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International Biodeterioration & Biodegradation 59 (2007) 97–102

www.elsevier.com/locate/ibiod

MTBE biodegradation and degrader microbial community dynamics inMTBE, BTEX, and heavy metal-contaminated water

Chi-Wen Lina,�, Hung-Chun Lina, Chi-Yung Laib

aDepartment of Environmental Engineering, Da-Yeh University, 112 Shan-Jiau Rd., Da-Tsuen, Changhua 515, Taiwan, ROCbDepartment of Biology, National Changhua University of Education, Changhua 515, Taiwan, ROC

Received 12 March 2006; received in revised form 21 June 2006; accepted 31 July 2006

Available online 26 September 2006

Abstract

The aim of this investigation was to explore microbial community changes under various environmental groundwater conditions

(single substrate, mixed substrates, and the presence of heavy metals) and link the changes with simultaneously diminishing substrate

concentration in the microcosms. Most microorganisms from environmental microcosms or wastewater treatment plants cannot be

cultivated artificially. Capturing microbial community fingerprints, therefore, requires applying a molecular biological technique. By

using SSCP profiles of PCR-amplified 16S rDNA genes, it was demonstrated that with the repeated addition of substrates during long-

term acclimation, substrate-utilizing populations in a microcosm gradually increased to become the dominant constituent. Conversely,

the presence of metals inhibited community development and differentiation. It was also shown that substrate degradation rates

increased under co-substrate conditions, with substrate-degraders easily adapting to the environment and becoming the dominant

bacteria, a phenomenon attributed to the propensity of the fittest species to outgrow their competitors when presented with suitable

substrates.

r 2006 Elsevier Ltd. All rights reserved.

Keywords: 16S rDNA; Metal ions; Microbial community structure; Molecular profiling; Methyl tert-butyl ether

1. Introduction

Methyl tertiary butyl ether (MTBE) is one of several fueloxygenates added to gasoline to replace tetraethyl lead andreduce harmful tailpipe emissions. The present guidelinelimit established by the US Environmental ProtectionAgency (EPA) is 20–40 mg l�1 for MTBE in drinking water(Jacobs et al., 2001). The EPA also classifies MTBE as apossible human carcinogen (Johnson et al., 2000). Numer-ous studies have demonstrated that MTBE can bebiodegraded under aerobic or anaerobic conditions (Eweiset al., 1997; Mo et al., 1997; Steffan et al., 1997; Hansonet al., 1999; Bruns et al., 2001; Hristova et al., 2003). TheMTBE biodegradation rate may further diminish in thepresence of co-contaminants, such as benzene, ethylben-zene, toluene, and xylene (BTEX). However, little is known

e front matter r 2006 Elsevier Ltd. All rights reserved.

iod.2006.08.002

ing author. Tel.: +886 4 8511339; fax: +886 4 8511347.

ess: cwlin@mail.dyu.edu.tw (C.-W. Lin).

about the microbial community structure during aerobicMTBE degradation in the presence of BTEX.The inhibitory effects on MTBE biodegradation have

recently been investigated by several researchers. Thesestudies have focused mainly on substrate inhibition (Park,1999), by-product inhibition (Liu et al., 2001; Wilson et al.,2002) or competitive inhibition (Garnier et al., 1999;Sedran et al., 2002). However, little has been publishedon the effect of heavy metals on MTBE biodegradation.Many types of pollutants may coexist in a contaminatedsite. For instance, organic chemicals and heavy metals canappear simultaneously in a leaking oil tank. Reports byseveral researchers (Baldrian et al., 2000; Bruins et al.,2000) indicate that some heavy metals (Cu, Zn, Cd)produce toxicity and may also simultaneously affectorganic contaminant biodegradation (Said and Lewis,1991) owing to heavy metal accumulation in the microbialcells. Metal toxicity on microorganisms has been studied bygrowth inhibition, while heavy metal toxic effects onbacterial community structure have been studied to a

ARTICLE IN PRESSC.-W. Lin et al. / International Biodeterioration & Biodegradation 59 (2007) 97–10298

lesser extent. To fully exploit MTBE bioremediationtechnology for field application, the biodegradation andmicrobial community structure in the presence of heavymetals must be better understood.

Most microorganisms cannot be detected in soil andgroundwater samples using cultivation-based methods(Amann et al., 1995; Ranjard et al., 2000). Conventionalmicroorganism analyses cannot fully reveal the species andthe resulting microbial community composition in theenvironment. Hence, it is necessary to utilize molecularapproaches to provide clues to bacterial types involved inaerobic phenol degradation and determine the relationshipbetween growth and decline among bacterial populations.

Methods for exploring the microbial community dy-namics during in situ bioremediation has recently receivedincreased attention. Griffiths et al. (1999) employed aDNA–phospholipids fatty acid (PLFA) method to analyzethe effects of varying substrate concentrations on a soilmicrobial community. Their findings indicated that, in thesubstrate range 0–125 mgml�1, lower substrate concentra-tions resulted in less change in community structure, asevidenced by high similarity indices (0.8–0.9) between pre-and post-incubation samples. Peter et al. (2000) demon-strated that compost microbial communities are dominatedby a few species with the most favoured growth require-ments. Hristova et al. (2001) dealt with molecularfingerprinting techniques and detected the MTBE-degrad-ing strain PM1 by real-time TaqMan PCR. Deeb et al.(2001) showed the substrate interactions in BTEX andMTBE mixtures by an MTBE-degrading isolate. Theseauthors have proposed some tools for determining whethercertain degraders are present, and have shown inhibitionby o-xylene. However, they did not address the changingcomposition of microbial communities. Zhang et al. (2005)studied a sequencing batch reactor (SBR) system usingPCR–denaturing gradient gel electrophoresis (DGGE) andreported that the resident community reached a stablestructure dominated by nine species after operation for 200days.

Single-strand-conformation polymorphism (SSCP) isone of the methods frequently used, being inexpensiveand highly sensitive (Lee et al., 1996; Schwieger and Tebbe,1998). In recent years, it has been applied extensively todetect mixed culture diversity in the environment. Forinstance, Stach et al. (2001) showed that SSCP microbialcommunity analysis could be used to compare theefficiency of alternative methods for DNA extraction andpurification. Balcke et al. (2004) used PCR–SSCP toevaluate microbial community changes under consecutiveaerobic–anaerobic and various pH conditions. Dassonvilleet al. (2004) examined microbial dynamics in anaerobic soilslurry and analyzed their dependence on geochemicalprocesses by using the SSCP technique.

In this study MTBE biodegradation was analyzed in thepresence of other contaminants in batch culture experi-ments. Some of the contaminants served as co-substrateswhile the others functioned as growth inhibitors. Then the

SSCP method was combined with batch culture experi-ments to examine the microbial population and communitychanges during degradation.

2. Materials and methods

2.1. Bacterial strains and batch microcosm experiments

The bacterial cultures used in this study were obtained from a

petrochemical wastewater treatment plant. They were initially grown in

a liquid medium and adapted to the targeted compounds and working

conditions before the biodegradation experiments. After a series of

screening and isolation procedures, three pure cultures were identified as

Ralstonia sp. (YABE411), Pseudomonas sp. (YATO411), and Pseudomo-

nas sp. (YAET411) using 16S rDNA sequence analysis. The microorgan-

isms were grown at pH 6.871 in a mineral salts medium as described

elsewhere (Lin and Li, 2006). DNA from pure cultures was used as the

electrophoresis markers in some of the bacterial community studies.

For the experiments, 250-ml screw-capped amber glass bottles, each

containing 80-ml autoclaved phosphate-buffered mineral salts solution

(approx. pH 6.8) autoclaved at 121 1C for 20min and sealed with Teflon

Mininert valves, were shaken continuously in the dark at 150 rpm and

3071 1C. A stainless steel needle fitted to a gas-tight syringe was used to

inject MTBE, after which the bottles were re-sealed and shaken for 12 h.

This procedure ensured complete dissolution of the MTBE and

equilibrated the headspace with the solution. Subsequently, 20mg l�1

mixed-culture inoculum was diluted to 20ml in deionized water and then

introduced to give the same final biomass concentration in each bottle.

The inoculum containing the three species noted above, and other

organisms, was obtained from a biotrickling filter in a petrochemical

wastewater treatment plant treating MTBE and BTEX. MTBE and BTEX

solvents (499% pure, Aldrich, Steinheim, Germany) were used as carbon

source. Metal ions (Al3+, Zn2+) from concentrated stock solutions of the

metals in ionic form (1000mg l�1, Merck), were prepared in distilled water.

Substrates (MTBE, BTEX) and metal ions (Al3+ and Zn2+) were added

as needed to provide a determining concentration in the bottles. Test

cultures and uninoculated control cultures were prepared in triplicate.

2.2. Analytical methods for substrates

Because MTBE and BTEX are volatile organic compounds (VOCs),

equilibrium between their gas and liquid concentrations is maintained

through rapid mass transfer between the gas and aqueous phases. Changes

in liquid-phase concentration due to biological reactions are slow when

compared to the mass transfer rate between the gas and liquid phase.

Therefore, gas-phase measurements closely reflect liquid substrate

concentrations for biodegradation experiments. In the biodegradation

experiments gas samples were periodically collected from the headspace of

each bottle to monitor MTBE degradation using 250-ml gas-tight syringesequipped with Teflon Mininert valve fittings. Samples were then injected

onto a gas chromatograph equipped with an RTX-1 capillary column

(30m� 0.53mm) and a flame ionization detector (GC-FID, 14B,

Shimadzu, Japan). Helium (99.98% pure) was used as the carrier gas,

and nitrogen as a makeup gas. The temperature of the oven was controlled

at a constant 105 1C; the injector was at 200 1C; and the detector at 250 1C.

MTBE concentrations were quantified against primary standard curves.

Metal ion concentrations were determined by using an inductively coupled

plasma atomic emission spectrophotometer (ICP–AES, Perkin Elmer,

Optima 2000DV, USA).

2.3. DNA extraction

DNA was extracted using an improved bead-beating method. A

groundwater sample (200ml) was mixed with 0.8 g 0.106mm glass beads

(Biospec Products, 11079101), 600ml phenol/chloroform/isoamyl alcohol

ARTICLE IN PRESS

Fig. 1. (a) SSCP bacterial community profiles from 16S rDNA PCR

amplicons during batch tests under mixed substrate conditions for 30

days, and (b) cluster analysis. Box zones in (a) represent similar microbial

communities; L1: MTBE+toluene, L2: MTBE+ethylbenzene, L3 :

MTBE+benzene, L4: MTBE+m-xylene, L5: MTBE+p-xylene, L6:

MTBE+o-xylene.

0

20

40

60

80

100

0 5 10 15 20 25

Time (days)

Subs

trat

e re

mai

ning

(%

)

Fig. 2. Influence of mixed substrate conditions on removal different

substrates: (m) benzene; (Open triangle) m-xylene; (’) toluene; (&) p-

xylene; (K) o-xylene; (J) ethylbenzene.

C.-W. Lin et al. / International Biodeterioration & Biodegradation 59 (2007) 97–102 99

(25:24:1), and 200ml disruption buffer (50mMNaCl, 50mM Tris–HCl pH

8, 5% SDS) in a 1.5-ml screw-cap microcentrifuge tube, which was then

filled with the disruption buffer to remove the air. The mixture was

homogenized on a Mini Bead Beater (Biospec Products, 3110BX) at

2500 rpm for 2min. The homogenate was centrifuged at 30,000g for 5min,

and the supernatant was transferred to a fresh tube and extracted with

400-ml chloroform/isoamyl alcohol (24:1). The upper aqueous phase was

transferred to a fresh tube and mixed with 240-ml isopropanol. The DNA

was precipitated by centrifugation (32,600g, 3min), and washed with 240-

ml 70% ethanol. The pellet was air-dried for approx. 1 h at room

temperature and then allowed to dissolve in 50-ml TE buffer at 65 1C for

1 h. The concentration of the DNA solution was adjusted to 50 ngml�1,

after which the final solution was stored at 4 1C for use as the PCR

template.

2.4. Polymerase chain reaction

The microbial communities were analyzed by using the PCR–SSCP

method as described by Lee et al. (1996) and Schwieger and Tebbe (1998)

with modifications. The V3 region of the 16S rDNA, corresponding to the

nucleotide positions 334–514 of the Escherichia coli. gene, was amplified

with the primers EUB1 (50-CAGACTCCTACGGGAGGCAGCAG-30)

and U2 (50-GTATTACCGCGGCTGCTGGCAC-30). The PCR program

included an initial denaturation at 94 1C for 5min, 30 cycles at 94 1C for

30 s, 55 1C for 30 s, and 72 1C for 30 s, followed by a final extension of

72 1C for 5min. The PCR products of 200 bp were verified by gel

electrophoresis on 1.8% agarose gels and stored at 4 1C for further use.

2.5. SSCP gel electrophoresis

A vertical gel electrophoresis apparatus (Model SE600, Hoefer, San

Francisco, USA) was used for SSCP analysis. The electrophoresis was

conducted in 10% polyacrylamide gel for 6 h at a constant voltage of

300V. The gel temperature was maintained at 4 1C using a circulating

water bath. The DNA samples were mixed with equal volumes of a

denaturing solution (95% formamide, 10mM NaOH, 0.02% bromophe-

nol blue, 0.02% xylene cyanol, and 20mM EDTA), heated to 95 1C for

5min, and snap-frozen on ice before loading. The gels were visualized by

using the silver-stain method, sandwiched between two pieces of mylar

membrane and dried.

2.6. Statistical comparison of SSCP pattern

The relative positions of the DNA bands in the SSCP gels were

analyzed using LabWork software. Similarities between microbial groups

were calculated as Dice indices according to procedures appearing in

several reports (Dice, 1945; Eichner et al., 1999; Lapara et al., 2001).

Dendrograms were calculated using a clustering algorithm of a UPGMA

(unweighted pair group method using arithmetic average) with cluster

analysis of similarity indices, constructed by NTSYSpc software

(NTSYSpc, version 2.1e, Exeter Software, USA).

3. Results and discussion

3.1. Effects of acclimation periods on microbial community

Fig. 1a shows that, after a 30-day acclimation period,different types of substrate resulted in different microbialcommunity structures. Lanes L1, L2, and L3 in Fig. 1arepresent communities developed in bi-substrate cultures ofMTBE+toluene, MTBE+ethylbenzene and MTBE+ben-zene, respectively. The total number of bands in theseprofiles were similar, indicating similar species compositionin the different environments. However, the substrate

removal efficiency decreased in the order of MTBE+toluene4MTBE+ethylbenzene4MTBE+benzene (Fig. 2).The community grown with MTBE+toluene also

ARTICLE IN PRESSC.-W. Lin et al. / International Biodeterioration & Biodegradation 59 (2007) 97–102100

generated the most intense bands, followed by the culturewith MTBE+benzene. Removal efficiency is therefore agood indication of substrate quality in supporting domi-nant microbial species growth, as revealed by changes inband intensity.

3.2. Effects of mixed substrates on microbial community

Growth in bi-substrate systems of MTBE and o-, m- orp-xylene resulted in similar community structures. More-over, communities in xylene-containing cultures tended togenerate greater band numbers than benzene-containingcultures. Bands from the former also tended to have greaterintensity (Fig. 1a, L4–L6). Therefore, communities fromxylene-containing cultures can on the basis of theirstructural similarity be placed in this group (Fig. 1a,L4–L6), and those from benzene-containing cultures inanother (Fig. 1a, L1–L3). Degrees of similarity betweencommunities are summarized by a dendrogram constructedusing UPGMA (Fig. 1b). Communities from xylene- andbenzene-containing cultures again fall into two separategroups with similarity within each group as high as 0.88.

The substrate degradation rate has been known to affectcommunity structure (Balcke et al., 2004). Our results showthat toluene, ethylbenzene, and benzene are degraded athigher rates than o-, m- and p-xylene (Fig. 2). Aromaticcompounds with high degradation rates frequently gen-erate communities with simple structures, probably byfavouring the growth of a few dominant species, exempli-fied in Fig. 3 by the gradual increase in the intensity of

Fig. 3. SSCP bacterial community profiles obtained from 16S rDNA PCR

amplicons during batch tests under mixed substrate conditions: L1,

YATO411; L2, MTBE+toluene (10 days); L3, MTBE+toluene (30 days);

L4, YABE411; L5, MTBE+benzene (10 days); L6, MTBE+benzene (30

days); L7, YAET411; L8, MTBE+ethylbenzene (9 days); L9,

MTBE+ethylbenzene (30 days). Arrows denote dominant microbes.

bands a and b in L2/L3 and L8/L9. Such changesdemonstrate that the acclimation of communities to a bi-substrate system proceeds over an extended incubationperiod as indigenous species capable of utilizing thesubstrates gradually increase in proportion and becomethe dominant populations. Consistent with these observa-tions, the species represented by bands a and b in theMTBE+toluene culture during the 30-day acclimationperiod also appeared in pure cultures isolated by usingtoluene, benzene, or ethylbenzene as the sole carbon source(Fig. 3, L1, L4 and L7, respectively).

3.3. Microbial community structure in presence of metal

ions

The effects of both Al3+ and Zn2+ on MTBEdegradation, with metal concentrations of 5 and 10mg l�1,were that in all cases MTBE removal rates were lower inthe presence of metal ions in the solution (Fig. 4). At bothconcentrations, Zn2+ consistently showed greater adverseeffects than Al3+. The inhibitory effect was dose-depen-dent at concentrations from 5 to 10mg l�1, with the effectgreater at higher concentration. This result is in agreementwith Amor et al. (2001), who found that microorganismsexhibited various growth rate and degradation ratereductions in the presence of different types and concen-trations of heavy metals. In the present case, zinc exhibitedthe highest toxicity.Fig. 5a shows the influence of the metal ions on the

SSCP profiles of communities, and Fig. 5b the similarityindices and clustering relationships among these commu-nities. Communities grown in the presence and absence ofmetal ions exhibit a similarity of 0.65, revealing a smallchange in community structure due to the ions. This is instark contrast to the results of co-substrate addition, whichgenerally produces distinctively different community pro-files (Fig. 3). Nevertheless, this result agrees with Jacek andJan (2001) who demonstrated differences in microbialcommunity structure between a heavy metal-contaminatedand an uncontaminated site.The concentration appears to have a greater impact on

community structure than the type of metal present.

0

20

40

60

80

100

0 2 4 6 8 10 12 14

Time (days)

MT

BE

rem

aini

ng (%

)

Fig. 4. Influence of metal ions, at 5 and10mg l�1, on MTBE removal. (W)

MTBE alone; (&) MTBE+Zn (5mg l�1); (’) MTBE+Zn(10mg l�1); (J)

MTBE+Al(5mg l�1); (K) MTBE+Al(10mg l�1).

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L1 L5L4L3L2

Similarity

0.60 0.67 0.74 0.81 0.88

Al (5 mg l-1)

Zn (5 mg l-1)

Al (10 mg l-1)

Zn (10 mg l-1)

MTBE alone

(a)

(b)

Fig. 5. (a) SSCP bacterial community profiles from 16S rDNA PCR

amplicons during batch tests for mixed cultures containing MTBE and

metals, and (b) cluster analysis: L1, Al (5mg l�1); L2, Zn (5mg l�1); L3, Al

(10mg l�1); L4, Zn (10mg l�1); L5, MTBE alone.

C.-W. Lin et al. / International Biodeterioration & Biodegradation 59 (2007) 97–102 101

Community structures treated with metal ions can bedivided into two groups on the basis of ion concentrations.At 5 and 10mg l�1, the similarities between Al- and Zn-treated communities were 0.88 and 0.82, respectively.However, the similarity between communities at the twoconcentrations of Zn or Al was o0.66. Cultures treatedwith metal ions also contained a smaller number of bandsand lower band intensity than the untreated sample. Boththese features agree with those of Jacek and Jan (2000),who found that communities from sites contaminated byhigh concentrations of heavy metals (Pb, 1380mgkg�1;Cd, 23.3mg kg�1; Zn, 2390mgkg�1) were much lower inspecies diversity than communities from less-contaminatedsites (160, 4.4 and 330mgkg�1, respectively).

In conclusion, it can be said that, as revealed by bandintensity increases in PCR–SSCP community profileanalysis, removal efficiency is a good indication of thesubstrate quality in supporting growth of the dominant

microbial species. The acclimation of communities to a bi-substrate system involves gradual proportional increase ofindigenous substrate-utilizing species in relation to theircompetitors. Consequently, aromatic compounds withhigher degradation rates frequently generate communitieswith simpler structures, probably by favouring the growthof a few dominant species. Here, the presence of metalssignificantly reduced the diversity of the bacterial commu-nity. When the high similarities between SSCP patterns of16S rDNA fingerprints are judged, it is apparent thatgreater metal concentration resulted in less growth anddifferentiation of the microbial community.

Acknowledgments

The financial support of the National Science Council ofTaiwan, ROC (NSC 93-2211-E-212-007) is gratefullyacknowledged. The authors also wish to express apprecia-tion to Dr Cheryl J. Rutledge for her assistance inpreparing this manuscript.

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