The potential impact of the composition of maternal breast milk is poorly known in children who develop celiac disease (CD). The aim of our study was to compare the microbiota composition and the concentrations of immune markers in breast milk from mothers whose offspring carried the genetic predisposition to CD, and whether they did or did not develop CD during follow-up for the first 3 years of life. Maternal breast milk samples [CD children (n = 6) and healthy children (n = 18)] were collected 3 months after delivery. Enzyme-linked immunosorbent assays were used to measure TGF-β1, TGF-β2, sIgA, MFG-E8 and sCD14. For microbiota analysis, next generation (Illumina) sequencing, real-time PCR and denaturing gradient gel electrophoresis were used. Phylotype abundance and the Shannon ‘H’ diversity index were significantly higher in breast milk samples in the CD group. There was higher prevalence of the phyla Bacteroidetes and Fusobacteria, the classes Clostridia and Fusobacteriia, and the genera Leptotrichia, Anaerococcus, Sphingomonas, Actynomyces and Akkermansia in the CD group. The immunological markers were differently associated with some Gram-negative bacterial genera and species (Chryseobacterium, Sphingobium) as well as Gram-positive species (Lactobacillus reuteri, Bifidobacterium animalis). In conclusion, the microbiota in breast milk from mothers of genetically predisposed offspring who presented CD showed a higher bacterial phylotype abundance and diversity, as well as a different bacterial composition, as compared with the mothers of unaffected offspring. These immune markers showed some associations with bacterial composition and may influence the risk for development of CD beyond early childhood.
Celiac disease (CD) is a systemic immune-mediated disorder triggered by gluten and related prolamins in genetically susceptible individuals1,2. This disease is characterized by a variable combination of gluten-dependent clinical manifestations, CD-specific antibodies, HLA-DQ2/8 haplotypes and enteropathy1,2,3.
Previously, it has been demonstrated that breast feeding, its duration and the introduction of gluten into an infant’s diet may be determinants in the pathogenesis of CD4,5,6. Breast milk components promote oral tolerance to dietary antigens by modulating immune development and infant gut microbiota composition4,7,8,9,10. The latter is also thought to be involved in the pathogenesis of CD11. In contrast, some studies have reported no effect of breast-feeding on the development CD in children12,13.
Earlier work has shown that infants with an increased genetic risk for CD were colonized with a lower proportion of Actinobacteria and a higher proportion of Firmicutes and Proteobacteria than infants with a low genetic risk for the development of CD6. De Palma et al. demonstrated that the type of milk feeding in relation to HLA-genotype played a role in establishing infant gut microbiota14. The HLA-DQ genotype may specifically influence the colonization process of Bacteroides species3,15. In addition, it has been observed that breast milk from CD mothers have lower levels of TGF-β1, sIgA and a reduced abundance of Bifidobacterium sp. and B. fragilis compared with healthy women5.
As the frequency of CD has increased we hypothesized that the immunological composition and microbiota of breast milk might differ between the mothers whose offspring carry a genetic susceptibility to CD, and developed CD relative to those who remain unaffected. The current study set out to characterize breast milk microbiota composition and its possible relationship with immunological markers (TGF-β1, TGF-β2, sIgA, MFG-E8, and sCD14) in mothers whose offspring presented or did not present CD during the first 3 years of life.
Immune markers in breast milk from mothers of CD and control offspring
There were no significant differences between TGF-β1, TGF-β2, sIgA, MFG-E8, and sCD14 levels in the breast milk from mothers in the CD group and the breast milk from mothers in the control group (Table 1).
The microbiome of the maternal breast milk
We applied Illumina sequencing of the 16S rRNA V4 region to reveal the full microbiome of these investigated samples. A total of 6,475,320 high quality reads were obtained, or 269,805 ± 239,377 reads per milk sample. The OTUs were classified into known taxa (7 phyla, 17 classes, 44 genera, and more than 90 species) and unclassified groups.
Phylotype abundance and the Shannon ‘H’ diversity index were significantly higher in milk samples from the mothers in the CD group than in the maternal samples in the control group (p = 0.016; p = 0.008, respectively) (Table 2).
A principal coordinate analysis (PCoA) plot based on different taxonomic levels (phylum, class and genus) was generated to assess the relationships between the community structures of these samples. Phylum and class abundance data indicated significant inter-individual variability (Fig. S1A,B), while the PCoA plot of relative genus abundance demonstrated weighted clustering in the CD group (Fig. S1C).
In both study groups, the phylum Firmicutes displayed the highest relative abundance (medians of 48.9% and 60.2% for the CD group and the control group, respectively) (Fig. 1A, Supplementary Table S2). In addition, Proteobacteria and Actinobacteria were also quite abundant (median 29.0% and 9.8% for the CD group vs. 23.6% and 8.1% for the control group). A borderline statistical significance was detected for the relative abundance of Bacteroidetes and Fusobacterium phyla (p = 0.056 and p = 0.048; Fig. 1A, Supplementary Table S2).
At the class level, Bacilli displayed the highest relative abundance both in the CD group (median 30.9%) and in the control group (median 56.1%), followed by Gammaproteobacteria (20.8% and 14.5%), Actinobacteria (8.9% and 8%) and Alphaproteobacteria (4% and 3.7%) (Fig. 1B, Supplementary Table S3). Statistically significant differences were only found for the relative abundance of class Clostridium (p = 0.02) and Fusobacteriia (p = 0.05) (Fig. 1B, Supplementary Table S3).
The most abundant genera of bacteria both in the CD and the control groups was Lactobacillus (median 10.3% and 34%, respectively), followed by Streptococcus (10.9% and 7.5%), Staphylococcus (4.9% and 4.9%), Buttiauxella (4.8% and 4.4%), and Rothia (2.8% and 2.6%) (Fig. 1C, Supplementary Table S4). The prevalence of the genera Anaerococcus (p = 0.006), Shpingomonas (p = 0.01), Actinomyces (p = 0.006), Leptotrichia (p = 0.005), and Akkermansia (p = 0.01) were significantly higher in the CD group (Supplementary Table S4). Moreover, the relative abundances of Sphingomonas (p = 0.04) and Akkermansia (p = 0.05) were significantly higher in the CD group (Fig. 1C, Supplementary Table S4).
The most abundant species in the CD group in comparison to the control group were Actinomyces odontolyticus (p = 0.03), Anaerococcus hydrogenalis (p = 0.04), and A. octavius (p = 0.01) (Supplementary Table S5). The prevalence of A. muciniphilia was also increased in the CD group (p = 0.002). It is noteworthy that the milk samples in the control group had lower relative abundance of Faecalibacterium prausnitzii (p = 0.049) (Supplementary Table S5), while unclassified Leptotrichia and Akkermansia were detected only in the CD group. Lactobacillus salivarius (83 and 78%) and Bifidobacterium animalis (both 50%) were the most prevalent species among lactic acid bacteria in both study groups, although their mean counts were less than 1% (Supplementary Table S5).
A total of nineteen Lactobacillus species (L. plantarum, L. curvatus, L. iners, L. mucosae, L. casei, L. fermentum, L. salivarius, L. reuteri, L. rhamnosus, L. crispatus, L. hominis, L. sakei, L. zeae, L. jensenii, L. senmaizukei, L. paracasei, L. oligofermentas, L. gasseri, and L. acidophilus) (Fig. 2B) and four species of Bifidobacterium (B. bifidum, B. animalis, B. adolescentis, and B. longum) were detected by whole genome sequencing (Fig. 3B). Additionally, L. johhansonii (Fig. 2C) and B. pseudocatenulatum, B. infantis and B. breve were using denaturing gradient gel electrophoresis (DGGE) (Fig. 3C, Table S9).
A principal coordinate analysis (PCoA) plot of relative abundance data of Lactobacillus and Bifidobacterium species indicated quite different results, with weighted clustering of Lactobacillus in the CD group (Fig. 2A) and Bifidobacterium in the control group (Fig. 3A). No differences were found between the groups in quantitative counts for Lactobacillus or Bifidobacterium (Figs. 2D, 3D; Table S9). Milk samples in the CD group showed borderline significance for the lower relative abundance of L. fermentum when compared to the control group (p = 0.058, Supplementary Table S5).
Associations between breast milk microbiota and immune markers
A. muciniphila showed a positive association with TGF-β1 (p = 0.04) and TGF-β2 (p = 0.007) (Table 3). Higher counts of the genera Chryseobacterium (p = 0.04) and Sphingobium (p = 0.01) were seen in milk samples with lower TGF-β2 levels. MFG-E8 correlated positively with the bacterial classes Flavobacteriia (p = 0.03) and Bacilli (p = 0.05) and inversely with the species B. animalis (p = 0.03). In addition, the species L. reuteri was positively (p = 0.04) and B. animalis was inversely (p = 0.02) correlated with sCD14 levels (Table 3). All the correlation coefficients are presented in Supplementary Tables S6–S8.
In this case–control study, we wanted to identify whether there were differences in the immunological composition and microbiota of breast milk between mothers whose genetically predisposed offspring developed CD during the first 3 years of life or remained unaffected. To the best of our knowledge, the present study was the first where maternal breast milk samples were analyzed for both immunological markers and for microbiota composition.
We demonstrated that, compared to milk samples from mothers of unaffected offspring, the human milk microbiota in the mothers in the CD group had a higher phylotype abundance and diversity, with a higher abundance of the phyla Bacteroidetes and Fusobacteria, the classes Clostridia and Fusobacteria, as well as the genera Leptotrichia, Anaerococcus, Sphingomonas, Actinomyces, and Akkermansia. Increased relative abundance for species such as A. odontolyticus, A. hydrogenalis, A. octavius, and F. prausnitzii, and decreased abundance for L. fermentum were also observed in the CD group. We observed an association between certain immune markers (TGF-β1, TGF-β2, MFG-E8, and sCD14) and some Gram-negative (Chryseobacterium, Sphingobium, and A. muciniphila) and Gram-positive bacteria (L. reuteri and B. animalis).
Breast milk does not contain abundant culturable microbiota. However, some next generation sequencing studies have revealed Streptococcus, Staphylococcus, Gemella, Veillonella, Rothia, Lactobacillus, Propionibacterium, Corynebacterium, and Pseudomonas in milk samples from healthy woman8,16. In the current study, the most abundant genus in all milk samples was Lactobacillus, followed by Streptococcus, Staphylococcus, Moraxella, Acinetobacter, Enterobacter, and Corynebacterium. Some studies have indicated that the primary bacterial taxa in milk may vary across population, suggesting that geographic, genetic, and dietary factors could be influencing microbial diversity in breast milk. In Finland, Leuconostoc, Weisella, and Lactococcus were the most predominant genera in milk samples17, while in Mexican–American mothers the predominance of Streptococcus, Staphylococcus, Xanthomonadaceae, and Sediminibacterium was observed18.
Breast milk is a major source of bacterial diversity for the neonatal gut, including gut-associated obligate anaerobes, and therefore significantly influences gut colonization and the maturation of the immune system6,19. Different genetic and epigenetic factors may influence host microbiome interactions and shift the immune balance in subjects with CD20. In case of the intestinal microbiome, higher microbial richness and diversity values are considered to be more health-supporting21. However, this may not be true in case of some other biotopes (for example, the reproductive tract)22. Quite unexpectedly, we found that samples taken from mothers whose children developed CD had a higher phylotype abundance and Shannon diversity index than healthy samples. It is interesting that a recent study by Huang et al. also demonstrated a significantly higher alfa diversity in fecal samples from CD progressors compared to healthy controls at an age of one year23. This indicates that the increase of a few dominant bacteria would be expected in cases of infection and suggests that milk microbiota is not activating an immune response in the host. Unfortunately, in the current study inflammatory markers were not measured. In contrast, it has been shown that in case of allergy, feeding breast milk with lower bacterial diversity during the first month of life was related to an increased risk of allergy in childhood24. We cannot rule out the influence of anticommensal antibodies on microbiota composition, which may differ even between healthy individuals and may have an impact on microbiome composition25,26.
Olivares et al.6 showed that the breast milk of mothers with CD demonstrated significant reductions in the counts of Bifidobacterium spp. and Bacteroides fragilis in comparison to healthy mothers. In the present study we observed a significantly increased relative abundance of class Clostridia in the milk samples from the CD group. We also found differences in the relative abundance of the genus Anaerococcus (incl. A. hydrogenalis, A. octavius) and F. prausnitzii. The presence of A. octavius was reported previously in breast milk samples from healthy Spanish women. In the same study, an association was shown between the amounts of proteins in milk and Anaerococcus27. F. prausnitzii is the major butyrate producer in the gastrointestinal tract. It has anti-inflammatory properties important for gastrointestinal microbiota homeostasis and is associated with a range of metabolic processes in the human mucosa28. A Spanish study showed that the adherence to a gluten-free diet resulted in slight microbiota shifts in adults, including a decreased abundance of F. prausnitzii29. As F. prausnitzii produces the anti-inflammatory short-chain fatty acid butyrate30,31, this species may have a key role in maintaining the integrity and function of the mucus barrier, representing an important aberrant mechanism in the development of CD.
We also found that the prevalence of the genus Akkermansia was higher in the CD group samples. Interestingly, unclassified Akkermansia species were found only in that group. A study by Huang et al. observed the Akkermansia genus only in the control group and not in CD progressors23. However, they analyzed fecal samples and not breast milk samples. In contrast, some experiments in mice have demonstrated that a gluten-free diet can induce changes in the intestinal microbiota by increasing the number of Akkermansia32,33. It can be speculated that this species may translocate from its normal habitat (intestine) to an unusual habitat (breast) in cases of CD.
An increased relative abundance for the genus Actinomyces and for a species of this genus, A. odontolyticus, were observed in the CD group. Mucosal membranes of the oropharynx, gastrointestinal tract and female genital tract represent the normal habitat of Actinomyces34. They have also been observed to associate with breast infection35,36. Although being low in virulence, they may rely on the presence of co-pathogens, such as anaerobic bacteria, to enhance pathogenicity37. Previously, A. graevenitzii was identified in biopsy specimens from the proximal small intestine of children with CD, and an unidentified Actinomyces sp. was shown to be attached to the epithelial lining38.
For detection of Lactobacillus and Bifidobacterium, we applied two methods, next generation sequencing and PCR-DGGE, since Smidt et al., demonstrated the partial concordance of different molecular methods for the detection of lactic acid bacteria39. In our study, PCR-DGGE appeared to be more sensitive for the identification of Bifidobacterium sp., while more than 80% of Bifidobacterium sp. detected by Illumina sequencing were unclassified.
In line with our findings, other studies have also shown the presence of Lactobacillus sp. and Bifidobacterium sp. in breast milk6,40. A low relative abundance of L. fermentum was observed in breast milk in the CD group. The expected pathway by which lactobacilli may get into milk is entero-mammary transport through dendritic cells41. Production of short-chain fatty acids by lactobacilli strains may play a role in the pathogenesis of CD. L. fermentum produces both lactic and succinic acids, which in turn modulate the key functions of many main players in the innate immune response42. They mediate macrophage dependent anti-inflammatory effects43 and possess antioxidative capacities44. Additionally, it was demonstrated that L. fermentum isolated from breast milk may have an immunostimulatory effect by inducing proinflammatory cytokines in rodent bone marrow derived macrophages40. In vivo assays in mice revealed that the ingestion of L. fermentum enhanced the production of Th1 cytokines by spleen cells and increased the IgA concentration in feces45. Likewise, Lactobacillus strains have high amylase trypsin inhibitor-metabolizing capacity to ameliorate many adverse effects induced by wheat immunogenic proteins46. Accordingly, one may assume that L. fermentum might protect against the development of CD.
There were no significant differences in the levels of the immunological markers analyzed between the CD and the control group, but some significant associations with milk microbiota were observed. The presence of Chryseobacterium and Sphingobacterium (both being Gram negative environmental bacilli) correlated inversely with breast milk TGF- β2 levels, while A. muciniphila (anaerobic mucin-degrading bacterium) showed a positive correlation with both TGF- β1 and TGF- β2. TGF-β belongs to a growth factor family that is responsible for the development and maturation of the mucosal immune system47. In our study, we also observed that higher counts of L. reuteri and lower counts of B. animalis in breast milk were associated with higher sCD14 concentrations. The latter bacterial species were also associated with higher concentrations of MFG-E8. Vidal et al. demonstrated that milk sCD14 acts as a “sentinel” molecule and immune modulator in homeostasis and in the defense of the neonatal intestine48. Supplementation of maternal breast milk with the probiotic L. reuteri increased the TGF-β2 concentration level in breast milk and decreased sensitization in breast-fed infants49. In addition, the Bacilli and Flavobacteriia classes correlated positively with the concentration of MFG-E8, which has been shown to have antiviral effects and to protect against rotavirus both in a cultured cell model as well as in infants50,51,52. Furthermore, it prevented the tissue damage caused by prolonged inflammation by clearing apoptotic cells and thereby facilitating immune resolution53.
Our study had many strengths. All study subjects in both countries were monitored and diagnosed with identical protocols. All CD diagnoses were confirmed by a biopsy of the small intestine. Although patients came from two different countries, all the analyses were carried out in one laboratory. Two different methods in parallel were applied for investigating Lactobacillus sp. and Bifidobacteria sp. in breast milk microbiota.
Our study also had some limitations. The number of cases was quite small, limiting the statistical power of this study. The mothers in the CD group were younger than the mothers in the control group. When interpreting the results this may be important because it is known that maternal age may influence milk composition and microbiota54. The children were observed for only 3 years and we lack information about possible new CD cases beyond that age. As we analyzed only a small number of breast milk samples taken at the age of 3 months, larger longitudinal studies are needed for a better understanding of possible differences between these two groups.
In conclusion, this study provides new insights into the complex breast milk microbiota composition in mothers whose offspring carry a genetic susceptibility to CD but have different disease outcomes. The breast-milk microbiota of mothers of children who developed CD differed in terms of higher bacterial phylotype abundance and diversity, as well as in relation to bacterial composition when compared to the mothers of unaffected children. The immunological markers were differently associated with bacterial composition and could also influence the risk of development CD in later life. Future studies are needed to reveal whether differences in breast milk composition ultimately influence the development of CD.
Materials and methods
Participants in the current analysis were recruited from the DIABIMMUNE Study, which was originally carried out in Estonia, Finland, and Russian Karelia from September 2008 to October 2013. The exact recruitment criteria and number of participants have been described previously55. For the current analysis, samples and data from Estonia and Finland were used.
This study was conducted in accordance with the Declaration of Helsinki and local ethical committees respectively approved this study (Ethics Review Committee on Human Research of the University of Tartu in Estonia and Ethical Committee for Psychiatric Diseases and Diseases in Children and Adolescents, Helsinki and Uusimaa Hospital District in Finland). Written informed consent was obtained from all parents for the participation of their child in this study.
Children who were followed from birth to the age of 3 years and those who later developed CD (n = 6) were included in our analysis. The diagnosis of CD was confirmed according to the European Society for Pediatric Gastroenterology, Hepatology and Nutrition2, including the abnormal morphology of small intestinal biopsy in accordance with the Marsh classification modified by Oberhuber56. From the same DIABIMMUNE cohort we selected 18 controls based on the CD-specific HLA DR/DQ genotype, country of birth, time of birth and sex of each patient.
Five children in the CD group were exclusively breastfed, while one child was mostly bottle-fed during the time of breast milk collection. The corresponding numbers in the control group were 15 and 3. One CD patient and his controls were from Estonia, while all the others were from Finland. The proportions of boys and girls were equal in the CD and control groups. All offspring were HLA DQ 2.5 positive (two CD patients were HLA DQ 2.5 homozygotes). Each infant was full-term (gestational age 37–42 weeks) and born as a singleton. None of the mothers or fathers of the offspring had CD. The mean age was 2.4 years (range 1.5–3.0 years) at seroconversion to positivity for IgA- class tissue transglutaminase antibodies in the CD group. General characteristics of the CD and the control group children are summarized in Table 4.
Collection and processing of samples
Breast milk samples were collected from mothers in both groups 3 months after delivery. The milk samples were obtained by manual expression into sterile bags. Samples were immediately put into a −20 ºC freezer and further stored at −80 ºC until processing. Breast milk samples were thawed and centrifuged for 15 min at 1000g at 4 ºC and after that the aqueous fraction was collected. The latter was used for further analyses. Centrifugation was repeated twice.
All breast milk analyses were carried out in the Department of Immunology and Department of Microbiology at Institute of Biomedicine and Translational Medicine, University of Tartu, Estonia.
Immune markers in breast milk
We used enzyme-linked immunosorbent assay (ELISA) to measure TGF-β1, TGF-β2, sIgA, MFG-E8 (lactadherin), and sCD14. For measuring TGF-β1, TGF-β2, MFG-E8, and sCD14 we used commercial kits from R&D Systems (Minneapolis, MN, USA) according to the manufacturer’s instructions. For sIgA, we used a commercial kit from Bethyl Laboratories (Montgomery, AL, USA) according to the manufacturer’s instructions.
Bacterial DNA from human milk was extracted using the PureLink™ Microbiome DNA Purification Kit [Invitrogen, using an ELMI SkyLine instrument (ELMI Ltd., Riga, Latvia)]. Human milk samples were centrifuged at 4000g for 30 min, after which the supernatant was removed. Cell pellets were washed with phosphate-buffered saline, centrifuged at 13,000g × 3 min at room temperature and the pellets were resuspended in a kit-specific lysis buffer. The protocol was then continued as described by the manufacturer (Invitrogen, USA).
16S library preparation was performed using an in-house sequencing protocol with the V4 (F515/R806) primer pair57. Sequencing was performed using an iSeq 100 (SN FS10000643, Illumina) and an iSeq 100 i1 Reagent kit (Illumina) in the 2 × 150 bp mode and using the dual index setup with two-read sequencing protocol. 16S rDNA sequence data were analyzed using BION-meta (https://www.box.com/bion), according to the author’s instructions and in-house scripts. First, sequences were cleaned at both ends using a 99.5% minimum quality threshold for at least 18 of 20 bases for the 5′-end and 28 of 30 bases for 3′-end. The reads were then joined, followed by the removal of read pairs shorter than 150 bp. Then, sequences were cleaned from chimeras and clustered using a 95% oligonucleotide similarity (k-mer length of 8 bp and a step size of 2 bp). Lastly, consensus reads were aligned to the SILVA reference 16S rDNA database (v123) using a word length of eight and similarity cut-off of 90%. The bacterial designation was analyzed at different taxonomic levels down to the species when applicable.
Primers and probes
In order to establish a quantitative assay, plasmid standards were generated using the method described in Bartosch et al.62. The amplified 16S rRNA gene region (amplified with specific primers) from B. longum DSM20219 and L. acidophilus ATCC4356 was cloned into chemically component E. coli JM109 cells using the pGEM-T vector system (Promega, Madison, WI, USA). Plasmids were purified with NucleoSpin Plasmid QuickPure kit according to the manufacturer’s instructions (Macherey–Nagel, Düren, Germany). Multiple dilutions of purified plasmids were quantified by spectrophotometry (NanoDrop ND-1000, Thermo Fisher Scientific, Waltham, MA, USA). Quantification of target DNA was achieved by using serial tenfold dilution from 105 to 101 plasmid copies of previously quantified plasmid standards.
Multiplex TaqMan assay PCR reactions were performed in a total volume of 25 µl using TaqMan® Universal PCR Master Mix (Applied Biosystems, Foster City, CA, USA). Each reaction included 5 µl of template DNA, 12.5 µl of TaqMan® Universal PCR Master Mix (Applied Biosystems), 400 nM of (F_allbif_IS and R_alllact_IS primers), 600 nM of (F_alllact_IS and R_alllact_IS primers), 900 nM of (F_Eub and R_Eub primers) and 100 nM both (P_alllact_IS, P_alllbif_IS) and 100 nM (P_Eub) of probes. The real-time PCR conditions consisted of an initial denaturation step at 50 °C for 2 min and then 95 °C for 10 min, continued with an amplification step followed by 40 cycles including denaturation at 95 °C for 15 s, and an annealing-elongation step at 60 °C for 1 min. Amplification and detection of DNA samples by real-time PCR was performed with a 7500 Fast Real-Time PCR System (Applied Biosystems Europe BV, Zug, Switzerland) using optical-grade 96-well plates. Data from triplicate samples were analyzed using Sequence Detection Software version 1.6.3 (Applied Biosystems).
Denaturing gradient gel electrophoresis
PCR was performed in a reaction volume of 50 µl containing 25 µl 2× DreamTaq Hot Start PCR Master Mix (Thermo Fisher Scientific), 200 ng of DNA solution and primers at a concentration of 10 µM. The DGGE cycling parameters were 5 min at 94 °C, followed by 35 cycles of 30 s at 57 °C (for primer Im-3 and Im-26), 30 s at 94 °C, 30 s at 62 °C (for primers Bif 164 and Bif 662 + GC), and 45 s at 56 °C (for primer Lac-1 and Lac-2 + GC), and a final extension at 72 °C for 10 min.
The DGGE analysis of PCR amplicons was performed using a Dcode™ System apparatus (Bio-Rad, Hercules, CA, USA). Polyacrylamide gels 8% [wt/vol] involving acrylamide-bisacrylamide [37.5:1] in 0.5× Tris–acetic acid-EDTA buffers with a denaturing gradient was prepared with a gradient mixer and Econopump (Bio-Rad). Gradients from 30 to 60% were employed for the separation of the products amplified with specific primers for Lactobacillus spp. and from 45 to 60% for the products amplified with primers specific for Bifidobacterium spp.
Sequencing of DGGE amplicons
PCR amplicons (Bif164-r and Bif662-f) and (Lac-1 and Lac-2) were purified and concentrated with a QIAquick PCR purification kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. Purified amplicons were then cloned into the E. coli JM 109 strain using the pGEM-T vector system (Promega, Madison, WI, USA). Colonies of ampicillin-resistant transformants were randomly picked from each sample and were subjected to PCR with the pGEM-T specific primers T7 and SP6 (Supplementary Table S1) from lyzed cells to check the size of inserts. Plasmid DNA from selected transformants was isolated using a QIAprep Spin Miniprep kit (Qiagen).
Sequencing reactions were performed using the BigDye Terminator CA v3.1 Cycle Sequencing kit (Applied Biosystem) according to the manufacturer’s instructions. The sequences obtained were analyzed using an automatic LI-COR DNA Sequencer 4000L (Licor, Lincoln, NE, USA) and were corrected manually. All of the sequences were thereafter identified using BLASTN and the NCBI nucleotide database.
Statistical analysis of clinical and molecular data was performed with the R software for Windows 2018 (R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria). The Shapiro–Wilk normality test was used to determine whether the data had a normal distribution. Comparisons between the CD group and the control group were performed using a t-test for normally distributed data and with the Mann–Whitney-Wilcoxon test for data with skewed distribution. To compare the number of positive cases between groups, Fisher’s exact test was used. Correlations were analyzed with a non-parametric Spearman’s rank-order correlation test. Alpha diversity based on the Shannon index of the OTU level. Analysis of beta-diversity of CD and control groups including PCoA of weighted UniFrac distances were performed and visualized using PAST 4.0 version. Statistical significance was accepted as p < 0.05, adjusted for ties.
The datasets generated during and/or analysed during the current study are available from the corresponding author on reasonable request.
Caio, G. et al. Celiac disease: A comprehensive current review. BMC Med. 17, 142 (2019).
Husby, S. et al. Diagnosis EWGoCD, Committee EG, European Society for Pediatric Gastroenterology H, Nutrition. European Society for Pediatric Gastroenterology, Hepatology, and Nutrition guidelines for the diagnosis of coeliac disease. J. Pediatr. Gastroenterol. Nutr. 54, 136–160 (2012).
Benitez-Paez, A. et al. Breast-milk microbiota linked to celiac disease development in children: A pilot study from the preventCD cohort. Front. Microbiol. 11, 1335 (2020).
Ivarsson, A. et al. Prevalence of childhood celiac disease and changes in infant feeding. Pediatrics 131, e687–e694 (2013).
Olivares, M. et al. Human milk composition differs in healthy mothers and mothers with celiac disease. Eur. J. Nutr. 54, 119–128 (2015).
Olivares, M. et al. The HLA-DQ2 genotype selects for early intestinal microbiota composition in infants at high risk of developing coeliac disease. Gut 64, 406–417 (2015).
Bezirtzoglou, E., Tsiotsias, A. & Welling, G. W. Microbiota profile in feces of breast- and formula-fed newborns by using fluorescence in situ hybridization (FISH). Anaerobe 17, 478–482 (2011).
Drell, T. et al. The influence of different maternal microbial communities on the development of infant gut and oral microbiota. Sci. Rep. 7, 9940 (2017).
Pozo-Rubio, T. et al. Immune development and intestinal microbiota in celiac disease. Clin. Dev. Immunol. 2012, 654143 (2012).
Sepp, E., Naaber, P., Voor, T., Mikelsaar, M. & Björksten, B. Development of intestinal microflora during the first month of life in Estonian and Swedish infants. Microb. Ecol. Health Dis. 12, 22–26 (2000).
Verdu, E. F., Galipeau, H. J. & Jabri, B. Novel players in coeliac disease pathogenesis: Role of the gut microbiota. Nat. Rev. Gastroenterol. Hepatol. 12, 497–506 (2015).
Szajewska, H. et al. Systematic review with meta-analysis: Early infant feeding and coeliac disease–Update 2015. Aliment. Pharmacol. Ther. 41, 1038–1054 (2015).
Vriezinga, S. L. et al. Randomized feeding intervention in infants at high risk for celiac disease. N. Engl. J. Med. 371, 1304–1315 (2014).
Palma, G. D. et al. Influence of milk-feeding type and genetic risk of developing coeliac disease on intestinal microbiota of infants: the PROFICEL study. PLoS ONE 7, e30791 (2012).
Sanchez, E. et al. Influence of environmental and genetic factors linked to celiac disease risk on infant gut colonization by Bacteroides species. Appl. Environ. Microbiol. 77, 5316–5323 (2011).
Williams, J. E. et al. Human milk microbial community structure is relatively stable and related to variations in macronutrient and micronutrient intakes in healthy lactating women. J. Nutr. 147, 1739–1748 (2017).
Cabrera-Rubio, R. et al. The human milk microbiome changes over lactation and is shaped by maternal weight and mode of delivery. Am. J. Clin. Nutr. 96, 544–551 (2012).
Dave, V. et al. Bacterial microbiome of breast milk and child saliva from low-income Mexican–American women and children. Pediatr. Res. 79, 846–854 (2016).
Jost, T., Lacroix, C., Braegger, C. & Chassard, C. Assessment of bacterial diversity in breast milk using culture-dependent and culture-independent approaches. Br. J. Nutr. 110, 1253–1262 (2013).
Cenit, M. C., Olivares, M., Codoner-Franch, P. & Sanz, Y. Intestinal microbiota and celiac disease: Cause, consequence or co-evolution?. Nutrients 7, 6900–6923 (2015).
Mosca, A., Leclerc, M. & Hugot, J. P. Gut microbiota diversity and human diseases: Should we reintroduce key predators in our ecosystem?. Front. Microbiol. 7, 455 (2016).
Al-Nasiry, S. et al. The interplay between reproductive tract microbiota and immunological system in human reproduction. Front. Immunol. 11, 378 (2020).
Huang, Q. et al. Children developing celiac disease have a distinct and proinflammatory gut microbiota in the first 5 years of life. BioRxiv https://doi.org/10.1101/2020.02.29.971242:2020.02.29.971242 (2020).
Dzidic, M. et al. Allergy development is associated with consumption of breastmilk with a reduced microbial richness in the first month of life. Pediatr. Allergy Immunol. 31, 250–257 (2020).
Prangli, A. L. et al. Antigenic proteins of Lactobacillus acidophilus that are recognised by serum IgG antibodies in children with type 1 diabetes and coeliac disease. Pediatr. Allergy Immunol. 21, e772–e779 (2010).
Talja, I. et al. Antibodies to Lactobacilli and Bifidobacteria in young children with different propensity to develop islet autoimmunity. J. Immunol. Res. 2014, 325938 (2014).
Boix-Amoros, A., Collado, M. C. & Mira, A. Relationship between milk microbiota, bacterial load, macronutrients, and human cells during lactation. Front. Microbiol. 7, 492 (2016).
Sokol, H. et al. Faecalibacterium prausnitzii is an anti-inflammatory commensal bacterium identified by gut microbiota analysis of Crohn disease patients. Proc. Natl. Acad. Sci. U S A 105, 16731–16736 (2008).
De Palma, G., Nadal, I., Collado, M. C. & Sanz, Y. Effects of a gluten-free diet on gut microbiota and immune function in healthy adult human subjects. Br. J. Nutr. 102, 1154–1160 (2009).
Lopez-Siles, M. et al. Cultured representatives of two major phylogroups of human colonic Faecalibacterium prausnitzii can utilize pectin, uronic acids, and host-derived substrates for growth. Appl. Environ. Microbiol. 78, 420–428 (2012).
Ouyang, J. et al. The Bacterium Akkermansia muciniphila: A sentinel for gut permeability and its relevance to HIV-related inflammation. Front. Immunol. 11, 645 (2020).
Hansen, C. H. et al. A maternal gluten-free diet reduces inflammation and diabetes incidence in the offspring of NOD mice. Diabetes 63, 2821–2832 (2014).
Marietta, E. V. et al. Low incidence of spontaneous type 1 diabetes in non-obese diabetic mice raised on gluten-free diets is associated with changes in the intestinal microbiome. PLoS ONE 8, e78687 (2013).
Smego, R. A. Jr. & Foglia, G. Actinomycosis. Clin. Infect. Dis. 26, 1255–1261 (1998) ((quiz 1262-3)).
Edmiston, C. E. Jr., Walker, A. P., Krepel, C. J. & Gohr, C. The nonpuerperal breast infection: Aerobic and anaerobic microbial recovery from acute and chronic disease. J. Infect. Dis. 162, 695–699 (1990).
Kononen, E. & Wade, W. G. Actinomyces and related organisms in human infections. Clin. Microbiol. Rev. 28, 419–442 (2015).
Bing, A. U. et al. Actinomyces species isolated from breast infections. J. Clin. Microbiol. 53, 3247–3255 (2015).
Ou, G. et al. Proximal small intestinal microbiota and identification of rod-shaped bacteria associated with childhood celiac disease. Am. J. Gastroenterol. 104, 3058–3067 (2009).
Smidt, I. et al. Comparison of detection methods for vaginal lactobacilli. Benef. Microbes 6, 747–751 (2015).
Martin, R. et al. Human milk is a source of lactic acid bacteria for the infant gut. J. Pediatr. 143, 754–758 (2003).
Lyons, K. E., Ryan, C. A., Dempsey, E. M., Ross, R. P. & Stanton, C. Breast milk, a source of beneficial microbes and associated benefits for infant health. Nutrients 12, 1039 (2020).
Blad, C. C., Tang, C. & Offermanns, S. G protein-coupled receptors for energy metabolites as new therapeutic targets. Nat. Rev. Drug Discov. 11, 603–619 (2012).
Garrote, G. L., Abraham, A. G. & Rumbo, M. Is lactate an undervalued functional component of fermented food products?. Front. Microbiol. 6, 629 (2015).
Mikelsaar, M. & Zilmer, M. Lactobacillus fermentum ME-3—An antimicrobial and antioxidative probiotic. Microb. Ecol. Health Dis. 21, 1–27 (2009).
Diaz-Ropero, M. P. et al. Two Lactobacillus strains, isolated from breast milk, differently modulate the immune response. J. Appl. Microbiol. 102, 337–343 (2007).
Caminero, A. et al. Lactobacilli degrade wheat amylase trypsin inhibitors to reduce intestinal dysfunction induced by immunogenic wheat proteins. Gastroenterology 156, 2266–2280 (2019).
Oddy, W. H. & McMahon, R. J. Milk-derived or recombinant transforming growth factor-beta has effects on immunological outcomes: A review of evidence from animal experimental studies. Clin. Exp. Allergy 41, 783–793 (2011).
Vidal, K., Labeta, M. O., Schiffrin, E. J. & Donnet-Hughes, A. Soluble CD14 in human breast milk and its role in innate immune responses. Acta Odontol. Scand. 59, 330–334 (2001).
Bottcher, M. F., Abrahamsson, T. R., Fredriksson, M., Jakobsson, T. & Bjorksten, B. Low breast milk TGF-beta2 is induced by Lactobacillus reuteri supplementation and associates with reduced risk of sensitization during infancy. Pediatr. Allergy Immunol. 19, 497–504 (2008).
Kvistgaard, A. S. et al. Inhibitory effects of human and bovine milk constituents on rotavirus infections. J. Dairy Sci. 87, 4088–4096 (2004).
Newburg, D. S. et al. Role of human-milk lactadherin in protection against symptomatic rotavirus infection. Lancet 351, 1160–1164 (1998).
Stene, L. C. et al. Rotavirus infection frequency and risk of celiac disease autoimmunity in early childhood: A longitudinal study. Am. J. Gastroenterol. 101, 2333–2340 (2006).
Hanayama, R. et al. Identification of a factor that links apoptotic cells to phagocytes. Nature 417, 182–187 (2002).
Moossavi, S. et al. Composition and variation of the human milk microbiota are influenced by maternal and early-life factors. Cell Host Microbe 25(324–335), e4 (2019).
Simre, K. et al. Exploring the risk factors for differences in the cumulative incidence of coeliac disease in two neighboring countries: The prospective DIABIMMUNE study. Dig. Liver Dis. 48, 1296–1301 (2016).
Oberhuber, G., Granditsch, G. & Vogelsang, H. The histopathology of coeliac disease: Time for a standardized report scheme for pathologists. Eur. J. Gastroenterol. Hepatol. 11, 1185–1194 (1999).
Zeller, G. et al. Potential of fecal microbiota for early-stage detection of colorectal cancer. Mol. Syst. Biol. 10, 766 (2014).
Haarman, M. & Knol, J. Quantitative real-time PCR assays to identify and quantify fecal Bifidobacterium species in infants receiving a prebiotic infant formula. Appl. Environ. Microbiol. 71, 2318–2324 (2005).
Haarman, M. & Knol, J. Quantitative real-time PCR analysis of fecal Lactobacillus species in infants receiving a prebiotic infant formula. Appl. Environ. Microbiol. 72, 2359–2365 (2006).
Satokari, R. M., Vaughan, E. E., Akkermans, A. D., Saarela, M. & de Vos, W. M. Bifidobacterial diversity in human feces detected by genus-specific PCR and denaturing gradient gel electrophoresis. Appl. Environ. Microbiol. 67, 504–513 (2001).
Walter, J. et al. Detection and identification of gastrointestinal Lactobacillus species by using denaturing gradient gel electrophoresis and species-specific PCR primers. Appl. Environ. Microbiol. 66, 297–303 (2000).
Bartosch, S., Fite, A., Macfarlane, G. T. & McMurdo, M. E. Characterization of bacterial communities in feces from healthy elderly volunteers and hospitalized elderly patients by using real-time PCR and effects of antibiotic treatment on the fecal microbiota. Appl. Environ. Microbiol. 70, 3575–3581 (2004).
The project was funded by the European Commission (7th Framework Programme, project 202063), the Estonian Research Council (grants nos. IUT20-43; IUT34-19; PRG712; PRG1428; PRG1120), the Estonian Ministry of Education and Research (grant no. KOGU-HUMB), the Academy of Finland (the Centre of Excellence in Molecular Systems Immunology and Physiology Research, 2012–2017, Decision Nos. 250114 and 284597), and Enterprise Estonia (grant no. EU48695). We thank Irja Roots, Kristi Alnek, and Helis Janson for technical assistance.
Competing of interests
The authors declare no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
The original online version of this Article was revised: The original version of this Article contained an error in the Acknowledgments section. It now reads: “The project was funded by the European Commission (7th Framework Programme, project 202063), the Estonian Research Council (grants nos. IUT20-43; IUT34-19; PRG712; PRG1428; PRG1120), the Estonian Ministry of Education and Research (grant no. KOGU-HUMB), the Academy of Finland (the Centre of Excellence in Molecular Systems Immunology and Physiology Research, 2012–2017, Decision Nos. 250114 and 284597), and Enterprise Estonia (grant no. EU48695). We thank Irja Roots, Kristi Alnek, and Helis Janson for technical assistance.”
About this article
Cite this article
Štšepetova, J., Simre, K., Tagoma, A. et al. Maternal breast milk microbiota and immune markers in relation to subsequent development of celiac disease in offspring. Sci Rep 12, 6607 (2022). https://doi.org/10.1038/s41598-022-10679-x