Abstract
D-Allose, a C3 epimer of D-glucose, has potential to improve human health as a functional food. However, its effect on the intestinal environment remains unknown. Aged humans progressively express changes in the gut, some of which deleteriously affect gastrointestinal health. In this study, we profiled the intestinal microbiome in aged mice and analyzed organic acids produced by bacteria in cecum contents after long-term ingestion of D-allose. D-Allose did not significantly change organic acid concentration. However, long-term ingestion did significantly increase the relative abundance of Actinobacteria and reduce the relative abundance of Proteobacteria. These results suggest that oral D-allose improves the proportion of favorable intestinal flora in aged mice. D-Allose significantly decreased the relative abundance of Lachnospiraceae bacteria, but increased the relative abundance of Bacteroides acidifaciens and Akkermansia muciniphila. Thus, D-allose might serve as a nutraceutical capable of improving the balance of gut microbiome during aging.
Keywords: D-allose, intestinal microbiome, organic acid, Bacteroides acidifaciens, Akkermansia muciniphila
Abbreviations
CNS, Central Nervous System; RDP, Ribosomal Database Project.
D-Allose is a rare sugar. Rare sugars are monosaccharides that occur in limited quantities in nature.1) D-Allose is a C3-epimer of D-glucose,2) and as with all rare sugars, is expected to have potential as a functional food.2) In rats3) and humans,4) D-allose is largely absorbed via a conventional sodium-dependent glucose cotransporter 15) after ingestion and then is rapidly excreted into the urine.
Absorbed D-allose has some health benefits and disease modification properties.2) Its anti-hypertensive effect appears to occur through the modulation of glucose metabolism,6) and its anti-osteoporosis effect may occur by inhibiting osteoclast differentiation, as demonstrated in an in vitro study.7) D-Allose also appears to have antioxidative effects, as it has been demonstrated to suppress reactive oxygen species (ROS) production.8) The mechanisms underlying its antioxidative abilities may also contribute to its anti-hypertension and anti-osteoporosis effects.6),7) These beneficial effects led us to hypothesize that D-allose could potentially be a health-promoting dietary supplement. Intriguingly, majority of ingested D-allose is absorbed in the small intestine, and unabsorbed D-allose flows into the large intestine.3),4) The unabsorbed D-allose may act on the intestinal microbiome or intestinal epithelial cells and thereby play a physiological role. However, the effects of D-allose on the intestinal environment have yet to be determined.
Intestinal microbial flora comprises bacteria belonging to mainly four phyla: Firmicutes, Bacteroidetes, Actinobacteria, and Proteobacteria.9) The balance and proportion of colonizing species in the intestines influence overall health.10) For instance, maintaining a good microbiota balance and possessing an abundance of Actinobacteria are believed to contribute to a healthy intestinal environment.11) The proportion of Bifidobacterium species, which include beneficial bacteria of the phylum Actinobacteria, decreases with aging.11) Bacteroidetes species influence intestinal immunity and weight maintenance.12) The beneficial bacteria in the Actinobacteria and Bacteroidetes phyla produce organic acids that improve the intestinal environment and help to maintain good health.13) On the other hand, an increase in the relative abundance of bacteria of the phylum Firmicutes appears to be associated with obesity.14)
In the present study, we examined the effect of long-term oral D-allose administration on age-related changes in the intestinal environment. Cecal-microbiome profile and concentrations of selected organic acids were evaluated in aged mice (22 months old) after oral administration of D-allose for 10 months.
EXPERIMENTAL
All animal experiments were approved by the Animal Experiment Committee of the Tokyo Metropolitan Institute of Gerontology (Animal Protocol Approval no. 17012 and 20018) and by the Matsutani Chemical Industry Co., Ltd. (Animal Protocol Approval No. 200918). The experiments were performed in accordance with the Guide for the Care and Use of Laboratory Animals (National Research Council [United States] Committee for the Update of the Guide for the Care and Use of Laboratory Animals, 2011).
One-month-old male C57BL/6J mice were obtained from CLEA Japan, Inc. (Tokyo, Japan). Mice were housed under specific pathogen-free conditions in groups of 5 per cage; cages were lined with paper chip bedding (Japan SLC, Inc., Shizuoka, Japan). The vivarium was maintained at 22 ± 1 °C and 55 ± 5 % humidity under a 12/12-h light-dark cycle. At 12 months of age, the mice were assigned to either the control diet (10 mice) or D-allose diet group (12 mice), in such a way that the group's mean body weights were similar. The initial body weights of the control group and D-allose group were 39.7±1.3 g and 39.4±1.6 g, respectively (t (19.8) = 0.03, p = 0.976). The control group received regular mouse feed (CRF-1, Oriental Yeast Ltd., Tokyo, Japan), and the D-allose diet group received CRF-1 comprising 3 % (w/w) of D-allose (Matsutani Chemical Industry Co., Ltd., Hyogo, Japan). Animals had ad libitum access to water for 10 months. The dose of D-allose fed to mice in the D-allose diet group was based on a previous toxicity study using rats, which calculated the D-allose concentration for LD50.15) At 22 months of age, the mice were euthanized with isoflurane, and cecal contents were collected and immediately frozen (-80 °C) until analysis. The main outcomes variables we calculated were the type and abundance of bacteria present in the cecum for both groups of mice, and the organic acids present and their abundance.
DNA from thawed individual cecal samples was extracted using a previously described method.16) This method uses a fully automated DNA isolation system (GENE PREP STAR PI-480; Kurabo, Osaka, Japan). The V3-V4 regions of bacterial 16S rRNA were amplified using PCR.17),18) Then, the amplified fragments were paired-end sequenced on a 2 × 284-bp cycle using a MiSeq system with MiSeq Reagent Kit version 3 (600 cycle) chemistry (Illumina, San Diego, CA, USA). The paired-end sequencing reads were merged using the fastq-join program set at default settings.19) Only joined reads with a quality value score ≥ 20 for more than 99 % of the sequence were extracted using the FASTX-Toolkit.20) Chimeric sequences were deleted using the usearch6.1 tool.21) The taxonomy of the sequence reads was assigned using the Ribosomal Database Project (RDP) Multiclassifier tool (ver. 2.11) and the TechnoSurugaLab microbial identification database DB-BA13.0.20),22) The Multiclassifier tool is available from the RDP website (http://rdp.cme.msu.edu/classifier/).23) Bacterial identification was performed using Metagenome@KIN ver. 2.2.1 analysis software (World Fusion; Tokyo, Japan). Bacterial identification at the phylum level was performed using the RDP Multiclassifier tool (confidence, ≥ 0.8). Identification of bacterial genera and species was made possible by using data from the TechnoSuruga Lab microbial identification database DB-BA13.0 (homology, ≥ 97 %). For each taxonomic level of bacteria identified, we calculated the average values of the relative abundance of the identified taxon and the fold change for each group.24)
Organic acid content analysis was performed as described previously.25) Briefly, 0.1 g of the collected cecal sample was mixed with zirconia beads and 2 mL of Milli-Q Water. The samples were heated at 85 °C for 15 min, vortexed at 5 m/s for 45 s using FastPrep 24 5G (MP Biomedicals, LLC, Irvine, CA, USA), and centrifuged at 15,350 × G for 10 min at 24 °C. The resulting supernatant was filtered through a 0.2-µm polytetrafluoroethylene membrane. Select organic acids (acetic acid, propionic acid, butyric acid, succinic acid, and lactic acid) from the filtered cecum samples were analyzed using high-performance liquid chromatography (Prominence HPLC System; Shimadzu Co., Kyoto, Japan) with two tandem columns (Shim-pack SCR-102(H), Φ8 mm × 300 mm; Shimadzu). The eluted organic acids were detected with a post-column pH Buffered Electric Conductivity Detector (CDD-10A; Shimadzu).
All data are presented as means ± standard error of the mean (SEM). Statistical significance of group differences was evaluated using the two-sided Welch's t-test.26) Statistical results showed the degree of freedom, test statistic, and p-value. All statistical analyses were done with version 27 SPSS software (IBM Corp., Armonk, NY, USA). P < 0.05 was considered statistically significant.
No significant increase in body and cecum weight.
The final body weights of the control group and D-allose group after 10 months administration were 50.5 ± 1.7 g and 46.9 ± 2.1 g, respectively (t (18.8) = 1.12, p = 0.278). The cecum weights of the control group and D-allose group were 0.819 ± 0.055 g and 0.809 ± 0.046 g (t (18.5) = 0.14, p = 0.888).
Favorable alteration in microbiome.
Our analyses revealed several significant findings in aged mice after long-term ingestion of D-allose. At the phylum level, compared to the control group, the abundance of Firmicutes and Bacteroidetes did not change significantly after D-allose administration: Firmicutes (t (17.3) = 1.82, p = 0.086); Bacteroidetes (t (18.2) = 1.91, p = 0.072). Statistical difference in the Firmicutes / Bacteroidetes ratio, which is known to increase in obesity, also fell short of significance (the control group and D-allose group were 4.33 ± 0.86 and 2.81 ± 0.40, respectively (t (12.8) = 1.59, p = 0.136). However, the D-allose group of mice had a significantly higher relative abundance of Actinobacteria (t (14.5) = 2.42, p < 0.05) (Table 1), including the beneficial probiotic bacteria Bifidobacterium. In humans, the relative abundance of Actinobacteria decreases with increasing age.11) By contrast in aged mice, the D-allose group had a significantly lower relative abundance of Proteobacteria (t (17.5) = 2.94, p < 0.01) (Table 1), including Escherichia, Salmonella, and Vibrio.9) In both humans and C57BL/6J mice, Proteobacteria, which cause inflammation in the intestines, increases with aging.11),27) Our results indicate that D-allose could appear to improve intestinal microbiome balance at the phylum level in aged mice.
Table 1. Relative abundance of detected bacteria in aged mouse cecum after long-term ingestion of D-allose: Phylum level.
| Rank† | Phylum | Relative abundance (%) | Fold-change†† | |
| Control | D-Allose | |||
| 1 | Firmicutes | 74.4±3.0 | 66.7±2.4 | 0.897 |
| 2 | Bacteroidetes | 21.2±2.6 | 28.5±2.2 | 1.34 |
| 3 | Actinobacteria＊ | 1.38±0.24 | 3.09±0.60 | 2.24 |
| 4 | Proteobacteria＊＊ | 2.45±0.38 | 1.05±0.28 | 0.429 |
| 5 | Verrucomicrobia | 0.0132±0.013 | 0.494±0.26 | 37.4 |
| － | Other Phyla | 0.635±0.11 | 0.205±0.15 | － |
Control group (10 mice) and D-allose group (12 mice). Mean values ± SEM; ＊p < 0.05, ＊＊ p < 0.01 by Welch's t-test.
†Rank from highest to lowest percentage in total of control and D-allose group. ††Quantity of change between the D-allose group and control group (i.e., D-Allose ÷ Control).
Genus level.
The results of our analyses of the aged intestinal microbiome at the genus level are shown in Table 2. The D-allose group had a significantly lower relative abundance of g_Lachnospiraceae bacterium KNHs209_incertae_sedis (t (14.1) = 2.46, p < 0.05) and Blautia (t (11.3) = 3.39, p < 0.01). The presence of Lachnospiraceae bacteria in the human gut is associated with the development of metabolic syndrome, obesity, diabetes, liver disease, inflammatory bowel disease, and chronic kidney disease.28) While at the same time, Lachnospiraceae produces butyric acid that is involved in the improvement of atherosclerosis, maturation of the immune system, host histone epigenetic states, and fatty acid metabolism in colonic epithelial cells.28) Blautia bacteria have some beneficial effects in producing butyric acid and acetic acid in a host gut. On the other hand, Blautia bacteria are correlated with various physiological dysfunctions (e.g., obesity, diabetes, cancer, and various inflammatory diseases).29) Because of its dual nature, the reduction of Blautia bacteria after D-allose ingestion cannot be generalized to include all bacteria in the genus. Overall, our results suggest that D-allose might contribute to the above-mentioned physiological functions partially by altering the intestinal microbiome at the genus level. The D-allose group also had a significantly lower relative abundance of one kind of Acetatifactor bacteria (t (14.7) = 2.35, p < 0.05), specifically Acetatifactor muris (see Table 3).30)
Table 2. Relative abundance of detected bacteria in aged mouse cecum after long-term ingestion of D-allose: Genus level.
| Rank† | Genus | Relative abundance (%) | Fold-change†† | |
| Control | D-Allose | |||
| 1 | Lactobacillus | 26.9±4.6 | 24.8±5.4 | 0.922 |
| 2 | Bacteroides | 20.1±1.3 | 28.5±3.8 | 1.42 |
| 3 | g_Lachnospiraceae bacterium KNHs209_incertae_sedis＊ | 18.0±3.2 | 9.00±1.8 | 0.500 |
| 4 | Faecalibaculum | 7.37±2.0 | 7.11±3.4 | 0.964 |
| 5 | Bifidobacterium | 3.87±1.1 | 6.61±1.5 | 1.71 |
| 6 | Roseburia | 2.53±0.64 | 3.76±2.1 | 1.49 |
| 7 | Acutalibacter | 3.37±0.80 | 2.14±0.67 | 0.635 |
| 8 | Lachnoclostridium | 1.75±0.26 | 3.06±0.94 | 1.75 |
| 9 | Flintibacter | 2.45±0.24 | 1.82±0.48 | 0.743 |
| 10 | Enterorhabdus | 1.66±0.16 | 2.06±0.52 | 1.24 |
| 11 | Muribaculum | 1.85±0.27 | 1.69±0.19 | 0.914 |
| 12 | Clostridium | 0.199±0.13 | 2.28±1.0 | 11.5 |
| 13 | Blautia ＊＊ | 1.92±0.40 | 0.479±0.14 | 0.249 |
| 14 | Akkermansia | 0.0461±0.045 | 1.54±0.83 | 33.4 |
| 15 | Acetatifactor ＊ | 1.21±0.31 | 0.358±0.18 | 0.296 |
| 16 | Muricomes | 0.262±0.043 | 0.541±0.16 | 2.06 |
| 17 | Jeotgalicoccus | 0.240±0.053 | 0.388±0.11 | 1.62 |
| 18 | Harryflintia | 0.253±0.052 | 0.372±0.092 | 1.47 |
| 19 | Escherichia | 0.397±0.33 | 0.240±0.13 | 0.605 |
| 20 | Anaerotaenia | 0.376±0.17 | 0.371±0.13 | 0.987 |
| － | Classified others | 3.81±1.1 | 2.69±0.54 | － |
| － | Unclassified others | 1.24±0.22 | 0.625±0.14 | － |
Same conventions as in Table 1.
Table 3. Relative abundance of detected bacteria in aged mouse cecum after long-term ingestion of D-allose: Species level.
| Rank† | Species | Relative abundance (%) | Fold-change†† | |
| Control | D-Allose | |||
| 1 | Bacteroides acidifaciens ＊＊ | 8.17±0.98 | 17.9±3.0 | 2.19 |
| 2 | Lactobacillus gasseri | 10.4±3.9 | 13.1±3.8 | 1.26 |
| 3 | Faecalibaculum rodentium | 7.42±2.0 | 7.09±3.4 | 0.956 |
| 4 | Bacteroides caecimuris ＊＊ | 8.32±0.72 | 4.14±0.97 | 0.496 |
| 5 | Lactobacillus reuteri | 5.43±1.8 | 4.81±1.8 | 0.886 |
| 6 | Bifidobacterium pseudolongum | 3.42±1.1 | 6.40±1.5 | 1.87 |
| 7 | Roseburia faecis | 2.54±0.64 | 3.34±2.2 | 1.30 |
| 8 | Bacteroides sartorii | 2.99±0.36 | 2.39±0.54 | 0.799 |
| 9 | Acutalibacter muris | 3.38±0.80 | 1.50±0.41 | 0.443 |
| 10 | Flintibacter butyricus | 2.45±0.24 | 1.74±0.50 | 0.710 |
| 11 | Clostridium scindens | 1.41±0.26 | 1.95±0.86 | 1.38 |
| 12 | Muribaculum intestinale | 1.85±0.27 | 1.51±0.23 | 0.816 |
| 13 | Acidaminococcus intestini | 0.0100±0.0067 | 2.12±2.1 | 212 |
| 14 | Akkermansia muciniphila ＊ | 0.0458±0.045 | 1.95±0.86 | 42.6 |
| 15 | Ruminococcus gnavus ＊ | 1.54±0.41 | 0.391±0.13 | 0.254 |
| 16 | Acetatifactor muris ＊ | 1.21±0.31 | 0.212±0.11 | 0.175 |
| 17 | Enterorhabdus muris | 0.677±0.10 | 0.488±0.14 | 0.721 |
| 18 | Actinomyces meyeri | 0.00114±0.0011 | 1.10±1.1 | 965 |
| 19 | Enterorhabdus mucosicola | 0.618±0.066 | 0.527±0.16 | 0.853 |
| 20 | Enterorhabdus caecimuris | 0.361±0.10 | 0.683±0.20 | 1.89 |
| － | Classified other species | 0.0346±0.010 | 0.0406±0.012 | － |
| － | Unclassified other species | 31.4±5.6 | 19.5±1.8 | － |
Same conventions as in Table 1.
Species level. The results of our species-level analyses are shown in Table 3. The D-allose group had a significantly higher relative abundance of two species in the intestinal microflora. Bacteroides acidifaciens was more abundant compared to controls (t (13.2) = 3.05, p < 0.01). B. acidifaciens helps maintain the intestinal immune system and produces propionic acid.31) Akkermansia muciniphila, which resides in the human intestine as a typical mucin glycan-degrading bacteria, was more abundant compared to controls (t (11.1) = 2.21, p < 0.05). A. muciniphila is expected to be a potential probiotic, and its abundance is inversely correlated with metabolic disorders, such as diabetes, obesity, and inflammatory conditions.32) The D-allose group had a significantly lower relative abundance of Ruminococcus gnavus (t (10.7) = 2.70, p < 0.05), which produces an inflammatory polysaccharide. In patients with Crohn's disease, the relative abundance of R. gnavus increases during symptom flares.33) These results suggest that D-allose might contribute to maintaining intestinal health. The D-allose group had a significantly lower relative abundance of Bacteroides caecimuris (t (19.3) = 3.45, p < 0.01) and Acetatifactor muris (t (11.3) = 3.02, p < 0.05), which can be isolated from the cecal content of obese mice.30) The physiological function of these species is still unclear.
Organic acid composition in aged cecum after long-term ingestion of D-allose.
D-Allose altered the relative abundance of several intestinal bacterial species and genera associated with organic acid production (Table 4). We measured increased B. acidifaciens, a species that produces propionic acids, but decreased Lachnospiraceae, a family that produces butyric acids. However, compared to the control group, the total organic acid content and the overall organic acid composition in the aged mouse cecum did not change significantly after D-allose administration: succinic acid (t (18.8) = 0.351, p = 0.730); lactic acid (t (16.1) = 1.38, p = 0.187); acetic acid (t (12.2) = 0.623, p = 0.545); propionic acid (t (15.2) = 1.76, p = 0.099); butyric acid (t (20.0) = 0.548, p = 0.590); total organic acids (t (19.0) = 0.453, p = 0.656). Thus, it appears that long-term ingestion of D-allose does not change organic acid concentration in the aged cecum as a result of altering the abundance of various intestinal bacteria.
Table 4. Organic acid concentration (µmol/g) in aged mouse cecum after long-term ingestion of D-allose.
| Succinic acid | Lactic acid | Acetic acid | Propionic acid | Butyric acid | Total | |
| Control | 2.02±0.27 | 15.3±3.0 | 63.3±0.8 | 7.37±0.70 | 17.7±1.7 | 106±3.7 |
| D-Allose | 1.89±0.23 | 10.1±2.0 | 65.6±3.4 | 8.88±0.43 | 16.2±2.0 | 101±5.2 |
Control group (10 mice) and D-allose group (12 mice). Mean values ±SEM.
Possible mechanism for D-allose in altering intestinal bacteria.
Some sugar-related compounds and artificial sweeteners are reported to alter intestinal flora,34) and changes in inflammation and metabolism status and fermentability are reported to be involved. Although past studies showed that D-allose by itself is difficult to ferment,4) intestinal Escherichia coli K-12 has been found to possess the genes involved in metabolizing D-allose as an energy source,2) indicating that this species is capable of growing in the presence of D-allose. As for B. acidifaciens and A. muciniphila, which were elevated in mice that ingested D-allose in our study, it is unknown whether they metabolize D-allose. Since D-allose may suppress intestinal α-glucosidase,35) and thus prevent the breakdown of other carbohydrates in the cecum, these carbohydrates can potentially undergo fermentation, which can affect intestinal flora.
As another possible mechanism, brain-gut interaction is considered, that might account for the way D-allose alters intestinal microbiota is through the central nervous system (CNS). Recently, accumulating evidence has suggested the interaction between the CNS and gut microbiota.36) Anatomically, a rich network of bidirectional neuronal connections exists between the central and enteric nervous system.37),38) Such an anatomical interplay within the gut-brain axis prompted us to hypothesize that the D-allose-induced alterations in intestinal microbiota are achieved through the CNS. Although little is known about the action of D-allose in the CNS, D-allose is reported to protect the brain from ischemic damage caused by ROS.2) These results suggest that D-allose may exert beneficial effects on cells in the CNS via the reduction of oxidative stress.
Concluding remarks.
In this study, we investigated the effect of long-term D-allose administration on the intestinal environment in aged mice. D-Allose favorably altered the balance of bacterial phyla of the intestinal microbiome. This improvement implies that D-allose might favorably modulate one feature of aging; that is, age-related decrements in beneficial gut bacterial, perhaps ultimately protecting against various gut-related diseases. Moreover, long-term D-allose ingestion altered several genera and species of intestinal bacteria related to metabolic disorders or intestinal health. Our study revealed the beneficial effect of D-allose at the genus and species level, however, we need to explore the biological significance of the changes in the intestinal environment to establish the physiological roles of D-allose. Also, we need to assess the biological parameters related to specific intestinal bacteria, such as B. acidifaciens and A. muciniphila. Further, clinical trials will be essential to establish the effect of D-allose in humans, considering known differences in the kind and proportions of intestinal bacteria between mice and humans.39)
CONFLICTS OF INTEREST
The authors declare the following financial interests/personal relationships that may be considered as potential competing interests: Tomoya Shintani, Akane Kanasaki, Misuzu Tanaka, and Tetsuo Iida are employees of the Matsutani Chemical Industry Co., Ltd. Genki Ozawa and Tadao Kunihiro are employees of Techno Suruga Laboratory Co., Ltd.
ACKNOWLEDGMENTS
We thank Tomoko Arasaki and Kazuko Nakanishi (Tokyo Metropolitan Institute of Gerontology) for their technical support. This research was supported by Matsutani Chemical Industry Co., Ltd. This work was also supported in part by JSPS KAKENHI (24730642, 25560382, 26115532, 26430076, 25293331, 15H03103, 18K07460, 19K22834, 19H04044, and 21K19752).
REFERENCES
- 1).Granström T.B., Takata G., Tokuda M., and Izumori K.: Izumoring: a novel and complete strategy for bioproduction of rare sugars. J. Biosci. Bioeng., 97, 89-94 (2004). [DOI] [PubMed] [Google Scholar]
- 2).Lim Y.R. and Oh D.K.: Microbial metabolism and biotechnological production of D-allose. Appl. Microbiol. Biotechnol., 91, 229-235 (2011). [DOI] [PubMed] [Google Scholar]
- 3).Iga Y. and Matsuo T.: D-allose metabolism in rats. J. Jpn. Soc. Nutr. Food Sci., 63, 17-19 (2010). [Google Scholar]
- 4).Kitagawa M., Tanaka M., Yoshikawa Y., Iida T., and Kishimoto Y.: Evaluation of absorption and fermentability of D-mannose, D-sorbose, and D-allose in humans. Luminacoids Res., 22, 75-82 (2018). [Google Scholar]
- 5).Kishida K., Iida T., Yamada T., and Toyoda Y.: D-Allose is absorbed via sodium-dependent glucose cotransporter 1 (SGLT1) in the rat small intestine. Metabol. Open, 11, 100112 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 6).Kimura S., Zhang G., Nishiyama A., Nagai Y., Nakagawa T., Miyanaka H., Fujisawa Y., Miyatake A., Nagai T., Tokuda M., and Abe Y.: D-Allose, an all-cis aldo-hexose, suppresses development of salt-induced hypertension in Dahl rats. J. Hypertens., 23, 1887-1894 (2005). [DOI] [PubMed] [Google Scholar]
- 7).Yamada K., Noguchi C., Kamitori K., Dong Y., Hirata Y., Hossain M. A., Tsukamoto I., Tokuda M., and Yamaguchi F.: Rare sugar D-allose strongly induces thioredoxin-interacting protein and inhibits osteoclast differentiation in Raw264 cells. Nutr. Res., 32, 116-123 (2012). [DOI] [PubMed] [Google Scholar]
- 8).Ishihara Y., Katayama K., Sakabe M., Kitamura M., Aizawa M., Takara M., and Itoh K.: Antioxidant properties of rare sugar D-allose: Effects on mitochondrial reactive oxygen species production in Neuro2A cells. J. Biosci. Bioeng., 112, 638-642 (2011). [DOI] [PubMed] [Google Scholar]
- 9).Ottman N., Smidt H., De Vos W.M., and Belzer C.: The function of our microbiota: who is out there and what do they do? Front. Cell. Infect. Microbiol., 2, 104 (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 10).Lynch S.V. and Pedersen O.: The human intestinal microbiome in health and disease. N. Engl. J. Med., 375, 2369-2379 (2016). [DOI] [PubMed] [Google Scholar]
- 11).Odamaki T., Kato K., Sugahara H., Hashikura N., Takahashi S., Xiao J.Z., Abe F., and Osawa R.: Age-related changes in gut microbiota composition from newborn to centenarian: a cross-sectional study. BMC Microbiol., 16, 90 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 12).Yang J.Y., Lee Y.S., Kim Y., Lee S.H., Ryu S., Fukuda S., Hase K., Yang C.S., Lim H.S., Kim M.S., Kim H.M., Ahn S.H., Kwon B.E., Ko H.J., and Kweon M.N.: Gut commensal Bacteroides acidifaciens prevents obesity and improves insulin sensitivity in mice. Mucosal Immunol., 10, 104-116 (2017). [DOI] [PubMed] [Google Scholar]
- 13).Markowiak-Kopeć P. and Śliżewska K.: The effect of probiotics on the production of short-chain fatty acids by human intestinal microbiome. Nutrients, 12, 1107 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 14).Ley R.E., Bäckhed F., Turnbaugh P., Lozupone C.A., Knight R.D., and Gordon J.I.: Obesity alters gut microbial ecology. Proc. Natl. Acad. Sci. USA, 102, 11070-11075 (2005). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 15).Iga Y., Nakamichi K., Shirai Y., and Matsuo T.: Acute and sub-chronic toxicity of D-allose in rats. Biosci. Biotechnol. Biochem., 74, 1476-1478 (2010). [DOI] [PubMed] [Google Scholar]
- 16).Takahashi S., Tomita J., Nishioka K., Hisada T., and Nishijima M.: Development of a prokaryotic universal primer for simultaneous analysis of bacteria and archaea using next-generation sequencing. PLoS ONE, 9, e105592 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 17).Caporaso J.G., Lauber C.L., Walters W.A., Berg-Lyons D., Lozupone C.A., Turnbaugh P.J., Fierer N., and Knight R.: Global patterns of 16S rRNA diversity at a depth of millions of sequences per sample. Proc. Natl. Acad. Sci. USA, 108, 4516-4522 (2011). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 18).Hisada T., Endoh K., and Kuriki K.: Inter- and intra-individual variations in seasonal and daily stabilities of the human gut microbiota in Japanese. Arch. Microbiol., 197, 919-934 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 19).Aronesty E.: Comparison of sequencing utility programs. Open Bioinforma. J., 7, 1-8 (2013). [Google Scholar]
- 20).Gordon A. and Hannon G.J.: FASTX-Toolkit FASTQ/A short-reads preprocessing tools [software]. Available from: http://hannonlab.cshl.edu/fastx_toolkit/index.html.
- 21).Edgar R.C., Haas B.J., Clemente J.C., Quince C., and Knight R.: UCHIME improves sensitivity and speed of chimera detection. Bioinformatics, 27, 2194-2200 (2011). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 22).Kasai C., Sugimoto K., Moritani I., Tanaka J., Oya Y., Inoue H., Tameda M., Shiraki K., Ito M., Takei Y., and Takase K.: Comparison of the gut microbiota composition between obese and non-obese individuals in a Japanese population, as analyzed by terminal restriction fragment length polymorphism and next-generation sequencing. BMC Gastroenterol., 15, 100 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 23).Wang Q., Garrity G.M., Tiedje J.M., and Cole J.R.: Native Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl. Environ. Microbiol., 73, 5261-5267 (2007). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 24).Weiss S., Xu Z.Z., Peddada S., Amir A., Bittinger K., Gonzalez A., Lozupone C., Zaneveld J.R., Vázquez-Baeza Y., Birmingham A., Hyde E.R., and Knight R.: Normalization and microbial differential abundance strategies depend upon data characteristics. Microbiome, 5, 27 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 25).Mizuno H., Bamba S., Abe N., and Sasaki M.: Effects of an alginate-containing variable-viscosity enteral nutrition formula on defecation, intestinal microbiota, and short-chain fatty acid production. J. Funct. Foods, 67, 103852 (2020). [Google Scholar]
- 26).Delacre M., Lakens D., and Leys C.: Why psychologists should by default use Welch's t-test instead of student's t-test. Int. Rev. Soc. Psychol., 30, 92-101 (2017). [Google Scholar]
- 27).Wu C.S., Muthyala S., Klemashevich C., Ufondu A.U., Menon R., Chen Z., Devaraj S., Jayaraman A., and Sun Y.: Age-dependent remodeling of gut microbiome and host serum metabolome in mice. Aging, 13, 6330-6345 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 28).Vacca M., Celano G., Calabrese F.M., Portincasa P., Gobbetti M., and De Angelis M.: The controversial role of human gut lachnospiraceae. Microorganisms, 8, 573 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 29).Liu X., Mao B., Gu J., Wu J., Cui S., Wang G., Zhao J., Zhang H., and Chen W.: Blautia-a new functional genus with potential probiotic properties? Gut Microbes, 13, 1-21 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 30).Pfeiffer N., Desmarchelier C., Blaut M., Daniel H., Haller D., and Clavel T.: Acetatifactor muris gen. nov., sp. nov., a novel bacterium isolated from the intestine of an obese mouse. Arch. Microbiol., 194, 901-907 (2012). [DOI] [PubMed] [Google Scholar]
- 31).Yanagibashi T., Hosono A., Oyama A., Tsuda M., Suzuki A., Hachimura S., Takahashi Y., Momose Y., Itoh K., Hirayama K., Takahashi K., and Kaminogawa S.: IgA production in the large intestine is modulated by a different mechanism than in the small intestine: Bacteroides acidifaciens promotes IgA production in the large intestine by inducing germinal center formation and increasing the number of IgA+ B cells. Immunobiology, 218, 645-651 (2013). [DOI] [PubMed] [Google Scholar]
- 32).Zhang T., Li Q., Cheng L., Buch H., and Zhang F.: Akkermansia muciniphila is a promising probiotic. Microb. Biotechnol., 12, 1109-1125. (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 33).Henke M.T., Kenny D. J., Cassilly C.D., Vlamakis H., Xavier R.J., and Clardy J.: Ruminococcus gnavus, a member of the human gut microbiome associated with Crohn's disease, produces an inflammatory polysaccharide. Proc. Natl. Acad. Sci. USA, 116, 12672-12677 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 34).Javier Ruiz-Ojeda F., Plaza-Díaz J., Sáez-Lara M.J., and Gil A.: Effects of sweeteners on the gut microbiota: A review of experimental studies and clinical trials. Adv. Nutr., 10, S31-S48 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 35).Kizu J., Okabe K., Hosono A., and Yamada K.: Effect of D-allose and D-tagatose on the rats small intestinal sucrase and isomaltase activities. in Abstract Book of 70th Annual Meeting of Japan Society of Nutrition and Food Science, 3F-13p, Kobe, Japan (2016). [Google Scholar]
- 36).Fung T.C., Olson C.A., and Hsiao E.Y.: Interactions between the microbiota, immune and nervous systems in health and disease. Nat. Neurosci., 20, 145-155 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 37).Carabotti M., Scirocco A., Maselli M.A., and Severi C.: The gut-brain axis: interactions between enteric microbiota, central and enteric nervous systems. Ann. Gastroenterol., 28, 203-209 (2015). [PMC free article] [PubMed] [Google Scholar]
- 38).Drossman D. A.: The functional gastrointestinal disorders and the Rome II process. Gut, 45, II1-5 (1999). [DOI] [PMC free article] [PubMed] [Google Scholar]
- 39).Nguyen T.L.A., Vieira-Silva S., Liston A., and Raes J.: How informative is the mouse for human gut microbiota research? Dis. Model. Mech., 8, 1-16 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]