatives of this type mostly inhabit the distal intestine, where they help provide the host with energy obtained from the diet through the fermentation of indigestible polysaccharides. This process can produce short-chain fatty acids, which provide up to 10% of daily calories if the diet is rich in fiber [46]. B. acidifaciens is one of the dominant commensal bacteria in the gut [47]. A study in a mouse model showed that B. acidifaciens had a protective effect against concanavalin A-induced liver injury [48]. Nagahara et al. (2020) showed that B. acidifaciens can increase intestinal immunoglobulin A levels, protect the intestine from pathogen infection, and alleviate inflammatory bowel disease [49]. Another study found that alginate oligosaccharide and its compound supplement could enhance immune function and improve intestinal microecological disorders by increasing the abundance of B. acidifaciens [50]. A study also showed that B. acidifaciens can alleviate dextran sulfate sodium (DSS)-induced colitis by reducing colonic mucosal damage, reducing inflammatory response, promoting mucosal barrier repair, and restoring intestinal microbiota diversity and balance in mice [51].
Moreover, an increase in the abundance of the uncharacterized species GGB27849 SGB40283, belonging to the order Bacteroidales, relative to the control group, was identified. Species of the order Bacteroidales are the most common Gram-negative bacteria in the human gut microbiota [52]. Bacteroidales have unique and active carbohydrate utilization systems and create networks of syntrophic interactions [53]. In addition, an increase in antibiotic resistance has been noted for Bacteroidales species [54].
Lactococcus lactis is a Gram-positive, facultative anaerobic bacterium; hundreds of strains of this bacterium are known [55]. The main beneficial effect reported is the anti-inflammatory potential of L. lactis, indicating its potential use as a therapeutic agent for chronic intestinal diseases [56,57]. For example, in a study by L Steidler et al., it was shown that intragastric administration of Lactococcus Lactis, which secretes IL-10, caused a 50% reduction in the incidence of colitis in mice treated with dextran sulfate sodium [58]. Nevertheless, this characteristic has positioned L. lactis as a promising probiotic and therapeutic agent. However, the pathogenicity of some L. lactis strains has been shown in many studies in recent years. Thus, in a study by J Antolín et al., a case of liver abscess caused by Lactococcus Lactis cremoris in an immunocompetent adult patient was described for the first time [59]. Some studies have shown that L. lactis can cause mastitis in cows, but understanding the pathogenesis of such mastitis presents a new challenge [60]. Our study showed a decrease in the number of these bacteria in the drug group compared to the control group. This may be due to the pathogenicity of some strains, but further research is needed to make a definitive statement.
Muribaculaceae bacterium is a strictly anaerobic bacterium that is extremely difficult to culture. It is quite widely represented in the intestines and is capable of breaking down complex carbohydrates [61]. These bacteria were found to represent a new probiotic with anti-inflammatory effects [62]. Our study showed a significant increase in these bacteria in the drug group compared to the control group, which may indicate an increase in the probiotic effect.
Our study showed an increase in the relative abundance of the uncharacterized bacterium GGB27929 SGB40368, belonging to the Staphylococcaceae family, in the LPS, LP, and LA groups compared to the control group. It is known that many representatives of this family are opportunistic microorganisms [63]. LPS can stimulate the increase in bacteria of this family, and administration of probiotics in the LP and LA groups does not appear to affect this. It is interesting to note that in group B, such a difference compared to the control group was not observed, which may indicate the influence of this probiotic on the number of Staphylococcaceae bacteria. However, it is difficult to say this unequivocally, since this type of bacteria, as indicated, still needs to be characterized.
We showed that in the LP and LA groups, the uncharacterized species of bacteria GGB75046 SGB102341, belonging to the family Eggerthellaceae, was completely absent. Members of the Eggerthellaceae are common in the mammalian gut microbiota [64,65,66] but are rarely found in other environments. Despite having genes responsible for degrading host mucins, little is known about the role of Eggerthellaceae species in the gut [67]. Bacteria from the Eggerthellaceae family have been found to produce urolithin, have anti-inflammatory and antioxidant properties, and can also stimulate the AhR (Aryl hydrocarbon receptor) to activate tight junction proteins, thereby having anti-tumor activity [68,69]. The properties described above suggest that some members of this family could be classified as new-generation probiotics. However, we did not observe this family to become enriched in mice fed probiotic supplements.
Furthermore, the results of our experiment demonstrate that bacteria of the species GGB31189 SGB59906, belonging to the family Erysipelotrichaceae, were significantly reduced in the LA group compared to the control group. Members of the Erysipelotrichaceae family are widely distributed in the intestine and are associated with metabolic disorders and the development of inflammatory diseases [70]. An increase in these bacteria has been repeatedly shown in colorectal cancer [71], Crohn’s disease [72], IBD [73], and others. Thus, we can conclude that taking a mixture of lactobacilli can reduce the number of these bacteria in mice, helping to reduce the likelihood of developing inflammatory diseases, while other studied probiotics do not achieve this effect.
An increase in the abundance of Staphylococcus nepalensis compared to the control group was observed after the addition of Lactobacillus plantarum to the diet of mice. Staphylococcus nepalensis is a member of the genus Staphylococcus that was first isolated from the respiratory tract of goats with pneumonia in the Himalayan region of Nepal, and later from human urine samples, the gastrointestinal tract of squirrel monkeys, the oral cavity of domestic cats, the skin of pigs, and the environment [74,75,76]. Although Staphylococcus nepalensis is a normal part of mammalian flora, it needs to be studied more closely because it can cause infectious diseases and can also transfer drug-resistance genes to Staphylococcus aureus [74]. This study is the first to show the presence of this bacterium in the fecal microbiome of mice and its increase in response to feeding Lactobacillus plantarum after injection of lipopolysaccharide.
The abundance of Staphylococcus saprophyticus increased with Lactobacillus plantarum supplementation in the fecal microbiome of mice. Staphylococcus saprophyticus is a type of coagulase-negative staphylococci that frequently causes urinary tract infections due to its ability to produce urease [77,78]. It can also lead to the development of bacteremia, endophthalmitis, wound infections, eczema, and respiratory tract infections [79]. Peptidoglycan, isolated from staphylococci and streptococci, is also known to cause septic shock in laboratory animals. This is characterized by fever, inflammatory reactions, thrombocytopenia, and multiple-organ dysfunction syndrome [80,81]. Our study suggests the need for a more comprehensive investigation of bacterial interactions within the fecal microbiome of laboratory mice, due to an increase in the number of pathogenic representatives of the Staphylococcus genus when Lactobacillus plantarum is added to the diet.
An increase in some uncharacterized species was observed in the fecal microbiome of mice with Lactobacillus plantarum supplementation. The species included in the study were GGB24134 SGB35937 from the family Lactobacillaceae, GGB27892 SGB40326 from the family Selenomonadaceae, as well as GGB27925 SGB40363 and GGB3583 SGB21441 from the family Muribaculaceae. Also, after the injection of lipopolysaccharide, we observed an increase in the abundance of the uncharacterized species GGB3583 SGB21441, which belongs to the family Muribaculaceae.
Lactobacillaceae is a family of lactic acid bacteria that inhabit the intestinal microbiota of humans and many animals. These bacteria have beneficial effects on gut health, including modulation of the immune system and protection against pathogens. Many species are used as probiotics [82]. Supplementation with Lactobacillus plantarum increased in a previously uncharacterized species from this family. Selenomonadaceae is commonly found in various anaerobic environments and is frequently isolated from the rumen and caeca of mammals [83]. This family is thought to be linked to the development of obesity [84,85]. Some studies demonstrated a negative correlation between the abundance of Selenomonadaceae in the human gut microbiome and Lactobacillus plantarum KY1032 supplementation [86]. Bacteria belonging to the Muribaculaceae family are known to produce propionate as an end product of fermentation and are abundant and diverse in the mouse gut. Although these bacteria are also found in humans, only a few members of this clade have been cultured and described, but their ecological role is still uncharacterized [87,88,89]. The previous study showed that Muribaculaceae tended to increase in abundance when a member of the Lactobacillus genus was added to the diet [90], which we also observed in our study for an uncharacterized member of this genus.
The study of the functional profiles of the microbiome revealed a rich diversity of metabolic pathways in the groups studied, but most of them are not integrated into databases and not mapped. This again suggests that the mouse fecal microbiome is poorly understood. Nevertheless, we observed a difference in the representation of three metabolic profiles for the LP group compared to the control group.
The trans-sulfuration pathway in bacteria is a metabolic process that converts homocysteine to cysteine through the enzymatic action of cystathionine β-synthase (CBS) and cystathionine γ-lyase (CGL) [91]. The initial step in the trans-sulfuration pathway is the condensation of homocysteine and serine catalyzed by cystathionine β-synthase (CBS). Subsequently, cystathionine γ-lyase (CGL) catalyzes the breakdown of cystathionine into cysteine, ammonia (NH3), and α-ketobutyrate [92]. The conversion of homocysteine to cysteine via the reverse trans-sulfuration pathway facilitates the conversion of methionine to cysteine, thereby providing the requisite homocysteine. Recent evidence indicates that the S-ribosylhomocysteinase (LuxS) protein, which functions as a quorum-sensing molecule in numerous bacteria, plays a pivotal role in the reverse trans-sulfuration pathway [93]. The gut microbiota can metabolize sulfur-containing amino acids, which affects homocysteine and cysteine levels. As cysteine is essential for the growth of many bacteria, the trans-sulfuration pathway facilitates the growth of microorganisms without the need for exogenous cysteine. In addition, LuxS functions as a signaling and metabolic molecule in bacteria [94]. It has been observed that dysbiosis (imbalance of the gut microbiota) in IBD patients may disrupt these pathways [94,95,96]. Some gut bacteria are capable of producing hydrogen sulfide (H2S) from cysteine. In moderate quantities, H2S has anti-inflammatory effects; however, in excess, it can be toxic and exacerbate inflammation. Additionally, numerous studies have demonstrated a close correlation between homocysteine and cysteine metabolism in various pathological conditions [97,98]. Consequently, the trans-sulfuration pathway may be implicated in metabolic dysregulation, which may in turn contribute to the development of gastrointestinal pathologies.
The L-histidine degradation pathway was predominant in the LP group compared to the control group. The ability to cleave histidine is widespread and highly conserved among all bacteria. In many cases, the complete catabolism of histidine to glutamate, ammonia, and formate or formamide allows the use of histidine as a carbon or nitrogen source or both. In some cases, cells use intermediates and pathway components to generate important signaling molecules that influence properties such as virulence or biofilm formation. The strong conservation of the enzymes involved and the genetic clustering of histidine utilization pathway genes suggest horizontal transfer over evolutionary time [99,100]. The major contribution to the representation of this pathway was made by unclassified bacteria, but it was also found in the bacteria Parabacteroides distasonis, Staphylococcus nepalensis, Staphylococcus saprophyticus, and Staphylococcus xylosus in the LP group. Parabacteroides distasonis is a member of the intestinal microbiota that has recently received much attention due to the many beneficial properties it confers on its host, manifesting anti-inflammatory properties and activity in reducing body weight gain, hyperglycaemia, and hepatic steatosis [101].

5. Conclusions

The objective of this study was to investigate the changes that occur in the gut microbiome of mice following injections of LPS, which cause inflammation. At the same time, we observed changes in the microbiota, characterized by a significant increase in the abundance of uncharacterized species belonging to Staphylococcaceae and Muribaculaceae, caused by LPS injections.
We also investigated the potential anti-inflammatory effects of some probiotic bacteria as a dietary supplement against LPS injection. No significant changes were observed in the microbiome when probiotics were administered compared to the LPS group that did not receive probiotic correction. However, at the morphological level, positive changes were observed in the gut in the presence of probiotics. Lactobacillus plantarum demonstrated the greatest protective effect and microbiome changes. Additionally, the greatest impact of this probiotic species was indicated by a change in the enrichment of metabolic pathways. A decrease in the enrichment of the homocysteine and cysteine interconversion pathway was observed, accompanied by a simultaneous increase in the L-histidine degradation pathway. In addition, an anti-inflammatory effect was observed in the LA group of mice, which had been administered a commercial probiotic mix of probiotic lactic acid bacteria. This effect was observed at the morphological level, but it was less pronounced than that observed in mice administered with Lactobacillus plantarum. In addition, histological analysis revealed that the Weissella confusa group did not exhibit anti-inflammatory activity, and no significant microbiome alterations were observed.
This study identified microbiome imbalances associated with LPS-induced inflammation and explored how specific probiotic bacteria can improve gut mucosal healing.

Author Contributions

Conceptualization, M.S.; methodology, M.S., A.G. and E.C.; validation, M.G., I.B., M.S. and Y.S.; formal analysis, Y.S.; investigation, I.B., Y.S., P.M., M.G., E.C., O.K. and E.M.; resources, M.S.; data curation, M.S.; writing—original draft preparation, M.G., I.B., Y.S. and P.M.; writing—review and editing, O.K. and M.S.; visualization, M.G. and E.M.; supervision, M.S.; project administration, M.S.; funding acquisition, M.S. All authors have read and agreed to the published version of the manuscript.

Funding

The research was carried out within the state assignment of Ministry of Science and Higher Education of the Russian Federation (project FZGW-2024-0003).

Data Availability Statement

Metagenome data are available in the NCBI BioProject database (BioProject ID: PRJNA1109598).

Acknowledgments

This study is dedicated to the memory of the head of the laboratory of metagenomics and food biotechnology of Voronezh State University of Engineering Technologies Popov V.N., who always helped the laboratory staff in the implementation of their ideas. We also thank Evgeny Popov for providing probiotic bacteria cultures.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Rietschel, E.T.; Kirikae, T.; Schade, F.U.; Mamat, U.; Schmidt, G.; Loppnow, H.; Ulmer, A.J.; Zähringer, U.; Seydel, U.; Di Padova, F.; et al. Bacterial Endotoxin: Molecular Relationships of Structure to Activity and Function. FASEB J. 1994, 8, 217–225. [Google Scholar] [CrossRef]
  2. Kell, D.B.; Pretorius, E. On the Translocation of Bacteria and Their Lipopolysaccharides between Blood and Peripheral Locations in Chronic, Inflammatory Diseases: The Central Roles of LPS and LPS-Induced Cell Death. Integr. Biol. 2015, 7, 1339–1377. [Google Scholar] [CrossRef]
  3. Page, M.J.; Kell, D.B.; Pretorius, E. The Role of Lipopolysaccharide-Induced Cell Signalling in Chronic Inflammation. Chronic Stress 2022, 6, 1–18. [Google Scholar] [CrossRef]
  4. Freudenberg, M.A.; Kalis, C.; Chvatchko, Y.; Merlin, T.; Gumenscheimer, M.; Galanos, C. Role of Interferons in LPS Hypersensitivity. J. Endotoxin Res. 2003, 9, 308–312. [Google Scholar] [CrossRef]
  5. Kulp, A.; Kuehn, M.J. Biological Functions and Biogenesis of Secreted Bacterial Outer Membrane Vesicles. Annu. Rev. Microbiol. 2010, 64, 163–184. [Google Scholar] [CrossRef]
  6. Brown, G.C. The Endotoxin Hypothesis of Neurodegeneration. J. Neuroinflamm. 2019, 16, 180. [Google Scholar] [CrossRef]
  7. Cani, P.D.; Amar, J.; Iglesias, M.A.; Poggi, M.; Knauf, C.; Bastelica, D.; Neyrinck, A.M.; Fava, F.; Tuohy, K.M.; Chabo, C.; et al. Metabolic Endotoxemia Initiates Obesity and Insulin Resistance. Diabetes 2007, 56, 1761–1772. [Google Scholar] [CrossRef]
  8. Wiedermann, C.J.; Kiechl, S.; Dunzendorfer, S.; Schratzberger, P.; Egger, G.; Oberhollenzer, F.; Willeit, J. Association of Endotoxemia with Carotid Atherosclerosis and Cardiovascular Disease: Prospective Results from the Bruneck Study. J. Am. Coll. Cardiol. 1999, 34, 1975–1981. [Google Scholar] [CrossRef]
  9. Baucells, B.J.; Sebastiani, G.; Herrero-Aizpurua, L.; Andreu-Fernández, V.; Navarro-Tapia, E.; García-Algar, O.; Figueras-Aloy, J. Effectiveness of a Probiotic Combination on the Neurodevelopment of the Very Premature Infant. Sci. Rep. 2023, 13, 10344. [Google Scholar] [CrossRef]
  10. Hemarajata, P.; Versalovic, J. Effects of Probiotics on Gut Microbiota: Mechanisms of Intestinal Immunomodulation and Neuromodulation. Ther. Adv. Gastroenterol. 2013, 6, 39. [Google Scholar] [CrossRef]
  11. Venegas, D.P.; De La Fuente, M.K.; Landskron, G.; González, M.J.; Quera, R.; Dijkstra, G.; Harmsen, H.J.M.; Faber, K.N.; Hermoso, M.A. Short Chain Fatty Acids (SCFAs)Mediated Gut Epithelial and Immune Regulation and Its Relevance for Inflammatory Bowel Diseases. Front. Immunol. 2019, 10, 424615. [Google Scholar]
  12. Aghamohammad, S.; Sepehr, A.; Miri, S.T.; Najafi, S.; Pourshafie, M.R.; Rohani, M. Anti-Inflammatory and Immunomodulatory Effects of Lactobacillus Spp. as a Preservative and Therapeutic Agent for IBD Control. Immun. Inflamm. Dis. 2022, 10, e635. [Google Scholar] [CrossRef] [PubMed]
  13. Fairfax, M.R.; Lephart, P.R.; Salimnia, H. Weissella Confusa: Problems with Identification of an Opportunistic Pathogen That Has Been Found in Fermented Foods and Proposed as a Probiotic. Front. Microbiol. 2014, 5, 91780. [Google Scholar] [CrossRef] [PubMed]
  14. Lakra, A.K.; Domdi, L.; Hanjon, G.; Tilwani, Y.M.; Arul, V. Some Probiotic Potential of Weissella Confusa MD1 and Weissella Cibaria MD2 Isolated from Fermented Batter. LWT 2020, 125, 109261. [Google Scholar] [CrossRef]
  15. Candelli, M.; Franza, L.; Pignataro, G.; Ojetti, V.; Covino, M.; Piccioni, A.; Gasbarrini, A.; Franceschi, F. Interaction between Lipopolysaccharide and Gut Microbiota in Inflammatory Bowel Diseases. Int. J. Mol. Sci. 2021, 22, 6242. [Google Scholar] [CrossRef] [PubMed]
  16. Alzahrani, N.A.; Bahaidrah, K.A.; Mansouri, R.A.; Alsufiani, H.M.; Alghamdi, B.S. Investigation of the Optimal Dose for Experimental Lipopolysaccharide-Induced Recognition Memory Impairment: Behavioral and Histological Studies. J. Integr. Neurosci. 2022, 21, 49. [Google Scholar] [CrossRef] [PubMed]
  17. Batista, C.R.A.; Gomes, G.F.; Candelario-Jalil, E.; Fiebich, B.L.; de Oliveira, A.C.P. Lipopolysaccharide-Induced Neuroinflammation as a Bridge to Understand Neurodegeneration. Int. J. Mol. Sci. 2019, 20, 2293. [Google Scholar] [CrossRef] [PubMed]
  18. Burakova, I.; Smirnova, Y.; Gryaznova, M.; Syromyatnikov, M.; Chizhkov, P.; Popov, E.; Popov, V. The Effect of Short-Term Consumption of Lactic Acid Bacteria on the Gut Microbiota in Obese People. Nutrients 2022, 14, 3384. [Google Scholar] [CrossRef]
  19. Gryaznova, M.; Smirnova, Y.; Burakova, I.; Syromyatnikov, M.; Chizhkov, P.; Popov, E.; Popov, V. Changes in the Human Gut Microbiome Caused by the Short-Term Impact of Lactic Acid Bacteria Consumption in Healthy People. Probiot. Antimicrob. Proteins 2023, 1–12. [Google Scholar] [CrossRef]
  20. Ren, D.; Wang, D.; Rong, F.; Liu, H.; Shen, M.; Yu, H. Oral Administration of Lactobacillus Plantarum Attenuates Inflammatory Damage in Mice Challenged with Two Pathogens. Eur. J. Inflamm. 2019, 17, 1–8. [Google Scholar] [CrossRef]
  21. Bao, W.; He, Y.; Yu, J.; Liu, M.; Yang, X.; Ta, N.; Zhang, E.; Liang, C. Regulatory Effect of Lactiplantibacillus Plantarum 2-33 on Intestinal Microbiota of Mice With Antibiotic-Associated Diarrhea. Front. Nutr. 2022, 9, 921875. [Google Scholar] [CrossRef]
  22. Rosshart, S.P.; Herz, J.; Vassallo, B.G.; Hunter, A.; Wall, M.K.; Badger, J.H.; McCulloch, J.A.; Anastasakis, D.G.; Sarshad, A.A.; Leonardi, I.; et al. Laboratory Mice Born to Wild Mice Have Natural Microbiota and Model Human Immune Responses. Science 2019, 365, eaaw4361. [Google Scholar] [CrossRef]
  23. Shin, J.; Lee, S.; Go, M.J.; Lee, S.Y.; Kim, S.C.; Lee, C.H.; Cho, B.K. Analysis of the Mouse Gut Microbiome Using Full-Length 16S RRNA Amplicon Sequencing. Sci. Rep. 2016, 6, 29681. [Google Scholar] [CrossRef]
  24. Costea, P.I.; Zeller, G.; Sunagawa, S.; Pelletier, E.; Alberti, A.; Levenez, F.; Tramontano, M.; Driessen, M.; Hercog, R.; Jung, F.E.; et al. Towards Standards for Human Fecal Sample Processing in Metagenomic Studies. Nat. Biotechnol. 2017, 35, 1069–1076. [Google Scholar] [CrossRef]
  25. GitHub—s-Andrews/FastQC: A Quality Control Analysis Tool for High Throughput Sequencing Data. Available online: https://github.com/s-andrews/FastQC?ysclid=ljfeelfswd703292447 (accessed on 28 June 2023).
  26. Chen, S.; Zhou, Y.; Chen, Y.; Gu, J. Fastp: An Ultra-Fast All-in-One FASTQ Preprocessor. Bioinformatics 2018, 34, i884–i890. [Google Scholar] [CrossRef]
  27. Langmead, B.; Salzberg, S.L. Fast Gapped-Read Alignment with Bowtie 2. Nat. Methods 2012, 9, 357–359. [Google Scholar] [CrossRef]
  28. Blanco-Míguez, A.; Beghini, F.; Cumbo, F.; McIver, L.J.; Thompson, K.N.; Zolfo, M.; Manghi, P.; Dubois, L.; Huang, K.D.; Thomas, A.M.; et al. Extending and Improving Metagenomic Taxonomic Profiling with Uncharacterized Species Using MetaPhlAn 4. Nat. Biotechnol. 2023, 41, 1633–1644. [Google Scholar] [CrossRef]
  29. Beghini, F.; McIver, L.J.; Blanco-Míguez, A.; Dubois, L.; Asnicar, F.; Maharjan, S.; Mailyan, A.; Manghi, P.; Scholz, M.; Thomas, A.M.; et al. Integrating Taxonomic, Functional, and Strain-Level Profiling of Diverse Microbial Communities with BioBakery 3. eLife 2021, 10, e65088. [Google Scholar] [CrossRef]
  30. Wilcoxon, F. Individual Comparisons by Ranking Methods. In Breakthroughs in Statistics; Springer Series in Statistics; Kotz, S., Johnson, N.L., Eds.; Springer: New York, NY, USA, 1992; pp. 196–202. [Google Scholar] [CrossRef]
  31. Mallick, H.; Rahnavard, A.; McIver, L.J.; Ma, S.; Zhang, Y.; Nguyen, L.H.; Tickle, T.L.; Weingart, G.; Ren, B.; Schwager, E.H.; et al. Multivariable Association Discovery in Population-Scale Meta-Omics Studies. PLoS Comput. Biol. 2021, 17, e1009442. [Google Scholar] [CrossRef]
  32. Ng, Q.X.; Yau, C.E.; Yaow, C.Y.L.; Chong, R.I.H.; Chong, N.Z.Y.; Teoh, S.E.; Lim, Y.L.; Soh, A.Y.S.; Ng, W.K.; Thumboo, J. What Has Longitudinal “Omics” Studies Taught Us about Irritable Bowel Syndrome? A Systematic Review. Metabolites 2023, 13, 484. [Google Scholar] [CrossRef]
  33. Rašková, V.; Květoňová, D.; Sak, B.; McEvoy, J.; Edwinson, A.; Stenger, B.; Kváč, M. Human Cryptosporidiosis Caused by Cryptosporidium Tyzzeri and C. Parvum Isolates Presumably Transmitted from Wild Mice. J. Clin. Microbiol. 2013, 51, 360–362. [Google Scholar] [CrossRef]
  34. García-Livia, K.; Martín-Alonso, A.; Foronda, P. Diversity of Cryptosporidium Spp. In Wild Rodents from the Canary Islands, Spain. Parasites Vectors 2020, 13, 445. [Google Scholar] [CrossRef]
  35. Widmer, G.; Köster, P.C.; Carmena, D. Cryptosporidium Hominis Infections in Non-Human Animal Species: Revisiting the Concept of Host Specificity. Int. J. Parasitol. 2020, 50, 253–262. [Google Scholar] [CrossRef]
  36. Maruo, T.; Sakamoto, M.; Ito, C.; Toda, T.; Benno, Y. Adlercreutzia Equolifaciens Gen. Nov., Sp. Nov., an Equol-Producing Bacterium Isolated from Human Faeces, and Emended Description of the Genus Eggerthella. Int. J. Syst. Evol. Microbiol. 2008, 58, 1221–1227. [Google Scholar] [CrossRef]
  37. Shen, L.; Ji, H.F. Reciprocal Interactions between Resveratrol and Gut Microbiota Deepen Our Understanding of Molecular Mechanisms Underlying Its Health Benefits. Trends Food Sci. Technol. 2018, 81, 232–236. [Google Scholar] [CrossRef]
  38. Oñate, F.P.; Chamignon, C.; Burz, S.D.; Lapaque, N.; Monnoye, M.; Philippe, C.; Bredel, M.; Chêne, L.; Farin, W.; Paillarse, J.M.; et al. Adlercreutzia Equolifaciens Is an Anti-Inflammatory Commensal Bacterium with Decreased Abundance in Gut Microbiota of Patients with Metabolic Liver Disease. Int. J. Mol. Sci. 2023, 24, 12232. [Google Scholar] [CrossRef]
  39. Sheng, S.; Chen, J.; Zhang, Y.; Qin, Q.; Li, W.; Yan, S.; Wang, Y.; Li, T.; Gao, X.; Tang, L.; et al. Structural and Functional Alterations of Gut Microbiota in Males With Hyperuricemia and High Levels of Liver Enzymes. Front. Med. 2021, 8, 779994. [Google Scholar] [CrossRef]
  40. Miyake, S.; Ding, Y.; Soh, M.; Seedorf, H. Complete Genome Sequence of Duncaniella Muris Strain B8, Isolated from the Feces of C57/BL6 Mice. Microbiol. Resour. Announc. 2019, 8, e00566-19. [Google Scholar] [CrossRef]
  41. Wenderlein, J.; Böswald, L.F.; Ulrich, S.; Kienzle, E.; Neuhaus, K.; Lagkouvardos, I.; Zenner, C.; Straubinger, R.K. Processing Matters in Nutrient-Matched Laboratory Diets for Mice-Microbiome. Animals 2021, 11, 862. [Google Scholar] [CrossRef]
  42. Wang, H.; Dang, D.; Zhu, L.; Pan, M.; Zhu, J.; Lu, W.; Lu, S.; Zhao, J. Effects of Varied Sulfamethazine Dosage and Exposure Durations on Offspring Mice. Microorganisms 2024, 12, 381. [Google Scholar] [CrossRef]
  43. Ishnaiwer, M. Multimodal Treatment of Intestinal Carriage of Multi-drug Resistant Bacteria with Probiotics and Prebiotics. 2022. Available online: https://books.google.ru/books/about/Multimodal_Treatment_of_Intestinal_Carri.html?id=3q0e0AEACAAJ&redir_esc=y (accessed on 18 March 2024).
  44. Iljazovic, A.; Roy, U.; Gálvez, E.J.C.; Lesker, T.R.; Zhao, B.; Gronow, A.; Amend, L.; Will, S.E.; Hofmann, J.D.; Pils, M.C.; et al. Perturbation of the Gut Microbiome by Prevotella Spp. Enhances Host Susceptibility to Mucosal Inflammation. Mucosal Immunol. 2020, 14, 113–124. [Google Scholar] [CrossRef] [PubMed]
  45. Osuna-Prieto, F.J.; Xu, H.; Ortiz-Alvarez, L.; Di, X.; Kohler, I.; Jurado-Fasoli, L.; Rubio-Lopez, J.; Plaza-Díaz, J.; Vilchez-Vargas, R.; Link, A.; et al. The Relative Abundance of Fecal Bacterial Species Belonging to the Firmicutes and Bacteroidetes Phyla Is Related to Plasma Levels of Bile Acids in Young Adults. Metabolomics 2023, 19, 54. [Google Scholar] [CrossRef]
  46. Johnson, E.L.; Heaver, S.L.; Walters, W.A.; Ley, R.E. Microbiome and Metabolic Disease: Revisiting the Bacterial Phylum Bacteroidetes. J. Mol. Med. 2017, 95, 1–8. [Google Scholar] [CrossRef] [PubMed]
  47. Momose, Y.; Park, S.H.; Miyamoto, Y.; Itoh, K. Design of Species-Specific Oligonucleotide Probes for the Detection of Bacteroides and Parabacteroides by Fluorescence in Situ Hybridization and Their Application to the Analysis of Mouse Caecal Bacteroides-Parabacteroides Microbiota. J. Appl. Microbiol. 2011, 111, 176–184. [Google Scholar] [CrossRef]
  48. Wang, H.; Wang, Q.; Yang, C.; Guo, M.; Cui, X.; Jing, Z.; Liu, Y.; Qiao, W.; Qi, H.; Zhang, H.; et al. Bacteroides Acidifaciens in the Gut Plays a Protective Role against CD95-Mediated Liver Injury. Gut Microbes 2022, 14, 2027853. [Google Scholar] [CrossRef]
  49. Nakajima, A.; Sasaki, T.; Itoh, K.; Kitahara, T.; Takema, Y.; Hiramatsu, K.; Ishikawa, D.; Shibuya, T.; Kobayashi, O.; Osada, T.; et al. A Soluble Fiber Diet Increases Bacteroides Fragilis Group Abundance and Immunoglobulin A Production in the Gut. Appl. Environ. Microbiol. 2020, 86, e00405-20. [Google Scholar] [CrossRef]
  50. Yan, S.; Zhu, Y.; Li, L.; Qin, S.; Yuan, J.; Chang, X.; Hu, S. Alginate Oligosaccharide Ameliorates Azithromycin-Induced Gut Microbiota Disorder via Bacteroides Acidifaciens-FAHFAs and Bacteroides-TCA Cycle Axes. Food Funct. 2023, 14, 427–444. [Google Scholar] [CrossRef] [PubMed]
  51. Zheng, C.; Zhong, Y.; Xie, J.; Wang, Z.; Zhang, W.; Pi, Y.; Zhang, W.; Liu, L.; Luo, J.; Xu, W. Bacteroides Acidifaciens and Its Derived Extracellular Vesicles Improve DSS-Induced Colitis. Front. Microbiol. 2023, 14, 1304232. [Google Scholar] [CrossRef]
  52. Zitomersky, N.L.; Coyne, M.J.; Comstock, L.E. Longitudinal Analysis of the Prevalence, Maintenance, and IgA Response to Species of the Order Bacteroidales in the Human Gut. Infect. Immun. 2011, 79, 2012–2020. [Google Scholar] [CrossRef]
  53. Tan, H.; Zhao, J.; Zhang, H.; Zhai, Q.; Chen, W. Isolation of Low-Abundant Bacteroidales in the Human Intestine and the Analysis of Their Differential Utilization Based on Plant-Derived Polysaccharides. Front. Microbiol. 2018, 9, 1319. [Google Scholar] [CrossRef]
  54. Snydman, D.R.; Jacobus, N.V.; McDermott, L.A.; Golan, Y.; Hecht, D.W.; Goldstein, E.J.C.; Harrell, L.; Jenkins, S.; Newton, D.; Pierson, C.; et al. Lessons Learned from the Anaerobe Survey: Historical Perspective and Review of the Most Recent Data (2005–2007). Clin. Infect. Dis. 2010, 50 (Suppl. S1), S26–S33. [Google Scholar] [CrossRef] [PubMed]
  55. Parapouli, M.; Delbès-Paus, C.; Kakouri, A.; Koukkou, A.I.; Montel, M.C.; Samelis, J. Characterization of a Wild, Novel Nisin A-Producing Lactococcus Strain with an L. Lactis Subsp. Cremoris Genotype and an L. Lactis Subsp. Lactis Phenotype, Isolated from Greek Raw Milk. Appl. Environ. Microbiol. 2013, 79, 3476–3484. [Google Scholar] [CrossRef] [PubMed]
  56. Ballal, S.A.; Veiga, P.; Fenn, K.; Michaud, M.; Kim, J.H.; Gallini, C.A.; Glickman, J.N.; Quéré, G.; Garault, P.; Béal, C.; et al. Host Lysozyme-Mediated Lysis of Lactococcus Lactis Facilitates Delivery of Colitis-Attenuating Superoxide Dismutase to Inflamed Colons. Proc. Natl. Acad. Sci. USA 2015, 112, 7803–7808. [Google Scholar] [CrossRef] [PubMed]
  57. Luerce, T.D.; Gomes-Santos, A.C.; Rocha, C.S.; Moreira, T.G.; Cruz, D.N.; Lemos, L.; Sousa, A.L.; Pereira, V.B.; De Azevedo, M.; Moraes, K.; et al. Anti-Inflammatory Effects of Lactococcus Lactis NCDO 2118 during the Remission Period of Chemically Induced Colitis. Gut Pathog. 2014, 6, 33. [Google Scholar] [CrossRef]
  58. Steidler, L.; Hans, W.; Schotte, L.; Neirynck, S.; Obermeier, F.; Falk, W.; Fiers, W.; Remaut, E. Treatment of Murine Colitis by Lactococcus Lactis Secreting Interleukin-10. Science 2000, 289, 1352–1355. [Google Scholar] [CrossRef] [PubMed]
  59. Antolín, J.; Cigüenza, R.; Salueña, I.; Vázquez, E.; Hernández, J.; Espinós, D. Liver Abscess Caused by Lactococcus Lactis Cremoris: A New Pathogen. Scand. J. Infect. Dis. 2004, 36, 490–491. [Google Scholar] [CrossRef] [PubMed]
  60. Wu, F.; Xie, X.; Du, T.; Jiang, X.; Miao, W.; Wang, T. Lactococcus Lactis, a Bacterium with Probiotic Functions and Pathogenicity. World J. Microbiol. Biotechnol. 2023, 39, 325. [Google Scholar] [CrossRef]
  61. Seedorf, H.; Griffin, N.W.; Ridaura, V.K.; Reyes, A.; Cheng, J.; Rey, F.E.; Smith, M.I.; Simon, G.M.; Scheffrahn, R.H.; Woebken, D.; et al. Bacteria from Diverse Habitats Colonize and Compete in the Mouse Gut. Cell 2014, 159, 253–266. [Google Scholar] [CrossRef] [PubMed]
  62. Cai, Y.; Luo, Y.; Dai, N.; Yang, Y.; He, Y.; Chen, H.; Zhao, M.; Fu, X.; Chen, T.; Xing, Z. Functional Metagenomic and Metabolomics Analysis of Gut Dysbiosis Induced by Hyperoxia. Front. Microbiol. 2023, 14, 1197970. [Google Scholar] [CrossRef]
  63. Yue, X.; Zhou, H.; Wang, S.; Chen, X.; Xiao, H. Gut Microbiota, Microbiota-Derived Metabolites, and Graft-versus-Host Disease. Cancer Med. 2024, 13, e6799. [Google Scholar] [CrossRef]
  64. Almeida, A.; Mitchell, A.L.; Boland, M.; Forster, S.C.; Gloor, G.B.; Tarkowska, A.; Lawley, T.D.; Finn, R.D. A New Genomic Blueprint of the Human Gut Microbiota. Nature 2019, 568, 499–504. [Google Scholar] [CrossRef] [PubMed]
  65. Raimondi, S.; Musmeci, E.; Candeliere, F.; Amaretti, A.; Rossi, M. Identification of Mucin Degraders of the Human Gut Microbiota. Sci. Rep. 2021, 11, 11094. [Google Scholar] [CrossRef] [PubMed]
  66. Jin, D.; Wu, S.; Zhang, Y.G.; Lu, R.; Xia, Y.; Dong, H.; Sun, J. Lack of Vitamin D Receptor Causes Dysbiosis and Changes the Functions of the Murine Intestinal Microbiome. Clin. Ther. 2015, 37, 996–1009.e7. [Google Scholar] [CrossRef] [PubMed]
  67. Sisk-Hackworth, L.; Brown, J.; Sau, L.; Levine, A.A.; Tam, L.Y.I.; Ramesh, A.; Shah, R.S.; Kelley-Thackray, E.T.; Wang, S.; Nguyen, A.; et al. Genetic Hypogonadal Mouse Model Reveals Niche-Specific Influence of Reproductive Axis and Sex on Intestinal Microbial Communities. Biol. Sex Differ. 2023, 14, 79. [Google Scholar] [CrossRef] [PubMed]
  68. Selma, M.V.; Beltrán, D.; Luna, M.C.; Romo-Vaquero, M.; García-Villalba, R.; Mira, A.; Espín, J.C.; Tomás-Barberán, F.A. Isolation of Human Intestinal Bacteria Capable of Producing the Bioactive Metabolite Isourolithin A from Ellagic Acid. Front. Microbiol. 2017, 8, 1521. [Google Scholar] [CrossRef] [PubMed]
  69. Singh, R.; Chandrashekharappa, S.; Bodduluri, S.R.; Baby, B.V.; Hegde, B.; Kotla, N.G.; Hiwale, A.A.; Saiyed, T.; Patel, P.; Vijay-Kumar, M.; et al. Enhancement of the Gut Barrier Integrity by a Microbial Metabolite through the Nrf2 Pathway. Nat. Commun. 2019, 10, 89. [Google Scholar] [CrossRef] [PubMed]
  70. Wu, J.; Liu, M.; Zhou, M.; Wu, L.; Yang, H.; Huang, L.; Chen, C. Isolation and Genomic Characterization of Five Novel Strains of Erysipelotrichaceae from Commercial Pigs. BMC Microbiol. 2021, 21, 125. [Google Scholar] [CrossRef]
  71. Chen, W.; Liu, F.; Ling, Z.; Tong, X.; Xiang, C. Human Intestinal Lumen and Mucosa-Associated Microbiota in Patients with Colorectal Cancer. PLoS ONE 2012, 7, e39743. [Google Scholar] [CrossRef] [PubMed]
  72. Schaubeck, M.; Clavel, T.; Calasan, J.; Lagkouvardos, I.; Haange, S.B.; Jehmlich, N.; Basic, M.; Dupont, A.; Hornef, M.; Von Bergen, M.; et al. Dysbiotic Gut Microbiota Causes Transmissible Crohn’s Disease-like Ileitis Independent of Failure in Antimicrobial Defence. Gut 2016, 65, 225–237. [Google Scholar] [CrossRef]
  73. Labbé, A.; Ganopolsky, J.G.; Martoni, C.J.; Prakash, S.; Jones, M.L. Bacterial Bile Metabolising Gene Abundance in Crohn’s, Ulcerative Colitis and Type 2 Diabetes Metagenomes. PLoS ONE 2014, 9, e115175. [Google Scholar] [CrossRef]
  74. Ghimire, K.; Kasarla, R.R. Staphylococcus Nepalensis: A New Species from Nepal. J. Univers. Coll. Med. Sci. 2023, 11, 69–72. [Google Scholar] [CrossRef]
  75. Nováková, D.; Pantůček, R.; Petráš, P.; Koukalová, D.; Sedláček, I. Occurance of Staphylococcus Nepalensis Strains in Different Sources Including Human Clinical Material. FEMS Microbiol. Lett. 2006, 263, 163–168. [Google Scholar] [CrossRef] [PubMed]
  76. Andrade-Oliveira, A.L.; Lacerda-Rodrigues, G.; Pereira, M.F.; Bahia, A.C.; Machado, E.d.A.; Rossi, C.C.; Giambiagi-deMarval, M. Tenebrio Molitor as a Model System to Study Staphylococcus Spp Virulence and Horizontal Gene Transfer. Microb. Pathog. 2023, 183, 106304. [Google Scholar] [CrossRef]
  77. Kuroda, M.; Yamashita, A.; Hirakawa, H.; Kumano, M.; Morikawa, K.; Higashide, M.; Maruyama, A.; Inose, Y.; Matoba, K.; Toh, H.; et al. Whole Genome Sequence of Staphylococcus Saprophyticus Reveals the Pathogenesis of Uncomplicated Urinary Tract Infection. Proc. Natl. Acad. Sci. USA 2005, 102, 13272–13277. [Google Scholar] [CrossRef] [PubMed]
  78. Rupp, M.E.; Soper, D.E.; Archer, G.L. Colonization of the Female Genital Tract with Staphylococcus Saprophyticus. J. Clin. Microbiol. 1992, 30, 2975–2979. [Google Scholar] [CrossRef] [PubMed]
  79. Al-Mathkhury, H.J.F.; Flaih, M.T.; Fleih, A.S. Role of Peptidoglycan in the Pathogenesis of Staphylococcus Saprophyticus in Mice. Baghdad Sci. J. 2021, 4, 597–602. [Google Scholar] [CrossRef]
  80. Bera, A.; Biswas, R.; Herbert, S.; Götz, F. The Presence of Peptidoglycan O-Acetyltransferase in Various Staphylococcal Species Correlates with Lysozyme Resistance and Pathogenicity. Infect. Immun. 2006, 74, 4598–4604. [Google Scholar] [CrossRef]
  81. Myhre, A.E.; Stuestøl, J.F.; Dahle, M.K.; Øverland, G.; Thiemermann, C.; Foster, S.J.; Lilleaasen, P.; Aasen, A.O.; Wang, J.E. Organ Injury and Cytokine Release Caused by Peptidoglycan Are Dependent on the Structural Integrity of the Glycan Chain. Infect. Immun. 2004, 72, 1311–1317. [Google Scholar] [CrossRef]
  82. Huynh, U.; Zastrow, M.L. Metallobiology of Lactobacillaceae in the Gut Microbiome. J. Inorg. Biochem. 2023, 238, 112023. [Google Scholar] [CrossRef]
  83. Zhang, K.; Dong, X. Selenomonas Bovis Sp. Nov., Isolated from Yak Rumen Contents. Int. J. Syst. Evol. Microbiol. 2009, 59, 2080–2083. [Google Scholar] [CrossRef]
  84. Jeelani, A.; Afroz Ahmed, S.; Vaqar Momin, F. Obesity-Caused by a Germ? Int. J. Sci. Res. Publ. 2013, 3, 1–3. Available online: http://www.ijsrp.org/research-paper-1301.php?rp=P13639 (accessed on 27 June 2024).
  85. Harakeh, S.M.; Khan, I.; Kumosani, T.; Barbour, E.; Almasaudi, S.B.; Bahijri, S.M.; Alfadul, S.M.; Ajabnoor, G.M.A.; Azhar, E.I. Gut Microbiota: A Contributing Factor to Obesity. Front. Cell. Infect. Microbiol. 2016, 6, 95. [Google Scholar] [CrossRef] [PubMed]
  86. Mo, S.J.; Lee, K.; Hong, H.J.; Hong, D.K.; Jung, S.H.; Park, S.D.; Shim, J.J.; Lee, J.L. Effects of Lactobacillus Curvatus HY7601 and Lactobacillus Plantarum KY1032 on Overweight and the Gut Microbiota in Humans: Randomized, Double-Blinded, Placebo-Controlled Clinical Trial. Nutrients 2022, 14, 2484. [Google Scholar] [CrossRef] [PubMed]
  87. Lagkouvardos, I.; Pukall, R.; Abt, B.; Foesel, B.U.; Meier-Kolthoff, J.P.; Kumar, N.; Bresciani, A.; Martínez, I.; Just, S.; Ziegler, C.; et al. The Mouse Intestinal Bacterial Collection (MiBC) Provides Host-Specific Insight into Cultured Diversity and Functional Potential of the Gut Microbiota. Nat. Microbiol. 2016, 1, 16131. [Google Scholar] [CrossRef] [PubMed]
  88. Lagkouvardos, I.; Lesker, T.R.; Hitch, T.C.A.; Gálvez, E.J.C.; Smit, N.; Neuhaus, K.; Wang, J.; Baines, J.F.; Abt, B.; Stecher, B.; et al. Sequence and Cultivation Study of Muribaculaceae Reveals Novel Species, Host Preference, and Functional Potential of This yet Undescribed Family. Microbiome 2019, 7, 28. [Google Scholar] [CrossRef]
  89. Smith, B.J.; Miller, R.A.; Schmidt, T.M. Muribaculaceae Genomes Assembled from Metagenomes Suggest Genetic Drivers of Differential Response to Acarbose Treatment in Mice. mSphere 2021, 6, e0085121. [Google Scholar] [CrossRef] [PubMed]
  90. Zhong, X.; Zhao, Y.; Huang, L.; Liu, J.; Wang, K.; Gao, X.; Zhao, X.; Wang, X. Remodeling of the Gut Microbiome by Lactobacillus Johnsonii Alleviates the Development of Acute Myocardial Infarction. Front. Microbiol. 2023, 14, 1140498. [Google Scholar] [CrossRef] [PubMed]
  91. Fontecave, M.; Atta, M.; Mulliez, E. S-Adenosylmethionine: Nothing Goes to Waste. Trends Biochem. Sci. 2004, 29, 243–249. [Google Scholar] [CrossRef] [PubMed]
  92. Boukaba, A.; Sanchis-Gomar, F.; García-Giménez, J.L. Epigenetic Mechanisms as Key Regulators in Disease: Clinical Implications. In Epigenetic Biomarkers and Diagnostics; Academic Press: Cambridge, MA, USA, 2016; pp. 37–66. [Google Scholar] [CrossRef]
  93. Thomas, D.; Surdin-Kerjan, Y. Metabolism of Sulfur Amino Acids in Saccharomyces Cerevisiae. Microbiol. Mol. Biol. Rev. 1997, 61, 503–532. [Google Scholar] [CrossRef]
  94. Buret, A.G.; Allain, T.; Motta, J.P.; Wallace, J.L. Effects of Hydrogen Sulfide on the Microbiome: From Toxicity to Therapy. Antioxid. Redox Signal. 2022, 36, 211–219. [Google Scholar] [CrossRef]
  95. Frank, D.N.; St. Amand, A.L.; Feldman, R.A.; Boedeker, E.C.; Harpaz, N.; Pace, N.R. Molecular-Phylogenetic Characterization of Microbial Community Imbalances in Human Inflammatory Bowel Diseases. Proc. Natl. Acad. Sci. USA 2007, 104, 13780–13785. [Google Scholar] [CrossRef] [PubMed]
  96. Wallace, J.L.; Wang, R. Hydrogen Sulfide-Based Therapeutics: Exploiting a Unique but Ubiquitous Gasotransmitter. Nat. Rev. Drug Discov. 2015, 14, 329–345. [Google Scholar] [CrossRef] [PubMed]
  97. Chung, K.K.K.; Dawson, V.L.; Dawson, T.M. S-Nitrosylation in Parkinson’s Disease and Related Neurodegenerative Disorders. Methods Enzymol. 2005, 396, 139–150. [Google Scholar] [CrossRef] [PubMed]
  98. Müller, T. Catechol-O-Methyltransferase Enzyme: Cofactor S-Adenosyl-l-Methionine And Related Mechanisms. Int. Rev. Neurobiol. 2010, 95, 49–71. [Google Scholar] [CrossRef] [PubMed]
  99. Nieuwkoop, A.J.; Bender, R.A. Histidine Degradation in Bacteria. Handb. Microb. Metab. Amin. acids 2017, 1, 291–303. [Google Scholar] [CrossRef]
  100. Bender, R.A. Regulation of the Histidine Utilization (Hut) System in Bacteria. Microbiol. Mol. Biol. Rev. 2012, 76, 565–584. [Google Scholar] [CrossRef]
  101. Chamarande, J.; Cunat, L.; Pavlov, N.; Alauzet, C.; Cailliez-Grimal, C. Parabacteroides Distasonis Properties Linked to the Selection of New Biotherapeutics. Nutrients 2022, 14, 4176. [Google Scholar] [CrossRef]
Microorganisms 12 01341 g001
Figure 1. Morphological structure of a mouse intestine. Magnification: (A,C,D) 400×, (B,E) 100×. Hematoxylin–eosin staining: (A) control group, 1—villi, 2—crypts; (B) LPS group, 3—tissue infiltration, 4—crypts; (C) LP group, 5—villi, 6—vessel; (D) W group, 7—villi, 8—crypts; (E) LA group, 9—desquamation of tissue, 10—crypts.
Figure 1. Morphological structure of a mouse intestine. Magnification: (A,C,D) 400×, (B,E) 100×. Hematoxylin–eosin staining: (A) control group, 1—villi, 2—crypts; (B) LPS group, 3—tissue infiltration, 4—crypts; (C) LP group, 5—villi, 6—vessel; (D) W group, 7—villi, 8—crypts; (E) LA group, 9—desquamation of tissue, 10—crypts.
Microorganisms 12 01341 g001
Microorganisms 12 01341 g002
Figure 2. Bacterial composition of the studied samples at the species level.
Figure 2. Bacterial composition of the studied samples at the species level.
Microorganisms 12 01341 g002
Microorganisms 12 01341 g003
Figure 3. Microbiome alpha diversity of study groups.
Figure 3. Microbiome alpha diversity of study groups.
Microorganisms 12 01341 g003
Microorganisms 12 01341 g004
Figure 4. Principal coordinate analysis (PCoA) plot of the beta diversity based on the Bray–Curtis dissimilarity index.
Figure 4. Principal coordinate analysis (PCoA) plot of the beta diversity based on the Bray–Curtis dissimilarity index.
Microorganisms 12 01341 g004
Microorganisms 12 01341 g005
Figure 5. Differences in the composition of the fecal microbiome between the groups studied. * p < 0.05, ** p < 0.01, *** p < 0.001.
Figure 5. Differences in the composition of the fecal microbiome between the groups studied. * p < 0.05, ** p < 0.01, *** p < 0.001.
Microorganisms 12 01341 g005
Microorganisms 12 01341 g006
Figure 6. Differences in metabolic pathway abundance between study groups. * p < 0.05.
Figure 6. Differences in metabolic pathway abundance between study groups. * p < 0.05.
Microorganisms 12 01341 g006
Table 1. Mouse small intestine morphometric study.
Table 1. Mouse small intestine morphometric study.
Metrics, MicronsControlLPSLPWLA
Villi height144.23 ± 23.21121.24 ± 13.27102.27 ± 22.28 **143.24 ± 10.46131.37 ± 12.89
Villus thickness12.34 ± 2.2828.72 ± 2.2512.37 ± 1.2314.33 ± 0.2518.24 ± 1.47
Crypt depth62.89 ± 12.5935.66 ± 7.6557.81 ± 4.5658.43 ± 3.7755.67 ± 10.26
Crypt width17.64 ± 0.5514.67 ± 1.2118.25 ± 4.7517.01 ± 2.8725.49 ± 2.64
Brush border height0.87 ± 0,920.47 ± 0.440,85 ± 3,770.82 ± 0.320.52 ± 0.26
** p < 0.01. Statistically significant differences were found only compared to the control group.
Table 2. Fecal bacteria numbers and their diversity measures.
Table 2. Fecal bacteria numbers and their diversity measures.
Feed TreatmentsObserved SpeciesShannon Index
Control2763.15
LPS2953.21
LP3013.41
W3273.28
LA2693.12
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Gryaznova, M.; Burakova, I.; Smirnova, Y.; Morozova, P.; Chirkin, E.; Gureev, A.; Mikhaylov, E.; Korneeva, O.; Syromyatnikov, M. Effect of Probiotic Bacteria on the Gut Microbiome of Mice with Lipopolysaccharide-Induced Inflammation. Microorganisms 2024, 12, 1341. https://doi.org/10.3390/microorganisms12071341

AMA Style

Gryaznova M, Burakova I, Smirnova Y, Morozova P, Chirkin E, Gureev A, Mikhaylov E, Korneeva O, Syromyatnikov M. Effect of Probiotic Bacteria on the Gut Microbiome of Mice with Lipopolysaccharide-Induced Inflammation. Microorganisms. 2024; 12(7):1341. https://doi.org/10.3390/microorganisms12071341

Chicago/Turabian Style

Gryaznova, Mariya, Inna Burakova, Yuliya Smirnova, Polina Morozova, Egor Chirkin, Artem Gureev, Evgeny Mikhaylov, Olga Korneeva, and Mikhail Syromyatnikov. 2024. "Effect of Probiotic Bacteria on the Gut Microbiome of Mice with Lipopolysaccharide-Induced Inflammation" Microorganisms 12, no. 7: 1341. https://doi.org/10.3390/microorganisms12071341

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Article metric data becomes available approximately 24 hours after publication online.
Back to TopTop