PMCCPMCCPMCC

Search tips
Search criteria 

Advanced

 
Logo of aemPermissionsJournals.ASM.orgJournalAEM ArticleJournal InfoAuthorsReviewers
 
Appl Environ Microbiol. 2010 March; 76(5): 1456–1461.
Published online 2009 December 28. doi:  10.1128/AEM.01895-09
PMCID: PMC2832380

Evaluation of Lactobacillus sobrius/L. amylovorus as a New Microbial Marker of Pig Manure[down-pointing small open triangle]

Abstract

Based on a comparison of the dominant microbial populations in 17 pig manure samples and using a molecular typing method, we identified a species, Lactobacillus sobrius and Lactobacillus amylovorus (which now are considered a single species and are designated L. sobrius/amylovorus here), that was consistently found in manure. The aim of the present study was to confirm by real-time PCR the relevance of this species as a marker of pig fecal contamination. The specificity of L. sobrius/amylovorus was evaluated in human and animal DNA extracted from feces. The real-time PCR assay then was applied to water samples, including effluents from urban wastewater treatment plants, runoff water, and rivers. L. sobrius/amylovorus was consistently present in all samples of swine origin: 48 fecal samples, 18 from raw manure and 10 from biologically treated manure at mean concentrations of 7.2, 5.9, and 5.0 log10 cells/g, respectively. The species was not detected in any of the other livestock feces (38 samples from cattle and 16 from sheep), in the 27 human fecal samples, or in the 13 effluent samples from urban wastewater treatment plants. Finally, L. sobrius/amylovorus was not detected in runoff water contaminated by cattle slurry, but it was quantified at concentrations ranging from 3.7 to 6.5 log10 cells/100 ml in runoff water collected after pig manure was spread on soil. Among the stream water samples in which cultured Escherichia coli was detected, 23% tested positive for L. sobrius/amylovorus. The results of this study indicate that the quantification of L. sobrius/amylovorus using real-time PCR will be useful for identifying pig fecal contamination in surface waters.

Pig manure may contain pathogenic microorganisms that are harmful to humans and animals (11). These pathogens, which include bacteria, viruses, and protozoans, can survive for several weeks during the storage of manure and in the soil after manure is spread on the land (30). As a consequence, the large amount of manure that is produced and applied on land in many agricultural areas may impact water quality. It contributes to non-point source pollution, which is due partially to runoff from manured soil, especially when manure is spread just before rainfall. It is difficult to determine the origin of diffuse pollution, as it cannot be traced to a specific spot. Fecal indicators (Escherichia coli, fecal coliforms, and enterococci), which are commonly used to quantify fecal pollution, are present in the intestinal tracts of both humans and warm-blooded animals and thus cannot be used to distinguish contamination by pig manure from other sources of pollution. For this reason, alternative microbial indicators have been proposed for the identification of specific pollution sources.

During the past 10 years, a few library-independent methods have been developed for the identification of pig fecal contamination. They are based mostly on the PCR amplification of specific genes or sequences, such as the STII toxin gene from enterotoxigenic E. coli (16), the internal transcribed spacer (ITS) sequence from Bifidobacterium thermacidophilum subsp. porcinum (26), the 16S rRNA gene of Bacteroides-Prevotella (5, 27, 31), and the methyl coenzyme M reductase gene from a methanogenic Archaea member (41). However, some of these methods are only qualitative, like the detection of B. thermacidophilum subsp. porcinum or of the mcrA and STII toxin genes, and do not allow the level of contamination to be quantified. Furthermore, it is noteworthy that the archaeal mcrA gene was not detected in 16% of the pig feces analyzed (41), and that the presence of the STII toxin gene depends on the level of E. coli in the sample, which needs to be greater than 100 cells to avoid false positives (16). Okabe et al. (31) quantified a Bacteroides-Prevotella pig-specific marker (Pig-Bac2) in water samples using real-time PCR. However, this marker lacks specificity, as the Pig-Bac2 marker also was present in human and cow feces at a concentration of 7 and 8 log10 copies per g, respectively (31). Only one pig-specific Bacteroidales 16S rRNA gene marker (Pig-2-Bac), which was developed recently by Mieszkin et al. (27) using real-time PCR, appears to be suitable to quantify pig fecal contamination. However, one limit of targeting the Bacteroidales group could be their strictly anaerobic metabolism, which may influence their persistence in well-oxygenated water. Savichtcheva et al. (36) thus have reported that oxygen has a negative effect on the survival rate of Bacteroides fragilis. We thus consider it important to study biomarkers that are less sensitive to oxygen in order to extend the choice of tools for tracking sources of pollution by manure. Moreover, in the case of the downgrading of bathing or shellfish areas, when health and economic risks are involved, it could be useful to combine multiple markers to identify the source of pollution with certainty.

In the search for potential pig manure markers, we recently analyzed the dominant bacterial groups of 17 raw pig manure samples using 16S rRNA-targeted PCR and the CE-SSCP (capillary electrophoresis-single-strand conformation polymorphism) molecular typing method (26). Among the dominant bacterial groups (Bacteroidales, Bifidobacterium, Eubacterium-Clostridiaceae, and Bacillus-Streptococcus-Lactobacillus), we highlighted the presence of a microaerophilic species, Lactobacillus sobrius, which was isolated from piglet feces previously (19). Lactobacilli are known to establish a stable population in the intestinal tract of piglets soon after birth (28, 39) and to rapidly become a dominant population of their intestinal flora, at least in the first days after weaning (2, 14, 34). Their concentration in pig feces has been estimated at about 3 × 108 bacteria/g (9). Because of their protective effect against diarrhea, some species of Lactobacillus, including L. sobrius, particularly have been studied (20, 35). Konstantinov et al. (21) therefore designed a primer pair that specifically amplifies a fragment of the L. sobrius genome using real-time PCR. Finally, Jakava-Viljanen et al. (13) recently demonstrated very high similarity between the L. sobrius and L. amylovorus type and reference strains and representative porcine isolates based on their 16S rRNA gene sequence analysis. According to these results, L. sobrius and L. amylovorus constitute a single species and consequently are referred to as L. sobrius/amylovorus in this paper.

Given the abundance of L. sobrius/amylovorus in piglet feces (19, 37) and its systematic presence in raw manure (26), we tested this species as a new marker of pig fecal contamination. The aims of our study were (i) to confirm the specificity of L. sobrius/amylovorus to pig feces by analyzing five host groups (human, pig, cattle, poultry, and sheep), manure and by-products of manure treatment, runoff water, and urban wastewaters, and (ii) to estimate the suitability of this marker to identify pig fecal contamination found in surface waters. The concentrations of L. sobrius/amylovorus were estimated by real-time PCR using the primers designed by Konstantinov et al. (21). They were compared to the levels of E. coli, total lactobacilli, and, for river water samples, to the concentrations of the pig-specific Bacteroidales 16S rRNA genetic marker (Pig-2-Bac) developed by Mieszkin et al. (27).

MATERIALS AND METHODS

Bacterial strains and growth conditions.

The strains used in this study were Lactobacillus acetotolerans DSM20749T, L. acidophilus DSM9126T, L. amylolyticus DSM11664T, L. amylovorus DSM20531T, L. crispatus DSM20584T, L. delbrueckii subsp. delbrueckii DSM20074T, L. hamsteri DSM5661T, L. helveticus DSM 20075T, L. johnsonii DSM10533T, L. kalixensis DSM16043T, L. kefiranofaciens subsp. kefiranofaciens DSM5016T, L. kitasatonis DSM16761T, L. reuteri DSM8533T, L. sobrius DSM16698T, and Escherichia coli DSM30083T. Strains were cultured under anaerobic conditions at 37°C on Man Rogosa Sharpe (MRS) agar (Biokar, France), except for L. acetotolerans DSM20749T and L. hamsteri DSM5661T, which were cultured on modified MRS medium (with 0.05% cysteine-hydrochloride and pH adjusted to 5.2) as recommended by DSMZ. E. coli was grown at 37°C on TBX medium (Biokar, France).

Fecal samples.

A total of 132 samples of fresh animal feces (38 cowpats of cattle, 48 feces of pig, 34 droppings of poultry, and 16 feces of sheep) were collected from 64 farms across Brittany. Twenty-seven samples of human feces from healthy people were obtained from two French research institutes (IFREMER [Brest] and INRA [Jouy-en-Josas]). From each well-mixed sample of fresh feces, subsamples of approximately 250 mg (wet weight) were taken and transferred into a microtube and stored at −20°C.

Manure and lagoon water samples.

Manure and lagoon water samples were collected from 18 piggeries located across Brittany as previously described (26). Samples were centrifuged at 16,000 × g to form a pellet of approximately 250 mg (wet weight). The pellets were stored at −20°C.

Water samples.

Six independent samples of field runoff water were collected 40 to 50 min after six rainfall simulations on an experimental agricultural plot previously spread with either cattle or pig manure. A rainfall simulator was placed under a tent to prevent wind and natural rain perturbation. To represent an extreme storm event in spring, simulated rainfall was applied at an intensity of 67 mm·h−1. All samples were poured into 2-liter flasks. Volumes of approximately 200 ml of water were centrifuged at 4,000 × g for 30 min, and pellets were transferred into microtubes and stored at −20°C.

Thirteen samples of treated urban wastewater were collected from locations across Brittany and Pays de la Loire (France). Samples of river surface water (1 liter) were collected from 30 sites located in rural areas in Brittany. For these samples, 100 ml of river water was filtered through a 0.2-μm-pore-size polycarbonate filter. When it was not possible to filter due to clogging, the water samples were centrifuged as described above. Filtrates or pellets were transferred into microtubes and stored at −20°C.

Enumeration of E. coli.

E. coli cells were enumerated in all water samples using 3M E. coli petri film to estimate the level of fecal contamination as described previously (26). The concentration of E. coli was expressed in CFU/ml.

Extraction of genomic DNA of strains.

Genomic DNA was isolated from a 1-ml exponential-phase culture of each strain collected using the Promega Wizard purification kit (Promega). DNA pellets were suspended in 100 μl of rehydration solution (10 mM Tris, 1 mM EDTA). The integrity of genomic DNA was monitored by 1.5% agarose gel electrophoresis in 1× Tris-borate-EDTA and ethidium bromide staining and quantified using a Nanodrop ND1000 microspectrophotometer (Labtech, France).

Extraction of genomic DNA from samples.

DNA was extracted from the pellets, and filters were stored at −20°C using the QIAamp DNA stool kit (Qiagen) in accordance with the manufacturer's instructions, except that the samples were homogenized in buffer ASL and heated at 95°C for 5 min to lyse bacterial cells. The elution volume was 50 μl.

Quantitative PCR.

PCR amplification was performed using a Bio-Rad CFX96 real-time PCR instrument with Bio-Rad CFX Manager software, version 1.1 (Bio-Rad, Hercules, CA).

All target sequences and primer sequences used in this study are presented in Table Table1.1. Primer sequences and PCR programs used for the quantification of lactobacilli and L. sobrius/amylovorus are described in Su et al. (37) and Konstantinov et al. (21), respectively. The reaction mixture consisted of 12.5 μl of IQ SYBR green Supermix (Bio-Rad), a 200 nM concentration of each primer, 2 μl of 1/10 diluted DNA, and 9.5 μl of water to reach a final volume of 25 μl. E. coli was quantified by the real-time PCR protocol described by Khan et al. (15). The reaction mixture consisted of 12.5 μl of IQ SYBR green Supermix (Bio-Rad), a 350 nM concentration of each primer, 2 μl of 1/10 diluted DNA, and 9.5 μl of water to reach a final volume of 25 μl. Bacteroidales marker Pig-2-Bac was quantified using TaqMan chemistry according to the protocol described by Mieszkin et al. (27). The reaction mixture consisted of 12.5 μl of IQ supermix (Bio-Rad), 300 nM concentrations of each primer, a 200 nM concentration of Pig-2-Bac113MGB probe, 2 μl of 1/10 diluted DNA, and 8.5 μl of water to reach a final volume of 25 μl. Each assay was conducted in triplicate in 96-well plates (Bio-Rad).

TABLE 1.
Sequences of the primers and probes used in this study

Standard curves for DNA.

The standard curves for genomic DNA were constructed using serially diluted DNA from L. sobrius DSM16698T and E. coli DSM30083T. In brief, the concentrations of L. sobrius and E. coli were estimated using the standard plate count method to be 3.5 × 108 and 1 × 109 CFU/ml, respectively. Cells were enumerated during the exponential growth phase. Total DNA was extracted from the two reference strains and 10-fold serially diluted in sterile deionized water to yield 7 × 100 to 7 × 105 cells per reaction mixture for L. sobrius and lactobacilli and 2.7 × 101 to 2.7 × 106 cells per reaction mix for E. coli. Results of real-time PCR thus were expressed as cells per ml of water or per g of manure.

For the quantification of the Bacteroides Pig-2-Bac marker, standard curves were generated from serial dilutions of a known concentration of linear plasmid DNA (kindly supplied by S. Mieszkin, IFREMER, Brest, France) linearized with NotI enzyme (Roche Diagnostics) (27).

RESULTS

Primer specificity.

The set of primers developed by Konstantinov et al. (27) was tested on 14 reference strains of Lactobacillus. As expected, only L. sobrius DSM16698T and L. amylovorus DSM20531T, which were reclassified recently as the same species (13), gave a signal, whereas closely related strains such as L. kitasatonis DSM16761T or L. acidophilus DSM9126T did not give a signal (data not shown).

Detection of L. sobrius/amylovorus in feces.

To estimate the relative abundance of L. sobrius/amylovorus in feces of different hosts, we compared the concentrations of this species to those of lactobacilli and E. coli (Table (Table2).2). A total of 163 samples were analyzed, including 48 pig feces, 87 feces samples from other animals, and 28 human feces. L. sobrius/amylovorus was present in 100% of the pig feces samples at a mean concentration of 7.2 log10 cells/g. The primers did not amplify products in human, cattle, and sheep feces. They amplified 5 of the 34 poultry droppings, which represent closely the composition of the fecal matter excreted. In all the samples of pig feces, the concentrations of L. sobrius/amylovorus always were within the same order of magnitude as those observed for lactobacilli and E. coli, whereas this species represented less than 0.1‰ of lactobacilli and of E. coli in the five poultry feces.

TABLE 2.
Concentrations of L. sobrius/amylovorus, lactobacilli, and E. coli (log10 cells/gram) in feces of different origins

Influence of manure treatment on the L. sobrius/amylovorus population.

We then tested the persistence of L. sobrius/amylovorus during manure management and our ability to detect it in pig farm effluents intended for spreading or irrigation. Target bacterial genomes and cultured E. coli were quantified in raw manure and in manure stored in a tank for 6 to 9 months after a biological treatment to remove nitrogen. Lagoon water samples that consisted of supernatant from treated manure stored in tanks also were collected and analyzed (Table (Table33).

TABLE 3.
Concentrations of L. sobrius/amylovorus, lactobacilli, and E. coli (log10 cells/g or ml) in pig farm effluent

L. sobrius/amylovorus was consistently detected in raw manure at a mean concentration of 5.9 log10/g. It is interesting that despite the aerobic biological treatment and the ensuing anaerobic storage for several months, the decrease in the number of L. sobrius/amylovorus cells did not exceed 1 log10. These results underline the persistence of the biomarker throughout the aerobic biological treatment. In lagoon water samples that were slightly contaminated, as indicated by the very low level of E. coli (close to the detection limit of the cultural method), L. sobrius/amylovorus was quantified in 50% of the samples at a level of 1.7 log10 cells/ml. In the two lagoon water samples where E. coli was not found, we did not detect L. sobrius/amylovorus.

Quantification of L. sobrius/amylovorus in urban effluent and water samples.

Real-time PCR assays using L. sobrius/amylovorus, lactobacilli, and E. coli primer sets and the enumeration of cultured E. coli were performed on water samples. Nineteen water samples were impacted by human and animal fecal waste. They included six runoff water samples from a soil surface after the spreading of cattle slurry or pig manure and 13 treated effluent samples from urban wastewater treatment plants. Thirty surface water sources were sampled in rural areas with no prior knowledge of the level or the origin of pollution, except for one sample that was collected in a stream close to a field where manure had been spread 3 days prior to a rainy period. The quantification and the prevalence of L. sobrius/amylovorus were also compared to those of the Bacteroidales Pig-2-Bac marker developed by Mieszkin et al. (27) (Tables (Tables44 and and55).

TABLE 4.
Concentrations of L. sobrius/amylovorus, lactobacilli, and E. coli (log10 bacteria/ml) in water samples and concentrations of host-specific Pig-2-Bac marker in surface water (log10 copies/ml)
TABLE 5.
Prevalence of E. coli and the pig markers and among the 30 surface water samples

Contrary to lactobacilli that were consistently found in waters regardless of their origin and the level of E. coli, L. sobrius/amylovorus was not detected in any effluents of urban wastewater treatment plants. L. sobrius/amylovorus was quantified at concentrations ranging from 3.7 to 6.5 log10 cells/100 ml in runoff water samples collected after the spreading of pig manure, whereas it was not detected in runoff water contaminated by cattle slurry. Furthermore, L. sobrius/amylovorus was quantified in three samples of surface water in which the Pig-2-Bac marker also was present (Table (Table55).

DISCUSSION

A high concentration of piggeries in a restricted area, as is the case in Brittany, leads to the fecal contamination of surface waters after manure is spread on the soil. The transfer of pathogenic microorganisms from manure can pose a risk to human health, especially in coastal areas where shellfish farming and nautical activities are widespread. It is thus important to possess tools to accurately identify and quantify pig fecal contamination in sensitive areas to be able to act directly on the source of pollution. However, few quantitative methods are available to differentiate pig and other animal or human sources of pollution. Only one specific biomarker, Pig-2-Bac, which was developed by Mieszkin et al. (27) and does not require culture, can be used to quantify pig fecal contaminations in environmental water samples. The present study tested the relevance of a new pig biomarker, L. sobrius/amylovorus, using the set of primers designed by Konstantinov et al. (21). This biomarker targets two species, L. amylovorus and L. sobrius, which were described by Nakamura (29) and by Konstantinov et al. (19), respectively, and now are considered a single species due their high level of genomic similarity (13). To our knowledge, this is the first report that targets lactobacilli as a biomarker of fecal contamination.

The concentrations of lactobacilli estimated by real-time PCR in the 48 pig feces analyzed in this study ranged between 5.8 and 8.4 log10 cells per g of feces and were slightly lower (ca. 0.7 log10 less) than those of E. coli, as previously observed by Furet et al. (9). These data also are comparable to the concentrations observed in the intestinal tract of piglets that range between 5.6 and 9.3 log10 cells per g (18, 34, 37, 40). The variability of these concentrations may be explained by the differences in age and diet of the animals (17, 18).

We also observed variability of the concentrations of lactobacilli and E. coli in feces of humans and other animals. However, mean concentrations of E. coli were in the same order of magnitude as those reported for human feces by Furet et al. (9) and Firmesse et al. (8) with SYBR green PCR. They also were similar to the concentrations of E. coli in cattle feces reported by Furet et al. (9) and of Enterobacteriaceae in ileum and cecum of poultry observed by Olsen et al. (32) using the FISH method.

Our study showed that L. sobrius/amylovorus, which was detected consistently at concentrations ranging between 4.8 and 8.3 log10 cells per g in pig feces, represents at least 80% of the lactobacilli. These data are in agreement with those of Su et al. (37) and Pieper et al. (34), who observed between 5.3 and 8.6 log10 L. sobrius in the ileum and small intestine of weaning piglets. Our data thus highlight the fact that L. sobrius/amylovorus, which was previously quantified in piglets (18, 34, 37, 40), also is abundant in the feces of adult pigs. While the presence of L. sobrius has been described only in the intestinal tract of piglets (19, 34, 37), the natural habitat of L. amylovorus is less known. L. amylovorus was initially described in maize silage (29). However, recently Brusetti et al. (3), who used length heterogeneity-PCR to study the succession of the lactic bacteria during maize ensiling, did not report the presence of L. amylovorus.

In our study, L. sobrius/amylovorus was not found in the cattle, sheep, and human samples. These results are consistent with the fact that neither L. amylovorus nor L. sobrius has been found by molecular inventory methods in intestinal or fecal microflora of humans (6, 12, 38). More importantly, while we always found lactobacilli and E. coli in the urban wastewater effluents, the latter at concentrations similar to those reported by Wéry et al. (42), we never detected L. sobrius/amylovorus in such samples.

The presence of L. sobrius/amylovorus at low concentrations in 5 of the 34 poultry feces is in accordance with the study of Cauwerts et al. (4), who isolated L. amylovorus by culture in the intestine of chicken. However, it is important to underline that when detected in poultry, the concentration of L. sobrius/amylovorus was 4 log10 below the level observed in pig feces. Given its low prevalence and concentration, it is not surprising that L. sobrius/amylovorus has not been identified using molecular tools in the intestinal tracts of chicken and turkey, the dominant lactobacilli being L. delbrueckii and L. acidophilus or L. aviarius and L. salivarius for chicken (10, 24, 25) and L. aviarius for turkey (23). Because the natural habitat of L. sobrius/amylovorus is not well documented, it was important to test manure and surface water samples for the presence of this biomarker to show that it still was present in pig manure but not ubiquitous in the environment, and that we were able to detect it in contaminated samples.

Like lactobacilli and E. coli, L. sobrius/amylovorus persisted in raw manure and was only slightly impacted by the biological treatment and storage of manure. These data are in agreement with the studies of Leung and Topp (22) and Peu et al. (33), who observed very few changes in the bacterial communities during the anaerobic storage of manure. It is important to note that L. sobrius/amylovorus was quantified in lagoon waters stored for 9 to 12 months only in the samples in which cultured E. coli was detected. These results suggest that, like E. coli, L. sobrius/amylovorus has the ability to persist in water for extended periods of time. Furthermore, after pig manure was spread on a field before a rainfall event, the transfer of L. sobrius/amylovorus was similar to that of E. coli measured by real-time PCR, as both bacteria were found in runoff water at similar concentrations.

To validate the specificity of our marker, we quantified the Pig-2-Bac marker described by Mieszkin et al. (27) in samples of natural river water using TaqMan PCR technology and compared its presence to that of lactobacilli and E. coli. In samples of surface water collected in rural areas, E. coli and lactobacilli measured by real-time PCR always were quantified at similar concentrations ranging between 5.5 and 6.9 log10 cells/100 ml, whereas cultured E. coli was detected in 13 of the 30 surface water samples. The difference in concentration between E. coli measured by molecular and cultural methods could be explained by the fact that this bacteria can enter a viable but nonculturable state (1) or by the amplification of DNA from dead cells. The systematic detection of lactobacilli in surface waters could be due to the existence of viable but nonculturable cells or to the presence of ubiquitous species, given the considerable diversity of this group of bacteria in ecological habitats (7). Interestingly, L. sobrius/amylovorus and Pig-2-Bac were detected simultaneously in three surface water samples, one of them being collected after a rainfall event less than 10 m from a field where pig manure had been spread. Our data not only highlight the fact that L. sobrius/amylovorus cells are not ubiquitous in the environment but also confirm the specificity of our marker and its ability to be transferred via runoff into surface water. Furthermore, our results confirm that the contamination of rivers by pig manure appears to be relatively frequent in Brittany, as L. sobrius/amylovorus was present in 23% of the rivers in which cultured E. coli was detected.

This study showed the potential for using L. sobrius/amylovorus to identify the contamination of water by pig manure. One concern might be a high concentration of poultry farms producing L. sobrius/amylovorus in the environment. However, the prevalence and the concentration of L. sobrius/amylovorus in poultry droppings were very low. In addition, poultry farms more often produce manure diluted with litter material and largely dried prior to land application, thus substantially reducing the concentration of L. sobrius/amylovorus if present. Even if such dilution did not apply, we have observed in the runoff experiment that, regardless of the bacteria, the concentration in the runoff water was ca. 1,000th of that in pig manure. This result implies that the concentration of L. sobrius/amylovorus originating from poultry manure is less than 1 cell/ml in the subsequent runoff water.

The quantification of this biomarker by real-time PCR makes it a promising tool for microbial source tracking. Its usefulness is enhanced by its systematic presence in manure, whether treated or not, collected in various farms located in different areas of Brittany. The persistence of L. sobrius/amylovorus, its transfer via runoff, and its quantification in river waters mean that this bacterium is relevant for the detection of contamination by pig manure. However, since it is not possible to exclude the relatively minor presence of L. sobrius/amylovorus in the feces of wild animals, this new marker should be used with other pig markers such as Pig-2-Bac to improve the detection accuracy of the method. Furthermore, future studies will concentrate on the behavior of L. sobrius/amylovorus under different environmental conditions.

Acknowledgments

We thank M. Gourmelon from IFREMER (Brest) and M. Leclerc from INRA (Jouy-en-Josas), who kindly provided cattle and human fecal samples for the study.

This research was supported financially by the Agence Française de Sécurité Sanitaire de l'Environnement et du Travail (AFSSET) and by the French Environment and Energy Management Agency (ADEME). R. Marti is the recipient of a Cemagref-Ademe fellowship.

Footnotes

[down-pointing small open triangle]Published ahead of print on 28 December 2009.

REFERENCES

1. Arana, I., M. Orruno, D. Perez-Pascual, C. Seco, A. Muela, and I. Barcina. 2007. Inability of Escherichia coli to resuscitate from the viable but nonculturable state. FEMS Microbiol. Ecol. 62:1-11. [PubMed]
2. Bateup, J. M., S. Dobbinson, M. A. McConnell, K. Munro, and G. W. Tannock. 1998. Molecular analysis of the composition of Lactobacillus populations inhabiting the stomach and caecum of pigs. Microb. Ecol. Health Dis. 10:95-102.
3. Brusetti, L., S. Borin, D. Mora, A. Rizzi, N. Raddadi, C. Sorlini, and D. Daffonchio. 2006. Usefulness of length heterogeneity-PCR for monitoring lactic acid bacteria succession during maize ensiling. FEMS Microbiol. Ecol. 56:154-164. [PubMed]
4. Cauwerts, K., F. Pasmans, L. A. Devriese, A. Martel, F. Haesebrouck, and A. Decostere. 2006. Cloacal Lactobacillus isolates from broilers show high prevalence of resistance towards macrolide and lincosamide antibiotics. Avian Pathol. 35:160-164. [PubMed]
5. Dick, L. K., A. E. Bernhard, T. J. Brodeur, J. W. Santo Domingo, J. M. Simpson, S. P. Walters, and K. G. Field. 2005. Host distribution of uncultivated fecal Bacteroidales bacteria reveal genetic markers for fecal source identification. Appl. Environ. Microbiol. 71:3184-3191. [PMC free article] [PubMed]
6. Eckburg, P. B., E. M. Bok, C. N. Bernstein, E. Purdom, L. Dethlefsen, M. Sargent, S. R. Gill, K. E. Nelson, and D. A. Relman. 2005. Diversity of the human intestinal microbial flora. Science 308:1635-1638. [PMC free article] [PubMed]
7. Felis, G. E., and F. Dellaglio. 2007. Taxonomy of lactobacilli and bifidobacteria. Curr. Issues Intest. Microbiol. 8:44-61. [PubMed]
8. Firmesse, O., S. Rabot, L. G. Bermudez-Humaran, G. Corthier, and J. P. Furet. 2007. Consumption of Camembert cheese stimulates commensal enterococci in healthy human intestinal microbiota. FEMS Microbiol. Lett. 276:189-192. [PubMed]
9. Furet, J. P., O. Firmesse, M. Gourmelon, C. Bridonneau, J. Tap, S. Mondot, J. Dore, and G. Corthier. 2009. Comparative assessment of human and farm animal faecal microbiota using real-time quantitative PCR. FEMS Microbiol. Ecol. 68:351-362. [PubMed]
10. Gong, J., W. Si, R. J. Forster, R. Huang, H. Yu, Y. Yin, C. Yang, and Y. Han. 2007. 16S rRNA gene-based analysis of mucosa-associated bacterial community and phylogeny in the chicken gastrointestinal tracts: from crops to ceca. FEMS Microbiol. Ecol. 59:147-157. [PubMed]
11. Guan, T. Y., and R. A. Holley. 2003. Pathogen survival in swine manure environments and transmission of human enteric illness—a review. J. Environ. Qual. 32:383-392. [PubMed]
12. Heilig, H. G., E. G. Zoetendal, E. E. Vaughan, P. Marteau, A. D. Akkermans, and W. M. de Vos. 2002. Molecular diversity of Lactobacillus spp. and other lactic acid bacteria in the human intestine as determined by specific amplification of 16S ribosomal DNA. Appl. Environ. Microbiol. 68:114-123. [PMC free article] [PubMed]
13. Jakava-Viljanen, M., A. Murros, A. Palva, and K. J. Bjorkroth. 2008. Lactobacillus sobrius Konstantinov et al. 2006 is a later synonym of Lactobacillus amylovorus Nakamura 1981. Int. J. Syst. Evol. Microbiol. 58:910-913. [PubMed]
14. Janczyk, P., R. Pieper, H. Smidt, and W. B. Souffrant. 2007. Changes in the diversity of pig ileal lactobacilli around weaning determined by means of 16S rRNA gene amplification and denaturing gradient gel electrophoresis. FEMS Microbiol. Ecol. 61:132-140. [PubMed]
15. Khan, I. U., V. Gannon, R. Kent, W. Koning, D. R. Lapen, J. Miller, N. Neumann, R. Phillips, W. Robertson, E. Topp, E. van Bochove, and T. A. Edge. 2007. Development of a rapid quantitative PCR assay for direct detection and quantification of culturable and non-culturable Escherichia coli from agriculture watersheds. J. Microbiol. Methods 69:480-488. [PubMed]
16. Khatib, L. A., Y. L. Tsai, and B. H. Olson. 2003. A biomarker for the identification of swine fecal pollution in water, using the STII toxin gene from enterotoxigenic Escherichia coli. Appl. Micriobiol. Biotechnol. 63:231-238. [PubMed]
17. Konstantinov, S. R., A. Awati, H. Smidt, B. A. Williams, A. D. Akkermans, and W. M. de Vos. 2004. Specific response of a novel and abundant Lactobacillus amylovorus-like phylotype to dietary prebiotics in the guts of weaning piglets. Appl. Environ. Microbiol. 70:3821-3830. [PMC free article] [PubMed]
18. Konstantinov, S. R., A. Awati, A. G. Williams, B. G. Miller, P. Jones, C. R. Stokes, A. D. Akkermans, H. Smidt, and W. M. de Vos. 2006. Post-natal development of the porcine microbiota composition and activities. Environ. Microbiol. 8:1191-1199. [PubMed]
19. Konstantinov, S. R., E. Poznanski, S. Fuentes, A. D. Akkermans, H. Smidt, and W. M. de Vos. 2006. Lactobacillus sobrius sp. nov., abundant in the intestine of weaning piglets. Int. J. Syst. Evol. Microbiol. 56:29-32. [PubMed]
20. Konstantinov, S. R., H. Smidt, A. D. Akkermans, L. Casini, P. Trevisi, M. Mazzoni, S. De Filippi, P. Bosi, and W. M. de Vos. 2008. Feeding of Lactobacillus sobrius reduces Escherichia coli F4 levels in the gut and promotes growth of infected piglets. FEMS Microbiol. Ecol. 66:599-607. [PubMed]
21. Konstantinov, S. R., H. Smidt, and W. M. de Vos. 2005. Representational difference analysis and real-time PCR for strain-specific quantification of Lactobacillus sobrius sp. nov. Appl. Environ. Microbiol. 71:7578-7581. [PMC free article] [PubMed]
22. Leung, K., and E. Topp. 2001. Bacterial community dynamics in liquid swine manure during storage: molecular analysis using DGGE/PCR of 16S rDNA. FEMS Microbiol. Ecol. 38:169-177.
23. Lu, J., and J. S. Domingo. 2008. Turkey fecal microbial community structure and functional gene diversity revealed by 16S rRNA gene and metagenomic sequences. J. Microbiol. 46:469-477. [PubMed]
24. Lu, J., U. Idris, B. Harmon, C. Hofacre, J. J. Maurer, and M. D. Lee. 2003. Diversity and succession of the intestinal bacterial community of the maturing broiler chicken. Appl. Environ. Microbiol. 69:6816-6824. [PMC free article] [PubMed]
25. Lu, J., J. Santo Domingo, and O. C. Shanks. 2007. Identification of chicken-specific fecal microbial sequences using a metagenomic approach. Water Res. 41:3561-3574. [PubMed]
26. Marti, R., P. Dabert, and A. M. Pourcher. 2009. Selection of a pig manure contamination marker based on the influence of biological treatment on the dominant faecal microbial groups. Appl. Environ. Microbiol. 75:4967-4974. [PMC free article] [PubMed]
27. Mieszkin, S., J. P. Furet, G. Corthier, and M. Gourmelon. 2009. Estimation of pig fecal contamination in a river catchment by real-time PCR using two pig-specific Bacteroidales 16S rRNA genetic markers. Appl. Environ. Microbiol. 75:3045-3054. [PMC free article] [PubMed]
28. Naito, S., H. Hayashidani, K. Kaneko, M. Ogawa, and Y. Benno. 1995. Development of intestinal lactobacilli in normal piglets. J. Appl. Bacteriol. 79:230-236. [PubMed]
29. Nakamura, L. K. 1981. Lactobacillus amylovorus a new starch-hydrolyzing species from cattle waste-corn fermentations. Int. J. Syst. Bacteriol. 31:56-63.
30. Nicholson, F. A., S. J. Groves, and B. J. Chambers. 2005. Pathogen survival during livestock manure storage and following land application. Bioresour. Technol. 96:135-143. [PubMed]
31. Okabe, S., N. Okayama, O. Savichtcheva, and T. Ito. 2007. Quantification of host-specific Bacteroides-Prevotella 16S rRNA genetic markers for assessment of fecal pollution in freshwater. Appl. Micriobiol. Biotechnol. 74:890-901. [PubMed]
32. Olsen, K. N., M. Henriksen, M. Bisgaard, O. L. Nielsen, and H. Christensen. 2008. Investigation of chicken intestinal bacterial communities by 16S rRNA targeted fluorescence in situ hybridization. Antonie van Leeuwenhoek 94:423-437. [PubMed]
33. Peu, P., H. Brugere, A. M. Pourcher, M. Kerouredan, J. J. Godon, J. P. Delgenes, and P. Dabert. 2006. Dynamics of a pig slurry microbial community during anaerobic storage and management. Appl. Environ. Microbiol. 72:3578-3585. [PMC free article] [PubMed]
34. Pieper, R., P. Janczyk, A. Zeyner, H. Smidt, V. Guiard, and W. B. Souffrant. 2008. Ecophysiology of the developing total bacterial and Lactobacillus communities in the terminal small intestine of weaning piglets. Microb. Ecol. 56:474-483. [PubMed]
35. Roselli, M., A. Finamore, M. S. Britti, S. R. Konstantinov, H. Smidt, W. M. de Vos, and E. Mengheri. 2007. The novel porcine Lactobacillus sobrius strain protects intestinal cells from enterotoxigenic Escherichia coli K88 infection and prevents membrane barrier damage. J. Nutr. 137:2709-2716. [PubMed]
36. Savichtcheva, O., N. Okayama, T. Ito, and S. Okabe. 2005. Application of a direct fluorescence-based live/dead staining combined with fluorescence in situ hybridization for assessment of survival rate of Bacteroides spp. in drinking water. Biotechnol. Bioeng. 92:356-363. [PubMed]
37. Su, Y., W. Yao, O. N. Perez-Gutierrez, H. Smidt, and W. Y. Zhu. 2008. Changes in abundance of Lactobacillus spp. and Streptococcus suis in the stomach, jejunum and ileum of piglets after weaning. FEMS Microbiol. Ecol. 66:546-555. [PubMed]
38. Suau, A., R. Boonet, M. Sutren, J. J. Godon, G. Gibson, M. Collins, and J. Doré. 1999. Direct analysis of gene encoding 16S rRNA from complex communities reveals many novel molecular species within the human gut. Appl. Environ. Microbiol. 65:4799-4807. [PMC free article] [PubMed]
39. Tannock, G. W., R. Fuller, and K. Pedersen. 1990. Lactobacillus succession in the piglet digestive tract demonstrated by plasmid profiling. Appl. Environ. Microbiol. 56:1310-1316. [PMC free article] [PubMed]
40. Tzortzis, G., A. K. Goulas, J. M. Gee, and G. R. Gibson. 2005. A novel galactooligosaccharide mixture increases the bifidobacterial population numbers in a continuous in vitro fermentation system and in the proximal colonic contents of pigs in vivo. J. Nutr. 135:1726-1731. [PubMed]
41. Ufnar, J. A., D. F. Ufnar, S. Y. Wang, and R. D. Ellender. 2007. Development of a swine-specific fecal pollution marker based on host differences in methanogen mcrA genes. Appl. Environ. Microbiol. 73:5209-5217. [PMC free article] [PubMed]
42. Wéry, N., C. Lhoutellier, F. Ducray, J. P. Delgenes, and J. J. Godon. 2008. Behaviour of pathogenic and indicator bacteria during urban wastewater treatment and sludge composting, as revealed by quantitative PCR. Water Res. 42:53-62. [PubMed]

Articles from Applied and Environmental Microbiology are provided here courtesy of American Society for Microbiology (ASM)