anaerobic/aerobic conditions and biostimulation for enhanced chlorophenols degradation in biocathode...

ORIGINAL ARTICLE

Anaerobic/aerobic conditions and biostimulationfor enhanced chlorophenols degradation in biocathodemicrobial fuel cells

Liping Huang • Yinghong Shi • Ning Wang •

Yuesheng Dong

Received: 8 October 2013 / Accepted: 10 February 2014 / Published online: 3 April 2014

� Springer Science+Business Media Dordrecht 2014

Abstract Anaerobic/aerobic conditions affected

bacterial community composition and the subsequent

chlorophenols (CPs) degradation in biocathode micro-

bial fuel cells (MFCs). Bacterial communities accli-

mated with either 4-chlorophenol (4-CP) or 2,4-

dichlorophenol (2,4-DCP) under anaerobiosis can

degrade the respective substrates more efficiently than

the facultative aerobic bacterial communities. The

anaerobic bacterial communities well developed with

2,4-DCP were then adapted to 2,4,6-trichlorophenol

(2,4,6-TCP) and successfully stimulated for enhanced

2,4,6-TCP degradation and power generation. A 2,4,6-

TCP degradation rate of 0.10 mol/m3/d and a maxi-

mum power density of 2.6 W/m3 (11.7 A/m3) were

achieved, 138 and 13 % improvements, respectively

compared to the controls with no stimulation. Bacte-

rial communities developed with the specific CPs

under anaerobic/aerobic conditions as well as the

stimulated biofilm shared some dominant genera and

also exhibited great differences. These results provide

the most convincing evidence to date that anaerobic/

aerobic conditions affected CPs degradation with

power generation from the biocathode systems, and

using deliberate substrates can stimulate the microbial

consortia and be potentially feasible for the selection

of an appropriate microbial community for the target

substrate (e.g. 2,4,6-TCP) degradation in the biocath-

ode MFCs.

Keywords Microbial fuel cell � Biocathode �Biodegradation � Bacterial community �Stimulation � Chlorophenol

Introduction

Chlorophenols (CPs) have been extensively used as

intermediates of dyes, pesticides and herbicides, and

wood preservatives. Thus they are commonly found in

ground waters, sediment and surface soils from dry

areas near wood treatment plants, industrial wastewa-

ter effluents and treatment lagoons. CPs usually

require long attenuation periods and exploring highly

efficient methods for accelerating the transformation

and degradation rates in aquatic sediments and

groundwater is particularly important (Field and

L. Huang (&) � N. Wang

Key Laboratory of Industrial Ecology and Environmental

Engineering, Ministry of Education (MOE), School of

Environmental Science and Technology, Dalian

University of Technology, Dalian 116024, China

e-mail: [email protected];

[email protected]

Y. Shi

Weihai Supervision and Inspection Institute of Product

Quality, Weihai 264200, China

Y. Dong

School of Life Science and Biotechnology, Dalian

University of Technology, Dalian 116024, China

123

Biodegradation (2014) 25:615–632

DOI 10.1007/s10532-014-9686-1

Sierra-Alvarez 2008). Conventional biological pro-

cesses are environmentally sustainable and cost-

effective for CPs degradation. Multiple approaches

including optimizing either bacterial inoculum ratios

or stepwise pH reductions, and imposing appropriate

substrates (electron donors/acceptors, intermediates)

(Cao et al. 2012; Chandra et al. 2012; Chen et al. 2012;

Krumins et al. 2009; Mun et al. 2008; Park et al. 2011;

Puyol et al. 2011; Tyagi et al. 2011) have been

deliberately explored to stimulate the bacterial activ-

ities and accelerate CPs degradation in conventional

biological processes. The main challenges to these

approaches in practice, however, are the need of

extensive organics and excess sludge generation.

Alternative to organic electron donor/acceptor, elec-

trochemical processes, which use applied voltages can

directly provide electrons to the microbes and stim-

ulate the dechlorination of tetrachlorobenzene and the

removal of weathered PCBs from sediments (Chun

et al. 2013; Stuart et al. 1999; Sun et al. 2010).

Remaining challenges are extensive energy consump-

tion and high operation costs.

One new promising method for more efficient and

cost-effective CPs degradation is the use of microbial

fuel cells (MFCs), in which microbe communities on

the anode catalyze the conversion of reduced wastes

into electrical current (Li and Yu 2011; Logan and

Rabaey 2012; Winfield et al. 2013; Yuan et al. 2013).

The microbial cathode (biocathode), which uses

bacteria as biocatalysts to accept electrons from the

cathode substratum, provides a different path that

either avoids the use of noble catalysts for oxygen

reduction (Cai et al. 2013; Sun et al. 2012; Xia et al.

2012, 2013), or enables the use of alternative electron

acceptors, broadening the applicability of MFCs for

oxidative recalcitrant wastes treatment (Huang et al.

2011a, b). The electrotrophic capabilities of both

decreasing electrode overpotentials and removing

recalcitrant substrates enables the biocathode more

advantage over chemical cathode, particularly at a

neutral pH environment. In addition, compared to

conventional biological processes, no extensive

organics are required in the biocathode, resulting in

comparatively much low biomass production and thus

benefiting to the subsequent sludge treatment. The

cathodic electrons, on the other hand, are extracted

from organic wastewaters in the anode, where the

primary economic benefit for wastewater treatment

was reducing, or completely avoiding, the need for

electrical power consumption for aeration. These

multiple merits make biocathode MFCs more attrac-

tive than conventional biological processes, although

there have also been incremental advances in the latter

(Huang et al. 2011a; Logan and Rabaey 2012).

Oxidative recalcitrant substrates including Cr(VI),

perchlorate, 2-chlorophenol, chloroethene and penta-

chlorophenol have been successfully reduced on the

microbial cathodes (Aulenta et al. 2010; Butler et al.

2010; Huang et al. 2012, 2013; Strycharz et al. 2010;

Tandukar et al. 2009). Remaining challenges are the

recalcitrant substrate degradation rates still need to be

improved.

While appropriate substrates have been deliberately

used to stimulate the bacterial activities for efficient

CPs degradation in conventional biological processes

(Chen et al. 2012; Puyol et al. 2011; Tyagi et al. 2011)

and electrical stimulation is proved to be effective for

substrates removal in electrochemical/MFC processes

(Chun et al. 2013; Huang et al. 2011a, b; Sun et al.

2010), it is still unknown whether or not appropriate

substrate coupled with provided electrons was an

effective stimulation means for accelerating CPs

degradation rates in the newly developed biocathode

MFCs. On the other hand, CPs degradation is heavily

dependent on anaerobic/aerobic conditions. Under

anaerobic conditions, ortho-chlorines are removed at a

fast rate whereas dechlorination of para-chlorines

occurs at a slow rate (Field and Sierra-Alvarez 2008).

For example, 2,4,6-trichlorophenol (2,4,6-TCP) is

preferably transformed by ortho-dechlorinations to

4-chlorophenol (4-CP) via 2,4-dichlorophenol (2,4-

DCP), and further degraded to phenol, benzoate, and

CO2 and CH4 in anaerobic processes (Field and Sierra-

Alvarez 2008; Mun et al. 2008). Aerobic conditions,

however, result in the initial attachment of CPs with

monooxygenases and yield chlorocatechols as the first

intermediates, which are subject to ring cleavage prior

to dechlorination (Field and Sierra-Alvarez 2008). A

coupled anaerobic–aerobic condition is regarded to be

beneficial for forwarding reactions of CPs degradation

through sequential reduction and oxidation, and

consequently mineralizes CPs (Chen et al. 2010; Field

and Sierra-Alvarez 2008; Li et al. 2010; Sponza and

Ulukoy 2005). In terms of MFC systems, oxygen is

one preferred electron acceptor on the cathode because

of its availability and high redox potential (Cai et al.

2013; Logan and Rabaey 2012; Xia et al. 2013).

Presence of oxygen could affect MFC systems, also in

616 Biodegradation (2014) 25:615–632

123

case of CPs reduction. However, the effect of oxygen

on CPs degradation in the biocathode MFCs has not

been examined in detail. In addition, 2,4,6-TCP can be

successively de-chlorinated to 2,4-DCP and 4-CP, all

of which contain no meta-substitution. The choice of

2,4,6-TCP, 2,4-DCP and 4-CP as model CPs can thus

exclude the effect of these most energy demanding

positions on CPs biodegradation (Papazi and Kotzab-

asis 2013), and the experimental results will be

reasonably explained.

In this study, species of 4-CP, 2,4-DCP and 2,4,6-

TCP were respectively used to initially develop micro-

bial cathodes using the same wastewaters as inoculums

under either aerobic or anaerobic conditions. System

performances were evaluated in terms of CPs degrada-

tion, change of total organic carbon (TOC), maximum

power production, biocatalytic activity, and microbial

consortia composition. Appreciable degradation rates

of 0.15 mol/m3/d (4-CP) and 0.12 mol/m3/d (2,4-DCP)

were achieved in the anaerobic microbial cathodes. The

biocathodes well developed with 2,4-DCP under

anaerobic conditions were successfully stimulated for

efficient 2,4,6-TCP degradation at a rate up to 238 % of

the controls with no stimulation. These results provide

for the first time that the degradation rates of diverse

CPs in the developed biocathodes were greatly different

and heavily influenced by anaerobic/aerobic conditions,

and bacterial communities well developed with 2,4-

DCP can be stimulated by 2,4,6-TCP and efficiently

degrade the latter.

Materials and methods

Fuel cell assembly

A tubular two-chamber MFC (Huang et al. 2013) was

used here with graphite fiber (PANEX33 160K,

ZOLTEK) and graphite felt (a geometric surface area

of 73 cm2) as the anode and the cathode, respectively,

producing a net working volume of 43 mL in the

anode chamber and 85 mL in the cathode chamber.

Before installation, these electrode materials were

treated as previously described (Huang et al. 2013). A

reference electrode (Ag/AgCl electrode, 195 mV

versus standard hydrogen electrode [SHE]) was used

to obtain cathode and anode potentials, with all

potentials reported here versus SHE. All the anodes

and cathodes were filled with NaH2PO4–Na2HPO4

buffer (pH 7.0) and operated at room temperature

(22 ± 3 �C). Two controls (duplicate reactors) were

also operated: one was used as an abiotic control (no

inoculum) (chemical cathode and physical adsorption

processes); the other was run in an open circuit

condition (OCC) with a well developed biocathode to

examine changes in CPs in the absence of current

generation (conventional biological processes, bio-

adsorption and physical adsorption). All of the reac-

tors were wrapped in aluminum foil to exclude light.

Inoculation and operation

Seed sludge was obtained from an anaerobic digester

at a local sewage treatment plant receiving a combi-

nation of domestic and industrial wastewaters, and

used to initially inoculate both the anode and cathode

in tubular reactors. Wastewaters were initially com-

bined with an equivalent volume of nutrient solution.

Anode and cathode chambers shared the same nutrient

medium except for the addition of acetate (12.2 mM)

in the anode and CPs (ca. 0.08 mM) in the cathode.

The medium (pH 7.0) consisted of (g/L): KH2PO4, 4.4;

K2HPO4, 3.4; NaHCO3, 1.0; NH4Cl, 1.3; KCl, 0.78;

MgCl2, 0.2; CaCl2, 0.015; NaCl, 0.5; and 1.0 mL of

trace elements (Huang et al. 2013). The solutions were

sparged with ultrapure N2 gas for 20 min whereas the

headspace was filled with ultrapure N2 (anaerobic

condition). In order to create aerobic biocathode, the

cathodic headspace was filled with mixed gases

composed of N2 and O2 (60:40) and the catholyte

had no N2 sparging, resulting in 6.5–7.2 mg/L

dissolved oxygen in the catholyte. The anolyte was

renewed every 2–3 days in order to sustain the

corresponding stable potentials. In the initial acclima-

tion period, the reactor produced a gradual increase

current at each batch cycle. After 8–12 cycles with

each renewed using the mixed culture containing CPs

and wastewaters in the catholyte, and acetate and

wastewaters in the anolyte (totally lasting about

300–400 h), the voltage output increased to certain

values, and these valtages were repeatable over the

subsequent cycles, indicating the accomplishment of

electrotroph acclimation. The wastewaters were then

deleted from the electrolytes, resulting in the culture

with either CPs in the catholyte or acetate in the

anolyte. The reactors were thus subsequently run for

another 2–4 cycles before CPs degradation tests were

reported. The reactors were sealed to avoid gas losses

Biodegradation (2014) 25:615–632 617

123

and operated in fed-batch mode. All of the inoculation

and solution replacements were performed in an

anaerobic glove box (YQX-II, Xinmiao, Shanghai).

Analyses and calculation

Concentrations of 4-CP, 2,4-DCP and 2,4,6-TCP were

analyzed using a high performance liquid chromato-

graph (HPLC Agilent 1100), equipped with a C18

capillary column (4.6 mm in diameter and 250 mm in

length, ODS-2 Hypersil, Thermo). The mobile phase

was prepared by dissolving trifluoroacetic acid with

ultrapure water (pH 2.8) and the ratio of this solution

and methanol was 20:80 (v/v). TOC in the catholyte

was analyzed by SHIMADZU TOC-5000. TOC

contributed by CPs were analyzed after the sample

was filtered using 0.22 lm Millipore membrane. The

biomass in the catholyte was thus calculated, using a

mass balance, as the difference between TOC and that

after filtration. All measurements were taken over two

or three consecutive retention times.

The voltage across an external resistor of 550 Xwas recorded (30 min intervals) using a data acquisi-

tion board (PISO813, Taiwan). Linear sweep voltam-

metry (LSV) with a scan rate of 1.0 mV/s was

employed for the determination of maximum power

density, which was normalized by the volume of

catholyte. The bioelectrochemical behavior of catho-

dic biofilms was examined using cyclic voltammetry

(CV) and a three-electrode configuration with a

potentiostat (CHI 650A, Chenhua, Shanghai). The

scanned potential was between -0.8 and ?0.4 V (vs.

SHE), at a scan rate of 1.0 mV/s (Logan 2012).

The total charges transferred from the anode to the

cathode (QT) are calculated using Eq. (1):

QT ¼Z t

0

Idt ð1Þ

where current I (A) is the ratio of voltage output (V) at

operational time t (s) and the external resistance of R

(X). During CPs degradation, these electrons can be

distributed to CPs de-chlorination (Qp), oxygen

reduction (QO), bacterial growth (QG) and the lost

and unknown processes (QL) (Huang et al. 2013).

Therefore, QT = QP ? QO ? QG ? QL. Charges dis-

tributed for CP de-chlorination (this excludes contri-

bution from conventional biological processes

[OCCs]), oxygen reduction, bacterial growth and the

unknown processes are then calculated as the ratio of

the corresponding charge recoveries relative to the

total charges.

Community analysis was performed using a poly-

merase chain reaction (PCR) and denaturing gradient gel

electrophoresis (DGGE). Samples were collected from

MFCs at the end of a cycle. Electrodes were fragmented

using sterile scissors. Cells attached on the electrodes

were removed by rinsing three times with sterile water,

and concentrated by centrifugation. Genomic DNA

extraction, PCR amplification, and DGGE analyses

were performed as previously described (Huang et al.

2012; Sun et al. 2012; Xia et al. 2012).

Results and discussion

Time course of power generation and 4-CP

degradation under aerobic and anaerobic

conditions

When 4-CP was added in both aerobic and anaerobic

reactors, voltage output increased within 1.0–1.5 h

and reached the same ca 0.30 V in both aerobic

(Fig. 1a) and anaerobic (Fig. 1b) MFCs for a period of

1.0–1.5 h, and decreased thereafter (three repeatable

cycles were shown). The change of 4-CP in aerobic

and anaerobic catholytes exhibited a similar decrease

trend, which was in accordance with the decrease in

voltage output in each cycle. In terms of 4-CP

degradation, however, anaerobic MFCs exhibited

more efficient than aerobic reactors, reflecting the

adverse effect of oxygen on 4-CP degradation. This

result was in consistent with other electron acceptors

such as Cu(II), nitrate and diatrizoate, which exhibited

a competition with oxygen for the cathodic provided

electrons (Ter Heijne et al. 2010; Wrighton et al. 2010;

Radjenovic et al. 2013).

Comparison of 4-CP degradation under aerobic

and anaerobic conditions

At an identical operational period of 12 h and under

anaerobic conditions, the concentration of 4-CP

decreased from an initial 0.090 ± 0.001 to

0.014 ± 0.005 mM (0.15 mol/m3/d) whereas 4-CP

in the aerobic reactors was diminished from an initial

0.086 ± 0.001 to 0.033 ± 0.008 mM (0.11 mol/m3/

d), respectively (Fig. 2a), demonstrating the more

618 Biodegradation (2014) 25:615–632

123

efficiency of anaerobic biofilm for 4-CP degradation.

While this degradation rate was lower than the

0.345–0.597 mol/m3/d in conventional biological pro-

cesses (Milia et al. 2011; Murcia et al. 2012), the

specific 4-CP degradation rate based on a unit of

mg/g biomass/h here was 3–5 times as high as the

conventional biological processes due to the low

biomass in the present catholyte. In contrast, the

abiotic controls under either anaerobic or aerobic

conditions exhibited a similar decrease from the initial

0.081 ± 0.004 to the 0.054 ± 0.006 mM at 12 h,

mainly ascribed to both chemical reduction and

physical adsorption (Fig. 2a). The slow 4-CP degra-

dation in the abiotic controls, compared to that in the

biotic MFCs reflects the role of biofilm on 4-CP

removal. In the OCC controls and under aerobic

conditions, 4-CP was removed from the initial

0.075 ± 0.006 to 0.045 ± 0.008 mM at 12 h, slightly

higher than the net decrease of 0.027 mM in the

abiotic controls, demonstrating the role of aerobic

conventional biological processes (including biologi-

cal degradation, bio-adsorption and physical adsorp-

tion) on 4-CP removal. In the OCC controls and under

Fig. 1 Time course of power generation and 4-CP degradation

under a aerobic and b anaerobic conditions (external resistor:

550 X, three repeatable cycles)

Fig. 2 Comparison of a 4-CP degradation, b TOC decrease,

c polarization curves, and d CV tests under anaerobic and

aerobic conditions

Biodegradation (2014) 25:615–632 619

123

anaerobic conditions, however, a net 4-CP removal as

high as 0.046 mM was achieved at the same operation

time of 12 h, stressing the more beneficial anaerobic

conditions for conventional biological processes for

4-CP removal (Fig. 2a). Current generation enhanced

4-CP degradation rates under both anaerobic and

aerobic conditions, demonstrating the importance of

electrotrophic activities on 4-CP degradation. This

result was in agreement with other observations as

previously summarized (Huang et al. 2011a).

Similar to the change of 4-CP removal under

aerobic or anaerobic conditions, TOC also exhibited a

decrease trend accordingly, from an initial

0.548 ± 0.042 to 0.218 ± 0.035 mM (aerobic) and

0.551 ± 0.051 to 0.114 ± 0.041 mM (anaerobic),

respectively at 12 h (Fig. 2b). These results indicate

the ring cleavage of 4-CP and subsequent mineraliza-

tion. Both aerobic and anaerobic abiotic controls

however exhibited a slight TOC decrease, from an

initial 0.47 mM to the same 0.33 mM at 12 h

(Fig. 2b), mainly ascribed to physical adsorption and

chemical reduction, similar to the report by Gu et al.

2007. In the OCC controls, TOC exhibited a decrease

from an initial 0.526 ± 0.048 to 0.302 ± 0.044 mM

(aerobic) and 0.491 ± 0.057 to 0.222 ± 0.031 mM

(anaerobic), respectively at 12 h (Fig. 2b), demon-

strating the role of conventional biological processes

on TOC decrease.

Aerobic MFCs exhibited an OCP of 0.73 V and a

maximum power of 4.2 W/m3 (20.8 A/m3), both of

which were higher than the anaerobic reactors

(0.52 V, 2.2 W/m3, 6.7 A/m3) (Fig. 2c), implying

the dependence of OCP and maximum power on

aerobic/anaerobic conditions. In contrast, the abiotic

controls produced a low OCP of 0.15 V and a

maximum power of 0.30 W/m3 (Fig. 2c), reflecting

the electrotrophic catalysis for lowering electrode

overpotentials and thus improving power production

(Logan 2009; Sun et al. 2012). The catalytic activities

of the cathode biofilms were further confirmed using

CV. Only a single set of oxidation–reduction peaks

with different peak currents and potentials were

observed for all biofilms, compared to the weaker

peaks measured in abiotic controls (Fig. 2d), implying

the different electrotrophic capabilities under either

aerobic or anaerobic conditions. Calculated on the

basis of 4-CP removal, power productions amounting

to 0.83 kWh/mol (aerobic) and 0.38 kWh/mol (anaer-

obic) were achieved in the biocathode systems. These

power generations reflect more advantages over both

conventional electrochemical processes and microbial

electrolysis cells for 4-CP dechlorination, which

consumed energy of about 1.17 kWh/mol (Cheng

et al. 1997) and 0.097–0.55 kWh/mol (Wen et al.

2013), respectively.

Aerobic 4-CP de-chlorination attributed to elec-

tricity generation consumed 5.1 % of the total charges

transferred from the cathode whereas oxygen reduc-

tion utilized 73.3 % of the total charges, bacterial

growth consumed 20.1 % of the total charges and

1.5 % was lost to unknown processes (such as

intermediates that were not measured or unknown

extracellular polymeric substances). A higher ratio of

oxygen reduction here may indicate the role of aerobic

conditions for creating beneficial conditions for

microbial oxygen reduction with concomitant 4-CP

de-chlorination. Under anaerobic conditions, how-

ever, the charge distribution exhibited 9.3 % of total

charges in 4-CP de-chlorination, 30.2 % as bacterial

growth, with 60.5 % was lost to unknown processes.

The apparent differences in charge distribution under

aerobic and anaerobic conditions reflect the impor-

tance of aerobic and anaerobic conditions on system

performance.

Bacterial communities analyzed by DGGE showed

that the aerobic cathodes had greater richness and

more diversity than the anaerobic cathodes (Table 1;

Fig. 3), in good agreement with a previous report with

no presence of 4-CP, where aerobic instead of

anaerobic conditions were beneficial for bacterial

richness and diversity (Shehab et al. 2013). Anaerobic

less bacterial diversity was in correspondence with a

higher 4-CP degradation rate and a lower power

production, in comparison with more bacterial diver-

sity, a lower 4-CP degradation rate and a higher power

generation under aerobic conditions. A plausible

explanation is the presence of more oxygen under

aerobic conditions may have diversified electron

transfer processes and benefited to the appearance of

diverse bacteria with a capability of using oxygen (a

standard redox potential of Eh = 1.23 V) as a sole

terminal electron acceptor, and thereby generated a

higher power. This oxygen-directed process was more

or less similar to the oxygen-reducing biocathodes

(Xia et al. 2012, 2013). On the other hand, the

anaerobic biofilms developed with 4-CP may only

exhibit the specific ability of using 4-CP

(Eh = 0.21 V) as a terminal electron acceptor and

620 Biodegradation (2014) 25:615–632

123

thus a high 4-CP degradation rate with a less power

generation. In fact, little information is presently

available about the link between the electrons derived

from the cathodes and terminal electron acceptors

(Huang et al. 2011b). Even for an extensively inves-

tigated electron acceptor like oxygen, it has not yet

been demonstrated that the electron transfer is a

respiratory mechanism in which electrons derived

from the cathodes serve as an energy-yielding electron

donor for oxygen reduction. There are a variety of

other possible mechanisms by which cells might

catalyze enhanced oxygen reduction (Huang et al.

2011b). Thus, the complex electron transfer processes

involved in both oxygen reduction and 4-CP degrada-

tion catalyzed by the electrotrophs should be further

explored based on pure cultures isolated from this

mixed system. The aerobic and anaerobic cathodes

had one common and prominent band, and some

different bands (Table 1; Fig. 3), reflecting the differ-

ent microbial community structures and hence the

Table 1 DGGE 16S rRNA gene band identifications in the biocathodes acclimated with 4-CP

Condition Band Accession

no.

GenBank closest match Identity

(%)aIsolation source

Aerobic 36,31 JF800712 Uncultured bacterium 98 Degrading polyaromatic hydrocarbons

(Thavamani et al. 2012)

37 GQ458111 Uncultured bacterium 98 Bioanodes from sediment microbial fuel cells

powered by rhizodeposits of living rice plants

(De Schamphelaire et al. 2010)

38 AF423372 Uncultured CFB group

bacterium

100 A heavily polluted microbial mat and its

community changes following degradation of

petroleum compounds (Abed et al. 2002)

39 GQ451713 Nitrosomonas europaea 98 Effects of bioaugmentation on ammonia

oxidisers at a two-step WWTP (Podmirseg

et al. 2010)

40,6 HE583077 Uncultured bacterium 97 Acetate enhances startup of a H2-producing

microbial biocathode (Jeremiasse et al. 2012)

41,11,17 AJ007007 Azoarcus sp. 96 Microbial enrichments from microbial fuel cells

during wastewater treatment (Ishii et al. 2012)

42,25 DQ123737 Uncultured soil bacterium 98 Uncultivated Proteobacteria associated with

pyrene degradation in a bioreactor treating

PAH-contaminated soil (Singleton et al. 2006)

43,4,9,26,35 AF508103 Variovorax paradoxus 97 Bacteria capable of degrading phenol and

reducing nitrate (Baek et al. 2003)

Anaerobic 1 JN541141 Uncultured

Sphingobacteriales

97 Microbial community analysis in biocathode

microbial fuel cells packed with different

materials (Sun et al. 2012)

2 JN366641 Bacterium enrichment 99 Benzene degraders (van der Zaan et al. 2012)

3,7,18,34 JN674090 Comamonas sp. 96 Degrading monocyclic aromatic hydrocarbons

(Kim et al. 2002) and diclofenac/ibuprofen

(Kraigher et al. 2008)



a The values represent the similarities between the associated DGGE band sequence and the closest-match sequence from GenBank

Fig. 3 Cathode bacterial community profiles revealed by

DGGE

Biodegradation (2014) 25:615–632 621

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differences in the dechlorination activity (Fig. 2a).

Bands of 4 from anaerobic 4-CP and 43 from aerobic

4-CP shared sequences belonging to Variovorax

paradoxus, reported capable of degrading phenol

(Baek et al. 2003). This result implies both the

bacterial non-sensitivities to dissolved oxygen in the

catholyte and its potential contributions to forwarding

4-CP degradation. Five bands of 36, 38, 39, 42 and 43

from aerobic 4-CP, and two bands of 2 and 3 from

anaerobic 4-CP were closely related with bacteria

degrading recalcitrant organics of monocyclic aro-

matic hydrocarbons, diclofenac, ibuprofen or polyar-

omatic hydrocarbons (Abed et al. 2002; Kim et al.

2002; Kraigher et al. 2008; Singleton et al. 2006;

Thavamani et al. 2012). The presence of these diverse

bacteria capable of degrading multiple recalcitrant

organics explains the successful dechlorination and

mineralization of 4-CP (Fig. 2a). Bands of 37, 40 and

41 from aerobic 4-CP, and band 1 from anaerobic

4-CP were closely related to either the electrotrophs of

uncultured Sphingobacteriales (Sun et al. 2012) and

uncultured bacterium (HE583077) (Jeremiasse et al.

2012), or the exoelectrogens of Azoarcus sp. (Ishii

et al. 2012) and uncultured bacterium (GQ458111)

(De Schamphelaire et al. 2010), reflecting the non-

specific characters of these bacteria to the anode and

the cathode (Cheng et al. 2012; Huang et al. 2013; Xia

et al. 2012). The more abundance of microorganisms

in the aerobic biofilm (Table 1; Fig. 3) was not

correlated to capacities of these predominant species

for more efficient 4-CP degradation, but positively

resulted in a higher power production, partially

attributed to the presence of high redox oxygen as an

electron acceptor in the aerobic biocathodes (Sun et al.

2012; Xia et al. 2013).

Comparison of 2,4-DCP degradation under aerobic

and anaerobic conditions

Under anaerobic conditions, 2,4-DCP gradually

decreased from an initial 0.078 ± 0.0001 to

0.016 ± 0.003 mM at 12 h (0.12 mol/m3/d) (Fig. 4a),

much higher than the conventional anaerobic degra-

dation rates of 0.004 mol/m3/d (Mun et al. 2008) and

0.010–0.022 mol/m3/d (Cycon et al. 2011). The trend

of anaerobic degradation rate higher than aerobic

mode here (Fig. 4a) was similar to 4-CP (Fig. 2a) and

Fig. 4 Comparison of a 2,4-DCP degradation, b TOC decrease,

c polarization curves, and d CV tests under anaerobic and

aerobic conditions

622 Biodegradation (2014) 25:615–632

123

consistent with the conventional aerobic/anaerobic

processes (Chen et al. 2010; Field and Sierra-Alvarez

2008). Similar to 4-CP, current generation also

enhanced 2,4-DCP degradation rates under both

anaerobic and aerobic conditions, demonstrating again

the beneficial electrotrophs for 2,4-DCP removal. In

the abiotic controls, there was a slight 2,4-DCP

decrease with the prolonged operation time from an

initial 0.075 ± 0.005 to 0.061 ± 0.004 mM (anaero-

bic) and 0.074 ± 0.003 to 0.059 ± 0.005 mM (aero-

bic), respectively (Fig. 4a), mainly attributed to

chemical reduction and adsorption. 2,4-DCP was

more apparently decreased in the anaerobic OCC

controls than that in the aerobic OCC controls, from a

same initial 0.075 ± 0.005 to 0.043 ± 0.005 mM

(anaerobic) and 0.058 ± 0.004 mM (aerobic), respec-

tively. These results stress the more efficiency of

conventional anaerobic biological processes than

conventional aerobic biological processes for 2,4-

DCP removal.

Concomitant with 2,4-DCP removal, TOC exhib-

ited a gradual decrease trend under both aerobic and

anaerobic conditions, from an initial 0.51 ± 0.02 to

0.10 ± 0.03 mM (anaerobic) and 0.26 ± 0.05 mM

(aerobic) at 12 h (Fig. 4b), reflecting the ring cleavage

of 2,4-DCP and subsequent mineralization, consistent

with the extensive evidence that 2,4-DCP was miner-

alized in conventional biological processes (Field and

Sierra-Alvarez 2008). Both anaerobic and aerobic

reactors exhibited more apparent TOC decreases than

those under OCCs, stressing the importance of elec-

trotrophs on TOC removal (Fig. 4b). In the abiotic

controls, there was only slight TOC decrease under

both anaerobic and aerobic conditions compared to the

comparatively more TOC decrease in the OCC

controls, reflecting the microbial role on TOC

removal.

Aerobic MFCs exhibited a maximum power of

3.8 W/m3 (16.4 A/m3) and an OCP of 0.74 V, both of

which were apparently higher than those under

anaerobic conditions (1.7 W/m3, 4.2 A/m3, 0.56 V)

(Fig. 4c). These results imply again the dependence of

electrotrophic activities on aerobic/anaerobic condi-

tions. Power overshoot was observed under both

aerobic and anaerobic conditions (Fig. 4c), mainly

ascribed to the inability of the biofilm to produce

higher current densities, or to an inability of the

biofilm to respond to either the decreased cathode

potentials or the elevated anode potentials at higher

current densities (Zhu et al. 2013). A wider current

window up to 21.7 A/m3 was observed under aerobic

conditions, demonstrating the more efficient ability of

the biofilm to maintain the initial substrate concentra-

tion in the bulk fluid and to mass transport limitation

(Logan 2012). While electro-catalytic degradation of

2,4-DCP consumed about 3.22–4.99 kWh/mol (Chen

et al. 2011), powers of 1.3 kWh/mol (aerobic) and

0.33 kWh/mol (anaerobic) were produced from the

present systems, strongly supporting this environmen-

tal sustainable and cost-effective MFC technologies

for efficient 2,4-DCP removal.

Charges distribution in aerobic biocathodes exhib-

ited 9.4 % of the total charges for 2,4-DCP de-

chlorination, 66.2 % for oxygen reduction, 18.1 % for

bacterial growth and 6.3 % was lost to unknown

processes. Under anaerobic conditions, however, 2,4-

DCP de-chlorination consumed 17.5 % of the total

charges, bacterial growth utilized 24.8 %, and amount

to 57.7 % was lost to the unknown processes. The

considerable differences in charge distribution under

aerobic and anaerobic conditions demonstrate again

the importance of aerobic and anaerobic conditions on

system performance.

Only a single set of oxidation–reduction peaks in

the range of -0.05 to ?0.07 V were observed on the

aerobic biofilm whereas two set of oxidation–reduc-

tion peaks of -0.1 to ?0.3 V were measured on the

anaerobic biofilm (Fig. 4d), demonstrating the impor-

tance of aerobic/anaerobic conditions on the potentials

of the oxidation–reduction peaks. The sizes of oxida-

tion–reduction peaks were also affected by aerobic/

anaerobic conditions, with aerobic peaks larger than

anaerobic ones, consistent with the wider current

window under these conditions (Fig. 4c).

Bacterial communities developed under either

aerobic or anaerobic conditions showed both common

and different prominent bands (Table 2; Fig. 3).

Bands of 32 (aerobic) and 5 (anaerobic), 34 (aerobic)

and 7 (anaerobic), and 35 (aerobic) and 9 (anaerobic)

shared the similar sequences, respectively (Table 2;

Fig. 3), suggesting the non-sensitivities of these

microorganisms to oxygen. Bands of 32 (aerobic)

and 5 (anaerobic) were mostly similar to the electro-

trophs of uncultured Comamonas, implying not only

the bacterial non-sensitivities to oxygen but also its

contributions to power production (Cheng et al. 2012;

Huang et al. 2013; Sun et al. 2012). The other bands of

34 (aerobic) and 7 (anaerobic), and 35 (aerobic) and 9

Biodegradation (2014) 25:615–632 623

123

(anaerobic) were most similar to Comamonas sp. and

Variovorax paradoxus, respectively, both of which

were degraders of recalcitrant organics including

monocyclic aromatic hydrocarbons, diclofenac/ibu-

profen, and phenol (Baek et al. 2003; Kim et al. 2002;

Kraigher et al. 2008). These results imply the

independence of the bacterial survival on aerobic/

anaerobic conditions and reflect the bacterial contri-

bution to 2,4-DCP degradation. The extensive pre-

sence of Variovorax paradoxus in all of the aerobic

4-CP (band 43), the anaerobic 4-CP (band 4), the

aerobic 2,4-DCP (band 35) and the anaerobic 2,4-DCP

(band 9) (Fig. 3) implies the bacterial importance to

degradation of both 4-CP and 2,4-DCP. More exo-

electrogens including uncultured Bacteroidetes (band

10) (Sun et al. 2012) and uncultured bacterium

(HE583077) (band 6) (Jeremiasse et al. 2012) were

observed under anaerobic conditions (Table 2;

Fig. 3). This result illustrates the more abundance of

exoelectrogens in anaerobic biofilm was not correlated

to capacities of these predominant species for high

power production (Fig. 4c), in consistence with other

reports (Shehab et al. 2013; Sun et al. 2012). Band 31

from aerobic 2,4-DCP and band 36 from aerobic 4-CP

were highly similar to uncultured bacterium

(JF800712), degrader of polyaromatic hydrocarbons

(Thavamani et al. 2012). Similarly, band 34 (aerobic

2,4-DCP), band 7 (anaerobic 2,4-DCP) and band 3

(anaerobic 4-CP) identically shared the same

sequences with Comamonas sp., efficiently degrading

Table 2 DGGE 16S rRNA gene band identifications in the biocathodes acclimated with 2,4-DCP


no.

GenBank closest match Identity


Aerobic 30 FJ756565 Uncultured Sphingobacterium

sp.

98 An atrazine-degrading culture in response to

high atrazine input (Udikovic-Kolic et al.

2011)

31,36 JF800712 uncultured bacterium 96 Bacterial community in soils polluted with

polyaromatic hydrocarbons (Thavamani et al.

2012)

32,5,16,20 JN541134 Uncultured Comamonas 97 Microbial community analysis in biocathode



33,24 FR682925 Delftia sp. R-41380 98 Biofilm for species selection and pesticide

degradation (Verhagen et al. 2011)

34,3,7,18 JN674090 Comamonas sp. 98 Bacteria degrading monocyclic aromatic

hydrocarbons (Kim et al. 2002) and diclofenac/

ibuprofen (Kraigher et al. 2008)



Anaerobic 5,16,20,32 JN541134 Uncultured Comamonas 96 Microbial community analysis in biocathode



6,40 HE583077 Uncultured bacterium 97 Acetate enhances startup of a H2-producing

microbial biocathode (Jeremiasse et al. 2012)

7,3,18,34 JN674090 Comamonas sp. 98 Bacteria degrading monocyclic aromatic

hydrocarbons (Kim et al. 2002) and diclofenac/

ibuprofen (Kraigher et al. 2008)

8,13 AF250407 Cytophaga sp. D2 100 Genetic diversity of carbofuran-degrading soil

bacteria (Desaint et al. 2000)



10 JN541132 Uncultured Bacteroidetes 97 Microbial community in biocathode MFCs

packed with different materials (Sun et al.

2012)


624 Biodegradation (2014) 25:615–632

123

monocyclic aromatic hydrocarbons (Kim et al. 2002)

and diclofenac/ibuprofen (Kraigher et al. 2008). These

results demonstrate the bacterial possible roles in the

subsequent 2,4-DCP degradation (Field and Sierra-

Alvarez 2008). The sequences of 30 and 33 from

aerobic 2,4-DCP, and 9 from anaerobic 2,4-DCP

(Table 2; Fig. 3) were most similar to uncultured

Sphingobacterium sp., Delftia sp. R-41380, and

Cytophaga sp. D2, respectively, each of which was

dominant in a microbial community for treating

atrazine, pesticide, or carbofuran (Desaint et al.

2000; Udikovic-Kolic et al. 2011; Verhagen et al.

2011). The presence of these diverse bacteria that were

capable of degrading multiple recalcitrant organics or

exhibited exoelectrogenic/electrotrophic activities,

can therefore explain the efficient 2,4-DCP degrada-

tion with simultaneous power production from the

biocathode MFCs.

Comparison of 2,4,6-TCP degradation

under aerobic and anaerobic conditions

At an operational period of 12 h, 2,4,6-TCP was

degraded at similar rates of around 0.042 mol/m3/d

under aerobic/anaerobic conditions, only slightly

higher than 0.029 mol/m3/d in the abiotic controls

and 0.031 mol/m3/d in the OCC controls (Fig. 5a).

These results demonstrate the slight positive role of

electrotrophs on 2,4,6-TCP removal. These degrada-

tion rates were lower than the 0.069–0.17 mol/m3/d in

conventional biological processes (Karn and Balda

2013; Karn and Reddy 2012), implying less efficiency

of this acclimated microbial consortia for 2,4,6-TCP

degradation. Concomitant with 2,4,6-TCP degrada-

tion, aerobic reactors achieved an OCP of 0.77 V,

substantially higher than the 0.26 V in the aerobic and

abiotic controls (Fig. 5b), confirming the catalysis of

the involved microorganisms for high OCPs. Greatly

different from a maximum power of 8.7 W/m3

(15.6 A/m3) produced from aerobic reactors, anaero-

bic MFCs showed a comparatively low maximum

power of 2.3 W/m3 (10.7 A/m3), illustrating low

electrotrophic activities on the cathodes. CV analysis

of both aerobic and anaerobic reactors showed the

presence of two oxidation–reduction peaks with the

anaerobic biocathodes, one comparatively strong

oxidation–reduction peak with aerobic reactors, and

one very small peak in the abiotic controls (Fig. 5c).

The magnitude of the oxidation–reduction peaks in the

aerobic reactors reflects the more efficient electro-

chemical activities of the biocathodes (Fig. 5b).

The microbial communities developed under either

aerobic or anaerobic conditions were very different

(Table 3; Fig. 3). In the anaerobic reactors, the

sequences of bands 11 and 14 were most similar to

the exoelectrogens of Azoarcus sp. and Desulfovibrio

intestinalis, respectively (Ishii et al. 2012; Kim et al.

2006) whereas band 13 shared a phylogenetic relation

to Cytophaga sp. D2 and band 12 to uncultured

Bacteroidetes sp., which degraded either carbofuran or

industrial waste gases (Desaint et al. 2000; Friedrich

Fig. 5 Comparison of a 2,4,6-TCP degradation, b TOC

decrease, c polarization curves, and d CV tests under anaerobic

and aerobic conditions

Biodegradation (2014) 25:615–632 625

123

et al. 2002). In addition, band 13 shared the same

sequence as band 8 from anaerobic 2,4-DCP (Table 3;

Fig. 3), implying its potential contribution to 2,4,6-

TCP degradation via the formation of 2,4-DCP.

Compared to the anaerobic 2,4,6-TCP bacterial com-

munities, the predominant bacteria detected on the

aerobic cathodes were mostly affiliated with degraders

of recalcitrant organics including pyrene (Singleton

et al. 2006), phenol (Baek et al. 2003), terephthalic

acid (Perkins et al. 2011), trichloroethylene (Futamata

et al. 2005), 2,4-dinitroanisole, and n-methyl-4-nitro-

aniline (Arnett et al. 2009) (Table 3; Fig. 3). In

addition, bands 24 and 25 were also found in aerobic

2,4-DCP (band 33) and aerobic 4-CP (band 42),

respectively, implying their potential contributions to

2,4,6-TCP degradation via the formations of both 2,4-

DCP and 4-CP. Similarly, band 26 sharing most

similar sequences with phenol degrader of Variovorax

paradoxus (Baek et al. 2003) and frequently found in

aerobic/anaerobic 4-CP/2,4-DCP reactors by bands 4,

9, 35 and 43 (Fig. 3) in concert may also explain the

facultative anaerobe contribution to the successive

2,4,6-TCP degradation. Band 20 belonged to exoelec-

trogens (Ishii et al. 2012) and was also present in

aerobic/anaerobic 2,4-DCP reactors (band 32 and 16),

implying its potential contribution to both power

production (Fig. 5b) and 2,4,6-TCP degradation via

the formation of 2,4-DCP. Considering the similar

Table 3 DGGE 16S rRNA gene band identifications in the biocathodes acclimated with 2,4,6-TCP


no.

GenBank closest

match

Identity


Aerobic 19 GQ285919 Uncultured bacterium 98 Degrading 2,4-dinitroanisole and n-methyl-4-

nitroaniline (Arnett et al. 2009)

20,32,16,5 JN541134 Uncultured Comamonas 96 Microbial community analysis in biocathode

microbial fuel cells packed with different materials

(Sun et al. 2012)

21 GQ263620 Uncultured bacterium 97 Bacterial community at a simulated low-level-

radioactive-waste site (Field et al. 2010)

22 AB205617 Uncultured bacterium 95 Bacterial community for denitrification of saline

industrial wastewater (Yoshie et al. 2006)

23 GQ263300 Uncultured bacterium 99 Bacterial community at a simulated low-level-

radioactive-waste site (Field et al. 2010)

24,33 FR682925 Delftia sp. R-41380 96 Biofilm for species selection and pesticide

degradation (Verhagen et al. 2011)

25,42 DQ123737 Uncultured bacterium 98 Bacteria associated with pyrene degradation in a

bioreactor (Singleton et al. 2006)

26,4,9,35,43 AF508103 Variovorax paradoxus 96 Bacteria capable of degrading phenol and reducing

nitrate (Baek et al. 2003)

27 JQ607838 Bacterium NLAE-zl-C99 97 Bacterial community in swine feces and stored

manure (Ziemer et al. 2009)

28 GQ138996 Uncultured bacterium 97 From upflow anaerobic bioreactors treating

terephthalic acid wastewater (Perkins et al. 2011)

29 AB167229 Brevundimonas diminuta 98 Bacteria responsible for trichloroethylene

degradation (Futamata et al. 2005)

Anaerobic 11,17,41 AJ007007 Azoarcus sp. 98 Microbial enrichments from MFCs during

wastewater treatment (Ishii et al. 2012)

12 AJ318144 Uncultured Bacteroidetes

sp.

96 A waste gas-degrading community in an industrial

biofilter (Friedrich et al. 2002)

13,8 AF250407 Cytophaga sp. D2 97 Genetic diversity of carbofuran-degrading soil

bacteria (Desaint et al. 2000)

14 AJ630285 Desulfovibrio intestinalis 97 Exoelectrogens bacterial community in a MFC (Kim

et al. 2006)


626 Biodegradation (2014) 25:615–632

123

2,4,6-TCP degradation under aerobic and anaerobic

conditions (Fig. 5a) together with the anaerobic more

efficient 2,4-DCP degradation (Fig. 4b), microbial

consortia well developed with anaerobic 2,4-DCP was

used for the subsequent stimulation for efficient 2,4,6-

DCP degradation.

Stimulation of 2,4-DCP acclimated microbial

consortia for 2,4,6-TCP degradation

At an operational period of 12 h, microbial consortia

developed with anaerobic 2,4-DCP cultivation effi-

ciently degraded 2,4,6-TCP at a rate of 0.10 mol/m3/d,

faster than the 0.042 mol/m3/d in the controls with no

stimulation (138 % improvement) (Fig. 6a) and com-

parative to the highest value of 0.17 mol/m3/d in

conventional biological processes (Karn and Balda

2013; Karn and Reddy 2012). In the OCCs controls,

however, this anaerobic 2,4-DCP developed microbial

consortia had a similar degradation ability with the

2,4,6-TCP acclimated bacterial communities (no

stimulation) under CCCs conditions, and higher than

both no-stimulated OCCs and abiotic CCCs controls

(Fig. 6a), reflecting the importance of both cathodic

provided electrons and stimulated microbial consortia

for efficient 2,4,6-TCP degradation.

TOC in the stimulated reactors experienced more

apparent decrease than the OCCs controls (Fig. 6b),

illustrating the importance of electrotrophs in the

biofilm on TOC removal. This decrease trend was

consistent with the change of 2,4,6-TCP (Fig. 6a),

implying the ring cleavage and mineralization of

2,4,6-TCP in the systems. In addition, less TOC

decrease was observed in all three controls of no-

stimulated CCCs, no-stimulated OCCs and abiotic

CCCs, reflecting again the importance of both

cathodic provided electrons and stimulated microbial

consortia on TOC removal.

The maximum power produced by the anaerobic

stimulated MFCs reached 2.6 W/m3 (11.7 A/m3),

higher than 2.3 W/m3 (10.7 A/m3) produced by the

controls with no stimulation (13 % improvement)

(Fig. 6c). In the abiotic controls with substrate shift

from 2,4-DCP to 2,4,6-DCP, only around 0.09 W/m3

(0.99 A/m3) was obtained, reflecting the importance

of stimulated biofilm on electricity generation.

Two sets of oxidation–reduction peaks were

observed for the stimulated biofilm, more apparent

than the biofilm with no stimulation (Fig. 6d). For the

Fig. 6 a 2,4,6-TCP degradation, b TOC decrease, c polarization

curves, and d CV tests before and after 2,4-DCP developed

microbial consortia stimulated by 2,4,6-TCP

Biodegradation (2014) 25:615–632 627

123

stimulated biofilm, the oxidation peaks were at

-0.05 V (0.134 mA) and ?0.33 V (0.357 mA) whereas

the reduction peaks were at -0.10 V (-0.191 mA)

and ?0.01 V (-0.186 mA), respectively. For the no

stimulation controls, however, one set of oxidation–

reduction peaks substantially appeared at 0.04 V

(0.168 mA) and -0.06 V (-0.277 mA) along with

the other small set of peaks at 0.37 V (0.261 mA) and

0.07 V (-0.106 mA) (Fig. 6d). These results suggest

that this stimulation had a substantial impact on both

the potential and the size of the oxidation–reduction

peaks and thus changed the bacterial activities.

Bacterial community analysis showed band 16 in

the stimulated biofilm shared the same sequence with

band 5 in anaerobic 2,4-DCP biofilm and band 20 in

aerobic 2,4,6-TCP biofilm (Table 4; Fig. 3), all of

which were closely related with the Burkholderia sp.,

capable of degrading indole (Hong et al. 2010). These

results indicate the bacterial robust survival under

either anaerobic 2,4-DCP or aerobic 2,4,6-TCP con-

ditions, and its potential contribution to degrading 2,4-

DCP and 2,4,6-TCP. Similarly, band 17 in the

stimulated biofilm shared the same sequence of band

11 in anaerobic 2,4,6-TCP biofilm (Table 4; Fig. 3),

both of which were highly related with Azoarcus sp.,

constantly present in anaerobic microbial consortia

not only degrading multiple recalcitrant organics of

alkane, toluene, and penicillin (Ehrenreich et al. 2000;

Juteau et al. 1999; Li et al. 2009), but also exhibiting

exoelectrogenic activities (Ishii et al. 2012). In

addition, band 18 in the stimulated biofilm shared

the same sequence as band 34 in aerobic 2,4-DCP

biofilm, band 7 in anaerobic 2,4-DCP biofilm, and

band 3 in anaerobic 4-CP biofilm (Table 4; Fig. 3), all

of the four were affiliated with the domain of

Comamonas sp., degrading recalcitrant organics of

chlorinated aromatic hydrocarbons (Kim et al. 2002)

and diclofenac/ibuprofen (Kraigher et al. 2008). These

results strongly support the bacterial potential contri-

bution to 2,4,6-TCP degradation and mineralization.

It is not clear if the CPs degraders were growing

using electrons transferred from electrotrophic bacte-

ria on the cathodes. In addition to a preference for CPs

substrates in conventional biological processes, some

CPs degraders require growth factors provided by

other microorganisms although this factor has not

been identified (May et al. 2008). This mutualism and

interspecies cooperation as well as possible syner-

gisms among different types of bacteria (Chen et al.

2013) thus cannot be excluded in order to forward

stepwise CPs degradation, sustain bacterial growth,

and generate power. While electrically conductive pili

have been found to connect a fermentative bacterium

and a methanogen (Logan 2009; Logan and Rabaey

2012), the electrically conductive graphite felts here

may help provide an electrical conduit between these

microorganisms, allowing the growth of both CPs

degraders and electrotrophic bacteria. This graphite

felt may thus have been beneficial for the formation of

biofilms, similar to the role of sediment on reductive

dechlorination in conventional biological processes

(Tyagi et al. 2011). The electrotrophic activities and

their roles in CPs dechlorination processes, and

alternatively, the bacterial CPs-degrading activities

and their roles in electricity generation remain

unknown. In fact, not all the members of the commu-

nity were responsible for cathode respiration, since

fermentation and other respiratory processes allowed

Table 4 DGGE 16S rRNA gene band identifications in the 2,4,6-TCP stimulated biocathodes after developed with anaerobic 2,4-

DCP

Band Accession

no.

GenBank

closest match

Identity


15 EU136281 Uncultured

bacterium

98 Bacterial communities in indole-degrading bioreactors (Hong et al. 2010)

16,5,20,32 JN541134 Uncultured

Comamonas

97 Microbial community analysis in biocathode microbial fuel cells packed with

different materials (Sun et al. 2012)

17,11,41 AJ007007 Azoarcus sp. 98 Microbial enrichments from microbial fuel cells during wastewater treatment

(Ishii et al. 2012)

18,3,7,34 JN674090 Comamonas

sp.

96 Bacteria degrading monocyclic aromatic hydrocarbons (Kim et al. 2002) and

diclofenac/ibuprofen (Kraigher et al. 2008)


628 Biodegradation (2014) 25:615–632

123

different organisms to proliferate (Kiely et al. 2010;

Logan 2009; Yuan et al. 2013). It is also not clear what

role numerically less abundant bacteria might play in

current generation although pyrosequencing results

have been found to be generally consistent with clone

libraries and this approach provides an additional

opportunity to probe more deeply into the community

members (Logan 2012). In addition, some exoelec-

trogenic bacteria can also be missed in biofilm analysis

(Logan 2012; Yuan et al. 2013). Current knowledge of

bacterial communities contributing to biocathode

processes is limited to denitrifying (Wrighton et al.

2010), perchlorate (Butler et al. 2010), pentachloro-

phenol (Huang et al. 2012) and oxygen reduction (Xia

et al. 2012). Further investigation of the electrotrophic

activities of bacteria developed by the specific CPs

under aerobic/anaerobic conditions, as well as the

stimulated microbial consortia with pure cultures is

still needed.

Conclusions

Biocathodes developed under anaerobic conditions

achieved higher degradation rates of 0.15 mol/m3/d

(4-CP) and 0.12 mol/m3/d (2,4-DCP) than those in

aerobic MFCs whereas aerobic reactors exhibited

higher maximum powers of 4.2 W/m3 (20.8 A/m3,

4-CP) and 3.8 W/m3 (16.4 A/m3, 2,4-DCP) than

anaerobic MFCs. The activities of bacterial commu-

nities well developed with 2,4-DCP were successfully

stimulated for efficient 2,4,6-TCP degradation and

high power production, 138 and 13 % improvements,

respectively compared to the controls with no stimu-

lation. Bacterial communities developed by the spe-

cific CPs under aerobic/anaerobic conditions, as well

as the stimulated microbial consortia shared some

dominant genera and also exhibited great differences.

These results demonstrate that anaerobic/aerobic

conditions can affect CPs degradation with power

generation from biocathode MFCs, and using deliber-

ate substrates can stimulate the developed microbial

consortia and be potentially feasible for the selection

of an appropriate microbial community for the target

substrate (e.g. 2,4,6-TCP) degradation in the biocath-

ode MFCs.

Acknowledgments The authors gratefully acknowledge

financial support from the Natural Science Foundation of

China (Nos. 21077017 and 51178077) and Specialized Research

Fund for the Doctoral Program of Higher Education ‘‘SRFDP’’

(No. 20120041110026).

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