INTRODUCTION

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Although probiotics play an important role in animal nutrition, their modes of action have not been determined yet. Microbial probiotics have been reported to have many beneficial effects when they are used in animal feeds; these effects include competitive exclusion of pathogens (6, 12) and improved digestion and absorption of nutrients (6, 19, 21, 23). It has been proposed that the efficiency of probiotics could be enhanced by genetic modifications that increase the enzymatic capacity of the gut through improvements in plant cell wall hydrolase production. However, there are concerns about using recombinant organisms in animals, particularly with respect to the transfer of antibiotic resistance genes and the possibility that a recombinant gene may move up through the food chain (9, 16, 20). To be of benefit to an animal, a probiotic organism must have an impact on the microbiology of the gut. To evaluate the usefulness of probiotics and to assess the risks, if any, associated with their use, it is important to understand the nature of the interactions. Culture techniques have not been entirely successful in assessing these interactions (26). Molecular methods, such as nucleic acid probe analysis, enzyme-linked immunoassays, and PCR, have also been used to detect probiotic strains and their effects, but the studies have been limited to detection of specific bacteria (2, 4, 10). Recent advances in the development of methods to study microbial ecology (1, 5, 7, 18, 26) have extended the range of techniques available for studying the impact of probiotics on microbial community structure in the gastrointestinal (GI) tract. In this study we used a variation of the community DNA hybridization procedures introduced by Lee and Fuhrman (7) to investigate changes in bacterial community complexity following introduction of genetically modified (GM) and non-GM strains of Enterococcus faecium into the diet of 1-day-old chicks. The aims of this study were to evaluate the impact of a GM probiotic on the bacterial flora of the chicken GI tract and to establish whether the observed changes were distinct from the changes observed when the parental, nonengineered strain was used.

	MATERIALS AND METHODS

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Probiotic organisms. The GM probiotic (probiotic A) was constructed by transforming erythromycin-resistant (Ery^r) plasmid pVACMC1 containing the Ruminococcus flavefaciens -1,4-glucanase gene (27) into E. faecium NCIMB 11508. The parent strain, E. faecium NCIMB 11508, was used as the non-GM probiotic (probiotic B).

Impact of probiotics on the chicken GI tract. Three groups of chickens were grown from 1 day old to 4 weeks old by using the following three treatments: (i) commercial diet containing probiotic A (rifampin-resistant [Rif^r] E. faecium containing pVACMC1) at a concentration of 10⁵ CFU/g; (ii) commercial diet containing probiotic B (Rif^r E. faecium) at a concentration of 10⁵ CFU/g; and (iii) commercial diet containing no probiotic (control).

Bacterial culture. Viable counts were obtained by using serial 10-fold dilutions of the gut content samples in phosphate-buffered saline that were plated in triplicate onto the appropriate selective media. The counts were determined by using the dilution that produced between 30 and 300 CFU per plate. All of the selective media were supplied by Oxoid Ltd., Basingstoke, United Kingdom, and were prepared according to the manufacturer's instructions. All plates were incubated for 48 h as recommended by the manufacturer of the selective media, and the plates were used to determine viable counts as follows: Lactobacillus spp. were counted on MRS agar (De Man, Rogosa, Sharpe); Enterococcus spp. were counted on Slanetz-Bartley medium; coliforms were counted on cystine-lactose electrolyte-deficient medium; Salmonella spp. were counted directly on desoxycholate lactose sucrose (DCLS) agar and by using enrichment selective Rappaport-Vassiliadis broth cultures that were subcultured on brilliant green agar; Campylobacter spp. were counted on campylobacter selective media; and M17 medium containing antibiotics (50 µg of rifampin per ml and 200 µg of erythromycin per ml) was used to select for probiotic A (rifampin and erythromycin resistant) and probiotic B (rifampin resistant).

Community DNA analysis. (i) DNA extraction. DNA was extracted from bacterial cultures by using the method of Netherwood et al. (13). DNA was extracted from gut contents by using an additional bead-beating step. Sterile glass beads (25 µg; diameter, 0.17 to 0.18 mm) were added to a guanidine thiocyanate-gut content mixture, and the preparation was shaken for 2 min at 1,600 rpm with a Mikrodismembrator U instrument (B. Braun Biotech International) before the heating step was performed.

(ii) PCR amplification of 16S ribosomal DNA (rDNA). DNA extract (1 µl) was added to a PCR mixture containing 10 mM Tris-HCl (pH 8.8) at 25°C, 1.5 mM MgCl₂, 50 mM KCl, 0.1% Triton X-100, each deoxynucleoside triphosphate at a concentration of 500 µM, 20 pmol of primer Eub338 GC clamp (5'-CGC CCG CCG CGC CCC CGC CCC GGC CCG CCG CCC CCG CCC GCT GCC TCC CGT AGG AGT-3'), 20 pmol of primer Univ1390 (5'-ACG GGC GGT GTG TRC-3'), and 1 U of DyNAzyme II DNA polymerase (Flowgen). Amplification was carried out by using one thermal cycle consisting of 5 min at 95°C, followed by 30 cycles consisting of 1 min at 94°C, 1 min at 57°C, and 2 min at 72°C. The final cycle consisted of 72°C for 20 min. Negative controls containing no DNA and eukaryotic DNA were included in addition to a positive control containing Escherichia coli 16S ribosomal DNA.

(iii) Oligonucleotide probe hybridization. The oligonucleotide DNA probes were radiolabelled with a synthetic oligonucleotide 5' end labelling kit obtained from MBI Fermentas according to the manufacturer's instructions. Hybridization was carried out by using the instructions for a Hybond-N membrane supplied by Amersham Life Sciences, Little Chalfont, United Kingdom, at 65°C for 18 h. Negative and positive controls for the probes, which consisted of DNA extracts of Enterococcus faecalis, E. faecium, Campylobacter sp., Salmonella sp., and Escherichia coli cultures, salmon sperm DNA, and no DNA, were applied to each membrane.

Community hybridization. The amplified 16S rDNA was radiolabelled by using the nick translation method with a HexaLabel DNA labelling kit as recommended by the manufacturer (MBI Fermentas, Graiciuno, Lithuania). Reciprocal hybridizations were carried out by using the method of Lee and Fuhrman (7). Hybridization results were quantified by using a radioactivity imaging system (Instant Imager; Packard).

	RESULTS

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Culture analysis of the avian GI tract following addition of GM and non-GM probiotics. Both the GM and non-GM probiotics became established in the gut at an initial concentration of 10⁵ CFU/g of gut contents. After 4 weeks of feeding with the GM probiotic, the level increased to 10⁷ CFU/g of gut contents (14). Within 5 days after the probiotics were eliminated from the diet, neither probiotic could be recovered from gut content samples.

Community DNA analysis of the avian GI tract following addition of GM and non-GM probiotics. (i) Reciprocal hybridization. The three replicate PCR mixtures derived from the same target DNA and obtained at each specific sampling time were combined prior to the reciprocal hybridization analysis. This reduced the effect of PCR drift, the inherent difference between replicate PCR caused by slightly different reaction conditions that affect the annealing rates of probes to different DNA species.

View larger version (12K): [in this window] [in a new window]   FIG. 1.   Levels of similarity of the bacterial communities in the group treated with probiotic A (GM strain) and the control (untreated) group. The two values at each sampling time are means based on three replicates of two reciprocal hybridizations. a:c, the probiotic A group was the target and the untreated group was the probe; c:a, the untreated group was the target and the probiotic A group was the probe. The true level of similarity is indicated by the line connecting the lower similarity values. The probiotic was removed from the diet after 28 days. The values are means ± standard errors of the means of results from three replicate experiments.

View larger version (14K): [in this window] [in a new window]   FIG. 2.   Levels of similarity of the bacterial communities in the group treated with probiotic B (unmodified strain) and the control (untreated) group. The two values at each sampling time are means based on three replicates of two reciprocal hybridizations. b:c, the probiotic B group was the target and the untreated group was the probe; c:b, the untreated group was the target and the probiotic B group was the probe. The true level of similarity is indicated by the line connecting the lower similarity values. The probiotic was removed from the diet after 28 days. The values are means ± standard errors of the means of results from three replicate experiments.

View larger version (13K): [in this window] [in a new window]   FIG. 3.   Levels of similarity of the bacterial communities in the group treated with probiotic A (GM strain) and the group treated with probiotic B (unmodified strain). The two values at each sampling time are means based on three replicates of two reciprocal hybridizations. a:b, the probiotic A group was the target and the probiotic B group was the probe; b:a, the probiotic B group was the target and the probiotic A group was the probe. The true level of similarity is indicated by the line connecting the lower similarity values. Probiotics were removed from the diets after 28 days. The values are means ± standard errors of the means of results from three replicate experiments.

(ii) Oligonucleotide probe analysis of 16S rDNA. Specific probing of the bound rDNA with a probe specific for E. faecalis (which did not cross-react with E. faecium) suggested that there were significant differences between the effects of the GM and non-GM strains on the numbers of E. faecalis cells in the GI tract (Fig. 4). GM probiotic A resulted in a decrease in the relative amount of E. faecalis compared with the untreated control group, while with probiotic B the numbers (expressed relative to the binding of a bacterial probe) increased relative to the control group. No significant difference with time was observed for the hybridization results obtained with the probes for E. faecium, E. coli, Salmonella spp., and Campylobacter spp. either in individual treatment groups or between groups.

View larger version (16K): [in this window] [in a new window]   FIG. 4.   Change in the ratio of E. faecalis to bacteria in the three treatment groups. The ratio of E. faecalis to total bacteria was estimated by measuring the levels of probe hybridization to PCR-amplified regions of 16S DNA for samples treated with probiotic A (GM strain) and probiotic B (unmodified strain) and control (untreated) samples. The values are means ± standard errors of the means based on three replicates. Probiotics were removed from the diets after 28 days.

	DISCUSSION

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Viable counting revealed no significant differences between the groups treated with the GM and non-GM probiotics and the control group. Viable counting on selective media has been shown to recover less than 20% of bacteria (26); the remainder are viable but not culturable under the isolation conditions used (1). However, the use of molecular techniques enables the unculturable component to be studied, and here we used a modification of the community hybridization technique introduced by Lee and Fuhrman (7) to investigate changes in bacterial communities as reflected in their 16S rDNA amplicons. This is a useful modification of community analysis methods since unlike the original procedure, in which total community DNA was hybridized (7), it allows workers to analyze bacterial DNA separately from archaeal or eucaryal DNA. Furthermore, DNA extraction can be carried out more rapidly on a small scale compared with whole-community DNA hybridization analysis, and a strong hybridization signal can be produced reliably. However, there are problems with using PCR for microbial community analysis, such as errors due to variations in gene copy number, PCR drift, and primer binding bias (17, 22, 24). A recent analysis of these problems by Polz and Cavanaugh (15) indicated that gene copy number is unlikely to be a major cause of observed bias. Biases due to equipment variations, pipetting errors, and template composition can be reduced considerably by using high template concentrations, by using fewer cycles, and by mixing replicate reaction preparations. Thus, PCR error can be minimized. PCR variation may still occur, but it is likely to be reproducible, which means that comparative analyses, such as those described here, are less influenced by amplification bias since the same errors are likely to occur in both of the communities studied.

Reciprocal hybridization revealed major changes in the bacterial community structure following removal of the probiotic after 28 days. At that time, when the numbers of probiotic organisms in the gut were decreasing (14), one might have expected other bacteria to be competing for the niches previously occupied by the probiotic organisms (Fig. 1 and 2). Since major differences were not apparent when probiotic A was compared with probiotic B, the data suggest that the overall community responses to the two probiotics were similar. To resolve better the differences induced by probiotic treatment, we also used denaturing gradient gel electrophoresis to investigate the changes in 16S rDNA amplicon diversity observed during rDNA hybridization. However, with this technique, which separates DNA on a denaturing gel on the basis of sequence dissimilarities, we were not able to resolve all of the bands obtained with the consensus primers (data not shown), and the technique was not investigated further.

To determine whether the probiotics changed the composition of the gut microbial community, probes specific for E. faecium, E. faecalis, Salmonella spp., Campylobacter spp., and E. coli and a general probe for all bacteria were hybridized with the bound 16S rDNA. The results of these studies suggested that there were significant changes in the relative amounts of E. faecalis following addition of the GM probiotic and the non-GM probiotic to the diet. In the presence of the non-GM probiotic, the relative amount of E. faecalis (as assessed relative to the binding of a bacterial probe) increased compared with the control, while with the GM probiotic the relative amount of E. faecalis decreased (Fig. 4). This suggests that despite the similar responses observed at the community level, the structural changes actually involved different components of the bacterial population. Although not conclusive, the data suggest that E. faecalis and E. faecium may occupy similar niches or even have a synergistic relationship. Although this possibility is perhaps predictable due to similarities in the physiology and growth of the two organisms, it awaits confirmation by in situ hybridization. When the probiotics were removed from the diet, the gut flora shifted again. As the probiotic gradually left the GI tract, the relative amount of E. faecalis changed such that it returned to the level observed in the untreated control group. This indicates that the effects observed were reversible and dependent on the continued presence of the viable probiotic in the gut. Importantly, the data show that the probiotic does not become established in the GI tract and that it would have to be supplied continuously in the diet in order to provide any beneficial effect.

The probe for E. faecium did not reveal a significant difference in the signals obtained. This indicates either that the relative amount of E. faecium in the enterococcal component of the gut was too low to produce statistically significant differences, that the microbial composition of the gut was not changed by the probiotic treatments, or that the relatively low-level signals observed were due to a lack of access of the probe to the target site (1). The probes for Salmonella spp., Campylobacter spp., and E. coli produced very low-intensity results, which indicates that the proportions of these organisms in the microbial community of the avian GI tract are relatively low. This is reflected in the very low viable counts or the lack of viable counts for Salmonella spp. and Campylobacter spp., and although the coliform viable count was as high as 10⁹ CFU/ml of gut contents, it appears that E. coli accounts for a minor proportion of the coliform population. Thus, the community hybridization approach followed by specific probe analysis proposed here provides a useful and cost-effective means for screening PCR products of community DNA and for investigating changes in community structure during succession.

Finally, there was no evidence of any detrimental or beneficial effect on the health of the chickens used in these studies due to the changes in the microbial flora of the gut. Pathogens were not detected in greater numbers in the chickens fed diets supplemented with GM probiotics, and we observed no significant differences in the health or growth of the three trial groups. However, the chickens were kept under optimal conditions and were not under the sorts of stresses imposed by commercial production. Thus, any extrapolation to the safety of GM probiotics in the commercial rearing of chickens awaits further investigation.