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BioMed Research International / 2014 / Article
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Recent Advances and Future Perspective in Microbiota and Probiotics

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Research Article | Open Access

Volume 2014 |Article ID 301053 |

Marta Simone, Caterina Gozzoli, Andrea Quartieri, Giuseppe Mazzola, Diana Di Gioia, Alberto Amaretti, Stefano Raimondi, Maddalena Rossi, "The Probiotic Bifidobacterium breve B632 Inhibited the Growth of Enterobacteriaceae within Colicky Infant Microbiota Cultures", BioMed Research International, vol. 2014, Article ID 301053, 7 pages, 2014.

The Probiotic Bifidobacterium breve B632 Inhibited the Growth of Enterobacteriaceae within Colicky Infant Microbiota Cultures

Academic Editor: Borja Sánchez
Received20 Jun 2014
Revised29 Jul 2014
Accepted31 Jul 2014
Published28 Aug 2014


Infant colic is a common gastrointestinal disorder of newborns, mostly related to imbalances in the composition of gut microbiota and particularly to the presence of gas-producing coliforms and to lower levels of Bifidobacteria and Lactobacilli. Probiotics could help to contain this disturbance, with formulations consisting of Lactobacillus strains being the most utilized. In this work, the probiotic strain Bifidobacterium breve B632 that was specifically selected for its ability to inhibit gas-producing coliforms, was challenged against the Enterobacteriaceae within continuous cultures of microbiota from a 2-month-old colicky infant. As confirmed by RAPD-PCR fingerprinting, B. breve B632 persisted in probiotic-supplemented microbiota cultures, accounting for the 64% of Bifidobacteria at the steady state. The probiotic succeeded in inhibiting coliforms, since FISH and qPCR revealed that the amount of Enterobacteriaceae after 18 h of cultivation was 0.42 and 0.44 magnitude orders lower in probiotic-supplemented microbiota cultures than in the control ones. These results support the possibility to move to another level of study, that is, the administration of B. breve B632 to a cohort of colicky newborns, in order to observe the behavior of this strain in vivo and to validate its effect in colic treatment.

1. Introduction

In the first hours of life, the germ-free gastrointestinal tract of newborns is colonized by microorganisms deriving from the mother and from the environment, with the establishment of a microbial community that will evolve into one of the most complex microbial ecosystems [1]. The maintenance of a correct balance of gut bacterial population is extremely important since microbiota performs a variety of activities and functions that deeply influence the health status of the host, such as the metabolism of nondigestible compounds with supply of short chain fatty acids, vitamin biosynthesis, the regulation of immune system, and the prevention of pathogen colonization [2, 3].

Despite the fact that increasing information about microbiota composition in adults is arising from metagenomics and other culture-independent approaches, the dynamics of initial colonization and evolution of the bacterial community during the first days of life are poorly understood so far [4]. In newborns, microbiota composition is variable and unstable, and the establishment of the intestinal microbiota is highly dependent on many factors, such as the mode of birth, breast or formula feeding, and antibiotic intake [57]. Furthermore, factors affecting the tropism and host-microbe interactions, such as intestinal pH, body temperature, bile acids, peristalsis, mucosal immune response receptors, and internal synergy, exert a pivoting role in shaping the composition of bacterial population [8, 9]. Initially, culturing studies indicated that the pioneer bacteria colonizing the digestive tract of newborns are Enterobacteriaceae and Gram-positive cocci (e.g., Streptococcus, Staphylococcus), which lower the redox potential and generate an anoxic environment, favorable for the establishment of strictly anaerobic bacteria, such as Bacteroidetes, Bifidobacterium, and Clostridiales [8, 10]. Bifidobacteria are generally reported to prevail in the gut microbiota of naturally delivered breast-fed infants after a few days, at the expenses of Enterobacteriaceae and facultative aerobes [11]. However, culture independent investigations have provided evidence that infant colonization may be much more complex, since it may be primed by anaerobes as well (e.g. Clostridiales) and Bifidobacteria may not be among the first colonizers or may remain a numerical minority [12].

Infant colic is a common functional gastrointestinal disorder of newborns, characterized by long bouts of crying and hard-to-relieve behavior [13]. Crying peaks range between 6 and 12 weeks of age and cause considerable concern and distress to parents. The pathogenesis of infant colic is not well understood, and several underlying causes have been suggested [13]. Among them, the relationship between colonic microbiota and this disorder is emerging as a major determinant. Culturing studies revealed higher counts of Gram-negative bacteria and a less numerous population of Lactobacilli and Bifidobacteria in the feces of colicky infants compared with healthy infants [14]. Molecular global investigation of the microbiota composition through phylogenetic microarray analysis demonstrated that gut microbiota differentiate much more slowly in colicky infants than in healthy ones and that colic correlated positively with the presence of specific genera of Gammaproteobacteria (such as Escherichia, Klebsiella, Serratia, Vibrio, Yersinia, and Pseudomonas) and negatively with bacteria belonging to the Bacteroidetes and Firmicutes [15, 16]. Consistently, it is known that Enterobacteriaceae, such as bacteria belonging to Escherichia and Klebsiella, produce gas from mixed acid fermentation and proinflammatory lipopolysaccharides, both these mechanisms being proposed to favor colic [17, 18].

The microbiota of colicky infants also presents lower amounts of Bifidobacteria and Lactobacilli, which are known to be anti-inflammatory and to exert various healthy properties [1921]. The intake of probiotic Lactobacilli during the first months of life can contribute to containing colic [22, 23]. On the contrary, in vivo studies utilizing probiotic Bifidobacteria for the treatment of colic are lacking. The strain Bifidobacterium breve B632 possesses antimicrobial activity against gas-producing coliforms isolated from the stools of infants suffering from colic [24].

In order to obtain preliminary results that could support an in vivo trial, the present study challenged B. breve B632 against the Enterobacteriaceae within cultures of microbiota from a 2-month-old colicky infant. A continuous culture fermentation simulating the gut microbiota of a colicky infant was performed to examine the time-course of E. coli and Enterobacteriaceae populations.

2. Methods

2.1. Chemicals and Bacterial Strain

All the chemicals were supplied by Sigma (Stenheim, Germany), unless otherwise stated. Bifidobacterium breve B632 was obtained from BUSCoB strain collection (Scardovi Collection of Bifidobacteria, Dept. of Agro-Environmental Science and Technology, University of Bologna, Italy). The strain was accepted for deposit by DSMZ for patent purposes and named B. breve DSMZ 24706. It was cultured anaerobically at 37°C in Lactobacilli MRS broth (BD Difco, Sparks, USA) containing 0.5 g/L L-cysteine hydrochloride (hereinafter called MRS).

2.2. Cultures of Gut Microbiota

The cultures of gut microbiota were performed in a microbiota medium MM [25], where the carbon source was substituted with 6.0 g/L of a mixture of galactooligosaccharides (GOS, Domo Vivinal, Needseweg, The Netherlands) and fructooligosaccharides (FOS, Beneo-Orafti P95, Oreye, Belgium). The mixture was composed of 90% GOS and 10% FOS (w/w), in agreement with the composition of prebiotic infant formula [26]. Oligosaccharides were filter-sterilized (0.22 μm) and added to the medium after autoclaving.

Fresh feces from a breast-fed colicky infant, born by natural delivery and not treated with antibiotics or probiotics, were utilized to prepare the inoculum for single-stage continuous cultures. Inoculum preparation was performed in anaerobic cabinet under an 85% N2, 10% CO2, and 5% H2 atmosphere. Feces were diluted to the ratio of 1 : 10 (w/v) in MM, supplemented with 10% glycerol (v/v), and stored at −80°C until use.

In control microbiota cultures (MC), 5 mL of fecal suspension was thawed at 37°C and utilized to inoculate bench-top bioreactors (Sixfors V3.01, Infors, Bottmingen, Swiss) containing 250 mL of MM. Fresh MM was fed at the dilution rate of 0.042 h−1, corresponding to one turnover per day. The medium was flushed with CO2 to maintain anaerobiosis. The culture was kept in anaerobiosis at 37°C, under gentle agitation. Automatic titration with 4 M NaOH maintained pH at 6.5.

In probiotic-supplemented microbiota cultures (PMC), fecal cultures were supplemented with 5.0  cfu/mL of B. breve B632. Concentrated stock cultures of B. breve B632 were supplemented with glycerol (10%, v/v), enumerated onto MRS-agar plates, and stored at −80°C until an appropriate volume was thawed and used for bioreactor inoculation.

Samples from MC and PMC were periodically collected to analyze fermentation products, to examine the microbiota composition, and to enumerate and isolate bifidobacteria.

2.3. Fluorescent In Situ Hybridization (FISH)

FISH enumeration of total bacteria, bifidobacteria, and Enterobacteriaceae was based on the procedure of Harmsen et al. [27], with slight modifications. Culture samples were diluted to the ratio of 1 : 4 with 40 g/L paraformaldehyde and incubated overnight at 4°C. Fixed cells were washed with PBS at pH 7.4 and then dehydrated with PBS-ethanol 1 : 1 solution for 1 h at 4°C. The probes Eub 338, Bif 164, and Enterobact D, were used for total bacteria, bifidobacteria, and Enterobacteriaceae, respectively [28]. To perform hybridization, 10 μL of cell suspension, 1 μL of the specific FITC-labeled probe, and 100 μL of hybridization buffer (20 mM TRIS-HCl, 0.9 M NaCl, and 0.1% SDS) were mixed and incubated for 16 h at the temperature specific for each probe [28].

A proper amount of the cell suspension was diluted in 4 mL of washing buffer (20 mM TRIS-HCl, 0.9 M NaCl) and maintained at hybridization temperature for 10 min before being filtered onto 0.2 μm polycarbonate filters (Millipore, Ettenleur, The Netherlands). Filters were mounted on microscope slides with Vectashield (Vector Labs, Burlingame, California). The slides were evaluated with a fluorescence microscope (Eclipse 80i, Nikon Instruments) equipped with mercury arc lamp, FITC specific filter, and digital camera. Depending on the number of fluorescent cells, 30 to 100 microscopic fields were counted and averaged in each slide. Each sample was enumerated in triplicate.

2.4. qPCR

Biomass samples from MC and PMC cultures were collected by centrifugation, suspended in PBS (pH 7.8), and extracted with QIAmp DNA Stool Mini Kit (Qiagen, Hilden, Germany) to obtain bacterial gDNA. gDNA was quantified with NanoPhotometer P-Class (Implen GmbH, Munchen, Germany), diluted to 2.5 ng/μL in TE buffer pH 8, and subjected to qPCR analysis with primers targeting Enterobacteriaceae and Escherichia coli [2931]. The set of primers Eco-F (GTTAATACCTTTGCTCATTGA)/Eco-R (ACCAGGGTATCAATCCTGTT) and Ent-F (ATGGCTGTCGTCAGCTCGT)/Ent-R (CCTACTTCTTTTGCAACCCACTC) were used for Enterobacteriaceae and Escherichia coli, respectively. The mixture contained 10 μL of SsoFast EvaGreen Supermix, 4 μL of each 2 μM primer, and 2 μL of template. qPCR reaction was carried out with the CFX96 Real-Time System (Bio-Rad Laboratories, Redmond, WA, USA), according to the following protocol: 98°C for 2 min; 45 cycles at 98°C for 0.05 min, 60°C for 0.05 min, and 95°C for 1 min; 65°C for 1 min.

2.5. RAPD-PCR Tracing of Bifidobacterium breve B632

Fresh culture samples were serially diluted in Wilkins-Chalgren anaerobe broth (Oxoid) in the anaerobic cabinet and plated on RB selective medium, in order to count and isolate Bifidobacteria [32]. Genomic DNA was extracted from 200 colonies isolated from the PMC processes, using Instagene matrix (Bio-Rad). RAPD-PCR was carried out in a 15 μL reaction mixture: 10X Dream Taq Buffer (including MgCl2 2 mM), 1.5 μL; dNTPs mixture 0.10 mM, 0,15 μL; 2 μM M13 primer (GAGGGTGGCGGTTCT), 3.75 μL; genomic DNA, 3 μL; and PCR water 5.25 μL. DNA amplification was performed with the following protocol: 94°C for 4 min (1 cycle), 94°C for 1 min, 34°C for 1 min, 72°C for 2 min (45 cycles); 72°C for 7 min (1 cycle). The PCR products were electrophoresed in a 2% agarose gel (25 × 25 cm) for 4 h at a constant voltage (160 V) in TAE buffer (40 mM Tris-acetate, 1 mM EDTA, and pH 8.0). RAPD-PCR profiles were visualized under ultraviolet light after staining with ethidium bromide, followed by digital image capturing. The resulting fingerprints were analyzed by the Gene Directory 2.0 (Syngene, UK) software package. The similarity among digitalized profiles was calculated and a dendrogram was derived with an unweighted pair-group method using arithmetic means (UPGMA).

2.6. Analysis of Fermentation Products

The samples were clarified through centrifugation (13,000 ×g, 5 min, 4°C) and filtration (0.22 μm cellulose acetate filter) and stored at −20°C until analyzed. Fermentation products (formic, acetic, lactic, propionic, butyric, and succinic acids and ethanol) were analyzed using a HPLC device (Agilent technologies, Waldbronn, Germany) equipped with refractive index detector and Aminex HPX-87 H ion exclusion column. Isocratic elution was carried out with 0.005 M H2SO4 at 0.6 mL/min [33].

2.7. Statistical Analysis

All values are means of four separate experiments. Comparisons were carried out according to Student’s -test. Differences were considered statistically significant for .

3. Results

3.1. Evolution of Fecal Microbial Groups and Fermentation Products

Single-stage continuous fermentation of the colonic microbiota from a colicky newborn was carried out for 24 h to study whether the addition of B. breve B632 could affect the growth of Enterobacteriaceae. Bifidobacteria, Enterobacteriaceae, and total bacteria were enumerated in MC and in PMC, the latter supplemented with 5.0  cfu/mL of B. breve B632 (Figures 1(a) and 1(b)). After 18 h of cultivation, FISH bacterial counts became steady in both MC and PMC cultures. Eubacteria increased up to 9.0–9.4  cfu/mL, without statistically significant difference between PMC and MC (). At all the time points, bifidobacteria were more abundant in PMC than in MC (). Enterobacteriaceae were negatively affected by the presence of B. breve B632 and were always less numerous in PMC than in MC ().

The evolution of Enterobacteriaceae and E. coli was determined also with q-PCR during the whole process. Enterobacteriaceae were significantly lower in PMC than in MC (), consistently with FISH results. On the other hand, statistically significant difference was not observed in the levels of E. coli (), with the exception of 18 h, when E. coli was less numerous in MC than in PMC (Figure 2).

The presence of B. breve B632 in PMC cultures was traced using RAPD-PCR fingerprinting at all the time points. Colonies were isolated using the Bifidobacterium selective medium RB and those positive to Bifidobacterium-specific PCR were subjected to RAPD-PCR analysis. At the beginning of the fermentation, B. breve B632 represented the 85% of bifidobacterial isolates in PMC, then decreased to 73% after 6 h, and stabilized at 64% at the steady state (, SD < 34%). The relative amount of B. breve B632 tended to decrease, albeit differences at the diverse time points were not statistically significant. Considering that at the steady state Bifidobacteria accounted for approximately 38% of total eubacteria according to FISH enumeration, B. breve B632 can be estimated as approximately the 24% of total bacterial population in PMC. In these samples, 2 biotypes of Bifidobacteria represented the autochthonous component. The same two biotypes were identified also at the inoculum in MC cultures, together with two other minor ones, none of them exhibiting a RAPD-PCR profile similar to that of B. breve B632.

Formate, acetate, lactate, propionate, butyrate, and ethanol originated by microbiota metabolism during the processes (Figures 3(a) and 3(b)). Like the bacterial counts, the concentrations of microbial products became stationary after approximately 18 h. Ethanol, formate, lactate, and acetate were the first to increase at the beginning of the fermentation. Propionate, 2,3-butanediol, and butyrate accumulated later, while lactate decreased as the steady state was approached.

During the growth phase, the major differences between MC and PMC processes were acetate and ethanol, accumulating at different levels during the first hours of the process: after 12 h, in MC and PMC, ethanol was 1.6 and 0.8 g/L, while acetate 0.8 and 2.4 g/L, respectively. At the steady state (18 h), MC had higher levels of butyrate and ethanol than PMC, while acetate and lactate were higher in PMC (). The other metabolites exhibited similar steady-state concentrations in PMC and MC processes ().

4. Discussion

Literature reports the use of Lactobacillus spp. strains to alleviate the symptoms of infant colic [22, 23]. On the other hand, no information is available on this specific use of bifidobacteria, although in vitro results showed that strains of Bifidobacterium can exert antimicrobial activity against gas forming coliforms [24]. Among a panel of Bifidobacterium strains that were selected as potential candidates for probiotic use against colic in infants, B. breve B632 appeared particularly promising because of its strong antimicrobial activity against coliforms, coupled to the lack of transmissible antibiotic resistance traits and cytotoxicity for the gut epithelium. Moreover, the strain is capable of adhering to gut epithelium cell lines and could stimulate gut health by increasing metabolic activity and immune response of epithelial cells [24].

In the present work, the antagonistic effect of B. breve B632 against coliforms was challenged within gut microbiota cultures of a colicky newborn, simulating in vivo conditions, in order to propose its use as anticolic probiotic. B. breve B632 survived well within the fecal culture, exhibiting a high viability during the process. At all the time points, Enterobacteriaceae were significantly less numerous in presence of the probiotic. These results indicate that B. breve B632 exerted antimicrobial activity against coliforms in fecal cultures as well, consistently with previous observation with spot agar tests and cocultures [24].

Unlike Enterobacteriaceae, E. coli counts were not affected by the presence of the probiotic. This observation can be ascribed to the different specificity of the primer sets utilized in qPCR quantification, since the primers for Enterobacteriaceae recognize a broader spectrum of species than the ones for E. coli (Table 1).

Probe or primer setGenus

Enterobact DCitrobacter



Based on the list of species that align with qPCR primers and FISH probes, it is likely that Gammaproteobacteria other than E. coli are involved in infant colic. For example, the qPCR primers for Enterobacteriaceae should recognize Yersinia, whereas the FISH probe for Enterobacteriaceae is expected to miss it.

Fecal samples have a microbial composition that does not exactly correspond to that of the colonic content, where major microbial-host interactions occur, and richness and diversity seem underrepresented [34]. However, systems as the one herein described are currently the best tools to investigate the external factors that could influence the intestinal microbial composition such as antibiotics or to test novel potential probiotics, before carrying out expensive in vivo trials. The data herein presented indicate that the potential probiotic strain B. breve B632 was able to survive in a complex microbial environment and restrained Enterobacteriaceae population.

5. Conclusions

The present study demonstrated the ability of a properly selected probiotic Bifidobacterium strain B. breve B632 to inhibit the growth of Enterobacteriaceae in an in vitro model system simulating the intestinal microbiota of a 2-month-old colicky infant. These results support the possibility to move to another level of study, that is, the administration of B. breve B632 to a cohort of colicky newborns, in order to observe the behavior of this strain in vivo and to validate its effect in colic treatment.

Conflict of Interests

The authors certify that there is no actual or potential conflict of interests in relation to this paper.


  1. L. Morelli, “Postnatal development of intestinal microflora as influenced by infant nutrition,” Journal of Nutrition, vol. 138, no. 9, pp. 1791S–1795S, 2008. View at: Google Scholar
  2. F. Guarner and J. R. Malagelada, “Gut flora in health and disease,” The Lancet, vol. 361, no. 9356, pp. 512–519, 2003. View at: Publisher Site | Google Scholar
  3. I. Sekirov, S. L. Russell, L. Caetano M Antunes, and B. B. Finlay, “Gut microbiota in health and disease,” Physiological Reviews, vol. 90, no. 3, pp. 859–904, 2010. View at: Publisher Site | Google Scholar
  4. D. di Gioia, I. Aloisio, G. Mazzola, and B. Biavati, “Bifidobacteria: their impact on gut microbiota composition and their applications as probiotics in infants,” Applied Microbiology and Biotechnology, vol. 98, no. 2, pp. 563–577, 2014. View at: Publisher Site | Google Scholar
  5. M. G. Dominguez-Bello, E. K. Costello, M. Contreras et al., “Delivery mode shapes the acquisition and structure of the initial microbiota across multiple body habitats in newborns,” Proceedings of the National Academy of Sciences of the United States of America, vol. 107, no. 26, pp. 11971–11975, 2010. View at: Publisher Site | Google Scholar
  6. E. Bezirtzoglou, A. Tsiotsias, and G. W. Welling, “Microbiota profile in feces of breast- and formula-fed newborns by using fluorescence in situ hybridization (FISH),” Anaerobe, vol. 17, no. 6, pp. 478–482, 2011. View at: Publisher Site | Google Scholar
  7. F. Fouhy, R. P. Ross, G. F. Fitzgerald, C. Stanton, and P. D. Cotter, “Composition of the early intestinal microbiota: knowledge, knowledge gaps and the use of high-throughput sequencing to address these gaps,” Gut Microbes, vol. 3, no. 3, pp. 203–220, 2012. View at: Publisher Site | Google Scholar
  8. R. I. Mackie, A. Sghir, and H. R. Gaskins, “Developmental microbial ecology of the neonatal gastrointestinal tract,” The American Journal of Clinical Nutrition, vol. 69, no. 5, pp. 1035S–1045S, 1999. View at: Google Scholar
  9. L. V. Hooper, D. R. Littman, and A. J. Macpherson, “Interactions between the microbiota and the immune system,” Science, vol. 336, no. 6086, pp. 1268–1273, 2012. View at: Publisher Site | Google Scholar
  10. C. F. Favier, E. E. Vaughan, W. M. de Vos, and A. D. L. Akkermans, “Molecular monitoring of succession of bacterial communities in human neonates,” Applied and Environmental Microbiology, vol. 68, no. 1, pp. 219–226, 2002. View at: Publisher Site | Google Scholar
  11. K. Orrhage and C. E. Nord, “Factors controlling the bacterial colonization of the intestine in breastfed infants,” Acta Paediatrica, Supplement, vol. 88, no. 430, pp. 47–57, 1999. View at: Google Scholar
  12. Y. Vallès, M. J. Gosalbes, L. E. de Vries, J. J. Abellán, and M. P. Francino, “Metagenomics and development of the gut microbiota in infants,” Clinical Microbiology and Infection, vol. 18, no. 4, pp. 21–26, 2012. View at: Publisher Site | Google Scholar
  13. M. L. Cirgin Ellett, “What is known about infant colic?” Gastroenterology Nursing, vol. 26, no. 2, pp. 60–65, 2003. View at: Google Scholar
  14. F. Savino, F. Cresi, S. Pautasso et al., “Intestinal microflora in breastfed colicky and non-colicky infants,” Acta Paediatrica, vol. 93, no. 6, pp. 825–829, 2004. View at: Publisher Site | Google Scholar
  15. C. de Weerth, S. Fuentes, and W. M. de Vos, “Crying in infants: on the possible role of intestinal microbiota in the development of colic,” Gut Microbes, vol. 4, no. 5, pp. 416–421, 2013. View at: Publisher Site | Google Scholar
  16. C. de Weerth, S. Fuentes, P. Puylaert, and W. M. de Vos, “Intestinal microbiota of infants with colic: development and specific signatures,” Pediatrics, vol. 131, no. 2, pp. e550–e558, 2012. View at: Publisher Site | Google Scholar
  17. F. Savino, L. Cordisco, V. Tarasco, R. Calabrese, E. Palumeri, and D. Matteuzzi, “Molecular identification of coliform bacteria from colicky breastfed infants,” Acta Paediatrica, vol. 98, no. 10, pp. 1582–1588, 2009. View at: Publisher Site | Google Scholar
  18. F. Savino, L. Cordisco, V. Tarasco et al., “Antagonistic effect of Lactobacillus strains against gas-producing coliforms isolated from colicky infants,” BMC Microbiology, vol. 11, article 157, 2011. View at: Publisher Site | Google Scholar
  19. T. A. Oelschlaeger, “Mechanisms of probiotic actions—a review,” International Journal of Medical Microbiology, vol. 300, no. 1, pp. 57–62, 2010. View at: Publisher Site | Google Scholar
  20. M. Rossi, A. Amaretti, A. Leonardi, S. Raimondi, M. Simone, and A. Quartieri, “Potential impact of probiotic consumption on the bioactivity of dietary phytochemicals,” Journal of Agricultural and Food Chemistry, vol. 61, no. 40, pp. 9551–9558, 2013. View at: Google Scholar
  21. M. Rossi, A. Amaretti, and S. Raimondi, “Folate production by probiotic bacteria,” Nutrients, vol. 3, no. 1, pp. 118–134, 2011. View at: Publisher Site | Google Scholar
  22. F. Savino, L. Cordisco, V. Tarasco et al., “Lactobacillus reuteri DSM 17938 in infantile colic: a randomized, double-blind, placebo-controlled trial,” Pediatrics, vol. 126, no. 3, pp. e526–e533, 2010. View at: Publisher Site | Google Scholar
  23. J. Anabrees, F. Indrio, B. Paes, and K. Alfaleh, “Probiotics for infantile colic: a systematic review,” BMC Pediatrics, vol. 13, p. 186, 2013. View at: Google Scholar
  24. I. Aloisio, C. Santini, B. Biavati et al., “Characterization of Bifidobacterium spp. strains for the treatment of enteric disorders in newborns,” Applied Microbiology and Biotechnology, vol. 96, no. 6, pp. 1561–1576, 2012. View at: Publisher Site | Google Scholar
  25. F. Tomas-Barberan, R. García-Villalba, A. Quartieri et al., “In vitro transformation of chlorogenic acid by human gut microbiota,” Molecular Nutrition and Food Research, vol. 58, no. 5, pp. 1122–1131, 2014. View at: Google Scholar
  26. E. Bruzzese, M. Volpicelli, V. Squeglia et al., “A formula containing galacto- and fructo-oligosaccharides prevents intestinal and extra-intestinal infections: an observational study,” Clinical Nutrition, vol. 28, no. 2, pp. 156–161, 2009. View at: Publisher Site | Google Scholar
  27. H. J. M. Harmsen, P. Elfferich, F. Schut, and G. W. Welling, “A 16S rRNA-targeted probe for detection of lactobacilli and enterococci in faecal samples by fluorescent in situ hybridization,” Microbial Ecology in Health and Disease, vol. 11, no. 1, pp. 3–12, 1999. View at: Publisher Site | Google Scholar
  28. A. W. Walker, S. H. Duncan, E. C. M. Leitch, M. W. Child, and H. J. Flint, “pH and peptide supply can radically alter bacterial populations and short-chain fatty acid ratios within microbial communities from the human colon,” Applied and Environmental Microbiology, vol. 71, no. 7, pp. 3692–3700, 2005. View at: Publisher Site | Google Scholar
  29. T. Rinttilä, A. Kassinen, E. Malinen, L. Krogius, and A. Palva, “Development of an extensive set of 16S rDNA-targeted primers for quantification of pathogenic and indigenous bacteria in faecal samples by real-time PCR,” Journal of Applied Microbiology, vol. 97, no. 6, pp. 1166–1177, 2004. View at: Publisher Site | Google Scholar
  30. E. Malinen, A. Kassinen, T. Rinttilä, and A. Palva, “Comparison of real-time PCR with SYBR green I or 5′-nuclease assays and dot-blot hybridization with rDNA-targeted oligonucleotide probes in quantification of selected faecal bacteria,” Microbiology, vol. 149, no. 1, pp. 269–277, 2003. View at: Publisher Site | Google Scholar
  31. M. Castillo, S. M. Martín-Orúe, E. G. Manzanilla, I. Badiola, M. Martín, and J. Gasa, “Quantification of total bacteria, enterobacteria and lactobacilli populations in pig digesta by real-time PCR,” Veterinary Microbiology, vol. 114, no. 1-2, pp. 165–170, 2006. View at: Publisher Site | Google Scholar
  32. R. Hartemink, B. J. Kok, G. H. Weenk, and F. M. Rombouts, “Raffinose-Bifidobacterium (RB) agar, a new selective medium for bifidobacteria,” Journal of Microbiological Methods, vol. 27, no. 1, pp. 33–43, 1996. View at: Publisher Site | Google Scholar
  33. A. Amaretti, T. Bernardi, A. Leonardi, S. Raimondi, S. Zanoni, and M. Rossi, “Fermentation of xylo-oligosaccharides by Bifidobacterium adolescentis DSMZ 18350: kinetics, metabolism, and β-xylosidase activities,” Applied Microbiology and Biotechnology, vol. 97, no. 7, pp. 3109–3117, 2013. View at: Publisher Site | Google Scholar
  34. A. Durbán, J. J. Abellán, N. Jiménez-Hernández et al., “Assessing gut microbial diversity from feces and rectal mucosa,” Microbial Ecology, vol. 61, no. 1, pp. 123–133, 2011. View at: Publisher Site | Google Scholar

Copyright © 2014 Marta Simone et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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