Background

Bifidobacteria are universally distributed in organisms that raise offspring by parental care including mammals, birds and social insects. Bifidobacteria are highly specialized organisms in using non-digestible oligosaccharides and a major part of their genomes is devoted to the utilization of carbon sources [1–5]. The proportion of genes related to carbohydrate transport and metabolism is higher in bifidobacteria than in Bacteroides, which are also characterized by their ability to utilize a variety of polysaccharides [6]. Host-specific adaption in regard to carbohydrate degradation has been suggested [7, 8]. Adult species, such as Bifidobacterium adolescentis and Bifidobacterium longum subsp. longum, are well equipped to degrade plant derived polysaccharides [4, 9]. Infant species, such as Bifidobacterium longum subsp. infantis and Bifidobacterium bifidum, are adapted to utilize human milk oligosaccharides (HMOs), one of the major glycan sources of breast milk [3, 10–12]. Primary components of HMOs are D-glucose, D-galactose, L-fucose, N-acetylglucosamine, and sialic acid. Lactose constitutes the reducing end, its galactose moiety can be fucosylated or sialylated to form 2′- or 3′-fucosyllactose (2′-FL or 3′-FL) or 3′- and 6′-sialyl-lactose (3′-SL or 6′-SL). Lactose can also be elongated with units of N-acetyllactosamine (Gal-β1-4GlcNAc) with its simplest form being Lacto-N-neotetraose (LNnT) [13].

The degradation of HMOs relies on a complex network of extracellular solute binding proteins, transporters and intra- or extracellular glycosyl hydrolases (GH). Both B. longum subsp. infantis and B. bifidum harbour several α-fucosidase and sialidases, hexosaminidases, lacto-N-biosidases, α- and β-galactosidases to degrade HMOs. B. longum subsp. infantis degrades HMOs intracellularly, while B. bifidum harvests HMOs extracellularly through the activity of membrane bound enzymes [3, 12]. Other species recovered from infant stool, such as Bifidobacterium breve, have a limited capacity to degrade HMOs, however, they can profit from cross-feeding of HMO constituents released by B. bifidum [14–16].

Bifidobacteria metabolize hexoses via the ‘bifid shunt’ with fructose-6-phosphoketolase being the key enzyme to theoretically yield 1.5 mol acetate, 1 mol lactate and 2.5 ATP from 1 mol glucose [17]. The ratios of lactate and acetate formed may vary with carbohydrate source and species, depending on whether the intermediate pyruvate is cleaved to acetyl phosphate and formate, or reduced to lactate [18]. Pentoses, such as xylose, are also fermented to lactate, acetate and possible formate [19]. There is little information available about bifidobacterial metabolism of desoxyhexoses, and rhamnose was not used by various species tested [6].

In a previous study, several Bifidobacterium strains were isolated from Kenyan infant stool, that were identified as B. longum, B. bifidum, B. breve, Bifidobacterium kashiwanohense, and Bifidobacterium pseudolongum [20]. B. kashiwanohense has only been isolated from a healthy Japanese infant [21] and from Kenyan anaemic infants [20]. B. pseudolongum has been frequently recovered from animal feces [22]. B. kashiwanohense, and B. pseudolongum are species not commonly associated with the infant bifidobacteria community, however, the presence of additional species in infant feces from developing countries, might reflect variations in diet and sanitary status. Little is known about the ability of these species to utilize HMOs.

Therefore it was the aim of this study to investigate HMO degradation by these newly obtained isolates. As we observed that beside B. longum subsp. infantis and B. bifidum, an isolate of B. longum subsp. suis, and strains of B. kashiwanohense were able to metabolise fucosyllactose, we further investigated metabolite formation during growth on L-fucose and fucosyllactose. Furthermore we analyzed genomes of the studied strains to elucidate possible genes and pathways involved in fucosyllactose degradation and L-fucose utilization through genome comparison.

Methods

Bacterial strains

Twenty-nine bifidobacterial strains were included in the initial HMO utilization screening (Table 1). Nineteen strains originated from stool samples of Kenyan infants [20] and ten reference strains were obtained from the Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (DSMZ, Braunschweig, Germany, Table 1). Kenyan isolates that had been previously typed to species level were additionally characterized on subspecies level using the (partial) 16S rRNA gene as marker. Briefly, DNA was extracted from overnight cultures using the PrepMan® Ultra protocol for pure culture (Thermo Fisher Scientific, Reinach, Switzerland). PCR amplification of partial 16S rRNA genes was performed using universal primers 518 F (5′-CCAGCAGCCGCGGTAATACG–3′) and 1391R (5′– GACGGGCGGTGTGTRCA–3′). PCR reaction mixtures (25 μL) contained 12.5 μL of 2× PCR MasterMix (Thermo Fisher Scientific), 0.2 μM of primers (Microsynth AG, Balgach, Switzerland) and 1 μl of template DNA. The cycling programme consisted of an initial denaturation of 5 min at 95 °C, followed by 32 cycles of denaturation for 30 s at 95 °C, annealing for 30 s at 52 °C, and extension for 1 min at 72 °C. Amplicons were sequenced by GATC Biotech (Konstanz, Germany).

Table 1 Strains used for HMO utilization screening
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For generation of phylogenetic trees, 16S rRNA gene sequences were aligned and cut using CLUSTALW implemented in BioEdit Version 7. Phylogenetic analysis of partial 16S rRNA gene sequences (772 bp) was performed using Maximum Likelihood Analysis implemented in MEGA6 [23], applying the Jones-Taylor-Thornton substitution model and default settings. Bootstrap support was calculated for 500 replicates, strains of Lactobacillus were applied as outgroup. Sequences are listed in the Additional file 1.

Based on partial 16S rRNA gene sequences, the Kenyan isolates were characterized as B. bifidum (n = 6), B. breve (n = 2), Bifidobacterium kashiwanohense (n = 3), Bifidobacterium pseudolongum subsp. globosum (n = 2), B. longum subsp. infantis (n = 5) and B. longum subsp. suis (n = 1) (Table 1).

Growth conditions

Bifidobacterium strains were routinely cultured at 37 °C in Wilkins-Chalgren Anaerobe Broth (Oxoid, Basel, Switzerland) supplemented with soya peptone (5 g L−1, Oxoid), Tween 80 (1 mL L−1, Sigma-Aldrich, Buchs, Switzerland), and fresh sterile filtered l-cysteine hydrochloride (0.5 g L−1, Sigma-Aldrich). Carbohydrate utilization profile of bifidobacteria was investigated in API 50CHL Medium (10 g L−1 bovine/porcine origin polypeptone, 5 g L−1 yeast extract, 1 mL−1 Tween 80, 2 g L−1 dipotassium phosphate, 5 g L−1 sodium acetate, 2 g L−1 di-ammonium citrate, 0.2 g L−1 magnesium sulphate heptahydrate, 0.05 g L−1 manganese sulphate monohydrate, 0.17 g L−1 bromocresol purple; BioMérieux, Genève, Switzerland). The pH of the API medium was adjusted to 7.5 to obtain a final pH of 7 after autoclaving at 121 °C for 15 min. Carbohydrates (concentration as indicated) were filter sterilized and added after autoclaving. Fresh sterile filtered l-cysteine hydrochloride was always added before cultivation (0.5 g L−1). Glucose, lactose, and L-fucose were obtained from Sigma-Aldrich, 2′-fucosyllactose (2′-FL, Fucα1-2Galβ1-4Glc), 3′-fucosyllactose (3′-FL, Fucα1-3Galβ1-4Glc), 3′-sialyl-lactose (3′-SL, NeuAcα2-3Galβ1-4Glc), 6′-sialyl-lactose (6′-SL, NeuAcα2-6Galβ1-4Glc), Lacto-N-neotetraose LNnT (Galβ1-4GlcNacβ1-3Galβ1-4Glc) were donated by Glycom A/S (Lyngby, Denmark).

Utilization of selected sugars and metabolite formation

Isolates derived from −80 °C stock cultures were streaked on supplemented Wilkins-Chalgren agar and were incubated anaerobically at 37 °C for two days. Single colonies of each isolate were subsequently incubated twice in supplemented Wilkins-Chalgren broth (10 ml, 1:10) at 37 °C for 20 h. To obtain working cultures, the supernatant was removed from overnight cultures, cells were washed, and re-suspended in same volume of 50 mM phosphate buffer, pH 6.5 (PB).

The initial HMO utilization screening was conducted in 96-well microtiter plates. Cell suspensions (20 μl) were added to 180 μl carbohydrate supplemented API medium (2′-FL and 3′-FL: 4 mM, 3′-SL and 6′-SL: 2 mM, LNnt: 1 mM; glucose: 6 mM; lactose: 3 mM). HMOs were also alone or supplied together (HMO mixture). Glucose and lactose were used to verify suitability of the assay. Strains were grown in independent triplicates under anaerobic condition (GENbag anaer; BioMérieux, Genève, Switzerland) at 37 °C for 48 h.

To investigate growth on fucosyllactose and L-fucose of selected strains, cell suspensions (50 μl) were added to 950 μl API medium supplied with 30 mM L-fucose, 2′-FL, or 3′-FL (28.0 and 27.0 mM, respectively). Strains were grown in independent triplicates under anaerobic condition at 37 °C for 48 h.

L-fucose utilization and metabolite analysis using high performance liquid chromatography with refractive index detection (HPLC-RI)

L-fucose, lactate, acetate and 1,2-PD concentrations were determined using HPLC (Merck-Hitachi, Darmstadt, Germany) equipped with an Aminex HPX-87H column (300 × 7.8 mm; BioRad, Cressier, Switzerland) and a RI detector. Samples were centrifuged at 13 000 g for 5 min at 4 °C. Supernatants (40 μL injection volume) were eluted with 10 mM H2SO4 at a flow rate of 0.6 ml min−1 at 40 °C. L-fucose, lactate, acetate, and 1,2-PD were quantified using external standards.

Analysis of HMO degradation using high performance anion exchange chromatography with pulsed amperometric detection (HPAEC-PAD)

HMO degradation was investigated using HPAEC-PAD, on Dionex IC3000 equipped with a CarbopacPA20 column (Thermo Fisher Scientific, Reinach, Switzerland) and an electrochemical detector with a gold electrode. Water (A), 200 mM NaOH (B), and 1 M Na-acetate (C) were used as solvents at a flow rate of 0.25 mL min−1. For HMO separation, a gradient of: 0 min 30.4 % B, 1.3 % C, 22 min 30.4 % B, and 11.34 % C followed by washing and regeneration was applied. HMOs and L-fucose were identified using external standards.

Genome sequencing

DNA was isolated from overnight culture of B. longum subsp. infantis TPY12-1 and B. longum subsp. suis BSM11-5 using the FastDNA SPIN Kit for Soil (MP Biomedicals, France) including a bead-beating procedure for cell disruption. Genome libraries of B. longum subsp. infantis TPY12-1 were sequenced with an Illumina HiSeq 2500 to obtain paired-end reads of 2x100 bp. Genome libraries of B. longum subsp. suis BSM11-5 were sequenced with an Illumina MiSeq to obtain paired-end reads of 2x150 bp. Sequencing was conducted at the Functional Genomic Center Zürich (FGCZ, Zürich, Switzerland).

Genome assembly and annotation

Genomes were assembled using Abyss v.1.9.0 for paired-end libraries implemented in Bio-Linux 8. The partial genomes were functionally annotated with RAST using default settings [24]. RAST annotations of genes of interest were verified using the BLAST tool implemented in RAST. Average nucleotide identity (ANI) was calculated using the online tool supplied by Rodriguez-R and Konstantinidis [25]. Carbohydrate-active enzymes were selectively confirmed based on similarity to the carbohydrate active enzyme (CAZy) database entries, and Pfam alignments implemented at the CAZymes Analysis Toolkit (CAT) [26]. Additionally, dbCAN was used for identification of carbohydrate active proteins which is based on a search for signature domains of every CAZyme family [27].

Results and discussion

Utilization of HMOs

We investigated growth of 19 bifidobacterial isolates of Kenyan infants and 10 culture collection strains (Table 1) in the presence of individual HMOs: 2′-FL, 3′-FL, 3′-SL, 6′-SL, and LNnT, and combined HMOs in API medium. All isolates were able to grow in the presence of glucose or lactose confirming the suitability of the assay (Table 2). Growth correlated with the degradation of the supplied HMOs as determined with HPAEC-PAD for selected strain-HMO combinations (Table 2).

Table 2 Degradation of HMOs by selected strains
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B. longum subsp. infantis utilized of 2′-FL, 3′-FL, 3′-SL and LNnT and degraded all HMOs when supplied together. B. bifidum grew in the presence of 2′-FL, 3′-FL and LNnT and also utilized 3′-SL and 6′-SL in HMO mixtures confirming adaptation of both species to HMO utilization, as reported before [3, 12]. Strains of B. bifidum liberated L-fucose and a second degradation product (Fig. 1, peak y) in the supernatant when grown in the presence of fucosyllactose while L-fucose accumulation or the release of any other degradation intermediate was not observed for B. longum subsp. infantis strains [28] (Fig. 1).

Fig. 1
figure 1

Degradation of 2′-FL (a) and 3′-FL (b) and accumulation of L-fucose. Shown are (1) unfermented control, (2) B. longum subsp. infantis DSM 20088, (3) B. kashiwanohense PV20-2, and (4) B. bifidum BSM28-1 as representatives of B. longum, B. kashiwanohense and B. bifidum isolates investigated. x, undefined media components; y, intermediate degradation compound of fucosyllactose metabolism; fuc, L-fucose

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All B. breve isolates were able to utilize LNnT as shown previously [15]. B. breve DSM 20213 also degraded 2′-FL and 3′-FL when grown with HMO mixtures.

B. longum subsp. suis has not been shown to utilize of HMOs [11]. Here we identified an isolate B. longum subsp. suis BSM11-5 able to metabolize 3′-FL and 2′-FL (Fig. 1). L-fucose was not accumulated when B. longum subsp. suis BSM11-5 was grown with 4 mM fucosyllactose.

Also, B. kashiwanohense DSM 21854 and the Kenyan isolates grew in the presence of 2′-FL and 3′-FL, thereby accumulating L-fucose and releasing compound y (Fig. 1). The amount of L-fucose released by B. kashiwanohense isolates was only about 12 % compared to the complete release of B. bifidum.

Strains of B. pseudolongum did not metabolize with any of the HMOs tested.

The ability to use fucosyllactose was thus identified as being a trait of several bifidobacteria species. B. longum subsp. suis and B. kashiwanohense have not considered infant bifidobacteria species, yet, the ability to utilize fucosyllactose points at adaptation to the infant gut.

L-fucose metabolism of bifidobacteria

Similar to B. bifidum, B. kashiwanohense excreted L-fucose into the supernatant [28]. L-fucose accumulation was not observed when B. longum subsp. infantis isolates and B. longum subsp. suis BSM 11–5 were grown in the presence of fucosyllactose.

Several clostridia and E. coli are capable of metabolizing L-fucose to 1,2 propanediol (1,2-PD) [29–33]. To investigate whether bifidobacteria are able to form 1,2-PD from L-fucose, B. longum subsp. infantis DSM 20088 and TPY12-1, B. longum subsp. suis BSM11-5, and B. kashiwanohense DSM 21854 were cultivated in API medium supplied with 30 mM L-fucose. As growth with L-fucose as sole carbohydrate source was unreliable, trace amounts of glucose (0.4 mM) were added to enable initial growth. When glucose was present, L-fucose was partially used and equimolar amounts of 1,2-PD were formed by the two strains of B. longum subsp. infantis, and by B. longum subsp. suis BSM11-5 (Table 3). B. longum subsp. suis BSM11-5 also formed equimolar amounts of acetate (Table 3). In contrast B. kashiwanohense DSM 21854 did not grow with L-fucose and trace amounts of glucose.

Table 3 L-fucose utilization and metabolite formation
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We also investigated whether 1,2-PD was formed from 3′-FL and 2′-FL (27 and 28 mM, respectively) (Table 4). In the presence of 2′-FL and 3′-FL, B. longum subsp. infantis DSM 20088 produced a lactate:acetate ratio of 2:3 as expected of the metabolism of hexoses through the bifid shunt [17] in addition, this strain produced 1,2-PD (Table 4).

Table 4 Fucosyllactose utilization, metabolite formation and L-fucose accumulation
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In contrast, the ratio of lactate:acetate of B. longum subsp. suis BSM 11–5 grown with 2′-FL and 3′-FL was approx. 1:1 and 1:3 respectively. B. longum subsp. suis BSM 11–5 synthesized 1,2-PD mainly from 3′-FL, and accumulated 5 mM L-fucose when grown in the presence of 2′-FL. L-fucose might have been accumulated during growth in the presence of 28 mM 2′-FL as glucose and galactose became also available after fucosyllactose degradation.

B. kashiwanohense DSM 21854 grew in the presence of 2′-FL and 3′FL and accumulated approximately 10 mM L-fucose but did not produce any 1,2-PD (Table 4). The ratio of lactate:acetate was approx. 1:2.

B. longum subsp. infantis degrades HMOs internally [3, 34]. The gap in substrate consumption, L-fucose release and/or 1,2-PD formation observed for B. longum subsp. infantis and B. longum subsp. suis might be due to the intracellular which were not released in the supernatant. In contrast, fucose and an additional compound were detected in supernatants of B. bifidum which harvests fucosyllactose extracellularly [28, 34].

We here identified 1,2-PD as a metabolite of bifidobacteria fucosyllactose respective L-fucose degradation. L-fucose derived 1,2-PD can be further metabolized to propionate and propanol by other gut microbes such as Eubacterium hallii [35, 36]. It was estimated that in adults approximately 30 % of propionate might derive from 1,2-PD, but no data exists for infants [36, 37]. Nevertheless, the bifidobacterial formation of lactate and 1,2-PD as precursors of short chain fatty acids butyrate and propionate, respectively, contributes to the trophic interactions of the infant gut microbiota [38].

Genome comparison

Strains of B. longum subsp. infantis, and B. longum subsp. suis used the L-fucose moiety of 2′-FL and 3′-FL to form 1,2-PD, but the enzymatic pathways of bifidobacterial L-fucose metabolism are not known [3]. To further elucidate putative fucosyllactose and L-fucose utilization pathways, we generated draft genomes of B. longum subsp. infantis TPY12-1 and B. longum subsp. suis BSM11-5 for comparing genome data with type strain B. longum subsp. infantis DSM 20088 [3], and to B. kashiwanohense DSM 21854 [39] and PV-20 [40]. Abyss assembly yielded 72 and 105 contigs (>500 bp) from 8.1 to 1.9 Mio reads for B. longum subsp. suis BSM11-5 and B. longum subsp. infantis TPY12-1, respectively (Table 5). For B. longum subsp. suis BSM11-5, the N50 was of 135.581 bp, and the largest contig had 298.614 bp. The N50 of B. longum subsp. infantis TPY12-1 was 94.696 bp with the largest contigs of 163.755 bp. ANI of B. longum subsp. infantis TPY12-1 compared to type strain was 98.4 %. B. longum subsp. suis type strain DSM 20211 and BSM11-5 had an ANI of 98.3 %, whereas ANI of B. longum subsp. suis BSM11-5 and B. longum subsp. infantis DSM 20088, or B. longum subsp. longum DSM 20019 was 96.6 and 96.7 %, respectively, confirming phylogenetic placement of this isolate based on partial 16S rRNA gene analysis (Additional file 1: Figure S1).

Table 5 Genome characteristics of B. longum subsp. infantis, suis and B. kashiwanohense strains used in this study
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Lo Cascio et al. [11] defined six gene regions related to gut adaption and HMO utilization which distinguished subspecies B. longum subsp. longum and infantis (urease, H1-H5). All six regions were present in the genome of B. longum subsp. infantis TPY12-1. B. longum subsp. suis BSM11-5 possessed an urea operon similar to B. longum subsp. infantis. This strain also harboured the LNB region (H5) as reported before for two B. longum subsp. suis isolates, and possessed parts of HMO utilization operon H1. However, B. longum subsp. suis BSM11-5 lacked additional α-fucosidase and sialidase containing gene regions H2-H4. Surprisingly, B. kashiwanohense DSM 21854 and PV20-2 also partly harboured H1 (Fig. 3), and B. kashiwanohense PV20-2 possessed an urea uptake and degradation operon with >90 % homology to B. longum subsp. infantis DSM 20088.

Presence of α-fucosidase encoding genes in B. longum subsp. infantis TPY 12–1, B. longum subsp. suis BSM11-5, and B. kashiwanohense

Alpha-fucosidases, which catalyze the release of α-1-2, α-1-3, and α-1-4 linked fucose, are assigned to GH families 29 and 95 (GH29 and GH95). B. longum subsp. infantis DSM 20088 harbours four α-fucosidases: BLON_0248 (GH29), BLON_0426 (GH29), BLON_2335 (GH95) and BLON_2336 (GH29). B. longum subsp. infantis DSM 20088 also possesses BLON_0346, which is assigned to GH42 but can degrade Fucα1-2Gal [41]. Paralogs BLON_0248 and BLON_0426 are 95 % identical [41].

BlastP was used to identify homologues of the α-fucosidases of B. longum subsp. infantis DSM 20088 in the genomes of B. longum subsp. infantis TPY12-1, B. longum subsp. suis BSM11-5 and the two B. kashiwanohense strains (Table 6).

Table 6 Presence of α-fucosidases
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B. longum subsp. infantis TPY12-1 possessed homologues of BLON_0248/BLON_0426, BLON_0346, BLON_2335, and BLON_2336. B. longum subsp. suis BSM11-5 harboured two α-fucosidases highly similar to BLON_2335, and BLON_2336. B. kashiwanohense DSM 21854 and PV20-2 possessed three and one α-fucosidases, respectively, with high homology (>78 %) to B. longum subsp. infantis DSM 20088. BBKW_1714 of B. kashiwanohense DSM 21584 was flanked by a mobile element and was 100 % identical to BLON_0248/BLON_0426. B. longum subsp. suis BSM11-5, and B. kashiwanohense α-fucosidases were highly similar to each other (98–99 %) (Table 6). A search for conserved protein domains of every CAZyme family [27], identified no further GH29 or GH95 the genomes of B. longum subsp. infantis TPY12-1, B. longum subsp. suis BSM11-5, and B. kashiwanohense DSM 21854 and PV20-2.

Heterologously expressed BLON_0248, BLON_426, and BLON_0346 previously showed no activity on 2′-FL and 3′-FL [41], and BLON_2336 degraded only 3′-FL. BLON_2335 was able to degrade 2′-FL and 3′-FL [41] and homologues of BLON_2335 were present B. longum and B. kashiwanohense strains analysed here.

B. kashiwanohense has been rarely isolated infant stool and not yet from adults [20, 21], while B. longum subsp. infantis is considered an infant bifidobacteria species due to its adaption to degrade HMOs. The presence of highly similar genetic elements (region H1, α-fucosidases, urease operon) flanked by mobile elements indicate possible gene transfer between B. longum and B. kashiwanohense which suggests a co-evolutionary history between the two species that adapted B. kashiwanohense to the infant gut environment.

Putative L-fucose degradation pathways in B. longum subsp. infantis, and B. longum subsp. suis BSM11-5

Strains of B. longum subsp. infantis, and B. longum subsp. suis BSM 11–5 used the L-fucose moiety of 2′-FL and 3′-FL to form 1,2-PD. However, the enzymatic pathway of Bifidobacterium sp. L-fucose metabolism is not known. Microorganisms can metabolize desoxyhexoses such as fucose by two pathways involving phosphorylated and non-phosphorylated intermediates (Fig. 2). For example in E. coli, Roseburia inulivorans, and Lactobacillus rhamnosus fucose isomerase FucI, fucose kinase FucK and fucose aldolase FucA are responsible for L-fucose to L-lactaldehyde transformation under anaerobic conditions [29, 30, 42]. L-rhamnose is metabolized via a similar enzymatic pathway [30]. For Campylobacter jejuni and Xanthomonas campestris, another pathway has been described with non-phosphorylated intermediates [43, 44]. Here L-fucose is internalized via the activity of a permease and metabolized to L-lactate and pyruvate by the activity of L-fucose mutarotase, L-fucose dehydrogenase, L-fuconolactone hydrolase, L-fuconate dehydratase, L-2-keto-3-deoxy-fuconate hydrolase, and a L-2,4-diketo-3-deoxy-fuconate hydrolase (Fig. 2, [43, 44]).

Fig. 2
figure 2

Comparison of L-fucose (a, b) and L-rhamnose (c) dedegradation pathways in E. coli (a), X. campestris (b), and Sphingomonas sp. (c) extracted from Boronat and Aguilar [30], Yew et al. [44], and Watanabe and Makino [45]

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B. longum subsp. infantis DSM 20088 lacks the genes encoding proteins to use fucose via phosphorylation [3]. FucI, FucK and FucA were also not detected in the other genomes analysed here. To investigate whether bifidobacteria might utilize L-fucose similar to X. campestris, we searched for the corresponding proteins of X. campestris in bifidobacteria genomes using BlastP, and also collected enzymes related to fucose metabolism that were annotated by RAST.

B. longum subsp. infantis DSM 20088 and TPY11-1, and B. longum subsp. suis BSM11-5 possessed putative L-fucose dehydrogenases, a L-fuconolactone hydrolase, L-fuconate dehydratases, L-2-keto-3-deoxy-fuconate hydrolases, and a L-2,4-diketo-3-deoxy-fuconate hydrolase with homologies ranging from 24 to 56 % AA similarity to the enzymes of the X. campestris (Table 7). A homologue of the L-fucose mutarotase of X. campestris (XCC4070) was not detected; however, putative L-fucose mutarotases encompassing the conserved RbsD/FucU transport protein family domain were identified by RAST (Table 7).

Table 7 Identification of B. longum and B. kashiwanohense genes related to L-fucose metabolism
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The majority of genes was located on two genomic regions (Fig. 3). In contrast, in X. campestris all responsible genes were located on an operon XCC4065-XCC4070 [44]. Region 1 encompassed a L-fucose mutarotase, a L-2-keto-3-deoxy-fuconate hydrolase, and a L-fuconate dehydratase (Fig. 3). The gene cluster of region 1 also contained genes encoding the α-fucosidases BLON_2335 and BLON_2336 and is part of the B. longum subsp. infantis HMO utilization operon H1 [3, 11]. A possible L-fuconolactone hydrolase, and paralogs of fuconate dehydratase and L-fucose mutarotase were located elsewhere on the genome in close proximity to a putative fucose permease (Fig. 3).

Fig. 3
figure 3

Genomic regions encompassing genes putatively involved in fucose degradation in B. longum and B. kashiwanohense strains. The gene cluster of region 1 also contained genes encoding the α-fucosidases BLON_2335 and BLON_2336 and is part of the B. longum subsp. infantis HMO utilization operon H1 [3, 11] (not drawn according to scale)

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Expression of the fuconate dehydratase of region 1, and of the putative L-2-keto-3-deoxy-fuconate hydrolase was recently reported to be upregulated when B. longum subsp. infantis DSM 20088 was grown in the presence of fucosyllactose providing strong support to the proposition that L-fucose is metabolized via this pathway [34].

B. kashiwanohense DSM 21854 and PV20-2 only possessed a gene segment similar to region 1 with a truncated L-fucose-mutarotase, and lacked region 2 which encompassed the fucose permease. This might be the reason why strains of B. kashiwanohense were not able to utilize L-fucose.

For X. campestris, lactate and pyruvate, but not 1,2-PD, were determined as final metabolites of fucose fermentation. Watanabe and Makino [45] however described a modified non-phosphorylated L-rhamnose pathway which would yield L-lactaldehyde from L-2-keto-3-deoxyrhamnonate. The transformation of L-2-keto-3-deoxyrhamnonate to L-lactaldehyde and pyruvate was catalyzed by a L-2-keto-3-deoxyrhamnonate aldolase. A possible candidate gene could be the RAST annotated 4-hydroxy-tetrahydrodipicolinate synthase possessing a structurally conserved phosphate binding motif. 4-hydroxy-tetrahydrodipicolinate synthase encoding genes were located in region 1 of all strains investigated, B. longum subsp. infantis strains possessed a paralog in region 2. 4-hydroxy-tetrahydrodipicolinate synthases reversibly catalyze the formation of 4-hydroxy-2,3,4,5-tetrahydrodipicolinate from pyruvate and L-aspartate-4-semialdehyde [46].

Taken together these results suggest that B. longum subsp. infantis TPY12-1, DSM 20088 and B. longum subsp. suis BSM11-5 metabolize fucose via a pathway with non-phosphorylated intermediates as previously described for Campylobacter sp. and X. campestris.

Conclusion

Here we observed that the ability to degrade fucosyllactose is a trait of various bifidobacteria species. We identified strains of B. longum subsp. infantis and an isolate of B. longum subsp. suis were able to degrade fucosyllactose and L-fucose to form 1,2-PD. We propose that bifidobacteria degrade L-fucose via a pathway with non-phosphorylated intermediates as described for desoxyhexoses before.