Role of the Small Intestinal Microbiota in Gastrointestinal Disorders
Article information
Abstract
The small intestinal microbiota plays a crucial role in maintaining the health and pathogenesis of various gastrointestinal disorders. Despite extensive research on gut microbiota, studies focusing on the small intestine are limited owing to methodological challenges. This review discusses the taxonomic composition, microbial load, and diversity of normal small intestinal microbiota. Additionally, it highlights the role of small intestinal microbiota in gastrointestinal disorders, such as functional dyspepsia, small intestinal bacterial overgrowth, and nonsteroidal anti-inflammatory drug-induced enteropathy. The impact of proton pump inhibitors on small intestinal microbiota dysbiosis underscores the importance of the appropriate use of strong acid suppressants in clinical practice. Future research should focus on both the luminal and mucosal microbiota of the small intestine to explore the taxonomic changes and functional differences.
INTRODUCTION
Unlike in the past, when microorganisms were studied using traditional culture methods, the introduction of next-generation sequencing over the past two decades has enabled extensive research on gut microbiota. To date, most studies have focused on bacteria using 16S rRNA sequencing, and the primary measure used is the diversity of the sample. α-diversity assesses the variation in microbiome diversity within a single sample, capturing both the richness (the number of species within a sample) and evenness (the relative abundance of different species within a sample). β-diversity, on the other hand, compares the taxonomic diversity between different samples and can be represented either with (weighted) or without (unweighted) accounting for the relative abundance of individual taxa [1]. Most studies for gut microbiota have focused on fecal samples or colonic mucosal biopsies obtained through colonoscopy, while research on the small intestinal microbiota is limited. The small intestine plays a vital role in digestion, nutrient absorption, metabolism, and immune regulation, and the small intestinal microbiota is believed to be integral to these functions [2,3]. In this article, we aimed to explore the normal small intestinal microbiota and the relationship between small intestinal dysbiosis and gastrointestinal diseases.
NORMAL SMALL INTESTINAL MICROBIOTA
Understanding normal gut microbiota is crucial for recognizing its role in various diseases. However, this information on the small intestine is limited compared to that on the colon. The small intestine has many different characteristics from the colon in terms of the environment in which microbiota survive. The small intestine is characterized by faster transit times and various secretions such as bile and pancreatic fluids. It is an exceedingly long organ that extends from the duodenum to the ileum, with varying pH levels across different regions. The oxygen concentration in the small intestine, although relatively higher than that in the colon, gradually decreases from the duodenum to the ileum. Unlike the colon, which is mainly populated by strict anaerobes, the small intestine harbors facultative anaerobic bacteria at varying concentrations depending on the region. Furthermore, the dietary nutrients consumed by the bacteria vary in different regions of the small intestine. These characteristics create a unique environment for microbial communities, resulting in lower microbial diversity and density in the small intestine compared to the colon [2]. The stomach and duodenum have a similar bacterial concentration of about 103 cells/mL, which increases distally in the small intestine, reaching up to 108 cells/mL in the distal ileum [3].
Compared to the relatively straightforward investigation of the colonic microbiota using non-invasive fecal samples or tissue biopsies collected through colonoscopy, investigating the normal small intestinal microbiota remains more challenging owing to methodological limitations. The study of the small intestinal microbiota involves methods such as aspiration of intestinal fluid, biopsy, and luminal brushing [2]. However, accessing all areas of the small intestine is challenging [4]. Most studies have been conducted under abnormal conditions, such as in patients with various diseases or in individuals with stomas. Therefore, defining the composition of a “healthy” small intestinal microbiota remains challenging.
Despite these limitations, certain general trends emerged in these studies. At the phylum level, the small intestinal microbiota is composed of Bacillota (formerly Firmicutes), Bacteroidota (formerly Bacteroidetes), Pseudomonadota (formerly Proteobacteria), Fusobacteriota (formerly Fusobacteria), and Actinomycetota (formerly Actinobacteria) in varying proportions depending on the individual and study. Other phyla such as Verrucomicrobiota (formerly Verrucomicrobia) and Saccharibacteria (formerly TM7) were occasionally detected. At the genus level, core microbiota, such as Streptococcus, Veillonella, Prevotella, Fusobacterium, and Haemophilus have been consistently identified across different regions of the small intestine [2].
The microbial communities of the duodenum and jejunum are similar in composition, while they differ from those of the ileum [5]. In addition to the core microbiota, Neisseria, Granulicatella, Gemella, Rothia, and Actinomyces are commonly found in the duodenum and jejunum, whereas Bacteroides, Escherichia, Shigella, Ruminococcus, Bifidobacterium, Clostridium, and Lactobacillus are predominant in the ileum (Fig. 1) [2].
Normal small intestinal microbiota and compositional change in gastrointestinal disorders reported in human studies. FD, functional dyspepsia; SIBO, small intestinal bacterial overgrowth; NSAIDs, nonsteroidal anti-inflammatory drugs.
Notably, microbial differences exist between the lumen and mucosa of the small intestine. Studies have observed the presence of specific microbial taxa that appear to be quite similar; however, the relative abundance of each taxon, such as Streptococcus, Prevotella, Fusobacterium, Actinomyces, Bacteroides, and Acinetobacter, differs between the lumen and mucosa. Streptococcus is consistently higher in luminal aspirates than in mucosal specimens across studies [2,6,7]. Mucosa-associated microbiota (MAM) are more conserved than those in luminal samples [6]. However, studies comparing the microbial composition in mucosal and luminal samples are limited. Further research is required to understand how this niche-specific community varies in the small intestine of human. In addition to bacteria, the gut microbiota includes non-bacterial microorganisms, such as viruses, archaea, and fungi. Research on colonic microbiota is expanding to include these nonbacterial components. However, studies on the small intestinal microbiota have been limited to bacteria, indicating the need for further exploration of these diverse microbiota.
FACTORS AFFECTING SMALL INTESTINAL MICROBIOTA
The composition of the small intestinal microbiota is influenced by various external factors such as nutrients, smoking, and acid suppressants [8,9].
Diet significantly influences the composition and function of the small intestinal microbiota. A study of ileostomy effluent samples revealed that the small intestine harbors a greater abundance of genus related to carbohydrate metabolism than the feces. While the colon is more suited to degrade complex carbohydrates, the small intestine microbiota rapidly adapts to nutrient availability, efficiently metabolizing simple carbohydrates [4]. Streptococcus plays a key role in primary digestion within the small intestine, producing lactic acid that supports the growth of secondary fermenters such as Veillonella and Clostridium [10]. The small intestinal microbiota also play a crucial role in regulating lipid digestion and absorption and are susceptible to changes caused by a high-fat diet [11]. In mouse experiments, feeding a high-fat diet, high-sugar diet, and high-protein diet changes small intestinal microbiota in distinct ways compared to a standard diet, with these changes being more pronounced than those in colonic microbiota [12].
Smoking has been shown to disrupt the mucosa-associated and luminal microbiota in the duodenum. In the lumen, smokers exhibited a decreased abundance of Prevotellaceae, Neisseriaceae, and Porphyromonadaceae, whereas the abundance of Enterobacteriaceae and Lactobacillaceae increased [8].
The duodenum harbors fewer microbiota than distal regions; however, it shows higher intra-individual variability and compositional dynamics [5]. This variation in the duodenal microbiota is largely affected by pH [5]. Therefore, proton pump inhibitors (PPI), which strongly and persistently reduce gastric acid, significantly impact the microbiota in the duodenum and small intestine. PPI administration increases Streptococcus and decreases Porphyromonas and Prevotella in the duodenal mucosa of health volunteers, which is correlated with duodenal eosinophils [13]. Conversely, PPI treatment in patients with gastric ulcers increases Akkermansia muciniphila and Porphyromonas dontalis and decreases Enterococcaceae, Enterobacteriaceae, Coprococcus, and Synergistes [9].
The impact of PPI on the small intestinal microbiota is presumed to be due to increased gastric pH [14]. Acid suppression permits an increase in gastric microbiota, exerting a downstream effect on the small intestinal microbial composition [15]. However, the small intestinal pH values observed in wireless motility capsules were similar between PPI users and non-users [16]. This finding indicates that the effects of PPI on the small intestinal microbiota may be driven by a pH-independent mechanism. For instance, PPI can modify the lumen content, disrupt nutrient absorption, and alter the quantity or location of bacterial food substrates. In addition, PPIs have been found to bind to nongastric H+/K+-ATPases in commensal bacteria and fungi. The P-type ATPase family, which includes H+/K+-ATPases, is found in the fungi Helicobacter pylori and Streptococcus pneumoniae [14,17].
PPI decrease symptoms and duodenal immune cells in patients with functional dyspepsia (FD) who have increased mucosal permeability; however, they increase immune cells and mucosal permeability in healthy volunteers [18]. PPI are particularly associated with small intestinal bacterial overgrowth (SIBO) [19] and aggravate nonsteroidal anti-inflammatory drug (NSAID)-induced small intestinal damage. These small intestinal conditions associated with PPI and small intestinal dysbiosis will be addressed later.
FUNCTIONAL DYSPEPSIA
FD is a complex and heterogeneous disorder characterized by chronic or recurrent upper abdominal pain or discomfort, including postprandial fullness and early satiety not explained by structural or biochemical abnormalities [20]. Since an increase in eosinophils in the duodenal mucosa has been reported in postprandial distress syndrome [21], subsequent studies have observed impaired duodenal mucosal integrity and increased infiltration of eosinophils and mast cells, indicating low-grade duodenal inflammation [22,23].
Several small-scale studies recently reported changes in the duodenal microbiota of patients with FD. MAM is hypothesized to play a more direct role in the pathogenesis of FD [24]. The mucus layer acts as a barrier against pathogenic microbiota, preventing their translocation to host tissues. Therefore, the microbiota residing in the mucus layer and those capable of penetrating this layer are more likely to induce mucosal inflammation [25,26]. Duodenal MAM is taxonomically distinct from those in other regions of the gastrointestinal tract, with Streptococcus as the dominant genus and lower levels of Prevotella, Veillonella, and Neisseria [27,28].
Most studies on the duodenal mucosal microbiota in patients with FD have observed an increased abundance of Bacillota and Streptococcus compared to that in healthy volunteers. The abundance of Streptococcus positively correlated with the severity of gastrointestinal symptoms. Prevotella, Veillonella, and Actinomyces are significantly reduced and show an inverse correlation with symptoms [23,29-31]. Additionally, mucosa-associated Neisseria, Porphyromonas, Selenomonas, Haemophilus, and Fusobacterium were lower in patients with FD compared to controls, and Porphyromonas inversely correlated with symptoms and duodenal eosinophils [13]. Furthermore, no significant difference was observed in MAM α-diversity; however, β-diversity differed between patients with FD and healthy controls [30,31]. Moreover, the higher bacterial load observed in patients with FD showed a negative correlation with bacterial diversity and quality of life scores and a positive correlation with the severity of upper gastrointestinal meal-related symptoms [29]. Collectively, the microbial changes in the duodenum observed in patients with FD suggest distinct alterations in microbial load and diversity, including increased Streptococcus and decreased other microbiota. This finding suggests a shift in duodenal microbial composition towards the oral microbiota to some extent or the alteration of native duodenal microbiota [24]. A recent study in Australia isolated a new Streptococcus salivarius, designated strain AGIRA0003, from the duodenal tissue of patients with dyspeptic symptoms. Given the significant differences in β-diversity between patients and controls, despite no differences in α-diversity [13,30], specific microbial structural changes related to the FD may be more important than changes in overall bacterial counts in the duodenum.
SMALL INTESTINAL BACTERIAL OVERGROWTH
SIBO is the most well-known and extensively studied condition in which the small intestinal microbiota contributes to the pathogenesis of gastrointestinal diseases. The traditional gold standard for diagnosis involves the measurement of bacterial counts in jejunal aspirates. Historically, the diagnostic criterion has been set to >105 CFU/mL of bacteria. However, recent AGA guidelines have recommended a new lower cutoff of >103 CFU/mL of coliforms in duodenal aspirates for the diagnosis of SIBO [32]. Despite being the gold standard, culture-based methods are invasive, expensive, and prone to contamination, which has led to the more common use of breath tests [33]. SIBO is linked to conditions such as dysmotility disorders, altered anatomy, and PPI use, and causes symptoms such as bloating, diarrhea, and malabsorption [32]. It has also been extensively studied in irritable bowel syndrome [34], and some associations have been reported in FD [35].
Several studies have reported that changes in small intestinal microbiota are associated with SIBO. Patients with SIBO had lower α-diversity of small intestinal microbiota than those without SIBO across the studies, regardless of diagnostic methods [36-38]. Additionally, β-diversity was different between patients with SIBO and non-SIBO individual. However, changes in microbial composition are inconsistent and vary among studies. In a study defining SIBO as >103 CFU/mL, patients with SIBO had a 4.31-fold higher relative abundance of Pseudomonadota and a 1.64-fold lower relative abundance of Bacillota than non-SIBO participants [36]. Additionally, patients with SIBO exhibited higher relative abundances of Gammaproteobacteria and Enterobacteriaceae, which were associated with bloating [36]. In another study, patients with SIBO showed an increased relative abundance of Enterobacteriaceae, Escherichia coli, Klebsiella pneumoniae, Klebsiella aerogenes, and Enterobacter. Shotgun sequencing revealed that two Escherichia coli strains and two Klebsiella species comprised 40.24% of all duodenal bacteria in patients with SIBO, compared to 5.6% in non-SIBO patients. These strains are associated with abdominal pain, diarrhea, and excess gas [38]. The magnitude of dysbiosis may play a more significant role in the pathogenesis of SIBO than an increase in microbial numbers within the small intestine. A study defining SIBO as >105 CFU/mL in duodenal aspirates found no correlation between SIBO and symptoms. Instead, dysbiosis in the small intestine was strongly associated with symptoms [39]. Symptomatic patients exhibited a decreased abundance of Porphyromonas, Prevotella, and Fusobacterium.
Microbial metabolic alterations may also play significant roles in this process. In patients with SIBO, symptoms were linked to increased microbial metabolic pathways involved in carbohydrate fermentation, hydrogen production, and hydrogen sulfide production [38]. Therefore, with the future availability of high-throughput sequencing and metabolomics for the small intestinal microbiota, the definition and diagnostic criteria for SIBO may evolve.
NSAID-INDUCED ENTEROPATHY
NSAID are a major cause of gastrointestinal mucosal damage, whereas PPI can prevent gastric mucosal injury caused by NSAID [40]. However, unlike in the stomach, the combined use of PPI and NSAID has been demonstrated to exacerbate small intestinal mucosal damage compared to NSAID alone in animal and human studies using capsule endoscopy [41,42]. This paradox occurs because the gut microbiota plays a crucial role in the development of NSAID-induced small intestinal injury [43]. Gut microbiota has long been recognized to be involved in the pathogenesis of NSAID-induced mucosal damage in the small intestine. In 1977, a study revealed that germ-free rats were found to be more resistant to indomethacin-induced small intestinal injury than conventional rats, particularly in males [44]. Similarly, NSAID caused small intestinal mucosal damage in gnotobiotic rats transplanted with Escherichia coli or Eubacterium; however, this effect was not observed in gnotobiotic rats transplanted with Bifidobacterium or Lactobacillus [45].
NSAIDs disrupt the intestinal mucus layer and the phospholipid bilayer on the cell surface while simultaneously decoupling mitochondrial oxidative phosphorylation in epithelial cells [46]. These changes lead to increased intestinal permeability, facilitating bacterial translocation and resulting in submucosal immune activation, which ultimately causes mucosal damage [47]. As discussed previously, PPI induces intestinal dysbiosis; thus, concomitant administration of PPI exacerbates NSAID-induced small intestinal mucosal damage [42].
Most studies on NSAID-induced changes in small intestinal microbiota have been conducted in animal models, with limited studies in humans. In a human study examining the effects of indomethacin on microbiota composition, duodenal aspirates showed no significant change in α-diversity; however, β-diversity analysis revealed considerable compositional shifts [48]. Taxonomically, a decrease in Pseudomonadota at the phylum level and in Alphaproteobacteria, Rhizobiales, and Pseudomonadaceae after indomethacin administration was observed [48]. These findings contrast with another animal study, which observed an increase in Pseudomonadota after diclofenac administration [49].
CONCLUSION
Research on small intestinal microbiota remains in its early stages owing to methodological limitations, leaving both normal and disease-associated states relatively unexplored compared to other parts of the gastrointestinal tract. However, studies have demonstrated that alterations in the composition of small intestinal microbiota contribute to the pathogenesis of several gastrointestinal disorders, including FD, SIBO, and NSAID-induced enteropathy. PPIs significantly affect the microbiota in the small intestine, leading to dysbiosis, which plays a key role in these disorders. With the introduction of more potent potassium-competitive acid blockers in clinical practice, further research is needed to determine whether these medications may cause more severe dysbiosis than PPIs [50]. Moreover, avoiding the unnecessary use of strong acid suppressants is crucial [51]. In the future, research should focus on both the luminal and mucosal microbiota of the small intestine, exploring not only taxonomic changes but also functional differences.
Notes
Availability of Data and Material
Data sharing not applicable to this article as no datasets were generated or analyzed during the study.
Conflicts of Interest
Yong Sung Kim, a contributing editor of the Korean Journal of Helicobacter and Upper Gastrointestinal Research, was not involved in the editorial evaluation or decision to publish this article. All remaining authors have declared no conflicts of interest.
Funding Statement
This work was supported by Wonkwang University 2015–2016 (JK).
Authors’ Contribution
Conceptualization: Jungnam Kwon, Yong Sung Kim. Data curation: Jungnam Kwon, Yong Sung Kim. Formal analysis: Dong Han Yeom, Moon Yong Lee, Yong Sung Kim. Investigation: all authors. Writing—original draft: Jungnam Kwon, Yong Sung Kim. Writing—review & editing. Dong Han Yeom, Moon Yong Lee, Yong Sung Kim. Approval of final manuscript: all authors.
Acknowledgements
None