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1 Laboratory of Microbiology, Department of Biochemistry, Physiology and Microbiology, Ghent University, K.L. Ledeganckstraat 35, B-9000 Gent, Belgium
2 Nestlé Research Centre, Route du Jorat 57, Vers-Chez-Les-Blanc, 1000 Lausanne 26, Switzerland
3 BCCM/LMG Bacteria Collection, Department of Biochemistry, Physiology and Microbiology, Ghent University, K.L. Ledeganckstraat 35, B-9000 Gent, Belgium
Correspondence
Liesbeth Masco
liesbeth.masco{at}ugent.be
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has to be considered as a junior synonym of B. animalis (Mitsuoka 1969) Scardovi and Trovatelli 1974, our data indicate that the latter species should be split into two new subspecies, i.e. Bifidobacterium animalis subsp. animalis subsp. nov. (type strain R101-8T=LMG 10508T=ATCC 25527T=DSM 20104T=JCM 1190T) and Bifidobacterium animalis subsp. lactis subsp. nov. (type strain UR1T=LMG 18314T=DSM 10140T=JCM 10602T).
Published online ahead of print on 16 January 2004 as DOI 10.1099/ijs.0.03011-0.
The GenBank/EMBL/DDBJ accession numbers for the atpD and groEL gene sequences determined in this study are given in Fig. 3
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Tables of additional descriptive data of the strains used in this study, and DNA base compositions and levels of DNA relatedness of B. lactis and B. animalis are available as supplementary material in IJSEM Online.
Present address: Department of Microbiology, National University of Ireland, Cork, Ireland. 
Present address: Nutrition & Health, Cognis, Düsseldorf, Germany.
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, and several studies have investigated its affiliation with the closely related but earlier described Bifidobacterium animalis (Scardovi & Trovatelli, 1974). Based on phenotypic characteristics, 16S rDNA sequence analysis and DNA–DNA hybridization, Cai et al. (2000) proposed that B. lactis should be considered as a junior synonym of B. animalis. However, new genotypic evidence, recently reported by Ventura & Zink (2002, 2003) and Zhu et al. (2003), suggested that B. lactis and B. animalis should still be considered to be two separate taxonomic entities, not at the species level but at the subspecies level.
Compared to B. animalis, strains of B. lactis exhibit elevated oxygen tolerance, which is a remarkable trait within the bifidobacteria that allows them to reach high numbers in commercial products under non-anaerobic conditions. Because of this, B. lactis strains are frequently applied in probiotic dairy products, food supplements and pharmaceutical preparations (Prasad et al., 1998). To guarantee the quality and the correct labelling of such products, it is thus very important that the taxonomic position of this industrially applied micro-organism is clear.
The aim of this polyphasic study was to investigate the taxonomic relationship between B. animalis and B. lactis on the basis of DNA–DNA hybridization, mol% G+C determination, sugar fermentation patterns, the ability to grow in milk, protein profiling, BOX-PCR and Fluorescent Amplified Fragment Length Polymorphism (FAFLP) fingerprinting and atpD and groEL gene sequence analysis.
The 16 Bifidobacterium strains used in this study, namely B. animalis LMG 10508T, LMG 18900 and NCC 273, and B. lactis LMG 18314T, LMG 11615, LMG 18906, LMG 11580, NCC 239, NCC 282, NCC 311, NCC 330, NCC 362 (=Bb12, Chr. Hansen, Denmark), NCC 363, NCC 383, NCC 387 and NCC 402 were obtained from the BCCM/LMG Bacteria Collection, Ghent University, Belgium (http://www.belspo.be/bccm/lmg.htm) or from the Nestlé Culture Collection (NCC), Nestlé Research Centre, Lausanne, Switzerland (additional descriptive data are available as supplementary Table 1 in IJSEM Online). All strains were grown overnight at 37 °C under anaerobic conditions (80 % N2, 10 % H2, 10 % CO2) on modified Columbia agar comprising 23 g special peptone (Oxoid), 1 g soluble starch, 5 g NaCl, 0·3 g cysteine-HCl.H2O (Sigma), 5 g glucose and 15 g agar dissolved in 1 l distilled water (BCCM/LMG, Medium 144).
Strains were phenotypically characterized using the AN MicroPlate system (Biolog) according to the instructions of the manufacturer. Cells were subcultured twice on modified Columbia agar, after which the MicroPlates were incubated under a hydrogen-free anaerobic atmosphere (100 % CO2) for 24 h. The MicroPlates were spectrophotometrically read using the Biolog Micro Station-reader. The ability to ferment starch was tested separately by inoculation of the strains on modified Columbia agar depleted of the usual carbon sources and subsequently supplemented with an equal amount (w/v) of soluble starch. After incubation under anaerobic conditions at 37 °C for 72 h, Lugol's solution (0·5 % I2+1 % KI in distilled water) was poured on the growth zone and visually checked for a hydrolysis halo.
High-molecular-mass DNA for DNA–DNA hybridizations and mol% G+C determination was prepared using a combination of the protocols of Marmur (1961) and Pitcher et al. (1989), as described by Goris et al. (1998). DNA base compositions were determined by the method of Mesbah et al. (1989). DNA was enzymically digested into deoxyribonucleosides and separated by HPLC using a Waters Symmetry Shield C8 column thermostatted at 37 °C. The solvent used was 0·02 M NH4H2PO4, pH 4·0, with 1·5 % acetonitrile. Unmethylated 
phage DNA (Sigma) was used as the calibration reference. DNA–DNA hybridizations were performed with biotin-labelled probes in microplate wells (Ezaki et al., 1989), using a HTS7000 Bio Assay Reader (Perkin Elmer) for the fluorescence measurements. The hybridization temperature was 45 °C in the presence of 50 % formamide. Reciprocal experiments were performed for every pair of strains.
SDS-PAGE analysis of whole-cell proteins, using standardized conditions for comparison with the laboratory-based protein-pattern database, was performed according to the methods of Pot et al. (1994).
Micro-scale DNA extraction was based on the method of Pitcher et al. (1989) with slight modifications as described previously (Masco et al., 2003). Micro-scale DNA extracts were used for BOX-PCR and FAFLP fingerprinting. Repetitive DNA element (rep-) PCR fingerprinting using the BOXA1R primer was carried out as described previously (Masco et al., 2003). FAFLP template preparation was carried out essentially as described by Thompson et al. (2001) with slight modifications. High-molecular-mass DNA was digested with TaqI (Westburg) and EcoRI (Amersham Pharmacia Biotech). For the pre-selective PCR the E00 primer (5'-GACTGCGTACCAATTC-3', 1 µM) and T00 primer (5'-CGATGAGTCCTGACCGA-3', 5 µM) (Sigma Genosys) were used. The initial denaturation step was performed at 94 °C. In the selective PCR, the E01-6FAM primer (5'-6FAM-GACTGCGTACCAATTCA-3', 1 µM) and T01 primer (5'-CGATGAGTCCTGACCGAA-3', 5 µM) (Sigma Genosys) were used. The selective PCR products were separated on a denaturing polyacrylamide gel (10·6 %, v/v, acrylamide; 36 %, w/v, urea; 1 %, w/v, resin; and 10 %, v/v, 1x TBE in HPLC water) in 1x TBE buffer. Numerical analysis was performed with BioNumerics V2.5 software (Applied Maths).
For sequencing of the atpD and groEL genes, DNA was prepared as described previously (Ventura et al., 2001). A 1133 bp fragment of atpD and a 1158 bp fragment of groEL were amplified using oligonucleotide primers atp-1 (5'-CACCCTCGAGGTCGAAC-3', position 180 of Bifidobacterium longum NCC 2705) and atp-2 (5'-CTGCATCTTGTGCCACTTC-3', position 1313 of B. longum NCC 2705), and gro-1 (5'-GACCATCACCAACGATG-3', position 138 of B. longum NCC 2705) and gro-2 (5'-GCTCCGGCTTGTTGGC-3', position 1296 of B. longum NCC 2705), respectively. Each PCR mixture (50 µl) contained 20 mM Tris/HCl, 50 mM KCl, 200 µM each dNTP, 50 pmol each primer, 1·5 mM MgCl2 and 1 U Taq DNA polymerase (Gibco-BRL). The PCR cycling profile consisted of an initial denaturation step of 3 min at 95 °C, followed by amplification for 30 cycles as follows: denaturation (30 s at 95 °C), annealing (30 s at 50 °C) and extension (2 min at 72 °C), and completed with an elongation phase (10 min at 72 °C). The resulting amplicons were separated on a 1 % agarose gel followed by ethidium bromide staining. PCR fragments were purified using the PCR purification kit (Qiagen) and were subsequently cloned in the pGEM-T Easy plasmid vector (Promega) following supplier's instructions. Nucleotide sequencing of both strands from cloned DNA was performed using the fluorescent-labelled primer cycle-sequencing kit (Amersham Buchler) following supplier's instructions. The primers used were atp-1, atp-2 and gro-1, gro-2 labelled with IRD800 (MWG Biotech). Sequence alignment was done using the MULTIALIGN program and the CLUSTAL W program. Dendrograms from gene sequences were drawn using the CLUSTAL X program. All atpD and groEL gene sequences reported in this study have been deposited at GenBank and their accession numbers are indicated in Fig. 3.
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According to the DSMZ Bacterial Nomenclature Up-to-date website (http://www.dsmz.de/bactnom/bactname.htm), B. lactis is considered to be a heterotypic synonym of B. animalis based on the proposal of Cai et al. (2000). In spite of this proposal, both names are still regularly used. In the period from January 2001 to August 2003, following the proposal of Cai et al. (2000) to unify B. lactis and B. animalis, the species name B. lactis was cited in at least 37 papers. A recent genotypic study of Ventura & Zink (2002) supported this unification, but also concluded from their Enterobacterial Repetitive Intergenic Consensus (ERIC)-PCR fingerprinting and 16S–23S internally transcribed spacer analysis that strains formerly classified as B. lactis should be allocated in a subspecies of B. animalis.
The current study was initiated to collect more polyphasic evidence in support of the subdivision of B. animalis at the subspecies level. DNA G+C content ranged from 60·3 to 61·4 mol%, with means of 61·3 and 61·0 mol% for representatives of B. animalis and B. lactis, respectively (additional data are available as supplementary Table 2 in IJSEM Online). DNA–DNA hybridizations were performed using seven strains of which some were also included in the study of Cai et al. (2000). Consistent with their findings, all DNA–DNA reassociation values were above 70 %, ranging from 76 to 100 %, and the type strains of B. lactis and B. animalis displayed at least 90 % DNA relatedness. This is in contrast to the findings of Meile et al. (1997) who found only 27 % DNA homology between the type strain of B. lactis and B. animalis using a rather unusual technique based on hybridization of uniformly labelled EcoRI-restricted chromosomal DNA of the B. lactis type strain followed by Southern hybridization with the same amounts of EcoRI-restricted DNA of other Bifidobacterium strains. Based on the narrow G+C content range and the high DNA reassociation values, our data reinforce the proposal of Cai et al. (2000) to join B. lactis and B. animalis in one single species for which the name of the oldest description, i.e. B. animalis, should be maintained according to Rule 42 of the Bacteriological Code (1990 Revision) (Lapage et al., 1992).
Following a polyphasic approach, all B. animalis and B. lactis strains were subjected to a number of techniques that have the potential to unravel relationships at the subspecific level which included protein and DNA (BOX-PCR and FAFLP) fingerprinting as well as atpD and groEL gene sequence typing. Furthermore, the ability of some strains to grow in milk was determined. As further discussed below, the overall result of this approach showed that each of these methods allowed the unambiguous separation of B. animalis from B. lactis. In case of SDS-PAGE protein profiling, BOX-PCR and FAFLP, the resulting B. lactis and B. animalis clusters exhibited similarity levels that were comparable to those between clusters of other Bifidobacterium species (data not shown). The results of the numerical analysis of the SDS-PAGE protein patterns are shown in Fig. 1. After numerical comparison of the digitized protein electrophoretic fingerprints, two well delineated clusters were observed which corresponded to strains previously assigned to B. animalis and B. lactis, respectively. Given the fact that protein profiling displays a lower taxonomic resolution compared to genotypic techniques such as rep-PCR and FAFLP, these findings indicate that both species are distinct from each other on a phenotypic basis. As shown previously by BOX-PCR fingerprinting (Masco et al., 2003), reference strains of B. lactis and B. animalis group in two separate clusters indicating their pronounced genotypic heterogeneity. In the present study, BOX-PCR was performed on additional strains from the NCC, which demonstrated the robustness of these genotypic subgroups (data not shown). Recently, Ventura & Zink (2002) reported that rep-PCR targeting the ERIC element also allowed differentiation between type and reference strains of B. animalis and B. lactis, respectively. FAFLP exhibits a slightly higher resolution than BOX-PCR fingerprinting and is considered, along with PFGE, as the most discriminating genotypic technique. Clustering of the FAFLP banding patterns of 14 strains studied resulted in two clusters representing B. animalis and B. lactis at a cut-off level of 59 % (Fig. 2).
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). In these trees, Bifidobacterium strains were grouped into two clusters. Cluster I contained only the type strain of B. animalis and the reference strain ATCC 27672, whereas cluster II contained six representatives of B. lactis including its type strain. Twenty-eight nucleotide substitutions were observed between the atpD gene sequences of B. lactis DSM 10140T and B. animalis ATCC 25527T. Likewise, 31 synonymous nucleotide substitutions were noticed between the groEL gene sequences of the two type strains. The phylogenetic distances calculated from the nucleotide substitution ratios at synonymous positions in the atpD and groEL genes were examined for all possible combinations of these Bifidobacterium genes. A significant correlation between the phylogenetic distances in the atpD genes and those in the groEL genes was observed. This result was not unexpected, because it has been demonstrated that a synonymous substitution rate is constant for many chromosomal genes in many organisms, and can thus serve as a molecular clock of their evolution (Lawrence et al., 1991). It is noteworthy that a clear separation of B. animalis ATCC 25527T and B. lactis DSM 10140T was not possible based on 16S rDNA sequence analysis since their sequences displayed at least 98·8 % homology (Cai et al., 2000).
Twelve B. lactis and B. animalis strains used in this work were tested for their ability to grow on a milk-based medium during which growth was monitored by measuring changes in conductance. When the milk medium was inoculated with 106 c.f.u. ml–1, only B. lactis strains DSM 10140T, NCC 363, NCC 383, NCC 311, NCC 387, NCC 402, NCC 239, ATCC 27673, ATCC 27674 and ATCC 27536 showed an increase in conductivity, whereas B. animalis ATCC 25527T and ATCC 27672 did not reveal any changes in the impedance values of the milk medium. All B. lactis strains maintained viable counts greater than 2x108 c.f.u. ml–1 throughout the 24 h fermentation and displayed differences in growth of 1·5 log (Fig. 4a). On the other hand, B. animalis ATCC 25527T and ATCC 27672 did not reveal any significant growth and their viable counts dropped steadily to below the value of 5x107 c.f.u. ml–1 with small differences (below 0·5 log) in relative growth (Fig. 4b). Collectively, these findings indicate that only B. lactis has the potential to grow in milk or milk-based media.
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, the carbohydrate-fermentation patterns of B. animalis and B. lactis were very similar based on the examination of 95 different carbon sources. As some characters varied from strain to strain, it was not possible to define a species-specific pattern for representatives of B. animalis and B. lactis. This is clearly illustrated by the fact that, of all tested carbon sources, only dextrin, 
-D-glucose, maltose, maltotriose, D-raffinose and sucrose were fermented by all tested strains. At the individual strain level, only strains NCC 311 and NCC 362 displayed an identical fermentation behaviour. In addition to the AN MicroPlate characterization, the ability to ferment starch was verified based on the formation of a hydrolysis halo on M144 medium depleted of the usual sugars and supplemented with 0·6 % soluble starch. Meile et al. (1997) asserted that the non-utilization of starch by B. lactis was a major difference between both species. However, consistent with the findings of Cai et al. (2000) and Lauer & Kandler (1983), we observed that neither species was able to use this carbon source.
In support of the proposal of Cai et al. (2000), the DNA–DNA hybridization data and phenotypic results reported in this study are evidence that B. animalis and B. lactis belong to one single species. However, results of protein profiling, genotypic analyses and growth evaluation in milk indicate that both these taxa are clearly different. Based on the fact that members of both species share more than 70 % DNA–DNA relatedness, B. lactis should be reclassified as B. animalis, as required by Rule 42 of the Bacteriological Code (1990 Revision) (Lapage et al., 1992). Taking into account that strains formerly assigned to B. animalis and B. lactis can be clearly distinguished at the intraspecific level, we propose two subspecies in B. animalis, for which the names B. animalis subsp. animalis and B. animalis subsp. lactis are suggested, respectively.
The following descriptions are based on data obtained from the present study and on previously reported data (Scardovi & Trovatelli, 1974; Meile et al., 1997).
Description of Bifidobacterium animalis emend.
Strains display the following characteristics typical for the genus Bifidobacterium: Gram-positive, non-motile, non-spore forming, irregular rod-shaped anaerobes. Glucose is fermented using the characteristic enzyme fructose-6-phosphate phosphoketolase in the so-called Bifidus-shunt. Dextrin, -D-glucose, maltose, maltotriose, D-raffinose and sucrose are fermented; starch is not fermented.
Description of Bifidobacterium animalis subsp. animalis subsp. nov.
Strains display characteristics typical for the species B. animalis as described above. The optimum growth temperature is 39–41 °C. No growth occurs in slants incubated in air or in air enriched with carbon dioxide. No growth occurs in milk or milk-based media. Lactate and acidic acids are produced in a molar ratio of 1 : 3·6±0·3. Strains originate from the faeces of rats. The DNA G+C content is 61·3±0·0 mol%. Type strain: Bifidobacterium animalis subsp. animalis R101-8T (LMG 10508T=ATCC 25527T=DSM 20104T=JCM 1190T).
Description of Bifidobacterium animalis subsp. lactis subsp. nov.
Strains display characteristics typical for the species B. animalis as described above. The optimum growth temperature is 39–42 °C. No growth occurs on agar plates exposed to air, but 10 % oxygen in the headspace atmosphere above liquid media is tolerated. Growth occurs in milk or milk-based media. The molar ratio of acetate to lactate from glucose metabolism is about 10 to 1 under anaerobic conditions, e.g. lactate production is replaced by formate production. Strains have been isolated from fermented milk samples, human and infant faeces, rabbit and chicken faeces and from sewage. The DNA G+C content is 61·0±0·5 mol%. Type strain: Bifidobacterium animalis subsp. lactis UR1T (LMG 18314T=DSM 10140T=JCM 10602T).
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ACKNOWLEDGEMENTS |
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is a subjective synonym of Bifidobacterium animalis (Mitsuoka 1969) Scardovi and Trovatelli 1974. Microbiol Immunol 44, 815–820.[Medline]Ezaki, T., Hashimoto, Y. & Yabuuchi, E. (1989). Fluorometric deoxyribonucleic acid- deoxyribonucleic acid hybridization wells as an alternative to membrane filter hybridization in which radioisotopes are used to determine genetic relatedness among bacterial strains. Int J Syst Bacteriol 39, 224–229.[CrossRef]
Goris, J., Suzuki, K., De Vos, P., Nakase, T. & Kersters, K. (1998). Evaluation of a microplate DNA-DNA hybridisation method compared with the initial renaturation method. Can J Microbiol 44, 1148–1153.[CrossRef]
Lapage, S. P., Sneath, P. H. A., Lessel, E. F., Skerman, V. B. D., Seeliger, H. P. R. & Clark, W. A. (1992). International Code of Nomenclature of Bacteria (1990 Revision). Bacteriological Code. Washington, DC: American Society for Microbiology.
Lauer, E. & Kandler, O. (1983). DNA-DNA homology, murein types and enzyme patterns in the type strains of the genus Bifidobacterium. Syst Appl Microbiol 4, 42–64.
Lawrence, J. G., Hartl, D. L. & Ochman, H. (1991). Molecular considerations in the evolution of bacterial genes. J Mol Evol 33, 241–250.[CrossRef][Medline]
Marmur, J. (1961). A procedure for the isolation of deoxyribonucleic acid from micro-organisms. J Mol Biol 3, 208–218.
Masco, L., Huys, G., Gevers, D., Verbrugghen, L. & Swings, J. (2003). Identification of Bifidobacterium species using rep-PCR fingerprinting. Syst Appl Microbiol 26, 557–563.[CrossRef][Medline]
Meile, L., Ludwig, W., Rueger, U., Gut, C., Kaufmann, P., Dasen, G., Wenger, S. & Teuber, T. (1997). Bifidobacterium lactis sp. nov., a moderately oxygen tolerant species isolated from fermented milk. Syst Appl Microbiol 20, 57–64.
Mesbah, M., Premachandran, U. & Whitman, W. B. (1989). Precise measurement of the G+C content of deoxyribonucleic acid by high-performance liquid chromatography. Int J Syst Bacteriol 39, 159–167.
Pitcher, D. G., Saunders, N. A. & Owen, R. J. (1989). Rapid extraction of bacterial genomic DNA with guanidium thiocyanate. Lett Appl Microbiol 8, 151–156.
Pot, B., Vandamme, P. & Kersters, K. (1994). Analysis of electrophoretic whole-organisms protein fingerprints. In Modern Microbial Methods, Chemical Methods in Prokaryotic Systematics, pp. 493–521. Edited by M. Goodfellow & A. G. O'Donnell. Chichester: Wiley.
Prasad, J., Gill, H., Smart, J. & Gopal, P. K. (1998). Selection and characterization of Lactobacillus and Bifidobacterium strains for use as probiotics. Int Dairy J 8, 993–1002.[CrossRef]
Scardovi, V. & Trovatelli, L. D. (1974). Bifidobacterium animalis (Mitsuoka) comb. nov. and the "minimum" and "subtile" groups of new bifidobacteria found in sewage. Int J Syst Bacteriol 24, 21–28.
Thompson, F. L., Hoste, B., Vandemeulebroecke, K. & Swings, J. (2001). Genomic diversity amongst Vibrio isolates from different sources determined by Fluorescent Amplified Fragment Length Polymorphism. Syst Appl Microbiol 24, 520–538.[CrossRef][Medline]
Ventura, M., Elli, M., Reniero, M. & Zink, R. (2001). Molecular microbial analysis of Bifidobacterium isolates from different environments by species-specific amplified ribosomal DNA restriction analysis (ARDRA). FEMS Microbiol Ecol 36, 113–121.[CrossRef][Medline]
Ventura, M. & Zink, R. (2002). Rapid identification, differentiation, and proposed new taxonomic classification of Bifidobacterium lactis. Appl Environ Microbiol 68, 6429–6434.
Ventura, M. & Zink, R. (2003). Comparative sequence analysis of the tuf and recA genes and restriction fragment length polymorphism of the internal transcribed spacer region sequences supply additional tools for discriminating Bifidobacterium lactis from Bifidobacterium animalis. Appl Environ Microbiol 69, 7517–7522.
Zhu, L., Li, W. & Dong, X. (2003). Species identification of genus Bifidobacterium based on partial HSP60 gene sequences and proposal of Bifidobacterium thermacidophilum subsp. porcinum subsp. nov. Int J Syst Evol Microbiol 53, 1619–1623.
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