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1 Biotechnology Department, Research and Development, Roche Vitamins AG, CH-4070 Basel, Switzerland
2 BCCMTM/LMG Bacteria Collection, Laboratory for Microbiology, Ghent University, K.-L. Ledeganckstraat 35, 9000 Ghent, Belgium
3 TNO Nutrition and Food Research, PO Box 360, 3700 AJ Zeist, The Netherlands
Correspondence
Alan Berry
alan.berry{at}roche.com
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The GenBank accession numbers for the 16S rRNA gene sequences of ATCC 21588T and R-1506 are AF461158 and AF461159, respectively.
Results of DNA fingerprinting (AFLP) analysis and the carotenoid production profile for all pigmented strains used in this work are available as supplementary data in IJSEM Online (http://ijs.sgmjournals.org).
Present address: Natural Products Microbiology, Infectious Diseases Division, Wyeth-Ayerst Research, 401 N. Middletown Road, Pearl River, NY 10965, USA. 
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-carotene, canthaxanthin and astaxanthin) or by extraction from natural sources (lutein/zeaxanthin from marigold, capsanthin/capsorubin from paprika). Commercial production of carotenoids using micro-organisms has been achieved in some cases. For instance, -carotene is produced by fermentation with the fungus Blakeslea trispora (Sibeyn & de Pater, 1998
) or by pond culture using the halotolerant alga Dunaliella salina (Borowitzka, 1999). Lycopene production has also been reported in B. trispora (Marcos et al., 2000). Astaxanthin is produced by fermentation using the red yeast Xanthophyllomyces dendorous (formerly named Phaffia rhodozyma) (Jacobson et al., 2000) or in photobioreactors or open ponds using the alga Haematococcus pluvialis (Lorenz & Cysewski, 2000; Olaizola, 2000).
Zeaxanthin (3,3'-dihydroxy--carotene) is a yellow carotenoid that has application in poultry pigmentation and in the prevention of age-related macular degeneration in humans. In the mid-1960s, scientists at Hoffman-La Roche isolated several marine bacteria that produced zeaxanthin (Schocher & Wiss, 1975). One bacterium, given the strain designation R-1512, was deposited at the American Type Culture Collection as strain ATCC 21588. Using the accepted taxonomic standards of that time, the zeaxanthin-producing bacterium was classified as a member of the genus Flavobacterium, but no species designation was assigned. An extensive mutagenesis and screening programme was conducted in the late 1960s/early 1970s to isolate mutants of strain R-1512 with higher zeaxanthin productivities. With respect to the present work, two such mutants are significant. These mutants, listed in ascending order of their zeaxanthin productivities, are R1534 and R114. A variety of other mutants have been used over the years for biochemical studies of carotenoid biosynthesis (Britton et al., 1977; Goodwin, 1972; McDermott et al., 1974; Mohanty et al., 2000).
In this report, we present the results of an extensive taxonomic analysis of strain ATCC 21588 (=R-1512) that shows that this bacterium and two other independent isolates comprise a new species in the genus Paracoccus. The name Paracoccus zeaxanthinifaciens is proposed, with ATCC 21588T (=R-1512T =LMG 21293T) being designated as the type strain.
The bacterial strains used are listed in Table 1. Strain R-1512T (=ATCC 21588T) and strain R-1506 are two independent isolates from the same initial screening of marine bacteria for zeaxanthin-producing strains. Strains R114 and R1534 are mutants derived from strain R-1512T by classical mutagenesis and screening for improved zeaxanthin production. Strains R114, R1534, and R-1506 were recently deposited (according to the terms of the Budapest Treaty) at the patent depository at the ATCC and were given the strain designations PTA-3335PP, PTA-3336PP and PTA-3431PP, respectively. Strains MBIC 3024, MBIC 3966, MBIC 4017 and MBIC 4020 were identified as members of the genus Paracoccus by their 16S rDNA gene sequences [deposited in EMBL by T. Hamada, Marine Biotechnology Institute (MBI), Kamaishi Laboratories, Kamaishi, Iwate, Japan], and were obtained from H. Kasai, MBI. It should be noted that MBIC 3024 is another designation for [Alcaligenes] sp. strain PC-1, a previously described ketocarotenoid-producing bacterium (Misawa et al., 1995; Yokoyama et al., 1994). Paracoccus marcusii DSM 11574T and Paracoccus carotinifaciens E-396T are type strains of carotenoid-producing species (Harker et al., 1998; Tsubokura et al., 1999a) and were obtained from DSMZ, the German Collection of Microorganisms and Cell Cultures and The National Institute of Advanced Industrial Science and Technology (Japan), respectively. Paracoccus solventivorans DSM 6637T (Siller et al., 1996) was also obtained from DSMZ, and was included in some comparisons as a representative non-carotenogenic species of Paracoccus.
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Genomic DNA was prepared according to the protocol of Niemann et al. (1997). Genes encoding 16S rRNA were amplified from genomic DNA from strains ATCC 21588T, R114, R1534 and R-1506 by PCR. Forward primer 16F27 [5'-AGA GTT TGA TCC TGG CTC AG-3'] was used for strains R1534 and R-1506, while forward primer 16F38 [5'-CTG GCT CAG GAC/T GAA CGC TG-3'] was used for strains ATCC 21588T and R114. The reverse primer 16R1522 [5'-AAG GAG GTG ATC CAG CCG CA-3'] was used for all strains. Purification of PCR products, DNA sequencing using five forward and three reverse primers, and sequence assembly were performed as described by Coeyne et al. (1999), but using the ABI Prism BigDye Terminator Cycle Sequencing Ready Reaction kit. Phylogenetic analysis was performed using the software package GeneCompar version 2.0 (Applied Maths) after including the consensus sequences from strains ATCC 21588T, R114, R1534 and R-1506 in an alignment of small-subunit ribosomal sequences collected from the EMBL database. A similarity matrix was created by homology calculation with a gap penalty of 0 % and after discarding unknown bases. A tree was constructed using the neighbour-joining method.
Genomic DNA was prepared according to the protocol of Wilson (1987). The G+C content of the DNAs was determined by HPLC according to the method of Mesbah et al. (1989) as modified by Logan et al. (2000). Reported values are the mean of three measurements on the same DNA sample. DNA–DNA hybridizations were performed using the initial renaturation rate method (De Ley et al., 1970). The hybridization temperature was 81·5 °C. Values are the mean of at least two determinations.
The same purified DNA as was used for the DNA–DNA hybridization experiments was used for AFLP analysis (Vos et al., 1995). The experimental procedures, the adaptors ligated to the sticky ends generated by cleavage with the restriction enzymes ApaI and TaqI and the primers A01 and T01 were as described by Huys et al. (1996). The other primers used were: A02, 5'-GACTGCGTACAGGCCCC-3'; A03, 5'GACTGCGTACAGGCCCG3'; T02, 5'-CGATGAGTCCTGACCGAC-3'; T03, 5'-CGATGAGTCCTGACCGAG-3'. Primers A01, A02 and A03 were labelled at their 5' end with [32P]ATP (Amersham International) using T4 kinase (Pharmacia Biosciences). The six primer combinations (PCs) used for AFLP were: PC A, A01 + T01; PC B, A01 + T02; PC D, A02 + T01; PC I, A03 + T03; PC G, A03 + T01; PC H, A03 + T02. The electrophoretic patterns were scanned, numerically analysed with GelCompar version 4.2 software (Applied Maths), compared using the Pearson product–moment correlation coefficient and clustered using the unweighted pair group method with averages (UPGMA) linking.
Extraction of cellular fatty acids and determination of the fatty acid compositions by gas chromatography was carried out using the Microbial Identification System (MIDI) according to the instructions of the manufacturer. For this test, bacteria were grown for 24 h at 28 °C on Trypticase Soy Agar.
For testing the aerobic utilization of carbon sources, Biolog-SF-N MicroPlate microtitre plates containing 95 substrates were used (Harker et al., 1998), with the exception that the substrate in well E6 was DL-lactic acid methyl ester instead of the usual sodium salt of DL-lactic acid. Cells were grown for 24 h at 28 °C on Marine Agar. A cell suspension with a density equivalent to 0·5 McFarland units was prepared in sterile distilled water. From this suspension, 18 drops were transferred into 21 ml AUX medium (API 20NE; bioMérieux) and mixed gently. One hundred microlitres of the suspension was transferred to each well of the Biolog MicroPlates, and the plates were incubated at 30 °C. Wells were visually checked for growth after 48 h and after 6 days. Also, at 6 days the visual scoring was confirmed by reading the microtitre plates using the Biolog plate reader.
Nine selected tests [reduction of nitrate to nitrite, denitrification, production of indole from tryptophan, acid production from glucose (fermentation), arginine dihydrolase, urease, aesculin hydrolysis (-glucosidase), gelatin hydrolysis and -galactosidase] were performed using API 20NE strips (bioMérieux). For these tests, cells were grown for 24 h at 28 °C on Marine Agar. Cell suspensions were prepared and strips inoculated according to the instructions of the manufacturer. Strips were incubated at 28 °C and results determined after 24 and 48 h. For testing the temperature, pH and salt concentration ranges for growth, cells were first grown for 24 h at 28 °C on Marine Agar. Cell suspensions having a density of between 1 and 2 McFarlandunits were then prepared in sterile distilled water. These suspensions were used as inocula for each test. For determining the temperature and salt concentration ranges for growth, three drops of cell suspension were transferred onto the surface of each Marine Agar plate. One drop was diluted by streaking, the other two drops were left undisturbed. In the case of the temperature test, plates were incubated at 10, 25, 30, 33, 37 and 40 °C. In the case of the salt tolerance test, the Marine Agar plates had been supplemented with NaCl to reach final concentrations of (w/v) 3, 6 and 8 % and all were incubated at 28 °C. For both tests, plates were checked for growth after 24 h, 48 h and 5 days. Growth was determined as visual growth (confluent in the drops and as colonies in the streaks with diluted inoculum) and scored relative to the growth under the control condition (30 °C for the temperature test and no added NaCl for the salt tolerance test). To test the pH range for growth, three drops of cell suspension were transferred into tubes containing 10 ml Marine Broth having final pH values after autoclaving of pH 6·1, 6·3, 7·0, 7·7, 8·1 and 9·1. The cultures were incubated with shaking at 28 °C. Growth was checked at 24 h, 48 h, 3 days and 6 days. Growth was determined as increased turbidity (measured as % transmission using the Biolog turbidimeter) and scored relative to growth at pH 7·0 (control). Anaerobic growth was tested by streaking a loopful of freshly grown cells on Marine Agar plates, with or without addition of 0·1 % (w/v) KNO3, and incubating at 30 °C in an atmosphere of 10 % CO2 and 90 % N2. To test starch hydrolysis, a loopful of freshly grown cells was applied to Marine agar plates supplemented with 0·2 % (w/v) soluble starch and plates were checked for clear zones after 48 h incubation at 30 °C by flooding of the plates with lugol. Production of poly--hydroxybutyrate (PHB) was tested using the crotonic acid method (Daniels et al., 1994). Escherichia coli LMG 2092T and Ralstonia eutrophus LMG 1201 served as negative and positive controls, respectively, for PHB production.
Colony pigmentation was observed visually and recorded after 5 days growth at 28 °C on Marine Agar. Cell morphology and motility were observed using an Olympus light microscope equipped with phase-contrast optics (magnification x1000). Cells were grown for 24 h at 28 °C on Marine Agar, and cell suspensions were made in sterile saline for microscopic examination.
For analysis of carotenoids by HPLC, 50 ml Marine Broth was inoculated with approximately 0·5 ml of an overnight liquid culture (grown in Marine Broth), and incubated for 24 h at 28 °C with shaking. Cells (approx. 0·35 g wet wt) were collected by centrifugation, washed with saline, and again collected by centrifugation. Carotenoids were extracted from the cells by suspending the pellet in 4 ml tetrahydrofuran (Fluka 87370). Solids were removed from the extracts by centrifugation. Samples were analysed by a reversed phase HPLC method that allowed simultaneous determination of astaxanthin, adonixanthin, zeaxanthin, canthaxanthin, -carotene and lycopene. The method is also able to separate the main cis-isomers of zeaxanthin. Chromatography was performed using an Agilent 1100 HPLC system equipped with a temperature-controlled autosampler and a diode array detector. The column was a YMC Carotenoid C30 column (5 micron, steel, 250 mm longx4·6 mm i.d.; Waters). The guard column was a Pelliguard LC-18 cartridge (20 mm; SUPELCO). The mobile phase was a methanol/methyl tert-butyl ether (MeOH/TBME) gradient having the following parameters (all percentages expressed as v/v): start, 80 % MeOH/20 %TBME; 10 min, 65 % MeOH/35 % TBME; 20 min, 10 % MeOH/90 % TBME. The flow rate was 1·0 ml min-1. The injection volume and column temperature were 10 µl and 15 °C, respectively. Carotenoids were detected by absorbance at 450 nm. Quantification of carotenoids was performed with a two level calibration using external standards. Calculations were based on peak areas. The selectivity of the method was checked by injecting standard solutions of the relevant carotenoid reference compounds. The target compounds (all-trans-carotenoids) were completely separated and showed no interference. Some minor cis isomers may co-elute, although these potentially interfering isomers are rare and may be neglected in routine analysis. The retention times (given in min) for the different carotenoids using this HPLC method were as follows: astaxanthin, 6·99; adonixanthin, 7·50; 15-cis-zeaxanthin, 7·80; 13-cis-zeaxanthin, 8·23; all-trans-zeaxanthin, 9·11; canthaxanthin, 9·95; cryptoxanthin, 13·45; -carotene, 17·40 and lycopene, 21·75. The linearity, sensitivity and reproducibility of the method with respect to detection of zeaxanthin were tested. A linear range was found from 0·1 µg ml-1 to 250 µg ml-1 zeaxanthin (correlation coefficient 0·9998). The lower limit of detection for zeaxanthin was determined to be 60 µg l-1. Using a higher injection volume and optimization of the integration parameters, it is possible to lower the detection limit to approximately 5 µg l-1. The retention time for all-trans-zeaxanthin was very stable [relative standard deviation (RSD), 0·2 %]. The peak area reproducibility, based on ten repetitive analyses of the same culture sample, was determined to be 0·17 % RSD for all-trans-zeaxanthin.
The complete sequence of the 16S rRNA genes from the zeaxanthin-producing isolate, strain ATCC 21588T (=R-1512T), its two mutant derivatives, R114 and R1534, and the independent zeaxanthin-producing isolate, strain R-1506, were determined (1404, 1404, 1415 and 1415 nucleotides, respectively). The sequences from ATCC 21588T, R114 and R1534 were identical. The sequences from R-1506 differed in only one nucleotide (position 1350) from the sequence from ATCC 21588T. This demonstrated that the two independent zeaxanthin-producing isolates are phylogenetically highly related and are likely to belong to the same species. Comparison of the ATCC 21588T and R-1506 sequences with those publicly available at the EMBL library showed that these bacteria, formerly classified as unidentified species of Flavobacterium, should be reclassified as members of the genus Paracoccus (for a review of the taxonomy of the genus Paracoccus, see Baj, 2000). However, the sequence similarities observed with all 14 currently validly described Paracoccus species was <97 %, the limit for a possible relatedness at the species level (Stackebrandt & Goebel, 1994). This indicated that strains ATCC 21588T and R-1506 belonged to one or two new species of Paracoccus. Sequence similarities of >97 % were observed between the 16S rDNA sequences of strains ATCC 21588T and R-1506 and several unnamed Paracoccus strains, suggesting that one or more of the unnamed (MBIC) strains could be related at the species level to strains ATCC 21588T and/or R-1506. Based on cluster analysis (Fig. 1), strains ATCC 21588T, R114, R1534, R-1506 and MBIC 3966 were selected for DNA–DNA hybridization experiments to analyse species relatedness. The sequence similarity of the selected MBIC strains and any of the 14 validly described Paracoccus species was <97 % (data not shown).
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. Strains ATCC 21588T, R114, R1534, R-1506 and MBIC 3966 showed a DNA reassociation value of 
88 %, clearly above the generally accepted limit (70 %) for species delineation (Wayne et al., 1987), and therefore belong to a new species within the genus Paracoccus, for which the name P. zeaxanthinifaciens is proposed. The G+C content of these five strains ranged from 66·9 to 67·7 mol%, thus remaining within 1 mol%, characteristic for a well defined species (Vandamme et al., 1996). On the other hand, the low DNA relatedness between strains MBIC 3024, MBIC 4017 and MBIC 4020 themselves, and between these strains and strains ATCC 21588T, R114, R1534, R-1506 and MBIC 3966 showed that MBIC 3024, MBIC 4017 and MBIC 4020 each belong to a different additional new species within the genus Paracoccus.
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Table 3 summarizes the mean cellular fatty acid composition of the 3 wild-type strains constituting P. zeaxanthinifaciens and the fatty acid composition of P. marcusii DSM 11574T, P. carotinifaciens E-396T and P. solventivorans DSM 6637T. All four species showed a comparable cellular fatty acid profile, with 18 : 1
7c as the major compound. This is consistent with fatty acid compositions reported previously for the three type strains (Harker et al., 1998; Tsubokura et al., 1999a; Siller et al., 1996), and indeed with the fatty acid composition characteristic of the entire genus Paracoccus (Baj, 2000). No attempt was made to determine the significance of the minor differences in fatty acid composition between P. zeaxanthinifaciens and the three type strains, as variability of these minor compounds within the latter Paracoccus species could not be assessed since these species currently contain only the type strain analysed.
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). Strains ATCC 21588T, R-1506 and MBIC 3966 were incapable of nitrate reduction or denitrification. Colony pigmentation further distinguishes the latter strains from all other species of Paracoccus. Among the established species of Paracoccus, only P. marcusii DSM 11574T, P. carotinifaciens E-396T and P. thiocyanatus THIO11T exhibit pigmented colonies (Baj, 2000). In the case of P. thiocyanatus THIO11T, the reddish pigment was determined to not be a carotenoid(s) (Harker et al., 1998). Although P. marcusii DSM 11574Tand P. carotinifaciens E-396T do produce carotenoids, they produce mainly ketocarotenoids (Hirschberg & Harker, 1999; Tsubokura et al., 1999b), hence the orange-pink colony pigmentation. Strains ATCC 21588T (R114, R1534), R-1506 and MBIC 3966 produce exclusively zeaxanthin (see below), giving rise to the yellow-orange colony pigmentation.
Strains ATCC 21588T, R-1506 and MBIC 3966 were subjected to a battery of other phenotypic tests and compared more specifically to the two known carotenoid-producing species P. marcusii DSM 11574T and P. carotinifaciens E-396T. Of the 95 carbon sources tested as growth substrates using the Biolog SF-N system, 12 could be used, and 53 could not be used by all strains of P. zeaxanthinifaciens (these carbon sources are listed below in the description of the new species). The strains gave variable growth responses to the remaining 25 substrates. Based on this test, P. zeaxanthinifaciens could be distinguished from P. marcusii DSM 11574T and P. carotinifaciens E-396T by the inability of P. zeaxanthinifaciens to use seven carbon sources (adonitol, erythritol, gentiobiose, methyl -glucoside, D-sorbitol, xylitol and quinic acid). Two additional carbon sources that were utilized by P. zeaxanthinifaciens (L-asparagine and L-aspartic acid) were not used for growth by P. marcusii DSM 11574T. The growth characteristics of P. marcusii DSM 11574T on the nine carbon sources mentioned above are the same as those reported by Harker et al. (1998), with the exception of methyl -glucoside, which in their hands did not support growth of the organism. Tsubokura et al. (1999a), in the original description of P. carotinifaciens E-396T, did not report whether these nine carbon sources can be used by this strain, so no comparison with the present results can be made.
P. zeaxanthinifaciens grew weakly or not at all at 10 °C, but grew well from 25 to 40 °C. Neither P. marcusii DSM 11574T nor P. carotinifaciens E-396T could grow at 40 °C. P. zeaxanthinifaciens grew on Marine Agar supplemented with 3, 6 or 8 % NaCl, and in Marine Broth having a final pH ranging from 6·1 to 9·1. In contrast, P. marcusii DSM 11574T and P. carotinifaciens E-396T grew very poorly in the presence of 8 % NaCl, and in addition P. marcusii DSM 11574T was not able to grow at pH 9·1. Urease was positive for P. zeaxanthinifaciens (although weak for the two mutants) and negative for P. marcusii DSM 11574T and P. carotinifaciens E-396T.
Cells of P. zeaxanthinifaciens strains were coccoid to short rods with dimensions of 0·8–0·9x1·1–1·5 µm. P. marcusii DSM 11574T, P. carotinifaciens E-396T and P. solventivorans DSM 6637T all appeared as short rods. The shape and size of these cells was similar to the original reports (Harker et al., 1998; Tsubokura et al., 1999a; Siller et al., 1996).
Table 4 summarizes the results of the physiological tests that differentiated between P. zeaxanthinifaciens, P. marcusii and P. carotinifaciens. Other characteristics that are shared by all members of P. zeaxanthinifaciens but not discriminative towards P. marcusii or P. carotinifaciens, are given in the species description.
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) and that P. marcusii DSM 11574T and P. carotinifaciens E-396T produce primarily ketocarotenoids (Hirschberg & Harker, 1999; Tsubokura et al., 1999b). Based on the work reported here, we propose that strains ATCC 21588T (=R-1512T), R-1506 and MBIC 3966 be classified as a new species of the genus Paracoccus, having the name Paracoccus zeaxanthinifaciens sp. nov., and further propose that ATCC 21588T (=R-1512T =LMG 21293T) be designated as the type strain.
Description of Paracoccus zeaxanthinifaciens sp. nov.
Paracoccus zeaxanthinifaciens (ze.a.xan.thi.ni.fa'ci.ens. N.L. neut. n. zeaxanthinum zeaxanthin; L. part. pres. faciens making/producing; N.L. adj. zeaxanthinifaciens zeaxanthin-producing).
Short rods to cocci, 0·8–0·9x1·1–1·5 µm, growing singly, in pairs or in short chains. Non-motile and Gram-negative. Colonies on Marine Agar are circular, convex, smooth and deep yellow to orange due to accumulation of the carotenoid zeaxanthin. Astaxanthin and other ketocarotenoids are not produced. Growth is strictly aerobic. Catalase, oxidase and -galactosidase are positive. -Glucosidase is positive, but weak for the type strain. Arginine dihydrolase is negative. Indole is not produced from tryptophan and glucose is not fermented. Starch and gelatin are not hydrolysed. Nitrate is not reduced to nitrite, and denitrification is not performed. PHB is produced. Grows weakly or not at all at 10 °C, but grows well from 25 to 40 °C. Grows on Marine Agar containing 3–8 % NaCl, and grows in Marine Broth at pH 6·1–9·1. Utilizes D-arabitol, D-galactose, 
-D-glucose, myo-inositol, -lactose, D-mannitol, D-melibiose, D-trehalose, L-asparagine, L-aspartic acid, L-glutamic acid and L-pyroglutamic acid for growth, but does not utilize -cyclodextrin, dextrin, glycogen, Tween 40, Tween 80, N-acetyl-D-galactosamine, N-acetyl-D-glucosamine, adonitol, L-arabinose, i-erythritol, L-fucose, gentobiose, methyl -glucoside, D-raffinose, L-rhamnose, D-sorbitol, xylitol, acetic acid, cis-aconitic acid, citric acid, formic acid, D-galacturonic acid, D-glucosaminic acid, 
-hydroxybutyric acid, p-hydroxyphenylacetic acid, itaconic acid, -ketobutyric acid, -ketovaleric acid, DL-lactic acid methyl ester, malonic acid, propionic acid, quinic acid, succinic acid, bromosuccinic acid, succinamic acid, alaninamide, D-alanine, L-histidine, hydroxy-L-proline, L-phenylalanine, D-serine, L-threonine, DL-carnitine, urocanic acid, inosine, uridine, thymidine, phenylethylamine, putrescine, 2-aminoethanol, 2,3-butanediol, glucose 1-phosphate or glucose 6-phosphate. The major non-hydroxyl fatty acid is C18 : 1(7c) and the major hydroxyl fatty acid is 3-OH C10 : 0. The G+C content of the DNA from the strains currently within the species ranges from 66·9 to 67·7 mol%. The type strain is ATCC 21588T =R-1512T =LMG 21293T, and it was isolated from seaweed collected from the coast of the African Red Sea. Reference strains are R-1506 and MBIC 3966.
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