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Open AccessReview

Bifidobacterium bifidum: A Key Member of the Early Human Gut Microbiota

1
Laboratory of Probiogenomics, Department of Chemistry, Life Sciences and Environmental Sustainability, University of Parma, 43124 Parma, Italy
2
Microbiome Research Hub, University of Parma, 43124 Parma, Italy
3
School of Microbiology, University College Cork, T12 YT20 Cork, Ireland
4
APC Microbiome Institute, University College Cork, T12 YT20 Cork, Ireland
*
Author to whom correspondence should be addressed.
Microorganisms 2019, 7(11), 544; https://doi.org/10.3390/microorganisms7110544
Received: 2 October 2019 / Revised: 29 October 2019 / Accepted: 7 November 2019 / Published: 9 November 2019

Abstract

Bifidobacteria typically represent the most abundant bacteria of the human gut microbiota in healthy breast-fed infants. Members of the Bifidobacterium bifidum species constitute one of the dominant taxa amongst these bifidobacterial communities and have been shown to display notable physiological and genetic features encompassing adhesion to epithelia as well as metabolism of host-derived glycans. In the current review, we discuss current knowledge concerning particular biological characteristics of the B. bifidum species that support its specific adaptation to the human gut and their implications in terms of supporting host health.
Keywords: Bifidobacterium bifidum; bifidobacteria; probiotics; genomics; microbiota Bifidobacterium bifidum; bifidobacteria; probiotics; genomics; microbiota

1. General Features of the Genus Bifidobacterium

The genus Bifidobacterium belongs to the Actinobacteria phylum and this genus together with nine other genera constitute the Bifidobacteriaceae family [1]. Currently, the genus Bifidobacterium is comprised of 80 (sub)species, which are distributed across seven different ecological niches, encompassing the Gastro Intestinal Tract (GIT) of humans, non-human mammals, birds, and social insects; waste water; and the oral cavity [2,3]. Remarkably, these ecological origins may represent the biological niche that is common to all these habitats, which is characterized by the fact that a large number of bifidobacterial hosts are submitting their child to parental care. Thus, their ecological origins are perhaps enabled by maternal inheritance of bifidobacterial cells. Interestingly, this supposition has recently been corroborated by the mapping of (near) identical bifidobacterial strains in mothers and their corresponding children (see below) [4,5].
A small number of bifidobacterial (sub)species, such as Bifidobacterium pseudolongum, Bifidobacterium adolescentis, Bifidobacterium pseudocatenulatum, and Bifidobacterium bifidum, have been isolated from various animal/mammalian hosts and for this reason are acknowledged as cosmopolitan bifidobacterial taxa [6]. In contrast, other taxa like Bifidobacterium breve appear to be much less widely distributed, perhaps due to adaptative behavior that is host-specific [6]. Among those bifidobacteria that are found in primates, certain bifidobacterial species are commonly identified in adults, such as B. adolescentis and B. catenulatum, while others, like B. bifidum, B. breve, and B. longum subsp. infantis, are more typically found in the fecal samples from breast-fed infants [7]. However, there does not seem to be an absolute infant versus adult division of bifidobacterial (sub)species. Such findings make sense from the perspective of vertical transmission of bifidobacterial species from mother to child, which also encompasses adult-type species like B. adolescentis [8,9]. The different preponderance of one species over another in the adult/newborn GIT is influenced by the different composition of the intestinal microbiota in terms of complexity, which in turn is very much determined by host diet [10].

2. Bifidobacterial Communities of the Human Gut

Bifidobacteria rapidly colonize the gut of infants within the first weeks following birth, a phenomenon that is believed to be driven in no small part by the bifidogenic activities of specific mother milk-derived oligosaccharides, commonly referred to as Human Milk Oligosaccharides (HMOs). Metagenomic-based analyses revealed a high abundance of a small number of bifidobacterial species such as B. breve, followed by “adult-type” bifidobacterial taxa such as B. longum and B. adolescentis [11]. The level of bifidobacterial abundance in the human gut drops with aging, even if microbial profiling experiments that are based on FISH and metagenomic analyses have predicted that their relative load in the adult large intestine is about 4.3 ± 4.4% of total microorganisms [12,13].
In recent years, several metagenomics as well as culture-dependent investigations of the human gut have allowed a detailed dissection of the bifidobacterial biodiversity that is present in this environment [6,7,14,15,16]. Notably, such analyses revealed that the most abundant and prevalent bifidobacterial taxa existing in the human colon are strains of B. breve, B. bifidum, B. longum, B. adolescentis, B. pseudolongum, B. pseudocatenulatum, and B. animalis subsp. lactis. These studies have highlighted that the varying composition of bifidobacterial gut communities in different individuals underline both an inter-subject and an intra-subject variability, which is in agreement with the large inter-variability of the overall intestinal microbiota as described previously [12,17].
Recently, detailed cataloguing of bifidobacterial communities that are present during infancy has been performed through the sequencing of the Internally Transcribed Spacer (ITS) region of the bifidobacterial rRNA locus, which when compared to the 16S rRNA gene, allows a substantially higher taxonomic resolution [18]. In this study, the Hierarchical clustering that was built on bifidobacterial community profiles that are present in infant stool samples highlighted the occurrence of four bifidobacterial groups, i.e., bifidotypes, characterized by specific, commonly co-occurring taxa such as B. breve, B. bifidum, B. adolescentis, or B. longum [19]. In this context, B. bifidum was demonstrated to be an important taxon among these bifidotypes in which its occurrence correlates with numerous other bifidobacterial species that were recovered from infant fecal samples.
Interestingly, a study that compared the fecal microbiota composition of infants that were fed with bovine milk formula or with breast milk by 16S rRNA gene microbial profiling and qPCR showed that both B. breve and B. bifidum abundance was greater in the feces of breast milk-fed infants than in that of formula-fed infants [20]. Furthermore, it was observed that samples with a higher load of bifidobacteria are characterized by the presence of B. bifidum at greater than 10% of the total bifidobacterial population [20]. The link between the highest abundance of total bifidobacteria with appreciable B. bifidum populations in the stools of breast milk-fed babies suggests that trophic factors play a role in determining this association [20].
In addition, bifidobacterial communities that are present in milk samples frequently contain strains of B. bifidum, suggesting that milk represents a key vector for vertical transmission of bifidobacteria [19,21] (Figure 1).

3. Maternal Inheritance of Bifidobacteria

In recent years, bifidobacteria have been shown to be subject to mother–infant transfer via a vertical transmission route that seems to be operational both in human beings [4,22] as well as in other mammalian species [6]. The most frequently shared bifidobacterial species between mothers and their corresponding children are strains of B. bifidum and B. breve, which were displayed to persist in the human gut for up to one year of life [5]. Similar results were found in another study that aimed at tracing maternally inherited bifidobacterial strains [22]. These investigations involved a complex set of analyses such as culturomics, shotgun metagenomics, and ITS bifidobacterial profiling experiments [18,23,24,25] and resulted in the identification of B. longum subsp. longum BLOI2 and B. breve BBRI4, which were isolated in two mother–infant couples and were also shown to be maintained in the intestine of babies until six months of age [22]. Similarly, a detailed cataloguing of bifidobacterial communities in 25 mother–offspring couples based on ITS bifidobacterial profiling trials followed by cultivation experiments identified several bifidobacterial strains that occur in the gut of both the mother and her child and of which are also present in the matching human milk sample [14]. Such findings are corroborated by a similar situation in other primates and certain mammals, where bifidobacteria were displayed to be vertically inherited from the mother to her newborn and where the mother’s milk appears to constitute a key vector that drives such events [6]. Even if the intestinal establishment of bifidobacteria is believed to be (partially) maternally determined, some studies postulate that their acquisition occurs during pregnancy [21].
The precise biological/evolutionary consequences of horizontal transfer are still obscure, although one may argue that according to the holobiont concept [21], the first microbial colonizers, which include bifidobacteria, exploit crucial physiological/immunological/metabolic functions in driving host development. Another important speculation that is linked to the formation of the infant gut microbiota is that bifidobacteria, in spite of their drop in abundance upon the weaning stage, carry on after their initial transmission to the infant intestine and are then preserved, though perhaps at very low abundances, in the large intestine of the adult to ultimately be vectored to the following generation [22]. The vertical transmission events of bifidobacteria may thus represent the consequence of long-term co-evolution between bifidobacteria and their mammalian hosts.

4. Genomics Features of the B. bifidum Species

Many bifidobacterial species have had their genomes sequenced in recent years, in particular those that are exploited by the food industry as probiotic microorganisms [21]. In this context, genome sequencing was performed for multiple strains belonging to the species B. animalis subsp. lactis [26,27], B. breve [28], and B. longum subsp. longum [28]. Similarly, various strains harboring the B. bifidum taxon have been subjected to whole genome sequencing [29], allowing comparative genomic analyses, which has shown a closed pan-genome structure. Such in silico analyses uncovered specific genetic strategies, allowing members of the B. bifidum species to establish and persevere in the human gut, for example thanks to the synthesis of different types of pili [30,31] or through metabolic abilities pertaining to host-derived glycans [29,32].
The NCBI-deposited reference genome of the B. bifidum species is that of the infant stool isolate PRL2010 [32], which was sequenced and published in 2010. A functional classification of the genes occuring in B. bifidum chromosomes following the Cluster of Orthologous Genes (COG) families revealed that a large proportion (approximately 14%) are ascribed to the COG family of carbohydrate metabolism and transport [33,34].

5. The Glycobiome of B. bifidum Species

The glycobiome represents the overall genes that are expected to be responsible for the carbohydrate metabolism and in the B. bifidum species, consists of more than 3000 genes that encode putative carbohydrate-active enzymes, comprising glycosyl hydrolases (GHs), glycosyl transferases (GTs), and carbohydrate esterases (CEs). In particular, according to the Carbohydrate Active Enzymes (CAZy) database [35], there are many genes coding members of family GH13, which are largely identified in bifidobacterial chromosomes, commonly for those bifidobacterial strains that are identified from the intestine of mammals. GH13 are common of bacteria and are responsible for the breakdown of an extensive range of glycans, involving plant-derived complex carbohydrates like starch and correlated substrates (e.g., amylose and amylopectin and/or (cyclo)maltodextrins), trehalose, stachyose, raffinose, and melibiose [33]. Interestingly, the B. bifidum glycobiome also includes members of GH families that are crucial in host glycan metabolisms, like those harboring GH33 and GH34, representing exo-sialidases, GH29, and GH95, which characterize fucosidases, and GH20 involving hexosaminidase and lacto-N-biosidase activities, which are genes that are predicted to be responsible for the metabolism of host-glycans (see below) [34]. In this context, a comparative genomic analysis based on 15 different strains highlighted the presence of a conserved gene set that appears to uniquely occur in B. bifidum strains [29]. This gene set represents core genome sequences encoding various GHs that are specifically involved in mucin breakdown, which is indicative of a nutritional acquisition strategy targeting host-derived glycans [29].
Furthermore, an exhaustive analysis of the genetic backgrounds for carbohydrate uptake in B. bifidum species revealed that this taxon contains a relatively small number of genes coding for carbohydrate carriers when associated with other bifidobacterial taxa that are present in the infant intestine, like B. breve, B. longum subsp. longum, and B. longum subsp. infantis [7,36]. In fact, only 25 genes are estimated to synthetize carrier systems in the B. bifidum PRL2010 chromosome and be committed to glycan uptake. However, the other human gut bifidobacterial chromosomes are predicted to encompass between 35 and 68 such genes, as characterized by B. adolescentis ATCC 15703 and B. longum subsp. infantis ATCC 15697, respectively. This suggests that carbohydrate metabolism in B. bifidum taxon is limited to a reasonably low amount of carbohydrates.

6. The Ability of B. bifidum to Metabolize Host-Derived Glycans

The main carbohydrates that are produced by human beings, i.e., host-derived glycans, which may represent a carbon source for bifidobacteria, are mucin and HMOs. Notably, these sugars are found in the gut in high amounts during two different stages of human life, i.e., the infant period (HMOs) and the adult life (mucin). Interestingly, the B. bifidum taxon is present during both of these life stages, though its prevalence and abundance are higher during infancy. However, we may argue that its occurrence in the adult gut is supported by the action of glycan substrates such as mucin (see below).
Mucin is a glycoprotein that constitutes the key component of the mucus gel coating, which covers the epithelial surface of the human GIT [37]. The key glycan monomers that were identified in this glycoprotein encompass N-acetylglucosamine, N-acetylgalactosamine, fucose, galactose, and sialic acid [38]. Notably, B. bifidum species is the only member among all the recognized Bifidobacterium (sub)species that is capable of growth by means of mucin metabolism [32,39,40].
As mentioned before, a large portion of the predicted GH-encoding genes that were identified in the B. bifidum PRL2010 genome represent enzymes that are estimated to be associated with the breakdown of mucin-derived oligosaccharides, most of which are exclusively occurring in the B. bifidum genome [29]. In addition, according to the CAZy database [35], the B. bifidum PRL2010 chromosome encompasses members of two carbohydrate-binding module (CBM) families, i.e., CBM32 and CBM51, which are predicted to assist in binding to carbohydrates that are encountered in the mucin core structure.
Further information on how B. bifidum metabolizes mucin has been obtained from functional genomic approaches, including proteomics- as well as transcriptomics-based analyses [32]. The main B. bifidum enzymes that are responsible for mucin breakdown include extracellular enzymes, like putative exo-α-sialidases, as well as a predicted 1,2-α-L-fucosidase and 1,3/4-α-L-fucosidase, and a putative cell wall-anchored endo-α-N-acetylgalactosaminidase [41,42,43]. Further enzymes that are predicted to be involved in mucin metabolism encompass N-acetyl-β-hexosaminidases and β-galactosidases. The mucin-breakdown features of B. bifidum species is also supported by the occurrence of glycan carriers harboring several families, like the ATP-binding cassette (ABC-type), phosphoenolpyruvate phosphotransferase system (PEP-PTS), and major facilitator superfamily (MFS).
Specifically, the chromosome of B. bifidum PRL2010 contains a DNA region encompassing eight genes that are predicted to code for enzymes responsible for the hydrolysis of galacto-N-biose, which constitutes one of the key structures of mucin-oligosaccharides. The possible metabolic scenario employed by B. bifidum PRL2010 involves the action of extracellular enzymes such as exo-α-sialidases and 1,2-α-/α-1,3/4-L-fucosidases that are responsible for de-sialylidation and de-fucosylation, respectively, thus allowing B. bifidum access to mucin-derived galacto-N-biose [42]. This mucin-derived substrate is then further metabolized by other enzymes coded by PRL2010 like lacto-N-biosidase and endo-α-N-acetylgalactosaminidase [42,44,45].
Comparative genomic analyses encompassing other public accessible B. bifidum chromosomes showed that the estimated genetic background that is responsible for mucin metabolism is well conserved in this bifidobacterial taxon [29].
Mucin breakdown is predicted to diminish the mucin layer and, therefore, decrease the defensive barrier of the intestinal mucosa, rendering this activity as a potentially unwanted occurrence. However, one may also argue that mucin breakdown has evolved as a “host-settler mechanism”. In fact, mucin synthesis in the gut typically starts upon delivery, then occurs for several months and arrives at its mature level at about one year of life [46]. Fascinatingly, mucin metabolism as performed by B. bifidum could activate enhanced production of mucin, thus enhancing the depth of the mucus layer wrapping the mucosa and, thus, strengthening the epithelial barrier function [47].

7. The Capability of B. bifidum to Interact with its Host by Extracellular Structures

Commensal gut bacteria are known to establish a cross-talk with the human host through various structures including pili, nanotubes, capsular structures, S-layers proteins, and flagella [48,49,50,51]. Interestingly, many of these structures are also used to drive the microbe–microbe dialogue occurring between the different players of the human gut microbiota, comprising bifidobacterial strains [52].
Recently, the genetic variety of predicted sortase-dependent pili of bifidobacteria was assessed, allowing the construction of a bifidobacterial sortase-dependent fimbriome database [31]. Interestingly, the identified bifidobacterial fimbrione was shown to be highly variable among the various bifidobacterial (sub)species, possibly as a result of horizontal gene transfer events [31]. Transcriptome experiments and binding assays involving different substrates has demonstrated the importance of bifidobacterial pili in modulating several adhesion capabilities of bifidobacteria to glycans and to extracellular matrix proteins [31], thus corroborating the ecological fitness of bifidobacteria in the human intestine.
Amongst the fimbriome of members of the Bifidobacterium genus, the best experimentally and functionally characterized sortase-dependent pili are those of B. bifidum PRL2010 [30,53,54]. Such extracellular structures are composed of major and minor subunits, which are covalently assembled through the action of a sortase. These proteins are typically encoded by three adjacent genes, referred to as a sortase-dependent pilus locus. The genome of B. bifidum PRL2010 includes three distinct sortase-dependent pilus loci of which only two were predicted to be genetically intact, while the third appears to be non-functional due to a frameshift within the coding region of the gene encoding major pilus subunit [55].
Microarray-mediated transcriptome analysis of B. bifidum PRL2010 following murine colonization and following interaction with human cell lines revealed a significant increase in the transcription of the genes including two sortase-dependent pili, named pil2 and pil3 [55]. Heterologous expression of pil2 and pil3 in the non-piliated Lactococcus lactis demonstrated that both pili are promoting adhesion to human gut mucosa by extracellular matrix (ECM) proteins. ECM de-glycosylation triggered a drastic drop in PRL2010 pilus-based binding ability compared to untreated ECM [55], suggesting that N- and/or O-linked glycoproteins are involved in the adhesion of B. bifidum PRL2010 pili to ECM.
Recombinant piliated L. lactis cells were also displayed to induce an increased tumor necrosis factor alpha (TNFα) response following administration to mice when associated with their non-piliated parent, suggesting that PRL2010 sortase-dependent pili also elicit immunomodulatory activity [55]. The biological significance of this finding may be that triggering TNFα production by pili synthetized by B. bifidum PRL2010 represents a cross-talk signal by an early colonizer of the human gut [7,14,53,56]. In this framework, it is worth mentioning that TNFα superfamily members are not only connected to the manifestation of inflammatory diseases [57], but also exert a key role in the rejection of tumors and the response to infections [58,59]. Furthermore, TNFα induction may be crucial for the establishment of cross-talk among immune cells and commensal bacteria without causing inflammation or other harmful effects [60].
In addition, sortase-dependent pili of PRL2010 were shown to promote self-aggregation as well as aggregation of PRL2010 cells with cells of other gut microorganisms including bifidobacteria and lactobacilli [55,61]. This suggests that these extracellular proteins are responsible for the generation of macro-colonies or biofilm in the environment.
Interestingly, the B. bifidum species is the only Bifidobacterium taxon without capsular polysaccharides genes which seems to be absent due to a recent deletion event [62].
Nevertheless, a small number of isolates of the B. bifidum species have recently been shown to produce a complex mixture of polysaccharides, i.e., a negatively charged phospho-glycero-β-galactofuranan (PGβG), four neutral polysaccharides being β-(1-6)-glucan, β-(1-4)-galactan, β-(1-6)-galactan and β-galactofuranan and finally, starch. Interestingly, these two fractions were shown to exert distinct immune responses when assayed on dendritic cells. PGβG increased pro-inflammatory immune responses by enhancing the levels of TNFα, whereas CSGG induced immunosuppressive regulatory T cells and interleukin-10 [63].
Many enteric bacteria possess the ability to transform bile salts biochemically through the deconjugation of bile salts via the action of a Bile Salt Hydrolase (BSH). The detergent action of bile salts helps in the reduction of intestinal absorption and increased fecal elimination [64], with an overall hypolipidemic in vivo effect [65]. Recent screening analyses revealed that BSH activity is widely distributed in many bifidobacterial species (i.e., B. longum subsp. longum, B. longum subsp. infantis, B. animalis, and B. bifidum) [66,67]. B. bifidum PRL2010 shows noteworthy BSH action on conjugated secondary bile salts (glycodeoxycholate and taurodeoxycholate) [68,69] in line with what has been observed in other Bifidobacterium species, such as B. animalis and B. longum [70]. The importance of this enzyme for microorganisms is also so far not well characterized. In fact, various suppositions have been suggested, but the specific biological meaning of bile salt hydrolases at this time remains unidentified. It has been theorized that the conjugation of bile salts may be a tool that bacteria use in order to produce bile detoxication and thus, these enzymes may exert a key function in microbial bile tolerance and survival in the human gut [71].

8. The Ecological Role of B. bifidum Species in the Infant Gut Microbiota

As described above, bifidobacteria are amongst the first microbial colonizers of the infant gut [21]. Upon delivery, the gut environment is subject to massive bacterial colonization, in particular bifidobacteria, which is acquired through vertical transmission involving direct mother–baby contact at birth and is supported through breastfeeding [21]. Notably, under “non-natural” conditions, such as a caesarian delivery or baby feeding with reconstituted or formula milk, the newborn gut may undergo rapid colonization by environmental microorganisms that normally do not occur at that stage or in such abundance, which may provoke long lasting health effects on the host [21]. Supplementation of health-promoting bacteria with, for example, bifidobacteria that are typically found in the infant gut during this very critical window of time may be crucial in terms of preventing the establishment of an aberrant microbiota with its associated negative health implications.
In this context, it has been shown that members of the B. bifidum species are able to exert a cooperative syntrophic effect to benefit other elements of the gut microbiota, particularly with regards to other members of (healthy) infant-associated bifidobacterial communities [52]. Notably, in vivo murine experiments involving B. bifidum PRL2010 showed that in contrast to other bifidobacterial strains like B. longum subsp. infantis ATCC15697 representing another member of the infant gut microbiota, PRL2010 possesses cross-feeding properties that support the growth of other bifidobacteria and exhibits a high interaction index, which measures the level of microbe–microbe interaction elicited by a bacterial strain [52]. The cross-feeding features of B. bifidum PRL2010 have been further analyzed by in vitro tests aimed at investigating how co-cultivation of PRL2010 cells with other strains is able to increase the growth abilities of these later as compared to growth yields obtained when these strains were cultivated independently. Remarkably, the cross-feeding characteristics of B. bifidum PRL2010 were evident when this strain was cultivated on host-derived glycans like mucin or HMOs [72,73] or on plant-derived carbohydrates such as starch and xylan [74]. In this context, it has been shown that the metabolism of these complex carbohydrates by PRL2010 causes the release of rather simple carbohydrates which then become accessible to other members of the (bifido)bacterial community. The trophic interaction of B. bifidum with other bifidobacterial species and thus its capability to sustain the growth of other bifidobacteria has recently been highlighted for B. bifidum ATCC 15696—notably, ATCC 15696 hydrolyzes 2’-O-fucosyl-lactose, a major fucosylated HMO, though it does not employ fucose that is left in the culture broth [75]. Nevertheless, fucose is a growth substrate for Bifidobacterium breve 24b. Notably, the release of fucose by B. bifidum (the donor), thereby allowing the growth of B. breve (the beneficiary), supports the concept of syntropy [75].
The presence of B. bifidum PRL2010 cells in the murine gut has been shown to provoke an expansion of the murine gut glycobiome, i.e., the overall genetic arsenal that is involved in carbohydrate metabolism, toward enzymatic degradation of plant- and host-derived glycans [52].
Another interesting activity of the members of the B. bifidum species in relation to their interaction with other members of the human gut microbiota, i.e., microbe–microbe cross-talk, is represented by the ability of strains of this species to produce sortase-dependent pili, particularly when residing in their natural ecological niche, the mammalian gut [30,53]. Pili also appear, apart from playing a role in host interaction as described above, to be crucial for the establishment of physical contacts with other bifidobacterial/gut commensal cells through cell aggregation [61].

9. Conclusions

Many B. bifidum strains have been described to elicit beneficial effects, including antibacterial features against pathogens like Helicobacter pylori [76,77], drop of apoptosis in the intestinal mucosa of premature babies affected by necrotizing enterocolitis [78], modulation of the host-immune system [79,80], and alleviation of inflammatory activities associated with certain chronic gut dysfunctions [81,82]. Nevertheless, none of these characteristics have been investigated in any detail within their natural ecological context, the infant gut. In such an environment, bifidobacterial cells are expected to drive important functions in terms of priming the immune system, enhancing the mucus layer, and modulating the establishment of a correct microbiota homeostasis. Nevertheless, in order to exploit a possible probiotic activity in the human gut, bacterial cells must be viable in this body-compartment. Furthermore, they should also colonize, compete, and persist in the human intestine. Thus, B. bifidum probiotic strains need to survive under gastrointestinal challenges [83] and should be able to colonize the human gut and consequently impact on the intestinal resident microbial communities. Another notable feature that is shown by human health promoting microorganisms is their ability to displace and compete with pathogens. In this context, in vitro experiments have demonstrated that B. bifidum PRL2010 cells strongly inhibit the adhesion of enteropathogens including Escherichia coli and Cronobacter sakazakii [83], which are frequently implicated in severe gastrointestinal diseases in infants.
Notably, for most bifidobacterial strains that are currently commercialized as probiotics, little information is available about the molecular mechanisms supporting their claimed probiotic actions.
Apart from the exploitation of B. bifidum strains as probiotic bacteria for infants, another possible use of these microorganisms includes the treatment of women during pregnancy. Such a probiotic administration strategy might be key in order to modulate the mother’s gut engraftment by B. bifidum cells prior to delivery and, therefore, to assure the establishment of an appropriate bifidobacterial community that will then be vertically inherited by the newborn. Nonetheless, various clinical testing involving B. bifidum must be carried out before meaningful probiotic claims (for humans) can be made.

Funding

This research was funded by GenProbio srl to Marco Ventura and Francesca Turroni and by EU Joint Programming Initiative—A Healthy Diet for a Healthy Life (JPI HDHL, http://www.healthydietforhealthylife.eu/) to Prof. Douwe van Sinderen (in conjunction with Science Foundation Ireland [SFI], Grant number 15/JP-HDHL/3280). D.v.S. is member of APC microbiome Ireland, which is funded by SFI through the Irish Government’s National Development Plan (Grant Numbers SFI/12/RC/2273-P1 and SFI/12/RC/2273-P2). The study is supported by Fondazione Cariparma, under the TeachInParma Project (DV).

Acknowledgments

This research benefited from the HPC (High Performance Computing) facility of the University of Parma, Italy.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Goodfellow, M.; Kampfer, P.; Busse, H.; Trujillo, M.E.; Suzuki, K.; Ludwig, W.; Whitman, W.B. Bergey’s Manual of Systematic Bacteriology, 2nd ed.; Springer: New York, NY, USA, 2012. [Google Scholar]
  2. Ventura, M.; Canchaya, C.; Tauch, A.; Chandra, G.; Fitzgerald, G.F.; Chater, K.F.; van Sinderen, D. Genomics of Actinobacteria: Tracing the evolutionary history of an ancient phylum. Microbiol. Mol. Biol. Rev. MMBR 2007, 71, 495–548. [Google Scholar] [CrossRef] [PubMed]
  3. Turroni, F.; van Sinderen, D.; Ventura, M. Genomics and ecological overview of the genus Bifidobacterium. Int. J. Food Microbiol. 2011, 149, 37–44. [Google Scholar] [CrossRef] [PubMed]
  4. Avershina, E.; Lundgard, K.; Sekelja, M.; Dotterud, C.; Storro, O.; Oien, T.; Johnsen, R.; Rudi, K. Transition from infant- to adult-like gut microbiota. Environ. Microbiol. 2016, 18, 2226–2236. [Google Scholar] [CrossRef] [PubMed]
  5. Milani, C.; Mancabelli, L.; Lugli, G.A.; Duranti, S.; Turroni, F.; Ferrario, C.; Mangifesta, M.; Viappiani, A.; Ferretti, P.; Gorfer, V.; et al. Exploring vertical transmission of bifidobacteria from mother to child. Appl. Environ. Microbiol. 2015, 81, 7078–7087. [Google Scholar] [CrossRef] [PubMed]
  6. Milani, C.; Mangifesta, M.; Mancabelli, L.; Lugli, G.A.; James, K.; Duranti, S.; Turroni, F.; Ferrario, C.; Ossiprandi, M.C.; van Sinderen, D.; et al. Unveiling bifidobacterial biogeography across the mammalian branch of the tree of life. ISME J. 2017, 11, 2834–2847. [Google Scholar] [CrossRef] [PubMed]
  7. Turroni, F.; Peano, C.; Pass, D.A.; Foroni, E.; Severgnini, M.; Claesson, M.J.; Kerr, C.; Hourihane, J.; Murray, D.; Fuligni, F.; et al. Diversity of bifidobacteria within the infant gut microbiota. PLoS ONE 2012, 7, e36957. [Google Scholar] [CrossRef]
  8. Duranti, S.; Turroni, F.; Lugli, G.A.; Milani, C.; Viappiani, A.; Mangifesta, M.; Gioiosa, L.; Palanza, P.; van Sinderen, D.; Ventura, M. Genomic characterization and transcriptional studies of the starch-utilizing strain Bifidobacterium adolescentis 22L. Appl. Environ. Microbiol. 2014, 80, 6080–6090. [Google Scholar] [CrossRef]
  9. Duranti, S.; Milani, C.; Lugli, G.A.; Mancabelli, L.; Turroni, F.; Ferrario, C.; Mangifesta, M.; Viappiani, A.; Sanchez, B.; Margolles, A.; et al. Evaluation of genetic diversity among strains of the human gut commensal Bifidobacterium adolescentis. Sci. Rep. 2016, 6, 23971. [Google Scholar] [CrossRef]
  10. Turroni, F.; Milani, C.; Duranti, S.; Ferrario, C.; Lugli, G.A.; Mancabelli, L.; van Sinderen, D.; Ventura, M. Bifidobacteria and the infant gut: An example of co-evolution and natural selection. Cell. Mol. Life Sci. CMLS 2018, 75, 103–118. [Google Scholar] [CrossRef]
  11. Ventura, M.; Turroni, F.; Lugli, G.A.; van Sinderen, D. Bifidobacteria and humans: our special friends, from ecological to genomics perspectives. J. Sci. Food Agr. 2014, 94, 163–168. [Google Scholar] [CrossRef]
  12. Eckburg, P.B.; Bik, E.M.; Bernstein, C.N.; Purdom, E.; Dethlefsen, L.; Sargent, M.; Gill, S.R.; Nelson, K.E.; Relman, D.A. Diversity of the human intestinal microbial flora. Science 2005, 308, 1635–1638. [Google Scholar] [CrossRef] [PubMed]
  13. Mueller, S.; Saunier, K.; Hanisch, C.; Norin, E.; Alm, L.; Midtvedt, T.; Cresci, A.; Silvi, S.; Orpianesi, C.; Verdenelli, M.C.; et al. Differences in fecal microbiota in different European study populations in relation to age, gender, and country: A cross-sectional study. Appl. Environ. Microbiol. 2006, 72, 1027–1033. [Google Scholar] [CrossRef] [PubMed]
  14. Duranti, S.; Lugli, G.A.; Mancabelli, L.; Armanini, F.; Turroni, F.; James, K.; Ferretti, P.; Gorfer, V.; Ferrario, C.; Milani, C.; et al. Maternal inheritance of bifidobacterial communities and bifidophages in infants through vertical transmission. Microbiome 2017, 5, 66. [Google Scholar] [CrossRef] [PubMed]
  15. Alessandri, G.; Milani, C.; Duranti, S.; Mancabelli, L.; Ranjanoro, T.; Modica, S.; Carnevali, L.; Statello, R.; Bottacini, F.; Turroni, F.; et al. Ability of bifidobacteria to metabolize chitin-glucan and its impact on the gut microbiota. Sci. Rep. 2019, 9, 5755. [Google Scholar] [CrossRef] [PubMed]
  16. Turroni, F.; Marchesi, J.R.; Foroni, E.; Gueimonde, M.; Shanahan, F.; Margolles, A.; van Sinderen, D.; Ventura, M. Microbiomic analysis of the bifidobacterial population in the human distal gut. ISME J. 2009, 3, 745–751. [Google Scholar] [CrossRef]
  17. Palmer, C.; Bik, E.M.; DiGiulio, D.B.; Relman, D.A.; Brown, P.O. Development of the human infant intestinal microbiota. PLoS Biol. 2007, 5, e177. [Google Scholar] [CrossRef]
  18. Milani, C.; Lugli, G.A.; Turroni, F.; Mancabelli, L.; Duranti, S.; Viappiani, A.; Mangifesta, M.; Segata, N.; van Sinderen, D.; Ventura, M. Evaluation of bifidobacterial community composition in the human gut by means of a targeted amplicon sequencing (ITS) protocol. FEMS Microbiol. Ecol. 2014, 90, 493–503. [Google Scholar] [CrossRef]
  19. Duranti, S.; Lugli, G.A.; Milani, C.; James, K.; Mancabelli, L.; Turroni, F.; Alessandri, G.; Mangifesta, M.; Mancino, W.; Ossiprandi, M.C.; et al. Bifidobacterium bifidum and the infant gut microbiota: An intriguing case of microbe-host co-evolution. Environ. Microbiol. 2019. [Google Scholar] [CrossRef]
  20. Tannock, G.W.; Lawley, B.; Munro, K.; Gowri Pathmanathan, S.; Zhou, S.J.; Makrides, M.; Gibson, R.A.; Sullivan, T.; Prosser, C.G.; Lowry, D.; et al. Comparison of the compositions of the stool microbiotas of infants fed goat milk formula, cow milk-based formula, or breast milk. Appl. Environ. Microbiol. 2013, 79, 3040–3048. [Google Scholar] [CrossRef]
  21. Milani, C.; Duranti, S.; Bottacini, F.; Casey, E.; Turroni, F.; Mahony, J.; Belzer, C.; Delgado Palacio, S.; Arboleya Montes, S.; Mancabelli, L.; et al. The First Microbial Colonizers of the Human Gut: Composition, Activities, and Health Implications of the Infant Gut Microbiota. Microbiol. Mol. Biol. Rev. 2017, 81. [Google Scholar] [CrossRef]
  22. Ferrario, C.; Duranti, S.; Milani, C.; Mancabelli, L.; Lugli, G.A.; Turroni, F.; Mangifesta, M.; Viappiani, A.; Ossiprandi, M.C.; van Sinderen, D.; et al. Exploring Amino Acid Auxotrophy in Bifidobacterium bifidum PRL2010. Front. Microbiol. 2015, 6, 1331. [Google Scholar] [CrossRef] [PubMed]
  23. Makino, H.; Kushiro, A.; Ishikawa, E.; Muylaert, D.; Kubota, H.; Sakai, T.; Oishi, K.; Martin, R.; Ben Amor, K.; Oozeer, R.; et al. Transmission of intestinal Bifidobacterium longum subsp. longum strains from mother to infant, determined by multilocus sequencing typing and amplified fragment length polymorphism. Appl. Environ. Microbiol. 2011, 77, 6788–6793. [Google Scholar] [CrossRef] [PubMed]
  24. Ferretti, P.; Pasolli, E.; Tett, A.; Asnicar, F.; Gorfer, V.; Fedi, S.; Armanini, F.; Truong, D.T.; Manara, S.; Zolfo, M.; et al. Mother-to-Infant Microbial Transmission from Different Body Sites Shapes the Developing Infant Gut Microbiome. Cell Host Microbe 2018, 24, 133–145.e135. [Google Scholar] [CrossRef] [PubMed]
  25. Yassour, M.; Jason, E.; Hogstrom, L.J.; Arthur, T.D.; Tripathi, S.; Siljander, H.; Selvenius, J.; Oikarinen, S.; Hyoty, H.; Virtanen, S.M.; et al. Strain-Level Analysis of Mother-to-Child Bacterial Transmission during the First Few Months of Life. Cell Host Microbe 2018, 24, 146–154.e144. [Google Scholar] [CrossRef]
  26. Milani, C.; Duranti, S.; Lugli, G.A.; Bottacini, F.; Strati, F.; Arioli, S.; Foroni, E.; Turroni, F.; van Sinderen, D.; Ventura, M. Comparative genomics of Bifidobacterium animalis subsp. lactis reveals a strict monophyletic bifidobacterial taxon. Appl. Environ. Microbiol. 2013, 79, 4304–4315. [Google Scholar] [CrossRef]
  27. Lugli, G.A.; Mancino, W.; Milani, C.; Duranti, S.; Mancabelli, L.; Napoli, S.; Mangifesta, M.; Viappiani, A.; Anzalone, R.; Longhi, G.; et al. Dissecting the Evolutionary Development of the Species Bifidobacterium animalis through Comparative Genomics Analyses. Appl. Environ. Microbiol. 2019, 85. [Google Scholar] [CrossRef]
  28. Bottacini, F.; O’Connell Motherway, M.; Kuczynski, J.; O’Connell, K.J.; Serafini, F.; Duranti, S.; Milani, C.; Turroni, F.; Lugli, G.A.; Zomer, A.; et al. Comparative genomics of the Bifidobacterium breve taxon. BMC Genom. 2014, 15, 170. [Google Scholar] [CrossRef]
  29. Duranti, S.; Milani, C.; Lugli, G.A.; Turroni, F.; Mancabelli, L.; Sanchez, B.; Ferrario, C.; Viappiani, A.; Mangifesta, M.; Mancino, W.; et al. Insights from genomes of representatives of the human gut commensal Bifidobacterium bifidum. Environ. Microbiol. 2015, 17, 2515–2531. [Google Scholar] [CrossRef]
  30. Bottacini, F.; van Sinderen, D.; Ventura, M. Omics of bifidobacteria: Research and insights into their health-promoting activities. Biochem. J. 2017, 474, 4137–4152. [Google Scholar] [CrossRef]
  31. Milani, C.; Mangifesta, M.; Mancabelli, L.; Lugli, G.A.; Mancino, W.; Viappiani, A.; Faccini, A.; van Sinderen, D.; Ventura, M.; Turroni, F. The Sortase-Dependent Fimbriome of the Genus Bifidobacterium: Extracellular Structures with Potential to Modulate Microbe-Host Dialogue. Appl. Environ. Microbiol. 2017, 83. [Google Scholar] [CrossRef]
  32. Turroni, F.; Bottacini, F.; Foroni, E.; Mulder, I.; Kim, J.H.; Zomer, A.; Sanchez, B.; Bidossi, A.; Ferrarini, A.; Giubellini, V.; et al. Genome analysis of Bifidobacterium bifidum PRL2010 reveals metabolic pathways for host-derived glycan foraging. Proc. Natl. Acad. Sci. USA 2010, 107, 19514–19519. [Google Scholar] [CrossRef] [PubMed]
  33. Milani, C.; Lugli, G.A.; Duranti, S.; Turroni, F.; Mancabelli, L.; Ferrario, C.; Mangifesta, M.; Hevia, A.; Viappiani, A.; Scholz, M.; et al. Bifidobacteria exhibit social behavior through carbohydrate resource sharing in the gut. Sci. Rep. 2015, 5, 15782. [Google Scholar] [CrossRef] [PubMed]
  34. Milani, C.; Turroni, F.; Duranti, S.; Lugli, G.A.; Mancabelli, L.; Ferrario, C.; van Sinderen, D.; Ventura, M. Genomics of the Genus Bifidobacterium Reveals Species-Specific Adaptation to the Glycan-Rich Gut Environment. Appl. Environ. Microbiol. 2016, 82, 980–991. [Google Scholar] [CrossRef] [PubMed]
  35. Cantarel, B.L.; Coutinho, P.M.; Rancurel, C.; Bernard, T.; Lombard, V.; Henrissat, B. The Carbohydrate-Active EnZymes database (CAZy): An expert resource for Glycogenomics. Nucleic Acids Res 2009, 37, D233–D238. [Google Scholar] [CrossRef] [PubMed]
  36. Pokusaeva, K.; Fitzgerald, G.F.; van Sinderen, D. Carbohydrate metabolism in Bifidobacteria. Genes Nutr. 2011, 6, 285–306. [Google Scholar] [CrossRef] [PubMed]
  37. Podolsky, D.K. Oligosaccharide structures of isolated human colonic mucin species. J. Biol. Chem. 1985, 260, 15510–15515. [Google Scholar]
  38. Forstner, J.F.; Oliver, M.G.; Sylvester, F.A. Production, Structure and Biologic Relevance of Gastrointestinal Mucins; Raven Press: New York, NY, USA, 1995; pp. 71–88. [Google Scholar]
  39. Turroni, F.; Milani, C.; van Sinderen, D.; Ventura, M. Genetic strategies for mucin metabolism in Bifidobacterium bifidum PRL2010: An example of possible human-microbe co-evolution. Gut Microbes 2011, 2, 183–189. [Google Scholar] [CrossRef]
  40. Ruas-Madiedo, P.; Gueimonde, M.; Fernandez-Garcia, M.; de los Reyes-Gavilan, C.G.; Margolles, A. Mucin degradation by Bifidobacterium strains isolated from the human intestinal microbiota. Appl. Environ. Microbiol. 2008, 74, 1936–1940. [Google Scholar] [CrossRef]
  41. Ashida, H.; Maki, R.; Ozawa, H.; Tani, Y.; Kiyohara, M.; Fujita, M.; Imamura, A.; Ishida, H.; Kiso, M.; Yamamoto, K. Characterization of two different endo-alpha-N-acetylgalactosaminidases from probiotic and pathogenic enterobacteria, Bifidobacterium longum and Clostridium perfringens. Glycobiology 2008, 18, 727–734. [Google Scholar] [CrossRef]
  42. Ashida, H.; Miyake, A.; Kiyohara, M.; Wada, J.; Yoshida, E.; Kumagai, H.; Katayama, T.; Yamamoto, K. Two distinct alpha-L-fucosidases from Bifidobacterium bifidum are essential for the utilization of fucosylated milk oligosaccharides and glycoconjugates. Glycobiology 2009, 19, 1010–1017. [Google Scholar] [CrossRef]
  43. Kiyohara, M.; Nakatomi, T.; Kurihara, S.; Fushinobu, S.; Suzuki, H.; Tanaka, T.; Shoda, S.; Kitaoka, M.; Katayama, T.; Yamamoto, K.; et al. alpha-N-acetylgalactosaminidase from infant-associated bifidobacteria belonging to novel glycoside hydrolase family 129 is implicated in alternative mucin degradation pathway. J. Biol. Chem. 2012, 287, 693–700. [Google Scholar] [CrossRef] [PubMed]
  44. Wakinaka, T.; Kiyohara, M.; Kurihara, S.; Hirata, A.; Chaiwangsri, T.; Ohnuma, T.; Fukamizo, T.; Katayama, T.; Ashida, H.; Yamamoto, K. Bifidobacterial alpha-galactosidase with unique carbohydrate-binding module specifically acts on blood group B antigen. Glycobiology 2013, 23, 232–240. [Google Scholar] [CrossRef] [PubMed]
  45. Shimada, Y.; Watanabe, Y.; Wakinaka, T.; Funeno, Y.; Kubota, M.; Chaiwangsri, T.; Kurihara, S.; Yamamoto, K.; Katayama, T.; Ashida, H. alpha-N-Acetylglucosaminidase from Bifidobacterium bifidum specifically hydrolyzes alpha-linked N-acetylglucosamine at nonreducing terminus of O-glycan on gastric mucin. Appl. Microbiol. Biotechnol. 2015, 99, 3941–3948. [Google Scholar] [CrossRef] [PubMed]
  46. Hooper, L.V.; Xu, J.; Falk, P.G.; Midtvedt, T.; Gordon, J.I. A molecular sensor that allows a gut commensal to control its nutrient foundation in a competitive ecosystem. Proc. Natl. Acad. Sci. USA 1999, 96, 9833–9838. [Google Scholar] [CrossRef] [PubMed]
  47. Rokhsefat, S.; Lin, A.F.; Comelli, E.M. Mucin-Microbiota Interaction During Postnatal Maturation of the Intestinal Ecosystem: Clinical Implications. Dig. Dis. Sci. 2016, 61, 1473–1486. [Google Scholar] [CrossRef] [PubMed]
  48. Danne, C.; Dramsi, S. Pili of gram-positive bacteria: Roles in host colonization. Res. Microbiol. 2012, 163, 645–658. [Google Scholar] [CrossRef]
  49. Dubey, G.P.; Ben-Yehuda, S. Intercellular nanotubes mediate bacterial communication. Cell 2011, 144, 590–600. [Google Scholar] [CrossRef]
  50. Gerbino, E.; Carasi, P.; Mobili, P.; Serradell, M.A.; Gomez-Zavaglia, A. Role of S-layer proteins in bacteria. World J. Microbiol. Biotechnol. 2015, 31, 1877–1887. [Google Scholar] [CrossRef]
  51. Rajagopal, M.; Walker, S. Envelope Structures of Gram-Positive Bacteria. Curr. Top. Microbiol. Immunol. 2017, 404, 1–44. [Google Scholar] [CrossRef]
  52. Turroni, F.; Milani, C.; Duranti, S.; Mancabelli, L.; Mangifesta, M.; Viappiani, A.; Lugli, G.A.; Ferrario, C.; Gioiosa, L.; Ferrarini, A.; et al. Deciphering bifidobacterial-mediated metabolic interactions and their impact on gut microbiota by a multi-omics approach. ISME J. 2016. [Google Scholar] [CrossRef]
  53. Turroni, F.; Duranti, S.; Bottacini, F.; Guglielmetti, S.; Van Sinderen, D.; Ventura, M. Bifidobacterium bifidum as an example of a specialized human gut commensal. Front. Microbiol. 2014, 5, 437. [Google Scholar] [CrossRef] [PubMed]
  54. Foroni, E.; Serafini, F.; Amidani, D.; Turroni, F.; He, F.; Bottacini, F.; O’Connell Motherway, M.; Viappiani, A.; Zhang, Z.; Rivetti, C.; et al. Genetic analysis and morphological identification of pilus-like structures in members of the genus Bifidobacterium. Microb. Cell Factories 2011, 10 (Suppl. 1), S16. [Google Scholar] [CrossRef] [PubMed]
  55. Turroni, F.; Serafini, F.; Foroni, E.; Duranti, S.; O’Connell Motherway, M.; Taverniti, V.; Mangifesta, M.; Milani, C.; Viappiani, A.; Roversi, T.; et al. Role of sortase-dependent pili of Bifidobacterium bifidum PRL2010 in modulating bacterium-host interactions. Proc. Natl. Acad. Sci. USA 2013, 110, 11151–11156. [Google Scholar] [CrossRef] [PubMed]
  56. Mancino, W.; Duranti, S.; Mancabelli, L.; Longhi, G.; Anzalone, R.; Milani, C.; Lugli, G.A.; Carnevali, L.; Statello, R.; Sgoifo, A.; et al. Bifidobacterial Transfer from Mother to Child as Examined by an Animal Model. Microorganisms 2019, 7, 293. [Google Scholar] [CrossRef]
  57. Yasutake, N.; Matsuzaki, T.; Kimura, K.; Hashimoto, S.; Yokokura, T.; Yoshikai, Y. The role of tumor necrosis factor (TNF)-alpha in the antitumor effect of intrapleural injection of Lactobacillus casei strain Shirota in mice. Med. Microbiol. Immunol. 1999, 188, 9–14. [Google Scholar] [CrossRef]
  58. Wajant, H.; Pfizenmaier, K.; Scheurich, P. Tumor necrosis factor signaling. Cell Death Differ. 2003, 10, 45–65. [Google Scholar] [CrossRef]
  59. Lebeer, S.; Vanderleyden, J.; De Keersmaecker, S.C. Host interactions of probiotic bacterial surface molecules: Comparison with commensals and pathogens. Nat. Rev. Microbiol. 2010, 8, 171–184. [Google Scholar] [CrossRef]
  60. Galdeano, C.M.; de Moreno de LeBlanc, A.; Vinderola, G.; Bonet, M.E.; Perdigon, G. Proposed model: Mechanisms of immunomodulation induced by probiotic bacteria. Clin. Vaccine Immunol. CVI 2007, 14, 485–492. [Google Scholar] [CrossRef]
  61. Turroni, F.; Serafini, F.; Mangifesta, M.; Arioli, S.; Mora, D.; van Sinderen, D.; Ventura, M. Expression of sortase-dependent pili of Bifidobacterium bifidum PRL2010 in response to environmental gut conditions. FEMS Microbiol. Lett. 2014, 357, 23–33. [Google Scholar] [CrossRef]
  62. Ferrario, C.; Milani, C.; Mancabelli, L.; Lugli, G.A.; Duranti, S.; Mangifesta, M.; Viappiani, A.; Turroni, F.; Margolles, A.; Ruas-Madiedo, P.; et al. Modulation of the eps-ome transcription of bifidobacteria through simulation of human intestinal environment. FEMS Microbiol. Ecol. 2016, 92, fiw056. [Google Scholar] [CrossRef]
  63. Speciale, I.; Verma, R.; Di Lorenzo, F.; Molinaro, A.; Im, S.H.; De Castro, C. Bifidobacterium bifidum presents on the cell surface a complex mixture of glucans and galactans with different immunological properties. Carbohydr. Polym. 2019, 218, 269–278. [Google Scholar] [CrossRef] [PubMed]
  64. Ooi, L.G.; Liong, M.T. Cholesterol-lowering effects of probiotics and prebiotics: A review of in vivo and in vitro findings. Int. J. Mol. Sci. 2010, 11, 2499–2522. [Google Scholar] [CrossRef] [PubMed]
  65. Kumar, M.; Nagpal, R.; Kumar, R.; Hemalatha, R.; Verma, V.; Kumar, A.; Chakraborty, C.; Singh, B.; Marotta, F.; Jain, S.; et al. Cholesterol-Lowering Probiotics as Potential Biotherapeutics for Metabolic Diseases. Exp. Diabetes Res. 2012, 2012, 902917. [Google Scholar] [CrossRef] [PubMed]
  66. Kim, G.B.; Miyamoto, C.M.; Meighen, E.A.; Lee, B.H. Cloning and characterization of the bile salt hydrolase genes (bsh) from Bifidobacterium bifidum strains. Appl. Environ. Microbiol. 2004, 70, 5603–5612. [Google Scholar] [CrossRef]
  67. Kim, G.B.; Yi, S.H.; Lee, B.H. Purification and characterization of three different types of bile salt hydrolases from Bifidobacterium strains. J. Dairy Sci. 2004, 87, 258–266. [Google Scholar] [CrossRef]
  68. Zanotti, I.; Turroni, F.; Piemontese, A.; Mancabelli, L.; Milani, C.; Viappiani, A.; Prevedini, G.; Sanchez, B.; Margolles, A.; Elviri, L.; et al. Evidence for cholesterol-lowering activity by Bifidobacterium bifidum PRL2010 through gut microbiota modulation. Appl. Microbiol. Biotechnol. 2015, 99, 6813–6829. [Google Scholar] [CrossRef]
  69. Jarocki, P.; Targonski, Z. Genetic diversity of bile salt hydrolases among human intestinal bifidobacteria. Curr. Microbiol. 2013, 67, 286–292. [Google Scholar] [CrossRef]
  70. Noriega, L.; Cuevas, I.; Margolles, A.; Los Reyes-Gavilan, C.G.D. Deconjugation and bile salts hydrolase activity by Bifidobacterium strains with acquired resistance to bile. Int. Dairy J. 2006, 16, 850–855. [Google Scholar] [CrossRef]
  71. De Smet, I.; Van Hoorde, L.; Vande Woestyne, M.; Christiaens, H.; Verstraete, W. Significance of bile salt hydrolytic activities of lactobacilli. J. Appl. Bacteriol. 1995, 79, 292–301. [Google Scholar] [CrossRef]
  72. Egan, M.; Motherway, O.C.M.; Kilcoyne, M.; Kane, M.; Joshi, L.; Ventura, M.; van Sinderen, D. Cross-feeding by Bifidobacterium breve UCC2003 during co-cultivation with Bifidobacterium bifidum PRL2010 in a mucin-based medium. BMC Microbiol. 2014, 14, 282. [Google Scholar] [CrossRef]
  73. Egan, M.; O’Connell Motherway, M.; Ventura, M.; van Sinderen, D. Metabolism of sialic acid by Bifidobacterium breve UCC2003. Appl. Environ. Microbiol. 2014, 80, 4414–4426. [Google Scholar] [CrossRef] [PubMed]
  74. Turroni, F.; Ozcan, E.; Milani, C.; Mancabelli, L.; Viappiani, A.; van Sinderen, D.; Sela, D.A.; Ventura, M. Glycan cross-feeding activities between bifidobacteria under in vitro conditions. Front. Microbiol. 2015, 6, 1030. [Google Scholar] [CrossRef] [PubMed]
  75. Centanni, M.; Ferguson, S.A.; Sims, I.M.; Biswas, A.; Tannock, G.W. Bifidobacterium bifidum ATCC 15696 and Bifidobacterium breve 24b Metabolic Interaction Based on 2′-O-Fucosyl-Lactose Studied in Steady-State Cultures in a Freter-Style Chemostat. Appl. Environ. Microbiol. 2019, 85, e02783-18. [Google Scholar] [CrossRef]
  76. Chenoll, E.; Casinos, B.; Bataller, E.; Astals, P.; Echevarria, J.; Iglesias, J.R.; Balbarie, P.; Ramon, D.; Genoves, S. Novel probiotic Bifidobacterium bifidum CECT 7366 strain active against the pathogenic bacterium Helicobacter pylori. Appl. Environ. Microbiol. 2011, 77, 1335–1343. [Google Scholar] [CrossRef] [PubMed]
  77. Shirasawa, Y.; Shibahara-Sone, H.; Iino, T.; Ishikawa, F. Bifidobacterium bifidum BF-1 suppresses Helicobacter pylori-induced genes in human epithelial cells. J. Dairy Sci. 2010, 93, 4526–4534. [Google Scholar] [CrossRef]
  78. Khailova, L.; Mount Patrick, S.K.; Arganbright, K.M.; Halpern, M.D.; Kinouchi, T.; Dvorak, B. Bifidobacterium bifidum reduces apoptosis in the intestinal epithelium in necrotizing enterocolitis. Am. J. Physiol. Gastrointest. Liver Physiol. 2010, 299, G1118–G1127. [Google Scholar] [CrossRef]
  79. Philippe, D.; Heupel, E.; Blum-Sperisen, S.; Riedel, C.U. Treatment with Bifidobacterium bifidum 17 partially protects mice from Th1-driven inflammation in a chemically induced model of colitis. Int. J. Food Microbiol. 2011, 149, 45–49. [Google Scholar] [CrossRef]
  80. Fu, Y.R.; Yi, Z.J.; Pei, J.L.; Guan, S. Effects of Bifidobacterium bifidum on adaptive immune senescence in aging mice. Microbiol. Immunol. 2010, 54, 578–583. [Google Scholar] [CrossRef] [PubMed]
  81. Guglielmetti, S.; Mora, D.; Gschwender, M.; Popp, K. Randomised clinical trial: Bifidobacterium bifidum MIMBb75 significantly alleviates irritable bowel syndrome and improves quality of life—A double-blind, placebo-controlled study. Aliment Pharm. Ther. 2011, 33, 1123–1132. [Google Scholar] [CrossRef] [PubMed]
  82. Mouni, F.; Aissi, E.; Hernandez, J.; Gorocica, P.; Bouquelet, S.; Zenteno, E.; Lascurain, R.; Garfias, Y. Effect of Bifidobacterium bifidum DSM 20082 cytoplasmic fraction on human immune cells. Immunol. Investig. 2009, 38, 104–115. [Google Scholar] [CrossRef] [PubMed]
  83. Serafini, F.; Strati, F.; Ruas-Madiedo, P.; Turroni, F.; Foroni, E.; Duranti, S.; Milano, F.; Perotti, A.; Viappiani, A.; Guglielmetti, S.; et al. Evaluation of adhesion properties and antibacterial activities of the infant gut commensal Bifidobacterium bifidum PRL2010. Anaerobe 2013, 21, 9–17. [Google Scholar] [CrossRef] [PubMed]
Figure 1. Schematic representation of the main properties exerted by B. bifidum in the human gut. Specifically, there is much evidence in the literature that shows that B. bifidum is able to interact with the host and with other members of gut microbiota by different mechanisms, activate the host immunity, adhere to gut mucosa with its extracellular structures, and metabolize host glycans, such as mucin. See the main text for further explanation.
Figure 1. Schematic representation of the main properties exerted by B. bifidum in the human gut. Specifically, there is much evidence in the literature that shows that B. bifidum is able to interact with the host and with other members of gut microbiota by different mechanisms, activate the host immunity, adhere to gut mucosa with its extracellular structures, and metabolize host glycans, such as mucin. See the main text for further explanation.
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