In recent years, the prevalence of chronic diseases such as atopy (1,2) and chronic intestinal inflammation (IBD) is increasing in the Western world (3). Although there is a strong hereditary component in the development of these diseases, heredity cannot entirely account for the recent increases seen (4). There is little consistent evidence that risk factors such as increased exposure to indoor allergens or pollution are solely responsible for the increase in atopic diseases. It has been suggested that a high turnover of the appropriate bacteria provides a potent, continuous immune stimulation necessary to prevent atopic diseases (5–10,44). It is interesting therefore to note that longitudinal studies of the gut microflora of Pakistani infants (low prevalence of allergies) and Swedish infants (higher prevalence of allergies) show that most infants in both countries harbor various Escherichia coli strains; however, Pakistani infants were colonized earlier and more frequently by new E. coli strains while the strains in the microflora of Swedish infants were stable for months and years (11–13). The influence of modern hygiene on IBD has been demonstrated by a clear connection between family size, the access to hot water, and an indoor toilet (3,14,15). Using animal models, the central role of the intestinal microflora in the development of IBD is even more evident. Experimental murine colitis does not occur in any of several mutant strains when they are maintained in a germ-free environment but develops rapidly when these mice are colonized by commensal bacteria (16). Although the immune system is functional at birth, it is thought that increased hygienic standards, by depriving the neonate of the immune stimulus of microbial infection, may produce an imbalance among the different components of the immune system and result in chronic inflammatory or atopic disorders (17–20,45).
The acquisition of the gut microflora by neonates has been studied thoroughly (21). During and shortly after birth, infants born vaginally are exposed mainly to microbes that originate from the mother (22–24). Infants delivered by cesarean section acquire intestinal flora mainly from the environment but also, as pointed out by Lennox-King et al. (25), from the fecal microbial strains of their mothers. The proximity of the birth canal and the anus, as well as intimate parental care of the neonate ensure transmission of microbes from mother to child.
Feeding type also influences infant gut colonization. Human milk is thought to create an environment favorable for the growth of bifidobacteria; however, study results are conflicting. Within the first month, bifidobacteria levels are equal in both breast milk and formula groups (26). However, the populations of bacteroides and lactobacilli do not appear to be influenced by feeding type (27). Furthermore, it has long been known that breastfeeding protects the infant against various infectious diseases (19,28,29).
The purpose of this study was to determine if oral administration of the probiotic micro-organism Lactobacillus rhamnosus strain GG (L. GG) colonizes the maternal gastrointestinal tract during late pregnancy and whether it results in colonization the infant.
MATERIALS AND METHODS
Oral Administration of L. GG
We identified six mothers taking L. GG (ATCC 53103, ConAgra Functional Foods, Omaha, NE, U.S.A.) orally, 2 × 109 CFU per day during late pregnancy (weeks 30–36). Administration was terminated at the time of delivery, and stool samples from the neonates and the mothers were taken at several points and analyzed for the L. GG content. Three women who did not take probiotics during pregnancy served as controls and fecal samples were taken from their infants 1 month after delivery.
Identification of L. GG in Fecal Samples
DeMan-Rogosa-Sharp agar (MRS, Remel, Lenexa, KS, U.S.A.) was used to culture L. GG-like colonies. Stool samples were serially diluted by placing 1 g (or 1 mL if liquid) of the stool into 9 mL of physiologic saline, centrifuging for 1 minute, and then serially diluting 101 through 108. A 0.1-mL sample of each dilution was plated onto MRS agar. The plates were incubated under anaerobic conditions at 37°C for approximately 48 hours. After incubation, all colonies with the characteristic creamy white appearance suggestive of L. GG were counted (30). Six randomly selected colonies from each child were picked and subcultured on MRS agar for species confirmation. All of these colonies were subsequently submitted for molecular testing after 48 hours of incubation (all colonies stained as gram-positive rods, and all had a characteristic buttery odor after incubation). Random colonies with morphologies inconsistent with L. GG that gram stained as gram-positive rods were tested using the API 20 Strep test (bio Merieux, Hazelwood, MO, U.S.A.). None of these colonies had a phenotypic pattern characteristic of the referenced strain of L. rhamnosus (ferment trehalose, sorbitol, ribose, and mannitol but did not ferment arabinose, glycogen, lactose, or raffinose).
DNA was extracted from the subcultures using the Puregene DNA Purification System (Gentra Sysems, Minneapolis, MN, U.S.A.) following the manufacturer's instructions for DNA isolated from 0.5-mL gram-positive bacterial cultures.
Molecular Identification of L. GG
Two oligonucleotide bacterial primers described by Kostman et al. (31) were used for amplification. The sequences of the primers are 5´-TTGTACACACCGCCCGT CA-3´ and 5´-GGTACCTTAGATGTTTCAGTTC-3´, which correspond to the 16S ribosomal RNA gene and the 23S ribosomal RNA gene, respectively. Primers were synthesized by the University of Nebraska Medical Center, Eppley Molecular Biology Core Laboratory. The PCR assay was performed with 5 μL of test sample DNA in a total reaction volume of 50 μL consisting of PCR buffer (20 mM Tris-HCl [pH 8.4] and 50 mM KCl); 0.1 mM (each) dATP, dGTP, dCTP, and dTTP; 1.5 mM MgCl2; 0.3 mcM (each) primers; and 1.5 U of platinum Taq DNA polymerase. Thirty-five cycles of amplification were performed in a Stratagene Robocycler (Robocycler Gradient 96 Thermal Cycler, Stratagene, La Jolla, CA) model 96 thermocycler after initial denaturation of DNA at 95°C for 4.5 minutes. Each cycle consisted of a denaturation step at 95°C for 40 seconds, an annealing step at 55°C for 40 seconds, and an extension step at 72°C for 1 minute, with a final extension at 72°C for 1 minute after the last cycle. A 581 base pair product was produced corresponding to a partial sequence of the 16S gene, a complete sequence of the internal transcribed spacer region, and a partial sequence of the 23 S gene. Included with the unknown patient samples was a referenced isolate of L. rhamnosus (ATCC 53103) as a positive control and for sequence analysis comparison. After amplification, the products were stored at 4°C until evaluated.
DNA sequencing of the amplified products was performed at the Eppley Molecular Biology Core Laboratory on a Perkin-Elmer/ABI model 373 DNA sequencer with protocols supplied by the manufacturer. For direct sequencing of the amplicons, PCR products were purified using the QIAquick PCR purification kit (Qiagen, Hilden, Germany), and directly sequenced using both 16S and 23S primers (for the original unknown patient isolates and the referenced strain of L. rhamnosus) and subsequently with the 16S primer only.
The nucleotide sequences of the unknowns were aligned with the referenced isolate sequence (581-bp) using the MacVector sequence analysis software, version 6.5 (Oxford Molecular Group, Inc., Campbell, CA, U.S.A.) alignment application. Identity of the unknown patient isolates was made visually in all cases with a >99% pairwise nucleotide sequence alignment between the referenced strain sequence and the patient unknowns. An initial BLAST search in GenBank of the unknown sequences and the referenced sequence did not yield an acceptable identity (<95% homology to sequences available in the database). The referenced strain partial 16S and 23S sequences and complete internal transcribed spacer region sequence was subsequently deposited into the National Center for Biotechnology Information (NCBI) GenBank (Washington, DC, U.S.A.) as accession number AF217610 for future BLAST search pairwise nucleotide analysis.
Successful Colonization of Neonates
No side effects were reported by the six women taking L. GG capsules twice daily. Four of six children were delivered vaginally (Infants 1–4) and two by cesarean section (Infants 5 and 6). At 1 and 6 months, all of the vaginally delivered children and one of the children delivered by cesarean section were colonized with L. GG, as documented by molecular testing. Three children were still colonized at 12 months (Infants 1, 2, and 4) and two children remained colonized as long as 24 months (Infants 1 and 4). At 3 years, no L. GG was detected in the stool of any of the previously colonized children (Infants 1–3)(Table 1).
In the stool of the three mothers tested, no L. GG was detected 1 month after delivery (Mothers 1, 4, and 6), and in one mother no L. GG was detected 24 months after delivery (Mother 4). In addition, no L. GG was identified in the stool of the siblings of two children after 2 (Infant 4) and 3 years (Infant 1). In the three children delivered vaginally to mothers not taking L. GG during pregnancy, no L. GG was identified in the stools (Infants 7–9) (Table 1).
In recent years, probiotic micro-organisms have been a focus of scientific research because of clinical observations suggesting that they are useful in treating some infectious diseases (32–34) and allergic disorders (35). Both prophylactic and therapeutic effects have been identified in infants and children (35,36). The impression that the intestinal flora plays a role in human health and disease has led to different strategies to manipulate the flora increase its health promoting impact. During parturition and rapidly thereafter, microbes from the mother and the surrounding environment colonize the gastrointestinal tract of the infant until a dense, complex microflora develops. The composition of the microflora diversifies shortly before and particularly after weaning and is influenced by the method and composition of feeding (37). The normal colonization of the human intestine is presumed to govern some aspects of the development of humoral and cellular mucosal immune function in the neonate and to maintain the normal balance between inflammation and tolerance in the gut throughout life. The addition of probiotics to the intestinal flora has some reported impact on the developing immune system (35,38).
This study shows that oral administration of L. GG to women during pregnancy results in colonization of the intestine of the woman and subsequently of the infants born to the woman, even if the infant takes no probiotics. All women taking L. GG during the last weeks of pregnancy conferred this strain to their vaginally delivered infants. Colonization was stable for at least 6 months after delivery in all children, and in two children for as long as 24 months. Our results suggest that infants born by cesarean section are exposed to maternal intestinal microflora (Infant 5) even though past experience suggests that the initial exposure after cesarean section is most likely environmental organisms from equipment, air, and other infants with the nursing staff serving as vectors for transfer (23–25,39). As documented by Lennox-King et al., however, these environmental sources do not exclude the exposure of these infants to their mothers' microflora. The first infant fecal sample was taken 1 month after birth, so colonization of the infant with L. GG could have taken place after delivery by close post-partum contact with the mother and not necessarily at the time of birth (40). Not surprisingly, L. GG was not detectable in all mothers' fecal samples shortly after delivery. This result is in accordance with results by other investigators showing only a temporary dose-response colonization of 2 to 4 days after oral administration of L. GG (41).
The effect of breastfeeding on the colonization capabilities of probiotics was not investigated in this study. It is known that bifidobacteria and other lactic acid bacteria dominate the flora of breastfed infants, in contrast to the more diverse flora of bottle-fed infants (19,38,42,43). In our study, all vaginally delivered infants were breastfed between 2 and 7 months. Thus, it is interesting that in one child, breast fed for only 2 months (Infant 3), L. GG was detectable for only 6 months, whereas all other infants breast fed for at least 5 months showed colonization with L. GG for at least 12 months. Thus, it is possible that breastfeeding created an intestinal environment favorable for the colonization of L. GG. However, with the small number of individuals studied, the effect of breastfeeding on the colonization capabilities of orally administered probiotics remains a matter of speculation.
Whether health benefits of L. GG are conferred to the infant colonized in this fashion was not examined in this study. We have shown however, that temporary colonization of an infant with L. GG may be possible by colonizing the pregnant woman before delivery. However, colonization is stable for only 6 months but in unexplained circumstances (possibly related to feeding type) may persist for as long as 24 months. Future studies should be designed to analyze these favorable factors and to examine whether health benefits of probiotics are conferred to the infant using this route of colonization.
The authors thank all of the participating mothers and infants taking part in this study and Soile Tynkkynen of Valio Ltd., Helsinki, Finland, for his help with the identification of Lactobacillus GG in the stool samples. The medication was kindly provided by ConAgra Functional Foods, Omaha, NE, U.S.A.
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