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Biodegradation ISSN 0923-9820Volume 24Number 2 Biodegradation (2013) 24:215-225DOI 10.1007/s10532-012-9578-1

Biodegradation of DDT by stimulation ofindigenous microbial populations in soilwith cosubstrates

Irmene Ortíz, Antonio Velasco, Sylvie LeBorgne & Sergio Revah

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ORIGINAL PAPER

Biodegradation of DDT by stimulation of indigenousmicrobial populations in soil with cosubstrates

Irmene Ortız • Antonio Velasco •

Sylvie Le Borgne • Sergio Revah

Received: 10 February 2012 / Accepted: 17 July 2012 / Published online: 31 July 2012

� Springer Science+Business Media B.V. 2012

Abstract Stimulation of native microbial popula-

tions in soil by the addition of small amounts of

secondary carbon sources (cosubstrates) and its effect

on the degradation and theoretical mineralization of

DDT [l,l,l-trichloro-2,2-bis(p-chlorophenyl)ethane]

and its main metabolites, DDD and DDE, were

evaluated. Microbial activity in soil polluted with

DDT, DDE and DDD was increased by the presence of

phenol, hexane and toluene as cosubstrates. The

consumption of DDT was increased from 23 % in a

control (without cosubstrate) to 67, 59 and 56 % in the

presence of phenol, hexane and toluene, respectively.

DDE was completely removed in all cases, and DDD

removal was enhanced from 67 % in the control to

*86 % with all substrates tested, except for acetic

acid and glucose substrates. In the latter cases, DDD

removal was either inhibited or unchanged from the

control. The optimal amount of added cosubstrate was

observed to be between 0.64 and 2.6 mg C g�1drysoil. The

CO2 produced was higher than the theoretical amount

for complete cosubstrate mineralization indicating

possible mineralization of DDT and its metabolites.

Bacterial communities were evaluated by denaturing

gradient gel electrophoresis, which indicated that

native soil and the untreated control presented a low

bacterial diversity. The detected bacteria were related

to soil microorganisms and microorganisms with

known biodegradative potential. In the presence of

toluene a bacterium related to Azoarcus, a genus that

includes species capable of growing at the expense of

aromatic compounds such as toluene and halobenzo-

ates under denitrifying conditions, was detected.

Keywords Organochlorine pesticides � DDT �Biostimulation � Cosubstrates � Biodegradation

Introduction

DDT [l,l,l-trichloro-2,2-bis(p-chlorophenyl)ethane]

was first synthesized in the late 1930s, and its

remarkable insecticidal activity promptly resulted in

large-scale worldwide production (Zitko 2003). DDT

production in Mexico was around 1,200 tons per year

in the 1980s and 1990s when production was prohib-

ited. Presently, DDT use is restricted to malaria

control by the Mexican Health authorities; however,

DDT was used between 1950 and 1990 in tropical

areas for agricultural and health purposes. Sites

I. Ortız (&) � S. Le Borgne � S. Revah

Departamento de Procesos y Tecnologıa, Universidad

Autonoma Metropolitana-Cuajimalpa, Artificios 40,

Col. Hidalgo, Delegacion Alvaro Obregon,

01120 Mexico, DF, Mexico

e-mail: irmene@correo.cua.uam.mx

A. Velasco

Centro Nacional de Investigacion y Capacitacion

Ambiental, Instituto Nacional de Ecologıa,

Mexico, DF, Mexico

123

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DOI 10.1007/s10532-012-9578-1

Author's personal copy

contaminated with DDT remain as a result of its

extremely long persistence in soils ([10 years), which

presents a serious pollution problem in some areas of

Mexico (Dıaz-Barriga et al. 2003). The lipophilic

properties of DDT allow it to accumulate in the food

chain and in the fatty tissues of organisms, affecting

the central neural system, liver and kidneys, and

causing reproductive disorders (US-DHS 2002).

In natural environments, DDT can be degraded

both by physicochemical and biological processes. As

DDT is transformed, increased levels of DDD [l,1-

dichloro-2,2-bis(p-chlorophenyl)ethane] and DDE [1,1-

dichloro-2,2-bis(p-chlorophenyl)ethylene], the main

intermediates of DDT transformation by anaerobic

and aerobic microbial processes, are observed (Xiao

et al. 2011; Fang et al. 2010). These compounds are

considered ubiquitous in the environment and are often

reported as more recalcitrant and toxic than DDT

(Sinclair and Boxall 2003). Soil solidification/stabiliza-

tion, flushing, incineration, absorption, treatment using

chemical (e.g. Fenton reaction) and biochemical (e.g.

surfactants and nutrients addition) techniques, have

been reported to reduce the content of DDT and its

metabolites in soils (Castelo-Grande et al. 2010; Dalla

Villa and Pupo Nogueira 2006; Purnomo et al. 2010;

Shareef and uz Zaman 2010; Tian et al. 2008; Walters

and Aitken 2001). The biological degradation of these

compounds under aerobic and anaerobic conditions by

bacterial and fungal communities has also been

described (Aislabie et al. 1997; Castelo-Grande et al.

2010; Foght et al. 2001; Kamanavalli and Ninnekar

2004; Kantachote et al. 2004; Purnomo et al. 2008;

Purnomo et al. 2010; Zhao et al. 2010).

DDT and other polychloroaromatic compounds

have been described as recalcitrant molecules and

researches often face difficulty when trying to isolate

organisms capable of utilizing them as sole sources of

carbon and energy (Fang et al. 2010). However, co-

metabolism of these compounds has been shown to

occur under laboratory conditions and could be a

significant process in the removal of these pesticides

from the environment (Aislabie et al. 1997; Singh et al.

1999). In addition, co-metabolism of pesticides by

natural microbial communities may result in their

complete degradation (Fetzner 1998; Castelo-Grande

et al. 2010; Deepthi and Manonmani 2007). Thus, the

rationale behind the use of more soluble or biode-

gradable cosubstrates to promote the degradation of

persistent molecules is that these cosubstrates can

foster microbial growth and induce the activation of

enzymes that participate in pollutant degradation

(Ortız et al. 2003; Ortız et al. 2006; Purnomo et al.

2010; Harder and Dijkhuizen 1982).

For instance, toluene is a cosubstrate able to support

the co-metabolism of trichloroethylene (TCE) and other

chlorinated compounds by soil microbial communities

and it has been reported that toluene monooxygenases

with broad substrate specificity have the ability to

degrade these compounds (Chauhan et al. 1998; Mu and

Scow 1994; Parales et al. 2000). Furthermore, toluene

injection has been successfully evaluated in full-scale

for in situ cometabolic degradation of TCE in ground-

water (McCarty et al. 1998). Also, some studies have

demonstrated the possible degradation of chlorinated

compounds by toluene-oxidizing bacteria in the absence

of aromatic cosubstrates (Ortız et al. 2003; Ortız et al.

2006; Yeager et al. 2004).

Moreover, organic amendments such as glucose,

plant matter or analogues compounds may allow

microflora to proliferate, and under proper conditions,

oxidation of the amendment generates a reducing

environment under which pollutants can be trans-

formed (Foght et al. 2001).

The objective of the present study was to evaluate

the degradation and mineralization of DDT, DDD

and DDE in microcosm experiments using different

cosubstrates to stimulate indigenous microbial popu-

lations in soils. The cosubstrates tested (Table 1) were

a linear hydrocarbon (hexane); an aromatic hydrocar-

bon (toluene); a phenol (phenol); a carboxylic acid

(acetic acid); an alcohol (ethanol) and a monosaccha-

ride (glucose). Furthermore, the bacterial diversity in

soil before and after cosubstrate addition was evaluated

by denaturing gradient gel electrophoresis (DGGE).

Materials and methods

Soil

Soil samples were collected from rural areas in the

state of Chiapas, Mexico. The initial concentrations

of DDT, DDE and DDD, in mg kg�1drysoil, were 7.46 ±

0.66, 3.31 ± 0.04 and 0.42 ± 0.01, respectively. The

soil composition in % (w/w) was 6.9 ± 0.34 carbon,

0.38 ± 0.09 hydrogen and 0.69 ± 0.03 nitrogen,

and the soluble organic carbon was 0.47 g kg�1drysoilas

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quantified by elemental analysis (CHNS analyzer

2400 series II, Perkin Elmer, USA). The soil was

sieved and the particle size used in experiments was

\0.64 mm.

For abiotic controls, soil was sterilized by auto-

claving at 120 �C for 15 min; this operation was

repeated for three consecutive days. After this steril-

ization process the concentrations of DDT, DDE and

DDD were evaluated in order to quantify losses

produced by heat, recovery after sterilization were

94.04, 98.7 and 93.84 %, respectively; these values

were in the range of extraction recovery.

Mineral medium

Commercial fertilizer (Triple 17�, Nutrimentos Mine-

rales de Hidalgo, Mexico) containing 17 % nitrogen

(N), 17 % phosphorous (P2O5) and 17 % potassium

(K2O) at a concentration of 0.6 g l-1 and a pH of 7

was used as a source of inorganic nutrients.

Microcosm experiments

DDT biodegradation was studied in triplicate in

microcosms (125-ml hermetic flasks) containing slur-

ries of 5 g polluted soil and 10 ml mineral medium

with the addition of one cosubstrate. Reactive grade

toluene, hexane, phenol, acetic acid, ethanol and

glucose were tested as cosubstrates. The incubation

conditions were 30 �C and 150 rpm. Controls con-

sisted of experiments with mineral medium but

without cosubstrate addition.

Abiotic controls were also prepared with and

without volatile cosubstrates (hexane and toluene)

using sterilized soil and with the addition of 1.3 g l-1

of sodium azide as microbial inhibitor (Ortız et al.

2003). These experiments were conducted for 7 weeks

(1,200 h). The cosubstrates were added every week,

which amounts to a total addition of cosubstrate

equivalent to 1.48 ± 0.25 mgcarbon g�1drysoil.

Influence of the cosubstrate concentration

Further slurry experiments varying the dose (amount

of cosubstrate added) were conducted with toluene

and phenol, which showed the best degradation

results. In order to verify if the effect could be

sustained with a lower amount of cosubstrate or if the

effect could be enhanced with a higher amount of

cosubstrate, the doses used were approximately dou-

ble and half of that used in previous experiments.

The cosubstrates were added weekly. For the low

dose, the equivalent of 0.21 mgcarbon g�1drysoil was added

Table 1 Effect of different cosubstrates tested on DDT degradation and CO2 production

aCosubstrate DDT elimination (%) DDD elimination (%) bCarbon recovered as CO2 [%]

Toluene 56.17 ± 8.36 85.27 ± 8.20 85.37 ± 5.71

Hexane 59.63 ± 7.36 85.71 ± 7.27 72.96 ± 5.70

Phenol 67.16 ± 8.06 86.14 ± 1.26 88.96 ± 3.9

Acetic acid ND 47.57 ± 7.58 47.06 ± 10.9

Ethanol 49.63 ± 1.41 85.86 ± 1.23 63.46 ± 0.7

Glucose 50.48 ± 11.40 69.17 ± 5.01 54.58 ± 2.7

Controld 23.5 ± 8.70 67.57 ± 4.20 c

Abiotic controld 1.87 ± 0.14e ND ND

Time of treatment: 50 days. The initial DDT, DDE and DDD concentration, in mg kg�1drysoil: 7.46 ± 0.66, 3.31 ± 0.04 and

0.42 ± 0.01, respectively. Standard deviation calculated with n = 6

ND Not Detecteda 1.43 mg of carbon added per g of dry soil in all casesb Calculated from degradation of DDE, DDT and DDD, assuming 100 % cosubstrate degradation and subtracting the endogenous

respirationc Endogenous respirationd Without cosubstratee Abiotic losses

Biodegradation (2013) 24:215–225 217

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on weeks 1, 2, and 3; for the intermediate dose,

0.36 mgcarbon g�1drysoil on weeks 1, 2, 3 and 4 was added;

and for the high dose, 0.47 mgcarbon g�1drysoil each week

for the 7 week duration of the experiment was added.

The total amounts added were 0.64, 1.43 and 3.32 mgcar-

bon g�1drysoil for the low, intermediate and high dose

experiments, respectively. The higher amount of

cosubstrates was fixed in the rage of reported degrada-

tion to guarantee their complete degradation (Mendonca

et al. 2004; Ortız et al. 2006).

DDT degradation rate

These experiments were prepared in triplicate for both

low and intermediate doses of toluene to evaluate only

DDT degradation. Two sets of controls were per-

formed, (a) control experiments without addition of

cosubstrate and, (b) abiotic controls with and without

cosubstrates. Three bottles were sacrificed for DDT

quantification, weekly for bottles amended with

toluene and every 2 weeks for control experiments.

The complete contents of each bottle were used for

DDT determination. At time zero, three bottles were

used to quantify initial DDT concentration. The

experiment was performed for 7 weeks with the

addition of toluene on a weekly basis. Toluene

consumption and CO2 production in the gas phase

were quantified as an indirect measurement of aerobic

microbial activity. The CO2 production in experiments

with cosubstrate was estimated as follows, the amount

of CO2 measured in test experiments minus the CO2 by

endogenous activity (control without cosubstrate). In

order to estimate a possible amount of CO2 attributed

to degradation of DDT, DDD and DDE, the theoretical

amount of CO2 that would be produced from miner-

alization of 100 % of the cosubstrate was deduced and

the production of biomass was not considered (Ortız

et al. 2003).

Analytical methods

Toluene and hexane concentrations were measured in

100 ll gaseous samples in duplicate using headspace

by gas chromatography with flame ionization detector

(HP 5890 Series II, USA) on a HP624 column

(Agilent, USA). The temperatures of the detector

and injector were 200 and 220 �C, respectively, while

the oven temperature was 120 and 80 �C for toluene

and hexane, respectively. Nitrogen was used as the

carrier gas.

CO2 concentrations were measured in 100 ll

gaseous samples in duplicate by gas chromatography

equipped with a thermal conductivity detector (GOW

MAC Series 550, USA) and a concentric column

CTR1 (Alltech, USA). The oven, detector and injector

temperatures were fixed at 40, 100 and 30 �C,

respectively. Helium was used as carrier gas (Ortız

et al. 2003).

Residual DDT, DDE and DDD were extracted by

sonication (US-EPA 2007b) using the complete con-

tents of each flask, the extraction recovery from soil

was above 96 % with this methodology. The quanti-

fication was performed by gas chromatography (US-

EPA 2007a) with an electron capture detector (Varian

3400, USA) equipped with a J&W Scientific DB-5

column (Agilent, USA). The temperatures of the

detector and injector were 300 and 250 �C, respec-

tively, while oven initial temperature was 160 �C and

increased up to final temperature of 240 �C, at a rate of

5 �C min-1. Nitrogen was used as carrier gas.

DNA extraction, PCR primers and DGGE analysis

Total DNA was extracted from soils (250 mg) treated

for 7 weeks, in triplicate using the UltraClean Soil

DNA isolation kit (MoBio Laboratories, USA). DNA

concentrations were determined with a NanoDrop

spectrophotometer (ND-1000; Nanodrop Technology,

USA). A nested PCR approach was used. The primers

Bac8f (50-AGA GTT TGA TCC TGG CTC AG-30)and 1492r (50-GGT TAC CTT GTT ACG ACT T-30)were used to amplify the complete 16S rRNA gene.

The primers gc-338f (50-CGC CCG CCG CGC GCG

GCG GGC GGG GCG GGG GCA CGG GGG GCC

TAC GGG AGG CAG CAG-30) and 907r (50-CCG

TCA ATT CCT TTG AGT TT-30) were used to

amplify the bacterial V3-V5 region for DGGE. For the

first PCR, each 25 ll reaction mixture contained 5 ng

DNA template, 19 ThermoPol reaction buffer (New

England Biolabs, Ipswich, MA), 0.2 mM of each

dNTP, 0.5 lM of each primer, and 1.25 U Taq DNA

polymerase (New England Biolabs). The reaction

conditions were (i) 2 min at 95 �C, (ii) 35 cycles of

45 s at 94 �C, 30 s at 56 �C, 1 min at 72 �C, and (iii)

5 min at 72 �C in a TC-3000 thermal cycler (Techne,

USA). The PCR products were purified with the

UltraClean PCR Clean-up DNA purification kit

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(MoBio Laboratories) and used for a second round of

PCR. Each 25 ll reaction mixture contained 10 ng

purified PCR product, 19 ThermoPol II reaction

buffer (New England Biolabs), 0.2 mM of each dNTP,

0.5 lM of each primer, 2 mM MgSO4, 0.67 mg ml-1

BSA and 1.25 U Taq DNA polymerase (New England

Biolabs). The reaction conditions were (i) 2 min at

95 �C, (ii) 10 cycles of 30 s at 94 �C, 30 s of

touchdown primer annealing from 56 to 51 �C,

1 min at 72 �C, (iii) 25 cycles of 45 s at 94 �C, 30 s

at 51 �C, 1 min at 72 �C, and (iv) 5 min at 72 �C. PCR

products were loaded onto a 6 % (w/v) polyacryl-

amide gel with a 30–60 % denaturing gradient and

was run in 19 Tris–acetate-EDTA buffer in a Dcode

System apparatus (Bio-Rad, USA) for 12 h at 75 V

and 60 �C. The gels were stained in the presence of

1 mg l-1 of ethidium bromide and destained in Milli-

Q water for 15 min each. Major bands were excized,

eluted with 50 ll of sterile Milli-Q water and 2.5 ll

were used as DNA templates for reamplification with

the gc-338f and 907r primers as described above. The

PCR products were cloned into a pGEM-T vector

(Promega, Madson, WI, USA) and transformed into

Escherichia coli XL1-Blue by electroporation. Clones

were sequenced at the sequencing facilities of the

Instituto de Biotecnologıa-UNAM (Cuernavaca, Mex-

ico). Sequences were analyzed with Blastn at NCBI

and Blast at Greengenes. The CLUSTAL W software

(Thomson et al. 2004) was used to perform the

alignment of multiple sequences and a phylogenetic

tree was constructed using the neighbour-joining

method (Saitou and Nei 1987) and the Molecular

evolutionary Genetics Analysis (MEGA 3.1) software

program (Kumar et al. 2004). Tree topology was

evaluated by bootstrap analysis (Felsenstein 1985)

based on 1,000 replicates.

Results and discussion

DDT, DDE and DDD degradation in presence

of cosubstrates

After 7 weeks of treatment, complete inhibition of

DDT degradation was observed in the presence of

acetic acid. For all other compounds, DDT degrada-

tion was enhanced by the presence of cosubstrate as

compared with the control (without cosubstrate). As

seen in Table 1, DDT degradation with ethanol,

glucose, toluene, hexane and phenol was 49, 50, 56,

59 and 67 %, respectively, while in the control

degradation amounted to 23.5 % and the abiotic losses

were 1.87 %, this percentage is lower than the

standard deviations obtained indicating that abiotic

losses were negligible. On the other hand, the differ-

ences between experiments with addition of cosub-

strates and controls can be attributed to the utilization

of more easily available substrates for growth and the

consequent increase of microbial activity (Guerin and

Jones 1988; Castelo-Grande et al. 2010; Harder and

Dijkhuizen 1982). Fang et al. (2010) reported similar

results, where glucose enhanced DDT degradation in

liquid cultures using an isolated bacterial strain. Also,

some bacterial strains were found to degrade TCE

most effectively when glutamate, lactate, glucose, and

fructose were supplied as the carbon source (Yeager

et al. 2004).

In the present study, final DDE concentrations were

under the detection limit, which was 0.21 lg kg�1drysoil,

indicating more than 99.9 % elimination. DDD

removal was also inhibited by the presence of acetic

acid; but in contrast to DDT with glucose as a

substrate, degradation of DDD was indistinguishable

from the control. Other substrates tested increased the

degradation of DDD from 67 % in control to approx-

imately 86 % (Table 1).

The mineralization of pollutants was estimated as

described in methodology. In abiotic controls, neither

DDT, nor DDD nor DDE consumption was detected,

in concordance with undetectable CO2 production. In

all the experiments conducted with cosubstrates, the

CO2 produced was higher than that in the controls

without cosubstrates, in these controls the CO2

produced can be attributed to endogenous respiration

of microbial community or degradation of soil organic

matter. While, in the experiments with cosubstrates

CO2 measured also includes the production due to

degradation of both, cosubstrates and soil pollutants.

The results indicated highest theoretical minerali-

zation values with toluene and phenol (85–89 %),

which were also the cosubstrates eliciting the highest

DDT and DDD degradation. However, mineralization

of DDT, DDD and DDE should be confirmed by using

labeled molecules (14C). Hexane also enhanced DDT

and DDD degradation, but theoretical mineraliza-

tion was lower compared with that found with toluene

and phenol. Glucose and ethanol increased DDT

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degradation, and ethanol also had a positive effect on

DDD degradation. However, in both cases estimated

mineralization was significantly lower than that cal-

culated for toluene and phenol. The treatments with

toluene and phenol were the most efficient when the

degradation and theoretical mineralization results

were considered. Although a variety of carbon

substrates appear to enhance DDT degradation, an

energy source that most closely resembles its chemical

structure may be most successful additive (Foght et al.

2001).

Volatile cosubstrates degradation

Degradation of volatile cosubstrates was quantified in

the gas phase, and results are shown in Fig. 1. In the

case of toluene, complete degradation was achieved in

66, 16 and 8 h for the first, second and third addition,

respectively. While, complete degradation of hexane

was achieved in 200, 80 and 60 h for the first, second

and third addition, respectively. The increased micro-

bial activity due to the consumption of toluene and

hexane can be correlated with the higher consumption

of DDT compared with the control (Table 1). The

strategy of weekly addition of cosubstrates established

periods of high carbon loads (from the cosubstrate).

The microbial population consumed this carbon

source within the first few days, and once this source

was depleted, the active microbial population was

exposed to a low load of carbon for a few days, where

the pesticide consumption could be favored. In abiotic

controls, reductions of 3 and 7 % of initial concentra-

tion of hexane and toluene were observed. As CO2

production was not detectable this losses were attrib-

uted to adsorption into the slurry.

Effect of cosubstrate dose on DDT degradation

The results of the experiments with variable amounts

of phenol and toluene are shown in Fig. 2. The low

dose tested, 0.64 mgcarbon g�1drysoil (corresponding to

0.7 mg toluene or 0.84 mg of phenol per kg of dry

soil), resulted in approximately 25 % DDT degrada-

tion, which is comparable to that observed in the

controls. With the intermediate and high doses of

cosubstrate: 1.43 and 3.32 mgcarbon g�1drysoil (1.57 and

3.72 mgtoluene g�1drysoil and 1.87 and 4.33 mgphenol

g�1drysoil), an enhancement of DDT degradation was

observed in all cases. However, the strongest effects

on DDT degradation (25–69 %) were obtained with

the intermediate concentration of toluene and with the

higher concentration of phenol. The highest dose of

toluene resulted in lower DDT degradation, but was

still higher than that seen in the controls. For phenol,

the highest dose resulted in a marginal enhancement of

DDT degradation. Therefore, additions of cosubstrate

Fig. 1 Removal of volatile cosubstrates. (Black square)

Toluene; (white circle) hexane; (long downwards arrow)

toluene addition; (long downwards double arrow) hexane

addition. Error bars represent standard deviation, n = 6

Fig. 2 Degradation of DDT with different cosubstrate dose.

(White rectangle) Control; (white rectangle with upper right tolower left fill) phenol; (white rectangle with horizantal fill)toluene. Time of treatment: 1,200 h (7 weeks) with weekly

additions. Error bars represent standard deviation, n = 6. Initial

DDT concentration: 7.46 ± 0.66 mg kg�1drysoil

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should lie in the range between low and intermediate

doses.

The CO2 produced with low, intermediate and high

doses (Fig. 3) were calculated as mentioned above,

deducting the amount of CO2 produced in the control.

Final CO2 production (after 1,200 h) in the control

(endogenous respiration) without toluene addition was

6.4 ± 0.4 g kg�1drysoil. As expected, higher CO2 pro-

duction was observed in the treatments with higher

amounts of toluene added. However, this higher

microbial activity did not result in a higher DDT

consumption, as previously discussed. In abiotic

controls CO2 was not detected in the first 6 weeks.

At the final time point (1,200 h) 0.24 ± 0.03 g

kg�1drysoil was quantified, this amount was considerably

lower than the endogenous respiration.

DDT degradation rate

Based on the results with variable toluene concentra-

tions, DDT degradation was performed using toluene

as the cosubstrate at low and intermediate concentra-

tions, and the results are shown in Fig. 4. Each point

on the graph corresponds to the average DDT

concentration from three bottles. The abiotic losses

evaluated in the final samples (1,200 h) were

4.26 % ± 0.18 of the initial concentration of DDT.

The degradation started after 400 h, with the enhance-

ment of DDT degradation observed after 800 h of

treatment. This effect decreased, confirming that there

is an optimal concentration range for the added

cosubstrate. The cosubstrate addition affected the

Fig. 3 Carbon dioxide

production as a function of

different amounts of toluene

added. CO2 produced in

control without toluene was

subtracted. Toluene added

(mg C g�1drysoil): 0.79 (black

circle); 1.43 (whitetriangle); 3.32 (blacksquare). Error bars represent

standard deviation, n = 6

Fig. 4 DDT degradation in the presence of toluene. Toluene

addition: (long downwards arrow) only in the intermediate dose

experiments and (long downwards double arrow) in both, the

low and intermediate dose experiments. (White circle) Low

dose, 0.64 mg C g�1drysoil. (Black circle) Intermediate dose,

2.6 mg C g�1drysoil. Error bars represent standard deviation, n = 6

Biodegradation (2013) 24:215–225 221

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activity of the microbial community and no further

degradation after 800 h was observed, indicating that

after that period, mass transport may be the major

mechanism that limits the degradation process instead

of the microbial reaction.

Analysis of bacterial populations by DGGE

As toluene was one of the most effective compounds

in enhancing DDT degradation, final samples after

7 weeks of treatment with addition of toluene were

used to evaluate potential changes in the bacterial

populations in presence of this cosubstrate compared

with the control without toluene. DGGE analysis of

the bacterial communities was conducted in the native

soil (S), native soil in mineral medium as a control

(C) and native soil with toluene in mineral medium

(T). Figure 5 shows the obtained profiles. All samples

generated a simple banding pattern with 4–5 promi-

nent bands suggesting that bacterial diversity was low

in this soil, probably due to the toxicity of DDT and its

metabolites. These populations might be resistant to

DDT, it has been reported that this type of community

tends to develop in older contaminated soils although

it is not clear if these organisms have better DDT

degradation capabilities (Kantachote et al. 2001).

Bands 1, 2 and 3 were present in all samples. Band 1

was related to Steroidobacter, a rod-shaped bacterium

that degrades steroids. The strong intensity of bands 2

and 3 suggests that the corresponding microorganisms

might be dominant in the tested soil. They were related

to Burkholderiales and Sphingobacteriales, respec-

tively. These bacterial groups encompass degraders of

aliphatic and aromatic hydrocarbons including tolu-

ene. A faint band appeared in the presence of toluene

(Band 4). Band 5 was detected in the native soil in

mineral medium (C) and in the native soil with toluene

in mineral medium (T) but not in the native soil (S).

All closest relatives matched with uncultured bacteria

according to the Blastn analysis performed at NCBI,

most of them were related to soil habitats. Band 4

strongly matched with Azoarcus sp. str. PH002, a

phenol-degrading denitrifying bacterium (Van Schie

and Young 1998). The Azoarcus genus comprises

species able to degrade toluene and other aromatic

compounds (including halobenzoates like 3-chloro-

benzoate) in the presence of oxygen or nitrate (Song

et al. 2000; Van Schie and Young 1998; Zhou et al.

1995). Band 2 was distantly related to an unclassified

Comamonadaceae isolate from a TCE contaminated

groundwater aquifer while band 5 was closely related

to a Xanthomonadaceae (Panagrimonas). These

1

23

45

Fig. 5 DGGE analysis of bacterial communities. (M) 100 bp

molecular weight marker; (S) native soil; (C) native soil in

mineral medium as a control and (T) native soil in mineral

medium with toluene. Excised bands are labeled 1, 2, 3, 4 and 5

222 Biodegradation (2013) 24:215–225

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bands were present in the native soil as well as in the

native soil with mineral medium and native soil with

mineral medium and toluene. The detected microor-

ganisms were probably tolerant to DDT and its

congeners, however their exact participation in the

degradation process is not known. Figure 6 presents a

phylogenetic tree relating the obtained sequences. The

recently described Sphingobacterium (DDT-6), iso-

lated from an agricultural field and able to completely

mineralize DDT, through DDD and DDE, was

included in the tree (Fang et al. 2010).

Conclusions

There was a stimulatory effect of phenol, hexane and

toluene as cosubstrates on microbial populations that

were able to degrade DDT, DDE and DDD in soil.

Furthermore, inhibitory effects were observed with

acetic acid. DDE degradation was complete in all

cases, and DDD removal was also enhanced with the

addition of all substrates tested at the intermediate

dose, except for acetic acid and glucose (where DDD

removal was inhibited or equal to the control, respec-

tively). An optimal range for the amount of cosub-

strate added was determined. Above that range, the

effect in degradation was not sustained. The values in

the range lie between the low and medium concentra-

tions tested (0.64 and 2.6 mg C g�1drysoil). The process

had a lag phase of approximately 350 h; after that

period, degradation was observed and correlated with

CO2 evolution, indicating that a biological reaction

took place until 800 h of treatment. No further

degradation was obtained after 800 h, indicating that

the microbial reaction was not the responsible mech-

anism after that period and, possibly mass transport

limits the degradation process. The enhancement of

pollutant theoretical mineralization was related to the

higher CO2 production in the presence of cosubstrate,

as this value was higher than the theoretical amount for

complete cosubstrate mineralization. Bacterial diver-

sity was low in the soil suggesting the presence of

DDT-resistant microorganisms. Most microorganisms

were related to degradation processes. An Azoarcus-

like bacterium appeared in the presence of toluene;

this bacterium could provide broad-specificity oxy-

genases enhancing the degradation of DDT.

Acknowledgments The authors thank Dra. Leticia Yanez

from Universidad Autonoma de San Luis Potosı for the donation

of soil samples and Dra. Maribel Hernandez from Universidad

Autonoma Metropolitana for her suggestions to improve the

manuscript. This work was financed by Consejo Nacional de

Ciencia y Tecnologıa (Project CONACYT CB-61218) and by

Secretarıa de Educacion Publica (Project SEP-PROMEP UAM-

PTC-067).

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