废水处理系统中重要功能类群Thauera属种群结构与功能的研究
详细信息    本馆镜像全文|  推荐本文 |  |   获取CNKI官网全文
摘要
在废水生物处理系统中,污染物的去除主要是由其中的微生物群落完成的,微生物群落的结构和功能往往决定了整个装置的性能。Thauera属细菌就是一类广泛存在于各种类型的废水处理装置中并具有多种芳香族污染物降解能力的重要功能类群,深入研究其种群组成与降解功能,具有重要的理论意义和应用价值。
     由于分离比较困难、缺少专一性的分子检测方法,人们对于废水处理系统中Thauera属种群的结构与功能的了解还非常有限。
     为此,本论文建立了Thauera特异性、嵌套式PCR-DGGE (denaturing gradient gel electrophoresis)的方法用于环境样品中Thauera种群结构的研究。对一个来自废水处理系统的生物膜样品以Thauera特异性PCR扩增产物建立了克隆文库,所有测序的克隆(91个)都属于Thauera属,说明该PCR方法具有很高特异性。为了快速简便的比较不同样品中Thauera的结构,以特异性PCR扩增产物为模板进行16S rRNA基因V3区嵌套式扩增后用于DGGE分析,建立了Thauera特异性PCR-DGGE方法。13个带有不同V3序列的克隆在DGGE上形成了10条条带,说明该方法具有较高的分辨率,可以用于检测样品中的Thauera属的组成。
     在一个大型焦化废水处理装置由于回流泵机械故障而导致化学需氧量(COD)去除率逐渐降低时,我们使用建立的特异性PCR-DGGE方法对系统中Thauera的结构变化进行了追踪监测。对DGGE指纹图谱进行主成份分析(PCA)后,发现当系统COD的去除率从前4周的84.1±2.7%降到第5和第6周的低于75%时,Thauera种群结构呈现出相应的时间演替轨迹,表明Thauera种群结构的变化与系统COD去除功能的波动密切相关。
     为了分离焦化废水处理系统中常规方法难以分离的Thauera细菌,我们设计了一套以Thauera特异性16S rRNA基因为靶向的分离方法。根据已知Thauera的代谢特性及目标菌的生活环境,设计了6种培养基,并使用Thauera特异性PCR-DGGE方法对焦化废水处理装置反硝化池样品在不同培养条件下获得的Thauera多样性进行了分析。选取多样性较高的培养基1/10 NB与MMQ在好氧条件下进行分离培养。以Thauera特异性PCR方法筛选阳性菌落,并用DGGE方法检验菌落的纯度。将含有Thauera的混合菌落在不同的选择性培养基上多次划线,使用特异性PCR及DGGE方法追踪含有Thauera的菌落并分析其纯度,最终从反硝化池样品中纯化获得了3株Thauera菌株(Q4、3-35和Q20-C)。这种以特异性分子标记为靶向分离培养细菌的方法,提高了细菌筛选的灵敏度,可协助分离常规方法难以分离的细菌。
     随后,对这3株Thauera菌株的基因及其污染物降解能力进行了比较分析。测序结果表明这3株Thauera细菌具有相同的16S rRNA基因,但是它们的ERIC-PCR基因组指纹图谱却存在显著差异(相似性<65%)。使用气相色谱-质谱(GC/MS)联用技术对这3株菌的污染物降解能力进行了测定。好氧条件下,它们能降解焦化废水中除喹啉外的所有主要有机污染物(苯酚、甲酚、二甲酚及吲哚等),但苯酚降解速度却各不相同(Q4>3-35> Q20-C)。在已知的8个Thauera种中,只有T. phenylacetia能在好氧条件下降解其中的一种污染物(苯酚)。虽然Q20-C含有亚硝酸盐还原酶基因(nirS),但这3株菌都没有显现出反硝化能力,而已知的Thauera细菌则都是反硝化菌。以上结果表明,这3个从焦化废水处理装置中分离到的细菌可能代表了一种新的Thauera类型,具有广泛的芳香族污染物降解能力。
     为了了解Thauera属细菌在废水处理脱氮过程中的作用,我们研究了8个不同来源的Thauera菌株(T. aminoaromatica、T. linaloolentis、T. phenylacetica、T. terpenica、Thauera sp. DNT-1、Thauera sp. 27、Thauera sp. 28及Thauera sp. 63)的反硝化功能及基因。反硝化过程中,所有Thauera均只产生少量的NO(<50 nmol/flask;< 32.9 nM in liquid)。除T. phenylacetica外,其余Thauera菌株都可将硝态氮彻底转化为氮气。PCR分析后发现T. phenylacetica缺失了N2O还原酶(nos)基因,使它的反硝化终产物为N2O。对T. aminoaromatica在不同pH、O2及NOx条件下的反硝化过程进行分析后发现,在pH7-9范围内,随着pH的升高,积累的中间产物(NO和N2O)会减少。相比于NO3-,以NO2-作为电子受体,不仅使得积累的N2O量显著升高(6-40倍),还使其开始反硝化的O2浓度从<0.8μM提高到了>4μM。这说明NO2-能诱导T. aminoaromatica反硝化基因的表达。在反硝化进行时,只有N2O还原酶的活性会被O2立刻抑制,而其他反硝化酶的活性基本不受的影响。这进一步阐明了氧气调控反硝化过程的机制。
     本论文建立了废水处理系统中重要功能类群Thauera属种群结构的专一性检测方法,并在该方法的协助下从废水处理系统中分离到了常规方法难以分离的Thauera菌株;揭示了Thauera属细菌在废水处理系统的有毒有机污染物降解及脱氮过程中的作用,并阐述了不同环境条件对Thauera反硝化功能的影响,为维持及强化废水处理装置的功能提供了菌种资源和理论依据。
It has been well recognized that the performance of a wastewater treatment plant (WWTP) is mainly determined by the structure and activity of its microbial community. Thauera genus has been known as one of the functionally important groups, which has been widely found in WWTPs and mostly shown high versatile organic substrate degrading capacity. However, our understanding of the structure and function of Thauera genus in WWTPs was still limited, due to its resistance to isolation and lack of group-specific analysis method.
     Therefore, a Thauera-specific nested-PCR denaturing gradient gel electrophoresis (DGGE) method was firstly developed, and its usefulness was demonstrated by monitoring the structural shifts of Thauera spp. in an anaerobic-anoxic-oxic fixed-biofilm coking wastewater treatment plant (WWTP) responding to operational perturbations. The specificity of the PCR method was demonstrated by the fact that all 16S rRNA gene sequences, which were cloned from the amplicons of a biofilm sample, belonged to Thauera genus. 16S rRNA gene V3 region was then amplified from the first round Thauera-specific PCR product and applied for DGGE analysis. Different amplified fragments of Thauera clones, with 13 different V3 regions, migrated into 10 positions on DGGE gel, which demonstrated the high resolution of this DGGE method.
     When the WWTP experienced a gradual deterioration in chemical oxygen demand (COD) removal function due to a mechanical failure of the recirculation pump, biofilm samples were collected from the reactor and analyzed by this method. Principal component analysis (PCA) of the DGGE fingerprinting data showed that the composition of Thauera group exhibited a time related trajectory when the plant’s COD removal rate decreased from 84.1±2.7% in the first 4 weeks to less than 75% at week 5 and 6, suggesting the structural shift of Thuaera genus was closely related with the system’s COD removal function.
     A new isolation method that was guided by Thauera-specific PCR and DGGE fingerprinting was developed to get purified cultures of the Thauera spp. from the WWTP. According to the physiological characteristics of known Thauera strains and the living environment of the target bacteria, six types of media were designed. The biofilm from the denitrifying bioreactor of the coking WWTP was inoculated to these media and cultured under both aerobic and anaerobic conditions, the diversity of Thauera spp. that grew under different conditions were analyzed by Thauera-specific PCR-DGGE. Two media (1/10 NB and MMQ) which recovered higher diversity of Thauera spp. and lower number of colonies were used to isolate Thauera sp. under aerobic condition. The colonies were screened by Thauera-specific PCR. The homogeneity of colonies showing positive PCR signals was then checked by DGGE. The colonies with multiple species were further purified by being streaked on different selective media. The positive colonies were tracked by Thauera-specific PCR, and their homogeneity analyzed by DGGE. Finally, three Thauera strains (Q4, Q20-C and 3-35) were isolated to pure cultures. Guided with group-specific PCR and DGGE method, the efficiency and sensitivity of bacterial isolation can thus be significantly improved.
     The functional genes and pollutants-degrading capacity of these Thauera isolates were then characterized. Although sequencing analysis showed they had identical 16S rRNA genes, their ERIC-PCR fingerprinting patterns were significantly different (similarity <65%), indicating wide variation of genomic structures. The degradation of organic pollutants in coking wastewater by these strains was studied with gas chromatography-mass spectrometry (GC/MS). All the main organic pollutants (phenol, methylphenol, quinoline and indole) in the coking wastewater, except quinoline, were degraded by them under aerobic condition. Their phenol degradation rates were different (Q4> 3-35> Q20-C). However, within all the eight Thauera species, only T. phenylacetia has ability to degrade one of these aromatic compounds (phenol) under aerobic condition. Nitrite reductase gene (nirS) was detected in Q20-C, but none of these strains showed denitrification capacity. However, all the known Thauera species were denitrifiers. These results suggested the Thauera strains that isolated from coking WWTP may represent a new Thauera species, and have high versatile aromatic compounds degrading capacity.
     To understand the roles of Thauera spp. played in nitrogen removal in WWTPs, the denitrification genes and functions of 8 Thauera strains (T. aminoaromatica, T. linaloolentis, T. phenylacetica, T. terpenica, Thauera sp. DNT-1, Thauera sp. 27, Thauera sp. 28 and Thauera sp. 63) that originated from different environments were studied. All the strains emitted little NO (<50 nmol per flask; < 32.9 nM in liquid) during the denitrification process. All of them were able to transform nitrate to N2, except T. phenylacetica, which produced N2O as the final product and was found to be lack of N2O reductase (nos) gene. Analysis of the denitrification of T. aminoaromatica under different pH、O2 and NOx condition, indicated that in its activity pH range (pH 7-9), less intermediates (NO and N2O) were produced at higher pH level. In comparison to nitrate, the N2O production was significantly increased (6-40 times) under nitrite condition; the O2 level that denitrification started was also increased from < 0.8μM (in nitrate) to > 4μM (in nitrite), showed the induction effect of nitrite on denitrification of T. aminoaromatica. The activity of N2O reductase was inhibited by O2, but the other denitrification enzymes were not affected. It improved our knowledge on the regulation mechanism of O2 on denitrification.
     In conclusion, a Thauera specific PCR-DGGE method was developed for analyzing the structure of this functionally important population in WWTPs. The Thauera strains in the coking WWTP, which were known as difficult to be isolated by conventional methods, were isolated under the guidance of this group-specific method. This study demonstrated the important roles of Thauera spp. in degradation of toxic organic pollutants and nitrogen removal in WWTPs, illustrated the impacts of different conditions on the denitrification of Thauera, and provided new insights, strains and methods for optimizing the function of WWTPs.
引文
1. Cook, K.L., et al., Effect of microbial species richness on community stability and community function in a model plant-based wastewater processing system. Microb Ecol, 2006. 52(4): p. 725-37.
    2. Wagner, M. and A. Loy, Bacterial community composition and function in sewage treatment systems. Curr Opin Biotechnol, 2002. 13(3): p. 218-27.
    3. Jaspers, E. and J. Overmann, Ecological significance of microdiversity: identical 16S rRNA gene sequences can be found in bacteria with highly divergent genomes and ecophysiologies. Appl Environ Microbiol, 2004. 70(8): p. 4831-9.
    4. Amann, R.I., W. Ludwig, and K.H. Schleifer, Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol Rev, 1995. 59(1): p. 143-69.
    5. Hugenholtz, P., B.M. Goebel, and N.R. Pace, Impact of culture-independent studies on the emerging phylogenetic view of bacterial diversity. J Bacteriol, 1998. 180(18): p. 4765-74.
    6. Wagner, M., et al., Probing activated sludge with oligonucleotides specific for proteobacteria: inadequacy of culture-dependent methods for describing microbial community structure. Appl Environ Microbiol, 1993. 59(5): p. 1520-5.
    7. Ward, D.M., R. Weller, and M.M. Bateson, 16S rRNA sequences reveal numerous uncultured microorganisms in a natural community. Nature, 1990. 345(6270): p. 63-5.
    8. Moyer, C.L., F.C. Dobbs, and D.M. Karl, Estimation of diversity and community structure through restriction fragment length polymorphism distribution analysis of bacterial 16S rRNA genes from a microbial mat at an active, hydrothermal vent system, Loihi Seamount, Hawaii. Appl Environ Microbiol, 1994. 60(3): p. 871-9.
    9. Barns, S.M., et al., Remarkable archaeal diversity detected in a Yellowstone National Park hot spring environment. Proc Natl Acad Sci U S A, 1994. 91(5): p. 1609-13.
    10. Berthelet, M., L.G. Whyte, and C.W. Greer, Rapid, direct extraction of DNA from soils forPCR analysis using polyvinylpolypyrrolidone spin columns. FEMS Microbiol Lett, 1996. 138(1): p. 17-22.
    11. Borneman, J. and E.W. Triplett, Molecular microbial diversity in soils from eastern Amazonia: evidence for unusual microorganisms and microbial population shifts associated with deforestation. Appl Environ Microbiol, 1997. 63(7): p. 2647-53.
    12. Felske, A., et al., Phylogeny of the main bacterial 16S rRNA sequences in Drentse A grassland soils (The Netherlands). Appl Environ Microbiol, 1998. 64(3): p. 871-9.
    13. Kogure, K., U. Simidu, and N. Taga, A tentative direct microscopic method for counting living marine bacteria. Can J Microbiol, 1979. 25(3): p. 415-20.
    14. Kogure, K., U. Simidu, and N. Taga, Distribution of viable marine bacteria in neritic seawater around Japan. Can J Microbiol, 1980. 26(3): p. 318-23.
    15. Ferguson, R.L., E.N. Buckley, and A.V. Palumbo, Response of marine bacterioplankton to differential filtration and confinement. Appl Environ Microbiol, 1984. 47(1): p. 49-55.
    16. Torsvik, V., J. Goksoyr, and F.L. Daae, High diversity in DNA of soil bacteria. Appl Environ Microbiol, 1990. 56(3): p. 782-7.
    17. Jones, J., The Effect of Environmental Factors on Estimated Viable and Total Populations of Planktonic Bacteria in Lakes and Experimental Enclosures. Freshwater Biology, 1977. 7(1): p. 69-71.
    18. Wagner, M., et al., Development of an rRNA-targeted oligonucleotide probe specific for the genus Acinetobacter and its application for in situ monitoring in activated sludge. Appl Environ Microbiol, 1994. 60(3): p. 792-800.
    19. Costerton, J.W., et al., Biofilms, the customized microniche. J Bacteriol, 1994. 176(8): p. 2137-42.
    20. Kaeberlein, T., K. Lewis, and S.S. Epstein, Isolating "uncultivable" microorganisms in pure culture in a simulated natural environment. Science, 2002. 296(5570): p. 1127-9.
    21. Liu, B., et al., Thauera and Azoarcus as functionally important genera in a denitrifying quinoline-removal bioreactor as revealed by microbial community structure comparison.FEMS Microbiol Ecol, 2006. 55(2): p. 274-86.
    22. Meyerhans, A., J.P. Vartanian, and S. Wain-Hobson, DNA recombination during PCR. Nucleic Acids Res, 1990. 18(7): p. 1687-91.
    23. Suzuki, M., M.S. Rappe, and S.J. Giovannoni, Kinetic bias in estimates of coastal picoplankton community structure obtained by measurements of small-subunit rRNA gene PCR amplicon length heterogeneity. Appl Environ Microbiol, 1998. 64(11): p. 4522-9.
    24. Suzuki, M.T. and S.J. Giovannoni, Bias caused by template annealing in the amplification of mixtures of 16S rRNA genes by PCR. Appl Environ Microbiol, 1996. 62(2): p. 625-30.
    25. Juretschko, S., et al., Combined molecular and conventional analyses of nitrifying bacterium diversity in activated sludge: Nitrosococcus mobilis and Nitrospira-like bacteria as dominant populations. Appl Environ Microbiol, 1998. 64(8): p. 3042-51.
    26. Snaidr, J., et al., Phylogenetic analysis and in situ identification of bacteria in activated sludge. Appl Environ Microbiol, 1997. 63(7): p. 2884-96.
    27. Juretschko, S., et al., The microbial community composition of a nitrifying-denitrifying activated sludge from an industrial sewage treatment plant analyzed by the full-cycle rRNA approach. Syst Appl Microbiol, 2002. 25(1): p. 84-99.
    28. Daims, H., et al., Nitrification in sequencing biofilm batch reactors: lessons from molecular approaches. Water Sci Technol, 2001. 43(3): p. 9-18.
    29. Bond, P.L., et al., Bacterial community structures of phosphate-removing and non-phosphate-removing activated sludges from sequencing batch reactors. Appl Environ Microbiol, 1995. 61(5): p. 1910-6.
    30. Christensson, M., L.L. Blackall, and T. Welander, Metabolic transformations and characterisation of the sludge community in an enhanced biological phosphorus removal system. Applied Microbiology and Biotechnology, 1998. 49(2): p. 226-234.
    31. Dabert, P., et al., Characterisation of the microbial 16S rDNA diversity of an aerobic phosphorus-removal ecosystem and monitoring of its transition to nitrate respiration. Applied Microbiology and Biotechnology, 2001. 55(4): p. 500-509.
    32. Liu, W.T., et al., In situ identification of polyphosphate- and polyhydroxyalkanoate-accumulating traits for microbial populations in a biological phosphorus removal process. Environmental Microbiology, 2001. 3(2): p. 110-122.
    33.严兴, A2/O固定生物膜法焦化废水处理系统群落空间演替模式的系统轨迹分析及应用(博士论文). 2006.
    34. Bond, P.L., et al., Identification of some of the major groups of bacteria in efficient and nonefficient biological phosphorus removal activated sludge systems. Applied and Environmental Microbiology, 1999. 65(9): p. 4077-4084.
    35. Manz, W., et al., Application of a suite of 16S rRNA-specific oligonucleotide probes designed to investigate bacteria of the phylum cytophaga-flavobacter-bacteroides in the natural environment. Microbiology, 1996. 142 ( Pt 5): p. 1097-106.
    36. Neef, A., et al., Population analysis in a denitrifying sand filter: conventional and in situ identification of Paracoccus spp. in methanol-fed biofilms. Appl Environ Microbiol, 1996. 62(12): p. 4329-39.
    37. Kampfer, P., et al., Characterization of Bacterial Communities from Activated Sludge: Culture-Dependent Numerical Identification Versus In Situ Identification Using Group- and Genus-Specific rRNA-Targeted Oligonucleotide Probes. Microb Ecol, 1996. 32(2): p. 101-21.
    38. Manz, W., et al., In situ characterization of the microbial consortia active in two wastewater. Water Res, 1994. 28: p. 1715?723.
    39. Amann, R., et al., In situ visualization of high genetic diversity in a natural microbial community. J Bacteriol, 1996. 178(12): p. 3496-500.
    40. Mao, Y.J., et al., Development of group-specific PCR-DGGE 1 GE fingerprinting for monitoring structural changes of Thauera spp. in an industrial wastewater treatment plant responding to operational perturbations. J Microbiol Methods, 2008.
    41. Carr, E.L., et al., Seven novel species of Acinetobacter isolated from activated sludge. Int J Syst Evol Microbiol, 2003. 53: p. 953-963.
    42. Wentzel, M.C., et al., Enhanced polyphosphate organism cultures in activated sludge systems. Part1. Enhanced culture development. Water S. A. (Pretoria), 1988. 14: p. 81-92.
    43. Lotter, L.H., The role of bacterial phosphate metabolism in enhanced phosphorus removal from the activated sludge process. Water Sci. Technol., 1985. 17(2): p. 127-138.
    44. Buchan, L., Possible biological mechanism of phosphorus removal. Water Sci. Technol., 1983. 15(3-4): p. 87-103.
    45. Beacham, A.M., R.J. Seviour, and K.C. Lindrea, Polyphosphate accumulating abilities of Acinetobacter isolates from a biological nutrient removal pilot plant. Water Res, 1992. 26: p. 121-122.
    46. Deinema, M.H., M.v. Loosdrecht, and A. Scholten, Some physiological characteristics of Acinetobacter spp. accumulating large amounts of phosphate. Water Sci. Technol., 1985. 119-125: p. 17.
    47. Ohtake, H., et al., Uptake and release of phosphate by a pure culture of Acinetobacter calcoaceticus. Water Res, 1985. 19: p. 1587-1594.
    48. Lorenz, P., et al., Screening for novel enzymes for biocatalytic processes: accessing the metagenome as a resource of novel functional sequence space. Curr Opin Biotechnol, 2002. 13(6): p. 572-7.
    49. Wagner, M., et al., In situ analysis of microbial consortia in activated sludge using fluorescently labelled, rRNA-targeted oligonucleotide probes and confocal scanning laser microscopy. J Microsc, 1994. 176(Pt 3): p. 181-7.
    50. Ginige, M.P., et al., Use of stable-isotope probing, full-cycle rRNA analysis, and fluorescence in situ hybridization-microautoradiography to study a methanol-fed denitrifying microbial community. Appl Environ Microbiol, 2004. 70(1): p. 588-96.
    51. Li, M., et al., Symbiotic gut microbes modulate human metabolic phenotypes. Proc Natl Acad Sci U S A, 2008. 105(6): p. 2117-22.
    52. Tschech, A. and G. Fuchs, Anaerobic degradation of phenol by pure cultures of newly isolated denitrifying pseudomonads. Arch Microbiol, 1987. 148(3): p. 213-7.
    53. Anders, H.J., et al., Taxonomic position of aromatic-degrading denitrifying pseudomonad strains K 172 and KB 740 and their description as new members of the genera Thauera, as Thauera aromatica sp. nov., and Azoarcus, as Azoarcus evansii sp. nov., respectively, members of the beta subclass of the Proteobacteria. Int J Syst Bacteriol, 1995. 45(2): p. 327-33.
    54. Song, B., et al., Characterization of halobenzoate-degrading, denitrifying Azoarcus and Thauera isolates and description of Thauera chlorobenzoica sp. nov. Int J Syst Evol Microbiol, 2001. 51(Pt 2): p. 589-602.
    55. Shinoda, Y., et al., Aerobic and anaerobic toluene degradation by a newly isolated denitrifying bacterium, Thauera sp. strain DNT-1. Appl Environ Microbiol, 2004. 70(3): p. 1385-92.
    56. Valle, A., et al., N-acyl-l-homoserine lactones (AHLs) affect microbial community composition and function in activated sludge. Environ Microbiol, 2004. 6(4): p. 424-33.
    57. Thomsen, T.R., Y. Kong, and P.H. Nielsen, Ecophysiology of abundant denitrifying bacteria in activated sludge. FEMS Microbiol Ecol, 2007. 60(3): p. 370-82.
    58. Manefield, M., et al., RNA stable isotope probing, a novel means of linking microbial community function to phylogeny. Appl Environ Microbiol, 2002. 68(11): p. 5367-73.
    59. Allen, M.S., et al., Analysis and glycosyl composition of the exopolysaccharide isolated from the floc-forming wastewater bacterium Thauera sp. MZ1T. Environ Microbiol, 2004. 6(8): p. 780-90.
    60. Macy, J.M., et al., Thauera selenatis gen. nov., sp. nov., a member of the beta subclass of Proteobacteria with a novel type of anaerobic respiration. Int J Syst Bacteriol, 1993. 43(1): p. 135-42.
    61. Macy, J.M., T.A. Michel, and D.G. Kirsch, Selenate reduction by a Pseudomonas species: a new mode of anaerobic respiration. FEMS Microbiol Lett, 1989. 52(1-2): p. 195-8.
    62. Song, B., N.J. Palleroni, and M.M. Haggblom, Description of strain 3CB-1, a genomovar of Thauera aromatica, capable of degrading 3-chlorobenzoate coupled to nitrate reduction. IntJ Syst Evol Microbiol, 2000. 50 Pt 2: p. 551-8.
    63. Foss, S. and J. Harder, Thauera linaloolentis sp. nov. and Thauera terpenica sp. nov., isolated on oxygen-containing monoterpenes (linalool, menthol, and eucalyptol) nitrate. Syst Appl Microbiol, 1998. 21(3): p. 365-73.
    64. Scholten, E., et al., Thauera mechernichensis sp. nov., an aerobic denitrifier from a leachate treatment plant. Int J Syst Bacteriol, 1999. 49 Pt 3: p. 1045-51.
    65. Mechichi, T., et al., Phylogenetic and metabolic diversity of bacteria degrading aromatic compounds under denitrifying conditions, and description of Thauera phenylacetica sp. nov., Thauera aminoaromaticasp. nov., and Azoarcus buckelii sp. nov. Arch Microbiol, 2002. 178(1): p. 26-35.
    66. Dagley, S., Catabolism of aromatic compounds by micro-organisms. Adv Microb Physiol, 1971. 6(0): p. 1-46.
    67. Heider, J. and G. Fuchs, Anaerobic metabolism of aromatic compounds. Eur J Biochem, 1997. 243(3): p. 577-96.
    68. Harwood, C.S., et al., Anaerobic metabolism of aromatic compounds via the benzoyl-CoA pathway. Fems Microbiology Reviews, 1998. 22(5): p. 439-458.
    69. Boll, M. and G. Fuchs, Benzoyl-coenzyme A reductase (dearomatizing), a key enzyme of anaerobic aromatic metabolism. ATP dependence of the reaction, purification and some properties of the enzyme from Thauera aromatica strain K172. Eur J Biochem, 1995. 234(3): p. 921-33.
    70. Boll, M. and G. Fuchs, Identification and characterization of the natural electron donor ferredoxin and of FAD as a possible prosthetic group of benzoyl-CoA reductase (dearomatizing), a key enzyme of anaerobic aromatic metabolism. Eur J Biochem, 1998. 251(3): p. 946-54.
    71. Laempe, D., et al., Cyclohexa-1,5-diene-1-carbonyl-CoA hydratase [corrected], an enzyme involved in anaerobic metabolism of benzoyl-CoA in the denitrifying bacterium Thauera aromatica. Eur J Biochem, 1998. 255(3): p. 618-27.
    72. Hartel, U., et al., Purification of glutaryl-CoA dehydrogenase from Pseudomonas sp., an enzyme involved in the anaerobic degradation of benzoate. Arch Microbiol, 1993. 159(2): p. 174-81.
    73. Breinig, S., E. Schiltz, and G. Fuchs, Genes involved in anaerobic metabolism of phenol in the bacterium Thauera aromatica. J Bacteriol, 2000. 182(20): p. 5849-63.
    74. Schmeling, S., et al., Phenylphosphate synthase: a new phosphotransferase catalyzing the first step in anaerobic phenol metabolism in Thauera aromatica. J Bacteriol, 2004. 186(23): p. 8044-57.
    75. Narmandakh, A., et al., Phosphorylation of phenol by phenylphosphate synthase: role of histidine phosphate in catalysis. J Bacteriol, 2006. 188(22): p. 7815-22.
    76. Hino, S., K. Watanabe, and N. Takahashi, Phenol hydroxylase cloned from Ralstonia eutropha strain E2 exhibits novel kinetic properties. Microbiology, 1998. 144 ( Pt 7): p. 1765-72.
    77. Smith, M.R., The biodegradation of aromatic hydrocarbons by bacteria. Biodegradation, 1990. 1(2-3): p. 191-206.
    78. Watanabe, K., H. Futamata, and S. Harayama, Understanding the diversity in catabolic potential of microorganisms for the development of bioremediation strategies. Antonie Van Leeuwenhoek, 2002. 81(1-4): p. 655-63.
    79. Zhang, X., et al., Microdiversity of phenol hydroxylase genes among phenol-degrading isolates of Alcaligenes sp. from an activated sludge system. FEMS Microbiol Lett, 2004. 237(2): p. 369-75.
    80. Baek, S.H. and J.P. Shapleigh, Expression of nitrite and nitric oxide reductases in free-living and plant-associated Agrobacterium tumefaciens C58 cells. Appl Environ Microbiol, 2005. 71(8): p. 4427-36.
    81. Bergaust, L., et al., Transcription and activities of NO(x) reductases in Agrobacterium tumefaciens: the influence of nitrate, nitrite and oxygen availability. Environ Microbiol, 2008.
    82. Dickinson, R.E. and R.J. Cicerone, Future global warming from atmospheric trace gases. Nature, 1986. 319: p. 109-115.
    83. IPCC, Climate change, the scientific basis. Cambridge University Press, Cambridge, United Kingdom.
    84. Kim, K.R. and H. Craig, Nitrogen-15 and Oxygen-18 Characteristics of Nitrous Oxide: A Global Perspective. Science, 1993. 262(5141): p. 1855-1857.
    85. Otte, S., et al., Nitrous oxide production by Alcaligenes faecalis under transient and dynamic aerobic and anaerobic conditions. Appl Environ Microbiol, 1996. 62(7): p. 2421-6.
    86. Tavares, P., et al., Metalloenzymes of the denitrification pathway. J Inorg Biochem, 2006. 100(12): p. 2087-100.
    87. Zumft, W.G., Cell biology and molecular basis of denitrification. Microbiol Mol Biol Rev, 1997. 61(4): p. 533-616.
    88. Kobayashi, M., et al., Denitrification, a novel type of respiratory metabolism in fungal mitochondrion. J Biol Chem, 1996. 271(27): p. 16263-7.
    89. Richardson, D. and G. Sawers, Structural biology. PMF through the redox loop. Science, 2002. 295(5561): p. 1842-3.
    90. Dodd, F.E., et al., X-ray structure of a blue-copper nitrite reductase in two crystal forms. The nature of the copper sites, mode of substrate binding and recognition by redox partner. J Mol Biol, 1998. 282(2): p. 369-82.
    91. Godden, J.W., et al., The 2.3 angstrom X-ray structure of nitrite reductase from Achromobacter cycloclastes. Science, 1991. 253(5018): p. 438-42.
    92. Williams, P.A., et al., Haem-ligand switching during catalysis in crystals of a nitrogen-cycle enzyme. Nature, 1997. 389(6649): p. 406-12.
    93. Fulop, V., et al., The anatomy of a bifunctional enzyme: structural basis for reduction of oxygen to water and synthesis of nitric oxide by cytochrome cd1. Cell, 1995. 81(3): p. 369-77.
    94. de Vries, S., et al., Purification and characterization of the MQH2:NO oxidoreductase fromthe hyperthermophilic archaeon Pyrobaculum aerophilum. J Biol Chem, 2003. 278(38): p. 35861-8.
    95. Cramm, R., A. Pohlmann, and B. Friedrich, Purification and characterization of the single-component nitric oxide reductase from Ralstonia eutropha H16. FEBS Lett, 1999. 460(1): p. 6-10.
    96. Hendriks, J., et al., Nitric oxide reductases in bacteria. Biochim Biophys Acta, 2000. 1459(2-3): p. 266-73.
    97. Householder, T.C., et al., Gonococcal nitric oxide reductase is encoded by a single gene, norB, which is required for anaerobic growth and is induced by nitric oxide. Infect Immun, 2000. 68(9): p. 5241-6.
    98. Suharti, H.A. Heering, and S. de Vries, NO reductase from Bacillus azotoformans is a bifunctional enzyme accepting electrons from menaquinol and a specific endogenous membrane-bound cytochrome c551. Biochemistry, 2004. 43(42): p. 13487-95.
    99. Simon, J., et al., The unprecedented nos gene cluster of Wolinella succinogenes encodes a novel respiratory electron transfer pathway to cytochrome c nitrous oxide reductase. FEBS Lett, 2004. 569(1-3): p. 7-12.
    100. Teraguchi, S. and T.C. Hollocher, Purification and some characteristics of a cytochrome c-containing nitrous oxide reductase from Wolinella succinogenes. J Biol Chem, 1989. 264(4): p. 1972-9.
    101. Krul, J.M., Dissimilatory nitrate and nitrite reduction under aerobic conditions by an aerobically and anaerobically grown Alcaligenes sp. and by activated sludge. J Appl Bacteriol, 1976. 40(3): p. 245-60.
    102. Lukow, T. and H. Diekmann, Aerobic denitrification by a newly isolated heterotrophic bacterium strain TL1. Biotechnology Letters, 1997. 19(11): p. 1157-1159.
    103. Etchebehere, C. and J. Tiedje, Presence of two different active nirS nitrite reductase genes in a denitrifying Thauera sp. from a high-nitrate-removal-rate reactor. Appl Environ Microbiol, 2005. 71(9): p. 5642-5.
    1. Yuan, Z. and L.L. Blackall, Sludge population optimisation: a new dimension for the control of biological wastewater treatment systems. Water Res, 2002. 36(2): p. 482-90.
    2. Sakano, Y., et al., Spatial distribution of total, ammonia-oxidizing, and denitrifying bacteria in biological wastewater treatment reactors for bioregenerative life support. Appl Environ Microbiol, 2002. 68(5): p. 2285-93.
    3. Schramm, A., et al., Structure and function of a nitrifying biofilm as determined by in situ hybridization and the use of microelectrodes. Appl Environ Microbiol, 1996. 62(12): p. 4641-7.
    4. Kortstee, G.J., et al., Biology of polyphosphate-accumulating bacteria involved in enhanced biological phosphorus removal. FEMS Microbiol Rev, 1994. 15(2-3): p. 137-53.
    5. Liu, B., et al., Thauera and Azoarcus as functionally important genera in a denitrifying quinoline-removal bioreactor as revealed by microbial community structure comparison. FEMS Microbiol Ecol, 2006. 55(2): p. 274-86.
    6. Valle, A., et al., N-acyl-l-homoserine lactones (AHLs) affect microbial community composition and function in activated sludge. Environ Microbiol, 2004. 6(4): p. 424-33.
    7. Schuhle, K. and G. Fuchs, Phenylphosphate carboxylase: a new C-C lyase involved in anaerobic phenol metabolism in Thauera aromatica. J Bacteriol, 2004. 186(14): p. 4556-67.
    8. Dibenedetto, A., et al., Structure-biodegradation correlation of polyphenols for Thauera aromatica in anaerobic conditions. Chemistry and Ecology, 2006. 22: p. 133-143.
    9. Song, B., N.J. Palleroni, and M.M. Haggblom, Isolation and characterization of diverse halobenzoate-degrading denitrifying bacteria from soils and sediments. Appl Environ Microbiol, 2000. 66(8): p. 3446-53.
    10. Shinoda, Y., et al., Aerobic and anaerobic toluene degradation by a newly isolated denitrifying bacterium, Thauera sp. strain DNT-1. Appl Environ Microbiol, 2004. 70(3): p. 1385-92.
    11. Biegert, T., G. Fuchs, and J. Heider, Evidence that anaerobic oxidation of toluene in thedenitrifying bacterium Thauera aromatica is initiated by formation of benzylsuccinate from toluene and fumarate. Eur J Biochem, 1996. 238(3): p. 661-8.
    12. Eschenhagen, M., M. Schuppler, and I. Roske, Molecular characterization of the microbial community structure in two activated sludge systems for the advanced treatment of domestic effluents. Water Res, 2003. 37(13): p. 3224-32.
    13. Loy, A., et al., 16S rRNA gene-based oligonucleotide microarray for environmental monitoring of the betaproteobacterial order "Rhodocyclales". Appl Environ Microbiol, 2005. 71(3): p. 1373-86.
    14. Kowalchuk, G.A., et al., Analysis of ammonia-oxidizing bacteria of the beta subdivision of the class Proteobacteria in coastal sand dunes by denaturing gradient gel electrophoresis and sequencing of PCR-amplified 16S ribosomal DNA fragments. Appl Environ Microbiol, 1997. 63(4): p. 1489-97.
    15. Kowalchuk, G.A., et al., Molecular analysis of ammonia-oxidizing bacteria of the beta subdivision of the class Proteobacteria in compost and composted materials. Appl Environ Microbiol, 1999. 65(2): p. 396-403.
    16. Greenberg AE, C.L.E.A., Standard Methods for the Examination of Water and Wastewater. 18th edn. American Public Health Association, American Water Works Association, and Water Environment Federation, Washington DC., 1992.
    17. Di Cello, F., et al., Biodiversity of a Burkholderia cepacia population isolated from the maize rhizosphere at different plant growth stages. Appl Environ Microbiol, 1997. 63(11): p. 4485-93.
    18. Muyzer, G., E.C. de Waal, and A.G. Uitterlinden, Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol, 1993. 59(3): p. 695-700.
    19. Eckburg, P.B., et al., Diversity of the human intestinal microbial flora. Science, 2005. 308(5728): p. 1635-8.
    20. Dyszynski, G.S., W.M., RDPquery: A Java program from the Sapelo Program MicrobialObservatory for automatic classification of bacterial 16S rRNA sequences based on Ribosomal Database Project taxonomy and Smith-Waterman alignment. 2006.
    21. Ludwig, W., et al., ARB: a software environment for sequence data. Nucleic Acids Res, 2004. 32(4): p. 1363-71.
    22. Schloss, P.D. and J. Handelsman, Introducing DOTUR, a computer program for defining operational taxonomic units and estimating species richness. Appl Environ Microbiol, 2005. 71(3): p. 1501-6.
    23. Good, I.J., The population frequencies of species and the estimation of population parameters. Biometrika, 1953. 40: p. 237 - 264.
    24. Mechichi, T., et al., Phylogenetic and metabolic diversity of bacteria degrading aromatic compounds under denitrifying conditions, and description of Thauera phenylacetica sp. nov., Thauera aminoaromaticasp. nov., and Azoarcus buckelii sp. nov. Arch Microbiol, 2002. 178(1): p. 26-35.
    25. Heylen, K., et al., The incidence of nirS and nirK and their genetic heterogeneity in cultivated denitrifiers. Environmental Microbiology, 2006. 8(11): p. 2012-2021.
    26. Thomsen, T.R., Y. Kong, and P.H. Nielsen, Ecophysiology of abundant denitrifying bacteria in activated sludge. FEMS Microbiol Ecol, 2007. 60(3): p. 370-82.
    27. Yamada, S., et al., Cloning and nucleotide sequence analysis of gyrB of Bacillus cereus, B. thuringiensis, B. mycoides, and B. anthracis and their application to the detection of B. cereus in rice. Appl Environ Microbiol, 1999. 65(4): p. 1483-90.
    28. De Clerck, E., et al., Isolation, characterization, and identification of bacterial contaminants in semifinal gelatin extracts. Appl Environ Microbiol, 2004. 70(6): p. 3664-72.
    29. Shen, J., et al., Molecular profiling of the Clostridium leptum subgroup in human fecal microflora by PCR-denaturing gradient gel electrophoresis and clone library analysis. Appl Environ Microbiol, 2006. 72(8): p. 5232-8.
    30. da Silva, K.R., et al., Application of a novel Paenibacillus-specific PCR-DGGE method and sequence analysis to assess the diversity of Paenibacillus spp. in the maize rhizosphere. JMicrobiol Methods, 2003. 54(2): p. 213-31.
    31. Garbeva, P., J.A. van Veen, and J.D. van Elsas, Predominant Bacillus spp. in agricultural soil under different management regimes detected via PCR-DGGE. Microb Ecol, 2003. 45(3): p. 302-16.
    32. Calvo, L., et al., Use of the ammonia-oxidizing bacterial-specific phylogenetic probe Nso1225 as a primer for fingerprint analysis of ammonia-oxidizer communities. Appl Microbiol Biotechnol, 2004. 63(6): p. 715-21.
    33. Freitag, T.E. and J.I. Prosser, Community structure of ammonia-oxidizing bacteria within anoxic marine sediments. Appl Environ Microbiol, 2003. 69(3): p. 1359-71.
    34. Cook, K.L., et al., Effect of microbial species richness on community stability and community function in a model plant-based wastewater processing system. Microb Ecol, 2006. 52(4): p. 725-37.
    35. Wagner, M., et al., Microbial community composition and function in wastewater treatment plants. Antonie Van Leeuwenhoek, 2002. 81(1-4): p. 665-80.
    36. Forney, L.J., et al., Structure of microbial communities in activated sludge: Potential implications for assessing the biodegradability of chemicals. Ecotoxicology and Environmental Safety, 2001. 49(1): p. 40-53.
    37. LaPara, T.M. and S. Ghosh, Population dynamics of the ammonia-oxidizing bacteria in a full-scale municipal wastewater treatment facility. Environmental Engineering Science, 2006. 23(2): p. 309-319.
    38. LaPara, T.M., et al., Effects of elevated temperature on bacterial community structure and function in bioreactors treating a synthetic wastewater. Journal of Industrial Microbiology & Biotechnology, 2000. 24(2): p. 140-145.
    39. Ayala-Del-Rio, H.L., et al., Correspondence between community structure and function during succession in phenol- and phenol-plus-trichloroethene-fed sequencing batch reactors. Appl Environ Microbiol, 2004. 70(8): p. 4950-60.
    40. Fernandez, A., et al., How stable is stable? Function versus community composition. ApplEnviron Microbiol, 1999. 65(8): p. 3697-704.
    41. Fernandez, A.S., et al., Flexible community structure correlates with stable community function in methanogenic bioreactor communities perturbed by glucose. Appl Environ Microbiol, 2000. 66(9): p. 4058-67.
    42. Franklin, R.B. and A.L. Mills, Structural and functional responses of a sewage microbial community to dilution-induced reductions in diversity. Microb Ecol, 2006. 52(2): p. 280-8.
    43. Hashsham, S.A., et al., Parallel processing of substrate correlates with greater functional stability in methanogenic bioreactor communities perturbed by glucose. Appl Environ Microbiol, 2000. 66(9): p. 4050-7.
    44. LaPara, T.M., et al., Stability of the bacterial communities supported by a seven-stage biological process treating pharmaceutical wastewater as revealed by PCR-DGGE. Water Research, 2002. 36(3): p. 638-646.
    45. Hartmann, M. and F. Widmer, Community structure analyses are more sensitive to differences in soil bacterial communities than anonymous diversity indices. Applied and Environmental Microbiology, 2006. 72(12): p. 7804-7812.
    1. Zhang M, Tay JH, Qian Y, et al. Coke plant wastewater treatment by fixed biofilm system for COD and NH3-N removal. Water Research, 1998. 32(2): p. 519-527.
    2. Chakraborty S, Veeramani H. Response of pulse phenol injection on an anaerobic-anoxic-aerobic system. Bioresource Technology, 2005. 96(7): p. 761-767.
    3. Liu BB, Zhang F, Feng XX, et al. Thauera and Azoarcus as functionally important genera in a denitrifying quinoline-removal bioreactor as revealed by microbial community structure comparison. Fems Microbiology Ecology, 2006. 55(2): p. 274-286.
    4. Thomsen TR, Kong Y, Nielsen PH. Ecophysiology of abundant denitrifying bacteria in activated sludge. Fems Microbiology Ecology, 2007. 60(3): p. 370-382.
    5. Valle A, Bailey MJ, Whiteley AS, et al. N-acyl-L-homoserine lactones (AHLs) affect microbial community composition and function in activated sludge. Environmental Microbiology, 2004. 6(4): p. 424-433.
    6. Dibenedetto A, Lo Noce RM, Narracci M, et al. Structure-biodegradation correlation of polyphenols for Thauera aromatica in anaerobic conditions. Chemistry and Ecology, 2006. 22: p. 133-143.
    7. Schuhle K, Fuchs G. Phenylphosphate carboxylase: a new C-C lyase involved in anaerobic phenol metabolism in Thauera aromatica. J Bacteriol, 2004. 186(14): p. 4556-67.
    8. Shinoda Y, Sakai Y, Uenishi H, et al. Aerobic and anaerobic toluene degradation by a newly isolated denitrifying bacterium, Thauera sp. strain DNT-1. Appl Environ Microbiol, 2004. 70(3): p. 1385-92.
    9. Zoetendal EG, Akkermans ADL, De Vos WM. Temperature gradient gel electrophoresis analysis of 16S rRNA from human fecal samples reveals stable and host-specific communities of active bacteria. Appl Environ Microbiol, 1998. 64(10): p. 3854-3859.
    10. Mao YJ, Zhang XJ, Yan X, et al. Development of group-specific PCR-DGGE fingerprinting for monitoring structural changes of Thauera spp. in an industrial wastewater treatmentplant responding to operational perturbations. J Microbiol Methods, 2008, DOI:
    10.1016/j.mimet.2008.06.005
    11. Di Cello F, Bevivino A, Chiarini L, et al. Biodiversity of a Burkholderia cepacia population isolated from the maize rhizosphere at different plant growth stages. Appl Environ Microbiol, 1997. 63(11): p. 4485-93.
    12. Loy A, Schulz C, Lucker S, et al. 16S rRNA gene-based oligonucleotide microarray for environmental monitoring of the betaproteobacterial order "Rhodocyclales". Appl Environ Microbiol, 2005. 71(3): p. 1373-86.
    13. Muyzer G, de Waal EC, Uitterlinden AG. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol, 1993. 59(3): p. 695-700.
    14. Costerton JW, Lewandowski Z, DeBeer D, et al. Biofilms, the customized microniche. J Bacteriol, 1994. 176(8): p. 2137-42.
    15. Amann RI, Ludwig W, Schleifer KH. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol Rev, 1995. 59(1): p. 143-69.
    16. Ferguson RL, Buckley EN, Palumbo AV. Response of marine bacterioplankton to differential filtration and confinement. Appl Environ Microbiol, 1984. 47(1): p. 49-55.
    17. Kogure K, Simidu U, Taga N. A tentative direct microscopic method for counting living marine bacteria. Can J Microbiol, 1979. 25(3): p. 415-20.
    18. Kogure K, Simidu U, Taga N. Distribution of viable marine bacteria in neritic seawater around Japan. Can J Microbiol, 1980. 26(3): p. 318-23.
    19. Jones J. The effect of environmental factors on estimated viable and total populations of planktonic bacteria in lakes and experimental enclosures. Freshwater Biology, 1977. 7(1): p. 69-71.
    20. Torsvik V, Goksoyr J, Daae FL. High diversity in DNA of soil bacteria. Appl Environ Microbiol, 1990. 56(3): p. 782-7.
    21. Wagner M, Amann R, Lemmer H, et al. Probing activated sludge with oligonucleotidesspecific for proteobacteria: inadequacy of culture-dependent methods for describing microbial community structure. Appl Environ Microbiol, 1993. 59(5): p. 1520-5.
    22. Wagner M, Erhart R, Manz W, et al. Development of an rRNA-targeted oligonucleotide probe specific for the genus Acinetobacter and its application for in situ monitoring in activated sludge. Appl Environ Microbiol, 1994. 60(3): p. 792-800.
    23. Kaeberlein T, Lewis K, Epstein SS. Isolating "uncultivable" microorganisms in pure culture in a simulated natural environment. Science, 2002. 296(5570): p. 1127-9.
    24. Ash C, Farrow JA, Dorsch M, et al. Comparative analysis of Bacillus anthracis, Bacillus cereus, and related species on the basis of reverse transcriptase sequencing of 16S rRNA. Int J Syst Bacteriol, 1991. 41(3): p. 343-6.
    25. Blattner FR, Plunkett G, Bloch CA, et al. The complete genome sequence of Escherichia coli K-12. Science, 1997. 277(5331): p. 1453-74.
    26. Jaspers E, Overmann J. Ecological significance of microdiversity: identical 16S rRNA gene sequences can be found in bacteria with highly divergent genomes and ecophysiologies. Appl Environ Microbiol, 2004. 70(8): p. 4831-9.
    27. Lehtimaki J, Lyra C, Suomalainen S, et al. Characterization of Nodularia strains, cyanobacteria from brackish waters, by genotypic and phenotypic methods. Int J Syst Evol Microbiol, 2000. 50 Pt 3: p. 1043-53.
    28. Perna NT, Plunkett G, Burland V, et al. Genome sequence of enterohaemorrhagic Escherichia coli O157:H7. Nature, 2001. 409(6819): p. 529-33.
    29. Wagner M, Loy A, Nogueira R, et al. Microbial community composition and function in wastewater treatment plants. Antonie Van Leeuwenhoek, 2002. 81(1-4): p. 665-80.
    30. Heylen K, Gevers D, Vanparys B, et al. The incidence of nirS and nirK and their genetic heterogeneity in cultivated denitrifiers. Environmental Microbiology, 2006. 8(11): p. 2012-2021.
    31. Mechichi T, Stackebrandt E, Gad'on N, et al. Phylogenetic and metabolic diversity of bacteria degrading aromatic compounds under denitrifying conditions, and description ofThauera phenylacetica sp. nov., Thauera aminoaromatica sp. nov., and Azoarcus buckelii sp. nov. Arch Microbiol, 2002. 178(1): p. 26-35.
    1. Liu, B., et al., Thauera and Azoarcus as functionally important genera in a denitrifying quinoline-removal bioreactor as revealed by microbial community structure comparison. FEMS Microbiol Ecol, 2006. 55(2): p. 274-86.
    2. Valle, A., et al., N-acyl-l-homoserine lactones (AHLs) affect microbial community composition and function in activated sludge. Environ Microbiol, 2004. 6(4): p. 424-33.
    3. Manefield, M., et al., RNA stable isotope probing, a novel means of linking microbial community function to phylogeny. Appl Environ Microbiol, 2002. 68(11): p. 5367-73.
    4. Song, B., N.J. Palleroni, and M.M. Haggblom, Description of strain 3CB-1, a genomovar of Thauera aromatica, capable of degrading 3-chlorobenzoate coupled to nitrate reduction. Int J Syst Evol Microbiol, 2000. 50 Pt 2: p. 551-8.
    5. Anders, H.J., et al., Taxonomic position of aromatic-degrading denitrifying pseudomonad strains K 172 and KB 740 and their description as new members of the genera Thauera, as Thauera aromatica sp. nov., and Azoarcus, as Azoarcus evansii sp. nov., respectively, members of the beta subclass of the Proteobacteria. Int J Syst Bacteriol, 1995. 45(2): p. 327-33.
    6. Shinoda, Y., et al., Aerobic and anaerobic toluene degradation by a newly isolated denitrifying bacterium, Thauera sp. strain DNT-1. Appl Environ Microbiol, 2004. 70(3): p. 1385-92.
    7. Mechichi, T., et al., Phylogenetic and metabolic diversity of bacteria degrading aromatic compounds under denitrifying conditions, and description of Thauera phenylacetica sp. nov., Thauera aminoaromaticasp. nov., and Azoarcus buckelii sp. nov. Arch Microbiol, 2002. 178(1): p. 26-35.
    8. Harwood, C.S., et al., Anaerobic metabolism of aromatic compounds via the benzoyl-CoA pathway. Fems Microbiology Reviews, 1998. 22(5): p. 439-458.
    9. Ash, C., et al., Comparative analysis of Bacillus anthracis, Bacillus cereus, and relatedspecies on the basis of reverse transcriptase sequencing of 16S rRNA. Int J Syst Bacteriol, 1991. 41(3): p. 343-6.
    10. Blattner, F.R., et al., The complete genome sequence of Escherichia coli K-12. Science, 1997. 277(5331): p. 1453-74.
    11. Jaspers, E. and J. Overmann, Ecological significance of microdiversity: identical 16S rRNA gene sequences can be found in bacteria with highly divergent genomes and ecophysiologies. Appl Environ Microbiol, 2004. 70(8): p. 4831-9.
    12. Lehtimaki, J., et al., Characterization of Nodularia strains, cyanobacteria from brackish waters, by genotypic and phenotypic methods. Int J Syst Evol Microbiol, 2000. 50 Pt 3: p. 1043-53.
    13. Perna, N.T., et al., Genome sequence of enterohaemorrhagic Escherichia coli O157:H7. Nature, 2001. 409(6819): p. 529-33.
    14. Zoetendal, E.G., A.D.L. Akkermans, and W.M. De Vos, Temperature gradient gel electrophoresis analysis of 16S rRNA from human fecal samples reveals stable and host-specific communities of active bacteria. Applied and Environmental Microbiology, 1998. 64(10): p. 3854-3859.
    15. Gillings, M. and M. Holley, Repetitive element PCR fingerprinting (rep-PCR) using enterobacterial repetitive intergenic consensus (ERIC) primers is not necessarily directed at ERIC elements. Letters in Applied Microbiology, 1997. 25(1): p. 17-21.
    16. Braker, G., A. Fesefeldt, and K.P. Witzel, Development of PCR primer systems for amplification of nitrite reductase genes (nirK and nirS) to detect denitrifying bacteria in environmental samples. Appl Environ Microbiol, 1998. 64(10): p. 3769-75.
    17. Futamata, H., S. Harayama, and K. Watanabe, Group-specific monitoring of phenol hydroxylase genes for a functional assessment of phenol-stimulated trichloroethylene bioremediation. Appl Environ Microbiol, 2001. 67(10): p. 4671-7.
    18. Zhang, M., et al., Coke plant wastewater treatment by fixed biofilm system for COD and NH3-N removal. Water Research, 1998. 32(2): p. 519-527.
    19. Etchebehere, C. and J. Tiedje, Presence of two different active nirS nitrite reductase genes in a denitrifying Thauera sp. from a high-nitrate-removal-rate reactor. Appl Environ Microbiol, 2005. 71(9): p. 5642-5.
    20. Zhang, X., et al., Microdiversity of phenol hydroxylase genes among phenol-degrading isolates of Alcaligenes sp. from an activated sludge system. FEMS Microbiol Lett, 2004. 237(2): p. 369-75.
    21. Moore, L.R., G. Rocap, and S.W. Chisholm, Physiology and molecular phylogeny of coexisting Prochlorococcus ecotypes. Nature, 1998. 393(6684): p. 464-7.
    22. Beja, O., et al., Comparative genomic analysis of archaeal genotypic variants in a single population and in two different oceanic provinces. Appl Environ Microbiol, 2002. 68(1): p. 335-45.
    23. Watanabe, K., H. Futamata, and S. Harayama, Understanding the diversity in catabolic potential of microorganisms for the development of bioremediation strategies. Antonie Van Leeuwenhoek, 2002. 81(1-4): p. 655-63.
    24. Watanabe, K., et al., Molecular detection, isolation, and physiological characterization of functionally dominant phenol-degrading bacteria in activated sludge. Appl Environ Microbiol, 1998. 64(11): p. 4396-402.
    25. Foss, S. and J. Harder, Thauera linaloolentis sp. nov. and Thauera terpenica sp. nov., isolated on oxygen-containing monoterpenes (linalool, menthol, and eucalyptol) nitrate. Syst Appl Microbiol, 1998. 21(3): p. 365-73.
    26. Scholten, E., et al., Thauera mechernichensis sp. nov., an aerobic denitrifier from a leachate treatment plant. Int J Syst Bacteriol, 1999. 49 Pt 3: p. 1045-51.
    27. Macy, J.M., et al., Thauera selenatis gen. nov., sp. nov., a member of the beta subclass of Proteobacteria with a novel type of anaerobic respiration. Int J Syst Bacteriol, 1993. 43(1): p. 135-42.
    1. Mechichi, T., et al., Phylogenetic and metabolic diversity of bacteria degrading aromatic compounds under denitrifying conditions, and description of Thauera phenylacetica sp. nov., Thauera aminoaromaticasp. nov., and Azoarcus buckelii sp. nov. Arch Microbiol, 2002. 178(1): p. 26-35.
    2. Foss, S. and J. Harder, Thauera linaloolentis sp. nov. and Thauera terpenica sp. nov., isolated on oxygen-containing monoterpenes (linalool, menthol, and eucalyptol) nitrate. Syst Appl Microbiol, 1998. 21(3): p. 365-73.
    3. Etchebehere, C. and J. Tiedje, Presence of two different active nirS nitrite reductase genes in a denitrifying Thauera sp. from a high-nitrate-removal-rate reactor. Appl Environ Microbiol, 2005. 71(9): p. 5642-5.
    4. Anders, H.J., et al., Taxonomic position of aromatic-degrading denitrifying pseudomonad strains K 172 and KB 740 and their description as new members of the genera Thauera, as Thauera aromatica sp. nov., and Azoarcus, as Azoarcus evansii sp. nov., respectively, members of the beta subclass of the Proteobacteria. Int J Syst Bacteriol, 1995. 45(2): p. 327-33.
    5. Shinoda, Y., et al., Aerobic and anaerobic toluene degradation by a newly isolated denitrifying bacterium, Thauera sp. strain DNT-1. Appl Environ Microbiol, 2004. 70(3): p. 1385-92.
    6. Macy, J.M., et al., Thauera selenatis gen. nov., sp. nov., a member of the beta subclass of Proteobacteria with a novel type of anaerobic respiration. Int J Syst Bacteriol, 1993. 43(1): p. 135-42.
    7. Tschech, A. and G. Fuchs, Anaerobic degradation of phenol by pure cultures of newly isolated denitrifying pseudomonads. Arch Microbiol, 1987. 148(3): p. 213-7.
    8. Liu, B., et al., Thauera and Azoarcus as functionally important genera in a denitrifying quinoline-removal bioreactor as revealed by microbial community structure comparison.FEMS Microbiol Ecol, 2006. 55(2): p. 274-86.
    9. Mao, Y., et al., Development of group-specific PCR-DGGE fingerprinting for monitoring structural changes of Thauera spp. in an industrial wastewater treatment plant responding to operational perturbations. J.Microbiol. Methods, 2008. doi:10.1016/j.mimet.2008.06.005.
    10. Lukow, T. and H. Diekmann, Aerobic denitrification by a newly isolated heterotrophic bacterium strain TL1. Biotechnology Letters, 1997. 19(11): p. 1157-1159.
    11. Baek, S.H. and J.P. Shapleigh, Expression of nitrite and nitric oxide reductases in free-living and plant-associated Agrobacterium tumefaciens C58 cells. Appl Environ Microbiol, 2005. 71(8): p. 4427-36.
    12. Bergaust, L., et al., Transcription and activities of NO(x) reductases in Agrobacterium tumefaciens: the influence of nitrate, nitrite and oxygen availability. Environ Microbiol, 2008.
    13. Dandie, C.E., et al., Analysis of denitrification genes and comparison of nosZ, cnorB and 16S rDNA from culturable denitrifying bacteria in potato cropping systems. Syst Appl Microbiol, 2007. 30(2): p. 128-38.
    14. Delorme, S., et al., Comparative genetic diversity of the narG, nosZ, and 16S rRNA genes in fluorescent pseudomonads. Appl Environ Microbiol, 2003. 69(2): p. 1004-12.
    15. Allison, C. and G.T. Macfarlane, Dissimilatory nitrate reduction by Propionibacterium acnes. Appl Environ Microbiol, 1989. 55(11): p. 2899-903.
    16. Scholten, E., et al., Thauera mechernichensis sp. nov., an aerobic denitrifier from a leachate treatment plant. Int J Syst Bacteriol, 1999. 49 Pt 3: p. 1045-51.
    17. Baker, S.C., et al., Molecular genetics of the genus Paracoccus: metabolically versatile bacteria with bioenergetic flexibility. Microbiol Mol Biol Rev, 1998. 62(4): p. 1046-78.
    18. Baumann, B., et al., Inhibition of denitrification activity but not of mRNA induction in Paracoccus denitrificans by nitrite at a suboptimal pH. Antonie Van Leeuwenhoek, 1997. 72(3): p. 183-9.
    19. Molstad, L., P. Dorsch, and L.R. Bakken, Robotized incubation system for monitoring gases(O(2), NO, N(2)O N(2)) in denitrifying cultures. J Microbiol Methods, 2007.
    20. Di Cello, F., et al., Biodiversity of a Burkholderia cepacia population isolated from the maize rhizosphere at different plant growth stages. Appl Environ Microbiol, 1997. 63(11): p. 4485-93.
    21. Eckburg, P.B., et al., Diversity of the human intestinal microbial flora. Science, 2005. 308(5728): p. 1635-8.
    22. Braker, G., A. Fesefeldt, and K.P. Witzel, Development of PCR primer systems for amplification of nitrite reductase genes (nirK and nirS) to detect denitrifying bacteria in environmental samples. Appl Environ Microbiol, 1998. 64(10): p. 3769-75.
    23. Throback, I.N., et al., Reassessing PCR primers targeting nirS, nirK and nosZ genes for community surveys of denitrifying bacteria with DGGE. Fems Microbiology Ecology, 2004. 49(3): p. 401-417.
    24. Kloos, K., et al., Denitrification within the genus Azospirillum and other associative bacteria. Australian Journal of Plant Physiology, 2001. 28(9): p. 991-998.
    25. Thompson, J.D., et al., The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res, 1997. 25(24): p. 4876-82.
    26. Kumar, S., et al., MEGA2: molecular evolutionary genetics analysis software. Bioinformatics, 2001. 17(12): p. 1244-5.
    27. Song, B.K. and B.B. Ward, Nitrite reductase genes in halobenzoate degrading denitrifying bacteria. Fems Microbiology Ecology, 2003. 43(3): p. 349-357.
    28. Scala, D.J. and L.J. Kerkhof, Nitrous oxide reductase (nosZ) gene-specific PCR primers for detection of denitrifiers and three nosZ genes from marine sediments. Fems Microbiology Letters, 1998. 162(1): p. 61-68.
    29. Nogales, B., et al., Detection and diversity of expressed denitrification genes in estuarine sediments after reverse transcription-PCR amplification from mRNA. Applied and Environmental Microbiology, 2002. 68(10): p. 5017-5025.
    30. Baranyi, J. and T.A. Roberts, A dynamic approach to predicting bacterial growth in food. Int J Food Microbiol, 1994. 23(3-4): p. 277-94.
    31. Goregues, C.M., V.D. Michotey, and P.C. Bonin, Molecular, biochemical, and physiological approaches for understanding the ecology of denitrification. Microbial Ecology, 2005. 49(2): p. 198-208.
    32. Thomas, K.L., D. Lloyd, and L. Boddy, Effects of oxygen, pH and nitrate concentration on denitrification by Pseudomonas species. FEMS Microbiol Lett, 1994. 118(1-2): p. 181-6.
    33. Korner, H. and W.G. Zumft, Expression of denitrification enzymes in response to the dissolved oxygen level and respiratory substrate in continuous culture of Pseudomonas stutzeri. Appl Environ Microbiol, 1989. 55(7): p. 1670-6.
    34. Bell, L.C. and S.J. Ferguson, Nitric and nitrous oxide reductases are active under aerobic conditions in cells of Thiosphaera pantotropha. Biochem J, 1991. 273(Pt 2): p. 423-7.
    35. Baumann, B., et al., Dynamics of denitrification activity of Paracoccus denitrificans in continuous culture during aerobic-anaerobic changes. J Bacteriol, 1996. 178(15): p. 4367-74.
    36. Rasmussen, T., Berks, B.C., Butt, J.N., and Thomson, A.J. (2002) Multiple forms of the catalytic centre, CuZ, in the enzyme nitrous oxide reductase from Paracoccus pantotrophus. Biochem J 364: 807-815.
    37. Coyle, C.L., Zumft, W.G., Kroneck, P.M., Korner, H., and Jakob, W. (1985) Nitrous oxide reductase from denitrifying Pseudomonas perfectomarina. Purification and properties of a novel multicopper enzyme. Eur J Biochem 153: 459-467.

© 2004-2018 中国地质图书馆版权所有 京ICP备05064691号 京公网安备11010802017129号

地址:北京市海淀区学院路29号 邮编:100083

电话:办公室:(+86 10)66554848;文献借阅、咨询服务、科技查新:66554700