Embed Size (px)
Citation preview
A
mpscfbPoFwp©
K
1
ias
Cof
y
1d
Biochemical Engineering Journal 39 (2008) 288–296
Changes of catabolic genes and microbial community structuresduring biodegradation of nonylphenol ethoxylates
and nonylphenol in natural water microcosms
Yu Zhang a,c,∗, Kazunari Sei b, Tadashi Toyama b, Michihiko Ike b,Jing Zhang a, Min Yang a,∗∗, Yoichi Kamagata c,d
a State Key Laboratory of Environmental Aquatic Chemistry, Research Center for Eco-Environmental Sciences,Chinese Academy of Sciences, Beijing 100085, China
b Department of Environmental Engineering, Graduate School of Engineering, Osaka University, 2-1 Yamadaoka, Suita, Osaka 565-0871, Japanc Institute for Biological Resources and Functions, National Institute of Advanced Industrial Science and Technology, Tsukuba, Ibaraki 305-8566, Japan
d Research Institute of Genome-based Biofactory, National Institute of Advanced Industrial Science and Technology, Sapporo 062-8517, Japan
Received 14 June 2007; received in revised form 30 August 2007; accepted 29 September 2007
bstract
Changes of possible key catabolic genes and microbial community structures during the degradation of NPEOs and NP in natural watericrocosms were investigated using the most-probable-number-polymerase chain reaction (MPN-PCR) and terminal restriction fragment length
olymorphism (T-RFLP). The copy number of catechol 2,3-dioxygenase (C23O) DNA increased significantly during NPEO and NP degradation,uggesting that meta-cleavage of the aromatic rings of NPEOs and NP might have happened. Catechol 1,2-dioxygenase (C12O) DNA, alkane-atabolic genes (alk), and 16S rDNA, on the other hand, did not change notably, suggesting that the two genes might not be the relevant genesor NPEOs and NP degradation. The 16S rRNA gene-based T-RFLP analysis results indicated that specific and different dominant (or degrading)acteria should be selected, depending on the substances. A strain with a DNA length of 78 bp, which might be affiliated with the beta subclass ofroteobacteria, became the dominant species for NPEO degradation, while strains at 88 and 198 bp were dominant in the NP microcosm. Diversityf microbial community structure tended to be simplified after NPEO degradation, while that in the NP microcosm remained relatively stable.
ive clusters were obtained according to the similarity in community structures of different microcosms by cluster analysis, which were consistentith the biodegradation behaviors of different microcosms. This is the first report on genetic evidence of a possible aromatic ring meta-cleavingathway of NPEOs and NP in an aquatic environment.2007 Elsevier B.V. All rights reserved.
Nony
art
eywords: Functional genes; Microbial community; Nonylphenol ethoxylates;
. Introduction
The environmental fate of nonylphenol ethoxylates (NPEOs),
mportant nonionic industrial surfactants, has received widettention because some of their biodegradation intermediates,uch as nonylphenol (NP), act as the mimic hormones to∗ Corresponding author at: State Key Laboratory of Environmental Aquatichemistry, Research Center for Eco-Environmental Sciences, Chinese Academyf Sciences, Beijing 100085, China. Tel.: +86 10 62923475;ax: +86 10 62923541.∗∗ Corresponding author. Tel.: +86 10 62923475; fax: +86 10 62923541.
E-mail addresses: [email protected] (Y. Zhang),[email protected] (M. Yang).
Eq
at(lcdbl
369-703X/$ – see front matter © 2007 Elsevier B.V. All rights reserved.oi:10.1016/j.bej.2007.09.015
lphenol; Biodegradation
quatic organisms [1,2]. Residues of NPEOs and NP have beeneported to be ubiquitous in river water, groundwater adjacento contaminated rivers, seawater, and tap water [3–5]. The U.S.nvironmental Protection Agency [6] has released draft wateruality criteria for NP.
NPEOs consist of three parts: an alkyl (C9), a phenyl ring,nd a polyoxyethylene chain (EO chain). It has been acceptedhat the aerobic metabolites of NPEOs are NP carboxylatesNPECs) and short chain NPEOs via the �-oxidation oxyethy-ene chain pathway, and the anaerobic ones are NP and short
hain NPEOs [2,7–9]. In addition to the above main biodegra-ation pathways, the oxidation of alkyl group of NPEOs haseen confirmed by identifying the alkyl-chain-oxidized metabo-ites [10,11]. As for the aromatic ring structure, no reports were
ineering Journal 39 (2008) 288–296 289
arsirwath
ttce(frd4werpoaoh
cOwoat
gpnm(gotitr
2
2
lptc(
Table 1Characteristics of the aquatic samples and conditions of the natural water micro-cosms used in this study (Zuion pond in Osaka University)
Parameter Value
Conductivity (�s cm−1) 10.1Ph 7.21DO (mg l−1) 7.0T (◦C) 18.9Turbidity (mg l−1) 18NO3
− (mg l−1) 0.41NO2
− (mg l−1) 0.014NP
ttca
N1wp53T2Smwo
2
aecmshwbmaw
am(sMu
Y. Zhang et al. / Biochemical Eng
vailable until Naylor et al. [12] verified a pathway of aromaticing cleavage using the radioactive label method. They synthe-ized NP10EO (NPEOs with average 10 EO) with a 14C labeln the aromatic ring and found over 40% of the [14C] aromaticing carbon was converted to 14CO2 under a simulated riverater environment, demonstrating the cleavage of the NPEOs’
romatic rings. However, no genes relevant to the biodegrada-ion of nonylphenol ethoxylates (NPEOs) and nonylphenol (NP)ave been identified.
Many studies have demonstrated that analysis of func-ional genes could play an important role in understandinghe biodegradation pathways of chemicals [13,14]. Alkane-atabolic genes (alk) have been reported to encode the keynzyme responsible for the degradation of alkane compounds<C16) [13]. C12O and C23O genes, which are responsibleor the ortho- and meta-cleavage pathways of aromatic rings,espectively, are two important metabolic genes in the biodegra-ation of aromatic compounds [14]. It has been reported that-n-alkyphenols (4-AP, C1–C5) are degraded via the C23O path-ay [15]. The authors claimed that the C23O pathway may be
xpanded further to accommodate larger side chains due to theelaxed specificities of catabolic enzymes, as well as a regulatoryrotein. It is therefore speculated that there might be a possibilityf the cleavage of aromatic rings for the degradation of NPEOsnd NP with the C23O or/and C12O. However, to date, microbi-logical proof of the aromatic ring cleavage of NPEOs and NPas yet to be provided.
Analysis of population dynamics during biodegradationould also be important to understand biochemical pathways.ne of the rRNA gene-based approaches is T-RFLP analysis,hich allows the rapid identification of ribotypes from a varietyf samples of environmental origin [16]. Due to the sensitivitynd high throughput of this method, it is considered as an idealechnique for comparative community analyses [17].
In this study, we focused on the changes of possible catabolicenes responsible for aromatic and alkane degradation andopulation dynamics during NPEO and NP biodegradation inatural water microcosms. Three kinds of catabolic genes wereonitored by most-probable-number-polymerase chain reaction
MPN-PCR) [18] using primer sets for general detection of theenes encoding C23O, C12O, and alk gene (C < 16). Changesf the microbial community structures were monitored usinghe T-RFLP of PCR-amplified 16S rDNA. The microbiologicalnformation of NPEO and NP biodegradation will give impor-ant clues to understand the environmental fate of NPEOs andelated compounds.
. Materials and methods
.1. Microcosms
Natural water samples were collected from Zuion Pondocated in Osaka University, Osaka, Japan. The source of the
ond is influent from the Yamada River, an urban river runninghrough Suita, Osaka. The collected natural water samples wereooled on ice, and brought back to the laboratory immediatelywithin 10 min). The samples were filtered using qualitative fil-(0am
H4+ (mg l−1) 0.41
O43− (mg l−1) 0.012
er paper (No. 2, pore size 5 mm, Advantec, Tokyo, Japan) inhe laboratory. The characteristics of the aquatic samples andonditions of the natural water microcosms used in this studyre shown in Table 1.
The river die-away method was used for biodegradation tests.P10EO (NPEO mixture with an average EO chain number of0) and 4-NP (Tokyo Chemical Industry Co. Ltd., Tokyo, Japan)ere respectively spiked in 400 ml of filtered natural water sam-les in 500 ml flasks. The final concentrations of NP10EO weremg l−1 (NPEO-5, 7.58 �mol l−1), and 25 mg l−1 (NPEO-25,7.9 �mol l−1), and that for NP was 5 mg l−1 (22.8 �mol l−1).hese microcosms were incubated together with a control for5 days at 28 ◦C on a rotary shaker set at 120 rpm in the dark.amples were taken every other day for chemical analyses andicrobial community structure analyses. All of the microcosmsere performed in triplicate. The data are shown as the averagef the triplicate trials.
.2. Determination of concentrations of NPEOs and NP
Aliquots (20 ml) of samples were taken from the microcosmsnd filtered (mixed cellulose ester filter, pore size 0.2 �m, diam-ter 25 mm, Advantec, Tokyo, Japan). NPEOs and the relatedhemicals (NP and NPECs) were analyzed without further treat-ent. To concentrate other metabolites, 10 ml portions of the
amples were passed through an Oasis HLB cartridge whichad been conditioned sequentially with 5 ml CH3OH and 5 mlater. After the cartridges had been dried for at least 30 miny a stream of nitrogen, elution was performed with 10 ml ofethanol. Resulted solution was completely dried with nitrogen,
nd then 0.5 ml of methanol or hexane (for GC–MS analysis)as added.Concentrations of NPEOs and the metabolites (e.g., NPECs
nd NP) were determined using a reverse phase liquid chro-atograph coupled with an electrospray mass spectrometry
LC–ES-MS) as reported elsewhere [19,20]. An LC–MSystem (Waters Alliance 2695 Separation Module, Waters
icromass ZQ 4000 and MassLynx V4.0 workstation) wassed with a Waters Symmetry ShieldTM RP-C18 column
2.1 mm × 150 mm) under a gradient elution at the flow rate of.2 ml min−1. The mobile phase was made up of a mixture ofmmonium acetate buffer (5 mM with 0.5‰ ammonia) and pureethanol. For the detection of NPEOs, positive ionization mode
2 gineer
wtiiastnimis8
l6mStsaq
2
apetoarrcs[ccs[gc3wwpw
2
brG(l
ppintiPkcwfcRg4Tctt
vUSA). The Dice’s coefficient of similarity [29] was calculatedon the basis of unweighted pair groups with the mathematicalaverage (UPGMA) method.
90 Y. Zhang et al. / Biochemical En
as used, while all of the metabolites were analyzed in nega-ive ionization mode. All analyses were performed in selectedon recording (SIR) mode, using the deprotonated molecularons of NPECs and NP in the negative ionization mode and themmonia adducts of NPEOs in the positive mode. The electro-pray interface was set at a probe temperature of 120 ◦C, andhe probe and cone voltages were −2.5 kV and −25 V in theegative ionization mode, and +3.0 kV and +30 V in the positiveonization mode, respectively. For identity confirmation of otheretabolites, full scans of product ions were obtained in negative
onization mode using LC separation as described above. Masspectra were obtained by scanning the quadrupoles from 150 to00m/z with a 2-s scan and setting the cone voltage at 20 V.
GC–MS analysis for detection of some possible metabo-ites of NPEOs and NP was performed using an Agilent890 GC equipped with an HP-5MS column (30 mm × 0.25m × 0.25 �m film thickness) using helium as the carrier gas.eparation on the column was achieved by using a tempera-
ure program from 50 to 300 ◦C (10 ◦C min−1). One-microlitreamples were injected in splitless mode with an injector temper-ture of 250 ◦C. The detector was an Agilent 5973 MSD withuadrapole and source settings of 150 and 280 ◦C, respectively.
.3. DNA enumeration by MPN-PCR
DNA templates were prepared by the proteinase K method,s previously described [21]. The DNA to be analyzed was inde-endently extracted in triplicate and serially diluted 10-fold atach step, and three samples of each dilution step were subjectedo PCR. The MPN number was determined as described previ-usly [22], based on the cut-off probability theory of Kohnond Fukunaga [23]. PCR primers were chosen from publishedeferences. The conditions of PCR were 30 cycles with denatu-ation at 94 ◦C for 60 s, annealing at 65 ◦C (6 cycles)/62 ◦C (6ycles)/59 ◦C (six cycles)/55 ◦C (12 cycles) for 30 s, and exten-ion at 72 ◦C for 30 s with the EUB-8f [24] and EUB-1387r25] primer sets for 16S rDNA. PCR was conducted for 40ycles, with denaturation at 94 ◦C for 60 s, annealing at 60 ◦C (10ycles)/57 ◦C (15 cycles)/55 ◦C (15 cycles) for 30 s, and exten-ion at 72 ◦C for 30 s with the C12Of/C12Or and C23Of/C23Or26] primer sets for C12O/C23O DNA. For determining the alkenes, PCR was conducted using ALK1 primers [27] for 30ycles, with denaturation at 94 ◦C for 60 s, annealing at 40 ◦C for0 s, and extension at 72 ◦C for 30 s. The PCR products (10 �l)ere analyzed by electrophoreses on a 1.2% agarose gel. The gelas stained with 0.5 �g ml−1 of ethidium bromide solution, andhotographed under UV light after being rinsed with distilledater.
.4. T-RFLP analysis of community structure
T-RFLP was used to examine variations of the micro-ial community structure of the microcosms. Eubacterial 16S
RNA genes were amplified with forward primer 27F (5′-AGTTTGATCCTGGCTCAG-3′) and reverse primer 1392R5′-ACGGGCGGTGTGTRC-3′) [28], where forward 27F wasabeled at the 5′ end with the phosphoramidite dye 6-FAM (phos-
Fc
ing Journal 39 (2008) 288–296
horamidite fluorochrome 5-carboxyfluorescein). The cyclerograms used were denaturation at 95 ◦C for 1 min, anneal-ng at 57 ◦C for 1 min, and extension at 72 ◦C for 3 min; theumber of cycles was 20–26. PCR products were subjectedo electrophoresis on 1.2% agarose gels, stained with ethid-um bromide (0.5 �g ml−1) and visualized by UV excitation.CR products were purified using a QIAquick PCR purificationit (QIAGEN, Japan) according to the manufacturer’s proto-ol. Then the PCR products were digested for 5 h at 37 ◦Cith HhaI, which is one of the most frequently used enzymes
or T-RFLP and it was able to determine the change in theommunity structure briefly. Terminal restriction fragments (T-Fs) were analyzed by electrophoresis on an ABI PRISM 310enetic analyzer (Applied Biosystems) with a GeneScan POP-TM capillary column (47 cm × 50 �m, Applied Biosystems).he size and the fluorescence intensity of each T-RF in a givenommunity fingerprint pattern were automatically calculated byhe GeneScan analysis software (version 3.7, Applied Biosys-ems).
Moreover, cluster analysis was carried out by using multi-ariate analysis software (NTSYS-pc.2.1, Exeter Software, NY,
ig. 1. Degradation profiles of NPEOs or NP (a) and NPECs (b) in each micro-osm.
Y. Zhang et al. / Biochemical Engineering Journal 39 (2008) 288–296 291
trol (
3
3
d5
Fm
5cd
Fig. 2. Change of 16S rDNA and catabolic genes in con
. Results and discussion
.1. NPEOs and NP degradation profile
Fig. 1(a) presents the changes of NPEOs and NP duringegradation. Fast degradation of NPEOs occurred in the NPEO-microcosm, and more than 95% removal was achieved on day
ig. 3. Ratios of MPN-DNA copies of C23O DNA to that of 16S rDNA in eachicrocosm.
mmacdtTp
3d
fc1CnAi
a), NPEO-5 (b), NPEO-25 (c) and NP (d) microcosms.
. Fast degradation of NPEOs occurred in the NPEO-25 micro-osm on day 9, and more than 90% removal was achieved onay 12. In contrast, NP demonstrated a slower decreasing rate.
LC–MS analysis results showed that NPECs gradually accu-ulated with the disappearance of NPEOs in the NPEOicrocosms (Fig. 1(b)). NPECs were formed, accounting for
pproximately 20% and 22% (molar basis) of the initial NPEOoncentrations in the NPEO-5 and NPEO-25 microcosms onay 18, respectively. NP2EC (NPEC with 2 ethoxy units) washe most abundant species in both microcosms (data not shown).he results found are in agreement with the NPEO metabolicathway evidenced by other authors [19].
.2. Variations of functional genes during NPEO and NPegradation
Fig. 2 presents variations of possible catabolic genes in dif-erent microcosms. In the control microcosm (Fig. 2(a)), theopy numbers of C12O and alk remained almost stable at01 copies ml−1 and zero, respectively. The copy numbers of
23O DNA and 16S rDNA (indicating the level of bacterialumber) in the control system, however, increased on day 9.s this experiment was performed in a natural water system,t is sometimes the case that the microbial community fluctu-
2 gineer
ao
ag(al
tdaB
92 Y. Zhang et al. / Biochemical En
tes [14]. It is speculated that certain substrates indigenous toriginal natural water might be degraded during this period.
For all of the three degradation microcosms, no signal for thelk genes was detected, suggesting that degradation of the alkyl
roup in NPEOs or NP might have not occurred during the periodFig. 2(b)–(d)). The existence of C12O DNA was confirmed inll of the microcosms from the beginning of the experiments. Theevels of C12O DNA in all of the NPEO and NP microcosmsicid
Fig. 4. Change of the bacterial community structure in control (a), NPEO
ing Journal 39 (2008) 288–296
hen increased slightly on day 2, but were nearly unchanged afteray 5, indicating that the existence of C12O DNA should not bettributed to the spiking of nonylphenolo compounds notably.y contrast with the copy numbers of C12O DNA, a significant
ncrease of the copy numbers of C23O DNA in all of the micro-osms appeared on day 2. The copy number levels of C23O DNAn the NPEO microcosms maintained an increasing trend untilay 13, while that in the NP microcosm was relatively stable
-5 (b), NPEO-25 (c) and NP (d) microcosms analyzed by T-RFLP.
Y. Zhang et al. / Biochemical Engineering Journal 39 (2008) 288–296 293
Conti
frim
tmtmt
wtCc[b
Fig. 4. (
rom day 2. The copy numbers of 16S rDNA, on the other hand,emained relatively stable in the NPEO and NP microcosms,ndicating that degradation of NPEOs and NP did not contribute
uch to the growth of bacterial populations.Fig. 3 shows that the ratio of the copy number of C23O DNA
o that of 16S rDNA in each degradation microcosm increased
arkedly with time during the degradation period, indicatinghat bacteria encoding C23O DNA might be enriched in theseicrocosms. Although the ratio of C23O DNA to 16S rDNA in
he control microcosm also increased, the timing for the increase
Cttc
nued ).
as much later and the level was much lower than those in thehree degradation microcosms. On the other hand, the ratios of23O DNA to 16S rDNA in the two NPEO degradation micro-osms were higher than that in the NP microcosm. Jeong et al.15] have reported that 3- and 4-alkylphenol (AP, C1–C5) cane degraded via a proximal (2, 3) ring cleavage pathway by
23O. They speculated that the C23O may be responsible forhe degradation of larger side chains due to the relaxed specifici-ies of catabolic enzymes. It is therefore speculated that partialleavage of aromatic rings with the C23O might have also hap-
2 gineering Journal 39 (2008) 288–296
pi
ttseaetoHpmtaNtam
cbpbot
3
d1fi2t2tabrtspct
t(Nsdfrmht
Fi
rqm
Stbhatdst(dnrltN
94 Y. Zhang et al. / Biochemical En
ened together with the �-oxidation oxyethylene chain pathwayn the three degradation microcosms.
It has been reported that NP could be degraded with fission ofhe phenol ring by some isolated strains, and it seems to be ableo further degrade the aromatic moiety of NP isomers as growthubstrates [30,31] proved the existence of a novel pathway thatnables bacteria to detach the alkyl substituent of an NP isomers C9 alcohol and to utilize the ring as a source of carbon andnergy. As for NPEOs, it is reported that ultimate biodegrada-ion of the metabolites occurs more slowly, if at all, becausef the need for a specific enzyme or bacterial population [32].owever, a recent study indicated that the benzene ring could beartially degraded within 10 days in an aerobic biodegradationicrocosm for NPEOs [33]. In a degradation study using syn-
hesized NP9EO with a 14C label in the aromatic ring, Naylor etl. [12] found that 4.7% (28 days) and 40.5% (128 days) of thePEO aromatic ring carbon were converted to 14CO2, respec-
ively, and some 14C was incorporated into the biomass. Thebove results support our assumption that partial ring cleavageight occur together with some primary degradation.Analysis of the cleavage-intermediates from the C23O ring
leavage pathway according to Jeong et al. [15] was attemptedy using GC–MS and LC–MS. However, no such intermediaryroducts could be observed. The cleavage-intermediates mighte immediately metabolized in the microcosms. Further studiesn the identification of the cleavage-intermediates are requiredo confirm the above speculation.
.3. Changes of microbial community structures by T-RFLP
The community structures in the control and different degra-ation microcosms were determined by T-RFLP, targeting the6S rRNA genes (Fig. 4). Though some changes could be identi-ed, it is clear that the T-RF peaks with DNA lengths of 674 and04 (or 202) bp were dominant in the control microcosm. In thewo NPEO degradation microcosms, the T-RF peaks at 674 and04 bp disappeared gradually with the increasing presence ofhe peak at 78 bp. Similarly, the T-RF peaks at 675 and 204 bplso disappeared gradually in the NP degradation microcosm,ut with the appearance of new peaks at 88 and 198 bp. Theseesults indicate that the population responsible for the degrada-ion of NPEOs and NP was clearly different. This is reasonable,ince primary transformation of NPEOs to NPECs together withartial ring cleavage might have occurred in the NPEO micro-osms, while the main reaction in the NP microcosm might behe decomposition of NP.
The possible strain candidate with T-RF of 78 bp is Betapro-eobacteria according to the Ribosomal Database Project IIMicrobial Community Analysis (MiCA) website) [34]. ManyPEO-degrading bacteria species belonging to the gamma
ubclass of the Proteobacteria have been isolated by culture-ependent methods [35-37]. However, Lozada et al. [38,39]ound that a high proportion of members of Betaproteobacte-
ia constitute the predominant group of bacteria in NPEO-fedicrocosms using dot-blot hybridization and fluorescent in situybridization. This finding together with our result suggestshat members of Betaproteobacteria might play an important
qkmt
ig. 5. Shannon–Weaver index of diversity (H′) (a) and Simpson index of dom-nance (D) (b) calculated from T-RLFP analyses of different microcosms.
ole in NPEO degradation in the natural environment, which isuite different from those obtained using the culture-dependentethods.The Shannon–Weaver index of diversity (H′) [40] and the
impson index of dominance (D) [41] were calculated fromhe results of T-RFLP to quantify the diversity of the micro-ial community (Fig. 5). High H′ and low D values indicateigh diversity of the microbial community. In the NPEO-5nd NPEO-25 degradation microcosms, the H′ values tendedo be lower and the D values higher than those of the NPegradation microcosms. This implied that microbial diver-ity in NPEO microcosms tended to decrease while that inhe NP microcosm remained relatively stable after day 2Fig. 5(b)). Thus, specific bacteria responsible for the degra-ation of NPEOs might have been selected. Because NP isot degraded as easily as NPEOs, the bacterial populationesponsible for the degradation could not be dominant, whicheads to relatively stable diversity. This is consistent withhe results that indicated that NP removal was slower thanPEO removal (Fig. 1(a)) and the community structures wereuite stable (Fig. 4(d)) during the degradation process. To our
nowledge, there are only a few reports about the change oficrobial community structures during NPEO and NP degrada-ion.
Y. Zhang et al. / Biochemical Engineer
Fb
cy(2N5mmdCCbtaccottoot
4
tidid
dpmsaabDbh1ciwimbr
A
F5Yhfo
R
ig. 6. Similarity in the bacterial community structure of different microcosmsy cluster analysis.
According to Dice’s coefficient of similarity in the bacterialommunity structure of different microcosms by cluster anal-sis, five major clusters can be identified, as follows (Fig. 6):1) Cluster 1: all the microcosms on day 0 and control on day; (2) Cluster 2: control on days 9, 13 and 18; (3) Cluster 3:P microcosm; (4) Cluster 4: NPEO-25 microcosm on days 2,, 9 and 13; (5) Cluster 5: NPEO-5 microcosm and NPEO-25icrocosm on day 18. This indicated that the bacterial com-unity structure was distinct depending on the substrates and
egradation processes. The control microcosm was divided intolusters 1 and 2. Populations on days 9–18 were assigned toluster 2. Changes of bacterial populations from day 9 mighte related to the sudden appearance of C23O DNA in the con-rol microcosm (Fig. 2(a)). The reason, however, is unknown. Inddition, days 2 and 13 are the critical points of Cluster 4, whichoincided with the degradation behavior in the NPEO-25 micro-osm in Fig. 1(a). It is interesting that the NPEO-25 microcosmn day 18 was categorized as Cluster 5, which contained all ofhe NPEO-5 microcosms. This might be attributed to the facthat the concentration of NPEOs in the NPEO-25 microcosmn day 18 was as low as that in the NPEO-5 microcosm. It wasbvious that the results of cluster analysis were consistent withhe different biodegradation processes as shown in Fig. 1.
. Conclusions
In this paper, the behavior of catabolic genes responsible forhe degradation of aromatic and alkane structures and changes
n microbial community structures in natural water microcosmsuring NPEO and NP degradation were analyzed. The signif-cant increase of C23O DNA occurred soon after the start ofegradation in NPEO and NP microcosms together with the[
[
ing Journal 39 (2008) 288–296 295
ecrease of NPEOs and NP, indicating that the existence ofotential aromatic ring-cleaving might have occurred in theicrocosms. This is the first report on genetic evidence of a pos-
ible aromatic ring meta-cleaving pathway of NPEOs and NP inn aquatic environment. The community structures in the controlnd degradation microcosms were determined by 16S rDNA-ased T-RFLP. The presence of a new dominant strain with aNA length of 78 bp in the NPEO microcosm suggested thatacteria affiliated with the beta subclass of Proteobacteria mayave an important role in NPEO degradation. Strains at 88 and98 bp were dominant in the NP microcosm. Five major clustersould be identified according to Dice’s coefficient of similarityn the bacterial community structures of different microcosms,hich were consistent with the different biodegradation behav-
ors. The monitoring of the microbial aspects involved in theetabolism of NPEOs and NP should be helpful for gaining a
etter understanding of the environmental fate of NPEOs andelated compounds.
cknowledgments
This work was supported by the National Natural Scienceoundation of China (Contract Nos. 50578153, 20521140076,0525824). The authors are also thankful to Dr. Inoue, Ms. Ningu and Mr. Kumada of Osaka University, Japan, for their kindelp. The authors are thankful to the Postdoctoral Fellowshipor Foreign Researchers by the Japan Society for the Promotionf Science (JSPS).
eferences
[1] E.J. Routledge, J.P. Sumpter, Estrogenic activity of surfactants and someof their degradation products assessed using a recombinant yeast screen,Environ. Toxicol. Chem. 15 (1996) 241–248.
[2] G.G. Ying, B. Williams, R. Kookana, Environmental fate of alkylphenolsand alkylphenol ethoxylates—a review, Environ. Int. 28 (2002) 215–226.
[3] K. Maruyama, M. Yuan, A. Otsuki, Seasonal changes in ethylene oxidechain length of poly(oxyethylene)alkylphenyl ether nonionic surfactantsin three main rivers in Tokyo, Environ. Sci. Technol. 34 (2000) 334–343.
[4] S.Y. Yuan, C.H. Yu, B.V. Chang, Biodegradation of nonylphenol in riversediment, Environ. Pollut. 127 (2004) 425–430.
[5] L. Wang, Y.H. Wu, H.W. Sun, J. Xu, S.G. Dai, Distribution and dissipa-tion pathways of nonylphenol polyethoxylates in the Yellow River: siteinvestigation and lab-scale studies, Environ. Int. 32 (2006) 907–914.
[6] USEPA (United States Environmental Protection Agency), Notice of avail-ability of final aquatic life ambient water quality criteria for nonylphenol,Fed. Reg. 71 (2006) 9337–9339.
[7] R. Renner, European bans on surfactant trigger transatlantic debate, Envi-ron. Sci. Technol. 31 (1997) 316A–320A.
[8] J. Ejlertsson, M.L. Nilsson, H. Kylin, A. Bergman, L. Karlson, M.Oquist, B.H. Svensson, Anaerobic degradation of nonylphenol mono- anddiethoxylates in digester sludge, landfilled municipal solid waste, and land-filled sludge, Environ. Sci. Technol. 33 (1999) 301–306.
[9] K.H. Langford, M.D. Scrimshaw, J.W. Birkett, J.N. Lester, Degradationof nonylphenolic surfactants in activated sludge batch tests, Water Res. 39(2005) 870–876.
10] W.H. Ding, S.H. Tzing, Analysis of nonylphenol polyethoxylates and theirdegradation products in river water, J. Chromatogr. A 824 (1998) 79–90.
11] A. Di Corcia, R. Cavallo, C. Crescenzi, M. Nazzari, Occurrence and abun-dance of dicarboxylated metabolites of nonylphenol ethoxylates in treatedsewage, Environ. Sci. Technol. 34 (2000) 3914–3919.
2 gineer
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
[
96 Y. Zhang et al. / Biochemical En
12] C.G. Naylor, C.A. Staples, G.M. Klecka, J.B. Williams, P.T. Varineau, C.Cady, Biodegradation of [14C] ring-labeled nonylphenol ethoxylate, Arch.Environ. Contam. Toxicol. 51 (2006) 11–20.
13] K. Sei, Y. Sugimoto, K. Mori, H. Maki, T. Kohno, Monitoring of alkane-degrading bacterial in a seawater microcosm during crude oil degradationby polymerase chain reaction based on alkane-catabolic genes, Environ.Microbiol. 5 (2003) 517–522.
14] K. Sei, D. Inoue, K. Wada, K. Mori, M. Ike, T. Kohno, M. Fujita, Monitoringbehaviour of catabolic genes and change of microbial community structuresin seawater microcosms during aromatic compound degradation, WaterRes. 38 (2004) 4405–4414.
15] J.J. Jeong, J.H. Kim, C.K. Kim, I. Hwang, K. Lee, 3- and 4-alkylphenoldegradation pathway in Pseudomonas sp. strain KL28: genetic organizationof the lap gene cluster and substrate specificities of phenol hydroxylase andcatechol 2,3-dioxygenase, Microbiology 149 (2003) 3265–3277.
16] A. Hiraishi, I. Mitsuru, S. Hisashi, Terminal restriction pattern analysisof 16S rRNA genes for the characterization of bacterial communities ofactivated sludge, J. Biosci. Bioeng. 90 (2000) 148–156.
17] T.L. Marsh, Terminal restriction fragment length polymorphism (T-RFLP):an emerging method for characterizing diversity among homologous popu-lations of amplification products, Curr. Opin. Microbiol. 2 (1999) 323–327.
18] C. Picard, C. Ponsonnet, E. Paget, X. Nesme, P. Simonet, Detection andenumeration of bacteria in soil by direct DNA extraction and polymerasechain reaction, Appl. Environ. Microbiol. 58 (1992) 2717– 2722.
19] N. Jonkers, T.P. Knepper, P. De Voogt, Aerobic biodegradationstudies of nonylphenol ethoxylates in river water using liquidchromatography–electrospray tandem mass spectrometry, Environ. Sci.Technol. 35 (2001) 335–340.
20] F. Houde, C. DeBlois, D. Berryman, Liquid chromatographic–tandem massspectrometric determination of nonylphenol polyethoxylates and nonylphe-nol carboxylic acids in surface water, J. Chromatogr. A 961 (2002) 245–256.
21] K. Sei, K. Asano, N. Tateishi, K. Mori, M. Ike, T. Kohno, M. Fujita, Devel-opment of simple methods of DNA extraction from environmental samplesfor monitoring microbial community based on PCR, Jpn. J. Water Treat.Biol. 36 (2000) 193–204.
22] T. Kohno, S. Fukunaga, Auto estimation of quantal improbability and deter-mination of MPN and its 95% limits with non-fixed dilution schemes, Jpn.J. Water Treat. 37 (1996) 381–397 (in Japanese).
23] T. Kohno, S. Fukunaga, The ‘cut-off probability’ as the measure of quantalimprobability in the dilution method, Water Res. 32 (1998) 3099–3107.
24] W.G. Weisburg, S.M. Barns, D.A. Pelletier, D.J. Lane, 16S ribosomal DNAamplification for phylogenetic study, J. Bacteriol. 173 (1991) 697–703.
25] T. Iwamoto, K. Tani, K. Nakamura, Y. Suzuki, M. Kitagawa, M. Eguchi,
M. Nasu, Monitoring impact of in situ biostimulation treatment on ground-water bacterial community by DGGE, FEMS Microbiol. Ecol. 32 (2000)129–141.26] K. Sei, K. Asano, N. Tateishi, K. Mori, M. Ike, M. Fujita, Design of PCRprimers and gene probes for the general detection of bacterial populations
[
[
ing Journal 39 (2008) 288–296
capable of degrading aromatic compounds via catechol cleavage pathways,J. Biosci. Bioeng. 88 (1999) 542–550.
27] T. Kohno, Y. Sugimoto, K. Sei, K. Mori, Design of PCR primers andgene probes for extensive detection of alkane-degrading bacteria, Microb.Environ. 17 (2002) 114–121.
28] R.I. Amann, W. Ludwig, K.H. Schleifer, Phylogenetic identification and insitu detection of individual microbial cells without cultivation, Microbiol.Rev. 59 (1995) 143–169.
29] C.A. Eichner, R.W. Erb, K.N. Timmis, I. Wagner-Dobler, Thermal gradientgel electrophoresis analysis of bioprotection from pollutant shocks in theactivated sludge microbial community, Appl. Environ. Microbiol. 65 (1999)102–109.
30] T. Tanghe, W. Dhooge, W. Verstraete, Isolation of a bacterial strain ableto degrade branched nonylphenol, Appl. Environ. Microbiol. 65 (1999)746–751.
31] F.L.P. Gabriel, A. Heidlberger, D. Rentsch, W. Giger, K Guenther, H-P.E. Kohler, A novel metabolic pathway for degradation of 4-nonylphenolenvironmental contaminants by Sphingomonas xenophaga Bayram: ipso-hydroxylation and intramolecular rearrangement, J. Biol. Chem. 280 (2005)15526–15533.
32] K.H. Langford, J.N. Lester, Fate and behavior of endocrine disruptersin wastewater treatment processes, in: J.W. Birkett, J.N. Lester (Eds.),Endocrine Disrupters in Wastewater and Sludge Treatment Processes, CRCPress, FL, USA, 2002.
33] J. Zhao, G. Zhang, Y. Qin, Y. Zhao, Aerobic biodegradation of alkyphenolethoxylates, Bioresour. Technol. 97 (2006) 2478–2480.
34] A.D. Kent, D.J. Smith, B.J. Benson, E.W. Triplett, Wed-based phylogeneticassignment tool for analysis of terminal restriction fragment length poly-morphism profiles of microbial communities, Appl. Environ. Microbiol.69 (2003) 6768–6776.
35] H. Maki, N. Masuda, Y. Fujiwara, M. Ike, M. Fujita, Degradation ofalkylphenol ethoxylates by Pseudomonas sp. strain TR01, Appl. Environ.Microbiol. 60 (1994) 2265–2271.
36] D.D. Gioia, L. Fambrini, E. Copponi, F. Fava, C. Barberio, Aggregation-based cooperation during bacterial aerobic degradation of polyethoxylatednonylphenols, Res. Microbiol. 155 (2004) 761–769.
37] D.M. John, G.F. White, Mechanism for biotransformation of nonylphenolpolyethoxylates to Xenoestrogens in Pseudomonas putida, J. Bacteriol. 180(1998) 4332–4338.
38] M. Lozada, R.F. Itria, E.L.M. Figuerola, P.A. Babay, L.A. De Tullio, L. Eri-jman, Bacterial community shifts in nonylphenol polyethoxyate-enrichedactivated sludge, Water Res. 38 (2004) 2077–2086.
39] M. Lozada, E.L.M. Figuerola, R.F. Itria, L. Erijman, Replicability of
dominant bacterial populations after long-term surfactant enrichment inlab-scale activated sludge, Environ. Microbiol. 8 (2006) 625–638.40] C.E. Shannon, W. Weaver, The Mathematical Theory of Communication,5th ed., University of Illinois Press, Urbana, IL, USA, 1963.
41] E.H. Simpson, Measurement of diversity, Nature 163 (1949) 688.
