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Self-bioremediation of cork-processing wastewaters by(chloro)phenol-degrading bacteria immobilised onto residualcork particles

I. del Castillo, P. Hernandez, A. Lafuente, I.D. Rodrıguez-Llorente, M.A. Caviedes,E. Pajuelo*

Departamento de Microbiologıa y Parasitologıa, Facultad de Farmacia, Universidad de Sevilla, c/Profesor Garcıa Gonzalez, 2,

41012 Sevilla, Spain

a r t i c l e i n f o

Article history:

Received 15 September 2011

Received in revised form

16 December 2011

Accepted 18 December 2011

Available online 30 December 2011

Keywords:

Phenol

Chlorophenols

Bacteria

Biofilms

Cork-processing wastewater

Bioremediation

a b s t r a c t

Cork manufacturing is a traditional industry in Southern Europe, being the main applica-

tion of this natural product in wine stoppers and insulation. Cork processing begins at

boiling the raw material. As a consequence, great volumes of dark wastewaters, with

elevated concentrations of chlorophenols, are generated, which must be depurated

through costly physicochemical procedures before discarding them into public water

courses. This work explores the potential of bacteria, isolated from cork-boiling waters

storage ponds, in bioremediation of the same effluent. The bacterial population present in

cork-processing wastewaters was analysed by DGGE; low bacterial biodiversity was found.

Aerobic bacteria were isolated and investigated for their tolerance against phenol and two

chlorophenols. The most tolerant strains were identified by sequencing 16S rDNA. The

phenol-degrading capacity was investigated by determining enzyme activities of the

phenol-degrading pathway. Moreover, the capacity to form biofilms was analysed in

a microtitre plate assay. Finally, the capacity to form biofilms onto the surface of residual

small cork particles was evaluated by acridine staining followed by epifluorescence

microscopy and by SEM. A low-cost bioremediation system, using phenol-degrading

bacteria immobilised onto residual cork particles (a by-product of the industry) is

proposed for the remediation of this industrial effluent (self-bioremediation).

ª 2011 Elsevier Ltd. All rights reserved.

Abbreviations: BLAST, Basic Local Alignment Search Tool; BOD, biological oxygen demand; BOX-PCR, PCR based on primers targetingthe highly conserved repetitive DNA sequences of BOX elements; CFU, colony forming unit; COD, chemical oxygen demand; DGGE,denaturing gradient gel electrophoresis; DTT, dithiothreitol; EPA, Environmental Protection Agency (US); MTC, maximum toleratedconcentration; NADH, nicotinamide-adenine-dinucletotide (reduced form); OD, optical density; PMSF, phenyl-methyl-sulfonyl fluoride;SEM, scanning electron microscopy; SP, storage pond (residual water of the storage pond); SPE, storage pond e “enriched” (residual waterof the storage pond after enrichment with 10% TSB); TAE, Tris-acetate-EDTA buffer for electrophoresis; TP, treatment pool (residualwater of the treatment pool); TPE, treatment pool e “enriched” (residual water of the treatment pool after enrichment with 10% TSB);TRITC, fluorescence filter for the detection of the fluorochrome tetramethylrhodamine isothiocyanate; TSA, trypticase-soya agar; TSB,trypticase-soya broth.* Corresponding author. Tel.: þ34 954556924; fax: þ34 954628162.E-mail address: [email protected] (E. Pajuelo).

Available online at www.sciencedirect.com

journal homepage: www.elsevier .com/locate/watres

wat e r r e s e a r c h 4 6 ( 2 0 1 2 ) 1 7 2 3e1 7 3 4

0043-1354/$ e see front matter ª 2011 Elsevier Ltd. All rights reserved.doi:10.1016/j.watres.2011.12.038


1. Introduction

Industrialisation is the main source of huge amounts of toxic

compounds being released to the biosphere, threatening

public health, wild life, and the environment. The list of toxic

compounds or elements published by US-EPA (Environmental

Protection Agency, www.epa.gov/oppt/p2home/) includes

over 6000 substances. Among them, phenolic compounds and

therein haloaromatic compounds (such as chlorophenols and

pentachlorophenol) are some of the most recalcitrant

organics, being degradation more difficult as the degree of

halogenation increases (Janssen et al., 2005; Solyanikova and

Golovleva, 2004).

Main uses of cork are the production of wine stoppers and

the utilisation as insulation material for thermal-, acoustic-

and electrical-insulation. The largest cork production and

manufacturing in the world is concentrated in the Mediter-

ranean basin. Spain is one of the most important cork

producers (22% of the total world production), together with

Portugal (the first one), Italy, Greece, Tunisia and Morocco. In

particular, in Southern Spain, around 40,000 tons cork per year

are produced. First step in cork manufacturing is focused on

cleaning and softening the raw material. For this purpose,

cork is boiled at 95�e100 �C for around 1 h in big treatment

pools. The same water can be used in 15e20 treatments, and

at the end, a dark wastewater resulting from this process is

collected in big storage ponds near the companies for later

depuration. Cork boiling water is characterised by high

chemical oxygen demand (COD), biological oxygen demand

(BOD) and phenol and polyphenols content, in the range of

4.5e5.5 g l�1, 1.1e1.8 g l�1 and 0.6e0.9 g l�1, respectively, and

by an acidic pH around 5 (Benitez et al., 2003; Domınguez et al.,

2007; Pintor et al., 2011). Phenolic fraction contains

compounds such as phenol, tannin fraction together with

gallic, protocatechuic, vanillic, syringic and ellagic acids, 2,4,6-

trichloroanisol and pentachlorophenol, (Benitez et al., 2003,

2006). Some of these compounds are among the most toxic

substances, as considered by EPA.

Chemical depuration treatments are expensive (Mendonca

et al., 2004), so little companies associate in order to share the

costs. Sometimes, wastewaters are stored in big ponds near

the manufacturing companies, and later on, transported to

a central treatment unit for reclamation using physicochem-

ical procedures. Physicochemical treatment of cork-boiling

water includes ozonation (Benitez et al., 2003; Lan et al.,

2008), Fenton oxidation, (Beltran de Heredia et al., 2004;

Guedes et al., 2003), flocculation (Domınguez et al., 2007) and

filtration techniques (Benitez et al., 2006, 2008; Bernardo et al.,

2011) or combinations of these methods, in order to decrease

the chemical oxygen demand (COD) in 75e85%. Any alterna-

tive treatment that can substitute all or part of these steps is of

great interest to the cork industry, since it can significantly

decrease the cost of wastewater treatment, especially if

alternative techniques can be applied in situ.

Bioremediation, i.e., the use of living organisms for envi-

ronmental cleaning, is an ecological and low-cost alternative

to the most traditional physicochemical remediation tech-

niques (Dıaz, 2004; Galvao et al., 2005; Stapleton and Singh,

2002). Major limitations are the bioavailability of organic

matter and finding efficient biodegraders. Many microorgan-

isms being able to degrade phenolic compounds have been

described, both in aerobic (Pseudomonas, Burkholderia, Sphin-

gomonas, Ralstonia, Arthrobacter, Acinetobacter, Alcaligenes) or

anaerobic conditions, (Desulfitebacterium, Dehalomonas, Deha-

lococcoides, Dehalobacter) (Adrian et al., 2000; Bunge et al., 2003;

Chang et al., 2003; Dıaz, 2004; Furukawa, 2003; Parvanov and

Topalova, 2008; Smidt and De Vos, 2004; Van der Meer, 2008).

Haloaromatic compounds are more difficult to degrade

because the high stability of the carbon-halogen bond.

However, some genera of bacteria have been also reported

being able to degrade (totally or partially) this kind or organics,

both in aerobic or anaerobic conditions, such as Sphingobium,

Sphingomonas, Dehalobacter, Dechloromonas, Dehalococcoides,

Rhodococcus (Furukawa, 2006; Solyanikova and Golovleva,

2004; Van der Meer, 2008; Cai and Xun, 2005).

To our knowledge, little information has been produced on

the biotechnological use of microorganisms in the treatment

of this effluent (Dias-Machado et al., 2006; Mendonca et al.,

2004). In this work, the complete bacterial population of

cork-processing wastewaters has been analysed. Also, culti-

vable bacteria have been tested for their ability to degrade

(chloro)phenols. A system is proposed by which phenol-

degrading bacteria from cork wastewaters storage ponds are

immobilised onto residual cork particles for wastewater

remediation.

2. Materials and methods

2.1. Total DNA isolation from cork-processingwastewaters

In this work, two kinds of water samples were utilised: the

water of the treatment pool (TP) had been used between 10

and 15 times, (the exact number depends on every particular

treatment), and it was close to exhaustion (just to be dis-

charged into the storage pond). The storage pond (SP) is a very

big pool containing the residual waters for long time. The

main difference between the two samples is the time that the

water remains in the pool. In the treatment pool the water

remains only the time needed for 10e15 treatments. However,

in the storage pond the water can remain for months, until it

is brought to the depuration central. The bacterial population

present in both wastewaters has been analysed using DGGE.

Samples of 2 L of the wastewaters of a local industry were

collected from the treatment pool (TP) and the storage pond

(SP) and divided into two flasks of 1 L each one. One of each

was supplementedwith 100mL (10%v/v) of TSB (trypticase soy

broth) medium (Vincent, 1970). These enriched samples,

designed TP-E and SP-E, were incubated at 28 �C for 48 h. Total

DNA was isolated from all the four samples using the Pow-

erSoil� DNA isolation kit (MoBio Laboratories, Inc. California)

following the instructions of the manufacturer with several

modifications, in order to adapt the protocol to a liquid

sample. 100 mL of samples TP-E and SP-E (containing higher

bacterial density after enrichment) were centrifuged at 1100 g

for 10 min at 4 �C and bacterial pellets were resuspended in

1 mL in sterile 0.9% NaCl. 1 L of samples TP and SP, with

wat e r r e s e a r c h 4 6 ( 2 0 1 2 ) 1 7 2 3e1 7 3 41724


a lower concentration of bacteria, were concentrated by

filtering through a 0.45 mm Millipore filter. The filter was

washed with 1 mL of sterile 0.9% NaCl. From this step, DNA

was extracted following the instructions of the manufacturer.

2.2. PCR-DGGE analysis

Amplification of partial 16S rDNAwas performed according to

Weisburg et al. (1991) using the modified primers fD1

(50CCGAATTCGTCGACAACAGACTTTGA TCCTGGCTCAG30)and rD1 (50CCCGGGATCCAAGCTTAAGGAGGTGATCCAG CC30)and Taq polymerase ExpandHigh Fidelity PCR System (Roche).

PCR reaction consisted in an initial denaturation step at 94 �Cfor 2 min, followed by 15 cycles (94 �C for 60 s, 55 �C for 45 s

and 72 �C for 2 min). The second PCR reaction was

performed according to Heuer et al. (1997) on 1e3 mL of the

previous PCR product using primers F984GC: (50

CGCCCGGGGCGCGCCCCGGGCGGGGCGGGGGCACGGGGGGAA

CGCGAAGAACCTTAC30) and R1378 (50GCGTGTGTACAAGACCC30). PCR conditions were as follows: an initial denatur-

ation step at 94 �C for 2min, followed by 10 cycles inwhich the

annealing temperature decreased from 65 to 55 �C (30 s at

94 �C, 1 min at [65e55 �C] and 2 min at 68 �C), followed by 25

cycles at low constant annealing temperature (30 s at 94 �C,1 min at 55 �C and 2min at 68 �C), and a final extension step at

68 �C for 10min. DGGE conditionswere as previously described

Heuer et al. (1997), using polyacrilamide gels with a gradient of

two denaturing agents: urea (from 3.0 to 4.5M) and formamide

(from 25 to 35%). Denaturing gradient gel electrophoresis of

PCR products was performed in a DCODE Universal Mutation

Detection System (Bio-Rad). at constant temperature (60 �C) at200 V for 15 min to compact the samples, followed by 16 h at

70 V. Gels were stained for 15 min with ethidium bromide and

washed with distilled water until obtain a clear image. Gels

banding patterns were derived from captured gel images and

analysed using Quantity One 1-D analysis software.

2.3. Comparison of bacterial communities based onDGGE pattern analysis

The richness, diversity, evenness, and stability of bacterial

communities at the respective habitats were estimated based

on the number and the relative intensity of the DGGE bands.

Phylotype (band) richness (R) was evaluated from total

different DGGE bands (Fromin et al., 2002). Diversity and

evenness of the communities were estimated by using Shan-

non’s diversity index with the formula, H0 ¼ �PPi ln Pi, for the

calculation of the index, where Pi is the relative intensity of

band i (Pi ¼ Ni/N; Ni, intensity of band i; N, sum of the inten-

sities of all bands) (Maggurran, 1988). Evenness was calculated

by using the formula, E ¼ H0/lnR, where R is the total different

number of DGGE bands (richness).

2.4. Extraction of the DGGE bands

The predominant bands in DGGE gel were excised and purified

frompolyacrilamide gel as described byWatanabe et al. (1998).

DNA was re-amplified with the second pairs of primers (F984

and R1378) without the GC-clamp (underlined in section 2.2)

and in a reaction with 35 cycles at 55 �C annealing

temperature. The products of the second PCR were electro-

phoresed through a 1.5% agarose gel in TAE buffer and purified

using the Ilustra GFX�PCR DNA and Gel Band Purification Kit

(GE Healthcare). The extracted DNA was quantified by

measuring the absorbance at 260 nm and sequenced.

2.5. Isolation of cultivable bacteria from wastewatersponds

Cultivable bacteria were isolated from all the four samples.

We have focused on the isolation of aerobic and facultative

anaerobic bacteria. Samples of wastewater were plated on

TSA plates and incubated at 28 �C for 4 days for bacteria

isolation. Colonies with different morphology and pigmenta-

tion were selected, purified in TSA plates, and stored at

�S80 �C.

2.6. Genetic diversity by BOX-PCR

BOX-PCR was performed according to the method of Johnson

et al. (2003) using the BOXA1R primer and 1 ml of DNA at 40 ng/

ml. The PCR products were electrophoresed through a 1.5%

agarose gel in TAE buffer, at constant voltage 70 V for 2 h and

30 min. BOX-PCR fingerprint similarities were determined by

calculating the Pearson’s product moment correlation coeffi-

cient (Jobson, 1991). In the current study, isolates were

considered to be identical if they exhibited BOX-PCR finger-

prints that were at least 90% similar, in order to discriminate

all repetitive strains.

2.7. Identification of cultivable bacteria

Bacteria with different BOX fingerprints were identified by

PCR-amplification and sequencing of the 16S rDNA. Primers

and PCR conditions used for 16S rDNA amplification are

described by Rivas et al. (2002). Biochemical characterisation

of some strains was performed using commercial systems

(API20E, API 20 NE from Biomerieux, RapID from Oxoid) [see

supplementary information Tables S1 and S2].

2.8. Resistance against phenolic compounds

The resistance of the isolates against phenol and chlor-

ophenols was determined on plates containing TSA medium

supplemented with increasing concentrations of these

compounds, ranging from 0 to 10mM (phenol), from 0 to 4mM

4-chlorophenol and 0e3 mM 2,4-dichlorophenol. Plates were

incubated at 28 �C for 3e4 days. The tolerance is expressed as

the MTC, maximum tolerated concentration, which is the

maximal concentration of the toxic allowing bacterial growth.

2.9. Phenol consumption

Phenol degrading capacitywas tested inmedia containing this

compound as the sole carbon source. Bacteria were grown in

minimalmedium (Foght et al., 1989) for 72 h at 28 �C at 200 rpm

constant shaking. 25 mL of liquid minimal medium supple-

mented with 2.5 mM phenol were inoculated with 100 mL of

the abovementioned cultures (approx. 108 C.F.U./mL). Aliquots

of 2mLwere taken at 1 h intervals during the first 6 h and up to

wat e r r e s e a r c h 4 6 ( 2 0 1 2 ) 1 7 2 3e1 7 3 4 1725


26 h. OD at 600 nm was measured. Aliquots were centrifuged

at 8000 g for 10min and phenol concentrationwas determined

in the supernatants using 4-amino-antipirine according to the

method previously described by Lacoste et al. (1959) using

minimal medium for establishing the background of the

colorimetric determination.

2.10. Enzymatic activities of phenol degradation pathway

Phenol hydroxylase activity (EC1.14.13.7) and catechol dioxy-

genases (catechol 1,2-dioxygenase (EC1.13.11.1) and catechol

2,3-dioxygenase (EC 1.13.11.2)) were determined in crude

extracts prepared from bacteria grown for 10 days in minimal

medium supplemented with 2.5mMphenol. 250mL of culture

were centrifuged and resuspended in 5 mL of 50 mM potas-

sium phosphate buffer, pH 7.5, containing 1 mM PMSF and

0.1 mM DTT. Bacterial cells were disrupted by sonication and

the lisate was centrifuged at 12,500 g for 10 min at 4 �C.Enzymatic activities were determined in the crude extract

according to Zaki (2006).

2.11. Adhesion assay on microtitre plates

Adhesion capacity in the presence of phenol wasmeasured as

theabilityof cultures toadhere to thesurfaceofwellsof 96-well

microtitre plates, as described by Favre-Bonte et al. (2007).

Cultures were grown in minimal medium supplemented with

2.5mMphenol at 28 �C for 48 h. All cultureswerenormalised at

OD600nm 1.0. 20 mL of the corresponding cultures were added to

12 wells of microtitre plates (a row for each microorganism).

For the control, 20 mL of minimal medium was added. The

plateswere sealedwith parafilm and incubated at 28 �C during

4 days. After this time, the medium was removed, the wells

were washed 5 times with 200 mL distilled water, and the bio-

films were stained with 200 mL 0.01% crystal violet for 20 min.

Plates were rinsed three times with 200 mL sterile water. The

dyewassolubilisedwith100mL95%ethanol 33%acetic acidand

quantified by measuring the absorbance at 570 nm/595 nm.

2.12. Biofilms formation onto cork particles

The formation of biofilms on the surface of cork particles was

examined. Cork particles were collected on the ground of the

cork industry (in fact, it is a residue of the industry). Cork

particles were homogenised in a coffee grinder and sieved

through a 0.5 mm sieve. They were then washed three times

with 25 mL 10 mM HCl for 10 min, and neutralised with the

appropriate volume of 1 N NaOH, rinsed five times in sterile

distilled water and air-dried. Flasks with 15 mL of minimal

medium supplemented with 2.5 mM phenol and 0.15 g of

treated cork particles were first autoclaved and then inocu-

lated with the selected microorganisms, previously grown on

2.5 mM phenol and normalised at OD600nm ¼ 1.0, and incu-

bated at 28 �C for 10 days. After that, particleswere transferred

to petri dishes, washed five times with distilled water, air-

dried and stained with Acridine Orange (Johansson et al.,

2008). Bacterial attachment to cork surface was examined

with an epifluorescence microscope (Olympus BX61) using

�100 objective lens and a TRITC green filter. Bacterial

attachment was also analysed by SEM (scanning electron

microscopy). Cork particles incubated in the presence of

bacteria as described above were fixed overnight in the pres-

ence of OsO4, followed by Au treatment, and observed under

a Jeol 6460LV microscope.

3. Results

3.1. Analysis of bacterial communities in cork-processing wastewaters

The biodiversity of bacteria present in cork-processing

wastewaters was analysed by DGGE in crude samples (TP,

TPE, SP and SPE) (Fig. 1A). Indices for diversity, richness and

evenness of bacterial communities are shown in Table 1. The

overall community diversity as a function of both the total

number of species (richness) present in the samples and their

relative distribution (evenness) was calculated from the data

obtained from the bacterial 16S rDNA DGGE profile. Diversity

(H0) in all samples was medium, between 2 and 2.6, and the

number of total different DGGE bands (ranging from 11 to 14

bands)was lower than that of other residual waters. Phylotype

(band) richness of the bacterial communities in the enriched

samples (TPE and SPE) was not significantly greater than that

of the initial samples (TP and SP). The algorithm for

computing similarity through this program was the Dice

coefficient. This coefficient has been used by other

researchers as the quantitative measurement of DGGE band-

ing patterns (Trevors et al., 2010). Fig. 1B shows the compar-

ison of the four profiles. TP and TPE were the samples whose

profiles showed the highest similarity, being SPE and TPE the

most different. Therewere common bands in the four profiles,

showing the presence of common microorganisms in all

pools, probably corresponding to cultivable (since they also

appeared in the enriched samples) and very competitive

bacteria, whereas some other bands disappeared in the

enriched samples or showed a lower intensity when

compared to the initial samples. The most intense bands in

DGGE profiles were excised, amplified and sequenced. These

organisms were coincident with bacteria described as phenol

degraders in previous works, such as Ralstonia, Steno-

trophomonas, Cupriavidus and Lysobacter (Zaki, 2006; Dong et al.,

2008; Basile and Erijman, 2010).

3.2. Isolation and identification of bacteria from cork-processing wastewaters

Aliquots of the four samples indicated above were stroked out

on TSA plates for colony isolation. Bacteria with different

colony morphology were selected. Initially, 35 isolates were

obtained in total; 4 apparently different colonies were

observed in SP and 6 in TP. Enriched samples showed more

diversity of cultivable microorganisms; 14 different colonies

were seen in SPE sample and 11 in TP-E. In the original

samples, both Gram negative and Gram positive bacteria were

isolated, while in the enriched samples, mainly Gramnegative

bacteria were identified.

In order to discriminate redundancy, PCR-BOX was per-

formed and bacteria with the same band profile were

considered as the same strain (Johnson et al., 2003). Some of

wat e r r e s e a r c h 4 6 ( 2 0 1 2 ) 1 7 2 3e1 7 3 41726


the isolates showed similar BOX-profiles (Fig. 2); for instance,

strains SPE-5A and SPE-8B showed the same PCR-BOX profile,

indicating that they probably were the same strain. Analogous

results were found for strains SPE-2A and SPE-5B or strains

TPE-3 and TPE-4. These results showed that the 35 isolates

could be grouped into 16 different clusters, according to the

BOX-PCR profiles. Nearly complete 16S gene sequences

(approx. 1500bp) from bacteria with different BOX fingerprint

were obtained (Table 2). Among the cultivable microorgan-

isms present in cork-processing wastewaters, both gram

positive bacilli (Arthrobacter, Bacillus) and gram negative bacilli

(some Enterobacteriaceae like Serratia and Enterobacter and

other gram negative fermentative bacilli like Aeromonas,), as

well as non fermentative gram negative bacilli (Comamonas,

Stenotrophomonas and Acinetobacter) were found.

3.3. Tolerance of bacteria against phenol andchlorophenols

All the isolated bacteria were tested for their tolerance against

phenolic compounds. Results are shown in Table 2. The

resistance towards phenol was variable, ranging from 2.5 to

10 mM. Four strains were able to grow at the maximum

concentration tested (SPE-5A, SPE-9A, TPE-5A and TP6) and

were selected for further analysis. Two other strains (TP3 and

TP4) were also selected, since they were able to grow better in

TSAmedium supplemented with 2.5mM phenol thanwithout

this compound. An additional strain (SP3) was selected as

a control with intermediate phenol tolerance. The resistance

against chlorophenols was also determined in these 7 strains.

Themost resistant strains concerning 4-chlorophenol and 2,4-

chlorophenol were TP4, TP6 and TPE-5A. These seven strains

were selected to carry out phenol degradation assays and

enzymatic determinations.

3.4. Phenol-degrading ability

The ability to degrade phenol was investigated at two levels:

first, the capacity of the 7 selected strains to use phenol as the

sole carbon and energy source was assayed, and secondly, the

activity of the first enzymes involved in phenol degradation

was determined. NADH-phenol hydroxylase, which is the

Fig. 1 e A. Analysis of bacterial populations present in cork-processing wastewaters. Samples corresponded to the

treatment pool (TP) and the storage pond (SP), as well as the corresponding samples after enrichment with TSB medium

(TPE and SPE). The numbered bands were extracted from the gel and sequenced. These bands showed the maximum

identity (BLAST) with the following organisms: (1, 3) Uncultured bacterium, (2, 12, 15) Ralstonia sp., (4, 7, 11)

Stenotrophomonas maltophilia, (5, 8, 9, 13, 14) Stenotrophomonas sp., (6) Cupriavidus tawanensis, (10) Lysobacter capcisy. B.

Dendrogram showing the similarity between the DGGE profiles corresponding to the four samples based on the Dice

coefficient.

wat e r r e s e a r c h 4 6 ( 2 0 1 2 ) 1 7 2 3e1 7 3 4 1727


enzyme involved in the first step of the phenol degradation

pathway, was determined. In addition, catechol 1,2-

dioxygenase and catechol 2,3-dioxygenase activities were

also measured, in order to clarify if the breakdown of the

aromatic ring occurred in the ortho or the meta position.

3.4.1. Phenol consumptionThe strains were cultivated in minimal medium supple-

mented with 2.5 mM phenol as the sole carbon and energy

source. The optical density at 600 nm was measured as

a parameter of bacterial growth. In the other hand, the

disappearance of phenol in the culture medium was evalu-

ated. Results are shown in Fig. 3. The ability to grow on phenol

as the sole carbon source varied among strains. Lag phases of

1 he2 h approx. were observed for some strains, such as TP3,

TP6, SPE-5A and SPE-9A. After this period, SPE-9A showed

a fast growth rate in the presence of phenol. Other strains like

TP3, TP4 and TP6 showed slower growth rates. Phenol was

determined every hour in the supernatant of these cultures

for 26 h (Fig. 3), and the rate of phenol degradation during the

first 6 h was calculated (Fig. 3). Final phenol concentration in

the supernatant of the cultures after 26 h incubation was also

measured (Table 3). SPE-9A was the most efficient phenol

degrader bacteria, with the highest phenol degradation rate

(216mg L�1 h�1), followed by TP3, TP4 and TP6 (around 100mg

phenol L�1 h�1). The other strains, SP3, SPE-5A and TPE-5A

degraded phenol less efficiently.

3.4.2. Determination of phenol hydroxylase activityThe metabolic pathway of phenol degradation is well known.

First step is the oxidation of the aromatic ring by phenol

hydroxylase to form the intermediate catechol (1,2-di-

hidroxibenzene). Then, the aromatic ring is opened by diox-

ygenases, either catechol-1,2-dioxygenase (ortho-cleavage) or

catechol-2,3-dioxygenase (meta-cleavage), followed by reduc-

tions finally leading to central metabolism intermediates

(Solyanikova and Golovleva, 2004).

The activity of the NADH-phenol hydroxylase was deter-

mined in all the resistant strains (Table 3). All the strains able

to grow on phenols showed phenol hydroxilase activity. The

strain showing the highest level was SPE-9A, coinciding with

the highest phenol consumption and tolerance, followed by

strains TP3 and TP4, which also were able to grow on phenol

and use it as the sole carbon and energy source (Fig. 3). Lower

phenol hydroxylase activity was found in SPE-5A and TPE-5A,

whereas TP6 and SP3 displayed almost undetectable levels. No

phenol hydroxilase activity was observed using NADPH as the

electron donor in any of the strains (not shown).

The activities of catechol-1,2- and 2,3-dioxygenases were

also measured. In all the strains, the activity of 2,3-

dioxygenase showed a higher value than that of catechol-

1,2-dioxygenase. However, in some strains (TP6, SP3, SPE-5A

and SPE-9A) the ratio between meta and ortho fission path-

ways was very high, indicating that they degraded phenol

mainly through the catechol meta fission pathway. On the

contrary, the other three strains (TP3, TP4 and TPE-5A) seemed

to perform both the ortho and meta fission pathways simulta-

neously, though at different rates. Furthermore, the absolute

values of the activities varied considerably. The strain SPE-9A

showed the highest levels of all the phenol degrading activi-

ties per mL culture, suggesting that this bacterium is an effi-

cient phenol degrader. These data were further confirmed by

determination of remaining phenol in the supernatant of the

cultures (Table 3), which showed that in fact SP-E9A degraded

Table 1 e Richness (R), diversity (H0) and evenness (E) ofbacterial communities at the respective samples.

Sample Richness (R) Diversity index (H0) Evenness (E)

TP 13 2.386 0.930

TPE 12 2.094 0.843

SP 11 2.309 0.963

SPE 14 2.606 0.987

Fig. 2 e BOX-PCR profiles of bacterial strains isolated from cork-processing wastewaters. MM, molecular markers.

wat e r r e s e a r c h 4 6 ( 2 0 1 2 ) 1 7 2 3e1 7 3 41728


phenol more efficiently, since only 22% phenol remained in

supernatant after 26 h.

3.5. Determination of biofilm forming capacity

3.5.1. Microtitre plates assayThe ability to form biofilms is a useful characteristic for

a bacterial strain to be used in bioremediation. Immobilisation

onto carriers increases the stability and viability of cultures

(El Fantroussi and Agathos, 2005; Tavares et al., 2010). Hence,

the capacity of selected strains for surface cover was evaluated

in a 96-well microtitre plate assay. Furthermore, cultures were

grown on 2.5mMphenol as the sole carbon and energy source,

in order to include the influence of this compound on the

formation of biofilms. The results of surface cover after 48 h are

shown in Fig. 4. The strains SP3, SPE-5A, TPE-5A and SPE-9A

showed the highest capacity to cover the surface of micro-

trite plates. However, when examined the tolerance against

phenolic compounds (Table 2) and the phenol degradation

capacity of these strains (Fig. 3 and Table 3), the best candidate

for wastewater bioremediation seemed to be SP-E9A, since it

couples high phenol tolerance and degradation ability and

a good surface cover. TP3 and TP4 showed an intermediate

surface cover capacity but they were also tolerant to phenolics

and moreover, some of the best phenol degraders, so they

could be also good candidates for wastewaters bioremediation.

By contrast, SPE-5A, TPE-5A and SP3, in spite of having good

cover surface, were not able to degrade phenol so efficiently.

3.5.2. Formation of biofilms onto cork particlesA residual layer of small cork particles and cork powder is

present on the floor of cork industries, as a result of the

manufacturing procedure. There are also cork pieces inade-

quated for industrial use (due to low quality, inadequate thick-

ness or other defects) thatmight be used as a carrier for bacterial

immobilisation. The ability to form a biofilm onto the surface of

cork particles was evaluated. As in the previous case, bacterial

strains were incubated in the presence of tiny cork particles and

grown in the presence of 2.5 mM phenol as the sole carbon

source, in order to test the effect of this compound on the

attachment of bacteria to cork particles. When observed under

the fluorescence microscope, the control particles incubated

without bacteria showed a background level of autofluorescence

(Fig. 5A), with green and dark zones. However, when cork

particles were incubated in the presence of bacteria, an intense

green fluorescence was observed (Fig. 5B). Close-up view of the

particles showed the attachment of a great number of bacteria

to the surface of a cork particle (Fig. 5C, D).

We have also observed the bacteria immobilised onto

residual cork particles under a scanning electron microscope

(Fig. 6). The structure of the cork, with a large surface of the

vegetal cells walls, seems to be a good material for bacterial

immobilisation (Fig. 6A, B). In fact, we have observed that the

strain SPE-9A, which is one of the best phenol degraders, is

able to profusely colonise and grow onto the surface of

residual cork particles, with a very high surface cover (Fig. 6C).

Other strains with intermediate degradation capacity, such as

TPE-5A showed lower density of bacteria on the surface of

cork (Fig. 6D). These results indicate that cork particles are

a suitable carrier for immobilise bacteria isolated from cork-

processing wastewaters, for the purpose of bioremediation.

4. Discussion

Cork manufacturing is an important industry in Extremadura

and western Andalucia, especially constituted by small

companies (SME’s). Cork boiling generates a toxic residuewith

an elevated content of phenol and chlorophenols (Benitez

et al., 2006; Domınguez et al., 2007; Pintor et al., 2011).

Despite of many microorganisms are to degrade phenolic

compounds, little information is available on the use of

microorganisms for the remediation of cork-processing

wastewaters, a technology that could be an alternative to

the expensive physico-chemical treatments used nowadays.

Some preliminary studies have been done on the isolation of

gram negative bacteria like Pseudomonas, Klebsiella,

Table 2 e Identification and tolerance against phenolic compounds of the 16 strains with different PCR-BOX fingerprint.

Strain Maximum identity (16S rDNA) MTC (mM)

Phenol 4-Chlorophenol 2,4-dichlorophenol

TP1 Bacillus cereus (99%) 2.5 n.d. n.d.

TP3 Serratia sp. (98%) 5.0 0.75 0.6

TP4 Enterobacter sp. (98%) 7.5 0.75 1.5

TP6 Bacillus cereus/thuriengiensis (98%) 10.0 2 1.5

SP1 Bacillus cereus (99%) 2.5 n.d. n.d.

SP2 Comamonas testosteroni (98%) 5.0 n.d. n.d.

SP3 Stenotrophomonas maltophilia (100%) 5.0 0.75 0.6

SP4 Comamonas testosteroni (99%) 5.0 n.d. n.d.

TPE-1A Acinetobacter sp. (98%) 5.0 n.d. n.d.

TPE-5A Arthrobacter arilaitensis (98%) 10.0 2 0.9

TPE-6 Aeromonas hydrophila/veronii (99%) 2.5 n.d. n.d.

SPE-1A Stenotrophomonas sp. (99%) 2.5 n.d. n.d.

SPE-4 Aeromonas sp. (97%) 5.0 n.d. n.d.

SPE-5A Acinetobacter sp. (98%) 10.0 0.75 0.6

SPE-8A Stenotrophomonas sp. (97%) 7.5 n.d. n.d.

SPE-9A Acinetobacter soli (99%) 10.0 0.75 0.6

Bold in Table 2 correspond to the strains that are selected for further analysis.

wat e r r e s e a r c h 4 6 ( 2 0 1 2 ) 1 7 2 3e1 7 3 4 1729


Stenotrophomonas and Burkholderia from this residue (Dias-

Machado et al., 2006). These bacteria have been used, in

combination with an ozonation treatment, for eliminating up

to 90% of total organic carbon from cork wastewaters.

In this work, the structure of the bacterial community of

cork-processing wastewaters was analysed by DGGE (Muyzer

et al., 1983). These samples contained a low richness and

biodiversity, based on the low number of different DGGE

Fig. 3 e Consumption of phenol by bacterial strains isolated from cork-processing wastewaters. Bacteria were grown in

minimal medium in the presence of 2.5 mM phenol. At 2 h intervals, bacterial growth was determined as OD at 600 nm and

phenol was determined in the supernatant.

wat e r r e s e a r c h 4 6 ( 2 0 1 2 ) 1 7 2 3e1 7 3 41730


bands. Low biodiversity could be due to two factors; the high

concentration of toxic compounds inwastewaters (and hence,

the high toxicity of this residue), and the relatively low total

carbon content in these samples, which are in fact a liquor

(Dias-Machado et al., 2006). Some of the most prominent

bands were sequenced, corresponding to well-known phenol-

degrading bacteria, such as Ralstonia, Stenotrophomonas,

Cupriavidus and Lysobacter (Zaki, 2006; Dong et al., 2008; Basile

and Erijman, 2010).

Aerobic cultivable bacteria were isolated from wastewa-

ters. A total of 35 strains were isolated, which were grouped in

16 different clusters according to PCR-BOX profiles. All the

strains were investigated for their ability to resist and degrade

phenol. The most resistant strains were identified, corre-

sponding to environmentally robust bacteria, including some

gram negative (Acinetobacter, Enterobacter, Serratia and Steno-

trophomonas) and some gram positive bacteria (Arthrobacter

and Bacillus). These genera of bacteria belong to classical

(chloro)phenol degraders (Sandhu et al., 2009; Solyanikova

and Golovleva, 2004). The resistance towards phenol was

variable, between 2.5 and 10 mM phenol; three strains were

able to tolerate 10 mM phenol and 1.5e2 mM chlorophenols.

Several strains were able to grow in the presence of phenol as

the sole carbon and energy source. All of these strains pre-

sented phenol hydroxylase activity. A preliminary analysis

was also performed in order to know if the degradation

pathway followed the ortho- or the meta- ring cleavage. Some

of the strains seemed to degrade phenol mainly by the meta

fission pathway (like TP6, SPE-5A or SPE-9A), whereas some

others (such as TP3 and TP4) seemed to use both pathways.

The capacity to form biofilms is an important character-

istic for a bacterial strain to be proposed for wastewaters

bioremediation purposes. Immobilised bacteria for bioreme-

diation are more stable than free living bacteria, and are able

to perform the pollutants degradation for longer periods and

in a more efficient manner. Immobilisation improves the

viability and stability of microbial cultures for bioremediation

(El Fantroussi and Agathos, 2005; Tavares et al., 2010).

Different residual products have been used as carriers for

immobilised cultures, such as tubings, packed columns,

granulated activated carbon, coconut shells, etc. (Amuda and

Ibrahim, 2006; Carvalho et al., 2001; Chang et al., 2003; Eker

and Kargi, 2008; Shieh et al., 1990; Wobus et al., 1995). Thus,

the ability to form biofilms was tested, both in a preliminary

test on a microtitre plate assay and also onto the surface of

cork particles. The cork powder particles have an inadequate

size distribution to be used in the manufacture of agglomer-

ates and they are mainly burned to produce energy (Gil, 1996).

Several strains were able to multiply and colonise the surface

of cork particles in the presence of phenol as the sole carbon

and energy source.

Considering all the previous results (resistance towards

phenol and chlorophenols, phenol degradation capacity and

cork surface cover), finally three bacterial strains (TP3, TP4

and SPE-9A) were considered as the best candidates to design

a self-bioremediation system, based on chloro(phenol)-

degrading bacteria immobilised onto cork particles.

When a bioremediation procedure is to be designed, it

would bemore attractive if a supplementary economical value

could be added. This is an important task, since a great

amount of small cork particles are usually on the floor of the

cork industries as a result of the different cork treatments and

trucks transit, which are not used, being in fact a residual

product of the industry. In our case, the self-bioremediation

system proposed has several advantages; the first one is

using native bacteria for wastewater treatment, and the

second one is using cork particles, a residue of the industry, as

the carrier for bacteria immobilisation. Additional studies are

being carried out in order to know the phenolics-degrading

ability of some of these strains in “real” cork-processing

wastewaters. Preliminary results show that degradation of

total polyphenols (expressed as tannic acid or caffeic acid)

reached 60e80%, depending on the strain and the

Table 3 e Enzyme activities of the phenol-degrading pathway in crude extract obtained from bacteria isolated from cork-processing wastewaters. Bacteria were cultivated for 10 days in the presence of 2.5 mM phenol as the sole carbon source.The last column represents the percentage of phenol remaining in the supernatant of the cultures after 26 h incubation inthe presence of phenol (see Fig. 3).

Bacteria Phenol hydroxylase(mU mL�1)

Catechol 1,2-dioxygenase(mU mL�1)

Catechol 2,3-dioxygenase(mU mL�1)

Ratio meta/orthofission

Phenol insupernatant (%)

TP3 2.20 0.40 2.10 5 63

TP4 3.50 0.30 2.00 7 61

TP6 0.07 0.10 5.70 57 53

SP3 0.20 0.05 1.90 38 60

TPE-5A 1.50 0.40 2.50 6 72

SPE-5A 1.80 0.11 4.60 42 49

SPE-9A 4.75 1.43 37.50 26 22

OD

a

t 5

75

nm

Fig. 4 e Evaluation of biofilms formation on a microtitre

plate assay. Bacterial strains were grown in the presence of

minimal medium supplemented with 2.5 mM phenol in

the wells of a 96-well microtitre plate. After 4 days, the

formation of biofilms on the surface of the plate was

evaluated. C: Control without bacteria. Bars represent

standard deviations (n [ 12).

wat e r r e s e a r c h 4 6 ( 2 0 1 2 ) 1 7 2 3e1 7 3 4 1731


Fig. 6 e Evaluation of biofilms formation on the surface of cork particles by SEM. A. Structure of a cork particle. B. Control

without bacteria. C. Colonisation of cork surface by strain SPE-9A. D. Colonisation of cork surface by strain TPE-5A.

Fig. 5 e Evaluation of biofilms formation on the surface of cork particles. Cork particles were incubated in the presence of

bacteria in minimal medium supplemented with 2.5 mM phenol. After washing, bacterial attachment was observed by

staining with acridine orange A. Control; cork particle incubated inmediumwithout bacteria, showing the background level

of autofluorescence, B. Cork particle incubated in the presence of bacterial strain SPE-5A, C, D. Close-up of bacterial

attachment to the surface of cork particles; C (TP6) and D (TP4).

wat e r r e s e a r c h 4 6 ( 2 0 1 2 ) 1 7 2 3e1 7 3 41732


experimental conditions. Different parameters for improving

phenol detoxification (pH, temperature, bacterial consortia,

nutrient addition, aeration, etc.), are being optimised at

present (manuscript in preparation). Self-bioremediation

system based on the immobilisation of bacteria onto cork

particles is a valuable tool to perform the self-remediation of

wastewaters. In addition, some other industrial activities in

our regions also generate wastewaters with a high content in

phenolics, such as the extraction of olive oil (Cabrera et al.,

1996; Hanafi et al., 2011) or papermill bleaching (Rahman

et al., 2007). It could be presumed that a similar system may

be designed for other phenolics-containing residual waters,

either using cork particles as carriers for bacteria immobili-

sation or selecting other residues, specific of each industry, as

alternative carriers.

Acknowledgements

This work has been financed by the Spanish Ministry of

Education and Science (project number BIO-2009-7766) and

FEDER funding. Authors want to thank the Microscopy Service

of the CITIUS (University of Sevilla) and ResBioAgro, S.L. for

technical support with DGGE experiments.

Appendix. Supplementary data

Supplementary data related to this article can be found online

at doi:10.1016/j.watres.2011.12.038.

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